Abstract

Background

   High dosage of dexamethasone (Dex) is an effective treatment for
   multiple diseases; however, it is often associated with severe side
   effects including muscle atrophy, resulting in higher risk of falls and
   poorer life quality of patients. Cell therapy with mesenchymal stem
   cells (MSCs) holds promise for regenerative medicine. In this study, we
   aimed to investigate the therapeutic efficacy of systemic
   administration of adipose-derived mesenchymal stem cells (ADSCs) in
   mitigating the loss of muscle mass and strength in mouse model of
   DEX-induced muscle atrophy.

Methods

   3-month-old female C57BL/6 mice were treated with Dex (20 mg/kg body
   weight, i.p.) for 10 days to induce muscle atrophy, then subjected to
   intravenous injection of a single dose of ADSCs (
   [MATH:
   <mrow><mn>1</mn><mo>×</mo><msup><mrow><mn>10</mn></mrow><mn>6</mn></msu
   p></mrow> :MATH]
   cells/kg body weight) or vehicle control. The mice were killed 7 days
   after ADSCs treatment. Body compositions were measured by animal DXA,
   gastrocnemius muscle was isolated for ex vivo muscle functional test,
   histological assessment and Western blot, while tibialis anterior
   muscles were isolated for RNA-sequencing and qPCR. For in vitro study,
   C2C12 myoblast cells were cultured under myogenic differentiation
   medium for 5 days following 100
   [MATH: <mi>μ</mi> :MATH]
   M Dex treatment with or without ADSC-conditioned medium for another
   4 days. Samples were collected for qPCR analysis and Western blot
   analysis. Myotube morphology was measured by myosin heavy chain
   immunofluorescence staining.

Results

   ADSC treatment significantly increased body lean mass (10–20%), muscle
   wet weight (15–30%) and cross-sectional area (CSA) (~ 33%) in
   DEX-induced muscle atrophy mice model and down-regulated muscle
   atrophy-associated genes expression (45–65%). Hindlimb grip strength
   (~ 37%) and forelimb ex vivo muscle contraction property were
   significantly improved (~ 57%) in the treatment group. Significant
   increase in type I fibres (~ 77%) was found after ADSC injection.
   RNA-sequencing results suggested that ERK1/2 signalling pathway might
   be playing important role underlying the beneficial effect of ADSC
   treatment, which was confirmed by ERK1/2 inhibitor both in vitro and in
   vivo.

Conclusions

   ADSCs restore the pathogenesis of Dex-induced muscle atrophy with an
   increased number of type I fibres, stronger muscle strength, faster
   recovery rate and more anti-fatigue ability via ERK1/2 signalling
   pathway. The inhibition of muscle atrophy-associated genes by ADSCs
   offered this treatment as an intervention option for muscle-associated
   diseases. Taken together, our findings suggested that adipose-derived
   mesenchymal stem cell therapy could be a new treatment option for
   patient with Dex-induced muscle atrophy.

Supplementary Information

   The online version contains supplementary material available at
   10.1186/s13287-023-03418-0.

   Keywords: Mesenchymal stem cell, Muscle atrophy, Dexamethasone, Cell
   therapy

Background

   Glucocorticoid and synthetic analogs such as dexamethasone (Dex) are
   potent anti-inflammatory and immunosuppressive drugs. The catabolic
   effects of glucocorticoid as endocrine hormones released in response to
   stress conditions or prolonged use at high dose have been reported to
   be associated with skeletal muscle atrophy [[39]1, [40]2]. For
   instance, patients with Cushing’s syndrome exhibit peripheral muscle
   weakness [[41]3]. In the past few decades, the number of long-term oral
   glucocorticoid prescription has increased by over 30%[[42]4]. Patients
   with congenital adrenal hyperplasia and Addison's disease might need to
   take long-term Dex treatment [[43]5–[44]7]. The resultant loss of
   skeletal muscle mass and muscle weakness might lead to impaired quality
   of life, increased risk of falls, decrease in wound healing,
   compromised lung function and so on.

