Canine Adipose MSC-Derived Exosomes Ameliorate Skeletal Muscle Injury in Mice

Simple Summary

Muscle injuries in dogs frequently lead to long-term complications such as muscle atrophy, with healthy muscle fibres being replaced by fatty or scar tissue. These changes will diminish the dog’s quality of life. However, current treatment options are limited. The aim of this study was to investigate whether the canine adipose mesenchymal stem cell-derived exosomes (cADMSC-Exos) possess therapeutic potential for muscle injury and dysfunction. cADMSCs and exosomes were successfully obtained and then identified. By establishing an in vitro model of C2C12 cell atrophy and a mouse muscle injury model, we demonstrated that cADMSC-Exos can ameliorate the adverse conditions of muscle atrophy, fibrosis, and fatty infiltration, exerting beneficial tissue-repairing effects. This lays the foundation for future research into the application of ‘cell-free’ medical technology for treating canine muscular disorders.

Abstract

Severe skeletal muscle injury in dogs can result in muscle atrophy, fibrotic remodeling, and fat accumulation, leading to skeletal muscle dysfunction and impaired quality of life. However, there is currently no effective treatment available. This study aims to investigate the potential of canine adipose mesenchymal stem cell-derived exosomes (cADMSC-Exos) as a novel acellular therapy for the repair of muscle atrophy and injury. cADMSCs and their derived exosomes were isolated and characterized. A dexamethasone-induced C2C12 myotube atrophy model was established to evaluate the effects of cADMSC-Exos on muscle atrophy by assessing myotube morphology and the expression of atrophy-related factors. Subsequently, a glycerol-induced mouse muscle injury model was constructed. Through histological analysis and Western blot, the efficacy and safety of cADMSC-Exos in vivo were systematically evaluated. Results indicated that cADMSC-Exos demonstrated significant anti-atrophic activity in both two models, ameliorating skeletal muscle atrophy and the upregulation of muscle RING finger 1 (MuRF1) and muscle atrophy F-box (Atrogin-1) (p < 0.05), consistent with morphological alterations. Moreover, cADMSC-Exos markedly alleviated fibrosis and fatty infiltration in injured muscle tissue (p < 0.0001). Overall, these findings indicate that cADMSC-Exos promote muscle repair and attenuate pathological remodeling by modulating the local microenvironment and protein expression, highlighting their potential as a therapeutic strategy for muscular disorders.

1. Introduction

Skeletal muscle constitutes the predominant tissue type in dogs, representing an estimated 44–57% of their overall body mass [1]. Skeletal muscle, together with the skeleton and joints, forms the structural basis for canine movement. Proper muscle structure and function are critical for physical performance and overall health. Muscle disorders are highly prevalent in veterinary medicine and may arise from acute trauma, overuse injuries associated with sporting or working activities, as well as chronic degenerative conditions related to metabolic disorders, such as cachexia, or aging, such as sarcopenia [2,3,4,5]. Severe muscle injury typically initiates a complex cascade of pathophysiological events, including impaired tissue regeneration, persistent inflammation, fibrosis, and fatty infiltration, which ultimately result in compromised muscle function [6,7,8]. However, treatment approaches for muscle injuries and dysfunction in veterinary clinical practice, including medication management, surgery, and rehabilitation therapy, present certain limitations [9]. The use of medications such as corticosteroids may not necessarily improve the condition and carries certain side effects. While surgical intervention combined with rehabilitation therapy can yield immediate improvements in gait, there remains a possibility of recurrence [10]. Consequently, there is currently a lack of intervention strategies capable of effectively promoting muscle repair and regeneration, necessitating the urgent exploration of novel therapies.

Mesenchymal stem cells (MSCs) have exhibited potential for tissue regeneration and repair in the treatment of various canine diseases [11,12]. Adipose-derived MSCs have demonstrated preliminary efficacy in treating semitendinosus myopathy in working dogs, with no subsequent recurrence [9]. However, the clinical application of MSC-based therapies faces limitations due to concerns over tumorigenicity, biological variability, and difficulties in treatment dose standardization [13,14]. Mesenchymal stem cell-derived exosomes (MSC-Exos), as extracellular vesicles measuring 30–150 nm, retain the immunomodulatory and regenerative functions of their parent cells while circumventing the risks associated with cell transplantation. Consequently, they have emerged as a highly regarded cell-free alternative therapy [15,16]. Notably, owing to their low immunogenicity and conserved cross-species bioactivity, exosomes can be evaluated in established murine models, thereby facilitating preclinical assessment and translational development [17,18,19].

