Amniotic fluid stem cell-conditioned media as a therapeutic adjunct in peripheral nerve injury repair: insights from a sciatic nerve mouse model

Introduction

Peripheral nerve injury presents a significant clinical challenge, often leaving patients with residual weakness, dysesthesias, and decreased function even after timely intervention. Treatment options span non-operative approaches, such as physical therapy and pain management, to surgical interventions (1). Direct end-to-end repair is recommended for acute peripheral nerve injuries with clean transections, where the tensionless approximation of nerve ends is possible. For defects greater than two centimeters, graft repair becomes necessary. This involves autografts harvested from other anatomical regions, nerve allografts, or tissue-engineered nerve grafts (TENGs) (2-7).

Autograft reconstruction remains the gold standard for peripheral nerve repair when direct end-to-end repair is not feasible. However, its application is limited by donor site availability and associated morbidity. Allografts offer a viable alternative but are constrained by cost and immunogenicity concerns. TENGs with biological adjuncts have emerged as a promising area of research to enhance the recovery of the injured peripheral nerve and its associated musculoskeletal unit.

Among the biological adjuncts, cell-based therapies have garnered attention for their regenerative potential. Amniotic fluid stem cells (AFS) have been increasingly investigated due to their pluripotent properties (8-15). However, studies have shown limited differentiation capabilities of implanted AFS (12,16), with current research suggesting that the regenerative effects of AFS are not solely reliant on direct cell differentiation but are mediated through their paracrine activity. AFS exerts paracrine effects by secreting bioactive factors that restore redox homeostasis and stimulate host stem cells, thereby promoting tissue regeneration. This understanding has led to the development of amniotic fluid stem cell-conditioned media (AFS-CM), which harnesses the paracrine potential of AFS without the risks associated with cellular therapies. By avoiding direct implantation of stem cells, AFS-CM circumvents the inflammatory response related to cell-based approaches, making it a safer and more practical alternative.

The paracrine effects of AFS-CM present a compelling opportunity for its application as a biological adjunct in peripheral nerve injury treatment. AFS-CM could enhance nerve regeneration and functional recovery by promoting redox balance and activating host stem cells.

This innovative approach offers the potential to complement existing surgical techniques, bridging the gap between current therapeutic limitations and optimal outcomes in peripheral nerve repair. There is a critical gap in knowledge on the use of the AFS-CM in peripheral nerve injury. This study uses a sciatic nerve mice injury model to assess the effects of AFS-CM in peripheral nerve recovery after injury. We hypothesize that AFS-CM will improve functional recovery, reduce muscle atrophy, and help improve peripheral nerve regeneration in CD1 mice after sciatic nerve transection. We present this article in accordance with the ARRIVE reporting checklist (available at https://atm.amegroups.com/article/view/10.21037/atm-25-17/rc).

Methods

Formulation of AFS conditioned media

AFS were procured from Nutech Inc. (Birmingham, AL, USA) and cultured in Dulbecco’s Modified Eagle Medium (DMEM; Corning, Product number 15-018-CV) supplemented with 20% fetal bovine serum (FBS) in T175 flasks (Thermo Fisher Scientific, Waltham, MA, USA). The cells were maintained under standard conditions until they reached approximately 80% confluence, at which point the medium was replaced with serum-free DMEM for 24 hours. Passage two to four AFS cultures were utilized in this study to ensure consistency.

Conditioned medium was collected and subjected to an initial centrifugation at 4 ℃, 3,000 g for five minutes to remove cellular debris. The supernatant was then purified using ultra-centrifugal filters with a 100 kDa semipermeable membrane (Thermo Fisher Scientific, PA, USA) at 4 ℃, 3,000 g for two hours. This additional step ensured the refinement of the AFS-conditioned medium (AFS-CM). The final supernatant was collected, aliquoted, and stored at −80 ℃ until use. Protein content was determined following the protocol provided by the manufacturer using a Bradford protein assay kit (Thermo Fisher Scientific, PA, USA, Product number A55866). Three independent samples from each batch of AFS-CM were analyzed to calculate the average protein concentration.

