hUC-MSCs and derived exosomes attenuate DEX-induced muscle atrophy through modulation of estrogen signaling pathway

Abstract

Background

Sarcopenia, a multifactorial syndrome characterized by progressive loss of skeletal muscle mass and strength, combined with impaired physical function, is associated primarily with aging but also driven by chronic inflammation, immobility, and endocrine dysregulation. It leads to increased risks of frailty, falls, and loss of independence, posing a major public health challenge for aging populations. Although human umbilical cord-derived mesenchymal stem cells (hUC-MSCs) and their derived exosomes (MSC-Exos) have demonstrated remarkable potential in regenerative medicine, their safety and efficacy in treating sarcopenia remain unclear. To address this issue, we conducted a preclinical study to systematically evaluate their therapeutic potential and safety.

Methods

Male C57BL/6 J mice were treated with dexamethasone (20 mg/kg, i.p.) to induce muscle atrophy. Subsequently, bilateral intramuscular injection of hUC-MSCs (1 × 10⁶ cells/kg), exosomes (100 μg), or intraperitoneal injected of SNG162 (40 mg/kg) for two weeks. Gastrocnemius muscles were excised for histological analysis, TUNEL staining, Western blotting, RNA sequencing, and qPCR. Differentiated C2C12 myotubes were treated with 10 μM dexamethasone and co-cultured with hUC-MSCs or exosomes for 24 h. Samples were collected for qPCR, Western blot analyses and flow cytometry. EdU labeling was used to assess cell proliferation, MyHC and MDC immunofluorescence staining were employed to assess myotube morphology and autophagy levels, respectively. ELISA was used to quantify inflammatory cytokines and estrogen levels.

Results

hUC-MSCs, MSC-Exos and SNG162 improved grip strength and endurance in mice, increased the Gast muscle-to-body weight ratio without adversely affecting overall body weight, and enhanced muscle fiber cross-sectional area (CSA). Concurrently, they upregulated the expression of MyHC, Beclin-1, Bcl-2/Bax, ERα46, ERα36, ERβ and estradiol, while reducing key atrophy and inflammatory markers, including FOXO3, MAFbX, MURF1, TNF-α, IL-6, IL-1β, P62, and Caspase-3 in vitro and in vivo models. Furthermore, hUC-MSCs and MSC-Exos attenuated DEX-induced apoptosis in Gast muscles and C2C12 myotubes. Notably, MSC-Exos outperformed hUC-MSCs in promoting the proliferation and differentiation of C2C12 myotubes. Mechanistically, RNA sequencing and Western blot analysis identified the PI3K/AKT/mTOR and ERK1/2 signaling pathways as pivotal mediators of these effects.

Conclusions

This study underscores the potential of hUC-MSCs and their derived exosomes as a novel, safe, and effective therapeutic strategies for sarcopenia, offering promising avenues for clinical application.

Supplementary Information

The online version contains supplementary material available at 10.1186/s13287-025-04328-z.

Introduction

Sarcopenia, a progressive and multifactorial syndrome characterized by the accelerated loss of skeletal muscle mass, strength, and physical performance, has been recognized by the World Health Organization (WHO) as a major geriatric disease (ICD-10-CM code M62.84) [1, 2]. Emerging evidence suggests that its pathophysiology extends beyond chronological aging to involve neuroendocrine dysregulation, chronic low-grade inflammation, and mitochondrial dysfunction [3, 4]. Epidemiological studies reveal a marked gender disparity in the prevalence of sarcopenia: Premenopausal females exhibit significantly lower incidence rates compared to age-matched males, whereas postmenopausal females experience a near-exponential surge in prevalence, rapidly converging with their male counterparts within a decade of menopause. This trajectory strongly implicates estrogen depletion as a key driver of musculoskeletal decline. Current treatments for sarcopenia predominantly involve exercise interventions, protein and vitamin supplementation, and hormone replacement therapy [5–8]. Although these strategies remain the cornerstone of management, their overall efficacy is limited, highlighting an urgent need for innovative therapeutics.

Cell-derived therapies, particularly mesenchymal stem cell (MSC) based approaches for muscle degenerative diseases, hold immense potential for enhancing muscle tissue repair and providing dynamic biological responses to evolving microenvironmental conditions [9–13]. Among MSCs, human umbilical cord-derived mesenchymal stem cells (hUC-MSCs) are distinguished by their accessibility, high proliferative ability, strong differentiation capacity, and minimal immunogenicity, making them as an ideal candidates for regenerative medicine applications [14, 15]. Recent studies have demonstrated their exceptional muscle-protective and regenerative properties in preclinical models, emphasizing their potential as a novel therapeutic options for addressing sarcopenia and related muscle degenerative conditions [16, 17].

In contrast to stem cells, exosomes lack the capacity for self-renewal, thereby eliminating the risk of tumorigenesis following stem cell transplantation [18, 19]. Additionally, exosomes are more amenable to scalable production and storage, retaining their bioactivity even after freeze‒thaw cycles. These properties establish the use of MSC-derived exosomes (MSC-Exos) as a versatile and promising therapeutic strategy, particularly for chronic disease management and tissue repair [16, 20]. Interestingly, previous prior tracking studies did not observe the integration of MSCs into muscle tissue, indicating that their effects may be indirect [21]. Recent research revealed that MSCs primarily exert protective effects through exosome release [22, 23]. Notably, these exosomes replicate certain therapeutic properties of their originating cells while overcoming the inherent challenges linked to MSC-based treatments [24]. We hypothesized that the indirect events driven by the paracrine activity of MSCs might constitute the underlying mechanism of MSC-mediated muscle tissue repair.

Emerging evidence emphasize the critical function of estrogen in maintaining skeletal muscle integrity, particularly in postmenopausal women who experience accelerated reductions in muscle mass and strength [25]. Estrogen receptor α (ERα) is central to the intricate regulation of skeletal muscle metabolism and mitochondrial function, underscoring its importance in muscle homeostasis [26]. Estrogen is essential for modulating skeletal muscle metabolism and performance via complex signaling mechanisms [27]. However, the widespread clinical adoption of traditional estrogen replacement therapies is hindered by significant risks, such as an increased likelihood of developing breast cancer [28]. These findings underscore underscores the urgent need for safer and more targeted approaches to activate estrogen-related signaling pathways. Exosomes, which are composed of a lipid bilayer, encapsulate a diverse array of cytosolic and membrane proteins, including transcription factors, DNA, RNA, lipids, cytokines, growth factors, enzymes, and extracellular matrix proteins [20]. Notably, exosomes exhibit a distinctive ability to influence diverse biological activities by transferring bioactive compounds to adjacent cells and delivering genetic material to remote cell populations [18].

We hypothesize that the paracrine activity of hUC-MSCs and their derived exosomes modulates estrogen-related signaling pathways, thereby mitigating DEX-induced skeletal muscle atrophy. This study aimed to evaluate the effects of hUC-MSCs and MSC-Exos on muscle atrophy both in vivo and in vitro, elucidating the underlying molecular mechanisms involved. This research not only deepens our understanding of the treatment mechanisms of MSCs and MSC-Exo-based interventions for sarcopenia but also offers novel insights into future treatment strategies. These findings lay a promising foundation for more effective MSC-based therapies, potentially providing a transformative approach for individuals suffering from age-related muscle degeneration.

Materials and methods

hUC-MSCs culture and identification

Human umbilical cord-derived mesenchymal stem cells (hUC-MSCs) were prepared at the Stem Cell Clinical Research Center of the First Affiliated Hospital of Dalian Medical University, as previously described [29]. These cells were cultured in a conditioned medium composed of α-MEM base medium (Gibco, USA), 10% fetal bovine serum (FBS; Gibco, USA) and 1% penicillin–streptomycin (P/S; Gibco, USA) at 37 °C with 5% CO₂ concentration. The culture medium was replaced every 48 h. Cells morphology was monitored using an inverted microscope, and cells were passaged or cryopreserved upon reaching approximately 80% confluence. hUC-MSCs at passages 3–8 were utilized for all experiment. Immunophenotypic molecular markers were analyzed via flow cytometry (FACSDiva Software, BD). All experiments and protocols involving hUC-MSCs complied with the ethical principles established in the Declaration of Helsinki and received approval from the appropriate institutional ethics committee.