Despite previous studies indicating that MSC-Exos can improve muscle atrophy in mouse models by promoting mitochondrial biogenesis, enhancing protein synthesis, and inhibiting apoptosis, limited evidence exists regarding the regenerative capacity of canine MSC-Exos in muscle repair and regeneration [20,21]. Canine adipose mesenchymal stem cells (cADMSCs) are readily obtainable from surgical waste materials such as ovariohysterectomy (OHE) procedures. They present the advantages of minimal ethical controversy and robust proliferative capacity, making them an ideal cellular source for the preparation of therapeutic exosomes [22,23]. Canine adipose mesenchymal stem cell-derived exosomes (cADMSC-Exos) exhibit considerable therapeutic potential owing to their rich content of regulatory factors, signaling molecules, genes, and proteins [24]. Given that human, rodent and canine skeletal muscle physiology are highly conserved, the evaluation of cADMSC-Exos in conventional preclinical animal models is a necessary step toward future clinical application in veterinary medicine [25,26].

Therefore, to evaluate the therapeutic potential of cADMSC-Exos and address gaps in this field, this study employed two complementary pathological models for validation: A dexamethasone-induced C2C12 myotube atrophy in vitro model, simulating metabolic or steroid-induced muscle wasting; and a glycerol-induced mouse skeletal muscle injury in vivo model, reproducing key pathological features such as myofibrillar atrophy, fibrosis, and fatty infiltration following severe injury [27,28]. This study aims to integrate in vivo and in vitro evidence, thereby providing robust experimental support for the use of cADMSC-Exos as a novel therapeutic agent for canine skeletal muscle injury.

2. Materials and Methods

2.1. Isolation and Culture of cADMSCs

cADMSCs were isolated from adipose tissue collected adjacent to the uterus and uterine horns of healthy female dogs (<6 years old) undergoing ovariohysterectomy (OHE) at the Veterinary Teaching Hospital of China Agricultural University. All tissue samples from surgical discards were obtained under sterile conditions and approved by the China Agricultural University Animal Welfare and Animal Experimental Ethical Inspection Committee under approval number AW03106202-2-1.

Adipose tissue was minced into small fragments (~1 mm³) and subjected to enzymatic digestion with 1 mg/mL type I collagenase (Solarbio, Beijing, China) at 37 °C for 1 h under constant agitation. To refine the digest, the suspension was passed through a 100 μm cell strainer, followed by centrifugation at 1000 rpm for 5 min. After discarding the supernatant, the obtained pellet was reconstituted in phosphate-buffered saline (PBS). Subsequently, the cells were incubated with an erythrocyte lysis reagent (Solarbio) for 5 min at ambient temperature to eliminate red blood cells. Following another centrifugation step, the pelleted cells were resuspended in complete growth medium and seeded into culture flasks. The complete growth medium consisted of α-minimum essential medium (α-MEM; Gibco, Thermo Fisher Scientific, Carlsbad, CA, USA) containing 10% fetal bovine serum (FBS; Bdbio, Hangzhou, China) and 1% penicillin–streptomycin (Biosharp, Beijing, China). Cells were cultured at 37 °C in a humidified atmosphere of 5% CO₂, with medium changes performed every 2–3 days. Upon reaching approximately 80% confluence, cells were passaged for subsequent experiments.

2.2. Multipotent Differentiation

Multipotency of cADMSCs was assessed by the induction with commercial differentiation kit. Osteogenic and adipogenic differentiation media were sourced from Cyagen Biosciences (Guangzhou, China), while chondrogenic induction medium was obtained from Procell (Wuhan, China). After 21–24 days of induction, differentiation was confirmed via lineage-specific staining: Alizarin Red S for osteogenic, Oil Red O for adipogenic, and Alcian Blue for chondrogenic differentiation.

2.3. Flow Cytometric Analysis

Flow cytometric analysis was performed to characterize the surface marker expression of cADMSCs using an LSRFortessa flow cytometer (BD Biosciences, San Jose, CA, USA). Passage 3 cADMSCs were stained with phycoerythrin (PE)-conjugated monoclonal antibodies or matched isotype controls at 37 °C for 30 min. The following antibodies were used: anti-CD29-PE (303003; BioLegend, San Diego, CA, USA), anti-CD44-PE (103007; BioLegend), anti-CD90-PE (12-5900-41; eBioscience, San Diego, CA, USA), anti-HLA-DR-PE (307605; BioLegend), IgG1-PE isotype control (400111; BioLegend), IgG2a-PE isotype control (400211; BioLegend), and IgG2b-PE isotype control (4006071; BioLegend). After washing with chilled PBS to remove unbound antibodies, a total of 2.5 × 10⁵ cells per sample were acquired and analyzed using the flow cytometer.

