Introduction

Sarcopenia is a progressive and generalized skeletal muscle disorder characterized by the degenerative loss of skeletal muscle strength and mass, involving the accelerated loss of muscle mass and function [1,2,3]. Studies have reported that aging appears to result in the disturbance of homeostasis in skeletal muscle and an imbalance of tissue regeneration, leading to an overall loss of skeletal muscle. Cellular changes in sarcopenic muscle include a reduction in the size and number of myofibers. This is due to the decreased number of fast myosin fibers with age, together with intramuscular and intermuscular fat infiltration, and a decreased number of satellite cells [4]. Sarcopenia could be divided into primary sarcopenia (or age-associated sarcopenia, AAS), caused by aging, and secondary sarcopenia (or disease-associated sarcopenia, DAS), caused by diabetes mellitus, cancer, chronic obstructive pulmonary disease, or heart failure [4,5,6], which also requires appropriate treatment of the underlying disease.

With the expansion of the aging population, the problem of AAS becomes increasingly severe, which seriously impacts the lifestyle of the elderly [7, 8]. Despite physical exercise is proven to be the most effective preventative measure for AAS in mouse models, it is often impractical or inefficient for elderly individuals with reduced functional capacities. Several clinical pharmaceuticals, such as testosterone, growth factors and dehydroepiandrosterone, have been reported to be helpful in treating AAS, but with controversial effects [3, 9]. Unfortunately, there are currently no approved therapies for AAS, making it imperative to develop new treatments.

Mesenchymal stem cells (MSCs) have gained widespread use in cell therapy due to their anti-fibrosis, immunomodulatory properties, as well as their ability to release various biologically active molecules [10,11,12]. It has been reported that MSCs could enhance muscular regeneration in animal and cellular models [13,14,15,16,17,18]. Particularly, human umbilical cord-derived mesenchymal stem cells (hUC-MSCs) are advantageous due to their higher yield without the invasive procedures or ethical issues, as well as their ability to secrete a wide range of multifunctional factors [19, 20]. Our group has previously developed a complete system for obtaining clinical-grade hUC-MSCs according to the current Good Manufacturing Practice (cGMP) guidelines. The hUC-MSCs obtained through this system have been shown to meet the quality criteria of the National Institute of Food and Drug Control (NIFDC) and possess good preclinical efficacy in the intervention or treatment of neurodegeneration associated with aging [21, 22]. Furthermore, therapeutic effects of hUC-MSCs have been demonstrated on muscular atrophy experimental models [15, 17, 23]. Although these studies lack a more comprehensive evaluation of preclinical efficacy, some positive effects bring new hope for the treatment of AAS. Therefore, we speculated that hUC-MSCs might be a superior source for reversing muscle dysfunction in AAS.

Here, for the first time, we comprehensively evaluated the preclinical efficacy of clinical-grade hUC-MSCs on two AAS mouse models, including the SAMP8 mice (a senescence-accelerated mouse commonly used as the AAS model) and D-galactose (D-gal)-induced aging model (a systemic and homogeneous aging model with the acceleration of senescence). Both models were proven to have the typical phenotype of AAS in our results. Based on the behavioral test, hematoxylin-eosin (H&E) staining, immunostaining and western blotting, we found that administration of hUC-MSCs effectively improved muscle strength, restored muscle morphology and performance of aging skeletal muscle in AAS mouse models. The mechanisms involved included raising the expression of crucial extracellular matrix proteins, activating skeletal satellite cells, enhancing autophagy, and impeding the cellular senescence by down-regulating p16/p53-p21 axis.

Collectively, our results demonstrated that hUC-MSCs transplantation could improve skeletal muscle dysfunction from multiple aspects, including cellular components, cell structure and cell function, and ultimately restore muscle strength in AAS mice. Additionally, our study clarified the underlying mechanism of systematically targeting AAS therapy, at least partially through reconstructing myocyte autophagy to provide self-energy supply and down-regulating the classic p16/p53-p21 axis to delay myocyte aging. More importantly, our study provided a promising strategy for the prevention and treatment of AAS and other age-associated muscle diseases.