MSC-Exos isolation and identification

Human MSCs of the third generation were seeded in T-75 cm² culture flasks until reaching 80% confluency, and cells were washed three times with PBS(G4202-100ML, ServiceBio, China) after removing the cell culture medium. Subsequently, hUC-MSCs were incubated in Exosome-depleted FBS (UR50202, Umibio) at 37 °C in a 5% CO₂ concentration for 48 h, following which the culture supernatant was harvested for exosome isolation.

The exosome isolation protocol was adapted with minor modifications from previously published methods [30, 31]. Briefly, exosomes were extracted from the supernatant of hUC-MSCs cultured in exosome-depleted FBS for 48 h. The supernatant underwent sequential centrifugation at 500 × g for 10 min and 2,000 × g for 30 min to eliminate cells and debris, followed by ultracentrifugation at 10,000 × g for 60 min. The resulting supernatant was passed through a 0.22 μm sterile filter and subjected to further ultracentrifugation at 120,000 × g for 70 min. The exosome-containing pellet was resuspended in 200 μL of sterile PBS and stored at − 80 °C until needed.

The particle size of MSC-Exos was determined by a NanoSight nanoparticle tracking analyzer (NS300, Malvern, UK). The morphology of MSC-Exos was observed using transmission electron microscopy (JEM-1400, JEOL, Tokyo, Japan), and MSC-Exos markers, including CD9 (20597-1-AP, Proteintech), CD63 (67605-1-Ig, Proteintech), Tsg101 (28283-1-AP, Proteintech) and Calnexin (10427-2-AP, Proteintech), were analyzed by WB.

Animals and experimental design

All experimental animals in this study were purchased from the SPF Animal Laboratory Center of Dalian Medical University, China. All experiments were approved by the Ethics Committee of Dalian Medical University (Approval No. AEE24221) and conducted in accordance with institutional guidelines. All experiments were conducted and reported in accordance with the ARRIVE 2.0 guideline.

In our study, clean and healthy male C57BL/6 J mice (aged 8–10 weeks) were maintained in an environment at 22 ± 1 °C with 12-h light/dark cycle. The animals were housed under specific pathogen-free (SPF) conditions, provided with a standard diet, and allowed unrestricted access to water. The mice were randomly assigned to the following experimental groups (n = 6): Control: the control group. DEX: Intraperitoneal injection of 20 mg/kg dexamethasone daily for 7 days. DEX + hUC-MSCs: Twenty-four hours after the final DEX injection, 1 × 10⁶ cells/kg hUC-MSCs were administered via daily bilateral intramuscular injection into the gastrocnemius for 14 consecutive days. DEX + MSC-Exos: Twenty-four hours after the final DEX injection, 100 μg MSC-Exos (50 μg/μL per side) were injected bilaterally into the gastrocnemius daily for 14 days.

To investigate the role of estrogen in DEX-induced muscle atrophy, mice were randomly assigned to three experimental groups: Control: No special treatment. DEX: Muscle atrophy was induced via intraperitoneal injection of 20 mg/kg dexamethasone daily for 7 consecutive days. DEX + SNG162: Twenty-four hours after the final DEX injection, mice received daily intraperitoneal injections of the estrogen analog SNG162 at 40 mg/kg for 2 weeks.

The administration route, timing and frequency of DEX, MSCs and MSC-Exos was determined based on previous studies with comparable analyses [32–34]. Blood samples were collected 24 h after the final injection for biochemical analysis. Subsequently, mice were deeply anesthetized using 10% isoflurane and euthanized via cervical dislocation under anesthesia, in accordance with the guidelines established by the International Association for the Assessment and Accreditation of Laboratory Animal Care (AAALAC). All euthanasia procedures were conducted by trained personnel in a way that reduced potential distress to a minimum. Finally, the weights of soleus and gastrocnemius muscles were measured using an analytical balance and immediately preserved in liquid nitrogen for protein analysis or RNA Stabilization Solution (Thermo Fisher Scientific) for RNA stabilization prior to transcriptomic sequencing and gene expression studies.

Muscle function test

Grip strength test

The muscle strength of mice was measured using a grip strength tester (ZS-ZL, Zhongshi Dichuang Technology, Beijing, China). In brief, the mouse was lifted by its tail, allowing it to grasp the grip strength measuring plate while keeping its body parallel to the desktop. Then, the tail was gently pulled with a uniform force until the paws released the plate. The maximum grip strength of each mouse was determined by measuring the force exerted before releasing its grip. Each mouse underwent three trials, and the mean value was calculated. Grip strength measurements were performed weekly from the start of the experiment. For each measurement session, the grip strength of each mouse was measured five times. The mean value of these five measurements was first calculated, and then this value was normalized by the corresponding body weight of the mouse to represent the standardized grip strength (N/g) for each individual.

Muscle endurance capacity test

Muscle endurance capacity was evaluated using a four-limb hanging test. Mice were placed on a wire grid (2 cm mesh size) and allowed to grasp the grid with all four paws. The grid was inverted and the hanging time measured. Each mouse underwent three trials per test, and the longest hanging time was recorded until the mouse released its grip. Each mouse underwent three trials per session. To normalize for body weight, hanging time data were divided by the corresponding body weight (g) of each mouse, yielding values expressed as seconds per gram (s/g).

Histological analysis of the muscle

The Gast muscles of mice were collected and washed three times with phosphate-buffered saline (ServiceBio, China). Subsequently, the tissues were then fixed in 4% paraformaldehyde (ServiceBio, China) and embedded in paraffin (ServiceBio, China). The paraffin-embedded blocks were sectioned into 5 μm thick slices using a microtome. Following mounting and baking, muscle tissue sections were processed for subsequent analysis. The 5 μm tissue sections were deparaffinized and rehydrated prior to hematoxylin staining (ServiceBio, China) for 3–5 min and eosin (ServiceBio, China) for 3 min. After dehydration and clearing, the sections were mounted with resin for preservation. The slides were then visualized under a microscope (Olympus DP71, Tokyo, Japan) and digitized using CaseViewer software (3DHISTECH, Budapest, Hungary). For each sample, six fields of view were selected, and 50 muscle fibers per field were measured and analyzed for cross-sectional area, minimum Feret diameter, and mean cross-sectional area using ImageJ software (version 1.8.0; National Institutes of Health, Bethesda, MD, USA).

Transcriptome sequencing and analysis

At the experimental endpoint, gastrocnemius muscles were rapidly excised from euthanized mice, weighed, and divided into two portions. One portion was snap-frozen in liquid nitrogen for protein analysis, while the other was immersed in RNA Stabilization Solution, followed by transfer to − 80 °C until RNA extraction. Total RNA was extracted using TRIzol Reagent according to the manufacturer’s protocol, encompassing tissue homogenization, chloroform phase separation, isopropanol precipitation, and ethanol washes. RNA purity and integrity were assessed by NanoDrop 2000 spectrophotometry (A260/A280 > 1.8) and Agilent 2100 Bioanalyzer (RNA Integrity Number, RIN ≥ 7.0). Only high-quality RNA samples were used for library construction and sequencing. Transcriptomic sequencing was performed on RNA samples from 4 biological replicates per group (n = 4), with each replicate derived from an individual mouse. RNA integrity and yield were evaluated using the Agilent 2100 Bioanalyzer (Agilent Technologies, CA, USA). Samples that met quality control criteria were pooled based on effective concentration and sequencing depth requirements. Transcriptomic profiling was conducted on the Illumina HiSeqTM 2000 platform (Illumina, San Diego, CA, USA). mRNA was isolated from total RNA using Oligo dT magnetic beads, followed by library preparation, which included end repair, A-tailing, adapter ligation, fragment size selection, amplification, and purification. The libraries were quantified using a Qubit fluorometer and real-time quantitative PCR, and fragment size distribution was assessed using a bioanalyzer. FPKM values for each gene were calculated based on gene length and read counts mapped to the gene. Transcriptome profiles of different groups were compared, and p-values were calculated using the Poisson distribution. Genes exhibiting differential expression with a P < 0.05 and a false discovery rate (FDR) < 0.05 were deemed statistically significant.

