Juvenile vs Adult Skeletal Muscle Transplants in the Treatment of Volumetric Muscle Loss Injury

Juvenile vs Adult Skeletal Muscle Transplants in the Treatment of Volumetric Muscle Loss Injury | Research Square window.SnipcartSettings = { analytics: { enabled: false } }; (function() { var accessVector = localStorage.getItem('access_vector') || ''; window.dataLayer = window.dataLayer || []; if (accessVector) { window.dataLayer.push({ user: { profile: { profileInfo: { snid: accessVector } } } }); } })(); (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0],j=d.createElement(s),dl=l!='dataLayer'?'&l='+l:'';j.async=true;j.src='https://www.googletagmanager.com/gtm.js?id='+i+dl;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-K279D39R'); Browse Preprints In Review Journals COVID-19 Preprints AJE Video Bytes Research Tools Research Promotion AJE Professional Editing AJE Rubriq About Preprint Platform In Review Editorial Policies Our Team Advisory Board Help Center Sign In Submit a Preprint Cite Share Download PDF Research Article Juvenile vs Adult Skeletal Muscle Transplants in the Treatment of Volumetric Muscle Loss Injury John J. Payne, Samuel R. Frandsen, Zachary H. Rasmussen, Matthew J. Mangus, and 7 more This is a preprint; it has not been peer reviewed by a journal. https://doi.org/ 10.21203/rs.3.rs-7437055/v1 This work is licensed under a CC BY 4.0 License Status: Published Journal Publication published 03 Dec, 2025 Read the published version in Stem Cell Research & Therapy → Version 1 posted 9 You are reading this latest preprint version Abstract Background Volumetric muscle loss (VML) causes irreversible structural and functional deficits by removing myofibers, nerves, vasculature, extracellular matrix, and satellite cells, the resident muscle stem cells essential for regeneration. Skeletal muscle transplantation can restore tissue volume and reintroduce regenerative cells, yet functional outcomes remain incomplete. Age of the donor muscle has not been evaluated, despite evidence that juvenile muscle contains higher satellite cell density and greater myogenic plasticity than adult muscle. We hypothesized that these features would yield superior regenerative outcomes when juvenile muscle is used as a transplant source. Methods Tibialis anterior (TA) muscles from juvenile (21 d), adolescent (34 d), and adult (~ 120 d) male Lewis rats were compared for myofiber morphology, satellite cell density, and in-vitro myogenic behavior. GFP⁺ juvenile or adult muscle was then transplanted into standardized VML defects (~ 15–20% TA volume) in adult rats. Seven weeks post-surgery, in-vivo isometric strength, donor fiber integration, satellite cell distribution, and centralized nuclei were assessed. Results Juvenile muscle exhibited ~ 15× greater satellite cell density than adult (122.8 ± 28.4 vs. 8.4 ± 3.3 cells/mm², p < 0.0001) with enhanced in-vitro differentiation (fusion index + 73% vs. adult, p = 0.0067). In-vivo, both juvenile and adult transplants restored myofiber number to control levels (juvenile: 11,369 ± 1,511; adult: 9,115 ± 1,274; controls: 10,316 ± 685) and improved strength versus untreated VML (juvenile: +50%, p = 0.0016; adult: +36%, p = 0.0299). No significant functional differences were observed between donor ages. Donor fibers integrated but remained small, with localized satellite cell enrichment and increased centralized nuclei in transplant regions, consistent with ongoing regeneration. Conclusions Juvenile skeletal muscle displays cellular and structural attributes favorable for regeneration and superior in-vitro myogenic behavior compared to adult muscle. However, these advantages did not translate into greater short-term in-vivo recovery following VML transplantation. Enhancing donor fiber hypertrophy, neuromuscular integration, and satellite cell expansion beyond the transplant region, potentially through rehabilitation or pharmaceutical interventions, may be necessary to realize the full therapeutic potential of juvenile donor muscle for regenerative medicine applications. volumetric muscle loss skeletal muscle transplantation satellite cells juvenile muscle donor age muscle regeneration tissue engineering stem cell therapy regenerative medicine Figures Figure 1 Figure 2 Figure 3 Figure 4 Figure 5 Figure 6 INTRODUCTION Volumetric muscle loss (VML) is a debilitating injury caused by trauma, tumor excision, or surgery that removes substantial amounts of skeletal muscle along with its cellular, vascular, neural, and extracellular matrix networks ( 1 , 2 ). This destructive injury eliminates structural support and depletes satellite cells, the resident muscle stem cells essential for regeneration ( 3 , 4 ). Because the satellite cell niche (basal lamina) is disrupted, the cells are unable to migrate into areas of damage and facilitate regeneration. Conversely, fibrotic scar tissue fills the void, overwhelming the intrinsic repair mechanisms, and results in chronic deficits in muscle strength and function ( 5 , 6 ). Clinically, there is no standard of care to treat VML, thus management is often focused on wound closure and limb salvage rather than restoration of muscle mass and function ( 7 ). Experimental regenerative approaches, including skeletal muscle transplantation and autologous minced muscle grafts, aim to fill the defect with donor tissue that provides both structural scaffolding and a reservoir of regenerative cells ( 7 , 8 ). Upon transplantation, donor satellite cells can activate, proliferate, differentiate, and fuse with host fibers to support local regeneration ( 4 ). However, even with successful donor integration, functional recovery is typically incomplete ( 5 , 9 ), highlighting the potential need to optimize donor tissue characteristics. One underexplored factor influencing transplant efficacy is the developmental age of donor muscle, as skeletal muscle undergoes pronounced structural and cellular changes across the lifespan ( 10 – 12 ). Juvenile muscle exhibits small-diameter myofibers, high nuclear and satellite cell densities, and a microenvironment supportive of fiber growth and remodeling ( 3 , 13 , 14 ). In contrast, adult muscle contains fewer satellite cells and larger, less plastic fibers ( 3 , 10 ). Such age-related differences may have critical implications for regenerative potential following transplantation. Evidence from other regenerative contexts supports this concept: juvenile tissue or juvenile-like progenitors often outperform adult counterparts in repair, likely due to a more abundant and responsive stem cell pool combined with a growth-permissive environment ( 3 , 10 , 15 ). Our group and others have shown that regenerative outcomes are constrained by the injury environment, including persistent denervation ( 16 , 17 ), extracellular matrix remodeling ( 2 ), and limited stem cell migration into the defect ( 5 ). These constraints suggest that the intrinsic advantages of juvenile donor tissue may still require complementary interventions, such as rehabilitation, to achieve full functional recovery ( 18 – 20 ). Here, we tested the hypothesis that juvenile skeletal muscle, by virtue of its cellular composition and myogenic capacity, would outperform adult muscle in restoring structure and function following VML injury. We first characterized the morphology, satellite cell content, and in-vitro myogenic behavior of tibialis anterior (TA) muscle from juvenile, adolescent, and adult rats. We then compared the in-vivo regenerative outcomes of juvenile versus adult TA muscle transplants in a standardized rat VML model. METHODS This study was conducted in two parts. Study 1 evaluated the structural and cellular properties of juvenile, adolescent, and adult skeletal muscle to determine age-dependent differences relevant to regeneration. Study 2 examined the in-vivo regenerative performance of juvenile and adult muscle transplants in a rat model of VML, using functional and histological outcome measures seven weeks post-VML surgery. The work herein has been reported in line with the ARRIVE guidelines 2.0. Ethical Approval All experiments were approved by the Institutional Animal Care and Use Committee (IACUC) at Brigham Young University (Protocol #23-0401) in accordance with National Institutes of Health guidelines for the care and use of laboratory animals. All animals were housed on a 12-12 h light-dark cycle, with food and water ad libitum. Study 1: Developmental Analysis of Donor Muscle Animals and Tissue Collection To evaluate regenerative properties of skeletal muscle by age, male Lewis rats were grouped as juvenile (21 ± 0 days), adolescent (34 ± 0 days), or adult (130 ± 4.6 days) (n = 4 per group, 12 total). The left tibialis anterior (TA) muscle was harvested for histological analysis, and the right TA was used for myoblast isolation. Sample size was based on prior reports that show large age-related differences in rodent skeletal muscle. All animals were euthanized following tissue collection while under heavy isoflurane anesthesia (5%) until loss of reflexes, then bilateral thoracotomy was used as a secondary method to ensure death. Immunohistochemistry and Imaging To assess age-related satellite cell content, myofiber number, and muscle morphology, TA muscles were collected, weighed, then embedded in tragacanth gum, snap frozen in isopentane, and cryosectioned (10 µm). Sections were fixed in 4% paraformaldehyde (Thermo Scientific), blocked with 10% normal goat serum (NGS; Thermo Fisher), and incubated overnight with Pax7 (mouse IgG1, 3 µg/mL; DSHB, AB_528428) and laminin (mouse IgG2a, 2 µg/mL; DSHB, AB_2618140). Secondary antibodies included Cy3 goat anti-mouse IgG1 (1:200; Jackson, 115-165-003) and Alexa Fluor 647 goat anti-mouse IgG2a (1:200; Jackson, 115-605-206). DAPI (1 µg/mL; Thermo Fisher, D1306) was used for nuclear counterstaining. Whole tissue cross-sections were used to count myofibers and to measure whole TA and myofiber cross-sectional area. Images were acquired on a tile-scanning Echo Revolution fluorescence microscope using a 20× PlanX Apo objective. The laminin images were converted to grayscale and a threshold set (10–30 range) using Fiji (ImageJ) software. Myofibers were segmented using “Analyze Particles” with a size range of 50–8000 µm² and circularity 0.30–1.00. Manual correction was performed for incomplete borders. Mean fiber CSA and total fiber number were calculated per sample. To assess satellite cell and nuclei content, five 1 mm² regions of interest were randomly imaged throughout each sample. Nuclei were counted based on images from the DAPI channel using the Otsu threshold parameters and the “watershed” feature to separate nuclei clusters. The “Analyze Particles” feature was set to a size threshold of 20-50µm. Satellite cells were identified and counted manually based on Pax7⁺/DAPI⁺ co-localization within or near the laminin border. Myoblast Isolation and In Vitro Assays To characterize age-dependent differences in satellite cell behavior and regenerative potential, primary myoblasts (activated satellite cells) were isolated from the right TA muscles of juvenile, adolescent, and adult rats. These cells were used in a series of in vitro assays to assess proliferative capacity, differentiation potential, and myogenic identity. Right TA muscles were digested using the Miltenyi Skeletal Muscle Dissociation Kit and gentleMACS system. Cell suspensions were filtered (70µm) and centrifuged, then incubated with PE-conjugated anti-CD106 (Miltenyi, 130-103-684) and sorted with Anti-PE MicroBeads (130-048-801) using LS columns. CD106⁺ myoblasts were seeded on Matrigel-coated plates and allowed to expand in DMEM (Gibco) with 10% FBS and 1% penicillin-streptomycin changed every 48 hours. Myogenic identity was confirmed via MyoD1 staining (mouse IgG2b, 2 µg/mL; DSHB, AB_2146602) with Cy3 anti-IgG2b secondary (1:200; Jackson, 115-165-207). DAPI was used for nuclei staining. The percentage of MyoD⁺ nuclei was assessed in three random fields per group. Myoblasts were seeded at a density of 25,000 cells/well on a Matrigel coated, 12-well plate and allowed to incubate overnight. Proliferation was assessed by EdU incorporation (Click-iT™ kit, Thermo Fisher, C10340) following manufacturer instructions at 4, 8, and 16 hours. Briefly, cells were fixed (4% PFA), permeabilized (0.5% TritonX-100 in PBS), and labeled with Alexa Fluor 647-conjugated azide and Hoechst. Hoechst-stained nuclei were counted, and the proliferation index was calculated as EdU⁺ cells/total nuclei. Data were averaged from four fields per well, in triplicate wells per group. For differentiation, cells were seeded at 100,000 cells per well on a Matrigel coated, 12-well plate and allowed to incubate overnight in growth media. Differentiation was induced by switching to DMEM + 2% horse serum for 72 hours. Cells were fixed and stained for Myogenin (mouse IgG1, 2 µg/mL; DSHB, AB_2146601) and MyHC (mouse IgG2b, 2 µg/mL; DSHB, AB_2147781). Secondary antibodies included Alexa Fluor 647 anti-IgG1 and Cy3 anti-IgG2b (1:200; Jackson). DAPI was used for nuclear staining. Three outcome measures were used to assess differentiation: fusion index, MyHC + area, and nucleation index. The fusion index was defined as the percentage of total nuclei located within MyHC + multinucleated myotubes, indicating the efficiency of myoblast fusion during early differentiation. Myotubes were defined as MyHC + structures containing two or more DAPI-stained nuclei. The MyHC + area was calculated as the total surface area occupied by MyHC + staining within each field of view, measured in ImageJ using a threshold binary masks. The nucleation index was calculated as the average number of nuclei contained within each individual MyHC + myotube. This index served as an indicator of myotube maturation. All quantifications were performed on at least four randomly selected, non-overlapping fields per well, and averaged across triplicate wells per condition. Study 2: VML Injury and Muscle Transplantation VML Surgery and Experimental Design To evaluate the regenerative performance of juvenile versus adult muscle transplants in a preclinical model of VML, a standardized full thickness VML injury and repair protocol was conducted in adult male rats. This approach was designed to mimic clinically relevant muscle trauma and assess functional and histological outcomes following transplantation of developmentally distinct donor tissue. Twenty-four adult male Lewis rats (3–4 months old) were randomly assigned to one of three treatment groups: VML No Treatment, VML + Adult Transplant, or VML + Juvenile Transplant (n = 8 per group). Sample size was based on prior muscle transplant studies in a rat VML model to detect functional improvements of 20–40% (17). All animals underwent unilateral VML surgery in the left TA muscle with the right leg serving as an uninjured, intra-animal control. To provide analgesia, a carprofen tablet (Bio-Serv, 5 g) was placed in the animal’s cage 24 hours prior to surgery and a single preoperative dose of sustained-release buprenorphine (1.2 mg/kg, SC; Wedgewood) was injected subcutaneously into the back of the neck at least one hour before surgery. Animals were anesthetized with 2–3% isoflurane in oxygen and placed in a supine position on a heated surgical platform. The lower left hindlimb was shaved and disinfected with alternating scrubs of chlorhexidine and 70% ethanol. A longitudinal skin incision (~1.5 cm) was made along the anterior surface of the lower leg to expose the underlying musculature. The skin and fascia were opened to expose the TA muscle, which was gently isolated from surrounding tissue using blunt dissection. To protect adjacent musculature, a sterile surgical spatula was inserted beneath the TA and full-thickness defect, approximately 6 mm in diameter (~15–20% of muscle volume), was created in the mid-belly region using a sterile biopsy punch (MedBlades, USA). Donor tissue was harvested immediately prior to transplantation from ubiquitously expressing GFP⁺ juvenile (21-day; n=4) or adult (~120-day; n=2) male Lewis rats (Rat Resource and Research Center; Strain: LEW-Tg(CAG-EGFP)YsRrrc; RRRC#: 00206) . TA muscles were dissected, placed in a sterile tissue culture dish on ice, and finely minced into ~1 mm³ fragments using sterile scissors. The total weight of minced tissue was adjusted to approximate the volume of the VML defect and then carefully implanted into the defect site. For animals receiving transplants, the fascia was sutured, followed by subcutaneous closure of the skin using interrupted sutures. VML No Treatment animals underwent the same surgical procedure without tissue implantation. In Vivo Muscle Strength Testing To evaluate muscle function following VML injury and transplantation, in-vivo isometric force testing was performed seven weeks post-surgery for all of the VML injured limbs (n=24) and most (n=19) of the uninjured control limbs. Under 2–3% isoflurane anesthesia, rats were placed supine on a temperature-controlled platform. The hindlimb was immobilized using a knee clamp, and the foot was secured to a force transducer footplate (Aurora Scientific 3-in-1 Muscle Test System). Subcutaneous needle electrodes were inserted near the peroneal nerve to stimulate the anterior compartment (TA/EDL) of the hindlimb. Stimulation parameters included 0.1 ms pulse width, 400 ms train duration, and increasing frequencies ranging from 10 to 200 Hz. Optimal voltage was determined for each animal to ensure maximal contractile response. Peak isometric tetanic force was recorded as the highest value generated during the frequency ramp. To account for inter-animal variability in body size, absolute force values were normalized to body weight (mN·m/kg). Tissue Collection and Histology Seven weeks post VML surgery, TA, EDL, soleus, and gastrocnemius muscles were dissected and weighed. Animals were then euthanized with an overdose of isoflurane (5%) and bilateral thoracotomy as a secondary measure to ensure death. TA sections were frozen in isopentane cooled in liquid nitrogen and stored at -80°C. The TA muscles were sectioned (10 µm) and stained for laminin, Pax7, and DAPI. High-resolution images were taken randomly from controls and from four regions in VML injured samples: defect (typically devoid of myofibers), transplant (identified by GFP expression), border (adjacent to the injury; smaller, disorganized host fibers), and distal (intact muscle away from the injury site). Satellite cells were counted based on Pax7⁺/DAPI⁺ co-localization. Myofiber CSA, total fiber count, and centrally nucleated fibers were quantified from regions of interest using ImageJ analysis software and the previously described segmentation thresholds. Donor Fiber Analysis To assess transplant integration and morphology, TA muscle sections were analyzed for GFP fluorescence and labeled with wheat germ agglutinin (WGA) to identify myofiber boundaries. Sections were cut at 10µm thickness and immediately fixed in 2% paraformaldehyde for 10 minutes at room temperature to preserve endogenous GFP signal. Following fixation, slides were rinsed once with PBS and incubated for 15 minutes with Alexa Fluor 647-conjugated wheat germ agglutinin (WGA; 1:500 dilution in PBS; Thermo Fisher Scientific, Cat# W32466). Sections were washed once in PBS and mounted in Fluoroshield (Sigma) and imaged using a 20X PlanX Apo objective on the Echo Revolution fluorescence microscope. GFP fluorescence was visualized directly without antibody amplification. Exposure settings were optimized to avoid saturation and kept constant between samples within each group. To quantify GFP⁺ donor fibers, high-resolution images were collected from well-defined GFP regions within the transplant zone. Fields