Open Access

20(s)‑ginseonside‑Rg3 modulation of AMPK/FoxO3 signaling to attenuate mitochondrial dysfunction in a dexamethasone‑injured C2C12 myotube‑based model of skeletal atrophy <em>in vitro</em>

Corrigendum in: /10.3892/mmr.2021.12259

  • Authors:
    • Manying Wang
    • Rui Jiang
    • Jianzeng Liu
    • Xiaohao Xu
    • Guang Sun
    • Daqing Zhao
    • Liwei Sun
  • View Affiliations

  • Published online on: February 26, 2021
  • Article Number: 306
  • Copyright: © Wang et al. This is an open access article distributed under the terms of Creative Commons Attribution License.

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Muscle atrophy, a side effect from administration of the anti‑inflammatory medication dexamethasone (DEX), is preventable by concomitant administration of the major monomeric constituent of Panax ginseng C.A. Meyer, 20(S)‑ginsenoside Rg3 (S‑Rg3). Putative S‑Rg3‑associated prevention of DEX‑induced muscle atrophy may involve S‑Rg3 mitigation of DEX‑induced mitochondrial dysfunction. In the present study, MTT assays revealed enhanced cell viability following S‑Rg3 treatment of DEX‑injured C2C12 myotubes. Subsequent PCR and western blotting results demonstrated S‑Rg3‑induced reduction of expression of muscle atrophy F‑box protein (atrogin‑1) and muscle RING‑finger protein‑1, proteins previously linked to muscle atrophy. Additionally, S‑Rg3 treatment of DEX‑injured myotubes led to aggregation of Rg3 monomers in cells and dose‑dependent increases in cellular mitochondrial basal respiratory oxygen consumption rate and intracellular ATP levels compared with their levels in untreated DEX‑injured myotubes. In addition, S‑Rg3 treatment significantly reversed DEX‑induced reductions of expression of key mitochondrial respiratory electron transport chain subunits of protein complexes II, III and V in DEX‑injured myotube cells. Furthermore, S‑Rg3 alleviation of mitochondrial dysfunction associated with DEX‑induced injury of C2C12 myotubes was linked to S‑Rg3‑associated decreases in both forkhead box O3 (FoxO3) protein expression and phosphorylation of AMP‑activated protein kinase (AMPK). Collectively, these results implicate S‑Rg3 modulation of signaling within the AMPK‑FoxO3 pathway as a putative mechanism underlying S‑Rg3 alleviation of DEX‑induced muscle atrophy.


In clinical settings, the synthetic glucocorticoid (GC) dexamethasone (DEX) and its derivatives are routinely used to treat inflammatory disorders (1). However, excessive use of DEX has been linked to negative human health effects that include glucose metabolic dysfunction, insulin resistance, mitochondrial dysfunction and muscle atrophy (2,3). Muscle atrophy may be linked to long-term DEX treatment, which leads to altered mitochondrial morphology, aggregation (4), enlargement and compromised mitochondrial oxidative capacity (5). Due to the fact that muscle atrophy can greatly affect the well-being of patients (6), alleviation of DEX-induced muscle atrophy would be of great benefit to patients with inflammatory diseases who rely on DEX treatment.

Chronic corticosteroid treatment has been shown in numerous clinical and animal studies to cause DNA oxidative damage resulting from disruption of mitochondrial morphology and oxidative capacity (7). Indeed, such effects have been reported frequently in conjunction with clinically apparent mitochondrial dysfunction and skeletal muscle atrophy linked to long-term and/or high-dose GC treatments (8,9). Muscle atrophy associated with DEX treatment has been experimentally linked to decreased protein synthesis accompanied by increased protein degradation (10), both of which appear to be triggered by ubiquitin-proteasome system activation involving muscle atrophy F-box protein (atrogin-1) and muscle RING finger-1 (MuRF1) protein (11). These two muscle-specific proteins function as E3 ubiquitin ligases that are expressed early during the muscle deterioration process before muscle loss becomes apparent and thus may participate as key participants in DEX-associated muscle atrophy.

