Human Mesenchymal Stem Cell-Derived Skeletal Muscle Cell Spheroids for Treating Dexamethasone-Induced Sarcopenia

Abstract

Background:

Sarcopenia, a musculoskeletal disease associated with aging or certain factors, is characterized by a reduction in muscle mass, strength, and performance. Dexamethasone (DEX)-induced muscular atrophy in animals, which shows a significant decrease in muscle mass, strength, and function, serves as a model for sarcopenia. Mesenchymal stem cell-based therapies, particularly those using 3D cultured spheroids, have emerged as a prominent area in muscle regeneration. Previous research has demonstrated that tonsil-derived mesenchymal stem cells (TMSCs) can differentiate into skeletal muscle cells (SKMCs) that exhibit attributes of skeletal muscles.

Methods:

Spheroids formed from TMSC-derived skeletal muscle cells (TMSC-SKMC-spheroids) were produced using microwells and subsequently transplanted into a sarcopenia model. This model utilized a dexamethasone (DEX)-induced muscular atrophy rat to mimic sarcopenia. The effectiveness of TMSC-SKMC-spheroid transplantation was assessed through grip strength tests, running fatigue tests, measurements of gastrocnemius muscle thickness and weight, and histopathological evaluations.

Results:

Post-transplantation, the rat models exhibited improvement in hind limb motor functions and gastrocnemius muscle regeneration. Additionally, the neuromuscular junctions in the gastrocnemius muscle of the transplantation group were restored.

Conclusion:

These findings demonstrate the therapeutic potential of TMSC-SKMC-spheroids in the DEX-induced atrophy rat model and suggest their promise as a valuable therapeutic resource for sarcopenia caused by various factors.

Supplementary Information

The online version contains supplementary material available at 10.1007/s13770-025-00753-6.

Introduction

Sarcopenia is a progressive systemic skeletal muscle disorder predominantly characterized by a loss of muscle mass and a decline in muscle function [1, 2]. It is classified into primary and secondary types. Primary sarcopenia, which is age-related [3, 4], is associated with an increased risk of falls, disability, and death [5, 6]. Secondary sarcopenia is linked to chronic conditions such as intensive care unit-acquired frailty, lack of exercise, and malnutrition [7–10], and often leads to metabolic disorders, reduced quality of life, prolonged hospitalization, and high mortality [11, 12].

Currently, there are no approved drugs for the treatment of sarcopenia; the main treatments remain exercise and diet. Efforts are underway to identify natural products with medicinal properties similar to those of foods to tackle this issue [13, 14]. Although there are no approved medications for sarcopenia other than exercise programs and nutritional support, several pharmacological agents have been suggested. Pharmacological options encompass myostatin (MSTN) inhibitors, anabolic or androgenic steroids, growth hormone (GH), angiotensin-converting enzyme inhibitors, troponin agonists, appetite stimulants, activated II receptor drugs, and β-receptor blockers; however, these treatments demonstrate variable efficacy and are associated with adverse effects. Specifically, testosterone administration is linked to increased cardiovascular risk and exacerbation of benign prostatic hyperplasia, and growth hormone therapy may cause fluid retention and orthostatic hypotension [15–18]. Consequently, various new treatment strategies for sarcopenia have been explored, including novel therapeutic formulations, drug delivery systems, tissue engineering techniques, and stem cell therapy [13, 14].

Research is underway employing various types of stem cells for treating sarcopenia, notably muscle stem cells, which are most naturally expected to regenerate skeletal muscle [19, 20]. Furthermore, Mesenchymal stem cells (MSCs) sourced from bone marrow, adipose tissue, amniotic fluid, umbilical cord, and Wharton's jelly, etc., have demonstrated effectiveness in several studies in improving sarcopenia, despite being non-muscle cells. These MSCs are recognized as promising therapeutic candidates for sarcopenia due to their capabilities for proliferation, differentiation into three germ layers, angiogenesis, immunomodulation, and paracrine secretion [19, 21]. Therapeutic effects have been observed in studies utilizing human umbilical cord MSCs (hUC-MSCs) in a mouse model of age-related sarcopenia (AAS), as well as in research employing human adipose-derived mesenchymal stem cells (hADSCs) in a mouse model of dexamethasone (DEX)-induced muscular atrophy [22, 23].

Tonsil-derived MSCs (TMSCs), isolated from discarded tissues post-tonsillectomy, exhibit superior proliferation capabilities compared to other MSC types, thus offering advantageous for therapeutic applications [24]. Previous research has demonstrated that TMSCs can differentiate into skeletal muscle (TMSC-SKMCs), and animals with muscle disease that received these cells exhibited muscle regeneration [25, 26]. The TMSC-SKMCs are expected to have therapeutic potential for sarcopenia. Furthermore, upon co-culture with motor neurons, TMSC-SKMCs exhibited expression of the acetylcholine receptor (AchR) and glucose transporter 4 (GLUT4), as well as detection of NMJs, which are functional characteristics akin to human skeletal muscle cells [27].

Despite the numerous advantages of stem cell therapy, a method for culturing and transplanting them in spheroid form rather than as isolated cells has been proposed to enhance therapeutic efficacy. MSC spheroids are reported to offer benefits such as enhanced pluripotency, increased differentiation potential, and improved anti-inflammatory properties [28–30]. While spheroids have been prepared from various cell types, the literature focusing on skeletal muscle spheroids remains scarce, with most studies concentrating on iPSC-derived spheroids [31]. In this study, we attempted to administer TMSC-SKMC-spheroids, induced from MSCs, to an animal model of sarcopenia.