with high signal-to-noise ratio and clear membrane borders were prioritized for analysis. Myofiber CSA was measured using ImageJ. GFP-positive fibers were defined as those showing continuous cytoplasmic GFP signal enclosed by WGA-labeled borders. A minimum of 200 GFP-positive fibers were analyzed per animal. The GFP area was also calculated as a percentage of total CSA using full-section scans in ImageJ, thresholded for GFP fluorescence. Statistical Analysis All statistical analyses were performed using GraphPad Prism version 10.5.0. One-way ANOVA with Tukey’s post hoc test was used for group comparisons. Data are presented as mean ± SD. A p-value < 0.05 was considered statistically significant. RESULTS Study 1: Comparison Between Juvenile, Adolescent, and Adult Male Lewis Rats Rat Characteristics Juvenile (21-day), adolescent (34-day), and adult (120-day) male Lewis rats were used to represent distinct developmental stages. As expected, body and TA muscle weights increased significantly with age (Table 1), consistent with known patterns of postnatal growth. Table 1: Rat Characteristics Group Age (days) Body Weight (grams) TA Wet Weight (grams) Juvenile 21 ± 0 53.85 ± 0.37 0.16 ± 0.0036 Adolescent 34 ± 0 144.5 ± 6.25 0.25 ± 0.0183 Adult 120 ± 5.7 396.5 ± 16.6 0.775 ± 0.031 Muscle Size and Myofiber Architecture Vary by Developmental Stage Whole-muscle CSA increased significantly with age (Figure 1A–B). Juvenile TAs averaged 3.95 ± 0.29 mm², adolescent TAs 9.64 ± 1.80 mm², and adult TAs 27.07 ± 5.36 mm². All pairwise comparisons were statistically significant (juvenile vs. adolescent p = 0.0012; juvenile vs. adult p < 0.0001; adolescent vs. adult p = 0.0003), confirming stepwise muscle growth. Despite these CSA increases, total myofiber number did not significantly differ between groups (juvenile: 8593 ± 525; adolescent: 10,101 ± 1484; adult: 10,126 ± 1282; p = 0.1621), indicating that muscle growth was driven primarily by fiber hypertrophy, not hyperplasia (Figure 1C). Analysis of individual myofiber CSA revealed age-related increases (Figure 1D). Juvenile fibers averaged 460 ± 38 µm², significantly smaller than adolescent (963 ± 147 µm²; p = 0.0267) and adult fibers (2798 ± 354 µm²; p < 0.0001). Adolescent fibers were also significantly smaller than adult fibers ( p < 0.0001). The CSA frequency distributions further emphasized this shift as juvenile muscle showed a narrow peak at small fiber sizes, whereas adolescent and adult profiles were broader and right-shifted, consistent with increased fiber size (Figure 1E). Nuclei, Myofiber, and Satellite Cell Density Decline with Advancing Age To assess how developmental age affects muscle, we quantified nuclei, myofiber density, and Pax7⁺ satellite cells per mm² (Figure 2A–B). Nuclear density was highest in juvenile muscle (2907.5 ± 99.4 nuclei/mm²), declined to 1550.4 ± 268.1 in adolescents, and dropped further in adults (704.3 ± 139.6; p < 0.0001 for all comparisons) (Figure 2C). Myofiber density also declined significantly with age ( p < 0.001), consistent with increasing fiber size (Figure 2D). Satellite cell density, measured by Pax7⁺ nuclei per mm², showed a dramatic age-related decline (Figure 2E). Juvenile muscles contained 122.8 ± 28.4 Pax7⁺ cells/mm², approximately 3.5 times higher than adolescents (36.1 ± 7.1; p < 0.001) and 15 times higher than adults (8.4 ± 3.3; p < 0.0001). Although adolescent satellite cell density appeared higher than adult, the difference did not reach statistical significance ( p = 0.1049). Together, these results confirm that juvenile skeletal muscle possesses a dense, more cellular microenvironment with elevated satellite cell content, nuclei and myofibers, which may enhance regenerative potential when used as a source of donor tissue to treat VML. Myoblast Proliferation and Differentiation To assess how donor age affects cellular behavior, myoblasts were isolated from juvenile, adolescent, and adult TA muscles and evaluated in-vitro. Myogenic Purity and Proliferation Immunostaining confirmed high myogenic purity across all groups, with MyoD⁺ nuclei accounting for 95.3 ± 2.7% in juvenile, 95.4 ± 0.2% in adolescent, and 95.2 ± 1.4% in adult cultures (Figure 3A–B). EdU incorporations over 4, 8, and 16 hours showed a time-dependent increase in proliferation, rising from ~15% at 4 hours to ~80% at 16 hours across all groups. However, no significant differences in proliferation rate were observed between age groups at any time point ( p = 0.9696), indicating that donor age did not affect baseline proliferative capacity (Figure 3C–D). Enhanced Differentiation in Juvenile Myoblasts In contrast, differentiation declined significantly with age. After 72 hours in differentiation medium, juvenile myoblasts showed a higher fusion index (17.49 ± 2.53%) than adolescent (13.86 ± 0.83%) and adult (10.12 ± 1.79%) cultures. The difference between juvenile and adult groups was statistically significant ( p = 0.0067), while differences between juvenile and adolescent ( p = 0.1161) and adolescent and adult ( p = 0.1066) were not (Figure 3E). Myosin Heavy Chain (MyHC) area per field, a measure of total myotube formation, was significantly larger in juvenile cultures (64,667 ± 10,161 µm²) than in adolescent (40,652 ± 2,331 µm²; p = 0.0191) and adult cultures (35,149 ± 7,968 µm²; p = 0.0074) (Figure 3F). No significant difference was observed between adolescent and adult groups ( p = 0.6658). The nucleation index, defined as the average number of nuclei per myotube, did not differ significantly between groups. Juvenile (13.27 ± 2.62), adolescent (9.89 ± 2.79), and adult (7.88 ± 1.07) cultures exhibited similar nuclear content per fiber, with no statistically significant differences (Figure 3G). Overall, these findings indicate that juvenile myoblasts have an enhanced capacity for differentiation and myotube formation, despite equivalent proliferation. Study 2: Comparison Between VML Treatment Groups Rat Characteristics at Time of VML Surgery To evaluate transplant performance, rats were randomized to VML No Treatment, VML + Adult Transplant, or VML + Juvenile Transplant groups (n = 8/group). All animals were of similar age and body weight at the time of surgery, with no significant differences between groups (Table 2). However, the VML tissue excised from the No Treatment group was significantly smaller than in the transplant groups, which may have contributed to milder deficits in untreated animals. Transplant weights did not differ significantly between juvenile and adult donor groups. Table 2: Rat Characteristics at time of surgery Group Age at Surgery (Days) Weight At Surgery (g) Weight of VML Piece (mg) VML Transplant Weight (mg) VML No Treatment 110.63 ± 3.62 385.13 ± 32.52 73.97 ± 12.89 N/A VML + Adult Transplant 113.00 ± 11.78 418.88 ± 42.29 85.79 ± 18.03 93.94 ± 20.79 VML + Juvenile Transplant 107.30 ± 14.95 404.20 ± 19.23 83.40 ± 14.17 97.60 ± 11.07 Note: Bold numbers indicate significance Rat Characteristics Post VML Surgery At tissue collection (seven weeks post-surgery), body weight was lower in the No Treatment group (415.3 ± 14.6 g) compared to the Adult (440.9 ± 38.8 g) and Juvenile (443.6 ± 25.0 g) transplant groups ( p < 0.05). Despite this, individual muscle wet weights, including the TA, EDL, soleus, and gastrocnemius, were not significantly different across groups (Table 3), suggesting that gross muscle mass was not markedly influenced by transplantation within the time window assessed. Table 3: Muscle Weights Wet Weight (g) Group Body Weight at Collection (g) Right TA Left TA Right EDL Left EDL Right Soleus Left Soleus Right Gastroc Left Gastroc VML No Treatment 415.29 ± 14.57 0.74 ± 0.07 0.70 ± 0.12 0.20 ± 0.04 0.20 ± 0.02 0.17 ± 0.04 0.19 ± 0.04 1.90 ± 0.12 1.99 ± 0.21 VML + Adult Transplant 440.86 ± 38.84 0.77 ± 0.03 0.73 ± 0.09 0.17 ± 0.02 0.17 ± 0.02 0.21 ± 0.03 0.17 ± 0.01 1.99 ± 0.14 1.96 ± 0.08 VML + Juvenile Transplant 443.56 ± 24.97 0.75 ± 0.11 0.78 ± 0.15 0.17 ± 0.01 0.20 ± 0.03 0.18 ± 0.04 0.19 ± 0.03 1.90 ± 0.15 2.00 ± 0.15 Notes: Tibialis Anterior (TA), Extensor Digitorum Longus (EDL), Gastrocnemius (Gastroc). Bold numbers indicate significance. In-Vivo Strength Measurements Functional Recovery To assess whether muscle transplantation improved force production, in-vivo isometric strength of the TA/EDL complex was measured in most of the uninjured right limbs (controls; n=19) and all of the VML injured limbs (n=24) seven weeks post-injury. Uninjured control limbs generated the highest absolute force (20.96 ± 3.04 mN·m, n=19), significantly greater than all VML-injured groups ( p < 0.0001). No Treatment animals exhibited the lowest force output (9.47 ± 0.67 mN·m, n=8). Adult transplants restored force to 12.94 ± 1.52 mN·m (n=8; p = 0.0299 vs. No Treatment), while juvenile transplants reached 14.25 ± 2.01 mN·m (n=8; p = 0.0016 vs. No Treatment), though not significantly greater than adult ( p = 0.6992). Normalized to body weight, a similar pattern emerged. Control limbs produced 48.12 ± 6.44 mN·m/kg, while VML No Treatment animals fell to 23.19 ± 2.50. Juvenile and adult transplants improved force to 32.16 ± 4.42 and 29.12 ± 4.53 mN·m/kg, respectively. Only juvenile transplants produced a statistically significant improvement over No Treatment ( p = 0.0141), while adult transplants did not ( p = 0.1670). No significant difference was observed between transplant groups ( p = 0.7025). These data suggest that both donor types support partial recovery of force, with juvenile transplants trending slightly higher. Histological Assessment of Regeneration Transverse TA sections were analyzed in a subset of animals from each group (n=4) to evaluate fiber number and muscle size, representing fiber maturity and regeneration. Total myofiber number was significantly reduced in the No Treatment group (6,952 ± 743) compared to uninjured controls (10,316 ± 685; p < 0.01). Transplants restored fiber number, with adult recipients averaging 9,115 ± 1,274 fibers and juvenile recipients 11,369 ± 1,511, which were comparable to controls (Figure 4E). TA muscle CSA was lowest in the No Treatment group (22.7 ± 5.5 mm²), significantly smaller than controls (34.3 ± 1.0 mm²; p = 0.0265). Transplantation preserved CSA (adult: 30.6 ± 3.6 mm²; juvenile: 32.7 ± 6.6 mm²), as neither group differed significantly from controls (Figure 4F). Despite improvements in fiber number and muscle size, the average myofiber CSA remained significantly reduced in all VML groups. Controls averaged 2,569 ± 334 µm², compared to 1,379 ± 290 (No Treatment), 1,258 ± 97 (adult transplant), and 923 ± 151 µm² (juvenile transplant) (Figure 4G). CSA distribution curves showed that juvenile transplants had the highest proportion of small-diameter fibers (<1000 µm²), suggesting ongoing regeneration or limited hypertrophy of transplanted fibers at this potentially early time point (Figure 4H). Integration of GFP⁺ Donor Fibers To assess integration of transplanted tissue, GFP fluorescence was analyzed. All but one transplant recipient displayed a clear GFP⁺ region within the injury zone (Figure 5A–C). GFP⁺ area occupied 10.78 ± 7.27% of the total muscle CSA in adult recipients (n=7) and 12.57 ± 7.39% in juvenile recipients (n=8), with no significant difference between groups ( p = 0.6412; Figure 5D). Average CSA of GFP⁺ fibers was smaller in juvenile transplants (518.6 ± 230.7 µm²; n=8) than in adult transplants (707.9 ± 246.6 µm²; n=7), though this difference did not reach statistical significance ( p =0.1735). Distribution profiles confirmed that juvenile transplants contained a higher proportion of small-diameter GFP⁺ fibers, while adult transplants included a broader range of sizes (Figure 5E). These patterns likely reflect developmental differences in donor fiber size and post-transplant remodeling. Satellite Cell Distribution To evaluate satellite cell content following VML, Pax7⁺ cells were quantified across four regions of interest: defect, transplant, border, and distal. In uninjured controls, satellite cells were evenly distributed (8.01 ± 1.02 cells/mm²). VML injury led to a mild decline in the defect region (3.61 ± 2.33), with increased cells in the border region (27.24 ± 8.31) and no change in distal (12.91 ± 5.31) regions. Transplantation did not significantly alter satellite cell density in host muscle regions compared to No Treatment. Juvenile transplant recipients had densities of 4.96 ± 5.21 (defect), 26.01 ± 10.40 (border), and 9.53 ± 5.40 (distal). Adult transplant values were similar: 2.24 ± 1.56, 31.29 ± 10.69, and 8.76 ± 4.55, respectively. As expected, satellite cell density was significantly elevated within the GFP⁺ transplant region: 27.11 ± 13.23 cells/mm² in juvenile and 33.63 ± 14.88 in adult recipients. These values were significantly higher than in control regions, suggesting that the transplants retained a rich satellite cell microenvironment. Surprisingly, satellite cell content was not different between adult and juvenile groups in the transplant region, nor did it alter content in the existing tissue regions. Centralized Nuclei The prevalence of centralized nuclei, an indicator of regenerating myofibers, was negligible in uninjured control muscle (0.60 ± 0.08%) but elevated in all VML injured muscle (Figure 6D). In No Treatment animals, centralized nuclei were highest in the border region (27.9 ± 4.6%) and lower in distal muscle (5.8 ± 1.2%). Transplantation produced similar regional patterns, with centralized nuclei enriched in the border and GFP⁺ regions, while distal muscle remained low. Adult transplants contained 20.7 ± 3.1% centralized nuclei in the border, 15.0 ± 2.8% in the GFP⁺ transplant region, and 4.4 ± 1.0% distally. Juvenile transplants exhibited higher regenerative activity, with 29.3 ± 5.0% centralized nuclei in the border and 23.3 ± 4.2% in the GFP⁺ region, while distal muscle remained low (6.5 ± 1.4%). Centralized nuclei were not quantified in the defect region due to the limited number of small regenerating fibers present in that region. Together, these results show that centralized nuclei and satellite cells were concentrated within and adjacent to the transplant, consistent with localized regeneration. Juvenile transplants displayed higher levels of centralized nuclei in both the border and GFP⁺ regions compared to adult transplants, suggesting enhanced but spatially restricted regenerative activity at seven weeks post-surgery. Of note, because cytosolic GFP is water-soluble, aqueous steps during the immunohistochemistry process can minimize GFP fluorescence, a process often referred to as “leaching”. To avoid over-interpretation, quantifications of GFP⁺ area, GFP⁺ fiber CSA, satellite cell and central nuclei counts related to GFP areas were performed only on sections with visually detectable GFP signal; samples where GFP was not clearly detected were excluded from GFP-specific analyses but were used for all other outcomes. DISCUSSION This study provides the first direct in-vivo comparison of juvenile versus adult skeletal muscle transplants for the treatment of VML. We found that juvenile donor tissue possessed markedly higher satellite cell density, smaller myofibers (allowing the transfer of more fibers per area), and enhanced in-vitro differentiation capacity compared to adult tissue. Despite these cellular advantages, both juvenile and adult transplants restored myofiber number and partially improved muscle strength to a similar extent seven weeks post-surgery. Functional recovery remained incomplete in both groups, and donor fibers were predominantly small at this time point. These findings indicate that donor muscle age influences intrinsic regenerative properties but does not necessarily translate into superior short-term, in-vivo outcomes in the absence of additional regenerative cues. In Study 1, our developmental analysis confirmed well-established age-related trends in muscle biology: nuclear and satellite cell densities were highest in juvenile muscle and declined progressively with maturation, consistent with prior studies in rodents and humans (10-12). The smaller fiber CSA and higher myogenic differentiation capacity of juvenile myoblasts support the concept that this tissue retains a more growth-permissive phenotype, which has been linked to increased adaptability and regeneration in other contexts (21-23). These properties provided a strong rationale for testing juvenile muscle as a donor source in VML repair. In Study 2, both juvenile and adult transplants integrated into the defect site, contributed GFP + donor fibers, and restored fiber counts to that of controls. Functional gains relative to untreated VML ranged from 35-50%, comparable to improvements reported in previous autologous or minced muscle transplant studies (7, 8). The lack of significant differences between juvenile and adult groups may be explained by several factors. First, the juvenile advantage in satellite cell number may have been transient; by seven weeks, many donor satellite cells could have differentiated into myofibers, leading to normalization between groups. Second, the immature size of donor fibers in both groups suggests that hypertrophy was still incomplete at this time point, limiting functional recovery. Finally, the restrictive microenvironment of the VML defect, characterized by altered extracellular matrix composition, persistent denervation, and limited vascularization (2, 5, 6), may have constrained the integration of juvenile tissue’s full regenerative potential. Satellite cell analyses reinforced this interpretation. In both donor age groups, Pax7⁺ cell enrichment was largely confined to the transplant region, with no apparent expansion into surrounding host muscle. This localized effect aligns with previous observations that satellite cells preferentially remain within areas containing intact basal lamina and supportive extracellular matrix (18, 24). Without such cues, migration into the broader defect is limited, which likely contributed to the absence of greater regenerative effects. Centralized nuclei findings further support this interpretation. Both transplant groups demonstrated localized enrichment of regenerating fibers within the border and GFP⁺ transplant regions, whereas distal muscle remained largely unaffected. Juvenile recipients exhibited higher proportions of centralized nuclei in both the border and transplant regions compared to adult recipients, suggesting more robust regenerative activity. However, these differences did not correspond to superior muscle size or force at seven weeks, indicating that the regenerative process was still incomplete and may require longer time frames or additional cues to translate into functional benefit. From a translational perspective, these results underscore that donor tissue composition, while important, is only one determinant of transplant efficacy. The persistence of small-diameter fibers and limited functional recovery in both groups suggest that mechanical loading and pro-hypertrophic stimuli are needed to maximize transplant benefit (18-20). Rehabilitation strategies such as voluntary wheel running, resistance exercise, or electrical stimulation have been shown to enhance myofiber hypertrophy, satellite cell activation, and neuromuscular integration after injury (19, 20). Incorporating such interventions into future transplant protocols could amplify the intrinsic advantages of juvenile-like donor tissue. Overall, this work is limited by several factors: first, we assessed the regenerative outcomes using a single time point of seven weeks. While this was intended to capture the regenerative window for muscle injury, it could be too early to detect long-term integration of the transplant, such as