Notably, GCs induction of muscle atrophy-associated protein breakdown appears to involve effects on transcription factor production, specifically of forkhead box (Fox) O1, FoxO3a (12) and GSK3β (13), that trigger ubiquitin proteasome system-dependent proteolysis of muscle proteins and subsequent autophagy (14). Meanwhile, studies of muscle atrophy induced by denervation have identified increased mitochondrial fission and resulting fragmentation as key signals responsible for triggering the AMP-activated kinase-forkhead box O3 (AMPK-FoxO3) signaling pathway (15,16). Mitochondria are organelles that provide energy to cells. Matching energy supply with energy demand is coordinated through various processes and is critical for cellular adaptation and survival under changing conditions (17). Damage to mitochondrial function can lead to insufficient ATP supply and activation of the AMPK/FoxO3 pathway to cause muscle atrophy. After AMPK-FoxO3 pathway activation is triggered via the metabolic sensor AMPK, FoxO3 is dephosphorylated then undergoes nuclear translocation. Once in the nucleus, it acts as a transcription factor that upregulates atrophy-inducing genes expression that engage in events culminating in protein degradation and eventual atrophy of skeletal muscle (18). In support of this mechanism, findings of a previous in vitro study suggested that nuclear translocation of FoxO3 led to upregulation of MuRF1 expression and subsequent myotube atrophy (19).

Numerous studies have shed light on several putative mechanisms that may be responsible for observed beneficial effects of Panax ginseng compounds on muscle health. For example, studies in rats have shown that treatment with traditional Chinese herbal medicine Panax ginseng prevented muscle atrophy (20), while a diabetic mouse study demonstrated that panaxatriols extracted from ginseng exerted a similar effect on skeletal muscle (21). In one study, treatment with the panaxatriol Rg1, a ginsenoside, prevented degradation of C2C12 myotube muscle protein through regulation of the AKT/mTOR/FoxO signaling pathway (22), with prevention of atrophy attributed in another study to AKT/mTOR pathway activation (23). Another panaxatriol, 20(s)-ginsenoside Rg3 (S-Rg3), a major monomeric ginsenoside also isolated from Panax ginseng, appeared to prevent TNF-α-induced myotube atrophy (24). Our previous study demonstrated that S-Rg3 could prevent myotube atrophy by promoting myoblast differentiation (25). However, no investigations of mechanisms involved in S-Rg3 prevention of myotube atrophy have focused on S-Rg3 effects on mitochondrial function, even though DEX is known to cause mitochondrial dysfunction. Therefore, the present study investigated whether S-Rg3 treatment of DEX-injured C2C12 myotubes could prevent muscle atrophy by alleviating mitochondrial dysfunction induced by DEX.

Materials and methods


Chemicals, reagents and kits were obtained from commercial sources as follows: S-Rg3 (Urchem Sinopharm Chemical Reagent Co., Ltd.), dexamethasone (DEX)and 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) (Sigma-Aldrich; Merck KGaA), mouse skeletal muscle-derived C2C12 myoblasts (American Type Culture Collection), fetal bovine serum (FBS; Clark BioScience), horse serum (Gibco; Thermo Fisher Scientific, Inc.), antibodies against cleaved caspase-3 (cat. no. 9661), phosphorylated-AMPK (p-AMPK; cat. no. 2535), AMPK (cat. no. 2532), Bcl-2 (cat. no. 3498), Bax (cat. no. 2772), FoxO3 (cat. no. 2497), histone H3 (cat. no. 4499) and β-tubulin (cat. no. 2146) were obtained from Cell Signaling Technology, Inc.; atrogin-1 antibody (cat. no. ab168372) and total OXPHOS Rodent WB Antibody Cocktail (anti-complex I, II, III, IV, V; cat. no. ab110413) were obtained from Abcam; antibody specific for muscle-specific RING finger 1 (MuRF1; cat. no. bs-2539R) was purchased from BIOSS. Chemiluminescence reagents was purchased from Santa Cruz Biotechnology.