The DEX-induced sarcopenia animal model is extensively utilized in research to explore muscle atrophy and sarcopenia because it effectively replicates certain characteristics of the human condition [32]. DEX, a glucocorticoid, promotes muscle atrophy by enhancing protein degradation and suppressing protein synthesis [33]. It has been reported that DEX treatment leads to significant muscle loss in animal models, a key characteristic of sarcopenia [34, 35]. Furthermore, comparative studies have demonstrated that DEX-induced muscle atrophy exhibits similarities with naturally aged sarcopenia models in terms of muscle mass loss and functional decline [32]. This similarity makes it an invaluable model for studying both primary and secondary sarcopenia. Overall, the DEX-induced sarcopenia animal model serves as a relevant and reliable platform for exploring the mechanisms and potential treatments of sarcopenia.

TMSC-SKMC has previously proven its functional effectiveness in research studies and was formed into a spheroid and administered to the gastrocnemius muscle of DEX-induced sarcopenic mice to confirm its efficacy. Grip strength tests, running fatigue tests, measurements of thickness and weight of the gastrocnemius muscle, and histopathological evaluations of the gastrocnemius muscle were conducted. The results were utilized to assess the treatment efficacy of DEX-induced sarcopenia in rats.

Materials and methods

Culture of tonsil-derived mesenchymal stem cells (TMSCs)

Human tonsils were obtained from patients undergoing tonsillectomy at Ewha Womans University Mokdong Hospital (Seoul, Korea). The investigation involving human tonsil tissue received approval from the institutional review board of Ewha Womans University Medical Center (IRB No. EUMC-2021-09-036). After tonsillectomy, the tissue was washed three times with saline, then minced in Dulbecco’s Modified Eagle’s Medium (DMEM; Hyclone, Logan, UT, USA) on cell culture dishes. The minced tissues were homogenized, transferred to a 15 mL conical tube, and incubated with Collagenase type I (210 U/mL, Invitrogen, Carlsbad, CA, USA) and DNase (10 μg/mL; Sigma-Aldrich, St. Louis, MO, USA) at 37 °C for 30 min. After digestion, the tissues were filtered through a cell strainer (BD Biosciences, San Jose, CA, USA), and the resultant cell suspension was centrifuged at 1000 rpm for 5 min. The cell pellets were then resuspended in high-glucose DMEM, augmented with 10% FBS and 1% penicillin–streptomycin (P/S; Sigma-Aldrich), and subsequently plated onto 100 mm cell culture dishes. Following 48 h of incubation in a humidified chamber with 5% CO₂, non-adherent cells were removed through washing, and fresh culture medium was administered to nurture the adherent cells. TMSCs used in this study were manufactured in a good manufacturing practice (GMP) facility.

Differentiation into TMSC-derived skeletal muscle cells (TMSC-SKMCs)

To induce differentiation of TMSCs into skeletal muscle cells (SKMCs), adherent cells were detached from dishes and cultured in suspension on 100 mm petri dishes within the culture medium for one day. The resulting spheroids were subsequently transferred to collagen-coated dishes and cultured in myoblast medium [25]. These TMSC-spheroids were maintained in this medium for four days to facilitate their differentiation into myoblasts. Following this, to further differentiate the TMSC-derived myoblasts (TMSC-myoblasts) into SKMCs (TMSC-SKMCs), the myoblast medium was replaced with the myocyte medium [25], and differentiation continued over a period of two weeks.

Culture and differentiation of human skeletal muscle cells (hSKMCs)

The progenitor cells of human skeletal muscle cells (hSKMCs), specifically skeletal muscle myoblasts (Cook MyoSite®, Cat. No. SK-1111, Pittsburgh, PA, USA), were cultured in MyoTonic Basal Medium enriched with MyoTonic Growth Supplement (Cook MyoSite®). These cells underwent differentiation into hSKMCs over a period of one week using MyoTonic Differentiation Medium (Cook MyoSite®).

Formation of spheroid

To form spheroids of uniform size and shape, cells were seeded at a density of 1 × 10⁶ (approximately 3,333 cells/microwell, AggreWellTM800, StemCell Technologies, British Columbia, Canada), and 300 spheroids were generated. TMSC-SKMC-spheroids were prepared from TMSC-derived myoblasts and cultured in TMSC-myocyte differentiation medium for 2 weeks [25]. In order to compare the in vitro properties of TMSC-SKMC-spheroids, TMSC-spheroids were prepared from TMSC and cultured in TMSC culture medium for 2 weeks.

RT-PCR

Reverse transcription polymerase chain reaction (RT-PCR) was performed as described previously [25]. The sequences of the forward and reverse primers used were as follows: PAX3 Forward: 5'–AAAGAGGAACAGCGCAGAA–3'Reverse: 5'–GAGGTCTCCGACAGCTGGT A–3'; myogenin forward, 5'-GTCTTCGCCGGGCATCCTTG-3' and reverse, 5'-GAGCTGGG GCATACACGAGG GG-3'; and glyceraldehyde 3-phosphate dehydrogenase (GAPDH) forward, 5'-TGGTATCGTGGA AGGACTCA-3' and reverse, 5'-CCTGCTTCACC ACCTTCT TG-3'.