hypertrophy, reinnervation, and matrix remodeling. A longer follow-up may reveal whether age-related advantages or transplant integration change as fibers mature. Second, we deliberately implemented a transplant-only strategy, to isolate donor-age effects under controlled conditions. This baseline now creates a platform to test whether targeted loading (rehabilitation) or additional biological cues can enhance the benefits of juvenile tissue. Third, donor integration was quantified using GFP-based histology, which is susceptible to leaking; future work could incorporate other tracking methods, such as tissue clearing with 3D quantification. Finally, experiments were performed in male Lewis rats to mirror the male predominance of combat-related VML; confirming these findings in females and across additional genetic backgrounds will be an important step toward broad translation. CONCLUSION Juvenile skeletal muscle displays cellular and structural characteristics consistent with high regenerative potential, including high satellite cell content, small myofibers, greater in-vitro differentiation capacity, and elevated proportions of centralized nuclei within transplant regions. In this VML model, both juvenile and adult donor tissue integrated into the defect, restored myofiber number, and partially improved muscle strength; however, functional outcomes were similar between groups at seven weeks post-surgery. These results indicate that optimizing donor age alone is insufficient for full functional recovery and that additional regenerative cues, or time, are required. The high regenerative profile of juvenile muscle provides a valuable biological benchmark for engineered muscle constructs. Myogenic progenitors derived from induced pluripotent stem cells or other stem cell sources could be directed toward a juvenile-like phenotype before transplantation (21, 22, 25). Additionally, emerging work with human–animal chimeric models offer a promising and scalable approach to sourcing juvenile-like transplant tissue (23, 26). Such strategies may reduce logistical barriers, including long-term animal care costs, and accelerate tissue availability. When paired with biomaterials and/or with rehabilitation interventions that promote hypertrophy and integration, these approaches could substantially improve structural and functional outcomes for patients with VML. Abbreviations CSA cross-sectional area DAPI 4′,6-diamidino-2-phenylindole EDL extensor digitorum longus FBS fetal bovine serum GFP green fluorescent protein IACUC Institutional Animal Care and Use Committee IHC immunohistochemistry MyHC myosin heavy chain NGS normal goat serum Pax7 paired box protein 7 PBS phosphate-buffered saline TA tibialis anterior VML volumetric muscle loss WGA wheat germ agglutinin Declarations Ethics Approval and Consent to Participate All animal procedures were approved on April 1, 2023, by the Institutional Animal Care and Use Committee (IACUC) at Brigham Young University under the title: A novel approach to improve skeletal muscle transplant efficiency following volumetric muscle loss injury, protocol number 23-0401. All methods were carried out in accordance with relevant guidelines and regulations for the care and use of laboratory animals. Consent for Publication Not applicable. Availability of Data and Materials The datasets generated and/or analyzed during the current study are available from the corresponding author on reasonable request. Competing Interests The authors declare that they have no competing interests. Funding This research was supported by internal funding from the College of Life Sciences at Brigham Young University. The funding body had no role in the design of the study, data collection, analysis, interpretation, or manuscript writing. Authors’ Contributions JRS conceived and designed the study. JJP, SRF, ZHR, MJM, ACT, KMJ, TKS, and MKK performed experiments and data collection. SSH, and EWE, contributed to data analysis. JRS drafted the manuscript. All authors contributed to the manuscript revision and approved the final version of the manuscript. Acknowledgements The authors would like to thank Anson Wood for piloting the cell culture model, Kayli Denmark for piloting IHC protocols, Chandler Bowen, and Dre Sorensen for assistance in image analysis, and the use of AI for assistance with readability, grammar and punctuation. References Greising SM, Rivera JC, Goldman SM, Watts A, Aguilar CA, Corona BT. Unwavering pathobiology of volumetric muscle loss injury. Scientific Reports. 2017;7(1):13179. Hoffman DB, Raymond-Pope CJ, Sorensen JR, Corona BT, Call JA, Greising SM. Temporal changes in the muscle extracellular matrix due to volumetric muscle loss injury. Connective Tissue Research. 2022;63(2):124–37. Brack AS, Rando TA. Intrinsic changes and extrinsic influences of myogenic stem cell function during aging. Stem Cell Reviews and Reports. 2007;3(3):226–37. Kuang S, Kuroda K, Le Grand F, Rudnicki MA. Asymmetric self-renewal and commitment of satellite stem cells in muscle. Cell. 2007;129(5):999–1010. Garg K, Ward CL, Hurtgen BJ, Wilken JM, Stinner DJ, Wenke JC, et al. Volumetric muscle loss: persistent functional deficits beyond frank loss of tissue. Journal of Orthopaedic Research. 2015;33(1):40–6. Sorensen JR, Hoffman DB, Corona BT, Greising SM. Secondary denervation is a chronic pathophysiologic sequela of volumetric muscle loss. Journal of Applied Physiology. 2021;130(5):1552–65. Grogan BF, Hsu JR. Volumetric muscle loss. Journal of the American Academy of Orthopaedic Surgeons. 2011;19(Suppl 1):S35–S7. Ward CL, Ji L, Corona BT. Autologous minced muscle grafts improve muscle strength in a porcine model of volumetric muscle loss injury. Journal of Applied Physiology. 2016;120(6):915–23. Corona BT, Rivera JC, Owens JG, Wenke JC, Rathbone CR. Volumetric muscle loss leads to permanent disability following extremity trauma. Journal of Rehabilitation Research and Development. 2015;52(7):785–92. García-Prat L, Muñoz-Cánoves P. Muscle stem cell aging: regulation and rejuvenation. Skeletal Muscle. 2021;11(1):4. Lexell J, Taylor CC, Sjöström M. What is the cause of the ageing atrophy? Total number, size and proportion of different fiber types studied in whole vastus lateralis muscle from 15‐ to 83‐year‐old men. Journal of the Neurological Sciences. 1988;84(2-3):275–94. Schiaffino S, Reggiani C. Fiber types in mammalian skeletal muscles. Physiological Reviews. 2011;91(4):1447–531. Smith LR, Barton ER. Collagen content does not alter the passive mechanical properties of fibrotic skeletal muscle in mdx mice. American Journal of Physiology-Cell Physiology. 2014;306(10):C889–C98. Snow MH. Satellite cell response in rat soleus muscle undergoing hypertrophy due to surgical ablation of synergists. Anatomical Record. 1977;189(2):479–97. Chal J, Oginuma M, Al Tanoury Z, Gobert B, Sumara O, Hick A, et al. Differentiation of pluripotent stem cells to muscle fiber to model Duchenne muscular dystrophy. Nature Biotechnology. 2015;33(9):962–9. Hoffman DB, Basten AM, Sorensen JR, Raymond-Pope CJ, Lillquist TJ, Call JA, et al. Response of terminal Schwann cells following volumetric muscle loss injury. Experimental Neurology. 2023;357:114431. Sorensen JR, Hoffman DB, Raymond-Pope CJ, Lillquist TJ, Russell AM, Corona BT, et al. Inhibition of ErbB2 mitigates secondary denervation after traumatic muscle injury. Journal of Physiology. 2025;0(0):1–18. Greising SM, Call JA. When is the right time to initiate rehabilitation? Time will tell…. Exp Physiol. 2024;109(6):889–91. Nakayama KH, Quarta M, Paine P, Alcazar C, Karakikes I, Garcia V, et al. Rehabilitation following skeletal muscle injury enhances tissue regeneration and function. NPJ Regenerative Medicine. 2019;4:3. Serrano AL, Baeza-Raja B, Perdiguero E, Jardí M, Muñoz-Cánoves P. Interleukin-6 is an essential regulator of satellite cell-mediated skeletal muscle hypertrophy. Cell Metabolism. 2008;7(1):33–44. Carosio S, Barberi L, Rizzuto E, Nicoletti C, Musarò A. Generation of eX vivo-vascularized Muscle Engineered Tissue (X-MET). Scientific Reports. 2013;3:1420. Chal J, Oginuma M, Al Tanoury Z, Gobert B, Sumara O, Hick A, et al. Differentiation of pluripotent stem cells to muscle fiber to model Duchenne muscular dystrophy. Nature biotechnology. 2015;33(9):962–9. Maeng G, Das S, Greising SM, Gong W, Singh BN, Kren S, et al. Humanized skeletal muscle in MYF5/MYOD/MYF6-null pig embryos. Nature biomedical engineering. 2021;5(8):805–14. Greising SM, Dearth CL, Corona BT. Regenerative and rehabilitative medicine: a necessary synergy for functional recovery from volumetric muscle loss injury. Cells Tissues Organs. 2016;202(3-4):237–49. Rao L, Qian Y, Khodabukus A, Ribar T, Bursac N. Engineering human pluripotent stem cells into a functional skeletal muscle tissue. Nature Communications. 2018;9:126. Choe Y-H, Das S, Ma X, Lee H, Sorensen JR, Hoffman DB, et al. Porcine myogenesis in cloned wildtype and MYF5/MYOD/MYF6-null porcine embryo. Communications biology. 2025;8(1):217. Additional Declarations No competing interests reported. Cite Share Download PDF Status: Published Journal Publication published 03 Dec, 2025 Read the published version in Stem Cell Research & Therapy → Version 1 posted Editorial decision: Revision requested 13 Oct, 2025 Reviews received at journal 09 Oct, 2025 Reviews received at journal 02 Oct, 2025 Reviewers agreed at journal 29 Sep, 2025 Reviewers agreed at journal 23 Sep, 2025 Reviewers invited by journal 22 Sep, 2025 Editor assigned by journal 18 Sep, 2025 Submission checks completed at journal 10 Sep, 2025 First submitted to journal 08 Sep, 2025 You are reading this latest preprint version Research Square lets you share your work early, gain feedback from the community, and start making changes to your manuscript prior to peer review in a journal. As a division of Research Square Company, we’re committed to making research communication faster, fairer, and more useful. We do this by developing innovative software and high quality services for the global research community. Our growing team is made up of researchers and industry professionals working together to solve the most critical problems facing scientific publishing. Also discoverable on Platform About Our Team In Review Editorial Policies Advisory Board Help Center Resources Author Services Accessibility API Access RSS feed Manage Cookie Preferences © Research Square 2026 | ISSN 2693-5015 (online) Privacy Policy Terms of Service Do Not Sell My Personal Information {"props":{"pageProps":{"initialData":{"identity":"rs-7437055","acceptedTermsAndConditions":true,"allowDirectSubmit":false,"archivedVersions":[],"articleType":"Research Article","associatedPublications":[],"authors":[{"id":523990846,"identity":"a495dcd0-6696-48e5-9685-6a5fe9bda6ca","order_by":0,"name":"John J. Payne","email":"","orcid":"","institution":"Brigham Young University","correspondingAuthor":false,"prefix":"","firstName":"John","middleName":"J.","lastName":"Payne","suffix":""},{"id":523990847,"identity":"a2dfea92-18b3-429a-8d02-8a526db7bb8c","order_by":1,"name":"Samuel R. Frandsen","email":"","orcid":"","institution":"Brigham Young University","correspondingAuthor":false,"prefix":"","firstName":"Samuel","middleName":"R.","lastName":"Frandsen","suffix":""},{"id":523990848,"identity":"54b4950c-b45e-47a7-9733-d46297790884","order_by":2,"name":"Zachary H. Rasmussen","email":"","orcid":"","institution":"Brigham Young University","correspondingAuthor":false,"prefix":"","firstName":"Zachary","middleName":"H.","lastName":"Rasmussen","suffix":""},{"id":523990849,"identity":"7ace9e28-63a1-4d8c-950c-41cedffc5946","order_by":3,"name":"Matthew J. Mangus","email":"","orcid":"","institution":"Brigham Young University","correspondingAuthor":false,"prefix":"","firstName":"Matthew","middleName":"J.","lastName":"Mangus","suffix":""},{"id":523990851,"identity":"fd8b8024-f2b7-41b6-8183-037a0245d894","order_by":4,"name":"Anna C. Taylor","email":"","orcid":"","institution":"Brigham Young University","correspondingAuthor":false,"prefix":"","firstName":"Anna","middleName":"C.","lastName":"Taylor","suffix":""},{"id":523990853,"identity":"a44ef1a1-4c66-4a43-9337-7fbe53d9b69b","order_by":5,"name":"Mason K. Kephart","email":"","orcid":"","institution":"Brigham Young University","correspondingAuthor":false,"prefix":"","firstName":"Mason","middleName":"K.","lastName":"Kephart","suffix":""},{"id":523990855,"identity":"021d8fe1-aa1c-44be-bf46-818ce015a718","order_by":6,"name":"Sandy S. Huang","email":"","orcid":"","institution":"Brigham Young University","correspondingAuthor":false,"prefix":"","firstName":"Sandy","middleName":"S.","lastName":"Huang","suffix":""},{"id":523990858,"identity":"596b6a1e-5e0c-45ad-9257-11e7bd9ef9ee","order_by":7,"name":"Thomas K. Schiefer","email":"","orcid":"","institution":"Brigham Young University","correspondingAuthor":false,"prefix":"","firstName":"Thomas","middleName":"K.","lastName":"Schiefer","suffix":""},{"id":523990861,"identity":"79981ee3-8f0e-41eb-a87d-89f8dd4bec2e","order_by":8,"name":"Kyndal M. Jones","email":"","orcid":"","institution":"Brigham Young University","correspondingAuthor":false,"prefix":"","firstName":"Kyndal","middleName":"M.","lastName":"Jones","suffix":""},{"id":523990862,"identity":"0282dfa6-6cd6-4ea1-be6f-ae696c7a0af9","order_by":9,"name":"Erastus W. Evans","email":"","orcid":"","institution":"Brigham Young University","correspondingAuthor":false,"prefix":"","firstName":"Erastus","middleName":"W.","lastName":"Evans","suffix":""},{"id":523990864,"identity":"c53274b8-7b25-4342-9edf-82d42058032d","order_by":10,"name":"Jacob R. Sorensen","email":"data:image/png;base64,iVBORw0KGgoAAAANSUhEUgAAAZAAAAAyAQMAAABI0h/eAAAABlBMVEX///8AAABVwtN+AAAACXBIWXMAAA7EAAAOxAGVKw4bAAAA0ElEQVRIiWNgGAWjYBACxgY4k/kYhD5AvBa2NOK0IAEeM+K0MM8+/vBxRcUde37pnm+PedsY5PhuJBBwWF+OseGZM88SZ845u90YqMVYkqCWHh42yca2wwkGN3K3SQO1JG4grIX9mWTjv8P29jdynoG01BOhhcFMsrHhMOMGiRw2kBagdYQdZmzYcOxw4owbaeaGc85JGM488wC/FsMe9ocPG2oO2/PPSH724E2ZjTzfcQK2GDYgcZh4GCTwKwcBeRRX/iCsYRSMglEwCkYgAACMAUVuhZ3SDgAAAABJRU5ErkJggg==","orcid":"","institution":"Brigham Young University","correspondingAuthor":true,"prefix":"","firstName":"Jacob","middleName":"R.","lastName":"Sorensen","suffix":""}],"badges":[],"createdAt":"2025-08-22 19:23:17","currentVersionCode":1,"declarations":"","doi":"10.21203/rs.3.rs-7437055/v1","doiUrl":"https://doi.org/10.21203/rs.3.rs-7437055/v1","draftVersion":[],"editorialEvents":[{"content":"https://doi.org/10.1186/s13287-025-04844-y","type":"published","date":"2025-12-03T15:57:57+00:00"}],"editorialNote":"","failedWorkflow":false,"files":[{"id":92845743,"identity":"56bced66-245c-4397-9154-6fe697477989","added_by":"auto","created_at":"2025-10-06 09:34:18","extension":"jpg","order_by":0,"title":"","display":"","copyAsset":false,"role":"acdc-reference","size":5495621,"visible":true,"origin":"","legend":"","description":"","filename":"Figure1.TibialisanteriorTAmusclestructureandmyofibersvarybyage.jpg","url":"https://assets-eu.researchsquare.com/files/rs-7437055/v1/c258085a57cbc281fe1e566b.jpg"},{"id":92845264,"identity":"b628d895-4d53-43b3-af00-1ebc87dd6a10","added_by":"auto","created_at":"2025-10-06 09:26:18","extension":"docx","order_by":1,"title":"","display":"","copyAsset":false,"role":"acdc-reference","size":89617,"visible":true,"origin":"","legend":"","description":"","filename":"JuvenilevsAdultSkeletalMuscleTransplantsintheTreatmentofVolumetricMuscleLossInjury.docx","url":"https://assets-eu.researchsquare.com/files/rs-7437055/v1/2fd83dab6fd9d359fb4cf5fd.docx"},{"id":92845265,"identity":"b6fdcfa7-6c02-4a74-ad7e-7003b671a749","added_by":"auto","created_at":"2025-10-06 09:26:18","extension":"jpg","order_by":2,"title":"","display":"","copyAsset":false,"role":"acdc-reference","size":8956439,"visible":true,"origin":"","legend":"","description":"","filename":"Figure2.Nucleimyofiberandsatellitecelldensitydeclinewithage.jpg","url":"https://assets-eu.researchsquare.com/files/rs-7437055/v1/6cef21cae085b2cf4d786674.jpg"},{"id":92845263,"identity":"ade90b15-ffd3-449c-bd49-419deaf15813","added_by":"auto","created_at":"2025-10-06 09:26:18","extension":"jpg","order_by":3,"title":"","display":"","copyAsset":false,"role":"acdc-reference","size":3767436,"visible":true,"origin":"","legend":"","description":"","filename":"Figure3.Myoblastpurityproliferationanddifferentiationacrossdonoragegroups.jpg","url":"https://assets-eu.researchsquare.com/files/rs-7437055/v1/82f1a79d12772cc9abf8ccb1.jpg"},{"id":92845259,"identity":"f2a17e02-7452-4bc6-81cd-605f0ca95e87","added_by":"auto","created_at":"2025-10-06 09:26:17","extension":"jpg","order_by":4,"title":"","display":"","copyAsset":false,"role":"acdc-reference","size":5788263,"visible":true,"origin":"","legend":"","description":"","filename":"Figure4.MuscleforceandmyofibermorphologyfollowingVMLinjuryandmuscletransplantation.jpg","url":"https://assets-eu.researchsquare.com/files/rs-7437055/v1/810371248ddc4bc576eb1ee8.jpg"},{"id":92845256,"identity":"f60ed2e7-d5c2-4473-af19-9dd1d7c1078e","added_by":"auto","created_at":"2025-10-06 09:26:17","extension":"jpg","order_by":5,"title":"","display":"","copyAsset":false,"role":"acdc-reference","size":4984040,"visible":true,"origin":"","legend":"","description":"","filename":"Figure5.DonorfiberGFPintegrationandmorphologyfollowingjuvenileandadultmuscletransplantation.jpg","url":"https://assets-eu.researchsquare.com/files/rs-7437055/v1/d1a0f77a47355a17faadeaf8.jpg"},{"id":92845272,"identity":"eaafb5f2-aff1-45cf-be3d-fe4fe14f2778","added_by":"auto","created_at":"2025-10-06 09:26:18","extension":"jpg","order_by":6,"title":"","display":"","copyAsset":false,"role":"acdc-reference","size":8087618,"visible":true,"origin":"","legend":"","description":"","filename":"Figure6.SatellitecelldensityacrossregionsofVMLinjuredmusclefollowingtransplantation.jpg","url":"https://assets-eu.researchsquare.com/files/rs-7437055/v1/ded77ce29bb53944b0efbc82.jpg"},{"id":92845267,"identity":"ec7d8305-5f4c-43e3-84e5-f147a974eb3c","added_by":"auto","created_at":"2025-10-06 09:26:18","extension":"json","order_by":7,"title":"","display":"","copyAsset":false,"role":"acdc-reference","size":12613,"visible":true,"origin":"","legend":"","description":"","filename":"134e6576e18c4823bc5970652df0b97e.json","url":"https://assets-eu.researchsquare.com/files/rs-7437055/v1/c1e551306af9771ad4d19db1.json"},{"id":92845745,"identity":"f23d529e-419d-4dbb-bce4-c4af38891650","added_by":"auto","created_at":"2025-10-06 09:34:19","extension":"xml","order_by":8,"title":"","display":"","copyAsset":false,"role":"acdc-reference","size":118043,"visible":true,"origin":"","legend":"","description":"","filename":"134e6576e18c4823bc5970652df0b97e1enriched.xml","url":"https://assets-eu.researchsquare.com/files/rs-7437055/v1/bb649e45c81f8a9b5c1a673e.xml"},{"id":92845274,"identity":"c3436d92-00a1-449a-8b0d-a9c680e78a60","added_by":"auto","created_at":"2025-10-06 09:26:18","extension":"jpg","order_by":9,"title":"","display":"","copyAsset":false,"role":"acdc-reference","size":5495621,"visible":true,"origin":"","legend":"","description":"","filename":"Figure1.TibialisanteriorTAmusclestructureandmyofibersvarybyage.jpg","url":"https://assets-eu.researchsquare.com/files/rs-7437055/v1/fdb171a367245b01035ee8de.jpg"},{"id":92845746,"identity":"f16c8364-671a-4077-b061-ff655f2d5883","added_by":"auto","created_at":"2025-10-06 09:34:19","extension":"jpg","order_by":10,"title":"","display":"","copyAsset":false,"role":"acdc-reference","size":8956439,"visible":true,"origin":"","legend":"","description":"","filename":"Figure2.Nucleimyofiberandsatellitecelldensitydeclinewithage.jpg","url":"https://assets-eu.researchsquare.com/files/rs-7437055/v1/0b6a024570edac9cf95242c9.jpg"},{"id":92845262,"identity":"df904a59-e369-43a2-bb95-1f966bc3dca0","added_by":"auto","created_at":"2025-10-06 