Results

Clinical-grade hUC-MSCs improved muscle strength and restored skeletal muscle morphology both in SAMP8 mice and D-gal-induced aging mice

Based on the detection of the cellular viability, morphology, differentiation potential and surface markers, the clinical-grade hUC-MSCs conformed to the quality standards of MSCs [21, 22] (Fig. S1). After treatment with hUC-MSCs, the behavior features of AAS mouse models were evaluated by grip test and anti-fatigue test, which are considered core metrics of sarcopenia [21, 22]. The results indicated that compared with the P8-PBS group, the mice treated with hUC-MSCs exhibited enhanced grip strength and anti-fatigue abilities (Fig. 1B, C), similar to the R1 group.

Fig. 1: Clinical-grade hUC-MSCs improved muscle strength and restored skeletal muscle morphology in SAMP8 mice.
figure 1

A Illustrated in schematic form is the chronological sequence used for hUC-MSCs or PBS treatment, behavioral tests, immunostaining, western blotting and others. B, C Evaluation of muscle strength was performed by measuring grip** (PR1&PBS = 0.0018; PMSC & PBS = 0.0012) and anti-fatigue (PR1 & PBS = 0.0001; PMSC & PBS = 0.0001) capacities. The time of Latency to fall in the Rota Rod system was used to reflect the muscle endurance, and grip** test was used to show the grip strength of SAMP8 mice (n = 6–7 per group). D The representative cross-sections of gastrocnemius muscle were stained with H&E to observe better the morphology of muscle cells in R1, P8-PBS, and P8-MSC groups (scale bar = 100 μm). E, F Quantitative analysis of muscle fiber cross-sectional area (PR1 & PBS < 0.0001; PMSC & PBS < 0.0001) (μm2) and muscle fiber diameter (PR1 & PBS = 0.0029; PMSC & PBS < 0.0001) (μm2) in SAMP8 mice. (n = 8 or 10 views per group from 5–6 male mice; all data shown as mean ± SEM, *P < 0.05, **P < 0.01, ***P < 0.001).

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The mice were euthanasia subsequently (Figs. 1A, S2A), and the gastrocnemius muscles were collected and subjected to histopathological tests by cross section and vertical section [24]. To better characterise the muscle size on different treatment, we conducted H&E staining on the cross section of gastrocnemius muscle and measured the area and diameter of muscle cells. Initially, we observed a significant reduction in the cross-sectional area and diameter in the P8-PBS group, which were restored in the P8-MSC group. A similar treatment effect was noted in the D-gal-induced aging model (Fig. S2). These results demonstrated the treatment of hUC-MSCs could restore muscle functions and morphology in AAS mouse models.

hUC-MSCs restored the ratio of slow and fast motor units of skeletal muscle in two mouse models

Skeletal muscle fibers can be classified into slow myosin (type I) fibers and fast myosin (type II) fibers [25], which play a critical role in accurately assessing the extent of muscle fiber impairment in aging muscle [26]. The primary cause of age-related loss of muscle mass is a decrease in the total number of both slow and fast myosin fibers, with the preferential atrophy of fast myosin fibers being secondary [27,28,29]. In order to estimate the potential of hUC-MSCs to prevent the slight decrease in fast fibers abundance, we examined the proportion of slow and fast myosin fibers in extensor digitorum longus (EDL). The EDL of skeletal muscle was then consecutively sectioned and stained for fast and slow muscle respectively. We observed that the unstained portion of the slow muscle section coincided with the deeply stained portion of the fast muscle section. Statistical results indicated that compared to the P8-PBS group, the ratio of slow and fast muscles in both the R1 group and P8-MSC group was lower (Fig. 2). Likewise, the same phenomenon of muscle contraction could also be found in the D-gal-induced aging model (Fig. S3). These results demonstrated that hUC-MSCs treatment restored the ratio of slow and fast motor units, thereby enhancing skeletal muscle performance.