Cell culture and induction of cell differentiation

C2C12 myoblasts were cultivated via incubation at 37°C in an atmosphere containing 5% CO2 with humidification in medium containing Dulbecco's modified Eagle's medium (DMEM) with 25 mM glucose, 10% FBS, 100 µg/ml streptomycin and 100 units/ml penicillin until 70–80% confluence. Next, myoblasts were seeded at 7.5×104 cells/well in 6-well plates or at 5×103 cells/well in 96-well plates. Myoblast fusion to form C2C12 myotubes was induced by culturing cells for 5 days in DMEM containing 2% horse serum and 25 mM glucose (26).

MTT cell viability assay

MTT assays were conducted to monitor cell viability (27,28) as follows: C2C12 myotubes treated for 24 to 48 h with various concentrations of DEX (0, 25, 50, 100 or 200 µM) (29) or with S-Rg3 (0, 0.02, 0.2 or 2 µM) and/or DEX (200 µM) for 24 h were incubated with MTT (0.5 mg/ml) for 4 h. All treatments were conducted at 37°C. Next, formazan dissolved in 150 µl DMSO was added to wells then measurements of absorbance levels of wells were taken at a wavelength of 490 nm at room temperature. Calculations of cell viability were performed that generated results as percentages of viable cells relative to vehicle control.

Intracellular ATP level measurement

After C2C12 myotubes were treated with various concentrations of DEX (0, 25, 50, 100 or 200 µM) for 24 h, 50 µM DEX for 0, 3, 6, 12 or 24 h or S-Rg3 (0, 0.02, 0.2 or 2 µM) and/or DEX (50 µM) for 6 h, all at 37°C, C2C12 myotubes were lysed using ATP lysis buffer composed of 0.5% Triton X-100, 100 mM glycine, pH 7.4 then lysates were centrifuged at 15,000 × g for 10 min at 4°C (30). Supernatants were collected and an ATP Bioluminescent Assay kit (Promega Corporation) was used to measure intracellular ATP levels in supernatants on ice according to the manufacturer's instructions.

Oxygen consumption rate (OCR) measurement

Basal respiration rates of mitochondria were measured via previously reported methods using a kit (MitoXpress® Xtra Oxygen Consumption Assay; Agilent Technologies, Inc.) (31). In brief, C2C12 myoblasts previously seeded into 96-well microplates (Corning, Inc.) were incubated for 5 day to allow them to differentiate. C2C12 myotubes were then treated with various concentrations of DEX (0, 25, 50, 100 or 200 µM) for 24 h, 50 µM DEX for 0, 3, 6, 12 or 24 h or S-Rg3 (0, 0.02, 0.2 or 2 µM) and/or DEX (50 µM) for 6 h, all at 37°C. OCRs were measured using a plate reader at room temperature (Cytation 5; BioTek Instruments, Inc.).

Transmission electron microscopy

C2C12 myotubes were treated with 50 µM DEX with or without 0.2 µM S-Rg3 for 6 h at 37°C. After cells were fixed by 2-h immersion in 2.5% glutaraldehyde fixative at 4°C, they were post-fixed in 1% OsO4 at room temperature for 2 h, then dehydrated in an ascending alcohol series and embedded in Epon resin. Using a microtome (Leica UC7; Leica Microsystems, Inc.), ultrathin sections (60–80 nm) were sliced from embedded cell blocks, then sections were stained with 2% uranyl acetate and lead citrate at room temperature for 15 min each, then examined using an electron microscope (Tecnai G2 20 TWIN, FEI; Thermo Fisher Scientific, Inc.). To measure DEX-induced changes in mitochondrial number and area, images were processed using Image-Pro Plus 6.0 software (Media Cybernetics, Inc.) and data were analyzed by a statistician who had no access to the images (blind data analysis).