Animal experiments

Animals

Fourteen-week-old male Sprague–Dawley (SD) rats were utilized. The sarcopenic rat model was established by administering 2.0 gastrocnemius muscles /kg DEX via intraperitoneal injection (IP) daily for 1 week. All experimental procedures and conditions complied with the approved guidelines and regulations. Twelve SD male rats were obtained from Raon Bio (Korea) and housed in a Specific Pathogen-Free facility with a 12 h light–dark cycle. Humidity and temperature were kept at 40–60% and 22 ± 0.5 °C, respectively. The protocols involving the SD rat strain received approval from the Institutional Animal Care and Use Committee at Ewha Womans University College of Medicine (IACUC No. EWHA MEDIACUC 22-043).

Transplantation

DEX-induced sarcopenic rats were randomly divided into two groups: a vehicle (PBS) only group (Sham; n = 5) and a group receiving transplantation with TMSC-SKMC-spheroids (Transplantation; n = 5). They were administered 1.0 gastrocnemius muscles /kg DEX twice a week for 8 weeks until the experiment concluded (Fig. S1A). Healthy rats of the same age served as the positive control group (Naïve; n = 5). For transplantation, 300 spheroids (1 × 10⁶ cells) per rat in a 200 uL volume of PBS were used (Table 1) (Fig. S1B and C). This mixture was equally divided and injected into the gastrocnemius muscles of both legs (Fig. S1D).

Table 1.

Experimental groups of rats in the TMSC-SKMC-spheroids transplantation test

Group	For transplantation		Rat	Number of animals (male)	Route of transplantation
	Number of spheroids (cells/spheroid)/rat	Total volume
Sham	(PBS only)	200 μL	DEX-induced muscle atrophy rat	5	• Intramuscular administration
Transplantation	60 TMSC-SKMC spheroids in PBS (1 × 106 cells/spheroid)	200 μL	DEX-induced muscle atrophy rats	5	• Gastrocnemius muscle • Treatment administered bilaterally, one site per leg Treatment administered bilaterally, one site per leg
Naïve	–	–	SD rat	5	–

Hind limb grip strength

Hind limb grip strength was assessed using a rat grip strength meter (BIO-GS3, Bioseb, Vitrolles, France). Each rat was gently held around the rib cage, allowing it to grasp the bar attached to the meter. The operator then gently pulled the rat in a rostral-to-caudal direction until it released its grip from the mesh frame. Each rat underwent 7 measurements spaced approximately 2 min apart, to establish the average grip force (g).

Running fatigue test

For the running fatigue test, rats were placed on a rodent treadmill set at a speed of 12 m/min to determine their running distance until reaching fatigue. The running time and distance were recorded automatically once the rats remained in the fatigue zone for 5 s. The treadmill was stopped after the rats met fatigue criteria five times.

Measurement of muscle thickness and weight

The thickness of the gastrocnemius muscles was measured in millimeters (mm) using a Vernier Caliper (Mitutoyo, Japan) one week before and after cell administration, as well as at intervals of 1, 2, 4, 6, and 8 weeks post-transplantation. Measurements were taken while the body was held stationary, and the right hind leg was extended to measure the gastrocnemius muscle.

Immunofluorescent staining

Cells and spheroids

The cells grown on microscope coverslips and spheroids collected in 1.7 ml Eppendorf tubes were fixed overnight at 4 °C using 4% (v/v) paraformaldehyde (Sigma-Aldrich) and 4% formaldehyde (Polysciences Inc, Warrington, PA), respectively. After fixation, the cells and spheroids underwent permeabilization using 0.5% Triton X-100 (Biosesang, Seongnam-si, Korea) for 20–30 min at room temperature (RT). Subsequently, blocking was performed with 1–2% bovine serum albumin (BSA; Bovogen, East Keilor, VIC, Australia) for 1 h at RT. They were then incubated with primary antibodies for 24 h at 4 °C, followed by incubation with secondary antibodies for 1 h at RT. The stained cells and spheroids were mounted with Vectashield containing 4',6-diamidino-2-phenylindole (DAPI; Vector Laboratories, Burlingame, CA, USA) and imaged using a fluorescence microscope (Nikon Ti2-U, Tokyo, Japan). Table S1 provides details on the antibodies used for immunofluorescence staining.

Neuromuscular junctions

After completing the behavioral experiments, the gastrocnemius muscles were harvested and immediately frozen in liquid nitrogen. The muscle tissues, stored at − 20 °C, were embedded in O.C.T. compound, oriented longitudinally. They were then sectioned at 40 µm thickness, attached to glass slides, and preserved at − 80 °C until analysis. The cryosections were warmed to RT for 30 min, fixed in 10% Neutral buffered formalin (NBF) (BBC, 0133) for 15 min, permeabilized with 0.5% Triton X-100 in phosphate-buffered saline for 15 min, and blocked in 5% bovine serum albumin with 0.5% Triton X-100 in 1X PBS for 2 h. After blocking, the sections were incubated overnight at room temperature on an orbital shaker with the primary antibody, Neurofilament-Heavy (NF-H, Santa Cruz, sc-20112). Following rinsing, the sections were incubated with Alexa Fluor™ 488 conjugated α-Bungarotoxin (α-BTX, Invitrogen, B13422) and AF568 Goat anti-Rabbit IgG (Invitrogen, A-11011) for 2 h at room temperature on an orbital shaker in the dark. The slides were subsequently covered with Vectashield mounting medium. In each selected field, imaging occurred at 1.14 µm intervals, reaching a depth of 20 µm, and a Z-stacked image (comprising approximately 15 images) was produced for each field of view using a confocal microscope (LSM800, ZEISS, Jena, Germany).