2.4. Cell Growth Curve and Population Doubling Time

cADMSCs from passages 2, 5, and 8 were seeded at a density of 1 × 10⁴ cells per well in 24-well plates and cultivated under routine conditions. Cell numbers were determined daily for 8 days using a hemocytometer. Growth curves were generated by plotting cell number against culture time. Population doubling time (PDT) was calculated using cell count data from the logarithmic growth phase (Day 1–4) based on an exponential growth model. Nonlinear regression analysis was conducted with GraphPad Prism software, and PDT was determined according to the formula:

$$\mathrm{P}\mathrm{D}\mathrm{T}=[\mathrm{T}\times (\mathrm{l}\mathrm{n}2)]/[\mathrm{l}\mathrm{n}(\mathrm{N}\mathrm{e}/\mathrm{N}\mathrm{b})]$$

(1)

where Nb is the initial cell count, Ne is the final cell count, and T indicates the time interval between the two measurements.

2.5. Isolation and Identification of cADMSC-Exos

cADMSCs were maintained under standard culture conditions. Within 24 h, the complete growth medium was replaced with conditional medium containing 10% exosome-depleted fetal bovine serum for subsequent exosome harvesting. After 48 h of incubation, the conditioned medium was collected and subjected to sequential centrifugation and filtration: initially centrifuged at 300× g for 10 min, succeeded by centrifugation at 3000× g for 30 min to exclude dead cells and cellular debris. The resulting supernatant was passed through a 0.22 μm membrane filter to remove particles exceeding 200 nm in size. Exosomes were subsequently isolated from the clarified supernatant using a membrane affinity-based method combined with magnetic bead enrichment, following the manufacturer’s directions (Omiget, Beijing, China). In brief, the filtered supernatant was combined with a proprietary binding buffer and subjected to incubation under mild rotary conditions to facilitate the binding of exosomes to the affinity matrix. Bead-bound exosomes were collected using a magnetic rack, washed with PBS, and eluted. The entire isolation procedure was performed at 4 °C. Following purification, cADMSC-Exos were preserved at −80 °C for further analysis.

2.6. Transmission Electron Microscopy (TEM)

The morphological features of cADMSC-Exos were analyzed by TEM (Hitachi, Ltd., Chiyoda, Tokyo, Japan). A 10 μL aliquot of exosome suspension was applied onto a copper grid and incubated for 2–10 min to allow adsorption. Residual liquid was blotted with filter paper, after which the sample was negatively stained with 10 μL of 2% phosphotungstic acid for 1 min. After air drying, samples were examined with a transmission electron microscopy (Hitachi, Ltd.) at an accelerating voltage of 80 kV.

2.7. Nanoparticle Tracking Analysis (NTA)

NTA was utilised to characterize the concentration and size distribution of cADMSC-Exos. Briefly, exosome suspensions were illuminated by a laser beam, and the scattered light from individual particles was recorded to analyze their Brownian motion. Measurements were conducted using a ZetaView PMX 110 instrument (Particle Metrix GmbH, Meerbusch, Germany) equipped with a 405 nm laser. Purified exosomes were diluted with PBS to achieve a final particle concentration of 1 × 10⁷ to 1 × 10⁹ particles per millilitre, after which they were subjected to analysis. Particle size, concentration, and motion trajectories were analyzed using the manufacturer’s software.

2.8. Culture and Differentiation of C2C12 Cell

The C2C12 mouse myoblast cell line was kindly provided by Prof. Yanjun Dong of the Key Laboratory of Veterinary Basic Sciences at China Agricultural University. This cell line was originally purchased from the Peking Union Medical College Cell Resource Center (National Biomedical Experimental Cell Resource Bank, Beijing, China). Cells were cultured in Dulbecco’s Modified Eagle Medium (DMEM; ServiceBio, Wuhan, China), supplied with 10% fetal bovine serum and 1% penicillin–streptomycin. Upon reaching approximately 80–90% confluence, the growth medium was replaced with differentiation medium consisting of DMEM supplemented with 2% horse serum (Gibco) to initiate myogenic differentiation. Cells were cultured under differentiation conditions for 5–7 days until the formation of mature multinucleated myotubes was observed.

2.9. Dexamethasone-Induced In Vitro Muscle Atrophy Model and Exosome Treatment

Based on previously published studies, an in vitro muscle atrophy model was established by treating differentiated C2C12 myotubes with 10 μM DEX (Lablead, Beijing, China) for 24 h [21]. The differentiated myotubes were assigned to three experimental groups: a control (CON) group, cultured in serum-free DMEM supplemented with 1% penicillin–streptomycin; a dexamethasone-treated (DEX) group, receiving 10 μM dexamethasone; and a combined treatment (DEX + Exos) group, in which myotubes were co-treated with 10 μM dexamethasone and cADMSC-Exos. All experiments were conducted under standardized conditions at 37 °C with 5% CO₂ over a 24 h period.