Silicone+ hydrogel and silicone+ hydrogel + AFS-CM formulation

Hydrogel was prepared using the proprietary HyStem kit and according to the manufacturer’s instructions. The hydrogel kit is based on cross-linking thiol-modified hyaluronan technology. Hyaluronic acid is a naturally occurring component of the extracellular matrix in connective, epithelial, and neural tissues. The System-HP kit includes Heprasil^® (thiol-modified hyaluronic acid and heparin), Gelin-S^® (thiol-modified gelatin), Extralink^® (PEGDA, polyethylene glycol diacrylate), and DG Water (degassed, deionized water). Briefly, Heprasil, Gelin-S, and Extralink were reconstituted with DG Water. Each vial was vortexed, after which Heprasil and Gelin-S were mixed in a 1:1 ratio. For silicone + hydrogel = AFS-CM formulation, AFS-CM was added to the hydrogel mix to obtain a 10 µg/mL concentration. The mean protein content of AFS-CM was calculated using a Bradford protein assay kit. Extralink vial was added to the Heprasil/Gelin-S mix and placed on 1 cm × 0.5 cm silicone tubes in a six-well plate.

Surgical procedure

The study protocol was approved by the Institutional Animal Care and Use Committee (IACUC) at Atrium Wake Forest Baptist Health (IACUC # A19-101), adhering to institutional guidelines for the care and use of animals. Thirty-six male CD1 mice were divided into three groups (n=12 per group): autograft, silicone + hydrogel (hydrogel group), and silicone + AFS-CM infused hydrogel (AFS-CM group). All mice underwent left sciatic nerve transection and repair surgery. Anesthesia was induced with 1.5–2.5% isoflurane in an induction chamber and maintained at 1.5% via a nose cone. The posterior left hind limb was shaved, cleansed with betadine scrub, and disinfected. Using aseptic techniques, a posterolateral incision was made, and the sciatic nerve was exposed through blunt dissection (Figure 1). A 1 cm segment of the nerve was excised to create a defect. In the hydrogel and AFS-CM groups, the defect was repaired by interposing a 1 cm silicone tubing filled with hydrogel or hydrogel + AFS-CM, secured with 10-0 nylon sutures using microsurgical techniques. In the Autograft group, the excised nerve segment was reversed and sutured back in place to bridge the defect. The muscle was closed with 5-0 Vicryl sutures, and the skin was closed using a combination of subdermal 5-0 Vicryl sutures and stainless-steel wound clips.

Figure 1 Sciatic nerve reconstruction with silicone + hydrogel and AFS-CM implant one-month post-surgery. AFS-CM, amniotic fluid stem cell-conditioned media.

Post-surgery, 1 mL of hydrogel (autograft and hydrogel groups) or hydrogel + AFS-CM (AFS-CM group) was injected subcutaneously into the experimental hind limb over the gastrocnemius and tibialis anterior muscles. This subcutaneous injection was repeated every two weeks. Postoperative analgesia was provided with buprenorphine (0.01 mg/kg, SC).

Gait analysis

The DigiGait Imaging System (Mouse Specifics Inc., Framingham, MA, USA) was used to evaluate motor function recovery in CD1 mice pre- and post-surgical procedures (11,17). This system utilizes a high-speed digital video camera, recording at 188 frames per second, to capture continuous imaging of the animal’s underside as it walks on a transparent treadmill. The system generates digital paw prints, which are then converted into dynamic gait signals, providing a temporal record of paw placement in relation to the treadmill belt. Motor functional recovery was assessed at two weeks, one month, and two months post-injury in rats that underwent sciatic autografts, silicone + hydrogel, or silicone + hydrogel and AFS-CM. Each mouse’s post-injury gait data was compared to its pre-injury baseline values, which were tabulated as the percent difference. This method allows for highly sensitive repeated measures analysis of variance, enabling the detection of subtle differences between treatment groups.