09:26:18","extension":"jpg","order_by":11,"title":"","display":"","copyAsset":false,"role":"acdc-reference","size":3767436,"visible":true,"origin":"","legend":"","description":"","filename":"Figure3.Myoblastpurityproliferationanddifferentiationacrossdonoragegroups.jpg","url":"https://assets-eu.researchsquare.com/files/rs-7437055/v1/8979e2b7a5ac7388f83bad8f.jpg"},{"id":92845278,"identity":"e440b472-9dee-45c9-9efb-409fed9d0989","added_by":"auto","created_at":"2025-10-06 09:26:19","extension":"jpg","order_by":12,"title":"","display":"","copyAsset":false,"role":"acdc-reference","size":5788263,"visible":true,"origin":"","legend":"","description":"","filename":"Figure4.MuscleforceandmyofibermorphologyfollowingVMLinjuryandmuscletransplantation.jpg","url":"https://assets-eu.researchsquare.com/files/rs-7437055/v1/5a6edd05ebf7e8d7128e42f3.jpg"},{"id":92845282,"identity":"9d950286-c503-47eb-954c-449d23f86b21","added_by":"auto","created_at":"2025-10-06 09:26:19","extension":"jpg","order_by":13,"title":"","display":"","copyAsset":false,"role":"acdc-reference","size":4984040,"visible":true,"origin":"","legend":"","description":"","filename":"Figure5.DonorfiberGFPintegrationandmorphologyfollowingjuvenileandadultmuscletransplantation.jpg","url":"https://assets-eu.researchsquare.com/files/rs-7437055/v1/2f7baff405e06b5aa5d26f7f.jpg"},{"id":92845280,"identity":"04c8beff-2715-4ae4-9312-73ca99b10c23","added_by":"auto","created_at":"2025-10-06 09:26:19","extension":"jpg","order_by":14,"title":"","display":"","copyAsset":false,"role":"acdc-reference","size":8087618,"visible":true,"origin":"","legend":"","description":"","filename":"Figure6.SatellitecelldensityacrossregionsofVMLinjuredmusclefollowingtransplantation.jpg","url":"https://assets-eu.researchsquare.com/files/rs-7437055/v1/c8a7db2e0445b8817abfc8ee.jpg"},{"id":92845285,"identity":"1925c15d-34ba-4cc5-ae93-7fb7a47df287","added_by":"auto","created_at":"2025-10-06 09:26:19","extension":"png","order_by":15,"title":"","display":"","copyAsset":false,"role":"acdc-reference","size":878483,"visible":true,"origin":"","legend":"","description":"","filename":"OnlineFigure1.TibialisanteriorTAmusclestructureandmyofibersvarybyage.png","url":"https://assets-eu.researchsquare.com/files/rs-7437055/v1/fb41225165f43d8ec4524a4e.png"},{"id":92845288,"identity":"8f423ba8-2b79-4b40-8bb4-9b88659f318b","added_by":"auto","created_at":"2025-10-06 09:26:20","extension":"png","order_by":16,"title":"","display":"","copyAsset":false,"role":"acdc-reference","size":1010791,"visible":true,"origin":"","legend":"","description":"","filename":"OnlineFigure2.Nucleimyofiberandsatellitecelldensitydeclinewithage.png","url":"https://assets-eu.researchsquare.com/files/rs-7437055/v1/8c369dbd20b40958113d1c11.png"},{"id":92845287,"identity":"8dcf3f07-5782-4c9f-b295-ab7e08aec5fc","added_by":"auto","created_at":"2025-10-06 09:26:19","extension":"png","order_by":17,"title":"","display":"","copyAsset":false,"role":"acdc-reference","size":307491,"visible":true,"origin":"","legend":"","description":"","filename":"OnlineFigure3.Myoblastpurityproliferationanddifferentiationacrossdonoragegroups.png","url":"https://assets-eu.researchsquare.com/files/rs-7437055/v1/c1e35a10095c108b27673ea5.png"},{"id":92845275,"identity":"a79d6ad8-f94d-4a92-984b-b213e3e49526","added_by":"auto","created_at":"2025-10-06 09:26:18","extension":"png","order_by":18,"title":"","display":"","copyAsset":false,"role":"acdc-reference","size":805703,"visible":true,"origin":"","legend":"","description":"","filename":"OnlineFigure4.MuscleforceandmyofibermorphologyfollowingVMLinjuryandmuscletransplantation.png","url":"https://assets-eu.researchsquare.com/files/rs-7437055/v1/1c0e8da91529c2767a973501.png"},{"id":92845276,"identity":"da7bd9e9-3756-42d8-a68b-516e9b52a6aa","added_by":"auto","created_at":"2025-10-06 09:26:19","extension":"png","order_by":19,"title":"","display":"","copyAsset":false,"role":"acdc-reference","size":691173,"visible":true,"origin":"","legend":"","description":"","filename":"OnlineFigure5.DonorfiberGFPintegrationandmorphologyfollowingjuvenileandadultmuscletransplantation.png","url":"https://assets-eu.researchsquare.com/files/rs-7437055/v1/b6afb3f51e15848d856d2e0f.png"},{"id":92845279,"identity":"3135b55c-25db-431b-a4b6-7279fb3765bd","added_by":"auto","created_at":"2025-10-06 09:26:19","extension":"png","order_by":20,"title":"","display":"","copyAsset":false,"role":"acdc-reference","size":852983,"visible":true,"origin":"","legend":"","description":"","filename":"OnlineFigure6.SatellitecelldensityacrossregionsofVMLinjuredmusclefollowingtransplantation.png","url":"https://assets-eu.researchsquare.com/files/rs-7437055/v1/c22d51599585003e2b4a9d45.png"},{"id":92845284,"identity":"f791943d-b227-4635-9748-a00f68aa5480","added_by":"auto","created_at":"2025-10-06 09:26:19","extension":"xml","order_by":21,"title":"","display":"","copyAsset":false,"role":"acdc-reference","size":122382,"visible":true,"origin":"","legend":"","description":"","filename":"134e6576e18c4823bc5970652df0b97e1structuring.xml","url":"https://assets-eu.researchsquare.com/files/rs-7437055/v1/9401fe1dfe1b46d34354eb27.xml"},{"id":92845744,"identity":"56fdae7d-b9fe-445f-a7c9-fb0dd38b80f3","added_by":"auto","created_at":"2025-10-06 09:34:19","extension":"html","order_by":22,"title":"","display":"","copyAsset":false,"role":"acdc-reference","size":132938,"visible":true,"origin":"","legend":"","description":"","filename":"earlyproof.html","url":"https://assets-eu.researchsquare.com/files/rs-7437055/v1/32c5958478f7b3b77fb0d76f.html"},{"id":92845742,"identity":"e365bf99-740c-4fce-83a1-e3a68bec2729","added_by":"auto","created_at":"2025-10-06 09:34:18","extension":"jpg","order_by":1,"title":"Figure 1","display":"","copyAsset":false,"role":"figure","size":5495621,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eTibialis anterior (TA) muscle structure and myofibers vary by age.\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(A)\u003c/strong\u003e Representative transverse cross-sections of the tibialis anterior (TA) muscle from juvenile (21-day), adolescent (34-day), and adult (120-day) male Lewis rats. Individual myofibers are pseudo-colored based on segmentation from laminin staining. \u003cstrong\u003e(B)\u003c/strong\u003e Whole-muscle CSA increased stepwise with age, more than doubling from juvenile to adolescent (+144%, p = 0.0012) and nearly tripling again from adolescent to adult (+181%, p = 0.0003). Adult CSA was ~7-fold greater than juvenile (p \u0026lt; 0.0001). \u003cstrong\u003e(C)\u003c/strong\u003e Total myofiber number per muscle did not significantly differ between groups (p=0.1621, indicating growth was primarily driven by fiber hypertrophy rather than hyperplasia. \u003cstrong\u003e(D)\u003c/strong\u003eAverage myofiber CSA increased ~2-fold from juvenile to adolescent (p = 0.0267) and nearly 3-fold again from adolescent to adult (p \u0026lt; 0.0001). Adult fibers were more than 6-fold larger than juvenile (p \u0026lt; 0.0001). \u003cstrong\u003e(E)\u003c/strong\u003eFrequency distribution of myofiber CSA further illustrates that juvenile muscle exhibited a narrow, left-shifted distribution, while adolescent and adult muscles showed progressively broader and right-shifted profiles. Data are presented as mean ± SD; n = 4 per group. Statistical significance: ****p \u0026lt; 0.0001, **p \u0026lt; 0.01, *p \u0026lt; 0.05. Scale bar = 1mm\u003c/p\u003e","description":"","filename":"Figure1.TibialisanteriorTAmusclestructureandmyofibersvarybyage.jpg","url":"https://assets-eu.researchsquare.com/files/rs-7437055/v1/12d6173a4173d28c4a857ee7.jpg"},{"id":92845271,"identity":"2d2da85e-a35b-4f13-9f0d-84fda0b9a440","added_by":"auto","created_at":"2025-10-06 09:26:18","extension":"jpg","order_by":2,"title":"Figure 2","display":"","copyAsset":false,"role":"figure","size":8956439,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eNuclei, myofiber, and satellite cell density decline with age.\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(A)\u003c/strong\u003e Representative whole-muscle cross-sections of the tibialis anterior (TA) from juvenile (21-day), adolescent (34-day), and adult (120-day) male rats. \u003cstrong\u003e(B)\u003c/strong\u003e High-magnification images show DAPI-stained nuclei (blue), laminin-labeled myofiber membranes (white), and Pax7⁺ satellite cells (red). \u003cstrong\u003e(C)\u003c/strong\u003eQuantification of nuclear density (nuclei/mm²) revealed a significant stepwise decline with age: adolescents had ~47% fewer nuclei than juveniles (p \u0026lt; 0.0001), and adults had ~55% fewer than adolescents (p \u0026lt; 0.0001), leaving adult muscle with less than one-quarter of juvenile density overall. \u003cstrong\u003e(D)\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eMyofiber density (fibers/mm²) also decreased significantly with age, dropping ~50% from juvenile to adolescent (p \u0026lt; 0.001) and another ~50% from adolescent to adult (p \u0026lt; 0.001), consistent with growth by fiber hypertrophy. (E) Satellite cell density (Pax7⁺ cells/mm²) was ~3.5-fold higher in juveniles compared to adolescents (p \u0026lt; 0.001) and ~15-fold higher than in adults (p \u0026lt; 0.0001). Although adolescent satellite cell density appeared greater than adult, this difference did not reach significance (p = 0.1049). Data are presented as mean ± SD, n = 4 animals per group. Statistical comparisons were made using one-way ANOVA followed by Tukey’s post hoc test. Statistical significance: ****p \u0026lt; 0.0001, ***p \u0026lt; 0.001, **p \u0026lt; 0.01. Scale bars = 1mm (A), 250µm (B).\u003c/p\u003e","description":"","filename":"Figure2.Nucleimyofiberandsatellitecelldensitydeclinewithage.jpg","url":"https://assets-eu.researchsquare.com/files/rs-7437055/v1/a36e73fc320d72e650edc5b4.jpg"},{"id":92845741,"identity":"3c0597d1-d2d5-43f2-bd45-f45c870ec6f6","added_by":"auto","created_at":"2025-10-06 09:34:18","extension":"jpg","order_by":3,"title":"Figure 3","display":"","copyAsset":false,"role":"figure","size":3767436,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eMyoblast purity, proliferation, and differentiation across donor age groups.\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(A)\u003c/strong\u003e Representative immunofluorescence image from an adult-derived myoblast culture showing co-localization of nuclear MyoD (red) with DAPI-stained nuclei (blue), confirming myogenic identity. \u003cstrong\u003e(B)\u003c/strong\u003eQuantification of MyoD⁺ nuclei indicated \u0026gt;95% purity in cultures from juvenile, adolescent, and adult donors, confirming successful isolation of committed myogenic cells. \u003cstrong\u003e(C)\u003c/strong\u003e EdU incorporation increased from ~15% at 4 h to ~80% by 16 h in all groups, with no age-related differences (p = 0.9696), indicating a similar proliferative capacity. \u0026nbsp;Representative images \u003cstrong\u003e(D)\u003c/strong\u003e of DAPI (blue) and EdU (white) staining illustrate comparable proliferative activity among juvenile, adolescent, and adult myoblasts at each time point. \u003cstrong\u003e(E)\u003c/strong\u003eAfter 72 h of differentiation, the fusion index (MyHC⁺ nuclei/total nuclei) was ~73% higher in juvenile vs adult myoblasts (p=0.0067). Adolescent cultures were intermediate and not significantly different from either group (p\u0026gt;0.1). \u003cstrong\u003e(F)\u003c/strong\u003e Total myotube formation, measured by MyHC⁺ area per field, was ~60–80% larger in juvenile compared to adolescent (p=0.0191) and adult (p=0.0074) myoblasts, respectively. No difference was observed between adolescent and adult groups (p = 0.6658). \u003cstrong\u003e(G)\u003c/strong\u003eThe nucleation index (nuclei/myotube) did not differ significantly across groups (p\u0026gt;0.05). \u003cstrong\u003e(H)\u003c/strong\u003e Representative images of differentiated myoblasts stained for DAPI (blue), Myogenin (white), and MyHC (red) highlight enhanced multinucleated myotube formation in juvenile cultures. Data are presented as mean ± SD. Values reflect the mean of 4–5 images per well, collected in triplicate wells (n = 3) for each group at each time point. \u0026nbsp;Statistical significance: **p \u0026lt; 0.01, *p \u0026lt; 0.05. Scale bars = 100 µm (A, H), 150 µm (D).\u003c/p\u003e","description":"","filename":"Figure3.Myoblastpurityproliferationanddifferentiationacrossdonoragegroups.jpg","url":"https://assets-eu.researchsquare.com/files/rs-7437055/v1/3882bbf2c788bbf675d7a7b2.jpg"},{"id":92845269,"identity":"63ccaa04-dae0-4bda-bc17-c8f8b75001cf","added_by":"auto","created_at":"2025-10-06 09:26:18","extension":"jpg","order_by":4,"title":"Figure 4","display":"","copyAsset":false,"role":"figure","size":5788263,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eMuscle force and myofiber morphology following VML injury and muscle transplantation.\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eIn vivo strength testing of the anterior compartment (tibialis anterior and extensor digitorum longus) revealed significant force deficits following VML, with partial recovery after transplantation. Peak isometric force \u003cstrong\u003e(A)\u003c/strong\u003e was highest in controls (20.96 ± 3.04 mN·m, n=19) and reduced by ~55% in untreated VML limbs (9.47 ± 0.67, n=8). Adult transplants restored force to 12.94 ± 1.52 (n=8; p = 0.0299 vs VML No Treatment), and juvenile transplants reached 14.25 ± 2.01 (n=8; p = 0.0016 vs VML No Treatment), representing a 36% and 50% increase, respectively (Adult vs Juvenile, p = 0.6992). Force traces \u003cstrong\u003e(B)\u003c/strong\u003e illustrate these differences. Body-weight–normalized force \u003cstrong\u003e(C)\u003c/strong\u003e showed a similar pattern. Histological analysis was performed on a subset of animals to explain differences in force (n = 4 per group). Representative TA cross-sections \u003cstrong\u003e(D)\u003c/strong\u003e show severe tissue loss in untreated VML, while both transplant groups showed improved fiber retention but smaller fibers. Each color represents a pseudo-colored myofiber based on laminin staining. Myofiber quantification \u003cstrong\u003e(E–H)\u003c/strong\u003esupported these observations. Fiber number \u003cstrong\u003e(E)\u003c/strong\u003e was significantly reduced in untreated VML muscle compared to controls (p \u0026lt; 0.01), while both transplant groups restored myofibers, with juvenile recipients ~25% higher than adults. Total TA CSA (F) was also reduced in the No Treatment muscle (p = 0.0265 vs control) but preserved in both transplant groups (≥85% of control). Mean myofiber CSA (G) remained significantly smaller in all VML groups vs controls (p \u0026lt; 0.001), reflecting persistent immature fibers. The myofiber CSA distributions (H) showed a leftward shift toward small fibers (\u0026lt;1,000 µm²) in all VML groups, most pronounced in juveniles, suggesting ongoing regeneration but with limited hypertrophy in the transplanted tissue. Data are mean ± SD. Statistical significance: ****p \u0026lt; 0.0001, ***p \u0026lt; 0.001, **p \u0026lt; 0.01, p \u0026lt; 0.05. Scale bar = 1 mm.\u003c/p\u003e","description":"","filename":"Figure4.MuscleforceandmyofibermorphologyfollowingVMLinjuryandmuscletransplantation.jpg","url":"https://assets-eu.researchsquare.com/files/rs-7437055/v1/3fded443dd5e88d0a0fedd66.jpg"},{"id":92845273,"identity":"1aa3e468-f716-40f4-ba4f-e21231275fa3","added_by":"auto","created_at":"2025-10-06 09:26:18","extension":"jpg","order_by":5,"title":"Figure 5","display":"","copyAsset":false,"role":"figure","size":4984040,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eDonor fiber (GFP\u003c/strong\u003e\u003csup\u003e\u003cstrong\u003e+\u003c/strong\u003e\u003c/sup\u003e\u003cstrong\u003e) integration and morphology following juvenile and adult muscle transplantation.\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eSeven weeks after volumetric muscle loss (VML) injury and transplantation with GFP⁺ donor muscle, cross-sectional imaging of the TA muscle revealed localized integration of donor-derived tissue within the transplant region. Whole-muscle sections \u003cstrong\u003e(A)\u003c/strong\u003e show GFP⁺ signal (green) corresponding to donor cells, overlaid with wheat germ agglutinin (WGA; white) to visualize overall muscle structure. Both adult and juvenile transplant groups demonstrated similar spatial localization and extent of GFP⁺ signal. Quantification\u003cstrong\u003e (B)\u003c/strong\u003e showed GFP⁺ tissue occupied 10.78 ± 7.27% of total TA CSA in adult recipients (n=7) and 12.57 ± 7.39% in juvenile recipients (n = 8), with no significant difference between groups (p=0.6412). High-magnification views\u003cstrong\u003e (C)\u003c/strong\u003e highlight individual GFP⁺ myofibers surrounded by WGA-stained extracellular matrix. Analysis of GFP⁺ myofiber CSA\u003cstrong\u003e (D)\u003c/strong\u003e indicated mean donor fibers were significantly smaller than control host fibers (p \u0026lt; 0.001), averaging \u0026lt;1,000 µm² in both transplant groups, with no significant difference between adult and juvenile means (p\u0026gt; 0.05). Distribution analysis\u003cstrong\u003e (E)\u003c/strong\u003e revealed \u0026gt;70% of GFP⁺ fibers were \u0026lt;1,000 µm², consistent with regenerating or immature myofibers. A modest leftward shift was detected in the juvenile group, reflecting a higher proportion of very small donor fibers. Data are presented as mean ± SD. Scale bars = 1 mm (A), 250 μm (C).\u003c/p\u003e","description":"","filename":"Figure5.DonorfiberGFPintegrationandmorphologyfollowingjuvenileandadultmuscletransplantation.jpg","url":"https://assets-eu.researchsquare.com/files/rs-7437055/v1/d3b8fd0780fcd37c2301b5c8.jpg"},{"id":92845277,"identity":"9e45a1cc-4109-4084-8948-631fa7de9439","added_by":"auto","created_at":"2025-10-06 09:26:19","extension":"jpg","order_by":6,"title":"Figure 6","display":"","copyAsset":false,"role":"figure","size":8087618,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eSatellite cell density across regions of VML-injured muscle following transplantation.\u003c/strong\u003e\u003cbr\u003e\n \u003cstrong\u003e(A)\u003c/strong\u003e Representative whole-muscle cross-sections from uninjured controls, VML No Treatment, VML + Adult Transplant, and VML + Juvenile Transplant groups. Colored boxes indicate regions of interest sampled for satellite cell analysis and centralized nuclei, including uninjured tissue (white), the VML defect zone (blue), the border region surrounding the defect (yellow), distal muscle remote from injury (red), and GFP-positive transplant regions (purple). High-magnification images in \u003cstrong\u003e(B)\u003c/strong\u003e display Pax7⁺ satellite cells (Pink) localized to the periphery of laminin-stained myofibers (white), with DAPI (blue) used for nuclear counterstaining. GFP (green) marks donor tissue in transplanted groups. Quantification of satellite cell \u003cstrong\u003e(C)\u003c/strong\u003e density (Pax7⁺ cells/mm²) revealed ~8 cells/mm² in uninjured muscle, declining modestly in defect regions (~4 cells/mm²) but increasing ~3-fold at the border (~27 cells/mm², p \u0026lt; 0.0009 vs control). Distal regions were unchanged (~9–13 cells/mm², p \u0026gt; 0.09). Satellite cell density was significantly enriched within GFP⁺ transplant regions (27–34 cells/mm² in juvenile (p=0.0198) and adult (0.0043) groups), but no differences were observed between donor ages (p=0.6789). \u003cstrong\u003e(D)\u003c/strong\u003e Centralized nuclei (% of fibers) were negligible in control muscle (\u0026lt;1%) but elevated in all injured groups. In No Treatment animals, centralized nuclei reached ~28% in border regions and ~6% distally. Adult transplants contained 21% centralized nuclei at the border, 15% in GFP⁺ regions, and 4% distally. Juvenile transplants displayed higher regenerative activity, with 29% centralized nuclei in the border (~40% higher than adult, p=0.0332) and 23% in GFP⁺ regions (~55% higher than adult, p=0.0338), while distal muscle remained low (6–7%). Centralized nuclei were not quantified in defect regions due to the lack of fibers. Data are presented as mean ± SD. Statistical significance: *p\u0026lt;0.05 compared to controls and #p\u0026lt;0.05 compared to VML + Adult Transplant. Scale bars = 1mm (A), 100µm (B).