Nicotinamide adenine dinucleotide nitro-blue tetrazolium (NADH-TR) staining

Frozen transverse Sects. (15 µm thick) of the gastrocnemius muscle were stained with NADH-TR to evaluate changes in oxidative state, which serves as an additional indicator of mitochondrial density. The sections were incubated in a staining solution with a 1:1 ratio of NBT (2gastrocnemius muscles /mL) and NADH (1.6gastrocnemius muscles /mL) in TRIS buffer (pH 7.6) for 30 min at 37 °C. After incubation, the sections were washed with tap water, followed by acetone-deionized water solutions of increasing concentrations, and then cleared in xylene. Sections remained in 90% acetone until a faint purple cloud was visible over the section; subsequently, they were mounted with DPX Mountant for microscopy (06522, Sigma-Aldrich Pty Ltd, Sydney, Australia), air-dried, and stored at room temperature.

Hematoxylin and Eosin staining

To analyze the cross-sectional area (CSA) of the gastrocnemius muscle, muscle tissue fixed with 10% formalin (Sigma Aldrich, USA) was embedded in paraffin, sectioned into 4 µm-thick slices, and stained with Hematoxylin and Eosin. The stained sections were magnified 200 × to obtain eight representative images per group. CSA was quantified using the ImageJ software version 1.53t (National Institutes of Health, Bethesda, MD, USA).

Western blot

The gastrocnemius muscle tissue was homogenized in PRO-PREP buffer on ice for 30 min. The muscle samples were centrifuged at 13,000 rpm at 4 °C for 10 min, and proteins from the supernatant were isolated using the TGX Stain-Free™ FastCast™ Acrylamide Kit (BIO-RAD, Berkeley, CA, USA) in 1 × Tris/Glycine/SDS Buffer (BIO-RAD) at 80 V for 2 h. Protein transfer was performed for 10 min using Trans-Blot Turbo (BIO-RAD) with a polyvinylidene fluoride membrane (BIO-RAD) using the 1.5 mm Gel protocol. Stained membranes were developed with Clarity Western ECL Substrate (BIO-RAD) and imaged with Amersham™ ImageQuant™ 800 (Cytiva Life Sciences, formerly GE Healthcare Life Sciences, Marlborough, MA, USA). Images were analyzed using ImageQuant TL (version 10.0.261). Quantitative analysis was performed using ImageJ software (version 1.49). The antibodies for western blot are summarized in Table S2.

Statistical analysis

All experiments were conducted at least three times. Statistical analyses were performed using Prism, version 10 (GraphPad Software, San Diego, CA, USA). Results are reported as the mean ± standard error of the mean (SEM). The result of RT-PCR analysis, and the statistical significance between the Sham vs. Transplantation groups or Sham vs. Naïve groups in behavior tests were assessed using Student’s t-test. One-way analysis of variance (ANOVA) was employed to evaluate significant differences among the Sham, Transplantation, and Naïve groups, while two-way ANOVA was utilized to analyze significant differences across groups and over time. Tukey's multiple comparisons test addressed differences in grip strength, the ratio of running distance per -1 w, quantification of the NADH-TR staining, mean CSA, and western blot. The Holm-Šídák multiple comparisons test was applied to the running distance and the thickness of the gastrocnemius muscle. A p-value of less than 0.05 was considered statistically significant.

Results

Spheroids derived from TMSCs or TMSC-SKMCs

Before forming spheroids, we compared the properties of TMSCs and TMSC-SKMCs with those of human skeletal muscle cells (SKMCs). To determine the differentiation of skeletal muscle cells, we utilized immunocytochemistry to detect the expression of MYOD (green) and myogenin (red). Both markers were minimally expressed in TMSCs and were present in TMSC-SKMCs, which displayed cell fusion morphology. In hSKMCs, the positive control, both markers were observed in the myofibers (Fig. 1A). We cultured TMSCs and TMSC-SKMCs in microwells for 2 weeks to create spheroids. Both types of spheroids expressed representative skeletal muscle markers, laminin and MYH1E. However, laminin expression was uniform throughout the TMSC-SKMC-spheroids, whereas it was concentrated on the periphery of TMSC-spheroids. Furthermore, in the merged images with DAPI expression, which confirmed cellular presence, laminin and MYH1E were co-expressed only in the TMSC-SKMC-spheroids. This co-expression was further substantiated at the cellular level in enlarged spheroid images. TMSC-spheroids showed distinct expression locations for the two skeletal muscle markers without co-expression (Fig. 1B). The expression of muscle-related genes such as PAX 3 and myogenin, was investigated by RT-PCR in both cells (Fig. 1C) and spheroids (Fig. 1D). PAX3, a gene frequently expressed at the muscle progenitor cell stage and maintained at a low level until differentiation into myoblasts, was strongly expressed in TMSC compared to TMSC-SKMC (*p < 0.05, Fig. 1C), and similarly, it was strongly expressed in TMSC-spheroids compared to TMSC-SKMC-spheroids (*p < 0.05, Fig. 1D). In addition, myogenin was highly expressed in TMSC-SKMC than in TMSC (**p < 0.01, Fig. 1C), and similarly, its expression was significantly increased in TMSC-SKMC-spheroids compared to TMSC-spheroids (**p < 0.01, Fig. 1D).