2.10. Cell Viability Assay

Cell viability was evaluated using the Cell Counting Kit-8 (CCK-8; Beyotime, Shanghai, China) in accordance with the manufacturer’s protocol. C2C12 myoblasts were plated in 96-well plates and exposed for 24 h to 10 μM dexamethasone, either administered alone or in combination with escalating doses of cADMSC-Exos (0, 25, 50, 100, or 200 μg/mL). Following the addition of CCK-8 reagent and incubation at 37 °C for 1 h, absorbance at 450 nm was recorded with a microplate reader for viability assessment.

2.11. Immunofluorescence Staining and Myotube Morphology Analysis

Differentiated C2C12 myotubes were subjected to 10 μM dexamethasone treatment for 24 h in the presence or absence of 50 μg/mL cADMSC-derived exosomes. Cells were then fixed, permeabilized, and incubated overnight at 4 °C with an anti-myosin heavy chain (MyHC) antibody (1: 300, sc-376157, Santa Cruz, Dallas, TX, USA), followed by incubation with a fluorescent secondary antibody and DAPI counterstaining. Images were captured using a fluorescence microscope (OPTEC, Chongqing, China). Myotube diameter, MyHC fluorescence intensity, and fusion index were quantified using ImageJ software (1.54g). Myotube diameter was assessed in no fewer than 50 myotubes derived from a minimum of 10 randomly selected microscopic fields per experimental group. The fusion index was derived from the ratio of nuclei within MyHC-positive multinucleated myotubes to the total number of nuclei, quantified across at least 10 randomly chosen microscopic field.

2.12. Western Blot

Total protein was harvested using RIPA lysis buffer supplemented with protease inhibitors (both from Solarbio), with concentrations subsequently determined via a bicinchoninic acid (BCA) assay kit (Beyotime). For each sample, 20 μg of protein was resolved by electrophoresis and then electroblotted onto polyvinylidene fluoride (PVDF) membranes (Epizyme, Shanghai, China). After a 1–2 h blocking period in 5% nonfat milk at ambient temperature, the membranes were probed with specific primary antibodies overnight at 4 °C. This was followed by a 1 h incubation with appropriate secondary antibodies. Finally, protein signals were captured using enhanced chemiluminescence reagents (Solarbio) and their optical densities were analyzed with ImageJ software.

The following primary antibodies were utilized: anti-TSG101 (1:1000, ab125011, Abcam, Cambridge, UK), anti-Alix (1:1000, ab186429, Abcam), anti-HSP70 (1:1000, ab181606, Abcam), anti-Calnexin (1:500, 10427-2-AP, Proteintech, Wuhan, China), anti-GAPDH (1:10,000, 60004-1-Ig, Proteintech), anti-Atrogin-1 (1:1500, bsm-5445R, Bioss), anti-MuRF1 (1:4000, 55456-1-AP, Proteintech). The secondary antibodies were anti-mouse IgG (1:10,000, bs-0296G-HRP, Bioss, Beijing, China) and anti-rabbit IgG (1:10,000, bs-0295G-HRP, Bioss).

2.13. RNA Extraction and Quantitative Real-Time Polymerase Chain Reaction (qRT-PCR)

Total RNA extraction was carried out with RNA-easy Isolation Reagent (Vazyme, Nanjing, China) following the manufacturer’s protocol. Complementary DNA (cDNA) was generated with ABScript III RT Master Mix for qPCR containing gDNA Remover (Abclonal, Wuhan, China). qRT-PCR was carried out using Genious 2 × SYBR Green Fast qPCR Mix (Abclonal, China). GAPDH served as the internal reference gene for mRNA expression analysis. Relative expression levels were determined using the 2^−ΔΔCt method.

The primer sequences were utilized: GAPDH-F: CTTTGTCAAGCTCATTTCCTGG, GAPDH-R: TCTTGCTCAGTGTCCTTGC; MuRF1-F: GTGTGAGGTGCCTACTTGCTC MuRF1-R: GCTCAGTCTTCTGTCCTTGGA; Atrogin-1-F: CAGCTTCGTGAGCGACCTC, Atrogin-1-R: GGCAGTCGAGAAGTCCAGTC.

2.14. Animals and Treatments

All animal procedures adhered to the Experimental Animal Administration Guidelines of China Agricultural University and were approved by the China Agricultural University Animal Welfare and Animal Experimental Ethical Inspection Committee under approval number AW050705202-2-1. The study was designed, performed, and reported in adherence to the ARRIVE 2.0 guidelines.