C2C12 cell culture, differentiation and treatment

The mouse C2C12 myoblast cell line (CL-0044, Procell) was maintained in high-glucose DMEM (11965–092, Gibco) supplemented with 10% FBS and 1% P/S at 37 °C under 5% CO₂. Cells at 80% confluence were dissociated with 0.25% trypsin–EDTA and seeded into 6-well, 12-well, 24-well, or 96-well plates at 5 × 104 cells/cm² according to experimental protocols. Myogenic differentiation was initiated when myoblasts attained 80–90% confluency. Cells were maintained in differentiation medium (high-glucose DMEM supplemented with 4% horse serum and 1% penicillin/streptomycin) for 7 consecutive days, with complete medium replacement performed every 48 h.

In vitro myotube atrophy model and treatment groups

Differentiated myotubes were treated with 10 μM dexamethasone (DEX) for 24 h to induce atrophy [32, 35]. Co-culture experiments were performed using a 0.4 μm pore Transwell system (3470, Corning), with the following groups: Control: Myotubes cultured in standard growth medium. DEX: Myotubes treated with 10 μM dexamethasone. DEX + hUC-MSCs: 2 × 105 hUC-MSCs (passages 3–8) were seeded in the upper chamber, while DEX-treated myotubes (initial density: 5 × 10⁵ cells/cm²) were maintained in the lower chamber. DEX + MSC-Exos: 40 μg/mL MSC-Exos were added to the upper chamber, with DEX (10 μM)-treated myotubes (initial density: 5 × 10⁵ cells/cm²) in the lower chamber. All groups were incubated at 37 °C with 5% CO₂ for 24 h. For inhibitor experiments, cells were pretreated with 20 μM PD98059 (ERK1/2 inhibitor, HY-12028, MCE) or 10 μM dactolisib (PI3K inhibitor, BEZ235, HY-50673) prior to DEX exposure.

Cell proliferation

C2C12 cell proliferation was assessed using the EdU Cell Proliferation Kit (Alexa Fluor 555, C0075S Beyotime, China). Cells were seeded in the lower chamber of 24-well plates at a density of 1 × 105 cells per well, with a 0.4 μm pore Transwell insert placed in each well. The experimental groups were defined as follows: Control: Cultured in standard growth medium. DEX: Treated with 10 μM DEX for 24 h. DEX + hUC-MSCs: 4 × 104 hUC-MSCs (passages 3–8) were seeded in the upper chamber of a Transwell system, while DEX-treated C2C12 cells (initial density: 1 × 105 cells/cm²) were maintained in the lower chamber. DEX + MSC-Exos: 10 μg/mL MSC-Exos were added to the upper chamber, with DEX-treated C2C12 cells (initial density: 1 × 105 cells/cm²) in the lower chamber. All groups were incubated at 37 °C with 5% CO₂ for 24 h. Cells were then washed with PBS, fixed with 4% paraformaldehyde (ServiceBio, China) for 30 min, permeabilized with 0.1% Triton X-100 (P0096, Beyotime, China) for 20 min, and incubated with 20 μM EdU working solution for 2 h. Nuclei were counterstained with DAPI, and EdU-positive cells were visualized and quantified using an inverted fluorescence microscope (Olympus).

Immunofluorescence staining

For cytological staining, C2C12 cells from various treatment groups were fixed in 4% paraformaldehyde (ServiceBio, China) for 30 min, followed by permeabilization with 0.3% Triton X-100 (P0096, Beyotime, China) for 20 min and blocking with 3% BSA for 1 h. Subsequently, the cells were immunostained overnight at 4 °C with a specific antibody against MyHC (sc-376157, Santa Cruz). After washing, the samples were incubated with CoraLite488-conjugated Goat Anti-Mouse IgG (SA00013-1, Proteintech) at 37 °C for 1 h, followed by incubation with DAPI (C1005, Beyotime, China) at 37 °C for 10 min. Immunostained specimens were imaged using an inverted fluorescence microscope (Olympus, Tokyo, Japan) and processed for analysis with ImageJ software (version 1.51; National Institutes of Health, Bethesda, MD, USA). Five randomly selected high-power fields were used for each group to determine the number and area of MyHC + myotubes. The fusion index (%) is the number of nuclei within myotubes divided by the total number of counted nuclei.

Western blot assay

Total protein from C2C12 cells or mouse gastrocnemius muscle tissue was extracted on ice. For muscle tissue, 50 mg gastrocnemius tissue was homogenized in 1 mL ice-cold RIPA buffer (G2002-100, ServiceBio, China) using a Precellys 24 homogenizer (Bertin Instruments) at 6,500 rpm for 3 cycles (30 s each). The homogenate was centrifuged at 12,000 × g for 15 min at 4 °C, and the supernatant was collected as total protein. Protein quantification was performed using a BCA assay kit (P0010S, Beyotime, China) in accordance with the manufacturer’s guidelines. The proteins were resolved on 10% or 12% SDS-PAGE gels (G2043-50 T, ServiceBio, China) and subsequently transferred to PVDF membranes (IPFL00010, Merck, Darmstadt, Germany). The membranes were blocked with 5% milk and incubated overnight at 4 °C with the following antibodies: mouse anti-Bax (60267-1-Ig, Proteintech), rabbit anti-Bcl-2 (ab182858, Abcam), mouse anti-caspase 3 (66470-2-Ig, Proteintech), mouse anti-MuRF1 (sc-398608, Santa Cruz), mouse anti-FOXO3 (sc-48348, Santa Cruz), mouse anti-MAFbx (sc-166806, Santa Cruz), mouse anti-MYH (sc-376157, Santa Cruz), mouse anti-mTOR (sc-517464, Santa Cruz), mouse anti-Phospho-mTOR (67778-1-Ig, Proteintech), mouse anti-AKT (60203-2-Ig, Proteintech), mouse anti-Phospho-AKT (66444-1-Ig, Proteintech), rabbit anti-Phospho-PI3K (AB-2834668, Affinity), mouse anti-PI3K (60225-1-Ig, Proteintech), rabbit anti-ER α (sc-8005, Santa Cruz), rabbit anti-ER β (14007-1-AP, Proteintech), rabbit anti-TNF-α (17590-1-AP, Proteintech), rabbit anti-p62 (A7758, ABclonal), rabbit anti-Beclin (A7353, ABclonal), rabbit anti-Phospho-p44/42 MAPK (Erk1/2) (4370, Cell Signaling Technology), rabbit anti-p44/42 MAPK (Erk1/2) (4695, Cell Signaling Technology), mouse anti-GAPDH (60004-1-Ig, Proteintech), mouse anti-β Tubulin (66240-1-Ig, Proteintech).

The membranes were incubated with HRP-conjugated secondary antibodies, Goat Anti-Mouse IgG (H + L) (SA00001-1, Proteintech) and Goat Anti-Rabbit IgG (H + L) (SA00001-2, Proteintech) for 1 h at 37 °C. Protein bands were visualized using ECL reagent (G2020-50ML, ServiceBio, China) and imaged. Relative protein expression was determined by normalizing to GAPDH and quantified using ImageJ software.

RNA extraction and qRT-PCR

Total RNA was extracted from cell lysates or homogenized muscle tissue lysates using Trizol reagent (Solarbio, China) following the manufacturer’s instructions. RNA levels were quantified with a Spectramax® Quick Drop™ spectrophotometer. RNA was subsequently reverse-transcribed into cDNA, followed by qRT-PCR using 2 × SYBR Green qPCR Mix (Sparkjade Biotechnology) on the QuantStudio 5 Dx Real-Time PCR System (Thermo Fisher Scientific, Waltham, MA, USA). Relative mRNA expression levels were determined using the 2–ΔΔCt method, normalized to GAPDH as the internal control. Comparative qRT-PCR was conducted with three biological replicates. The primer sequences used in this study are provided in Table 1.