Fig. 2: hUC-MSCs restored the ratio of slow and fast motor units of skeletal muscle in SAMP8 mice.
figure 2

A The representative immunohistochemical images of extensor digitorum longus (EDL) muscle cells illustrated the localization of fast myosin and slow myosin in R1, P8-PBS and P8-MSC mice. The square denoted the same muscle cell (scale bar = 100 μm). B The percentage of fast myosin and slow myosin in EDL muscle cells of SAMP8 mice (PR1 & PBS = 0.0173; PMSC & PBS = 0.0050) (n = 8 or 10 views per group from 5–6 male mice; all data shown as mean ± SEM, *P < 0.05, **P < 0.01, ***P < 0.001).

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hUC-MSCs regulated the extracellular matrix of muscle cells in AAS mouse models

The extracellular matrix (ECM) plays a crucial role in the growth of muscle cells by creating the cellular niche and mediating the signal transduction [30]. With aging, the ECM surrounding muscle cells could undergo change, such as decreased stability of the muscle fiber sarcolemma (MFS) and weakened cell adhesion function [31]. In order to describe the changes in AAS models, we detected the expression of dystrophin and laminin, which played critical roles in stabilizing the sarcolemma of muscle fiber and participate in cell communication [31,32,33,34]. The immunofluorescent images showed that the muscle cells in SAMP8 mice exhibited lower levels of dystrophin and laminin proteins expression than those in the R1 mice. Following treatment with hUC-MSCs, the expression levels of dystrophin and laminin were noticeably increased, implying restoration of the muscle cells ECM (Fig. 3). Similarly, the elevated expressions of dystrophin and laminin in the D-gal-induced aging model were also observed after hUC-MSCs treatment (Fig. S4). These findings indicated that hUC-MSCs preserved muscle cell adhesion, improved their microenvironment and advanced muscle toughness and tensile strength in AAS by restoring the muscle ECM.

Fig. 3: hUC-MSCs modulated the expression of important extracellular matrix proteins in SAMP8 mice.
figure 3

A The representative immunohistochemical images of ECM depicted dystrophin protein expression in the SAMP8 mouse model (scale bar = 100 μm), with individual cells expressing positive protein shown under high magnification within the square (scale bar = 25 μm). The percentage of dystrophin among different groups in the visual field area was quantified (PR1 & PBS = 0.0001; PMSC & PBS = 0.0001) (C) (n = 8 or 10 views per group from 5–6 male mice). B Representative immunohistochemistry images indicated the expression of Laminin protein in SAMP8 mice (scale bar = 50 μm). The average fluorescence value of Laminin protein expression was quantified according to the random visual field in different SAMP8 mice groups (PR1 & PBS = 0.0032; PMSC & PBS = 0.0001) (D) (n = 8 or 10 views per group from 5–6 male mice; all data shown as mean ± SEM, *P < 0.05, **P < 0.01, *** P < 0.001).

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hUC-MSCs restrained the decline in the number of muscle satellite cells in two mouse models

The decline in regenerative capacity of skeletal muscle with aging is attributed to the depletion and exhaustion of muscle stem cells (MuSCs), also known as satellite cells. To decipher whether hUC-MSCs could prevent this decline by reducing the depletion of MuSCs, we analyzed the expression of Pax7, the specific marker of MuSCs in skeletal muscle cells [35, 36]. Fluorescence image and Western blot revealed that the quantity of Pax-7+ cells in SAMP8 mice was lower than in R1 mice. Nevertheless, the number of MuSCs was significantly recovered after hUC-MSC treatment (Fig. 4). Moreover, in the D-gal-induced aging model, intraperitoneal injection of hUC-MSCs restored the number of MuSCs (Fig. S5). These results suggested that hUC-MSCs treatment can significantly recover the number of MuSCs in two AAS models, which maintained the stability of the stem cell pool, promoted proliferation and differentiation, and remodeled muscle fibers.

Fig. 4: hUC-MSCs restrained the decline in the number of muscle satellite cells in SAMP8 mice.
figure 4

A, B Representative immunohistochemical images of satellite cells in skeletal muscle showed the expression of Pax-7, and the number of Pax-7+ cells in different visual fields in SAMP8 mice was calculated (PR1 & PBS < 0.0001; PMSC & PBS < 0.0001; PR1 & MSC = 0.0006) (scale bar = 25 μm; n = 8 or 10 views per group from 5–6 male mice). C, D The expression of Pax-7 in gastrocnemius muscle of R1, P8-PBS and P8-MSC groups was detected by western blot and statistically analyzed (PR1 & PBS = 0.0691; PMSC & PBS = 0.0001; PR1 & MSC = 0.0008) (n = 3 per group; all data shown as mean ± SEM, *P < 0.05, **P < 0.01, ***P < 0.001).