Five-week-old male C57BL/6 mice weighing 15–17 g were obtained from Beijing SPF Biotechnology Co., Ltd. (Beijing, China). After a three-week acclimatization period under standard housing conditions (temperature: 22 ± 3 °C; humidity: 50–60%; 12 h light/dark cycle; ad libitum access to food and water), 32 mice were randomly assigned to two groups (n = 16 per group): the Model group and the Exos group. In the Model group, the left tibialis anterior (TA) muscle was injected with glycerol (Gly), while the contralateral right TA muscle received an equal volume of PBS as a control treatment (PBS). In the Exos group, the left TA muscle was co-treated with glycerol and cADMSC-Exos (Exos), and the right TA muscle was injected with PBS as a control treatment (PBS). The experimental procedures were performed as follows:

Day 1 (muscle injury induction): The left TA muscle of each mouse received an intramuscular injection of 50 μL of 50% glycerol (Sigma, Darmstadt, Germany) to induce injury, while the right TA was injected with an equivalent volume of PBS as an internal control.

Day 2 (treatment): Mice in the Exo group were administered a 100 μg intramuscular injection of cADMSC-Exos into the left TA muscle, whilst the right TA muscle was injected with an equivalent volume of PBS as an internal control. To control for injection-related variables, mice in the Model group received equal volumes of PBS injected into both the left and right TA muscles.

Day 7 and Day 14 (sample collection): At the designated time point, eight mice were selected from each group. Following anaesthesia with 10% isoflurane, euthanasia was performed by cervical dislocation. Muscle atrophy was assessed by calculating the proportion of muscle weight (mg) to body weight (g). After harvesting, each TA muscle was systematically divided into predefined portions to accommodate different downstream analyses. Samples from comparable mid-belly regions of the muscle were allocated for histological staining (H&E, Masson’s trichrome, and Oil Red O), while the remaining tissue was rapidly frozen in liquid nitrogen and stored at −80 °C for protein analysis by Western blotting. Tissue sampling was performed in a standardized manner across all experimental groups to minimize regional bias and ensure comparability.

2.15. Hematoxylin and Eosin (H&E) and Masson Staining

After harvesting, TA tissue was fixed in 4% paraformaldehyde overnight. Subsequently, the tissue underwent ethanol gradient dehydration and was embedded in paraffin. Transverse sections of 5 μm thickness were obtained from paraffin-embedded samples and stained with H&E and Masson’s Trichrome using commercial kits (Solarbio) according to the manufacturer’s protocols. Representative images were acquired under standardized conditions using an SDPTOP CX40 microscope (Sunny Optical, Ningbo, China). Muscle fiber cross-sectional area (CSA, μm²) was quantified using ImageJ software under unified threshold settings to ensure comparability across all samples. For each sample, multiple randomly selected fields were analyzed, and the mean CSA value was used for statistical analysis. The degree of fibrosis was assessed based on Masson’s Trichrome staining, and the fibrotic area percentage relative to total cross-section was automatically calculated using Fiji software (v10.8.1) with consistent image-processing parameters applied to all regions.

2.16. Oil Red O Staining

Freshly harvested TA muscles were stabilized in 4% paraformaldehyde prior to cryoprotection via an ascending sucrose gradient. Once encapsulated in optimal cutting temperature (OCT) medium, the specimens were sliced into 12 μm-thick cryosections. To assess lipid accumulation, an Oil Red O staining kit (Solarbio) was employed following the manufacturer’s protocol. In short, the sections underwent a preliminary rinse in 60% isopropanol, followed by a 20 min immersion in a saturated Oil Red O working solution and subsequent rapid differentiation. For nuclear visualization, slides were counterstained with Mayer’s hematoxylin for 3 min and then subjected to a 10 min blueing step under running tap water. Sections were mounted in 75% glycerol for observation. Quantitative analysis of lipid infiltration was performed in Fiji software using automated color-threshold segmentation with uniform settings across all images. The proportion of Oil Red O-positive area relative to total tissue area was calculated automatically, and the mean value from multiple random fields per section was used for group comparison.

2.17. Data Statistics and Analysis

Statistical analyses were conducted using GraphPad Prism software (v10.1.2; GraphPad Software, San Diego, CA, USA). Data are expressed as mean ± standard deviation (SD). For cells subjected to different treatments, statistical analysis was performed using one-way analysis of variance (ANOVA). For comparisons involving TA muscle, two-way ANOVA was employed to assess differences among multiple groups. All ANOVA analyses were followed by Tukey’s post hoc test. A p value of less than 0.05 was regarded as statistically significant.