   Healthy skeletal muscle consists of two types of muscle fibres,
   slow-twitch type I fibres and fast-twitch type II fibres. Type I
   fibres, with low ATPase activities, which have higher oxidative
   capacity and relatively more resistant to fatigue [[45]8]. Type II
   fibres can be further divided into type IIA, type IIB and IIX fibres.
   Type IIA fibres, with high ATPase activities, have both higher
   oxidative and glycolytic capacity. Type IIB and type IIX both have high
   glycolytic but low oxidative capacity. The muscle atrophy induced by
   glucocorticoid is generally characterised by reduction in the size of
   type II fibres [[46]9] and muscle force [[47]10]. Several signalling
   pathways have been proposed to explain the decreased rate of protein
   synthesis and increased rate of protein breakdown underlying muscle
   atrophy [[48]11, [49]12]. Muscle RING finger 1 (MuRF-1) and muscle
   atrophy F-box (Atrogin-1/MAFbx) are two muscle-specific E3 ubiquitin
   ligases playing important roles in muscle atrophy, which can be
   activated by glucocorticoid [[50]13]. As the underlying mechanisms are
   not fully understood, effective treatment for alleviating
   glucocorticoid-induced muscle atrophy is limited.

   The paracrine and immunomodulatory properties of mesenchymal stem cells
   (MSCs) make it one of the popular cell types in cell therapy [[51]14].
   Over a thousand clinical trials using MSCs were designed to test
   therapeutic interventions for various severe diseases such as treatment
   in orthopedics, degenerative, immune rejection, inflammation and
   autoimmunity [[52]15]. In addition to these known targeted medical
   conditions, recent study suggested that autologous transplantation of
   adipose-derived MSCs (ADSCs) with collagen hydrogel into crushed
   tibialis anterior muscle render beneficial effects in improving muscle
   function and regeneration [[53]16]. Moreover, systemic injection of
   human ADSCs in animal model of muscle injury induced expression of
   human dystrophin without immunosuppression [[54]17, [55]18]. With
   these, we hypothesised that that systemic injection of ADSCs could
   improve muscle mass and muscle functions in animal model of Dex-induced
   muscle atrophy. The signalling pathway involved in ADSC-mediated
   benefit would be explored to elucidate the underlying mechanisms.

Methods

Dex-induced muscle atrophy animal model

   Three-month-old female C57BL/6 mice were used to develop the muscle
   atrophy model. To induce muscle atrophy, mice received daily
   intraperitoneal injection of Dex (20 mg/ kg body weight, Sigma #D4902)
   [[56]19] for 10 days. Animals were housed in 12 h light/ 12 h dark
   cycles at 22–28 ℃ with standard chow diet and water. Five mice were
   kept in one standard small cage.

In vivo treatments, ADSC transplantation and ERK1/2 inhibitor injection

   Human adipose-derived mesenchymal stems (ADSC) were cultured following
   our previous publication using Iscove's modified Dulbecco's medium
   (IMDM) (Gibco #12200036) containing 10% foetal bovine serum (FBS)
   (Gibco #10270106) and 1% penicillin–streptomycin-glutamine (PSG) (Gibco
   #10378016) 10 ng/mL FGF2 (PeproTech #100-18B) [[57]20]. Mice were
   randomly divided into non-Dex-treated group (CON),
   dexamethasone-treated group (DEX) and ADSC treatment group
   (DEX + ADSC). After 10 days of dexamethasone induction, mice received a
   dose of ADSCs (
   [MATH:
   <mrow><mn>1</mn><mo>×</mo><msup><mrow><mn>10</mn></mrow><mn>6</mn></msu
   p></mrow> :MATH]
   /kg body weight in 200
   [MATH: <mi>μ</mi> :MATH]
   L PBS) or PBS through tail vein injection, respectively. Mice were
   killed with carbon dioxide asphyxiation 7 days after ADSC
   transplantation. For in vivo mechanistic study, mice received daily
   administration of MEK-ERK1/2 inhibitor U0126 (25 µM/kg body weight,
   i.p.) from day 11 for 7 days [[58]21–[59]24].

C2C12 cell culture and treatments

   C2C12 cells were cultured with high-glucose Dulbecco's modified Eagle
   medium (HG-DMEM) with 10% FBS and 1% PSG and followed with 5 days of
   differentiation (HG-DMEM medium with 2% horse serum (HS) (Gibco
   #16050122) and 1% PSG). Dex group (DEX) and ADSC treatment group
   (DEX + ADSC) received 4 days of 100 µM Dex (dissolved in DMSO and
   diluted with PBS) treatment with IMDM medium contained 10% FBS and
   1% PSG or 100 µM Dex treatment with ADSC-conditioned medium. Control
   group (CON) was cultured with IMDM medium contained 10% FBS, 1% PSG and
   DMSO (equal volume as DEX and DEX + ADSC groups) for 4 days.