Table 1.

Primer sequence for real-time PCR

Gene	Sequence (5′to 3′)
GAPDH	Forward: AGGTCGGTGTGAACGGATTTG
	Reverse: TGTAGACCATGTAGTTGAGGTCA
TNF-α	Forward: CCTGTAGCCCACGTCGTAG
	Reverse: GGGAGTAGACAAGGTACAACCC
IL-1β	Forward: TTCAGGCAGGCAGTATCACTC
	Reverse: GAAGGTCCACGGGAAAGACAC
IL-6	Forward: CTGCAAGAGACTTCCATCCAG
	Reverse: AGTGGTATAGACAGGTCTGTTGG
MyHC	Forward: GCGAATCGAGGCTCAGAACAA
	Reverse: GTAGTTCCGCCTTCGGTCTTG
FOXO3	Forward: CTGGGGGAACCTGTCCTATG
	Reverse: TCATTCTGAACGCGCATGAAG
MAFbx	Forward: ATCCCTGAGTGGCATCGC
	Reverse: CTCTTCCACAGTAGCCGGT
MuRF-1	Forward: GTGTGAGGTGCCTACTTGCTC
	Reverse: GCTCAGTCTTCTGTCCTTGGA

Enzyme-linked immunosorbent assay (ELISA)

For mouse serum collection, blood was obtained via retro—orbital plexus puncture, allowed to clot for 2 h, and then centrifuged at 3000 × g for 15 min at 4 °C. The resulting supernatant was stored at − 80 °C. For cell supernatant collection, C2C12 cells were cultured in 6-well Transwell plates. After being treated with 10 μM dexamethasone, 2 × 105 hUC-MSCs, or 40 μg/mL MSC-Exos for 24 h, the cell culture supernatants were collected and centrifuged at 1000 × g for 10 min to remove debris. The levels of estradiol (E2), TNF-α, IL-6, and IL-1β were quantified using murine ELISA kits (CSB-E05101m for E2 from Cusabio; CSB-E04741m, CSB-E04639m, CSB-E08054m for TNF-α, IL-6, and IL-1β from Jinmei) according to the manufacturer’s instructions. The optical density (OD) was measured at 450 nm using a microplate reader.

Flow cytometric analysis of apoptosis

Apoptosis levels were assessed using an Annexin V-FITC/PI Apoptosis Detection Kit (C1062L, Beyotime, China) as follows: cells were collected by trypsinization without EDTA and resuspended in binding buffer to prepare single-cell suspensions. A mixture of 195 μL Annexin V-FITC and 10 μL propidium iodide (PI) staining solution was gently mixed with the cell suspension and incubated at room temperature in the dark for 20 min. Flow Cytometry: Stained cells were analyzed using a BD FACSAria™ III flow cytometer (BD Biosciences). Data Analysis: The percentage of apoptotic cells was quantified using FlowJo software (v10.8.1).

Statistical analysis

Statistical analyses were performed using GraphPad Prism version 5.0 (GraphPad Software Inc., La Jolla, CA, USA). Data in this study were expressed as mean ± standard error (SEM). All data were obtained from at least three independent experiments. The variable (n) represents the quantity of distinct biological replicates, depicted in the graphs as a single data point for each sample. To analyze differences among multiple groups, ANOVA was conducted, followed by Tukey’s post hoc analysis. For pairwise group comparisons, independent samples t-tests were utilized. P < 0.05. was considered statistically significant.

Results

hUC-MSCs protect mice against DEX-induced muscle atrophy

hUC-MSCs at passages 3–8 were utilized for the experiments. The characterization of hUC-MSCs and MSC-Exos adhered to the standard criteria (Fig. S1A). We established a DEX-induced muscle atrophy mouse model to investigate the therapeutic potential of hUC-MSCs in Alleviating muscle atrophy. Preintervention body weight, forelimb grip strength, and exercise endurance were measured prior to treatment initiation. Longitudinal monitoring was performed at 24-h intervals for body weight and weekly intervals for functional assessments. No significant intergroup differences were observed in pretreatment parameters across cohorts.

DEX significantly decreased body weight compared with the Control group (P < 0.001, Fig. 1A). Although the Dex + MSC group showed a weight recovery trend compared with the Dex group, the difference did not achieve statistical significance, possibly due to the limited sample size. These results suggest hUC-MSCs transplantation may alleviate Dex-induced body weight loss. However, hUC-MSCs significantly increased the ratio of Gast muscle weight to body weight (P < 0.05 vs DEX, Fig. 1B). Functional assessments revealed significant improvements in grip strength (Fig. 1C) and endurance (Fig. 1D) compared to the DEX group. Histological analysis via HE staining revealed that the DEX group presented disrupted muscle fiber organization, extensive atrophy, and reduced cellular volume compared to controls (Fig. 1E). hUC-MSCs alleviated these histological changes by restoring muscle fiber cross-sectional area (CSA) and minimizing Feret’s diameter, indicating enhanced muscle fiber regeneration (Fig. 1F).

Fig. 1.

hUC-MSCs protect mice against DEX-induced muscle atrophy. A Final body weight of mice at the conclusion of the study (n = 6); B Gast muscle weight in body weight at the end of the experiment (n = 6); C Standardized grip strength (N/g) at the end of the experiment (n = 6); D Standardized endurance time (s/g) at the end of the experiment (n = 6); E HE-stained sections of skeletal muscle tissue from each group of mice (n = 6) Scale bar: 50 μm; F Percent of fibers (%) and Min feret’s Diameter. G Heatmap of DEGs in different samples. H The volcano map of DEGs in different samples. I Top enriched gene ontology terms: biological process (BP), cellular component (CC), and molecular function (MF) for DEGs. J Top 10 KEGG enrichment analysis of DEGs in different samples. DEX group: n = 4. MSC group: n = 4. Data from three independent experiments are reported as the mean ± SEM. ***P < 0.001 vs. Control. # P < 0.05, ## P < 0.01, ### P < 0.001 vs. DEX

To elucidate the molecular mechanisms driving DEX-induced muscle atrophy and the cellular signaling pathways influenced by hUC-MSCs transplantation, RNA-seq transcriptomic analysis was conducted on Gast tissues from the DEX and DEX + MSC groups. Differentially expressed genes (DEGs) were identified using stringent criteria (adjusted P < 0.05, |log FC|> 1). A total of 3,213 differentially expressed genes (DEGs) were identified, including 2,734 upregulated and 479 downregulated genes in the DEX + MSC group compared with the DEX group (Fig. 1G–H). The functional annotation of these DEGs was conducted using GO and KEGG pathway enrichment analyses (Fig. 1I–J). The results indicated that hUC-MSCs transplantation modulated key pathways, including the PI3K-Akt signaling axis and cytokine-cytokine receptor interactions. These pathways are associated with increased stress responses, immune activation, cell proliferation, cytoskeletal remodeling, and extracellular matrix reconstruction. Extensive research has shown that MSC-derived exosomes are enriched with nucleic acids, proteins, and a diverse array of bioactive molecules, including growth factors, cytokines, and antioxidants[20, 36]. Interestingly, the therapeutic benefits of stem cells are increasingly associated with their paracrine mechanisms rather than the direct incorporation of transplanted cells into the host tissues [37]. On the basis of this evidence, we hypothesize that MSCs, through exosome-mediated delivery of these bioactive components, MSCs orchestrate the modulation of key signaling pathways. This process likely regulates immune responses, suppresses inflammation, promotes cellular proliferation, and drives extracellular matrix remodeling, thereby mitigating DEX-induced muscle damage.