Mitochondrial membrane potential detection

C2C12 myotubes were treated with 50 µM DEX and 0, 0.02, 0.2 or 2 µM S-Rg3 for 6 h at 37°C. A JC-1 fluorescent probe (Beyotime Institute of Biotechnology) was used to estimate the effect of S-Rg3 on mitochondrial membrane potential using the manufacturer's instructions provided with the probe, but with modifications (32). After 5-days culture of C2C12 myoblasts seeded at 7.5×104 cells/well in 6-well plates, cells were treated for 6 h with S-Rg3 and DEX, each at various concentrations. Cells were next incubated in the dark with JC-1 stain for 20 min at 37°C then stained cells were immediately analyzed using flow cytometry (FACScan; BD Biosciences).

RNA extraction and reverse transcription-quantitative (RT-q) PCR

C2C12 myoblasts seeded at a density of 7.5×104 cells/well into 6-well plates were induced with DMEM containing 2% horse serum and 25 mM glucose for 5 days, then treated with 200 µM DEX for 24 h and collected on ice. After extraction of total RNA using TRIzol® reagent (Invitrogen; Thermo Fisher Scientific, Inc.) according to the manufacturer's protocols, cDNA was generated from total RNA (1 µg) via a reverse transcription kit (cat. no. KR118-02; Tiangen Biotech Co., Ltd.) on ice, then a real-time PCR system (Eppendorf) was used to conduct qPCR. The following thermocycling conditions were used for the qPCR: Initial denaturation at 95°C for 5 min; followed by 40 cycles of 95°C for 15 sec, 60°C for 30 sec and 72°C for 30 sec. Labeling of amplification products generated via qPCR was achieved using SYBR Green Master Mix (Takara Biotechnology Co., Ltd.) and gene-specific primers. Primer sequences were as follows: MuRF1, 5′-TGGAAACGCTATGGAGAACC-3′ (forward) and 5′-ATTCGCAGCCTGGAAGATG-3′ (reverse); Atrogin-1, 5′-CTGGCAGCAGCAGCTGAATAG−3′ (forward) and 5′-CACATGCAGGTCTGGGGCTGC−3′ (reverse); actin, 5′-AGGCCCAGAGCAAGAGAGGTA−3′ (forward) and 5′-CCATGTCGTCCCAGTTGGTAA−3′ (reverse). Amplification products generated from housekeeping gene actin mRNA were used to normalize the levels of the other products. Relative mRNA expression was calculated based on the 2−∆∆Cq method (33).

Western blot analysis

C2C12 myotubes were treated with 50 or 200 µM DEX and 0, 0.02, 0.2 or 2 µM S-Rg3 for 6 or 24 h at 37°C. After washing cells twice with phosphate-buffered saline, they were suspended in RIPA lysis buffer (Beyotime Institute of Biotechnology) and allowed to lyse for 30 min at 4°C. Nuclear and cytoplasmic proteins were isolated separately using a BestBio BB-3102 kit (BestBio). Protein concentration was determined using a BCA Protein Assay kit (Beyotime Institute of Biotechnology). Lysates (30 µg total/lane) were separated via 12% SDS-PAGE followed by electro-transfer to polyvinylidene fluoride (PVDF) membranes. After blocking of membranes in 5% bovine serum albumin (cat. no. 4240; BioFroxx) at room temperature for 1 h, membranes were incubated overnight with 1:1,000 dilution of primary antibody at 4°C followed by two washes and a 1-h incubation with 1:5,000 of horseradish peroxidase-conjugated goat anti-rabbit IgG (cat. no.) BA1054 and goat anti-mouse IgG (cat. no. BA1050) secondary antibodies (Boster Biological Technology) at room temperature. Proteins were visualized via chemiluminescence (ProteinSimple), then imaged using a FluorChem HD2 system (ProteinSimple). Densitometry was performed using AlphaView SA software (ProteinSimple) (34).

Statistical analysis

All statistical analyses were performed with GraphPad Prism 7 (GraphPad Software, Inc.) and using an unpaired Student's t-test or one-way analysis of variance (ANOVA) followed by Tukey's post-hoc tests. Data are expressed as the mean ± standard deviation of three independent experimental repeats. P<0.05 was considered to indicate a statistically significant difference.