   For ADSC-conditioned medium collection,
   [MATH:
   <mrow><mn>2</mn><mo>×</mo><msup><mrow><mn>10</mn></mrow><mn>5</mn></msu
   p></mrow> :MATH]
   ADSCs were seeded in 6-well plate and cultured with IMDM medium
   containing 10% FBS, 1% PSG and 10 ng/mL FGF2. Two days after cell
   culture, conditioned medium was collected and centrifuged to remove
   cell debris. For inhibitor treatment, all inhibitors were dissolved in
   DMSO and freshly diluted with culture medium. 10 µM JNK inhibitor
   (SP-600125) [[60]25, [61]26], 1 µM ERK1/2 inhibitor (PD-325901)
   [[62]27, [63]28] or 10 µM p38 inhibitor (SB-203580) [[64]29, [65]30]
   was added concurrently during ADSC-conditioned medium treatment.
   Meanwhile, equal volume of DMSO was added in DEX and DEX + ADSC groups.

Body composition

   Whole-body, hindlimb and forelimb lean mass and fat mass were measured
   by small animal dual-energy X-ray absorptiometry (DXA) (UltraFocus^DXA,
   Faxitron Bioptics, Tucson, Arizona, USA) 7 days after ADSC
   transplantation. Dedicated Bioptics Vision software was used for data
   analysis.

Muscle functional tests

   Grip strength metre was used to measure the forelimb grip strength.
   Each mouse was allowed to grasp the machine, and then its tail was
   gently pulled back and the tension was recorded automatically by the
   machine. Each mouse had five times to perform the test, and 5-min rest
   was given between each test. The highest and lower results were
   excluded, and the remainder values were recorded and normalised with
   whole body weight for analysis. Rodent Treadmill (Ugo basile #47303)
   was used to measure the fatigue-like behaviour in mice. Training
   section and fatigue test were performed by following the published
   protocol [[66]31]. Animals followed the 3-day-training protocol and one
   full day resting before starting the fatigue test. The test ended when
   the mouse remained in the fatigue zone for 5 times. Total running
   distance was recorded and present as the fatigue level. For the ex vivo
   muscle functional test, right hindlimb gastrocnemius (GA) muscles were
   carefully isolated from anaesthetised mice and tied at the Achilles’s
   tendon for mounting on Dynamic Muscle Control system (DMC v5.4; Aurora
   Scientific, Inc.) supplemented with Krebs buffer, 95% O[2] and 5%
   CO[2]. The optimal length of the GA muscle was measured, a single
   150 Hz stimulus was given for three times, and the responses were
   recorded for calculation of twitch force. For the determination of peak
   tetanic force, a continuous 150 Hz stimulus was given for three times
   and the responses were recorded. Normalised twitch force and peak
   tetanic force were achieved by dividing with GA muscle cross-sectional
   area (CSA). After resting for 5 min, the fatigability was measured by
   repeated stimulus every 5 s for a total of 300 s and the isometric
   contractions were recorded. A stimulus of 150 Hz was given to GA muscle
   5 min and 10 min after repeated stimulus, and the peak tetanic forces
   were recorded to determine the recovery rate. The results were analysed
   using the Dynamic Muscle Analysis system (DMA v3.2; Aurora Scientific,
   Inc.).

Histochemical and immunofluorescence staining

   Freshly isolated GA muscle tissues from the left hindlimb were snap
   frozen in liquid nitrogen and kept at -80℃ until further processing.
   Frozen muscles were embedded in OCT embedding medium and sectioned at
   10 µm for further staining. For immunofluorescence staining, samples
   were fixed in pre-ice methanol and blocked with 2% horse serum for 1 h
   at room temperature and followed by primary antibodies incubation
   overnight at 4℃ and secondary antibodies incubation at room temperature
   for 1 h. DAPI was used for nuclear staining. Hematoxylin and eosin
   (H&E) staining was used to obtain cross-sectional morphology of muscle
   fibre. ImageJ software was used for image data analysis. Fusion index
   was calculated as the number of nuclei presented inside each
   MyHC-positive myotube and divided by the total number of nuclei in a
   field of view.