\u003c/p\u003e","description":"","filename":"Figure6.SatellitecelldensityacrossregionsofVMLinjuredmusclefollowingtransplantation.jpg","url":"https://assets-eu.researchsquare.com/files/rs-7437055/v1/d7e8d39ca0d262685e106ca4.jpg"},{"id":97724100,"identity":"7b0a0412-3f14-44c6-96bd-ced2900bda88","added_by":"auto","created_at":"2025-12-08 16:11:55","extension":"pdf","order_by":0,"title":"","display":"","copyAsset":false,"role":"manuscript-pdf","size":24387269,"visible":true,"origin":"","legend":"","description":"","filename":"manuscript.pdf","url":"https://assets-eu.researchsquare.com/files/rs-7437055/v1/3cbd549e-0e6e-443e-abef-4102052172a0.pdf"}],"financialInterests":"No competing interests reported.","formattedTitle":"Juvenile vs Adult Skeletal Muscle Transplants in the Treatment of Volumetric Muscle Loss Injury","fulltext":[{"header":"INTRODUCTION","content":"\u003cp\u003eVolumetric muscle loss (VML) is a debilitating injury caused by trauma, tumor excision, or surgery that removes substantial amounts of skeletal muscle along with its cellular, vascular, neural, and extracellular matrix networks (\u003cspan citationid=\"CR1\" class=\"CitationRef\"\u003e1\u003c/span\u003e, \u003cspan citationid=\"CR2\" class=\"CitationRef\"\u003e2\u003c/span\u003e). This destructive injury eliminates structural support and depletes satellite cells, the resident muscle stem cells essential for regeneration (\u003cspan citationid=\"CR3\" class=\"CitationRef\"\u003e3\u003c/span\u003e, \u003cspan citationid=\"CR4\" class=\"CitationRef\"\u003e4\u003c/span\u003e). Because the satellite cell niche (basal lamina) is disrupted, the cells are unable to migrate into areas of damage and facilitate regeneration. Conversely, fibrotic scar tissue fills the void, overwhelming the intrinsic repair mechanisms, and results in chronic deficits in muscle strength and function (\u003cspan citationid=\"CR5\" class=\"CitationRef\"\u003e5\u003c/span\u003e, \u003cspan citationid=\"CR6\" class=\"CitationRef\"\u003e6\u003c/span\u003e).\u003c/p\u003e\u003cp\u003eClinically, there is no standard of care to treat VML, thus management is often focused on wound closure and limb salvage rather than restoration of muscle mass and function (\u003cspan citationid=\"CR7\" class=\"CitationRef\"\u003e7\u003c/span\u003e). Experimental regenerative approaches, including skeletal muscle transplantation and autologous minced muscle grafts, aim to fill the defect with donor tissue that provides both structural scaffolding and a reservoir of regenerative cells (\u003cspan citationid=\"CR7\" class=\"CitationRef\"\u003e7\u003c/span\u003e, \u003cspan citationid=\"CR8\" class=\"CitationRef\"\u003e8\u003c/span\u003e). Upon transplantation, donor satellite cells can activate, proliferate, differentiate, and fuse with host fibers to support local regeneration (\u003cspan citationid=\"CR4\" class=\"CitationRef\"\u003e4\u003c/span\u003e). However, even with successful donor integration, functional recovery is typically incomplete (\u003cspan citationid=\"CR5\" class=\"CitationRef\"\u003e5\u003c/span\u003e, \u003cspan citationid=\"CR9\" class=\"CitationRef\"\u003e9\u003c/span\u003e), highlighting the potential need to optimize donor tissue characteristics.\u003c/p\u003e\u003cp\u003eOne underexplored factor influencing transplant efficacy is the developmental age of donor muscle, as skeletal muscle undergoes pronounced structural and cellular changes across the lifespan (\u003cspan additionalcitationids=\"CR11\" citationid=\"CR10\" class=\"CitationRef\"\u003e10\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR12\" class=\"CitationRef\"\u003e12\u003c/span\u003e). Juvenile muscle exhibits small-diameter myofibers, high nuclear and satellite cell densities, and a microenvironment supportive of fiber growth and remodeling (\u003cspan citationid=\"CR3\" class=\"CitationRef\"\u003e3\u003c/span\u003e, \u003cspan citationid=\"CR13\" class=\"CitationRef\"\u003e13\u003c/span\u003e, \u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e). In contrast, adult muscle contains fewer satellite cells and larger, less plastic fibers (\u003cspan citationid=\"CR3\" class=\"CitationRef\"\u003e3\u003c/span\u003e, \u003cspan citationid=\"CR10\" class=\"CitationRef\"\u003e10\u003c/span\u003e). Such age-related differences may have critical implications for regenerative potential following transplantation.\u003c/p\u003e\u003cp\u003eEvidence from other regenerative contexts supports this concept: juvenile tissue or juvenile-like progenitors often outperform adult counterparts in repair, likely due to a more abundant and responsive stem cell pool combined with a growth-permissive environment (\u003cspan citationid=\"CR3\" class=\"CitationRef\"\u003e3\u003c/span\u003e, \u003cspan citationid=\"CR10\" class=\"CitationRef\"\u003e10\u003c/span\u003e, \u003cspan citationid=\"CR15\" class=\"CitationRef\"\u003e15\u003c/span\u003e). Our group and others have shown that regenerative outcomes are constrained by the injury environment, including persistent denervation (\u003cspan citationid=\"CR16\" class=\"CitationRef\"\u003e16\u003c/span\u003e, \u003cspan citationid=\"CR17\" class=\"CitationRef\"\u003e17\u003c/span\u003e), extracellular matrix remodeling (\u003cspan citationid=\"CR2\" class=\"CitationRef\"\u003e2\u003c/span\u003e), and limited stem cell migration into the defect (\u003cspan citationid=\"CR5\" class=\"CitationRef\"\u003e5\u003c/span\u003e). These constraints suggest that the intrinsic advantages of juvenile donor tissue may still require complementary interventions, such as rehabilitation, to achieve full functional recovery (\u003cspan additionalcitationids=\"CR19\" citationid=\"CR18\" class=\"CitationRef\"\u003e18\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR20\" class=\"CitationRef\"\u003e20\u003c/span\u003e).\u003c/p\u003e\u003cp\u003eHere, we tested the hypothesis that juvenile skeletal muscle, by virtue of its cellular composition and myogenic capacity, would outperform adult muscle in restoring structure and function following VML injury. We first characterized the morphology, satellite cell content, and in-vitro myogenic behavior of tibialis anterior (TA) muscle from juvenile, adolescent, and adult rats. We then compared the in-vivo regenerative outcomes of juvenile versus adult TA muscle transplants in a standardized rat VML model.\u003c/p\u003e"},{"header":"METHODS","content":"\u003cp\u003eThis study was conducted in two parts. Study 1 evaluated the structural and cellular properties of juvenile, adolescent, and adult skeletal muscle to determine age-dependent differences relevant to regeneration. Study 2 examined the in-vivo regenerative performance of juvenile and adult muscle transplants in a rat model of VML, using functional and histological outcome measures seven weeks post-VML surgery. The work herein has been reported in line with the ARRIVE guidelines 2.0.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eEthical Approval\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eAll experiments were approved by the Institutional Animal Care and Use Committee (IACUC) at Brigham Young University (Protocol #23-0401) in accordance with National Institutes of Health guidelines for the care and use of laboratory animals. All animals were housed on a 12-12 h light-dark cycle, with food and water ad libitum.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eStudy 1: Developmental Analysis of Donor Muscle\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eAnimals and Tissue Collection\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eTo evaluate regenerative properties of skeletal muscle by age, male Lewis rats were grouped as juvenile (21 \u0026plusmn; 0 days), adolescent (34 \u0026plusmn; 0 days), or adult (130 \u0026plusmn; 4.6 days) (n = 4 per group, 12 total). The left tibialis anterior (TA) muscle was harvested for histological analysis, and the right TA was used for myoblast isolation. Sample size was based on prior reports that show large age-related differences in rodent skeletal muscle. All animals were euthanized following tissue collection while under heavy isoflurane anesthesia (5%) until loss of reflexes, then bilateral thoracotomy was used as a secondary method to ensure death. \u0026nbsp; \u003cstrong\u003eImmunohistochemistry and Imaging\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eTo assess age-related satellite cell content, myofiber number, and muscle morphology, TA muscles were collected, weighed, then embedded in tragacanth gum, snap frozen in isopentane, and cryosectioned (10 \u0026micro;m). Sections were fixed in 4% paraformaldehyde (Thermo Scientific), blocked with 10% normal goat serum (NGS; Thermo Fisher), and incubated overnight with Pax7 (mouse IgG1, 3 \u0026micro;g/mL; DSHB, AB_528428) and laminin (mouse IgG2a, 2 \u0026micro;g/mL; DSHB, AB_2618140). Secondary antibodies included Cy3 goat anti-mouse IgG1 (1:200; Jackson, 115-165-003) and Alexa Fluor 647 goat anti-mouse IgG2a (1:200; Jackson, 115-605-206). DAPI (1 \u0026micro;g/mL; Thermo Fisher, D1306) was used for nuclear counterstaining.\u003c/p\u003e\n\u003cp\u003eWhole tissue cross-sections were used to count myofibers and to measure whole TA and myofiber cross-sectional area. Images were acquired on a tile-scanning Echo Revolution fluorescence microscope using a 20\u0026times; PlanX Apo objective. The laminin images were converted to grayscale and a threshold set (10\u0026ndash;30 range) using Fiji (ImageJ) software. Myofibers were segmented using \u0026ldquo;Analyze Particles\u0026rdquo; with a size range of 50\u0026ndash;8000 \u0026micro;m\u0026sup2; and circularity 0.30\u0026ndash;1.00. Manual correction was performed for incomplete borders. Mean fiber CSA and total fiber number were calculated per sample.\u0026nbsp;\u003c/p\u003e\n\u003cp\u003eTo assess satellite cell and nuclei content, \u0026nbsp;five 1 mm\u0026sup2; regions of interest were randomly imaged throughout each sample. Nuclei were counted based on images from the DAPI channel using the Otsu threshold parameters and the \u0026ldquo;watershed\u0026rdquo; feature to separate nuclei clusters. The \u0026ldquo;Analyze Particles\u0026rdquo; feature was set to a size threshold of 20-50\u0026micro;m. Satellite cells were identified and counted manually based on Pax7⁺/DAPI⁺ co-localization within or near the laminin border. \u003cstrong\u003eMyoblast Isolation and In Vitro Assays\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eTo characterize age-dependent differences in satellite cell behavior and regenerative potential, primary myoblasts (activated satellite cells) were isolated from the right TA muscles of juvenile, adolescent, and adult rats. These cells were used in a series of in vitro assays to assess proliferative capacity, differentiation potential, and myogenic identity.\u003c/p\u003e\n\u003cp\u003eRight TA muscles were digested using the Miltenyi Skeletal Muscle Dissociation Kit and gentleMACS system. Cell suspensions were filtered (70\u0026micro;m) and centrifuged, then incubated with PE-conjugated anti-CD106 (Miltenyi, 130-103-684) and sorted with Anti-PE MicroBeads (130-048-801) using LS columns. CD106⁺ myoblasts were seeded on Matrigel-coated plates and allowed to expand in DMEM (Gibco) with 10% FBS and 1% penicillin-streptomycin changed every 48 hours.\u003c/p\u003e\n\u003cp\u003eMyogenic identity was confirmed via MyoD1 staining (mouse IgG2b, 2 \u0026micro;g/mL; DSHB, AB_2146602) with Cy3 anti-IgG2b secondary (1:200; Jackson, 115-165-207). DAPI was used for nuclei staining. The percentage of MyoD⁺ nuclei was assessed in three random fields per group.\u003c/p\u003e\n\u003cp\u003eMyoblasts were seeded at a density of 25,000 cells/well on a Matrigel coated, 12-well plate and allowed to incubate overnight. Proliferation was assessed by EdU incorporation (Click-iT\u0026trade; kit, Thermo Fisher, C10340) following manufacturer instructions at 4, 8, and 16 hours. Briefly, cells were fixed (4% PFA), permeabilized (0.5% TritonX-100 in PBS), and labeled with Alexa Fluor 647-conjugated azide and Hoechst. Hoechst-stained nuclei were counted, and the proliferation index was calculated as EdU⁺ cells/total nuclei. Data were averaged from four fields per well, in triplicate wells per group.\u003c/p\u003e\n\u003cp\u003eFor differentiation, cells were seeded at 100,000 cells per well on a Matrigel coated, 12-well plate and allowed to incubate overnight in growth media. Differentiation was induced by switching to DMEM + 2% horse serum for 72 hours. Cells were fixed and stained for Myogenin (mouse IgG1, 2 \u0026micro;g/mL; DSHB, AB_2146601) and MyHC (mouse IgG2b, 2 \u0026micro;g/mL; DSHB, AB_2147781). Secondary antibodies included Alexa Fluor 647 anti-IgG1 and Cy3 anti-IgG2b (1:200; Jackson). DAPI was used for nuclear staining.\u0026nbsp;\u003c/p\u003e\n\u003cp\u003eThree outcome measures were used to assess differentiation: fusion index, MyHC\u003csup\u003e+\u003c/sup\u003e area, and nucleation index. The fusion index was defined as the percentage of total nuclei located within MyHC\u003csup\u003e+\u003c/sup\u003e multinucleated myotubes, indicating the efficiency of myoblast fusion during early differentiation. Myotubes were defined as MyHC\u003csup\u003e+\u003c/sup\u003e structures containing two or more DAPI-stained nuclei. The MyHC\u003csup\u003e+\u003c/sup\u003e area was calculated as the total surface area occupied by MyHC\u003csup\u003e+\u003c/sup\u003e staining within each field of view, measured in ImageJ using a threshold binary masks. The nucleation index was calculated as the average number of nuclei contained within each individual MyHC\u003csup\u003e+\u003c/sup\u003e myotube. This index served as an indicator of myotube maturation. All quantifications were performed on at least four randomly selected, non-overlapping fields per well, and averaged across triplicate wells per condition.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eStudy 2: VML Injury and Muscle Transplantation\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eVML Surgery and Experimental Design\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eTo evaluate the regenerative performance of juvenile versus adult muscle transplants in a preclinical model of VML, a standardized full thickness VML injury and repair protocol was conducted in adult male rats. This approach was designed to mimic clinically relevant muscle trauma and assess functional and histological outcomes following transplantation of developmentally distinct donor tissue.\u003c/p\u003e\n\u003cp\u003eTwenty-four adult male Lewis rats (3\u0026ndash;4 months old) were randomly assigned to one of three treatment groups: VML No Treatment, VML + Adult Transplant, or VML + Juvenile Transplant (n = 8 per group). Sample size was based on prior muscle transplant studies in a rat VML model to detect functional improvements of 20\u0026ndash;40% (17). All animals underwent unilateral VML surgery in the left TA muscle with the right leg serving as an uninjured, intra-animal control. To provide analgesia, a carprofen tablet (Bio-Serv, 5 g) was placed in the animal\u0026rsquo;s cage 24 hours prior to surgery and a single preoperative dose of sustained-release buprenorphine (1.2 mg/kg, SC; Wedgewood) was injected subcutaneously into the back of the neck at least one hour before surgery.\u0026nbsp;\u003c/p\u003e\n\u003cp\u003eAnimals were anesthetized with 2\u0026ndash;3% isoflurane in oxygen and placed in a supine position on a heated surgical platform. The lower left hindlimb was shaved and disinfected with alternating scrubs of chlorhexidine and 70% ethanol. A longitudinal skin incision (~1.5 cm) was made along the anterior surface of the lower leg to expose the underlying musculature. The skin and fascia were opened to expose the TA muscle, which was gently isolated from surrounding tissue using blunt dissection. To protect adjacent musculature, a sterile surgical spatula was inserted beneath the TA and full-thickness defect, approximately 6 mm in diameter (~15\u0026ndash;20% of muscle volume), was created in the mid-belly region using a sterile biopsy punch (MedBlades, USA).\u0026nbsp;\u003c/p\u003e\n\u003cp\u003eDonor tissue was harvested immediately prior to transplantation from ubiquitously expressing GFP⁺ juvenile (21-day; n=4) or adult (~120-day; n=2) male Lewis rats (Rat Resource and Research Center; Strain: \u003cstrong\u003eLEW-Tg(CAG-EGFP)YsRrrc; RRRC#: 00206)\u003c/strong\u003e. TA muscles were dissected, placed in a sterile tissue culture dish on ice, and finely minced into ~1 mm\u0026sup3; fragments using sterile scissors. The total weight of minced tissue was adjusted to approximate the volume of the VML defect and then carefully implanted into the defect site.\u003c/p\u003e\n\u003cp\u003eFor animals receiving transplants, the fascia was sutured, followed by subcutaneous closure of the skin using interrupted sutures. VML No Treatment animals underwent the same surgical procedure without tissue implantation.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eIn Vivo Muscle Strength Testing\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eTo evaluate muscle function following VML injury and transplantation, in-vivo isometric force testing was performed seven weeks post-surgery for all of the VML injured limbs (n=24) and most (n=19) of the uninjured control limbs. Under 2\u0026ndash;3% isoflurane anesthesia, rats were placed supine on a temperature-controlled platform. The hindlimb was immobilized using a knee clamp, and the foot was secured to a force transducer footplate (Aurora Scientific 3-in-1 Muscle Test System).\u003c/p\u003e\n\u003cp\u003eSubcutaneous needle electrodes were inserted near the peroneal nerve to stimulate the anterior compartment (TA/EDL) of the hindlimb. Stimulation parameters included 0.1 ms pulse width, 400 ms train duration, and increasing frequencies ranging from 10 to 200 Hz. Optimal voltage was determined for each animal to ensure maximal contractile response. Peak isometric tetanic force was recorded as the highest value generated during the frequency ramp. To account for inter-animal variability in body size, absolute force values were normalized to body weight (mN\u0026middot;m/kg).\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eTissue Collection and Histology\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eSeven weeks post VML surgery, TA, EDL, soleus, and gastrocnemius muscles were dissected and weighed. Animals were then euthanized with an overdose of isoflurane (5%) and bilateral thoracotomy as a secondary measure to ensure death. TA sections were frozen in isopentane cooled in liquid nitrogen and stored at -80\u0026deg;C. The TA \u0026nbsp;muscles were sectioned (10 \u0026micro;m) and stained for laminin, Pax7, and DAPI. High-resolution images were taken randomly from controls and from four regions in VML injured samples: defect (typically devoid of myofibers), transplant (identified by GFP expression), border (adjacent to the injury; smaller, disorganized host fibers), and distal (intact muscle away from the injury site). Satellite cells were counted based on Pax7⁺/DAPI⁺ co-localization. Myofiber CSA, total fiber count, and centrally nucleated fibers were quantified from regions of interest using ImageJ analysis software and the previously described segmentation thresholds.