Fig. 1.

Skeletal muscle properties of the spheroids used for transplantation are demonstrated. A Tonsil-derived mesenchymal stem cells (TMSCs) differentiated into skeletal muscle cells (TMSC-SKMCs) with characteristics of human skeletal muscle cells (hSKMCs) as confirmed by immunocytochemistry. MYOD and Myogenin, which are important for differentiation and development of skeletal muscle cells, were analyzed. Scale bars indicate 200 μm. B The morphology and external expression of each spheroid were examined using a fluorescence microscope following Immunohistochemistry. The analysis revealed a more pronounced co-expression of laminin and MYH1E in TMSC-SKMC-spheroids. Scale bars indicate 200 μm. Gene expression levels of myogenic markers in TMSC and TMSC-SKMC C and TMSC-spheroids and TMSC-SKMC-spheroids D were examined by RT-PCR. The expression levels were compared by quantification with GAPDH using ImageJ software. Statistical analysis using Student’s t-test was conducted for between-group comparisons, with data presented as mean ± SEM. *p < 0.05, **p < 0.01. Abbreviations: MYOD, myogenic differentiation; MYH1E, Myosin heavy chain 1E; DAPI, 4′,6-diamidino-2-phenylindole; myogenic differentiation 1; PAX3, paired box 3; GAPDH, glyceraldehyde 3-phosphate dehydrogenase

Behavioral tests and observations of the transplantation sites

Motor function, muscle thickness, and weight of the gastrocnemius, the site of transplant, as well as body weight were measured at − 1, 1, 2, 4, 6, and 8 weeks post-transplantation of TMSC-SKMC spheroids. Motor function is a crucial indicator in diagnosing the DEX-induced sarcopenia rat model, and both grip strength and running fatigue tests are well-established measures. Red, blue, and black bars in Fig. 2 represented the sham group, TMSC-SKMC-spheroids transplantation group, and naïve group, respectively (Fig. 2A–D). Grip strength in the transplant group was significantly higher starting from the first week when compared to the sham group (*p < 0.05, Fig. 2A). In the treadmill running fatigue test, there was an observable increase in the transplant group from the first week post-transplantation, but a statistically significant improvement was noted only at 6 weeks (*p < 0.05, Fig. 2B). Additionally, when analyzing treadmill test results as a ratio of values measured one week before transplantation, a statistically significant increase was observed only 6 weeks post-transplantation (*p < 0.05, Fig. 2C).

Fig. 2.

Behavioral tests and observations at the transplantation site were conducted to evaluate the therapeutic effects of TMSC-SKMC-spheroids in DEX-induced sarcopenic rats at weeks − 1, 1, 2, 4, 6, and 8 post-transplantation. A Maximum muscle strength was quantified using a grip test. B Exercise capacity was assessed by measuring the running distance over 300 s on a treadmill. C The ratio of the running distance at 1, 2, 4, 6, and 8 weeks to that at − 1 week post-transplantation was calculated. Statistical analysis using one-way ANOVA was conducted for between-group comparisons, with data presented as mean ± SEM. *p < 0.05, **p < 0.01, ***p < 0.001. Red, blue, and black bars represent the sham, transplantation, and naïve groups, respectively. D Muscle regeneration was assessed by measuring gastrocnemius muscle thickness. E The ratio of gastrocnemius muscle thickness at 8 weeks to that at − 1 week post-transplantation was calculated. (F) The ratio of gastrocnemius muscle weight to body weight was determined at the end of the experiment, 8 weeks post-transplantation. Blue squares represent the sham group, and red squares represent the transplantation group. Statistical analysis using Student’s t-test was conducted for between-group comparisons, with data presented as mean ± SEM. *p < 0.05. Abbreviations: TMSC, Tonsil-derived mesenchymal stem cells; SKMC, skeletal muscle cells; DEX, Dexamethasone

We also measured the thickness of the gastrocnemius muscle, which is the site for TMSC-SKMC-spheroids transplantation. Six weeks after transplantation, the transplant group demonstrated a statistically significant increase in muscle thickness compared to the sham group (*p < 0.05, Fig. 2D). Additionally, the muscle thickness ratio 8 weeks post-transplantation, relative to 1 week pre-transplantation, was mildly higher in the transplantation group (red squares; 1.29 ± 0.013) compared to the sham group (blue squares; 1.26 ± 0.031) (p = 0.2166, Fig. 2E). The ratio of gastrocnemius muscle weight to body weight 8 weeks post-transplantation showed a slight increase from the sham group (blue squares; 0.603 ± 0.030) to the transplantation group (red squares; 0.649 ± 0.014) (P = 0.0541, Fig. 2F).

Reversal of skeletal muscle fiber types

Muscle fiber type has been identified as crucial for controlling muscle functions. We analyzed fiber types I and II across three experimental groups, revealing distinctly different patterns between the two types (Fig. 3A–C). It is well-established that muscle function is influenced by muscle fiber type. Thus, we examined fiber types I and II, observing completely different patterns between them (Fig. 3D, ****p < 0.0001). Type I muscle fibers, prevalent in the sham group, significantly decreased to levels observed in the normal group at 8 weeks post-transplantation. Notably, type II muscle fibers, impacted by DEX-induced muscle atrophy, also significantly increased to normal group levels at this timepoint compared to the sham group (Fig. 3E, ****p < 0.0001).