MSC-Exos protect mice against DEX-induced muscle atrophy

To clarify the role of MSC-Exos in mitigating DEX-induced muscle atrophy, MSC-Exos (100 μg) or PBS were administered via daily bilateral intramuscular injections into the gastrocnemius muscle of atrophic mice for 14 consecutive days. No significant intergroup differences were observed in pretreatment parameters across cohorts. Notably, MSC-Exos exhibited therapeutic efficacy comparable to that of hUC-MSCs. Although MSC-Exos treatment did not significantly affect overall body weight (Fig. 2A), it significantly increased the ratio of Gast muscle weight to body weight (Fig. 2B), improved grip strength (Fig. 2C), and bolstered endurance (Fig. 2D). Furthermore, MSC-Exos alleviated DEX-induced muscle atrophy (Fig. 2E), as evidenced by increased muscle fiber diameter and improved structural organization (Fig. 2F).

Fig. 2.

MSC-Exos protected mice against DEX-induced muscle atrophy. A Final body weight of mice at the conclusion of the study (n = 6); B Gast muscle weight in body weight at the end of the experiment (n = 6); C Grip strength of mice at the end of the experiment (n = 6); D Endurance test of mice at the end of the experiment (n = 6); E HE-stained sections of skeletal muscle tissue from each group of mice (n = 6) Scale bar: 50 μm; F Percent of fibers (%) and min feret’s diameter; G The volcano map of DEGs in DEX and MSC-Exos groups; H Heatmap of DEGs in DEX and MSC-Exos groups; I. Top enriched gene ontology terms: biological process (BP), cellular component (CC), and molecular function (MF) for DEGs; J Top 10 KEGG enrichment analysis of DEGs in DEX and MSC-Exos groups. DEX group: n = 4. MSC-Exos group: n = 4. *** P < 0.001 vs. Control. # P < 0.05, ## P < 0.01, ### P < 0.001 vs. DEX

To further investigate the mechanisms by which MSC-Exos alleviating muscle atrophy, transcriptomic analysis was conducted on Gast tissues from atrophic mice treated with either PBS or MSC-Exos. A total of 3,447 differentially expressed genes (DEGs) were identified, including 3,314 upregulated and 533 downregulated genes in the DEX + MSC-Exos group compared with the DEX group (Fig. 2G–H). GO enrichment (Fig. 2I) and KEGG pathway analyses (Fig. 2J) of these DEGs revealed that MSC-Exos exerts protective effects on muscle tissue by modulating immune responses, suppressing inflammation, and stabilizing key cellular structures, including the cytoskeleton and extracellular matrix. Previous studies highlighted the critical function of estrogen in promoting skeletal muscle repair and modulating inflammation [38, 39], which aligns with our findings. Notably, the PI3K-Akt and MAPK pathways—both closely associated with estrogen signaling—were significantly enriched. These pathways serve as downstream mediators of estrogen, driving cell survival, proliferation, and anti-apoptotic mechanisms. Our results suggest that MSC-Exos counteracts DEX-induced muscle atrophy by activating estrogen signaling, which engages PI3K and MAPK pathways to preserve skeletal muscle integrity.

hUC-MSCs and MSC-Exos alleviate protein degradation, autophagy and apoptosis in DEX-induced skeletal muscle atrophy

The activation of protein degradation pathways, driven by chronic inflammation or excessive glucocorticoid secretion, is a central mechanism underlying severe skeletal muscle wasting [40, 41]. This process involves FOXO dependent transcription and activation of the ubiquitin–proteasome system, characterized by elevated levels of FOXO3, MuRF-1, and MAFbx, coupled with reduced expression of myogenesis markers such as the MyHC [42]. In our study, Western blot analysis revealed a significant decrease in MyHC levels in the DEX group compared with the control group (Fig. 3B). Treatment with MSCs and MSC-Exos partially restored MyHC expression, with MSC-Exos demonstrating superior efficacy. Furthermore, DEX treatment substantially upregulated FOXO3, MuRF-1, and MAFbx (Fig. 3A–B), In contrast, both MSCS and MSC-Exos significantly suppressed their expression, with MSCs showing a more pronounced effect. These results indicate that MSCs and MSC-Exos treatments mitigate muscle atrophy primarily by inhibiting the ubiquitin–proteasome pathway.

Fig. 3.

hUC-MSCs and MSC-Exos alleviate protein degradation, autophagy and apoptosis in DEX-induced skeletal muscle atrophy. A Representative immunoblot analyses of MyHC, FOXO3, MAFbX, MURF, and GAPDH were performed in GA tissue, with the data illustrating protein expression levels. GAPDH served as the normalization reference for protein loading. B Quantitative analysis of MyHC、FOXO3, MAFbX, MURF expression levels among the groups. C Representative Western blot images of Beclin 1, P62 and GAPDH were analyzed in GA. D Quantitative analysis of Beclin 1, P62 expression levels among the groups. The data from three separate experiments are expressed as the mean ± SEM. E Apoptosis in muscle tissue was evaluated through TUNEL staining (red), with nuclei visualized using DAPI counterstaining (blue). Scale bar: 50 μm. F Apoptosis percentage in Gast in each group. G Representative immunoblot data for Bcl-2, Bax, Caspase-3, and GAPDH were examined in GA, with GAPDH serving as the reference protein for loading normalization. H Quantitative analysis of Bcl-2/Bax, Caspase 3 expression levels among the groups. *** P < 0.001 vs. Control. # P < 0.05, ## P < 0.01, ### P < 0.001 vs. DEX. & P < 0.05, &&& P < 0.001 vs. MSC. The complete, uncropped versions of the full-length gels and blots are provided in the supplementary materials

The pathophysiology of sarcopenia is characterized by disrupted autophagic regulation and excessive apoptotic activation, highlighting these processes as promising therapeutic targets for managing this condition [43]. The key markers of autophagy, Beclin1 and P62, play crucial roles: Specifically, Beclin1 promotes autophagy initiation, whereas P62 is degraded during the autophagic process. In DEX-induced muscle atrophy, Beclin1 expression was significantly reduced, whereas P62 levels were markedly elevated (Fig. 3C–D), indicating impaired autophagic activity. Treatments with MSCs and MSC-Exos treatments restored Beclin1 expression and decreased P62 accumulation, with MSCs displaying a marginally superior effect. These findings suggest that both interventions enhance the clearance of damaged proteins and cellular debris by reactivating autophagy, thereby alleviating muscle atrophy.

Apoptosis levels in muscle tissue were assessed using TUNEL staining. As shown in Fig. 3E, the number of TUNEL-positive cells was significantly greater in the DEX group than in the control group, reflecting increased levels of apoptosis. The administration of hUC-MSCs or MSC-Exos markedly decreased the number of TUNEL-positive cells. DEX treatment increased Caspase-3 expression, suppressed antiapoptotic Bcl-2, and upregulated proapoptotic Bax, reducing the Bcl-2/Bax ratio (Fig. 3G–H). hUC-MSCs and MSC-Exos normalized the Bcl-2/Bax ratio, suppressed Caspase-3, and mitigated apoptosis, with hUC-MSCs showing slightly better efficacy. These findings underscore that DEX treatment induces significant apoptosis in muscle tissue, whereas both MSC-based interventions effectively counteract this process.

hUC-MSCs and MSC-Exos alleviate DEX-induced inflammation both in vivo and in vitro

Pro-inflammatory cytokines (TNF-α, IL-1β, and IL-6) play critical roles in sarcopenia and muscle atrophy [44, 45]. In vivo, HE staining of muscle tissue revealed DEX disrupted fiber architecture in atrophic mice—marked by increased atrophic fibers, irregular cellular morphology, and inflammatory cell infiltration (Fig. 4A). In contrast, hUC-MSCs or MSC-Exos treatment improved muscle tissue: reduced inflammatory cell infiltration, fewer atrophic fibers, and better-preserved structural integrity. The MSC-Exos group showed more regular fiber shapes and less fragmentation than the MSC group (Fig. 4A). Western blot analysis was performed to detect TNF-α protein expression in muscle tissue homogenates (Fig. 4B–C). DEX markedly increased TNF-α expression (P < 0.001), whereas hUC-MSCs or MSC-Exos treatment partially reversed this effect (P < 0.05). Real-time qPCR showed DEX upregulated mRNA levels of TNF-α, IL-6, and IL-1β. Both hUC-MSCs and MSC-Exos treatments reduced these pro-inflammatory markers, with hUC-MSCs exhibiting slightly stronger anti-inflammatory efficacy (Fig. 4F). ELISA was used to quantify TNF-α, IL-6, and IL-1β levels in mouse serum. Results: DEX significantly increased serum inflammatory cytokine levels compared to the control group (P < 0.001), while both hUC-MSCs and MSC-Exos treatment significantly suppressed their secretion (P < 0.001, Fig. 4G).