3. Results

3.1. Characteristics of cADMSCs

cADMSCs isolated from peri-ovarian and uterine horn adipose tissues exhibited a characteristic fibroblast-like, spindle-shaped morphology (Figure 1A). The multilineage plastic potential of these cells was confirmed by successful differentiation into adipogenic, chondrogenic, and osteogenic lineages (Figure 1B). Flow cytometric analysis demonstrated that the isolated cells were strongly positive for mesenchymal surface markers CD29 (91.8%), CD44 (98.8%), and CD90 (93.8%), while remaining negative for the hematopoietic/MHC-II marker HLA-DR (0.0%), consistent with the established immunophenotypic criteria for MSCs (Figure 1C). Furthermore, growth kinetic analysis revealed that cADMSCs at passages 2, 5, and 8 maintained progressive expansion over an 8-day culture period without reaching a definitive plateau (Figure 1D). Notably, P2 cells displayed the most robust proliferative capacity, with a PDT of 0.89 ± 0.05 d, which increased as the passage number progressed (P5: 1.72 ± 0.02 d; P8: 1.81 ± 0.02 d).

3.2. Characterization of cADMSC-Exos

The obtained cADMSC-Exos were characterized by morphology, size distribution, and protein composition. TEM imaging showed typical double-membrane, cup-shaped vesicles (Figure 2A). NTA measurements indicated a narrow size distribution with an average diameter of 131.1 nm and a final concentration of 2.5 × 10¹⁰ particles/mL (Figure 2B). Based on the protein concentration (0.98 mg/mL), the administered in vivo dose of 100 μg total protein corresponds to approximately 2.55 × 10⁹ particles. Western blot analysis further verified the presence of exosomal markers (TSG101, Alix, and HSP70) and the absence of the cellular contaminant Calnexin, confirming successful purification of cADMSC-Exos (Figure 2C).

3.3. cADMSC-Exos Protect C2C12 Cells Against Dexamethasone-Induced Cytotoxicity and Myotube Atrophy

Based on previous studies, an in vitro muscle atrophy model was generated by exposing C2C12 cells to 10 μM dexamethasone for 24 h [21]. The CCK-8 assay was employed to assess dexamethasone-induced cytotoxicity and to identify the optimal concentration of cADMSC-Exos (Figure 3A). Dexamethasone significantly reduced cell viability compared with the control group (p < 0.05). Co-treatment with cADMSC-Exos dose-dependently attenuated this reduction, with the maximal protective effect observed at 50 μg/mL (p < 0.001). No further increase in cell viability was detected at higher concentrations. Therefore, 50 μg/mL cADMSC-Exos was used in subsequent experiments.

To evaluate the impact of cADMSC-Exos on myotube morphology, differentiated C2C12 myotubes were exposed to dexamethasone with or without cADMSC-Exos and immunostained for MyHC (Figure 3B). Dexamethasone reduced MyHC fluorescence intensity, myotube diameter, and fusion index, whereas cADMSC-Exos co-treatment significantly increased MyHC fluorescence intensity (p < 0.05) and improved myotube diameter (p < 0.0001) and fusion index (p < 0.01) relative to the DEX group (Figure 3C–E).

Furthermore, Western blot and qRT-PCR analyses were performed to examine the expression of the muscle atrophy-related E3 ubiquitin ligases muscle RING finger 1 (MuRF1) and muscle atrophy F-box (Atrogin-1). Dexamethasone treatment significantly increased the protein and mRNA levels of MuRF1 and Atrogin-1 (p < 0.01), whereas co-treatment with cADMSC-Exos markedly reduced their expression (p < 0.05) (Figure 3F–J). Taken together, these results reveal that cADMSC-Exos protect C2C12 cells from dexamethasone-induced cytotoxicity and myotubular atrophy.

3.4. cADMSC-Exos Attenuate Glycerol-Induced Skeletal Muscle Injury In Vivo

An acute glycerol-induced TA muscle injury model was established in 8-week-old male C57BL/6 mice. An overall view is in Figure 4A. Gross examination showed apparent muscle atrophy in glycerol-injured muscles, particularly at 7 days post-injury, whereas Exos-treated muscles exhibited no obvious atrophic changes compared with contralateral controls (Figure 4B). Consistently, TA muscle weight was significantly reduced in the Gly treatment at 7 and 14 days post-injury (p < 0.0001 and p < 0.01, respectively), while Exos treatment significantly attenuated muscle weight loss at both time points (p < 0.001 and p < 0.01, respectively). No significant differences were observed between Exos-treated and contralateral PBS-treated muscles across groups, further indicating that Exos treatment can restore muscle mass (Figure 4C).