Electrophysiological assessment

Electrophysiological evaluations were performed using the Cadwell EMG Sienna Wave System. Mice were anesthetized at one month and two months with isoflurane, and both the regenerated sciatic nerve and the contralateral control nerve were surgically exposed. To assess nerve functionality, the tibial and peroneal branches distal to the reconstruction site were briefly stimulated to confirm plantar flexion and foot eversion, ensuring nerve viability before further analysis. For the electromyography recordings, the regenerated nerve was stimulated proximally near the suture site using a monopolar cathodic electrode delivering 1 mA of current, while the reference anode was placed on the thorax. The distance between the stimulating and recording electrodes was measured with a ruler to maintain consistency. Muscle responses were captured by placing recording electrodes into the medial and lateral gastrocnemius muscles, as well as the tibialis anterior muscle, on both the injured and uninjured limbs. Compound muscle action potentials (CMAP) were elicited with three consecutive stimulations, and the resulting signals were averaged to determine CMAP amplitude and delay. To minimize variability caused by anesthesia, CMAP results from the injured limb were normalized as a ratio to the contralateral, uninjured side.

Tissue harvesting and histomorphological analysis

The mice were humanely euthanized through an intracardiac injection of a saturated potassium chloride solution. Gastrocnemius and tibialis anterior muscles were harvested and weighed from both the experimental and contralateral sides. Harvested muscle was then fixed in 4% paraldehyde and later embedded in paraffin for tissue immunohistochemistry. Muscles harvested from experimental and contralateral limbs were sectioned and stained with muscle ring finger-1 (MURF-1) and malondialdehyde (MDA).

MURF-1 staining

Muscle RING-finger protein-1 (MuRF-1) is a muscle-specific E3 ubiquitin ligase that plays a central role in protein degradation and skeletal muscle atrophy. Its expression is a key marker of muscle wasting and atrophy (18,19). CD1 mice muscle tissues were fixed in 10% neutral-buffered formalin for 24 hours, processed, and embedded in paraffin. Sections (4–5 µm) were cut, mounted on charged slides, and dried at 60 ℃ for 1 hour. The sections were deparaffinized in xylene, rehydrated through graded ethanol, and subjected to antigen retrieval in sodium citrate buffer (pH 6.0) at 95 ℃ for 20 minutes. Endogenous peroxidase activity was blocked with 3% hydrogen peroxide for 10 minutes. To minimize nonspecific binding, slides were incubated in 10% normal goat serum for 1 hour. Tissue sections were incubated overnight at 4 ℃ with the primary antibody, MuRF-1 (ab183094, Abcam, Cambridge, MA, USA), diluted 1:200 in PBS with 1% BSA. After washing, a biotinylated secondary antibody was applied, followed by detection using the VECTASTAIN ABC Kit and DAB chromogen. Sections were counterstained with hematoxylin, dehydrated in graded ethanol, cleared in xylene, and mounted with a permanent medium.

MuRF-1 expression was evaluated under a brightfield microscope, with negative controls omitting the primary antibody to ensure specificity.

MDA staining

MDA is a marker of lipid peroxidation and oxidative stress, making it a key indicator of cellular damage in pathological conditions (20,21). To evaluate MDA expression in muscle tissues, CD1 mice samples were processed for immunohistochemistry.

Muscle tissues were fixed in 10% neutral-buffered formalin for 24 hours, processed, and embedded in paraffin. Sections (4–5 µm thick) were cut, mounted on charged slides, and dried at 60 ℃ for 1 hour. Sections were deparaffinized in xylene, rehydrated through graded ethanol, and subjected to antigen retrieval in citrate buffer (pH 6.0) at 95 ℃ for 20 minutes. Endogenous peroxidase activity was blocked by treating the sections with 3% hydrogen peroxide for 10 minutes. To prevent nonspecific antibody binding, sections were incubated with 10% normal goat serum in PBS for 1 hour at room temperature. The sections were incubated overnight at 4 ℃ with the primary antibody, anti-malondialdehyde (ab6463, Abcam), diluted 1:100 in PBS containing 1% BSA. After three PBS washes, a biotinylated secondary antibody was applied, and signal detection was performed using the VECTASTAIN ABC Kit. The chromogen 3,3'-diaminobenzidine (DAB) was used to visualize staining. Sections were counterstained with hematoxylin to identify nuclei, dehydrated in graded ethanol, cleared in xylene, and mounted with a permanent medium. Staining was evaluated using brightfield microscopy. Negative controls, omitting the primary antibody, were included to ensure the specificity of MDA staining.