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hUC-MSCs increased autophagy and delayed muscle cells senescence via p16-Rb/p53-p21 axis

It is widely acknowledged that cellular autophagy is closely intertwined with senescence. Decreased autophagy may hasten the aging process, whereas increased autophagy holds the potential for anti-aging effects [37]. To investigate how hUC-MSCs could ameliorate the muscle dysfunctions in AAS mouse models by regulating muscle autophagy, we evaluated the expression levels of Lamp2 [38] and LC3- II/I [39], which are autophagy-related biomarkers. The western blot images showed the expression levels of Lamp2 and LC3- II/I in the P8-MSC group (Fig. 5A, B) and the D-gal-MSC group (Fig. S6A, B) were remarkably elevated as compared to the aging group, indicating that autophagy was stimulated in the presence of hUC-MSCs.

Fig. 5: hUC-MSCs increased autophagy and delayed myocyte senescence through p16-Rb/p53-p21 axis in SAMP8 mice.
figure 5

A, B, C, D The expressions of Lamp2(PR1 & PBS = 0.0151; PMSC & PBS = 0.0243), LC3B II/I (PR1 & PBS = 0.0193; PMSC & PBS = 0.0016), p16(PR1 & PBS = 0.0011; PMSC & PBS = 0.0020) and p53(PR1 & PBS = 0.0001; PMSC & PBS = 0.0001) in muscle cells of R1, P8-PBS and P8-MSC mice were detected by western blot (n = 3 per group; all data shown as mean ± SEM, *P < 0.05, **P < 0.01, ***P < 0.001). E, F Representative immunofluorescence images showed the expression of p16(PR1 & PBS < 0.0001; PMSC & PBS < 0.0001; PR1 & MSC = 0.0221), p53(PR1 & PBS = 0.0005; PMSC & PBS = 0.0003) and p21(PR1 & PBS = 0.0001; PMSC & PBS = 0.0001) in skeletal muscle in R1, P8-PBS and P8-MSC mice (scale bar = 50 μm) and quantified its expression based on the mean fluorescence value in various perspectives of view (n = 8 or 10 views per group from 5–6 male mice; all data shown as mean ± SEM, ***P < 0.001).

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Several pathways, such as inhibition of p53, p16, and p21 activation, are implicated in the acceleration of senescence [40] and the inhibition of autophagy [41]. Western blot and immunofluorescence images demonstrated that the expression levels of p16, p53 and p21 were decreased in the P8-MSC group (Fig. 5C–F) and the D-gal-MSC group (Fig. S6C–F) as compared to the P8-PBS and the D-gal-PBS groups, respectively, suggesting that hUC-MSCs treatment could inhibit the activation of senescence-related pathways.

Collectively, hUC-MSCs played a pivotal role in revitalizing myocyte autophagy to furnish self-energy supply, and it could downregulate the classic p16 / p53-p21 axis to decelerate myocyte aging.

Discussion

AAS, the primary clinical malady affecting the elderly, poses a significant challenge for achieving healthy aging [Immunohistochemical experiment

The mice were perfused transcardially with ice-cold saline, followed by 50 mL of 4 % paraformaldehyde (PFA), took out their muscle tissue, and fixed in 4 % PFA overnight. Each tissue was embedded in paraffin and sectioned coronally into 5-μm-thick slices. A standard histological immunohistochemical protocol was performed, which involved dewaxing and rehydration of the sections, incubating them with 3% H2O2 in methanol, and retrieving the antigen by placing the slides in target retrieval solution at 95 °C for 20 min. The sections were then incubated overnight with Monoclonal Anti-Myosin (Skeletal, Fast) antibody (Sigma, Germany, M4276), Monoclonal Anti-Myosin (Skeletal, Slow) antibody (Sigma, Germany, M8421), Rabbit polyclonal to Anti-Dystrophin (Abcam, USA, ab15277), or Anti-Myosin Heavy Chain Antibody (Sigma, Germany, 05-716) at 4 °C. After incubation with secondary antibody at room temperature for 30 min, the slides were incubated with HRP-peroxidase complex at room temperature for 30 another minutes. Reaction products were visualized using 3,3-diaminobenzidine (DAB) for counterstaining.