H&E staining revealed that at day 7 post-injury, Gly treatment in the Model group exhibited inflammatory infiltration and a significant reduction in muscle fibre CSA compared with the PBS control treatment on the contralateral side (p < 0.0001) (Figure 5A,B). Compared with Gly treatment, Exos treatment reduced inflammatory infiltration and markedly increased CSA (p < 0.001). By day 14, inflammation in the TA muscles treated with glycerol and cADMSC-Exos had largely subsided. CSA further improved under Exos treatment, reaching levels comparable to the contralateral PBS treatment; whereas CSA remained minimal under Gly treatment. Moreover, at both time points, both Gly treatment and Exos treatment demonstrated a higher proportion of central nucleus fibres (CNFs) compared to the contralateral PBS treatment, indicating that the muscle regeneration process was still ongoing.

Masson staining revealed marked collagen deposition in the Gly-treated Model group at both days 7 and 14 post-injury, compared with the contralateral PBS-treated group (p < 0.0001) (Figure 5C,D). The area of fibrosis was notably reduced following Exos treatment, particularly at day 14 (p < 0.0001), with fibrosis levels approaching those of the contralatera PBS-treated Exos group. This indicates that cADMSC-Exos effectively mitigate injury-related fibrotic remodelling.

Oil Red O staining further confirmed that Gly treatment in the Model group resulted in substantial lipid deposition in the TA muscle at both time points (p < 0.0001) (Figure 5E,F), whereas Exos treatment markedly reduced intramuscular lipid droplet accumulation (p < 0.0001), restoring droplet levels by day 14 to near those observed in the PBS-treated group.

Finally, Western blot analysis showed that glycerol-induced muscle injury markedly upregulated the expression of MuRF1 and Atrogin-1 (p < 0.01) (Figure 6A–D). Treatment with cADMSC-Exos significantly suppressed this upregulation in injured muscles (p < 0.05). Collectively, these findings demonstrate that cADMSC-Exos effectively promote muscle repair and attenuate pathological remodeling following glycerol-induced skeletal muscle injury in mice.

4. Discussion

Skeletal muscle injury and dysfunction are common conditions in dogs, yet there is currently a lack of therapies capable of effectively promoting regeneration and repair of skeletal muscle [29,30]. This study for the first time confirms that cADMSC-Exos effectively alleviate muscle atrophy in both in vitro and in vivo models, while significantly suppressing pathological remodelling processes such as fibrosis and fatty infiltration following injury. These findings provide crucial preclinical evidence for developing cell-free therapies based on cADMSC-Exos for treating musculoskeletal disorders in dogs.

In the field of stem cell therapy, the selection of cell sources is pivotal in determining therapeutic efficacy and translational feasibility. Currently, canine MSCs can be obtained from multiple tissues including bone marrow, umbilical cord, amnion, placenta, and adipose tissue [22]. Among these, cADMSCs are particularly noteworthy due to their strongest proliferative activity and shortest PDT [23]. This highly efficient proliferative capacity, coupled with the advantage of being readily obtainable from discarded clinical surgical tissue, endows cADMSCs with considerable translational potential for the large-scale preparation of therapeutic exosomes [31]. The cADMSCs isolated in this study exhibited the typical MSC phenotype and multipotent differentiation capacity, maintaining vigorous proliferative activity during early passage stages. This establishes a robust cellular foundation for subsequent stable and reliable exosome production.

Compared to direct MSC transplantation, exosomes as a cell-free therapy effectively circumvent the risks of tumourigenicity and immune rejection associated with living cell applications, offering superior safety advantages [32]. However, despite the promising prospects of exosome therapy, the efficacy of canine-derived MSC-Exos in skeletal muscle injury restoration has previously lacked systematic investigation. This study not only successfully isolated and characterised high-purity cADMSC-Exos meeting international characterisation standards, but further evaluated their functional role in muscle injury recovery through dual-model validation for the first time, thereby filling a knowledge gap in this field.

Glucocorticoids such as dexamethasone, commonly employed to treat inflammatory or immune-mediated conditions in dogs, often necessitate long-term or high-dose administration [27]. However, this may lead to numerous adverse reactions, including muscle atrophy [33]. The mechanism underlying the dexamethasone-induced muscle atrophy model primarily involves the activation of the ubiquitin–proteasome system (UPS), specifically through the upregulation of E3 ubiquitin ligases such as MuRF1 and Atrogin-1, which mediate myofibrillar protein degradation [34]. MuRF1 directly targets contractile proteins such as MyHC and α-actin for degradation, whilst Atrogin-1 mediates the degradation of myogenic regulatory factors including myogenic differentiation protein D (MyoD), thereby impeding the formation of new muscle fibres [35]. In our study, cADMSC-Exos improved the survival rate of C2C12 cells impaired by dexamethasone, maintained the integrity of myotube morphology, and enhanced fusion capacity. Moreover, cADMSC-Exos markedly inhibited the upregulation of MuRF1 and Atrogin-1 at both the protein and transcriptional levels. These findings suggest that cADMSC-Exos may attenuate the atrophy process at the molecular level, likely through the downregulation of key E3 ubiquitin ligases (MuRF1 and Atrogin-1) involved in the UPS pathway, thereby reducing the potential for myofibrillar protein degradation and protecting the myogenic regulatory network. This provides a plausible mechanistic explanation for the anti-atrophy effects observed in our in vivo models.