Harvested muscles were stained with α-bungarotoxin at 1:2,000 (Thermo Fisher, NY, USA) to visualize neuromuscular junction (NMJ) replenishment following nerve injury and repair.

Nerve

Both the experimental and contralateral nerves, along with the proximal and distal nerve stumps, were collected. Nerve samples were fixed in either 4% paraformaldehyde or 2% osmium tetroxide, followed by dehydration and embedding in paraffin or resin. Serial 5 µm sections were obtained 1 mm distal to the distal suture within the nerve injury site. This was done to investigate the regenerating nerve at its reconstruction interface. Sections were stained with toluidine blue and analyzed under Zeiss light and electron microscopes (Thornwood, NY, USA) at magnifications of 200× and 3,700×. Regenerated axons were quantified using Python software. The G-ratio was calculated as the ratio of non-myelinated axonal diameter to the total axonal diameter (Figure 2).

Figure 2 Measurement of the G-ratio. A schematic representation illustrating the calculation of the G-ratio, defined as the ratio of the inner axonal diameter (d) to the total fiber diameter (D), which includes both the axon and the surrounding myelin sheath.

Mice grouping, timing, and treatment allocation

Thirty-six male CD1 mice were used for in vivo experimentation (Figure 3). Mice underwent surgical sciatic nerve transection and either underwent sciatic nerve apposition (Autograft; n=12), reconstruction with silicone infused with hydrogel (Hydrogel group; n=12), or reconstruction with silicone and hydrogel + AFS-CM (AFS-CM; n=12). All mice underwent subcutaneous injection of 1 ml of hydrogel (Autograft and Hydrogel group) vs. Hydrogel + AFS-CM (AFS-CM) during the initial surgical procedure and every two weeks after the procedure. Mice were humanely euthanized at two-week, one-month, and two-month time points.

Figure 3 Experimental design for time points and group allocation: schematic representation showing the allocation of mice into three experimental groups (AFS-CM, hydrogel, and autograft) at each of the three-time points (two weeks, one month, and two months). Each group consisted of four mice (N=4) at each time point. AFS-CM, amniotic fluid stem cell-conditioned media.

Statistical analysis

Continuous data were analyzed using a one-way analysis of variance (ANOVA). Post-hoc analysis was conducted using Tukey’s Honestly Significant Difference (HSD) test to evaluate multiple comparisons of mean differences between groups. Statistical significance was defined as a P value of less than 0.05. All statistical tests were two-sided and executed using Python’s SciPy library for statistical analysis and the Pandas library for efficient data manipulation.

For image-based analysis, the assessment of G-ratio, MURF-1, MDA, and alpha-bungarotoxin staining utilized Python’s OpenCV library for image processing and sci-kit-image for advanced feature extraction and analysis. Data visualization was executed through the Matplotlib and Seaborn libraries.

Results

Analysis of gastrocnemius muscle weight differences across experimental groups

Harvested muscles were obtained and weighed before paraffin fixation. The bilateral gastrocnemius muscles from each mouse were weighed three times, and the average of these measurements was calculated. The percentage weight difference between the left and right gastrocnemius muscles was tabulated. Statistical analysis revealed a significant difference in the left gastrocnemius muscle weights among the AFS-CM, Autograft, and Hydrogel groups at two weeks and one month (Figure 4). Post-hoc Tukey’s honestly significant difference (HSD) tests identified a significant difference between the left gastrocnemius muscle mean weights of the AFS-CM and Hydrogel groups at one month (P=0.02; Table 1).