Fig. 3.

Changes in fiber types of the gastrocnemius muscle after transplantation of TMSC-SKMC-spheroids. Cross section of gastrocnemius muscle stained with histochemistry for NADH-TR showing Type I (dark) and Type II (light) myofibers. Groups: Sham A and transplantation B consisting of DEX-induced sarcopenia rats at 8 weeks post-transplantation. Naïve group C: SD rats aged 24 weeks. Scale bars represent 100 μm. D Quantitative analysis of the ratio of type 1 D and type 2 E muscle fibers to total fiber count in the gastrocnemius muscles 8 weeks after transplantation. Statistical analysis utilizing one-way ANOVA was performed to compare groups, with results presented as mean ± SEM. ****p < 0.0001. The red, blue, and black boxes represent the sham, transplantation, and naïve groups, respectively. Abbreviations: TMSC, Tonsil-derived mesenchymal stem cells; SKMC, skeletal muscle cells; NADH-TR, nitro-blue tetrazolium

Myofibers and neuromuscular junctions in the cross-sectional area of gastrocnemius muscles

To evaluate the efficacy of TMSC-SKMC-spheroids through increased myofiber size, we assessed the CSA of gastrocnemius muscles using H&E staining at 8 weeks post-transplantation (Fig. 4A). Compared to the sham group, the transplantation group showed a significant increase in the myofiber ratio per CSA, although not to the extent seen in the naïve group (Fig. 4B). Furthermore, when these results were converted to the average CSA by experimental group, the transplantation group (3105.27 ± 82.12 μm²) and the naïve group (3688.81 ± 120.56 μm²) exhibited 1.4- and 1.6-fold increases, respectively, compared to the sham group (2288.59 ± 114.27 μm²), which was statistically significant (Fig. 4C, ****p < 0.0001).

Fig. 4.

Phenotypic analysis of the gastrocnemius muscle after transplantation of TMSC-SKMC-spheroids. A Histological changes in fiber diameter between DEX-induced sarcopenia rats (Sham and transplantation groups) and SD rats (Naïve group). Scale bars are set at 100 µm. B Comparison of histological changes in fiber diameters across experimental groups in 24-week-old rats. For each rat, 3 distinct fields per muscle sample were analyzed, and the fiber cross-sectional area was measured; data were normally distributed and assessed using Student’s t-test. Values are represented as mean ± SEM. **p < 0.01, ***p < 0.001, ††p < 0.01, †††p < 0.001. *: Sham vs Transplantation; †: Sham vs Naïve. C Increased cross-sectional area (CSA) of the gastrocnemius muscle at 8 weeks after transplantation of TMSC-SKMC-spheroids. Statistical comparison between groups was conducted using one-way ANOVA, with data presented as mean ± SEM. **p < 0.0001, ****p < 0.0001. The red, blue, and black bars represent the sham, transplantation, and naïve groups, respectively. Abbreviations: TMSC, Tonsil-derived mesenchymal stem cells; SKMC, skeletal muscle cells; DEX, Dexamethasone

Eight weeks after TMSC-SKMC-spheroids transplantation, we confirmed neuromuscular junction damage, which has been reported as one of the causes of sarcopenia using immunofluorescent staining of GM. NMJs were stained with α-BTX (green), binding specifically to acetylcholine receptors, while NF-H (red) was used to visualize axons. The NMJ between the axon and the muscle endplate was observed in both the transplantation and naïve groups, but was barely visible in the sham group. Furthermore, the endplates in the transplantation group displayed a normal, pretzel-like structure, although not to the extent seen in the naïve group, and were barely visible in the sham group (Fig. 5A, Fig. S2). The changes in NMJ morphology in the experimental groups indicated the regeneration of NMJs due to TMSC-SKMC-spheroids treatment. To further support these results, we confirmed the expression of phosphatidylinositol 3-kinase/protein kinase B signaling pathway (PI3K/Akt) pathway and muscle RING finger protein (MuRF1), which play an important role in regulating NMJ function, in GM (Fig. 5B). As a result, the expression of phospho-Akt (p-Akt), PI3K, and MuRF1 increased in the transplant group compared to the sham group, and in particular, the expression levels of pAkt (****p < 0.0001) and MuRF1 (**p < 0.0001) were similar between the transplant group and the sham group and were statistically significant.

Fig. 5.