Western blot

   RIPA buffer (Abcam #ab156034) with protease/phosphatase inhibitor
   (Thermo Scientific™ #1861821) was used to extract proteins from tissue
   and cell samples. Protein concentration was measured by using Pierce™
   BCA Protein Assay Kit (Thermo Scientific™ #23225). Samples were
   separated in 10% SDS–polyacrylamide gel and transferred to PVDF
   membranes at 100 V for 1 h. Membranes were then blocked in 5% BSA at
   room temperature and incubated with primary antibodies at 4℃ overnight,
   followed by secondary antibodies incubation at room temperature for
   1 h. Chemiluminescent substrate (Thermo ScientificTM #34095) was added,
   and membranes were placed in ChemiDoc MP imaging system (Bio-Rad) for
   signals detection. ImageJ software was used for image data analysis.
   Full blot images were provided in supplementary data (Additional file
   [67]1: Fig. S1).

RNA extraction and qPCR

   TRIzol (Invitrogen #15596018) was used for total RNA purification, and
   then High-Capacity cDNA Reverse Transcription Kit (Applied Biosystems™
   #4368813) was used to obtain cDNA. Gene expression quantification was
   performed by using the QuantStudio™ 7 Flex Real-Time PCR System
   (Applied Biosystems™). All gene expression results were normalised by
   housekeeping gene GAPDH.

Bulk RNA-sequencing

   Total RNA was extracted from GA muscles as mentioned above. RNA was
   then lysed into short fragments using a frag- mentation buffer.
   First-strand cDNA was synthesised using random N6 primers, followed by
   second-strand cDNA synthesis. The ends of double cDNA were repaired; 5′
   ends were phosphorylated, and 3′ ends formed cohesive ends with
   A-tailing. Then, cDNA was ligated to the sequencing adapters. The
   ligation products were amplified using BGISEQ platform to build a cDNA
   library and sequenced on the DNBSEQ PE100 platform. Raw data were
   preprocessed with quality control steps to remove the adapter signals
   during library preparation and low signal sequence quality reads. The
   quality control was performed with SOAPnuke (v 1.5.2). After quality
   control, the “clean data” were stored in FASTQ format for further
   analysis. Sequence alignment was performed with software HISAT2
   (v2.0.4) and mapped to template GRCm38 (NCBI,
   GCF_000001635.26_GRCm38.p6). After the mapping procedure, the total
   mapping ratio and the uniquely mapping ratio of each sequenced data
   were examined to monitor the between sample quality control. Finally,
   the sequenced samples that passed the quality check were obtained for
   further analysis.

Statistical analysis

   Sample size for each experiment was calculated by G-power software. All
   statistical analyses were performed using SPSS software and statistical
   significance was determined using one-way ANOVA (*P < 0.05, **P < 0.01,
   ***P < 0.005, #P < 0.001). Each experiment was performed at least three
   times.

Results

ADSC treatment improve muscle mass and muscle atrophy-associated gene and
protein expression

   The beneficial effects of ADSCs in skeletal muscle system were
   evaluated through multiple experiments. First, whole-body composition
   was measured by DXA machine. The results suggest that ADSC treatment
   reversed the loss of lean mass, including whole body, hindlimb and
   forelimb (Fig. [68]1A). Meanwhile, body fat which was increased by Dex
   can be alleviated by ADSC treatment. The wet weight of hindlimb muscle
   including tibialis anterior (TA), gastrocnemius (GA) quadriceps femoris
   (QA) and extensor digitorum longus (EDL) muscle was significantly
   increased by ADSC treatment (Fig. [69]1B). H&E staining results
   indicated that ADSCs are capable of reversing the GA muscle CSA
   (Fig. [70]1C). Dex is known to affect skeletal muscle through
   activating protein degradation and inhibiting protein synthesis. Thus,
   we performed the qPCR analysis for genes expressed in protein
   degradation and protein synthesis. Results showed that Dex activated
   genes involved in protein degradation (Mstn, Atrogin-1 and Murf-1) but
   had no effect on genes responsible for protein synthesis (p70s6k and
   e-IF4G-1) (Fig. [71]1D). To conclude, these results show that ADSCs
   have the ability in reversing the phenotypes in Dex-induced muscle
   atrophy.

Fig. 1.