Figure 4 Percent normalized change differences in gastrocnemius muscle weights are presented for the AFS-CM, autograft, and hydrogel groups at two weeks (A), one month (B), and two months (C). Statistically significant differences between groups are highlighted with corresponding P values. Error bars represent the 95% confidence intervals. Mean group differences were analyzed using analysis of variance, followed by post-hoc Tukey’s HSD test to evaluate multiple group comparisons. *, a Tukey’s HSD P value of <0.05. AFS-CM, amniotic fluid stem cell-conditioned media; HSD, honestly significant difference.

Gait analysis

DigiGait analysis results were obtained at one month and two months. At one month, no significant differences were observed in the ataxia coefficient (P=0.30) or midline distance (P=0.09) among the Hydrogel, AFS-CM, and Autograft groups. There was a significant difference in ataxia coefficient between the groups at two months, with post-hoc analysis revealing a significantly lower ataxia coefficient percent difference in the Autograft group when compared to the Hydrogel group (P=0.01). Additionally, the midline distance demonstrated a trend toward significance (P=0.053), favoring the Autograft group, suggesting enhanced spatial coordination (Figure 5A). Overlap distance at one month showed no significant differences among the groups (P=0.10). However, at two months, the Hydrogel group displayed a significantly higher percent difference in overlap distance than the AFS-CM and Autograft groups (P<0.001), highlighting inferior stride recovery in the Hydrogel group. For the Sciatic Functional Index (SFI), no significant differences were found at one month (P=0.057) or two months (P=0.18) (Figure 5B).

Figure 5 Percent differences in (A) ataxia coefficient (top row) and midline distance (bottom row) and (B) in overlap distance (top row) and SFI (SFIbottom row) among the hydrogel, AFS-CM, and autograft treatment groups in the animal cohort at one month (left column) and two months (right column) post-treatment. Statistically significant differences between groups are highlighted with corresponding P values. Error bars represent the 95% confidence intervals. Mean group differences were analyzed using analysis of variance, followed by post-hoc Tukey’s HSD test to evaluate multiple group comparisons. *, a Tukey’s HSD P value of <0.05. AFS-CM, amniotic fluid stem cell-conditioned media; HSD, honestly significant difference; SFI, sciatic functional index.

Electromyographical analysis

Electromyographical analysis showed no mean percent difference in cAMP and latency for both gastrocnemius and tibial anterior muscle between experimental and control limbs. Our study revealed no differences in nerve conduction latency for the tibialis anterior and gastrocnemius muscle experimental limb at one and two months (P>0.05; Figure 6A). Similarly, we found no difference in nerve conduction amplitude for the tibialis anterior and gastrocnemius muscle experimental limb at one and two months (P>0.05; Figure 6B).

Figure 6 Percent difference in EMG (A) latency for the left tibialis anterior (top row) and left gastrocnemius (bottom row) muscles and (B) EMG amplitude for the left gastrocnemius (top row) and left tibialis anterior (bottom row) muscles in the animal cohort at one month (left column) and two months (right column) post-treatment. Groups compared include hydrogel, AFS-CM, and autograft treatment groups. P values indicate no significant differences between treatment groups at any time point. Error bars signify a 95% confidence interval. Analysis of variance test assessed mean group differences. AFS-CM, amniotic fluid stem cell-conditioned media; EMG, electromyography.

Histomorphological analysis

The histomorphological analysis demonstrated a lower mean area percentage of muscle that stained positive with MURF-1 antibody for the AFS-CM group at the two-month mark compared to the Hydrogel group (P=0.02; Figure 7). There was no difference between the groups at two weeks (P=0.36). We found no difference in the mean area percentage of muscle that stained positive with MDA antibody between the groups for both the two-week (P=0.26) and the two-month (P=0.44) time points (Figure 8).