Induction of muscle atrophy and mitochondrial dysfunction by dexamethasone (DEX) treatment

Results of experiments whereby C2C12 myotubes received treatment with various doses of DEX (25, 50, 100 and 200 µM) for 24 and 48 h revealed no effect on cell viability at DEX concentrations below 200 µM (Fig. 1A). Meanwhile, increased levels of two muscle atrophy marker proteins, MuRF1 and atrogin-1, were observed in C2C12 myotubes treated with 200 µM DEX (Fig. 1B). Next, measurements of C2C12 myotube OCR and intracellular ATP level were performed to reveal the relationship between muscle atrophy and mitochondrial function. Subsequently it was observed that 24-h DEX treatment (50–200 µM DEX) led to ATP deprivation (Fig. 1C) and reduced OCR (Fig. 1D). Next, C2C12 myotubes were treated with DEX (50 µM) for various lengths of time, with the results revealing that 6-h DEX treatment of C2C12 myotubes led to ATP deprivation (Fig. 1E) and OCR reduction (Fig. 1F). When considered together, these results demonstrated that damage to mitochondria preceded myotube atrophy, since 50 µM DEX treatment of C2C12 myotubes for only 6 h led to mitochondrial dysfunction, while 24-h treatment with 200-µM DEX was necessary to induce myotube atrophy. Therefore, to generate a mitochondrial damage model the present study employed 6-h administration of 50 µM DEX and for the myotube atrophy model administered 24-h treatment with 200 µM DEX.

Figure 1.

Muscle atrophy and mitochondrial dysfunction induced by DEX. (A) After 24 or 48-h treatment with various doses of DEX (25, 50, 100 and 200 µM), no effect on C2C12 myotube viability was observed at DEX concentrations <200 µM. **P<0.01 vs. CON (24 h); ##P<0.01 vs. CON (48 h). Results were analyzed via one-way ANOVA with data expressed as the mean ± standard deviation (n=3). (B) DEX-treated C2C12 myotube atrogen-1 and MuRF1 mRNA levels were higher compared with corresponding levels in control cells. **P<0.01 vs. CON. (MuRF1); ##P<0.01 vs. CON (atrogin-1). Results were analyzed via Student's t-test, with data expressed as the mean ± standard deviation (n=3). (C) Intracellular ATP level was reduced following 24-h treatment with various DEX doses (25, 50, 100 and 200 µM). *P<0.05 and **P<0.01 vs. CON. Results were analyzed via one-way ANOVA, with data expressed as the mean ± standard deviation (n=3). (D) Basal respiration (OCR) measured by MitoXpress® Xtra Oxygen Consumption Assay was reduced at 24 h following administration of various doses of DEX (25, 50, 100 and 200 µM). *P<0.05 and **P<0.01 vs. CON. Results were analyzed via one-way ANOVA, with data expressed as the mean ± standard deviation (n=3). (E) Reduction of intracellular ATP level following DEX incubation for various time points in hours. *P<0.05 vs. CON. Results were analyzed via one-way ANOVA, with data expressed as the mean ± standard deviation (n=3). (F) Reduction of OCR following DEX administration for various time points in hours. *P<0.05 vs. CON. Results were analyzed via one-way ANOVA, with data expressed as the mean ± standard deviation (n=3). DEX, dexamethasone; CON, 0 µM DEX; MuRF1, muscle RING finger-1; atrogin-1, muscle atrophy F-box protein; OCR, oxygen consumption rate.