   [72]Fig. 1
   [73]Open in a new tab

   Assessment of muscle mass after ADSCs treatment in Dex-induced muscle
   atrophy mice. A Whole-body, hindlimb and forelimb composition including
   lean mass and fat mass measured by DXA machine. B Quantification of
   tibialis anterior (TA), gastrocnemius (GA), quadriceps femoris muscle
   (QA) extensor digitorum longus (EDL) and soleus muscle wet weight. C
   Quantification of muscle CSA in H&E stained GA muscle. D Analysis of
   muscle atrophy-related genes (Mstn, Atrogin-1 and Murf-1) and protein
   synthesis-related genes (p70s6k and e-IF4g-1) genes expression in TA
   muscles with ADSCs treatment. n = 6 per group. Quantitative data are
   presented as mean ± SD. Statistical analysis are performed using
   one-way ANOVA test, with significance set at P < 0.05 (*P < 0.05,
   **P < 0.01, ***P < 0.005, #P < 0.001)

ADSC treatment improves muscle functions in Dex-induced mice

   To further understand the beneficial effect of ADSCs on skeletal muscle
   functions, forelimb grip strength and ex vivo muscle functional test
   were performed. Forelimb grip strength and max grip strength were
   markedly increased in ADSC treatment group as compared with Dex-treated
   mice; similar results were also shown in the normalised grip strength
   (Fig. [74]2A). Walking distance is one of the muscle function
   parameters; our treadmill running fatigue test result suggested that
   total running distance can be significantly improved after ADSC
   treatment (Fig. [75]2B). Ex vivo muscle functional test showed that
   ADSC treatment group has a better force response to electrical
   stimulations (Fig. [76]2C). Compared with Dex-induced mice,
   ADSC-treated mice produced ~ 57% more peak tetanic force and ~ 58% more
   twitch force in GA muscle. The half-relaxation time (time taken for
   force to decline from 50 to 25% of the peak force) was found ~ 35%
   longer in Dex-induced mice compared with control group, but ADSC
   treatment group had ~ 31% shorter half-relaxation time (Fig. [77]2D).
   After receiving continuous stimulation, no difference between control
   mice and Dex-induced mice had found (Fig. [78]2E). However,
   ADSC-treated mice showed significantly improved anti-fatigue ability.
   Also, better recovery rates were observed in ADSC-treated mice
   (Fig. [79]2F). Taken together, these data suggest that ADSCs reverse
   the loss of muscle functions caused by Dex and enhance the resistance
   to fatigue ability. Skeletal muscle fibre type has been known to
   contribute to muscle contraction. Therefore, we next checked the fibre
   type population in each group. The number of oxidative type I muscle
   fibre significantly increased after ADSC treatment; meanwhile, ADSCs
   also reversed the change of type IIA and type IIB/X fibres
   (Fig. [80]3A). The distribution of CSA of type I muscle fibre also
   shifted towards a larger size compared to DEX group (Fig. [81]3B). As
   previous studies have mentioned, oxidative type I muscle fibres are
   more fatigue resistant than type II muscle fibres. Thus, ADSC increased
   type I muscle fibre number and CSA explain the enhancement in GA muscle
   anti-fatigue ability.

Fig. 2.

   [82]Fig. 2
   [83]Open in a new tab

   Effect of ADSCs treatment in improving skeletal muscle functions and
   regulating muscle fibre types. A Hindlimb grip strength were measured
   at 7 days after ADSCs treatment with or without normalised by whole
   body weight. n = 6 per group. B Treadmill fatigue were measured 6 days
   after ADSCs treatment. n = 3 per group. C–F Ex vivo GA muscle
   contraction test. n = 5 per group. Peak tetanic fore and twitch fore
   with or without normalised by GA muscle CSA C Half-relaxation time (D).
   GA muscle fatigability normalised by prefatigued developed tension (E).
   Contraction force of GA muscle after 5 min and 10 min of fatigue (F).
   n = 6 per group. Statistical analysis are performed using one-way ANOVA
   test, with significance set at P < 0.05 (*P < 0.05, **P < 0.01,
   ***P < 0.005, #P < 0.001)

Fig. 3.