Figure 7 Comparison of mean area percentage staining positive for MURF-1 across groups (AFS-CM, autograft, and hydrogel) and time points (two months and two weeks). The bar graph illustrates the significantly higher percentage of MURF-1 staining in the Autograft group at two months compared to the AFS-CM group (P=0.02). No significant differences were observed between groups at the two-week time point (P=0.36). Error bars signify a 95% confidence interval. Analysis of variance test assessed mean group differences. A post-hoc Tukey’s HSD was performed to assess multiple comparisons of mean differences between groups. *, a post hoc Tukey’s HSD P value of <0.05. AFS-CM, amniotic fluid stem cell-conditioned media; HSD, honestly significant difference.

Figure 8 Comparison of mean area percentage staining positive for MDA across groups (hydrogel, AFS-CM, and autograft) and time points (two months and two weeks). The bar graph shows no statistically significant differences in MDA staining between the groups at either point, with P=0.44 at two months and P=0.26 at two weeks. Error bars represent the standard deviation. Error bars signify a 95% CI—analysis of variance test assessed mean group differences. AFS-CM, amniotic fluid stem cell-conditioned media; CI, confidence interval; MDA, malondialdehyde.

G-ratio and NMJ

G-ratio was calculated for both the two-week and two-month mouse cohorts. Our findings revealed no significant difference between mice belonging to the AFS-CM, Autograft, and Hydrogel cohort at both two weeks (P=0.37) and two months (P=0.46) (Figure 9A-9C). Alpha-bungarotoxin-stained muscle tissue was analyzed to assess NMJ. Our findings revealed no difference in the ratio of alpha-bungarotoxin staining NMJ area to the cross-sectional experimental muscle area between the AFS-CM, autograft, and hydrogel groups at the two-week cohort (P=0.28). However, our study found a difference between the three groups at two months, demonstrating an increased ratio of alpha-bungarotoxin staining NMJ area to the cross-sectional experimental muscle area at two months in favor of the AFS-CM group (P=0.03; Figure 10).

Figure 9 Representative histologic and quantitative assessment of peripheral nerve myelination following treatment after sciatic nerve injury. (A) Toluidine blue–stained cross-sections of injured sciatic nerves demonstrate myelinated axons at two months following treatment (magnification 20×). (B,C) Quantitative analysis of mean G-ratios of myelinated fibers at two weeks and two months shows comparable myelination across treatment groups, with no statistically significant differences detected at either time point. Error bars represent 95% confidence intervals. Statistical comparisons were performed using analysis of variance. AFS-CM, amniotic fluid stem cell–conditioned media; CI, confidence interval.

Figure 10 The ratio of NMJ area to cross-sectional muscle area as identified by alpha-bungarotoxin staining. (A,B) Two-week and two-month time intervals, respectively. Statistically significant differences between groups are highlighted with corresponding P values. Error bars represent the 95% confidence intervals. Mean group differences were analyzed using analysis of variance, followed by post-hoc Tukey’s HSD test to evaluate multiple group comparisons. *, a Tukey’s HSD P value of <0.05. ANOVA, analysis of variance; AFS-CM, amniotic fluid stem cell-conditioned media; HSD, honestly significant difference; NMJ, neuromuscular junction.

Discussion

Traumatic injury to peripheral nerves remains an issue in the United States. Even with timely intervention, many patients will still suffer significant morbidity, incomplete nerve recovery, and muscle weakness (22,23). While surgical intervention can improve patient outcomes, peripheral nerve regeneration can be stifled by many factors. These factors include imbalance in inflammatory homeostasis, patient age, and injury severity. Biologic adjuncts such as AFS therapy have been explored with increased interest due to their non-immunogenic properties and relatively low-cost (12,24). AFSs are easily harvested from the placenta soon after delivery and can be stored for later use. Their regenerative capabilities and immunoregulatory properties can facilitate tissue recovery after injury. Our study explored the role of adjunctive AFS conditioned media in a sciatic nerve injury model entailing CD1 male mice. Our findings reveal higher muscle mass, improved gait, reduced MURF-1 expression, and a higher ratio of muscle cross-sectional area stained with alpha bungarotoxin for mice belonging to the AFS-CM group compared to Hydrogel control.