Treatment with S-Rg3 reverses C2C12 myotube atrophy induced by DEX

After verifying that DEX-induced myotube atrophy had occurred as the present study's model for skeletal muscle atrophy, effects of S-Rg3 treatment on DEX-induced injury were evaluated. DEX treatment alone decreased C2C12 myotube cell viability to 68.68±1.21%, while S-Rg3 treatment restored C2C12 myotube cell viability in a dose-dependent manner (Fig. 2A). The expression levels of muscle cell-specific major muscle atrophy-associated E3 ubiquitin ligases atrogin-1 and MuRF1 were examined to assess S-Rg3 effects on DEX-induced skeletal muscle cell atrophy. Notably, DEX increased expression of both atrogin-1 and MuRF1 proteins in C2C12 myotubes, while reduced expression of the two proteins was observed following S-Rg3 treatment (Fig. 2B-D). The effect of S-Rg3 on apoptosis caused by DEX was also tested. The expression levels of apoptosis-related proteins, such as cleaved caspase-3, Bcl-2 and Bax were examined by western blot analysis. The expression of cleaved caspase-3, which involved in the activation cascade of caspases responsible for apoptosis execution was decreased by S-Rg3 treatment for 24 h in DEX-induced C2C12 myotubes, and the ratio of the antiapoptotic protein Bcl-2 and proapoptotic protein Bax was increased following treatment with S-Rg3 for 24 h (Fig. 2E-F).

Mitochondrial morphological changes induced by DEX are reversed by S-Rg3 treatment

Previous studies conducted using a C2C12 skeletal muscle cell line have demonstrated that mitochondrial morphological changes occur during mitophagy, a necessary cellular process for removing damaged mitochondria (35,36). Using mitochondrial volume as a morphological indicator, DEX-induced mitochondrial injury was analyzed in C2C12 myotubes using electron microscopy (Fig. 3A). Following DEX treatment, relatively large mitochondrial structural changes were observed, including swollen and broken mitochondria or the absence of mitochondria. These changes were ameliorated by treatment with S-Rg3 as evidence that S-Rg3 restored mitochondrial function (Fig. 3B and C).

DEX-induced mitochondrial dysfunction is alleviated by S-Rg3 treatment

To reveal whether an association exists between DEX-induced muscle atrophy and mitochondrial dysfunction, intracellular ATP levels and OCRs were measured in DEX-injured C2C12 myotubes. Ultimately, low ATP levels were observed following treatment with DEX that were increased following subsequent S-Rg3 treatment (Fig. 4A). In addition, S-Rg3 treatment reversed DEX-induced mitochondrial respiratory dysfunction (Fig. 4B), reversed DEX-induced reduction of mitochondrial membrane potential (an early sign of apoptosis in C2C12 myotubes) (Fig. 4C) and reversed DEX-induced reductions of JC-1 aggregate levels from below control levels (as revealed via staining followed by flow cytometry) to levels above levels of untreated DEX-injured cells (Fig. 4D). Finally, as an additional assessment of whether mitochondrial dysfunction was induced by DEX, expression levels of key mitochondrial respiratory electron transport chain subunit proteins of complexes II, III and V were measured and significantly reduced expression levels of all key subunit proteins following DEX treatment were observed; these levels were increased following addition of S-Rg3 (Fig. 4E and F).

Figure 4.

S-Rg3 reversal of mitochondrial dysfunction induced by DEX. (A) DEX-injured C2C12 myotube intracellular ATP level was increased by S-Rg3 treatment. *P<0.05 vs. CON; #P<0.05 and ##P<0.01 vs. DEX. Results were analyzed via one-way ANOVA, with data expressed as the mean ± standard deviation (n=3). (B) OCR as measured by MitoXpress® Xtra Oxygen Consumption Assay measurement showing OCR decrease following 6-h DEX administration that was restored following administration of S-Rg3. *P<0.05 vs. CON; #P<0.05 and ##P<0.01 vs. DEX. Results were analyzed via one-way ANOVA, with data expressed as the mean ± standard deviation (n=3). (C) After 6-h DEX and/or S-Rg3 treatments, mitochondrial membrane potential was assessed following incubation of C2C12 myotubes with the JC-1 probe followed by flow cytometric analysis. (D) Bar graph representing cells staining positive for JC-1 monomer then analyzed as in (C). ***P<0.001 vs. CON; ##P<0.01 and ###P<0.001 vs. DEX. Results were analyzed via one-way ANOVA, with data expressed as the mean ± standard deviation (n=3). (E) Representative western blots of DEX-injured C2C12 myotube complex I, II, III, IV and V proteins. (F) Relative expression of complex I, II, III, IV and V proteins quantified by densitometric analysis, with β-tubulin serving as loading control. Data are expressed as the mean ± standard deviation (n=3). **P<0.01 vs. CON; #P<0.05 and ##P<0.01 vs. DEX. Results were analyzed via one-way ANOVA, with data expressed as the mean ± standard deviation (n=3). S-Rg3, 20(S)-ginsenoside Rg3; DEX, dexamethasone; OCR, oxygen consumption rate; CON, control group.