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eDonor Fiber Analysis\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eTo assess transplant integration and morphology, TA muscle sections were analyzed for GFP fluorescence and labeled with wheat germ agglutinin (WGA) to identify myofiber boundaries. Sections were cut at 10\u0026micro;m thickness and immediately fixed in 2% paraformaldehyde for 10 minutes at room temperature to preserve endogenous GFP signal. Following fixation, slides were rinsed once with PBS and incubated for 15 minutes with Alexa Fluor 647-conjugated wheat germ agglutinin (WGA; 1:500 dilution in PBS; Thermo Fisher Scientific, Cat# W32466). Sections were washed once in PBS and mounted in Fluoroshield (Sigma) and imaged using a 20X PlanX Apo objective on the Echo Revolution fluorescence microscope. GFP fluorescence was visualized directly without antibody amplification. Exposure settings were optimized to avoid saturation and kept constant between samples within each group.\u003c/p\u003e\n\u003cp\u003eTo quantify GFP⁺ donor fibers, high-resolution images were collected from well-defined GFP regions within the transplant zone. Fields with high signal-to-noise ratio and clear membrane borders were prioritized for analysis. Myofiber CSA was measured using ImageJ. GFP-positive fibers were defined as those showing continuous cytoplasmic GFP signal enclosed by WGA-labeled borders. A minimum of 200 GFP-positive fibers were analyzed per animal. The GFP area was also calculated as a percentage of total CSA using full-section scans in ImageJ, thresholded for GFP fluorescence.\u0026nbsp;\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eStatistical Analysis\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eAll statistical analyses were performed using GraphPad Prism version 10.5.0. One-way ANOVA with Tukey\u0026rsquo;s post hoc test was used for group comparisons. Data are presented as mean \u0026plusmn; SD. A p-value \u0026lt; 0.05 was considered statistically significant.\u003c/p\u003e"},{"header":"RESULTS","content":"\u003cp\u003e\u003cstrong\u003eStudy 1: Comparison Between Juvenile, Adolescent, and Adult Male Lewis Rats\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eRat Characteristics\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eJuvenile (21-day), adolescent (34-day), and adult (120-day) male Lewis rats were used to represent distinct developmental stages. As expected, body and TA muscle weights increased significantly with age (Table 1), consistent with known patterns of postnatal growth.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eTable 1: Rat Characteristics\u003c/strong\u003e\u003c/p\u003e\n\u003ctable border=\"1\" cellspacing=\"0\" cellpadding=\"0\" width=\"462\"\u003e\n \u003ctbody\u003e\n \u003ctr\u003e\n \u003ctd style=\"width: 90px;\"\u003e\n \u003cp\u003e\u003cstrong\u003eGroup\u003c/strong\u003e\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 84px;\"\u003e\n \u003cp\u003e\u003cstrong\u003eAge (days)\u003c/strong\u003e\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 138px;\"\u003e\n \u003cp\u003e\u003cstrong\u003eBody Weight (grams)\u003c/strong\u003e\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 150px;\"\u003e\n \u003cp\u003e\u003cstrong\u003eTA Wet Weight (grams)\u003c/strong\u003e\u003c/p\u003e\n \u003c/td\u003e\n \u003c/tr\u003e\n \u003ctr\u003e\n \u003ctd style=\"width: 90px;\"\u003e\n \u003cp\u003e\u003cstrong\u003eJuvenile\u003c/strong\u003e\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 84px;\"\u003e\n \u003cp\u003e21 \u0026plusmn; 0\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 138px;\"\u003e\n \u003cp\u003e53.85 \u0026plusmn; 0.37\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 150px;\"\u003e\n \u003cp\u003e0.16 \u0026plusmn; 0.0036\u003c/p\u003e\n \u003c/td\u003e\n \u003c/tr\u003e\n \u003ctr\u003e\n \u003ctd style=\"width: 90px;\"\u003e\n \u003cp\u003e\u003cstrong\u003eAdolescent\u003c/strong\u003e\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 84px;\"\u003e\n \u003cp\u003e34 \u0026plusmn; 0\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 138px;\"\u003e\n \u003cp\u003e144.5 \u0026plusmn; 6.25\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 150px;\"\u003e\n \u003cp\u003e0.25 \u0026plusmn; 0.0183\u003c/p\u003e\n \u003c/td\u003e\n \u003c/tr\u003e\n \u003ctr\u003e\n \u003ctd style=\"width: 90px;\"\u003e\n \u003cp\u003e\u003cstrong\u003eAdult\u003c/strong\u003e\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 84px;\"\u003e\n \u003cp\u003e120 \u0026plusmn; 5.7\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 138px;\"\u003e\n \u003cp\u003e396.5 \u0026plusmn; 16.6\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 150px;\"\u003e\n \u003cp\u003e0.775 \u0026plusmn; 0.031\u003c/p\u003e\n \u003c/td\u003e\n \u003c/tr\u003e\n \u003c/tbody\u003e\n\u003c/table\u003e\n\u003cp\u003e\u003cstrong\u003e\u0026nbsp;\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eMuscle Size and Myofiber Architecture Vary by Developmental Stage\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eWhole-muscle CSA increased significantly with age (Figure 1A\u0026ndash;B). Juvenile TAs averaged 3.95 \u0026plusmn; 0.29 mm\u0026sup2;, adolescent TAs 9.64 \u0026plusmn; 1.80 mm\u0026sup2;, and adult TAs 27.07 \u0026plusmn; 5.36 mm\u0026sup2;. All pairwise comparisons were statistically significant (juvenile vs. adolescent \u003cem\u003ep\u003c/em\u003e = 0.0012; juvenile vs. adult \u003cem\u003ep\u003c/em\u003e \u0026lt; 0.0001; adolescent vs. adult \u003cem\u003ep\u003c/em\u003e = 0.0003), confirming stepwise muscle growth. Despite these CSA increases, total myofiber number did not significantly differ between groups (juvenile: 8593 \u0026plusmn; 525; adolescent: 10,101 \u0026plusmn; 1484; adult: 10,126 \u0026plusmn; 1282; \u003cem\u003ep\u003c/em\u003e = 0.1621), indicating that muscle growth was driven primarily by fiber hypertrophy, not hyperplasia (Figure 1C).\u003c/p\u003e\n\u003cp\u003eAnalysis of individual myofiber CSA revealed age-related increases (Figure 1D). Juvenile fibers averaged 460 \u0026plusmn; 38 \u0026micro;m\u0026sup2;, significantly smaller than adolescent (963 \u0026plusmn; 147 \u0026micro;m\u0026sup2;; \u003cem\u003ep\u003c/em\u003e = 0.0267) and adult fibers (2798 \u0026plusmn; 354 \u0026micro;m\u0026sup2;; \u003cem\u003ep\u003c/em\u003e \u0026lt; 0.0001). Adolescent fibers were also significantly smaller than adult fibers (\u003cem\u003ep\u003c/em\u003e \u0026lt; 0.0001). The CSA frequency distributions further emphasized this shift as juvenile muscle showed a narrow peak at small fiber sizes, whereas adolescent and adult profiles were broader and right-shifted, consistent with increased fiber size (Figure 1E).\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eNuclei, Myofiber, and Satellite Cell Density Decline with Advancing Age\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eTo assess how developmental age affects muscle, we quantified nuclei, myofiber density, and Pax7⁺ satellite cells per mm\u0026sup2; (Figure 2A\u0026ndash;B). Nuclear density was highest in juvenile muscle (2907.5 \u0026plusmn; 99.4 nuclei/mm\u0026sup2;), declined to 1550.4 \u0026plusmn; 268.1 in adolescents, and dropped further in adults (704.3 \u0026plusmn; 139.6; \u003cem\u003ep\u003c/em\u003e \u0026lt; 0.0001 for all comparisons) (Figure 2C). Myofiber density also declined significantly with age (\u003cem\u003ep\u003c/em\u003e \u0026lt; 0.001), consistent with increasing fiber size (Figure 2D).\u003c/p\u003e\n\u003cp\u003eSatellite cell density, measured by Pax7⁺ nuclei per mm\u0026sup2;, showed a dramatic age-related decline (Figure 2E). Juvenile muscles contained 122.8 \u0026plusmn; 28.4 Pax7⁺ cells/mm\u0026sup2;, approximately 3.5 times higher than adolescents (36.1 \u0026plusmn; 7.1; \u003cem\u003ep\u003c/em\u003e \u0026lt; 0.001) and 15 times higher than adults (8.4 \u0026plusmn; 3.3; \u003cem\u003ep\u003c/em\u003e \u0026lt; 0.0001). Although adolescent satellite cell density appeared higher than adult, the difference did not reach statistical significance (\u003cem\u003ep\u003c/em\u003e = 0.1049). Together, these results confirm that juvenile skeletal muscle possesses a dense, more cellular microenvironment with elevated satellite cell content, nuclei and myofibers, which may enhance regenerative potential when used as a source of donor tissue to treat VML.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eMyoblast Proliferation and Differentiation\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eTo assess how donor age affects cellular behavior, myoblasts were isolated from juvenile, adolescent, and adult TA muscles and evaluated in-vitro.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eMyogenic Purity and Proliferation\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eImmunostaining confirmed high myogenic purity across all groups, with MyoD⁺ nuclei accounting for 95.3 \u0026plusmn; 2.7% in juvenile, 95.4 \u0026plusmn; 0.2% in adolescent, and 95.2 \u0026plusmn; 1.4% in adult cultures (Figure 3A\u0026ndash;B). EdU incorporations over 4, 8, and 16 hours showed a time-dependent increase in proliferation, rising from ~15% at 4 hours to ~80% at 16 hours across all groups. However, no significant differences in proliferation rate were observed between age groups at any time point (\u003cem\u003ep\u003c/em\u003e = 0.9696), indicating that donor age did not affect baseline proliferative capacity (Figure 3C\u0026ndash;D).\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eEnhanced Differentiation in Juvenile Myoblasts\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eIn contrast, differentiation declined significantly with age. After 72 hours in differentiation medium, juvenile myoblasts showed a higher fusion index (17.49 \u0026plusmn; 2.53%) than adolescent (13.86 \u0026plusmn; 0.83%) and adult (10.12 \u0026plusmn; 1.79%) cultures. The difference between juvenile and adult groups was statistically significant (\u003cem\u003ep\u003c/em\u003e = 0.0067), while differences between juvenile and adolescent (\u003cem\u003ep\u003c/em\u003e = 0.1161) and adolescent and adult (\u003cem\u003ep\u003c/em\u003e = 0.1066) were not (Figure 3E).\u003c/p\u003e\n\u003cp\u003eMyosin Heavy Chain (MyHC) area per field, a measure of total myotube formation, was significantly larger in juvenile cultures (64,667 \u0026plusmn; 10,161 \u0026micro;m\u0026sup2;) than in adolescent (40,652 \u0026plusmn; 2,331 \u0026micro;m\u0026sup2;; \u003cem\u003ep\u003c/em\u003e = 0.0191) and adult cultures (35,149 \u0026plusmn; 7,968 \u0026micro;m\u0026sup2;; \u003cem\u003ep\u003c/em\u003e = 0.0074) (Figure 3F). No significant difference was observed between adolescent and adult groups (\u003cem\u003ep\u003c/em\u003e = 0.6658).\u003c/p\u003e\n\u003cp\u003eThe nucleation index, defined as the average number of nuclei per myotube, did not differ significantly between groups. Juvenile (13.27 \u0026plusmn; 2.62), adolescent (9.89 \u0026plusmn; 2.79), and adult (7.88 \u0026plusmn; 1.07) cultures exhibited similar nuclear content per fiber, with no statistically significant differences (Figure 3G). Overall, these findings indicate that juvenile myoblasts have an enhanced capacity for differentiation and myotube formation, despite equivalent proliferation.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eStudy 2: Comparison Between VML Treatment Groups\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eRat Characteristics at Time of VML Surgery\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eTo evaluate transplant performance, rats were randomized to VML No Treatment, VML + Adult Transplant, or VML + Juvenile Transplant groups (n = 8/group). All animals were of similar age and body weight at the time of surgery, with no significant differences between groups (Table 2). However, the VML tissue excised from the No Treatment group was significantly smaller than in the transplant groups, which may have contributed to milder deficits in untreated animals. Transplant weights did not differ significantly between juvenile and adult donor groups.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eTable 2: Rat Characteristics at time of surgery\u003c/strong\u003e\u003c/p\u003e\n\u003ctable border=\"1\" cellspacing=\"0\" cellpadding=\"0\" width=\"630\"\u003e\n \u003ctbody\u003e\n \u003ctr\u003e\n \u003ctd style=\"width: 90px;\"\u003e\n \u003cp\u003e\u003cstrong\u003eGroup\u003c/strong\u003e\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 120px;\"\u003e\n \u003cp\u003e\u003cstrong\u003eAge at Surgery (Days)\u003c/strong\u003e\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 120px;\"\u003e\n \u003cp\u003e\u003cstrong\u003eWeight At Surgery (g)\u003c/strong\u003e\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 144px;\"\u003e\n \u003cp\u003e\u003cstrong\u003eWeight of VML Piece (mg)\u003c/strong\u003e\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 156px;\"\u003e\n \u003cp\u003e\u003cstrong\u003eVML Transplant Weight (mg)\u003c/strong\u003e\u003c/p\u003e\n \u003c/td\u003e\n \u003c/tr\u003e\n \u003ctr\u003e\n \u003ctd valign=\"bottom\" style=\"width: 90px;\"\u003e\n \u003cp\u003e\u003cstrong\u003eVML No Treatment\u003c/strong\u003e\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 120px;\"\u003e\n \u003cp\u003e110.63 \u0026plusmn; 3.62\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 120px;\"\u003e\n \u003cp\u003e385.13 \u0026plusmn; 32.52\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 144px;\"\u003e\n \u003cp\u003e\u003cstrong\u003e73.97 \u0026plusmn; 12.89\u003c/strong\u003e\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 156px;\"\u003e\n \u003cp\u003eN/A\u003c/p\u003e\n \u003c/td\u003e\n \u003c/tr\u003e\n \u003ctr\u003e\n \u003ctd valign=\"bottom\" style=\"width: 90px;\"\u003e\n \u003cp\u003e\u003cstrong\u003eVML + Adult Transplant\u003c/strong\u003e\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 120px;\"\u003e\n \u003cp\u003e113.00 \u0026plusmn; 11.78\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 120px;\"\u003e\n \u003cp\u003e418.88 \u0026plusmn; 42.29\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 144px;\"\u003e\n \u003cp\u003e85.79 \u0026plusmn; 18.03\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 156px;\"\u003e\n \u003cp\u003e93.94 \u0026plusmn; 20.79\u003c/p\u003e\n \u003c/td\u003e\n \u003c/tr\u003e\n \u003ctr\u003e\n \u003ctd valign=\"bottom\" style=\"width: 90px;\"\u003e\n \u003cp\u003e\u003cstrong\u003eVML + Juvenile Transplant\u003c/strong\u003e\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 120px;\"\u003e\n \u003cp\u003e107.30 \u0026plusmn; 14.95\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 120px;\"\u003e\n \u003cp\u003e404.20 \u0026plusmn; 19.23\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 144px;\"\u003e\n \u003cp\u003e83.40 \u0026plusmn; 14.17\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 156px;\"\u003e\n \u003cp\u003e97.60 \u0026plusmn; 11.07\u003c/p\u003e\n \u003c/td\u003e\n \u003c/tr\u003e\n \u003c/tbody\u003e\n\u003c/table\u003e\n\u003cp\u003e\u003cstrong\u003e\u003cem\u003eNote: Bold numbers indicate significance\u003c/em\u003e\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eRat Characteristics Post VML Surgery\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eAt tissue collection (seven weeks post-surgery), body weight was lower in the No Treatment group (415.3 \u0026plusmn; 14.6 g) compared to the Adult (440.9 \u0026plusmn; 38.8 g) and Juvenile (443.6 \u0026plusmn; 25.0 g) transplant groups (\u003cem\u003ep\u003c/em\u003e \u0026lt; 0.05). Despite this, individual muscle wet weights, including the TA, EDL, soleus, and gastrocnemius, were not significantly different across groups (Table 3), suggesting that gross muscle mass was not markedly influenced by transplantation within the time window assessed.\u003c/p\u003e\n\u003ctable border=\"1\" cellspacing=\"0\" cellpadding=\"0\" width=\"624\"\u003e\n \u003ctbody\u003e\n \u003ctr\u003e\n \u003ctd colspan=\"2\" style=\"width: 192px;\"\u003e\n \u003cp\u003e\u003cstrong\u003eTable 3: Muscle Weights\u003c/strong\u003e\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd colspan=\"8\" style=\"width: 432px;\"\u003e\n \u003cp\u003e\u003cstrong\u003eWet Weight (g)\u003c/strong\u003e\u003c/p\u003e\n \u003c/td\u003e\n \u003c/tr\u003e\n \u003ctr\u003e\n \u003ctd style=\"width: 96px;\"\u003e\n \u003cp\u003e\u003cstrong\u003eGroup\u003c/strong\u003e\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 96px;\"\u003e\n \u003cp\u003e\u003cstrong\u003eBody Weight at Collection (g)\u003c/strong\u003e\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 48px;\"\u003e\n \u003cp\u003e\u003cstrong\u003eRight TA\u003c/strong\u003e\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 42px;\"\u003e\n \u003cp\u003e\u003cstrong\u003eLeft\u0026nbsp;\u003c/strong\u003e\u003c/p\u003e\n \u003cp\u003e\u003cstrong\u003eTA\u003c/strong\u003e\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 54px;\"\u003e\n \u003cp\u003e\u003cstrong\u003eRight EDL\u003c/strong\u003e\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 48px;\"\u003e\n \u003cp\u003e\u003cstrong\u003eLeft EDL\u003c/strong\u003e\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 54px;\"\u003e\n \u003cp\u003e\u003cstrong\u003eRight Soleus\u003c/strong\u003e\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 54px;\"\u003e\n \u003cp\u003e\u003cstrong\u003eLeft Soleus\u003c/strong\u003e\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 66px;\"\u003e\n \u003cp\u003e\u003cstrong\u003eRight Gastroc\u003c/strong\u003e\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 66px;\"\u003e\n \u003cp\u003e\u003cstrong\u003eLeft Gastroc\u003c/strong\u003e\u003c/p\u003e\n \u003c/td\u003e\n \u003c/tr\u003e\n \u003ctr\u003e\n \u003ctd style=\"width: 96px;\"\u003e\n \u003cp\u003e\u003cstrong\u003eVML No Treatment\u003c/strong\u003e\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 96px;\"\u003e\n \u003cp\u003e\u003cstrong\u003e415.29 \u0026plusmn; 14.57\u003c/strong\u003e\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 48px;\"\u003e\n \u003cp\u003e0.74\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e\u0026plusmn;\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e0.07\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 42px;\"\u003e\n \u003cp\u003e0.70\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e\u0026plusmn;\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e0.12\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 54px;\"\u003e\n \u003cp\u003e0.20\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e\u0026plusmn;\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e0.04\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 48px;\"\u003e\n \u003cp\u003e0.20\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e\u0026plusmn;\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e0.02\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 54px;\"\u003e\n \u003cp\u003e0.17\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e\u0026plusmn;\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e0.04\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 54px;\"\u003e\n \u003cp\u003e0.19\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e\u0026plusmn;\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e0.04\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 66px;\"\u003e\n \u003cp\u003e1.90\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e\u0026plusmn;\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e0.12\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 