An injured peripheral nerve undergoes Wallerian degeneration (25) soon after nerve transection. Critical to this process is the activity of the peripheral nerve Schwann cells, which undergo de-differentiation to repair Schwann cell phenotype soon after traumatic injury. The repair Schwann cell phenotype facilitates the clearing of the distal debris via phagocytosis and activation of the innate immune system. After clearance, repair Schwann cells form a hollow tube, which re-differentiates into its myelinating phenotype for final remyelination and regeneration of the injured peripheral nerve (26). Critical to this process is redox homeostasis, as prolonged inflammation can stifle peripheral nerve regeneration and impair the re-differentiation of Schwann cells back to their myelinating phenotype (27-29). AFSs are pluripotent cells obtained from the maternal amnion. Its regenerative power emerges from its immunomodulatory capability and ability to replenish cellular proteins necessary for tissue regeneration (30-34). As such, its use has been explored as a therapeutic adjunct due to its regenerative capabilities.

Our findings demonstrate that denervated gastrocnemius muscles of mice treated with AFS-CM exhibited higher wet muscle mass and a lower proportion of MURF-1-positive stained cross-sectional muscle area compared to untreated groups. Skeletal muscle degeneration and atrophy rapidly follow denervation, as neuronal input from healthy peripheral nerves is essential for maintaining calcium signaling critical for muscle contraction and protein synthesis. The loss of this input shifts muscle homeostasis toward protein degradation, primarily driven by the ubiquitin-proteasome pathway (35,36). Additionally, the upregulation of atrophy-related genes, such as MURF-1 and atrogin-1, accelerates muscle-specific protein breakdown. A significant contributor to this degradation process is the increased generation of reactive oxygen species (ROS). Persistent ROS production impairs the activity of muscle satellite cells, which are vital for muscle regeneration following injury (37-40). In our study, the AFS-CM-treated mice received direct administration of AFS-CM to the distal muscle groups of the surgically transected sciatic nerve, as described in the methods section. The regenerative and immunomodulatory properties of AFS-CM may have offered protection against denervation-induced muscle atrophy compared to other groups. Interestingly, the groups showed no significant difference in MDA expression. MDA, an end-product of prolonged oxidative stress found in cell membranes, is a marker for oxidative damage in affected cells (21,41-43). This finding suggests that while AFS-CM may mitigate muscle atrophy, it does not significantly alter oxidative stress levels in denervated muscle tissue as measured by MDA staining.

Treadmill-based computerized gait analysis captures the locomotion of running animals to obtain dynamic gait signals. This method allows for accurate gait kinematic analysis of each animal, offering insights into nerve recovery as observed through gait parameters. Gait parameters, such as overlap distance and ataxia coefficient, measure gait coordination and recovery. Overlap distance measures the distance between the placement of the forepaw and the hind paw on the same side of the body during walking, whereas the ataxia coefficient measures variability in paw placement during animal gait. Both parameters are measures of animal coordination during gait. Mice in our AFS-CM cohort demonstrated improved recovery in overlap distance and ataxia coefficient at two months compared to mice in our Hydrogel cohort, consistent with improvements in gait coordination. Furthermore, we found an increase in alpha-bungarotoxin-stained tissue among the AFS-CM-treated mouse cohort compared to the Hydrogel cohort. Preservation of the NMJ is a critical factor in preserving skeletal muscle and restoring function and coordination after traumatic peripheral nerve injury (44). As such, our findings suggest that the gait recovery witnessed in our study may be due to AFS-CM’s ability to recover NMJs after peripheral nerve injury.