AMPK/FoxO3 signaling pathway inhibition by S-Rg3 in the DEX-injured C2C12 myotube-based muscle atrophy model

A critical cellular energy sensor, AMPK (37), is regulated by the intracellular AMP to ATP ratio, with robust activation of AMPK induced by ATP deprivation (38). In the present study, AMPK phosphorylation increased markedly following 6-h DEX treatment (Fig. 5A and B). In addition, increased nuclear and decreased cytoplasmic levels of transcription factor protein FoxO3, a known master regulator of muscle atrophy-associated E3 ligases MuRF1 and atrogin-1 (39), were observed. These changes were reversed by S-Rg3 treatment, which decreased AMPK phosphorylation (Fig. 5A and B) and reduced FoxO3 nuclear translocation (Fig. 5C and D). Therefore, these data suggested that the AMPK/FoxO3 pathway is involved in the protective effect of S-Rg3 on mitochondrial dysfunction.


In spite of their beneficial anti-inflammatory and other clinically useful properties, GCs are known for numerous adverse myopathic effects (40). For example, high-dose synthetic GC DEX treatment has been observed to increase degradation and decrease synthesis of muscle protein that together led to reduced muscle mass and muscle fiber thinning (41). These myopathic effects appear to involve triggering of catabolic signals, including ubiquitin E3 ligases atrogin-1 and MuRF1 (42). As demonstrated in previous studies, such effects may be counteracted by ginsenoside Rg1 (22) and ginseng protein (43), which are reported to possess anti-atrophy effects, as well as by ginsenoside S-Rg3, which has been shown to prevent myotube atrophy induced by TNF-α (24). The present study demonstrated that S-Rg3 prevention of myotube atrophy was linked to decreases in atrogin-1 and MuRF1 expression levels that subsequently corresponded to observed increase in viability of DEX-injured C2C12 myotube cells. The present study also demonstrated that the in vitro experimental model of myotube atrophy employed could be used to evaluate drugs, such as S-Rg3, for effectiveness when used for treatment of muscle atrophy.

Muscle atrophy, a clinical disorder commonly associated with diabetes (44), chronic obstructive pulmonary disease (45) and other chronic diseases, negatively affects the quality of life of patients and promotes progression of pathological disease (46). A growing body of evidence suggests that mitochondrial dysfunction is a key player in muscle atrophy caused by disuse and disease (47). Meanwhile, a previous study demonstrated that treatment with DEX can cause serious impairment of mitochondrial function manifesting as mitochondrial loss, dysfunctional mitochondrial respiration and disordered mitochondrial morphology and distribution (48). To counter such adverse DEX-induced effects, therapies targeting mitochondrial processes to increase mitochondrial biogenesis and/or enhance mitochondrial respiration are sought for prevention or treatment of muscle atrophy (49). In the present study, C2C12 myotube mitochondrial dysfunction induced by 50 µM DEX administration was associated with decreased ATP level, mitochondrial respiration rate, mitochondrial membrane potential and mitochondrial complex II, III and V subunit protein levels. Notably, these pathological changes were prevented by S-Rg3 treatment.