66px;\"\u003e\n \u003cp\u003e1.99\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e\u0026plusmn;\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e0.21\u003c/p\u003e\n \u003c/td\u003e\n \u003c/tr\u003e\n \u003ctr\u003e\n \u003ctd style=\"width: 96px;\"\u003e\n \u003cp\u003e\u003cstrong\u003eVML + Adult Transplant\u003c/strong\u003e\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 96px;\"\u003e\n \u003cp\u003e440.86 \u0026plusmn; 38.84\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 48px;\"\u003e\n \u003cp\u003e0.77\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e\u0026plusmn;\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e0.03\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 42px;\"\u003e\n \u003cp\u003e0.73\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e\u0026plusmn;\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e0.09\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 54px;\"\u003e\n \u003cp\u003e0.17\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e\u0026plusmn;\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e0.02\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 48px;\"\u003e\n \u003cp\u003e0.17\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e\u0026plusmn;\u003c/p\u003e\n \u003cp\u003e0.02\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 54px;\"\u003e\n \u003cp\u003e0.21\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e\u0026plusmn;\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e0.03\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 54px;\"\u003e\n \u003cp\u003e0.17\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e\u0026plusmn;\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e0.01\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 66px;\"\u003e\n \u003cp\u003e1.99\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e\u0026plusmn;\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e0.14\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 66px;\"\u003e\n \u003cp\u003e1.96\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e\u0026plusmn;\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e0.08\u003c/p\u003e\n \u003c/td\u003e\n \u003c/tr\u003e\n \u003ctr\u003e\n \u003ctd style=\"width: 96px;\"\u003e\n \u003cp\u003e\u003cstrong\u003eVML + Juvenile Transplant\u003c/strong\u003e\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 96px;\"\u003e\n \u003cp\u003e443.56 \u0026plusmn; 24.97\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 48px;\"\u003e\n \u003cp\u003e0.75\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e\u0026plusmn;\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e0.11\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 42px;\"\u003e\n \u003cp\u003e0.78\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e\u0026plusmn;\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e0.15\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 54px;\"\u003e\n \u003cp\u003e0.17\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e\u0026plusmn;\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e0.01\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 48px;\"\u003e\n \u003cp\u003e0.20\u003c/p\u003e\n \u003cp\u003e\u0026plusmn;\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e0.03\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 54px;\"\u003e\n \u003cp\u003e0.18\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e\u0026plusmn;\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e0.04\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 54px;\"\u003e\n \u003cp\u003e0.19\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e\u0026plusmn;\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e0.03\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 66px;\"\u003e\n \u003cp\u003e1.90\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e\u0026plusmn;\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e0.15\u003c/p\u003e\n \u003c/td\u003e\n \u003ctd style=\"width: 66px;\"\u003e\n \u003cp\u003e2.00\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e\u0026plusmn;\u0026nbsp;\u003c/p\u003e\n \u003cp\u003e0.15\u003c/p\u003e\n \u003c/td\u003e\n \u003c/tr\u003e\n \u003c/tbody\u003e\n\u003c/table\u003e\n\u003cp\u003e\u003cstrong\u003e\u003cem\u003eNotes: Tibialis Anterior (TA), Extensor Digitorum Longus (EDL), Gastrocnemius (Gastroc). Bold numbers indicate significance.\u003c/em\u003e\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eIn-Vivo Strength Measurements\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eFunctional Recovery\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eTo assess whether muscle transplantation improved force production, in-vivo isometric strength of the TA/EDL complex was measured in most of the uninjured right limbs (controls; n=19) and all of the VML injured limbs (n=24) seven weeks post-injury. Uninjured control limbs generated the highest absolute force (20.96 \u0026plusmn; 3.04 mN\u0026middot;m, n=19), significantly greater than all VML-injured groups (\u003cem\u003ep\u003c/em\u003e \u0026lt; 0.0001). No Treatment animals exhibited the lowest force output (9.47 \u0026plusmn; 0.67 mN\u0026middot;m, n=8). Adult transplants restored force to 12.94 \u0026plusmn; 1.52 mN\u0026middot;m (n=8; \u003cem\u003ep\u003c/em\u003e = 0.0299 vs. No Treatment), while juvenile transplants reached 14.25 \u0026plusmn; 2.01 mN\u0026middot;m (n=8; \u003cem\u003ep\u003c/em\u003e = 0.0016 vs. No Treatment), though not significantly greater than adult (\u003cem\u003ep\u003c/em\u003e = 0.6992).\u003c/p\u003e\n\u003cp\u003eNormalized to body weight, a similar pattern emerged. Control limbs produced 48.12 \u0026plusmn; 6.44 mN\u0026middot;m/kg, while VML No Treatment animals fell to 23.19 \u0026plusmn; 2.50. Juvenile and adult transplants improved force to 32.16 \u0026plusmn; 4.42 and 29.12 \u0026plusmn; 4.53 mN\u0026middot;m/kg, respectively. Only juvenile transplants produced a statistically significant improvement over No Treatment (\u003cem\u003ep\u003c/em\u003e = 0.0141), while adult transplants did not (\u003cem\u003ep\u003c/em\u003e = 0.1670). No significant difference was observed between transplant groups (\u003cem\u003ep\u003c/em\u003e = 0.7025). These data suggest that both donor types support partial recovery of force, with juvenile transplants trending slightly higher.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eHistological Assessment of Regeneration\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eTransverse TA sections were analyzed in a subset of animals from each group (n=4) to evaluate fiber number and muscle size, representing fiber maturity and regeneration. Total myofiber number was significantly reduced in the No Treatment group (6,952 \u0026plusmn; 743) compared to uninjured controls (10,316 \u0026plusmn; 685; \u003cem\u003ep\u003c/em\u003e \u0026lt; 0.01). Transplants restored fiber number, with adult recipients averaging 9,115 \u0026plusmn; 1,274 fibers and juvenile recipients 11,369 \u0026plusmn; 1,511, which were comparable to controls (Figure 4E).\u003c/p\u003e\n\u003cp\u003eTA muscle CSA was lowest in the No Treatment group (22.7 \u0026plusmn; 5.5 mm\u0026sup2;), significantly smaller than controls (34.3 \u0026plusmn; 1.0 mm\u0026sup2;; \u003cem\u003ep\u003c/em\u003e = 0.0265). Transplantation preserved CSA (adult: 30.6 \u0026plusmn; 3.6 mm\u0026sup2;; juvenile: 32.7 \u0026plusmn; 6.6 mm\u0026sup2;), as neither group differed significantly from controls (Figure 4F).\u003c/p\u003e\n\u003cp\u003eDespite improvements in fiber number and muscle size, the average myofiber CSA remained significantly reduced in all VML groups. Controls averaged 2,569 \u0026plusmn; 334 \u0026micro;m\u0026sup2;, compared to 1,379 \u0026plusmn; 290 (No Treatment), 1,258 \u0026plusmn; 97 (adult transplant), and 923 \u0026plusmn; 151 \u0026micro;m\u0026sup2; (juvenile transplant) (Figure 4G). CSA distribution curves showed that juvenile transplants had the highest proportion of small-diameter fibers (\u0026lt;1000 \u0026micro;m\u0026sup2;), suggesting ongoing regeneration or limited hypertrophy of transplanted fibers at this potentially early time point (Figure 4H).\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eIntegration of GFP⁺ Donor Fibers\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eTo assess integration of transplanted tissue, GFP fluorescence was analyzed. All but one transplant recipient \u0026nbsp;displayed a clear GFP⁺ region within the injury zone (Figure 5A\u0026ndash;C). GFP⁺ area occupied 10.78 \u0026plusmn; 7.27% of the total muscle CSA in adult recipients (n=7) and 12.57 \u0026plusmn; 7.39% in juvenile recipients (n=8), with no significant difference between groups (\u003cem\u003ep\u003c/em\u003e = 0.6412; Figure 5D).\u003c/p\u003e\n\u003cp\u003eAverage CSA of GFP⁺ fibers was smaller in juvenile transplants (518.6 \u0026plusmn; 230.7 \u0026micro;m\u0026sup2;; n=8) than in adult transplants (707.9 \u0026plusmn; 246.6 \u0026micro;m\u0026sup2;; n=7), though this difference did not reach statistical significance (\u003cem\u003ep\u003c/em\u003e=0.1735). Distribution profiles confirmed that juvenile transplants contained a higher proportion of small-diameter GFP⁺ fibers, while adult transplants included a broader range of sizes (Figure 5E). These patterns likely reflect developmental differences in donor fiber size and post-transplant remodeling.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eSatellite Cell Distribution\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eTo evaluate satellite cell content following VML, Pax7⁺ cells were quantified across four regions of interest: defect, transplant, border, and distal. In uninjured controls, satellite cells were evenly distributed (8.01 \u0026plusmn; 1.02 cells/mm\u0026sup2;). VML injury led to a mild decline in the defect region (3.61 \u0026plusmn; 2.33), with increased cells in the border region (27.24 \u0026plusmn; 8.31) and no change in distal (12.91 \u0026plusmn; 5.31) regions.\u003c/p\u003e\n\u003cp\u003eTransplantation did not significantly alter satellite cell density in host muscle regions compared to No Treatment. Juvenile transplant recipients had densities of 4.96 \u0026plusmn; 5.21 (defect), 26.01 \u0026plusmn; 10.40 (border), and 9.53 \u0026plusmn; 5.40 (distal). Adult transplant values were similar: 2.24 \u0026plusmn; 1.56, 31.29 \u0026plusmn; 10.69, and 8.76 \u0026plusmn; 4.55, respectively.\u003c/p\u003e\n\u003cp\u003eAs expected, satellite cell density was significantly elevated within the GFP⁺ transplant region: 27.11 \u0026plusmn; 13.23 cells/mm\u0026sup2; in juvenile and 33.63 \u0026plusmn; 14.88 in adult recipients. These values were significantly higher than in control regions, suggesting that the transplants retained a rich satellite cell microenvironment. Surprisingly, satellite cell content was not different between adult and juvenile groups in the transplant region, nor did it alter content in the existing tissue regions.\u0026nbsp;\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eCentralized Nuclei\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThe prevalence of centralized nuclei, an indicator of regenerating myofibers, was negligible in uninjured control muscle (0.60 \u0026plusmn; 0.08%) but elevated in all VML injured muscle (Figure 6D). In No Treatment animals, centralized nuclei were highest in the border region (27.9 \u0026plusmn; 4.6%) and lower in distal muscle (5.8 \u0026plusmn; 1.2%).\u003c/p\u003e\n\u003cp\u003eTransplantation produced similar regional patterns, with centralized nuclei enriched in the border and GFP⁺ regions, while distal muscle remained low. Adult transplants contained 20.7 \u0026plusmn; 3.1% centralized nuclei in the border, 15.0 \u0026plusmn; 2.8% in the GFP⁺ transplant region, and 4.4 \u0026plusmn; 1.0% distally. Juvenile transplants exhibited higher regenerative activity, with 29.3 \u0026plusmn; 5.0% centralized nuclei in the border and 23.3 \u0026plusmn; 4.2% in the GFP⁺ region, while distal muscle remained low (6.5 \u0026plusmn; 1.4%).\u003c/p\u003e\n\u003cp\u003eCentralized nuclei were not quantified in the defect region due to the limited number of small regenerating fibers present in that region. Together, these results show that centralized nuclei and satellite cells were concentrated within and adjacent to the transplant, consistent with localized regeneration. Juvenile transplants displayed higher levels of centralized nuclei in both the border and GFP⁺ regions compared to adult transplants, suggesting enhanced but spatially restricted regenerative activity at seven weeks post-surgery.\u003c/p\u003e\n\u003cp\u003eOf note, because cytosolic GFP is water-soluble, aqueous steps during the immunohistochemistry process can minimize GFP fluorescence, a process often referred to as \u0026ldquo;leaching\u0026rdquo;. To avoid over-interpretation, quantifications of GFP⁺ area, GFP⁺ fiber CSA, satellite cell and central nuclei counts related to GFP areas were performed only on sections with visually detectable GFP signal; samples where GFP was not clearly detected were excluded from GFP-specific analyses but were used for all other outcomes.\u0026nbsp;\u003c/p\u003e"},{"header":"DISCUSSION","content":"\u003cp\u003eThis study provides the first direct in-vivo comparison of juvenile versus adult skeletal muscle transplants for the treatment of VML. We found that juvenile donor tissue possessed markedly higher satellite cell density, smaller myofibers (allowing the transfer of more fibers per area), and enhanced in-vitro differentiation capacity compared to adult tissue. Despite these cellular advantages, both juvenile and adult transplants restored myofiber number and partially improved muscle strength to a similar extent seven weeks post-surgery. Functional recovery remained incomplete in both groups, and donor fibers were predominantly small at this time point. These findings indicate that donor muscle age influences intrinsic regenerative properties but does not necessarily translate into superior short-term, in-vivo outcomes in the absence of additional regenerative cues.\u003c/p\u003e\n\u003cp\u003eIn Study 1, our developmental analysis confirmed well-established age-related trends in muscle biology: nuclear and satellite cell densities were highest in juvenile muscle and declined progressively with maturation, consistent with prior studies in rodents and humans (10-12). The smaller fiber CSA and higher myogenic differentiation capacity of juvenile myoblasts support the concept that this tissue retains a more growth-permissive phenotype, which has been linked to increased adaptability and regeneration in other contexts (21-23). These properties provided a strong rationale for testing juvenile muscle as a donor source in VML repair.\u003c/p\u003e\n\u003cp\u003eIn Study 2, both juvenile and adult transplants integrated into the defect site, contributed GFP\u003csup\u003e+\u003c/sup\u003e donor fibers, and restored fiber counts to that of controls. Functional gains relative to untreated VML ranged from 35-50%, comparable to improvements reported in previous autologous or minced muscle transplant studies (7, 8). The lack of significant differences between juvenile and adult groups may be explained by several factors. First, the juvenile advantage in satellite cell number may have been transient; by seven weeks, many donor satellite cells could have differentiated into myofibers, leading to normalization between groups. Second, the immature size of donor fibers in both groups suggests that hypertrophy was still incomplete at this time point, limiting functional recovery. Finally, the restrictive microenvironment of the VML defect, characterized by altered extracellular matrix composition, persistent denervation, and limited vascularization (2, 5, 6), may have constrained the integration of juvenile tissue\u0026rsquo;s full regenerative potential.\u003c/p\u003e\n\u003cp\u003eSatellite cell analyses reinforced this interpretation. In both donor age groups, Pax7⁺ cell enrichment was largely confined to the transplant region, with no apparent expansion into surrounding host muscle. This localized effect aligns with previous observations that satellite cells preferentially remain within areas containing intact basal lamina and supportive extracellular matrix (18, 24). Without such cues, migration into the broader defect is limited, which likely contributed to the absence of greater regenerative effects.\u003c/p\u003e\n\u003cp\u003eCentralized nuclei findings further support this interpretation. Both transplant groups demonstrated localized enrichment of regenerating fibers within the border and GFP⁺ transplant regions, whereas distal muscle remained largely unaffected. Juvenile recipients exhibited higher proportions of centralized nuclei in both the border and transplant regions compared to adult recipients, suggesting more robust regenerative activity. However, these differences did not correspond to superior muscle size or force at seven weeks, indicating that the regenerative process was still incomplete and may require longer time frames or additional cues to translate into functional benefit.\u003c/p\u003e\n\u003cp\u003eFrom a translational perspective, these results underscore that donor tissue composition, while important, is only one determinant of transplant efficacy. The persistence of small-diameter fibers and limited functional recovery in both groups suggest that mechanical loading and pro-hypertrophic stimuli are needed to maximize transplant benefit (18-20). Rehabilitation strategies such as voluntary wheel running, resistance exercise, or electrical stimulation have been shown to enhance myofiber hypertrophy, satellite cell activation, and neuromuscular integration after injury (19, 20). Incorporating such interventions into future transplant protocols could amplify the intrinsic advantages of juvenile-like donor tissue.\u003c/p\u003e\n\u003cp\u003eOverall, this work is limited by several factors: first, we assessed the regenerative outcomes using a single time point of seven weeks. While this was intended to capture the regenerative window for muscle injury, it could be too early to detect long-term integration of the transplant, such as hypertrophy, reinnervation, and matrix remodeling. A longer follow-up may reveal whether age-related advantages or transplant integration change as fibers mature. Second, we deliberately implemented a transplant-only strategy, to isolate donor-age effects under controlled conditions. This baseline now creates a platform to test whether targeted loading (rehabilitation) or additional biological cues can enhance the benefits of juvenile tissue. Third, donor integration was quantified using GFP-based histology, which is susceptible to leaking; future work could incorporate other tracking methods, such as tissue clearing with 3D quantification. Finally, experiments were performed in male Lewis rats to mirror the male predominance of combat-related VML; confirming these findings in females and across additional genetic backgrounds will be an important step toward broad translation.\u003c/p\u003e"},{"header":"CONCLUSION","content":"\u003cp\u003eJuvenile skeletal muscle displays cellular and structural characteristics consistent with high regenerative potential, including high satellite cell content, small myofibers, greater in-vitro differentiation capacity, and elevated proportions of centralized nuclei within transplant regions. In this VML model, both juvenile and adult donor tissue integrated into the defect, restored myofiber number, and partially improved muscle strength; however, functional outcomes were similar between groups at seven weeks post-surgery. These results indicate that optimizing donor age alone is insufficient for full functional recovery and that additional regenerative cues, or time, are required.