NMJ regeneration in the gastrocnemius muscle of DEX-induced sarcopenia rat at eight weeks after TMCS-SKMC-spheroids transplantation. A Those muscle was examined by confocal microscopy. Before transplantation, the NMJs of the sham and transplantation groups were partially fragmented and showed a morphology similar to a nerve transection compared to the naïve group. After transplantation, the NMJs of the transplantation group showed reduced fragmentation compared to the sham group, showing a morphology similar to the naïve group. Axon terminals were visualized by immunostaining for AChR labeled with α-BTX and NF-H. The scale bar represents 20 μm. B Expression of p-Akt (/ t-Akt), PI3K, and MuRF1 in the gastrocnemius muscle, as assessed by western blot, and was quantified using Image J software (Version 1.49), and its level was normalized to that of GAPDH. One-way ANOVA and Tukey’s post hoc test were performed for comparison between groups. The data are presented as the mean ± SEM (**p < 0.01 and ****p < 0.0001). Abbreviations: NMJ: neuromuscular junction; DEX: dexamethasone; AChR: acetylcholine receptor; α-BTX: α-bungarotoxin; NF-H: neurofilament-midstream; p-Akt: phospho-protein kinase B; t-Akt: total Akt; PI3K: phosphatidylinositol 3-kinase; MuRF1: muscle RING finger protein-1

Discussion

Therapeutic approaches proposed for treating sarcopenia are currently being developed across three broad categories. The first category encompasses biological therapeutic strategies, which involve the development of monoclonal antibodies against MSTN, including Trevogrumab, Stamulumab, and Landogrozumab, as a means to treat sarcopenia, albeit with variable efficacy [36–38]. Additionally, hormonal products such as soluble activin receptor 2B (ACVR2B), GH/IGF-1 axis boosters, and selective androgen receptor modulators, including MK-0773, along with testosterone replacement therapy, have demonstrated potential in addressing sarcopenia [39, 40]. Moreover, ghrelin and anamorelin contribute to the treatment of sarcopenia by increasing appetite and weight gain, while melatonin assists by alleviating muscle dysfunction associated with sarcopenic obesity [41, 42]. Secondly, natural compounds such as curcumin have shown therapeutic potential by improving muscle fatigue, resveratrol by enhancing muscle protein synthesis and reducing muscle wasting, and ginkgo extract by bolstering muscle strength, for sarcopenia [43, 44]. However, available clinical trial results related to these therapeutics are currently limited, and comprehensive longitudinal studies are necessary for a better understanding and treatment of sarcopenia [17]. Finally, supplements have the potential to enhance physical performance and muscular health, but their effects on sarcopenia yield mixed results [45]. Key elements of supplementation, such as balanced energy intake, sufficient protein, essential amino acids, micronutrients like vitamin D, and probiotics to support gut health have been reported, yet further studies are essential to confirm their definitive benefits to muscle health [46–48].

Clinical diagnosis of sarcopenia primarily involves measuring skeletal muscle function and mass through both clinical assessments and mimicking non-clinical tests, such as the forelimb grip strength test and treadmill running test in animal model studies [1, 16]. In this context, other studies using the AAS mouse model have shown decreases of approximately ~ 35% in grip strength and ~ 65% in treadmill running distance [32, 49–51]. The DEX-induced sarcopenia rat model developed in this study exhibited reductions of ~ 43 and ~ 74% in grip strength and treadmill running distance respectively, indicating a more significant decline in muscle function than in previous studies. Moreover, the group receiving transplants of TMSC-SKMC-spheroids experienced decreases of ~ 21 and ~ 58% in these tests, respectively, which suggests partial recovery of the symptoms (Fig. 2A, B, Tables S3, S5). These outcomes appear to demonstrate a higher recovery rate compared to those from a study involving repeated administrations of hUC-MSCs in the AAS mouse model published [22].

Several studies have reported that both type I and type II muscle fibers are affected by DEX-induced muscle atrophy [32, 52], similar to our findings. Relative to the naïve group rats, there was a significant alteration in the number of type I and type II muscle fibers in the DEX-induced sarcopenia rats of the sham or transplant groups (Fig. 3). Compared to the sham group, the TMSC-SKMC-spheroid transplant group showed a reduction in type I muscle fibers but an increase in type II muscle fibers after 8 weeks, aligning closely with the baseline characteristics of the naïve group. Another study categorized type I skeletal muscle fibers as slow myosin fibers and type II as fast myosin fibers [53], which was referenced to assess the extent of myofiber damage in aged muscles. A similarly designed study that transplanted umbilical cord stem cells into AAS mice models reported results that paralleled ours [22].

Although Although there may be variations depending on species, age, and type of sarcopenia animal model generated, assessing the fiber size of hindlimb skeletal muscle can serve as a metric for measuring muscle quality. Moreover, previous studies have indicated that skeletal muscle CSA decreases by about 20–30% primarily in naturally aged mice [54, 55]. In contrast, the DEX-induced model in this study exhibited a reduction of approximately 38%, which more distinctly illustrates sarcopenia characteristics (Fig. 4). The CSA of the gastrocnemius muscle increased by about 22% 8 weeks post-transplantation of TMSC-SKMC-spheroids, akin to the observed increase in CSA of hindlimb skeletal muscles following testosterone administration in a prenatally androgenized rat model [56]. Additionally, a 33% increase in CSA was noted following the transplantation of hADMSCs into a DEX-induced muscle atrophy mouse model, but the observation period was short at 1 week. This study observed a therapeutic effect only related to the Erk1/2 signaling pathway [23]. In addition, the study involving repeated administrations of UC-MSCs in the AAS mouse model, which study observed a therapeutic effect related to the autophagy mechanism [22]. The therapeutic effect of this study such as normalization of type I muscle fiber ratio and regeneration of NMJ by TMSC-SKMC-spheroids transplantation in DEX-induced muscular atrophy rat are considered to be quite effective.