Mitochondria function as the main energy-producing organelles in cells, with mitochondrial functional disturbances leading to insufficient energy supply and activation of intracellular signaling pathways that culminates in AMPK activation, apoptosis and/or autophagy (50,51). Notably, intracellular ATP deprivation triggers immediate activation of AMPK, an important energy sensor, that subsequently causes mitochondrial biogenesis (via PGC-1α phosphorylation) or autophagy (via ULK1 phosphorylation) (52). As demonstrated in previous studies, AMPK participates in muscle atrophy in two ways, by phosphorylating FoxO3 and thereby directly controlling its nuclear translocation (53) and by participating in AMPK/FoxO3 pathway signaling triggered by mitochondrial fission (54). Our previous work demonstrated that S-Rg3 protects DEX-induced muscle atrophy probably by promoting AKT/mTOR phosphorylation and inhibiting FoxO3 nuclear transcription (25). The present study explored the role of mitochondrial function in muscle atrophy. The results showed that S-Rg3 treatment of DEX-injured myotubes decreased phosphorylation of AMPK and subsequently prevented nuclear translocation of FoxO3, effectively alleviating mitochondrial dysfunction and preventing muscle atrophy induced by DEX in vitro.

The present study studied GC-induced atrophy using an in vitro muscle myotube model to gain insights into underlying mechanisms for S-Rg3 prevention of muscle atrophy. It was discovered that S-Rg3 treatment restored mitochondrial function and promoted recovery from DEX-induced muscle atrophy by inhibiting the AMPK/FoxO3 pathway activated by DEX.


Not applicable.


The present study was supported by the National Natural Science Foundation of China (grant nos. U19A2013, U20A20402 and 81973813), the National Key Research and Development Program of China (grant no. 2017YFC1702106), the Administration of Traditional Chinese Medicine of Jilin Province (grant no. 2020171) and the Science and Technology Development Plan of Changchun City (grant no. 18YJ013).

Availability of data and materials

The data used and/or analyzed during the current study are available from the corresponding author on reasonable request.

Authors' contributions

MW analyzed data and wrote the manuscript. RJ and JL performed the experiments and conducted the analysis of data. XX and GS performed the statistical analysis of the data. DZ and LS designed the study and were involved in drafting the manuscript and confirm the authenticity of all the raw data. All authors read and approved the final manuscript.

Ethics approval and consent to participate

Not applicable.

Patient consent for publication

Not applicable.

Competing interests

The authors declare that they have no competing interests.



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Wang M, Jiang R, Liu J, Xu X, Sun G, Zhao D and Sun L: 20(s)‑ginseonside‑Rg3 modulation of AMPK/FoxO3 signaling to attenuate mitochondrial dysfunction in a dexamethasone‑injured C2C12 myotube‑based model of skeletal atrophy <em>in vitro</em> Corrigendum in /10.3892/mmr.2021.12259. Mol Med Rep 23: 306, 2021
Wang, M., Jiang, R., Liu, J., Xu, X., Sun, G., Zhao, D., & Sun, L. (2021). 20(s)‑ginseonside‑Rg3 modulation of AMPK/FoxO3 signaling to attenuate mitochondrial dysfunction in a dexamethasone‑injured C2C12 myotube‑based model of skeletal atrophy <em>in vitro</em> Corrigendum in /10.3892/mmr.2021.12259. Molecular Medicine Reports, 23, 306.
Wang, M., Jiang, R., Liu, J., Xu, X., Sun, G., Zhao, D., Sun, L."20(s)‑ginseonside‑Rg3 modulation of AMPK/FoxO3 signaling to attenuate mitochondrial dysfunction in a dexamethasone‑injured C2C12 myotube‑based model of skeletal atrophy <em>in vitro</em> Corrigendum in /10.3892/mmr.2021.12259". Molecular Medicine Reports 23.5 (2021): 306.
Wang, M., Jiang, R., Liu, J., Xu, X., Sun, G., Zhao, D., Sun, L."20(s)‑ginseonside‑Rg3 modulation of AMPK/FoxO3 signaling to attenuate mitochondrial dysfunction in a dexamethasone‑injured C2C12 myotube‑based model of skeletal atrophy <em>in vitro</em> Corrigendum in /10.3892/mmr.2021.12259". Molecular Medicine Reports 23, no. 5 (2021): 306.