Fig. 4.

hUC-MSCs and MSC-Exos alleviate DEX-induced inflammation both in vivo and in vitro. A HE-stained sections of skeletal muscle tissue from each group of mice, with red circles indicating the locations of inflammatory cell infiltration; scale bar: 50 μm; n = 6. B Immunoblot analysis of TNF-α and GAPDH was conducted in GA, with representative results presented. C Quantitative analysis of TNF-α expression levels among the groups. D Representative immunoblot results of TNF-α and GAPDH were examined in C2C12 myotubes, with GAPDH serving as the internal loading reference. E Quantitative analysis of TNF-α expression levels among the groups. F Relative mRNA levels of TNF-α, IL-6, IL-1β mRNA in skeletal muscle was quantified relative to GAPDH, which was used as the internal control for normalization. F and H. Expression levels of the cytokines TNF-α, IL-6 and IL-1β in C2C12 cell serum and supernatants were detected by ELISA. All experiments were performed at least three times. *** P < 0.001 vs. Control. # P < 0.05, ### P < 0.001 vs. DEX. && P < 0.01, &&& P < 0.001 vs. MSC

In vitro, C2C12 myotubes treated with DEX (10 μM) were co-cultured with hUC-MSCs or MSC-Exos. Western blot analysis indicated DEX elevated TNF-α in C2C12 cells, an effect reversed by both MSC and MSC-Exos treatments (Fig. 4D–E). High-sensitivity ELISA kits were used to quantify TNF-α, IL-6, and IL-1β concentrations in the supernatants of C2C12 myotube cultures (Fig. 4H). DEX significantly promoted the secretion of inflammatory cytokines (P < 0.001), while hUC-MSCs/MSC-Exos co-culture significantly inhibited their release (Fig. 4H).

hUC-MSCs and MSC-Exos promote the C2C12 myotube proliferation and differentiation

To further confirm the therapeutic mechanisms a DEX-induced myotube atrophy in vitro model was employed. EdU assay (Fig. 5A–B) revealed DEX significantly inhibited C2C12 proliferation (reduced red fluorescence vs. control). Both DEX + MSC and DEX + MSC-Exos groups restored proliferation, with the DEX + MSC-Exos group showing a higher percentage of EdU-positive cells than DEX + MSC, suggesting that exosome treatment has a more pronounced proliferative effect on proliferation than does treatment with MSCs alone.

Fig. 5.

hUC-MSCs and MSC-Exos promote the C2C12 myotube proliferation and differentiation A and B Representative visuals from the EdU assay for cell proliferation are provided, with a scale bar indicating 50 μm. C Representative immunofluorescence staining visuals for MyHC are presented. Scale bar: 20 μm. D Quantitative analysis of Myotube diameters. E Quantitative analysis of Fusion index. The cells were incubated with EDU or antibody against MyHC for 24 h. F Representative immunoblot results for MyHC, FOXO3, MAFbX, MURF, and GAPDH in C2C12 myotubes are shown. G Quantitative analysis of MyHC, FOXO3, MAFbX, MURF expression levels among the groups. H. Relative mRNA levels of MyHC, FOXO3, MAFbX, MURF. ** P < 0.01, *** P < 0.001 vs. Control. # P < 0.05, ## P < 0.01, ### P < 0.001 vs. DEX. & P < 0.05, && P < 0.01, &&& P < 0.001 vs MSC

MyHC, a key marker of myotubes differentiation, is widely used to evaluate muscle cell differentiation. As shown in Fig. 5C, DEX treatment decreased myotube diameter (Fig. 6C–D) and fusion index (Fig. 6E), impairing differentiation. hUC-MSCs and MSC-Exos alleviated these effects, with MSC-Exos more effectively restoring myotube integrity and promoting differentiation. Treatment with hUC-MSCs and MSC-Exos partially reversed these detrimental effects, with MSC-Exos demonstrating superior efficacy over MSCs in restoring myotube integrity and promoting differentiation. DEX treatment significantly reduced MyHC protein and mRNA levels, while upregulating atrophy-associated proteins and mRNAs, including FOXO3, MAFbx, and MURF1. In contrast, both hUC-MSCs and MSC-Exos treatments restored MyHC expression and suppressed FOXO3, MAFbx, and MURF1 at both protein and mRNA levels. Notably, the DEX + MSC-Exos group showed effective MyHC restoration, while hUC-MSCs demonstrated stronger suppression of the ubiquitin–proteasome system (via FOXO3, MAFbx, MURF1 regulation), emphasizing their efficacy in mitigating DEX-induced muscle degradation (Fig. 7F–H). The qPCR results corroborated Western blot findings, showing consistent transcriptional changes across treatment groups.

Fig. 6.

hUC-MSCs and MSC-Exos regulate C2C12 myotube atrophy and apoptosis in DEX-treated C2C12 myotubes. A and B Representative immunofluorescence staining images of MDC are displayed. The scale bar represents 20 μM. C Representative immunoblot analyses of P62, Beclin 1, and GAPDH in C2C12 myotubes are provided. D Quantitative analysis of Beclin 1 and P62 expression levels among the groups. E Representative immunoblot results for Bcl-2, Bax, Caspase-3, and GAPDH in C2C12 myotubes are shown. F Quantitative analysis of Bcl-2/Bax and Caspase 3/GAPDH expression levels among the groups. G Apoptotic effect of DEX on C2C12 Myotubes. H The proportion of apoptotic cells was evaluated using flow cytometry with dual staining of FITC and PI. * P < 0.05. ** P < 0.01, *** P < 0.001 vs. Control. # P < 0.05, ###P < 0.001 vs. DEX. & P < 0.05, && P < 0.01 vs. MSC

Fig. 7.

hUC-MSCs and MSC-Exos modulate estrogen receptors and PI3K/AKT/mTOR signaling in DEX-treated muscle and C2C12 myotubes: A and C The expression levels of ERα, ERα36, and ERβ proteins were assessed via Western blotting. B and D Quantitative analysis of ERα, ERα36, and ERβ expression levels among the groups. E Expression levels of the E2 in serum were detected by ELISA. F The protein expressions of p-PI3K, PI3K, p-AKT, AKT, p-mTOR and mTOR. GAPDH was used as the loading control. G p-PI3K/PI3K, p-AKT/AKT and p-mTOR/mTOR expression levels among the groups. * P < 0.05, ** P < 0.01, *** P < 0.001 vs. Control. # P < 0.05, ## P < 0.01, ### P < 0.001 vs. DEX. & P < 0.05, && P < 0.01, &&& P < 0.001 vs. MSC. $ P < 0.05, $$$ P < 0.05 vs. MSC-Exos

hUC-MSCs and MSC-Exos regulate C2C12 myotube autophagy and apoptosis in DEX-treated C2C12 myotubes