Immunofluorescent staining

The sections of the gastrocnemius muscle were subjected to dewaxing and rehydration, followed by fixation in pre-cooled acetone at 4 °C for 25 min, and permeabilization in PBS containing 0.1% Triton X-100 for 15 min. The sections were then blocked for 1 h with 10 % normal Donkey Serum, before incubation with Rabbit polyclonal to Anti-Laminin (Abcam, USA, ab11575) or Anti-Pax7 Rabbit pAb antibody (Servicebio, China, GB113190) and Anti-p21 Rabbit pAb (Servicebio, China, GB11153). Incubation was carried out overnight at 4 °C followed by washing with PBS three times and incubation with the secondary antibody (Invitrogen, USA) for one hour at room temperature, followed by another round of washing. The nucleus was observed using 2 μM DAPI (Servicebio, China, G1012).

Hematoxylin-Eosin staining

The sections of the gastrocnemius muscle underwent dewaxing and rehydration procedures, followed by hematoxylin staining for a duration of 10 min. The sections were then washed with running water, differentiated with 0.7% hydrochloric acid and ethanol for a few seconds, and rinsed with tap water. After 15 min, the sections were stained with 95% ethanol, alcohol-based eosin, 95% ethanol (I, II), 100% ethanol (I, II), xylene and xylene (I, II) for 30 seconds each, and finally covered a film. H&E staining was used for distinguishing skeletal muscle (red).

Western blot analysis

Western blots were performed and analyzed as previously described [22]. Briefly, gastrocnemius muscle samples were homogenized in RIPA buffer, and then centrifuged at 12000 × g for 30 min at 4 °C. The quantification of protein in supernatant was accomplished using a BCA kit (Beyotime Biotechnology, Wuhan, China). The protein samples were boiled in the presence of sample buffer at 95 °C for 3 min. The proteins were subjected to the separation by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE), and subsequently transferred onto a nitrocellulose membrane. The target protein was probed by a corresponding antibody and then visualized by enhanced chemiluminescence (ECL) reagent and imaged by the chemiluminescence imaging system Amersham Imager 680 (General Electric Company, USA). Antibodies used for Western blots were: Anti-Pax7 (Abcam, USA, ab199010), Anti-alpha smooth muscle Actin (Abcam, USA, ab5694), Anti-LAMP2-Lysosome Marker (Abcam, USA, ab25631), Anti-Beta actin (Abcam, USA, ab8226), Anti-Cleaved LC3B (Sigma, Germany, L7S43), Anti-ARPC5/p16 ARC (Abcam, USA, ab51243), p53 (1C12) Mouse mAb (Cell Signaling Technology, USA, 1C12). Origin images of all western blot have been uploaded as a single ‘Supplemental Material’ file.

Quantitative and statistical analysis

The cell density, nuclear area, cell area, and antibody expression in Immunohistochemical experiment, Immunofluorescent staining and Hematoxylin-Eosin staining were acquired via images by Tissue FAXS (Tissue Gnostics GmbH, Vienna Austria) with a Zeiss Axio Imager Z2 Microscope System at ×200 magnification. The cross-sectional area of 500 selected skeletal muscle fibers in the stained sections was measured and calculated using ImageJ software (version 1.8.0, National Institutes of Health) and Image-Pro Plus (version 6.0.0, media cybernetics).

Statistical analyses were executed with GraphPad Prism 9.0 software (GraphPad Software, San Diego, CA, USA) and presented as the mean value accompanied by the plus or minus standard error of the mean. The statistical significance of the differences among the three groups was determined using one-way analysis of variance (ANOVA), followed by Tukey post hoc tests, as depicted in the bar graph. A value of P <0.05 was considered to be statistically significant.