   [84]Fig. 3
   [85]Open in a new tab

   Effect of ADSCs treatment in controlling muscle fibre type switching
   (A-B) Immunofluorescence staining and quantification for fibre types in
   GA muscle sections. Blue: type I; green: type IIA; red: type IIB;
   black: type IIX staining (A). Fibre types CSA distribution (B). n = 6
   per group. Statistical analysis are performed using one-way ANOVA test,
   with significance set at P < 0.05 (*P < 0.05, **P < 0.01, ***P < 0.005,
   #P < 0.001)

Signalling pathways regulated by ADSCs in Dex-induced mice

   To determine the roles of ADSCs in improving muscle mass and muscle
   functions, we performed the RNA-Sequencing transcriptomic analysis.
   There were 1933, 2778 and 215 differentially expressed genes (DEGs)
   found in DEX vs CON, DEX + ADSC vs DEX and DEX + ADSC vs CON,
   respectively (Fig. [86]4A). The results suggested that DEX + ADSC group
   is more similar to the CON group, which has much lower DEGs compared
   with DEX group (Fig. [87]4B). There are 77.4% of the DEGs regulated by
   DEX could also affected by ADSCs (1497 out of the 1933 DEGs)
   (Fig. [88]4C). Next, we focused on the 2778 DEGs that were regulated by
   ADSCs. For the purpose of acquiring the functional classification of
   these 2778 DEGs, KEGG pathway enrichment analysis was performed
   (Fig. [89]4D). The results showed that PI3K-Akt, mitogen-activated
   protein kinases (MAPK), Rap1 and Ras signalling pathway were found to
   be listed in the top 20 functional enriched KEGG pathways. In skeletal
   muscle, MAPK was known as one of the major regulators in response to
   oxidative, energetic and mechanical stress [[90]32]. Thus, we further
   investigated the role of MAPK in regulating skeletal muscle functions
   by ADSCs.

Fig. 4.

   [91]Fig. 4
   [92]Open in a new tab

   RNA sequencing of GA muscle from control, Dex-induced muscle atrophy
   mice and ADSCs-treated mice. A Differentially expressed genes (DEGs)
   between DEX vs CON, DEX + ADSC vs DEX and DEX + ADSC vs CON. B Heatmap
   of DEGs in different samples. C Venn diagram of the overlapped DEGs
   between DEX vs CON and DEX + ADSC. D Top 20 functionally enriched KEGG
   pathway analysis of differentially expressed genes (DEGs) found in
   DEX + ADSC vs DEX

Effects of MAPK signalling in Dex-treated C2C12 myotubes

   To validate the RNA-seq results, we further used an in vitro model to
   study the necessity of MAPK for ADSC treatment. C2C12 myotube treated
   with high dosage of Dex was used as an in vitro muscle atrophy model
   [[93]33, [94]34]. After treated with ADSC-conditioned medium for
   4 days, myotube morphology and muscle atrophy gene expression was
   significantly reversed (Fig. [95]5A, B). To confirm the necessity of
   MAPK signalling pathway for ADSC treatment, Dex-induced C2C12 myotubes
   with ADSC-conditioned medium were cultured with JNK, p38 or ERK1/2
   inhibitor. After inhibiting ERK1/2 expression, the effect of ADSCs on
   improving myotube morphology was suppressed but no significant effect
   after treated with JNK or p38 inhibitor (Fig. [96]5C). Furthermore,
   muscle atrophy genes, Murf-1 and Atrogin-1, were significantly
   upregulated after inhibition of ERK1/2 compared with ADSC-treated group
   and no significant differences were found after treated with JNK or p38
   inhibitors (Fig. [97]5D). These in vitro results suggest that ADSCs
   might improve Dex-induced muscle atrophy through ERK1/2 pathway. Next,
   we measured the ERK1/2 protein expression level in both muscle tissues
   and C2C12 myotube samples to confirm our hypothesis (Fig. [98]5E,
   F, Fig. S1). Western blot results showed that higher levels of
   phosphorylated ERK1/2 protein were found in ADSC treatment group in
   both in vivo and in vitro samples. Ultimately, ERK1/2 was confirmed as
   one important pathway for ADSC in revering Dex-induced muscle atrophy.

Fig. 5.