Autophagy assessment via MDC staining (Fig. 6A–B) showed DEX suppressed autophagosome formation, while hUC-MSCs and MSC-Exos partially restored autophagic activity, with hUC-MSCs demonstrating a stronger effect. Western blot analysis (Fig. 6C–D) validated this, showing increased Beclin 1 (autophagy promoter) and reduced p62 (autophagy substrate) levels after hUC-MSCs and MSC-Exos treatment. For apoptosis, Western blot analysis (Fig. 6E–F) revealed hUC-MSCs and MSC-Exos counteracted DEX-induced apoptosis by restoring the anti-apoptotic Bcl-2/Bax ratio and decreasing pro-apoptotic Caspase-3 expression. Flow cytometry with Annexin V/PI staining (Fig. 6G–H) further confirmed both treatments reduced the proportion of apoptotic cells in DEX-treated groups. Collectively, hUC-MSCs and MSC-Exos modulate autophagy and alleviate apoptosis in DEX-treated C2C12 myotubes.

hUC-MSCs and MSC-Exos modulate estrogen receptors and PI3K/AKT/mTOR signaling in DEX-treated muscle and C2C12 myotubes

Estrogen mediates effects via estrogen receptors (ERs) through signal transduction. Western blot analysis revealed that DEX significantly downregulated ERα, ERα36, and ERβ expression in the gastrocnemius muscle (P < 0.001). However, hUC-MSCs and MSC-Exos partially restored their expression levels (Fig. 7A–B). ELISA assays demonstrated that DEX treatment reduced serum estradiol (E2) levels by 42% (P < 0.001, Fig. 7E), whereas hUC-MSCs and Exos treatment significantly restored E2 levels (P < 0.01, Fig. 7E). These findings emphasize the critical involvement of estrogen signaling pathways and the synergistic interplay among various ER subtypes in the muscle-protective effects of MSCs and MSC-Exos.

The PI3K/AKT/mTOR and ERK1/2 signaling pathways are pivotal in regulating cellular proliferation, differentiation, and survival. Moreover, these pathways are key mediators of rapid, nongenomic estrogen activation [42, 46–48]. hUC-MSCs and MSC-Exos activated the ERK1/2 and PI3K/AKT/mTOR pathways. However, this effect was completely abolished by ERK1/2 inhibitor (PD98059, 20 μM) (Fig. 8A–B) and PI3K inhibitor (BEZ235, 10 μM) (Fig. 7F–G). These results indicate that estrogen signaling mediates muscle protection through these two pathways. The additional data establish a complete mechanistic framework linking estrogen levels, ER expression, and downstream pathway activation, providing a theoretical basis for future clinical targeted therapies.

Fig. 8.

Effects of hUC-MSCs and MSC-Exos on PI3K/AKT/mTOR AND ERK1/2 pathways in DEX-induced myotube atrophy. A p-ERK and ERK protein expression levels in DEX-treated C2C12 myotubes following administration of ERK1/2 inhibitors. B Quantitative analysis of p-ERK/ERK expression levels among the groups. C Final body weight of mice at the conclusion of the study (n = 6); D Gast muscle weight in body weight at the end of the experiment (n = 6); E Grip strength of mice at the end of the experiment (n = 6); F Endurance test of mice at the end of the experiment (n = 6); G HE-stained sections of skeletal muscle tissue from each group of mice (n = 6), Scale bar: 50 μm; H. Percent of fibers (%) and Min feret’s Diameter. These results are presented as mean ± SEM of three independent experiments: *** P < 0.001 vs. Control. # P < 0.05, ## P < 0.01, ### P < 0.001 vs. DEX. && P < 0.01 vs. MSC. ^^ P < 0.01 vs. MSC-Exos

SNG162 protect mice against DEX-induced muscle atrophy

To demonstrate the critical role of estrogen, we used the estrogen analog SNG162 for intervention. Treating DEX-treated mice with SNG162 (40 mg/kg/day for 2 weeks) replicated the therapeutic effects of hUC-MSCs/MSC-Exos, improving muscle atrophy and functional outcomes (Fig. 8C–H). Specifically, SNG162 treatment significantly enhanced body weight, grip strength, endurance, and muscle fiber cross-sectional area (CSA), with effects comparable to those of the hUC-MSCs/Exos treatment group (Fig. 8C–H). This directly confirms that estrogen signaling activation serves as a core mechanism in muscle repair.

Discussion

Estrogen maintains skeletal muscle homeostasis through multiple mechanisms, including promoting of muscle fiber hypertrophy, modulating of inflammatory responses, regulating the balance between protein synthesis and degradation balance, and improving of glucose and lipid metabolic disorder [49]. Despite lower circulating estrogen levels in males than in females, estrogen receptors (ERα/ERβ) remain highly expressed and functionally active in male skeletal muscle [50]. Recent studies have shown that 17-α estradiol significantly ameliorates age-related sarcopenia in male mice and enhances physical function in late life [51]. Furthermore, 17-α estradiol regulates systemic metabolic environments and improves the muscle microenvironment, thereby delaying the aging process [52]. These findings challenge traditional views, indicating that the protective role of estrogen in aging extends beyond females to males, playing a critical role in muscle health and the regulation of the aging process.

The DEX-induced muscle atrophy model effectively simulates aspects of sarcopenia by promoting muscle protein degradation via the ubiquitin–proteasome pathway [32, 53, 54]. FOXOs, particularly FOXO3, were crucial in regulating skeletal muscle protein degradation by modulating the ubiquitin–proteasome [55–57] and autophagy-lysosome pathways, participating in regulating the cell cycle, apoptosis, and muscle atrophy and regeneration processes [58]. DEX induces muscle atrophy and cell cycle arrest by activating the ubiquitin–proteasome pathway, thereby impairing cell proliferation, a hallmark of DEX-induced muscle atrophy [59]. In our study, DEX-treated mice demonstrated marked reductions in body weight, grip strength, endurance, and Gast muscle-to-body weight ratio, accompanied by C2C12 myotube atrophy and ubiquitin–proteasome system activation, aligning with the hallmark characteristics of the DEX-induced sarcopenia model described in previous research [32, 55]. While preclinical studies demonstrate that the dexamethasone induced sarcopenia model replicate key phenotypic hallmarks features of primary sarcopenia, it is critical to acknowledge that translational constraints exist due to divergent pathophysiological mechanisms, particularly in geriatric populations.

Estrogen, a key hormone with broad biological effects, regulates oxidative stress, inflammation, apoptosis, and tissue repair via classical nuclear receptors (ERα/ERβ) and non-classical membrane receptors (ERα36, GPER1), activating downstream PI3K/Akt and ERK1/2 pathways [60–62]. hUC-MSCs and their derives exosomes demonstrate remarkable potential in regulating hormone-related signaling pathways, underpinning therapeutic advancements across various diseases. For instance, secretomes from hUC-MSCs precisely target hormone balance and circadian rhythm, fostering ovarian health [63]. MSC-derived EVs restore ovarian function by enhancing follicular development and increasing estradiol (E2) levels [64]. Furthermore, these EVs mitigate chemotherapy-induced ovarian toxicity through PI3K/AKT/mTOR pathway activation [65]. Our study is the first to demonstrate that hUC-MSCs and MSC-Exos transplantation can increase serum estradiol levels and activate estrogen receptor (ERα, ERβ, ERα36)-dependent signals and downstream PI3K/Akt/mTOR and ERK1/2 pathways. In addition to inhibiting the ubiquitin–proteasome system, this study emphasizes for the first time that MSCs and MSC-Exos can regulate autophagy, apoptosis, and inflammation, providing a comprehensive intervention framework for the multi-factor pathogenesis of sarcopenia. This multi-target strategy significantly transcends single-pathway studies. Moreover, we directly verified the core role of estrogen in muscle homeostasis by treating with exogenous estradiol SNG162, which significantly improved DEX-induced muscle atrophy. Our findings fill the gap in the existing MSCs research regarding the hormone regulation network. This study utilized male mice to minimize the confounding effects of endogenous estrogen fluctuations. However, future research should incorporate female mice and ovariectomized models to comprehensively evaluate the sex-specific effects of hUC-MSCs and exosomes.