Our study revealed no difference in comparison between the groups. Furthermore, we found no difference in sciatic nerve remyelination as measured by G-ratio between the groups. Our findings differ from previous studies exploring the effect of AFSs on peripheral nerve regeneration. Ma et al. explored the effect of AFSs in combination with acellular nerve allograft (AFS + ANA) for long-gap nerve repairs in a sciatic nerve injury rat model (11). The authors revealed the mice undergoing AFS + ANA demonstrated greater G-ratio for the AFS + ANA group at four months post-sciatic nerve transection. Still, the therapeutic effects of AFS-CM are noted in its ability to maintain wet muscle mass and NMJs, leading to improved gait coordination in our mouse model. While no clear signs of immune rejection were observed after administering human AFS-CM into mice, there remains potential for a xenogeneic immune response. Future studies should investigate the immunogenicity of AFS-CM more thoroughly.

Our study is not without limitations. We did not identify the optimal therapeutic intervention time for treatment groups after sciatic nerve transection. The time-point for subdermal injections of AFS-CM versus hydrogel was arbitrarily chosen concordant with the time for mice harvest. However, this interval administration may not demonstrate the optimal time for treatment delivery among the different groups. As such, the actual therapeutic effect of AFS-CM may differ based on timing. Furthermore, our study was performed on male CD1 mice. Thus, we are unable to explore the role of sex in the therapeutic effect of AFS-CM on peripheral nerve injury. Our study observed an increased cross-sectional area of muscle tissue staining positive for alpha-bungarotoxin in mice treated with AFS-CM, indicating a higher presence of NMJs. However, it’s important to note that alpha-bungarotoxin staining identifies nicotinic acetylcholine receptors at NMJs but does not distinguish between functional and non-functional synapses (45). Still, this study is one of the first to explore the role of AFS-CM on peripheral nerve recovery in a sciatic nerve injury model.

Conclusions

Our study highlights the therapeutic potential of AFS-CM in promoting recovery following peripheral nerve injury. AFS-CM treatment demonstrated the ability to mitigate muscle atrophy, preserve NMJs, and improve functional outcomes, such as gait coordination, underscoring its regenerative and immunomodulatory properties. While we observed increased alpha-bungarotoxin staining indicating greater NMJ recovery, this technique does not differentiate between functional and non-functional junctions, highlighting an area for further investigation. Additionally, no significant differences in oxidative stress markers or sciatic nerve remyelination were observed, suggesting AFS-CM’s therapeutic effects may be localized to muscle preservation and functional recovery rather than direct nerve repair. The limitations of our study, including the arbitrary timing of treatment and the exclusive use of male CD1 mice, underscore the need for future research to optimize treatment protocols, assess sex-specific responses, and evaluate the functional integrity of preserved NMJs. Despite these challenges, our findings provide a foundation for further exploration of AFS-CM as a promising adjunct for peripheral nerve regeneration, with the potential to improve patient outcomes in clinical applications.

Acknowledgments

None.

Footnote

Reporting Checklist: The authors have completed the ARRIVE reporting checklist. Available at https://atm.amegroups.com/article/view/10.21037/atm-25-17/rc

Data Sharing Statement: Available at https://atm.amegroups.com/article/view/10.21037/atm-25-17/dss

Peer Review File: Available at https://atm.amegroups.com/article/view/10.21037/atm-25-17/prf

Funding: None.

Conflicts of Interest: All authors have completed the ICMJE uniform disclosure form (available at https://atm.amegroups.com/article/view/10.21037/atm-25-17/coif). The authors have no conflicts of interest to declare.

Ethical Statement: The authors are accountable for all aspects of the work in ensuring that questions related to the accuracy or integrity of any part of the work are appropriately investigated and resolved. The study protocol was approved by the Institutional Animal Care and Use Committee (IACUC) at Atrium Wake Forest Baptist Health (IACUC # A19-101), adhering to institutional guidelines for the care and use of animals.

Open Access Statement: This is an Open Access article distributed in accordance with the Creative Commons Attribution-NonCommercial-NoDerivs 4.0 International License (CC BY-NC-ND 4.0), which permits the non-commercial replication and distribution of the article with the strict proviso that no changes or edits are made and the original work is properly cited (including links to both the formal publication through the relevant DOI and the license). See: https://creativecommons.org/licenses/by-nc-nd/4.0/.

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