Since spheroids were used in this study, a direct comparison with spheroids derived from MSCs from other tissues is not available. However, TMSCs have characteristics suitable for clinical applications because these cells proliferate faster, have superior differentiation ability, and can secure scalability of cells compared to MSCs from other tissues [24]. Based on the characteristics of TMSCs, TMSC-SKMCs and TMSC-SKMC-spheroids differentiated from TMSCs could provide an advantage that facilitates the clinical application of compared to using MSCs derived from other tissues.

In this study, NMJ regeneration was selected to investigate the mechanism underlying the effects of TMSC-SKMC-spheroids transplantation. Sarcopenia may result from multifactorial and interrelated pathophysiological mechanisms, including aging, neuromuscular junction impairment, mitochondrial dysfunction, insulin resistance, lipotoxicity, endocrine disturbances, oxidative stress, and inflammation [1, 57]. Among these, NMJ instability and dysfunction are prominent causes that could lead to muscle weakness or paralysis, and NMJ damage is likely one of the most significant pathophysiological mechanisms of sarcopenia [57, 58]. In Fig. 5A, NMJ regeneration between axons and sarcolemma was morphologically evident in the TMSC-SKMC-spheroids treated group. This study potentially marks the first documentation of sarcopenia recovery associated with NMJ regeneration via cell therapy in a DEX-induced muscle atrophy animal model. Although electrophysiological or muscle contraction assessment is mainly used to evaluate such NMJ regeneration [59], the regulatory mechanisms or molecules of the neuromuscular junction related to sarcopenia, such as the PI3K/pAkt signaling pathway or MuRF1 analyzed in in this study, also be utilized as additional evaluation methods. The PI3K/Akt pathway plays a crucial role in regulating various aspects of NMJ function, including synaptic structure, growth, and plasticity, and the MuRF1, plays a crucial role in NMJ stability and muscle atrophy [60, 61].

Another therapeutic advantage of TMSC-SKMC-spheroids identified in this study is the use of the 3D cultured spheroid form over traditional 2D cultures. Stem cell-based spheroids, due to their self-renewal, differentiation, and protein secretion capabilities, are emerging as promising approaches in tissue engineering and regenerative medicine for various tissues, including bone, cartilage, nerve, and skin [62, 63]. Several studies have reported the use of hydrogels such as sulfated alginate to produce bone marrow-derived MSC-spheroids that support the regeneration of injured animal models [64]. Conversely, our study demonstrated that TMSC-SKMC-spheroids can be therapeutically effective for sarcopenia without the use of hydrogels. Hydrogel synthesis methods rely on chemical crosslinking, which can involve potentially toxic chemicals such as initiators and crosslinkers [65]. In addition, non-hydrogel spheroid formation methods have the advantages of simple processes and safety, which may make them more readily applicable to various biomedical and regenerative medicine fields.

In this study, the TMSC-spheroid group was excluded, in part because a previous study showed that TMSC-SKMCs showed significantly better efficacy than TMSCs when transplanted into a muscle injury model [25]. In addition, the biochemical superiority of TMSC-SKMCs had been also demonstrated in an in vitro comparative study on protein expression of undifferentiated TMSCs and TMSC-SKMCs [27], suggesting that it would be much more effective to use muscle cells as material for making spheroids for sarcopenia. Furthermore, TMSC-SKMCs could promote skeletal muscle regeneration in partial myectomy model, and GFP-labeled TMSC-SKMCs were identified in the gastrocnemius muscle of mdx mice up to 4 weeks after transplantation and promoted skeletal muscle regeneration via paracrine effect [25, 26]. Therefore, verification of the direct contribution of TMSC-SKMC-spheroids using in vivo cell tracking methods is also required to strengthen the elucidation of the therapeutic mechanism in further studies.

In conclusion, we identified that TMSCs can be differentiated into SKMCs and formed into TMSC-SKMC-spheroids, which were subsequently transplanted into DEX-induced sarcopenic rats. The behavior tests assessing grip strength and running fatigue, along with several histopathological analyses, demonstrated recovery post-TMSC-SKMC-spheroid administration. Notably, changes in muscle fiber types were a predominant indicator of muscle function recovery in DEX-induced sarcopenic rats, and the mechanism of neuromuscular regeneration was also confirmed. Moreover, the TMSCs utilized in this study were produced in a GMP-compliant facility, and similar to the hUC-MSC study on the AAS animal model, this research approaches the stage of evaluating preclinical efficacy [22]. These results suggest that TMSC-SKMC-spheroids could be a promising therapeutic resource for a broad spectrum of sarcopenia, including AAS.

Supplementary Information

Below is the link to the electronic supplementary material.

Author contributions

S.-C.J. and S.P. conceived and designed the study; Y.Y., J.Y., and Y.H.N. performed experiments; Y.Y., D.-H.K., S.-C.J., and S.P. analyzed the data; Y.Y., S.-C.J., and S.P. wrote the paper. All authors have read and agreed to the final version of the manuscript.

Data availability

The datasets generated during and/or analyzed during the current study are available from the corresponding author upon reasonable request.

Declarations

Conflicts of Interest

The authors declare no conflict of interest.

Ethical approval

The study protocol was approved by the Ewha Womans University Medical Center (EUMC) institutional review board (IRB number: EUMC-2021-09-036). All the experimental procedures were reviewed and approved by the ethics committee for animal research at Ewha Womans University (IACUC No. EWHA MEDIACUC 22–043). Informed Consent Statement: Informed written consent was obtained from all the patients participating in the study.