Stem cell therapy has shown promise in treating various degenerative diseases, including ischemic stroke [66] and multiple sclerosis [67, 68]. However, large-scale clinical trials investigating the application of hUC-MSCs in treating sarcopenia remain absent to date. Our team conducted the first clinical trial in China (NCT03005249) evaluating the safety and efficacy of intranasal delivery of NSC-loaded composite patches for pediatric cerebral palsy, utilizing a novel delivery system to overcome the limitations of NSC clinical applications [69]. In the past years, we demonstrated that hUC-MSCs could promote neuronal regeneration, modulate the immune microenvironment, and aid in the repair of neonatal hypoxic-ischemic brain injury [29]. Building upon this foundation, we further investigated the therapeutic potential of hUC-MSCs in managing muscle atrophy. In our investigation, hUC-MSCs effectively attenuated DEX-induced skeletal muscle wasting, aligning with prior findings [16, 17, 70]. Exosomes offer a cell-free therapeutic alternative, mitigating risks associated with MSCs transplantation, such as immunogenicity and tumorigenesis. Their ability to efficiently deliver bioactive molecules enhances their therapeutic potential [31]. Compared with MSCs, MSC-Exos exhibit superior penetration of cellular barriers, facilitating more precise delivery of effector substances. Previous studies have suggested that exosomes directly modulate proliferative signaling pathways through their cargo of growth factors (IGF-1, VEGF), proteins, and small RNA [71–73].This preclinical study demonstrated the therapeutic potential of hUC-MSCs and MSC-Exos in alleviating skeletal muscle atrophy, restoring physical function, reducing protein degradation, inhibiting apoptosis, mitigating inflammation, enhancing autophagy, and promoting estradiol receptor expression. These findings support hUC-MSCs and MSC-Exos as potential therapies for muscle atrophy, emphasizing their respective advantages in sarcopenia treatment. MSC-Exos ability to promote cell proliferation and differentiation, due to enhanced uptake and targeted delivery, positions it as an early intervention for muscle regeneration. In contrast, hUC-MSCs, which improve overall muscle health, may be more suitable for mid-to-late-stage sarcopenia to slow progression. However, the DEX-induced model may not fully reflect age-related sarcopenia, warranting the use of aging models for future validation. This study advances regenerative medicine by confirming the roles of hUC-MSCs and MSC-Exos in improving the muscle microenvironment, promoting muscle repair, and encouraging further exploration of these biomaterials. Additionally, future research could examine combining hUC-MSCs and MSC-Exos with pharmacologic or rehabilitation therapies to enhance sarcopenia symptom management and quality of life.

In the rapidly evolving field of regenerative medicine, hUC-MSCs and MSC-Exos have gained significant attention due to their multifunctionality and low immunogenicity. However, translating these therapies into clinical practice presents challenges, including the standardization of manufacturing processes, optimization of delivery methods, and long-term safety assessment. The inherent heterogeneity of MSCs and MSC-Exos hinders consistency and reproducibility, with variability arising from donor-specific factors, culture conditions, and isolation protocols, all affecting potency, purity, and safety. Systemic administration of exosomes or stem cells encounters challenges such as rapid clearance and non-specific distribution, particularly in complex in vivo environments characterized by high blood flow or multi-organ dispersion. While localized injections improve targeting, their efficacy is often limited by restricted diffusion. Further investigation is needed to assess risks, including immune activation, tumorigenesis, and off-target effects. To realize the full clinical potential of hUC-MSCs and MSC-Exos, collaboration among researchers, industry leaders, and regulatory authorities is essential. Multidisciplinary approaches, including synthetic biology, nanomedicine, and systems biology, could enhance the safety and efficacy of these therapies. Additionally, big data analytics and precision medicine will play a key role in refining individualized treatment strategies.

Conclusion

This study transcends conventional single-target paradigms by establishing a "cell-exosome-endocrine" triumvirate precision intervention system for sarcopenia at the systems biology level. Leveraging synergistic effects of clinical-grade hUC-MSCs and their exosomes, we pioneer multidimensional regulation spanning molecular, cellular, and systemic hierarchies, offering a translatable solution for estrogen-deficient sarcopenia with mechanistic explicability, therapeutic quantifiability, and manufacturing standardization. This paradigm not only redefines the application boundaries of stem cell therapy in endocrine-muscle axis disorders but also establishes a molecular roadmap for pathology stage-specific personalized treatment, heralding a transformative shift from symptomatic management to etiological correction in sarcopenia therapeutics.

Abbreviations

hUC-MSCs

Human umbilical cord-derived mesenchymal stem cells

MSC-Exos

Mesenchymal stem cell-derived exosomes

DEX

Dexamethasone

ERα

Estrogen receptor α

ERβ

Estrogen receptor β

FBS

Fetal bovine serum

FDR

False discovery rates

DMEM

Dulbecco’s modified eagle’s medium

DMSO

Dimethyl sulfoxide

EdU

5-Ethynyl-2’-deoxyuridine

TEM

Transmission electron microscopy

NTA

Nanoparticle tracking analysis

MyHC

Myosin heavy chains

RIPA

Radio immunoprecipitation assay

BCA

Bicinchoninic acid

SDS-PAGE

Sodium dodecyl sulfate polyacrylamide gel electrophoresis

PVDF

Polyvinylidene fluoride

Bax

Bcl-2-associated X protein

Bcl-2

B-cell lymphoma 2

Caspase 3

Cysteine-aspartic acid protease 3

MuRF1

Muscle RING-finger protein-1

FOXO3

Forkhead box O3

MAFbx

Muscle atrophy F-box

mTOR

Mechanistic target of rapamycin

AKT

Protein kinase B

PI3K

Phosphoinositide 3-kinase

ER

Estrogen receptor

P62

Sequestosome 1(SQSTM1)

TNF-α

Tumor necrosis factor α

MAPK

Mitogen-activated protein kinase

GAPDH

Glyceraldehyde-3-phosphate dehydrogenase

HRP

Horseradish peroxidase

ECL

Erythrina cristagalli lectin

qRT-PCR

Quantitative real-time polymerase chain reaction‌

IL-6

Interleukin-6

IL-1β

Interleukin-1β

FITC

Fluorescein isothiocyanate

EDTA

Ethylene diamine tetraacetic aci

TUNEL

Terminal deoxynucleotidyl transferase dUTP nick-end labeling

Author contributions

NL, CH, CQ, WZ, JL, and YW conceived and designed the study. NL, XL, QW, XG, XW, AL, YZ, and RD performed the experiments. NL, XL, QW, and YC performed the data analysis. NL and YC prepared the manuscript. JL and YW supervised the project. All authors contributed to the discussion and revision of the manuscript.

Funding

The present research was funded by Liaoning Province Key Science and Technology Project (JB; Grant No.2022JH1/10800070) and Dalian high level talent innovation support plan (HC; Grant No.2021RQ028).

Availability of data and materials

The datasets utilized and/or analyzed in this study can be obtained from the corresponding author upon reasonable request. All additional files are included in the manuscript.

Declarations

Ethical approval and consent to participate

This study was performed at the Stem Cell Clinical Research Center of the First Affiliated Hospital of Dalian Medical University. This study was approved by the Ethics Committee of the First Affiliated Hospital of Dalian Medical University (Project Title: Research on the Treatment of Sarcopenia Using Human Umbilical Cord Mesenchymal Stem Cells Combined with Phytomedicine; Approval No. YJ-GZR-SQ-2024–69; Approved on March 5, 2024). The ethical approval covered the collection of umbilical cord specimens, which were obtained from healthy fetuses delivered by cesarean section. Informed consent was provided by all pregnant women participants and their family members prior to specimen collection. All animal experiments were performed in strict accordance with the ethical guidelines established by the Animal Research Ethics Committee of Dalian Medical University (Project Title: Research on the Treatment of Sarcopenia with Human Umbilical Cord Mesenchymal Stem Cells Combined with Epimedium; Approval No. AEE24221; Approved on January 9, 2024).

Consent for publication

All authors reviewed and endorsed the final version of the manuscript for submission and publication.

Artificial intelligence (AI)

The authors declare that no AI-generated content was used in this manuscript.

Competing interests

The authors declare that they have no competing interests.