\u003c/p\u003e\n\u003cp\u003eThe high regenerative profile of juvenile muscle provides a valuable biological benchmark for engineered muscle constructs. Myogenic progenitors derived from induced pluripotent stem cells or other stem cell sources could be directed toward a juvenile-like phenotype before transplantation (21, 22, 25). Additionally, emerging work with human\u0026ndash;animal chimeric models offer a promising and scalable approach to sourcing juvenile-like transplant tissue (23, 26). Such strategies may reduce logistical barriers, including long-term animal care costs, and accelerate tissue availability. When paired with biomaterials and/or with rehabilitation interventions that promote hypertrophy and integration, these approaches could substantially improve structural and functional outcomes for patients with VML.\u003c/p\u003e"},{"header":"Abbreviations","content":"\u003cdiv class=\"DefinitionList\"\u003e\u003cdiv class=\"DefinitionListEntry\"\u003e\u003cdiv class=\"Term\"\u003eCSA\u003c/div\u003e\u003cdiv class=\"Description\"\u003e\u003cp\u003ecross-sectional area\u003c/p\u003e\u003c/div\u003e\u003c/div\u003e\u003cdiv class=\"DefinitionListEntry\"\u003e\u003cdiv class=\"Term\"\u003eDAPI\u003c/div\u003e\u003cdiv class=\"Description\"\u003e\u003cp\u003e4\u0026prime;,6-diamidino-2-phenylindole\u003c/p\u003e\u003c/div\u003e\u003c/div\u003e\u003cdiv class=\"DefinitionListEntry\"\u003e\u003cdiv class=\"Term\"\u003eEDL\u003c/div\u003e\u003cdiv class=\"Description\"\u003e\u003cp\u003eextensor digitorum longus\u003c/p\u003e\u003c/div\u003e\u003c/div\u003e\u003cdiv class=\"DefinitionListEntry\"\u003e\u003cdiv class=\"Term\"\u003eFBS\u003c/div\u003e\u003cdiv class=\"Description\"\u003e\u003cp\u003efetal bovine serum\u003c/p\u003e\u003c/div\u003e\u003c/div\u003e\u003cdiv class=\"DefinitionListEntry\"\u003e\u003cdiv class=\"Term\"\u003eGFP\u003c/div\u003e\u003cdiv class=\"Description\"\u003e\u003cp\u003egreen fluorescent protein\u003c/p\u003e\u003c/div\u003e\u003c/div\u003e\u003cdiv class=\"DefinitionListEntry\"\u003e\u003cdiv class=\"Term\"\u003eIACUC\u003c/div\u003e\u003cdiv class=\"Description\"\u003e\u003cp\u003eInstitutional Animal Care and Use Committee\u003c/p\u003e\u003c/div\u003e\u003c/div\u003e\u003cdiv class=\"DefinitionListEntry\"\u003e\u003cdiv class=\"Term\"\u003eIHC\u003c/div\u003e\u003cdiv class=\"Description\"\u003e\u003cp\u003eimmunohistochemistry\u003c/p\u003e\u003c/div\u003e\u003c/div\u003e\u003cdiv class=\"DefinitionListEntry\"\u003e\u003cdiv class=\"Term\"\u003eMyHC\u003c/div\u003e\u003cdiv class=\"Description\"\u003e\u003cp\u003emyosin heavy chain\u003c/p\u003e\u003c/div\u003e\u003c/div\u003e\u003cdiv class=\"DefinitionListEntry\"\u003e\u003cdiv class=\"Term\"\u003eNGS\u003c/div\u003e\u003cdiv class=\"Description\"\u003e\u003cp\u003enormal goat serum\u003c/p\u003e\u003c/div\u003e\u003c/div\u003e\u003cdiv class=\"DefinitionListEntry\"\u003e\u003cdiv class=\"Term\"\u003ePax7\u003c/div\u003e\u003cdiv class=\"Description\"\u003e\u003cp\u003epaired box protein 7\u003c/p\u003e\u003c/div\u003e\u003c/div\u003e\u003cdiv class=\"DefinitionListEntry\"\u003e\u003cdiv class=\"Term\"\u003ePBS\u003c/div\u003e\u003cdiv class=\"Description\"\u003e\u003cp\u003ephosphate-buffered saline\u003c/p\u003e\u003c/div\u003e\u003c/div\u003e\u003cdiv class=\"DefinitionListEntry\"\u003e\u003cdiv class=\"Term\"\u003eTA\u003c/div\u003e\u003cdiv class=\"Description\"\u003e\u003cp\u003etibialis anterior\u003c/p\u003e\u003c/div\u003e\u003c/div\u003e\u003cdiv class=\"DefinitionListEntry\"\u003e\u003cdiv class=\"Term\"\u003eVML\u003c/div\u003e\u003cdiv class=\"Description\"\u003e\u003cp\u003evolumetric muscle loss\u003c/p\u003e\u003c/div\u003e\u003c/div\u003e\u003cdiv class=\"DefinitionListEntry\"\u003e\u003cdiv class=\"Term\"\u003eWGA\u003c/div\u003e\u003cdiv class=\"Description\"\u003e\u003cp\u003ewheat germ agglutinin\u003c/p\u003e\u003c/div\u003e\u003c/div\u003e\u003c/div\u003e"},{"header":"Declarations","content":"\u003cp\u003e\u003cstrong\u003eEthics Approval and Consent to Participate\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eAll animal procedures were approved on April 1, 2023, by the Institutional Animal Care and Use Committee (IACUC) at Brigham Young University under the title: A novel approach to improve skeletal muscle transplant efficiency following volumetric muscle loss injury, protocol number 23-0401. All methods were carried out in accordance with relevant guidelines and regulations for the care and use of laboratory animals.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eConsent for Publication\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eNot applicable.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eAvailability of Data and Materials\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThe datasets generated and/or analyzed during the current study are available from the corresponding author on reasonable request.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eCompeting Interests\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThe authors declare that they have no competing interests.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eFunding\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThis research was supported by internal funding from the College of Life Sciences at Brigham Young University. The funding body had no role in the design of the study, data collection, analysis, interpretation, or manuscript writing.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eAuthors\u0026rsquo; Contributions\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eJRS conceived and designed the study. JJP, SRF, ZHR, MJM, ACT, KMJ, TKS, and MKK performed experiments and data collection. SSH, and EWE, contributed to data analysis. JRS drafted the manuscript. All authors contributed to the manuscript revision and approved the final version of the manuscript.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eAcknowledgements\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThe authors would like to thank Anson Wood for piloting the cell culture model, Kayli Denmark for piloting IHC protocols, Chandler Bowen, and Dre Sorensen for assistance in image analysis, and the use of AI for assistance with readability, grammar and punctuation.\u003c/p\u003e"},{"header":"References","content":"\u003col\u003e\n\u003cli\u003eGreising SM, Rivera JC, Goldman SM, Watts A, Aguilar CA, Corona BT. Unwavering pathobiology of volumetric muscle loss injury. Scientific Reports. 2017;7(1):13179.\u003c/li\u003e\n\u003cli\u003eHoffman DB, Raymond-Pope CJ, Sorensen JR, Corona BT, Call JA, Greising SM. Temporal changes in the muscle extracellular matrix due to volumetric muscle loss injury. Connective Tissue Research. 2022;63(2):124\u0026ndash;37.\u003c/li\u003e\n\u003cli\u003eBrack AS, Rando TA. Intrinsic changes and extrinsic influences of myogenic stem cell function during aging. Stem Cell Reviews and Reports. 2007;3(3):226\u0026ndash;37.\u003c/li\u003e\n\u003cli\u003eKuang S, Kuroda K, Le Grand F, Rudnicki MA. Asymmetric self-renewal and commitment of satellite stem cells in muscle. Cell. 2007;129(5):999\u0026ndash;1010.\u003c/li\u003e\n\u003cli\u003eGarg K, Ward CL, Hurtgen BJ, Wilken JM, Stinner DJ, Wenke JC, et al. Volumetric muscle loss: persistent functional deficits beyond frank loss of tissue. Journal of Orthopaedic Research. 2015;33(1):40\u0026ndash;6.\u003c/li\u003e\n\u003cli\u003eSorensen JR, Hoffman DB, Corona BT, Greising SM. Secondary denervation is a chronic pathophysiologic sequela of volumetric muscle loss. Journal of Applied Physiology. 2021;130(5):1552\u0026ndash;65.\u003c/li\u003e\n\u003cli\u003eGrogan BF, Hsu JR. Volumetric muscle loss. Journal of the American Academy of Orthopaedic Surgeons. 2011;19(Suppl 1):S35\u0026ndash;S7.\u003c/li\u003e\n\u003cli\u003eWard CL, Ji L, Corona BT. Autologous minced muscle grafts improve muscle strength in a porcine model of volumetric muscle loss injury. Journal of Applied Physiology. 2016;120(6):915\u0026ndash;23.\u003c/li\u003e\n\u003cli\u003eCorona BT, Rivera JC, Owens JG, Wenke JC, Rathbone CR. Volumetric muscle loss leads to permanent disability following extremity trauma. Journal of Rehabilitation Research and Development. 2015;52(7):785\u0026ndash;92.\u003c/li\u003e\n\u003cli\u003eGarc\u0026iacute;a-Prat L, Mu\u0026ntilde;oz-C\u0026aacute;noves P. Muscle stem cell aging: regulation and rejuvenation. Skeletal Muscle. 2021;11(1):4.\u003c/li\u003e\n\u003cli\u003eLexell J, Taylor CC, Sj\u0026ouml;str\u0026ouml;m M. What is the cause of the ageing atrophy? Total number, size and proportion of different fiber types studied in whole vastus lateralis muscle from 15‐ to 83‐year‐old men. Journal of the Neurological Sciences. 1988;84(2-3):275\u0026ndash;94.\u003c/li\u003e\n\u003cli\u003eSchiaffino S, Reggiani C. Fiber types in mammalian skeletal muscles. Physiological Reviews. 2011;91(4):1447\u0026ndash;531.\u003c/li\u003e\n\u003cli\u003eSmith LR, Barton ER. Collagen content does not alter the passive mechanical properties of fibrotic skeletal muscle in mdx mice. American Journal of Physiology-Cell Physiology. 2014;306(10):C889\u0026ndash;C98.\u003c/li\u003e\n\u003cli\u003eSnow MH. Satellite cell response in rat soleus muscle undergoing hypertrophy due to surgical ablation of synergists. Anatomical Record. 1977;189(2):479\u0026ndash;97.\u003c/li\u003e\n\u003cli\u003eChal J, Oginuma M, Al Tanoury Z, Gobert B, Sumara O, Hick A, et al. Differentiation of pluripotent stem cells to muscle fiber to model Duchenne muscular dystrophy. Nature Biotechnology. 2015;33(9):962\u0026ndash;9.\u003c/li\u003e\n\u003cli\u003eHoffman DB, Basten AM, Sorensen JR, Raymond-Pope CJ, Lillquist TJ, Call JA, et al. Response of terminal Schwann cells following volumetric muscle loss injury. Experimental Neurology. 2023;357:114431.\u003c/li\u003e\n\u003cli\u003eSorensen JR, Hoffman DB, Raymond-Pope CJ, Lillquist TJ, Russell AM, Corona BT, et al. Inhibition of ErbB2 mitigates secondary denervation after traumatic muscle injury. Journal of Physiology. 2025;0(0):1\u0026ndash;18.\u003c/li\u003e\n\u003cli\u003eGreising SM, Call JA. When is the right time to initiate rehabilitation? Time will tell\u0026hellip;. Exp Physiol. 2024;109(6):889\u0026ndash;91.\u003c/li\u003e\n\u003cli\u003eNakayama KH, Quarta M, Paine P, Alcazar C, Karakikes I, Garcia V, et al. Rehabilitation following skeletal muscle injury enhances tissue regeneration and function. NPJ Regenerative Medicine. 2019;4:3.\u003c/li\u003e\n\u003cli\u003eSerrano AL, Baeza-Raja B, Perdiguero E, Jard\u0026iacute; M, Mu\u0026ntilde;oz-C\u0026aacute;noves P. Interleukin-6 is an essential regulator of satellite cell-mediated skeletal muscle hypertrophy. Cell Metabolism. 2008;7(1):33\u0026ndash;44.\u003c/li\u003e\n\u003cli\u003eCarosio S, Barberi L, Rizzuto E, Nicoletti C, Musar\u0026ograve; A. Generation of eX vivo-vascularized Muscle Engineered Tissue (X-MET). Scientific Reports. 2013;3:1420.\u003c/li\u003e\n\u003cli\u003eChal J, Oginuma M, Al Tanoury Z, Gobert B, Sumara O, Hick A, et al. Differentiation of pluripotent stem cells to muscle fiber to model Duchenne muscular dystrophy. Nature biotechnology. 2015;33(9):962\u0026ndash;9.\u003c/li\u003e\n\u003cli\u003eMaeng G, Das S, Greising SM, Gong W, Singh BN, Kren S, et al. Humanized skeletal muscle in MYF5/MYOD/MYF6-null pig embryos. Nature biomedical engineering. 2021;5(8):805\u0026ndash;14.\u003c/li\u003e\n\u003cli\u003eGreising SM, Dearth CL, Corona BT. Regenerative and rehabilitative medicine: a necessary synergy for functional recovery from volumetric muscle loss injury. Cells Tissues Organs. 2016;202(3-4):237\u0026ndash;49.\u003c/li\u003e\n\u003cli\u003eRao L, Qian Y, Khodabukus A, Ribar T, Bursac N. Engineering human pluripotent stem cells into a functional skeletal muscle tissue. Nature Communications. 2018;9:126.\u003c/li\u003e\n\u003cli\u003eChoe Y-H, Das S, Ma X, Lee H, Sorensen JR, Hoffman DB, et al. Porcine myogenesis in cloned wildtype and MYF5/MYOD/MYF6-null porcine embryo. Communications biology. 2025;8(1):217.\u003c/li\u003e\n\u003c/ol\u003e"}],"fulltextSource":"","fullText":"","funders":[],"hasAdminPriorityOnWorkflow":false,"hasManuscriptDocX":true,"hasOptedInToPreprint":true,"hasPassedJournalQc":"","hasAnyPriority":false,"hideJournal":false,"highlight":"","institution":"","isAcceptedByJournal":true,"isAuthorSuppliedPdf":false,"isDeskRejected":"","isHiddenFromSearch":false,"isInQc":false,"isInWorkflow":false,"isPdf":false,"isPdfUpToDate":true,"isWithdrawnOrRetracted":false,"journal":{"display":true,"email":"[email protected]","identity":"stem-cell-research-and-therapy","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":false,"externalIdentity":"scrt","sideBox":"Learn more about [Stem Cell Research \u0026 Therapy](http://stemcellres.biomedcentral.com)","snPcode":"","submissionUrl":"https://www.editorialmanager.com/scrt/default.aspx","title":"Stem Cell Research \u0026 Therapy","twitterHandle":"@BioMedCentral","acdcEnabled":true,"dfaEnabled":true,"editorialSystem":"em","reportingPortfolio":"BMC/SO AJ","inReviewEnabled":true,"inReviewRevisionsEnabled":true},"keywords":"volumetric muscle loss, skeletal muscle transplantation, satellite cells, juvenile muscle, donor age, muscle regeneration, tissue engineering, stem cell therapy, regenerative medicine","lastPublishedDoi":"10.21203/rs.3.rs-7437055/v1","lastPublishedDoiUrl":"https://doi.org/10.21203/rs.3.rs-7437055/v1","license":{"name":"CC BY 4.0","url":"https://creativecommons.org/licenses/by/4.0/"},"manuscriptAbstract":"\u003ch2\u003eBackground\u003c/h2\u003e\u003cp\u003eVolumetric muscle loss (VML) causes irreversible structural and functional deficits by removing myofibers, nerves, vasculature, extracellular matrix, and satellite cells, the resident muscle stem cells essential for regeneration. Skeletal muscle transplantation can restore tissue volume and reintroduce regenerative cells, yet functional outcomes remain incomplete. Age of the donor muscle has not been evaluated, despite evidence that juvenile muscle contains higher satellite cell density and greater myogenic plasticity than adult muscle. We hypothesized that these features would yield superior regenerative outcomes when juvenile muscle is used as a transplant source.\u003c/p\u003e\u003ch2\u003eMethods\u003c/h2\u003e\u003cp\u003eTibialis anterior (TA) muscles from juvenile (21 d), adolescent (34 d), and adult (~\u0026thinsp;120 d) male Lewis rats were compared for myofiber morphology, satellite cell density, and in-vitro myogenic behavior. GFP⁺ juvenile or adult muscle was then transplanted into standardized VML defects (~\u0026thinsp;15\u0026ndash;20% TA volume) in adult rats. Seven weeks post-surgery, in-vivo isometric strength, donor fiber integration, satellite cell distribution, and centralized nuclei were assessed.\u003c/p\u003e\u003ch2\u003eResults\u003c/h2\u003e\u003cp\u003eJuvenile muscle exhibited\u0026thinsp;~\u0026thinsp;15\u0026times; greater satellite cell density than adult (122.8\u0026thinsp;\u0026plusmn;\u0026thinsp;28.4 vs. 8.4\u0026thinsp;\u0026plusmn;\u0026thinsp;3.3 cells/mm\u0026sup2;, p\u0026thinsp;\u0026lt;\u0026thinsp;0.0001) with enhanced in-vitro differentiation (fusion index\u0026thinsp;+\u0026thinsp;73% vs. adult, p\u0026thinsp;=\u0026thinsp;0.0067). In-vivo, both juvenile and adult transplants restored myofiber number to control levels (juvenile: 11,369\u0026thinsp;\u0026plusmn;\u0026thinsp;1,511; adult: 9,115\u0026thinsp;\u0026plusmn;\u0026thinsp;1,274; controls: 10,316\u0026thinsp;\u0026plusmn;\u0026thinsp;685) and improved strength versus untreated VML (juvenile: +50%, p\u0026thinsp;=\u0026thinsp;0.0016; adult: +36%, p\u0026thinsp;=\u0026thinsp;0.0299). No significant functional differences were observed between donor ages. Donor fibers integrated but remained small, with localized satellite cell enrichment and increased centralized nuclei in transplant regions, consistent with ongoing regeneration.\u003c/p\u003e\u003ch2\u003eConclusions\u003c/h2\u003e\u003cp\u003eJuvenile skeletal muscle displays cellular and structural attributes favorable for regeneration and superior in-vitro myogenic behavior compared to adult muscle. However, these advantages did not translate into greater short-term in-vivo recovery following VML transplantation. Enhancing donor fiber hypertrophy, neuromuscular integration, and satellite cell expansion beyond the transplant region, potentially through rehabilitation or pharmaceutical interventions, may be necessary to realize the full therapeutic potential of juvenile donor muscle for regenerative medicine applications.\u003c/p\u003e","manuscriptTitle":"Juvenile vs Adult Skeletal Muscle Transplants in the Treatment of Volumetric Muscle Loss Injury","msid":"","msnumber":"","nonDraftVersions":[{"code":1,"date":"2025-10-06 09:26:11","doi":"10.21203/rs.3.rs-7437055/v1","editorialEvents":[{"type":"communityComments","content":0},{"type":"decision","content":"Revision requested","date":"2025-10-13T22:23:08+00:00","index":"","fulltext":""},{"type":"editorInvitedReview","content":"","date":"2025-10-09T20:11:27+00:00","index":"hide","fulltext":""},{"type":"editorInvitedReview","content":"","date":"2025-10-02T10:42:09+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"133757983596855727148497108785689107525","date":"2025-09-29T17:56:28+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"32892143909883830321982256307021273","date":"2025-09-23T09:38:50+00:00","index":"hide","fulltext":""},{"type":"reviewersInvited","content":"","date":"2025-09-23T00:01:00+00:00","index":"","fulltext":""},{"type":"editorAssigned","content":"","date":"2025-09-18T18:30:02+00:00","index":"","fulltext":""},{"type":"checksComplete","content":"","date":"2025-09-11T03:29:25+00:00","index":"","fulltext":""},{"type":"submitted","content":"Stem Cell Research \u0026 Therapy","date":"2025-09-08T14:58:09+00:00","index":"","fulltext":""}],"status":"published","journal":{"display":true,"email":"[email protected]","identity":"stem-cell-research-and-therapy","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":false,"externalIdentity":"scrt","sideBox":"Learn more about [Stem Cell Research \u0026 Therapy](http://stemcellres.biomedcentral.com)","snPcode":"","submissionUrl":"https://www.editorialmanager.com/scrt/default.aspx","title":"Stem Cell Research \u0026 Therapy","twitterHandle":"@BioMedCentral","acdcEnabled":true,"dfaEnabled":true,"editorialSystem":"em","reportingPortfolio":"BMC/SO AJ","inReviewEnabled":true,"inReviewRevisionsEnabled":true}}],"origin":"","ownerIdentity":"e2063de8-e362-465a-a8aa-0dc0bfeac581","owner":[],"postedDate":"October 6th, 2025","published":true,"recentEditorialEvents":[],"rejectedJournal":[],"revision":"","amendment":"","status":"published-in-journal","subjectAreas":[],"tags":[],"updatedAt":"2025-12-08T16:06:17+00:00","versionOfRecord":{"articleIdentity":"rs-7437055","link":"https://doi.org/10.1186/s13287-025-04844-y","journal":{"identity":"stem-cell-research-and-therapy","isVorOnly":false,"title":"Stem Cell Research \u0026 Therapy"},"publishedOn":"2025-12-03 15:57:57","publishedOnDateReadable":"December 3rd, 2025"},"versionCreatedAt":"2025-10-06 09:26:11","video":"","vorDoi":"10.1186/s13287-025-04844-y","vorDoiUrl":"https://doi.org/10.1186/s13287-025-04844-y","workflowStages":[]},"version":"v1","identity":"rs-7437055","journalConfig":"researchsquare"},"__N_SSP":true},"page":"/article/[identity]/[[...version]]","query":{"redirect":"/article/rs-7437055","identity":"rs-7437055","version":["v1"]},"buildId":"8U1c8b4HqxoKbykW_rLl7","isFallback":false,"isExperimentalCompile":false,"dynamicIds":[84888],"gssp":true,"scriptLoader":[]}