   [99]Fig. 5
   [100]Open in a new tab

   Inhibition of MAPK signalling pathway in Dex-induced muscle atrophy
   C2C12 cells. A A schematic diagram for in vitro cell experiment. (B-C)
   The establishment of Dex-induced C2C12 cells as a muscle atrophy in
   vitro model. n = 4 per group. B Immunofluorescence staining for myosin
   heavy chain and the quantification for myotube fusion index. C
   Real-time PCR analysis of muscle atrophy genes. D The myosin heavy
   chain staining and fusion index calculation of ADSCs-treated atrophy
   C2C12 cells with MAPK inhibitors (JNK, p38 and ERK1/2). n = 3 per
   group. E Real-time PCR analysis of muscle atrophy genes for samples
   treated with inhibitors. n = 6 per group. F, G ERK1/2 protein
   expression for in vitro (F) and in vivo (G) samples. Quantitative data
   are presented as mean ± SD. Statistical analysis are performed using
   one-way ANOVA test (A-B, D) and unpaired t-test (C), with significance
   set at P < 0.05 (*P < 0.05, **P < 0.01, ***P < 0.005, #P < 0.001)

Ablation of ERK1/2 in ADSC-treated Dex-induced mice leads to a decrease
anti-fatigue ability

   To illustrate the role of ERK1/2 signalling in ADSC treatment, U0126
   was used to inhibit the activation of ERK1/2 in Dex-induced mice.
   First, we confirmed that ADSCs were not be able to activate ERK1/2
   expression after U0126 injection. Muscle wet weight of GA and QA were
   partially decreased (Fig. [101]6A). Forelimb grip strength was reduced
   after ERK1/2 inhibition (Fig. [102]6B). However, ex vivo muscle
   functional test showed that ERK1/2 inhibition barely affected the
   contraction of GA muscle (Fig. [103]6C). After continued stimulation,
   GA muscle from ERK1/2 inhibitor mice became fatigued faster and had
   slower recovery (Fig. [104]6D). As described above, fatigue resistant
   ability is highly controlled by oxidative type I muscle fibre. As
   expected, the number of type I muscle fibre decreased after ERK1/2
   inhibition (Fig. [105]6E). Distribution of fibre CSA showed that U0126
   shifted type I muscle fibre into a smaller size but not type II muscle
   fibres (Fig. [106]6F). In conclusion, U0126 can partially inhibit the
   effects of ADSC especially on improving the anti-fatigue ability and
   increasing type I muscle fibres.

Fig. 6.

   [107]Fig. 6
   [108]Open in a new tab

   The effect of ERK1/2 inhibition on muscle quality in ADSCs-treated
   muscle atrophy mice. A Quantification of muscle wet weight of TA, EDL,
   GA, Soleus and QA muscle wet weight. B Hindlimb grip strength and
   normalised by whole body weight grip strength. C Ex vivo GA muscle
   tetanic and twitch fore with or without normalised by GA muscle CSA. D
   GA muscle fatigability and recovery rate measured by the contraction
   force of GA muscle after 5 min and 10 min of fatigue. E
   Immunofluorescence staining and quantification of fibre types in GA
   muscle. F Distribution of CSA of GA fibre. n = 6 per group. Statistical
   analysis are performed using t-test, with significance set at P < 0.05
   (*P < 0.05, **P < 0.01, ***P < 0.005, #P < 0.001)

Discussion

   In this study, we reported that ADSCs could alleviate muscle wasting
   induced by Dex, especially the function of skeletal muscles. Hindlimb
   and forelimb muscle functions were enhanced by ADSC treatment, as
   demonstrated by the grip strength test and ex vivo muscle functional
   test. In the ex vivo muscle functional test, ADSC treatment improved
   the twitch force and tetanic force, which indicated better contraction
   response to force stimulation and showed better anti-fatigue ability.
   This correlation between anti-fatigue and muscle fibre types correspond
   with the results shown in previous study [[109]35]. As Type I fibres
   show more fatigue-resistance compared with fast-twitch fibres, the
   increase in size and number of type I fibres in GA muscle after ADSC
   treatment in this study explained the improvement of anti-fatigue
   ability and recovery rate.

   In addition to better anti-fatigue ability, transplantation of human
   MSC has been found to induce muscle regeneration in animals with muscle
   atrophy and reduce the expressions of muscle atrophy-associated genes
   such as Atrogin-1 and Murf-1 [[110]36]. Likewise, our results suggested
   that muscle mass and functions were significantly improved after ADSC
   transplantation in mice with Dex-induced muscle atrophy. Besides,
   findings of in vivo and in vitro experiments suggested that the
   expression of Atrogin-1 and Murf-1 was inhibited upon ADSC treatment,
   indicating that ADSC administration can rescue muscle atrophy by
   regulating the expression of muscle atrophy-associated genes. Both
   Atrogin-1 and Murf-1 are known to be regulated by the
   phosphoinositide-3-kinase (PI3K)/Akt signalling pathway [[111]37],
   which is one of the main signalling pathways involved in Dex-induced
   muscle atrophy [[112]33, [113]34]. PI3K/Akt pathway, activated by
   insulin and IGF1, regulates protein synthesis and degradation, cellular
   proliferation and survival. Akt inhibits the FoxO3a transcription
   factor, which is responsible for protein degradation. FoxO3a
   contributes to the loss of skeletal muscle protein regulated in
   lysosomal and proteasomal pathways [[114]38–[115]40]. In vivo studies
   have shown that the phosphorylated level of PI3K/Akt pathway was
   significantly decreased in Dex-induced muscle atrophy, whereas FoxO3a
   expression was increased, and E3 ubiquitin ligases were activated
   [[116]41, [117]42].