More importantly, in a glycerol model simulating severe skeletal muscle injury, cADMSC-Exos demonstrated multifaceted reparative functions. Not only did it promote myofibrillar regeneration, manifested by increased CSA and an increase in CNFs, but it also significantly alleviated fibrosis and lipid infiltration. This global improvement in pathological remodelling may stem from the systemic regulation of the injured microenvironment by exosomes. Existing research indicates that MSC-Exos can induce macrophages to polarise towards the anti-inflammatory M2 phenotype [36]. Exosomes derived from M2 macrophages can alter the abnormal differentiation trajectory of fibroblast/adipocyte precursor cells (FAPs), which may represent the key mechanism underlying the observed reduction in fibrosis and fat infiltration [37,38]. Consequently, the improvement in pathological remodelling observed in this study suggests that cADMSC-Exos may exert an indirect regulatory effect on the activity and differentiation fate of FAPs, thereby effectively curbing adverse tissue remodelling processes such as post-injury fibrosis and fatty infiltration. Furthermore, the downregulation of MuRF1 and Atrogin-1 at the protein level by cADMSC-Exos aligns with findings from in vitro models, further confirming the potent anti-atrophic activity of cADMSC-Exos.

Although the findings of this study provide valuable insights, several limitations should be acknowledged. First, the specific bioactive components mediating the regenerative effects of cADMSC-Exos remain unidentified. The therapeutic actions of MSC-Exos are widely attributed to their molecular cargo, particularly regulatory miRNAs and proteins. Previous studies have reported that exosomal miRNAs such as miR-486 and miR-132 play important roles in alleviating skeletal muscle atrophy by modulating proteostasis [21,33]. Whether similar regulatory molecules are enriched in cADMSC-Exos and contribute to the observed suppression of the UPS pathway warrants further investigation. Future studies will therefore require the application of multi-omics approaches, including miRNA sequencing and proteomic analyses, to elucidate the underlying molecular mechanisms in greater depth. Second, this study did not directly assess the biodistribution and retention of exosomes using in vivo imaging techniques. Given the intramuscular route of administration, we infer that the observed local therapeutic efficacy—manifested as reduced atrophy and alleviated pathological remodelling—along with the concurrent downregulation of UPS markers in the treated tissue, is primarily mediated by the direct action of exosomes at the injection site. However, without direct tracking data, the possibility of systemic redistribution or secondary systemic effects cannot be fully excluded. Future investigations utilizing high-fidelity tracking methods, such as genetic labelling (e.g., CD63-GFP or CD63-NanoLuc), are necessary to precisely map the spatiotemporal fate of cADMSC-Exos [39]. Third, although the high conservation of skeletal muscle physiology across mammals supports the cross-species bioactivity observed here, the use of murine models to evaluate canine-derived exosomes limits direct translational relevance. Potential species-specific variations in immune responses or ligand–receptor interactions cannot be fully ruled out in a xenogeneic setting [40]. Therefore, this research remains at the preclinical stage. Future translation into canine clinical applications necessitates further standardisation of exosome preparation protocols, optimisation of administration methods and dosage regimens, alongside systematic long-term safety assessments in target species (dogs). These steps are essential to ultimately advance its practical application in treating canine muscular disorders.

5. Conclusions

In this study, cADMSCs were obtained from adipose tissue discarded during routine OHE procedures in healthy female dogs, and cADMSC-Exos were subsequently isolated. The therapeutic efficacy of cADMSC-Exos was then evaluated using both a dexamethasone-induced myotube atrophy model and a glycerol-induced skeletal muscle injury model in mice. Our results demonstrate that cADMSC-Exos can alleviate pathological changes such as skeletal muscle atrophy, fibrosis, and fatty infiltration. These effects may be attributed to the ability of cADMSC-Exos to regulate local protein metabolism and the damaged microenvironment. Collectively, this preliminary evidence suggests that cADMSC-Exos hold potential for improving skeletal muscle injury, laying the groundwork for translational research applications in future veterinary treatments for musculoskeletal disorders.