   In the present study, gene expression analysis of ADSC treatment in
   Dex-induced muscle atrophy mouse model identified significant increase
   in activity of MAPKs, including JNK, ERK1/2 and p38. While the effect
   of JNK and p38 on various skeletal muscle-associated processes,
   including proliferation, differentiation and response to contraction
   and stretch were well documented [[118]43], the effect of ERK1/2 adds a
   new perspective for understanding the underlying mechanism of its
   beneficial effect on skeletal muscle qualities. There are few studies
   suggesting the function of ERK1/2 in regulating skeletal muscle mass
   and function [[119]44, [120]45]. However, the correlation between ADSCs
   and ERK1/2 still unknown. In this study, we linked the function of
   ADSCs, ERK1/2 signalling pathway and muscle qualities together. Our
   findings correlate to recent study showing similar results that ERK1/2
   signalling pathway has a positive effect on muscle mass and functions
   in muscle atrophy animal and cell models [[121]44]. We also found that
   the activation of ERK1/2 signalling pathway by ADSCs is associated with
   higher number of type I muscle fibres, which improved the anti-fatigue
   ability of GA muscle, which is consistent with previous study showing
   that ERK1/2 regulate fast to slow muscle fibre type switching
   [[122]45]. This is the first study showing the ability of ADSCs in
   regulating ERK1/2 signalling pathway and further controlling the muscle
   fibre type switching.

   Cell-free-based therapy has drawn interest for its advantages in
   overcoming the drawback of cell-based therapy [[123]46]. Recent study
   on cell-free-based therapy has shown that skeletal muscle regeneration
   is promoted ADSC-derived secretome [[124]47], which is a cocktail of
   multiple factors and extracellular vesicles, including small exosomes
   and large microvesicles [[125]48]. Hence, we investigated the effect of
   ADSC-derived exosome on Dex-induced in vitro sarcopenia model. However,
   no significant beneficial effects were found. Moreover, similar results
   as ADSCs were found after inhibition of ADSC to release exosome. Thus,
   ADSCs might exert their functions in Dex-induced muscle atrophy model
   through other secretome rather than exosome. According to previous
   studies, stromal-cell-derived factor 4 (SDF4) found in human ADSC
   secretome could activate ERK1/2 signalling pathway through the
   interaction with CXCR4 [[126]49–[127]51]. Besides, studies suggested
   that MSC secretomes, SDF4, FGF19, could also activate ERK1/2 signalling
   pathway and are key factors for regulating skeletal muscle mass and
   functions [[128]52]. These findings support the development of
   cell-free-based therapy. However, further studies are required to fully
   understand the mechanism of ADSC secretome on muscle recovery and
   regeneration. For future translational research, it is important to
   note that Dex is often used for patients with inflammatory conditions
   and other clinical problems, which may present confounding effect of
   pre-existing medical conditions.

Conclusion

   This study provided evidence that ERK1/2 signalling pathway
   participated in systemic administration of ADSCs for Dex-induced muscle
   atrophied mice, which alleviate muscle wasting including with increased
   number of type I fibre, stronger muscle strength, faster recovery rate
   and more anti-fatigue ability, which can further support the
   development of pharmaceutical intervention for muscle atrophy. However,
   there are other mechanisms that may contribute to these effects which
   cannot be excluded. Therefore, further studies are required to confirm
   the specificity and elucidate the exact mechanism of action of this
   pathway, in order to provide more conclusive evidence.

Supplementary Information

   [129]13287_2023_3418_MOESM1_ESM.pptx^ (1.5MB, pptx)

   Additional file 1. Figure S1. Original blot of ERK1/2 protein
   expression for in vitro muscle and in vivo cell samples.

Acknowledgements