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. 2023 Aug 18;33(5):1233–1243. doi: 10.1007/s10068-023-01407-w

Ginsenosides Rh1, Rg2, and Rg3 ameliorate dexamethasone-induced muscle atrophy in C2C12 myotubes

Xiao Men 1, Xionggao Han 1, Se-Jeong Lee 1, Geon Oh 1, Ji-Hyun Im 1, Kwi Sik Bae 2, Geum-Su Seong 3, Im-Joung La 4, Do-Sang Lee 4, Sun-Il Choi 1,5,✉, Ok-Hwan Lee 1,✉
PMCID: PMC10909033  PMID: 38440685

Abstract

High doses or prolonged use of the exogenous synthetic glucocorticoid dexamethasone (Dex) can lead to muscle atrophy. In this study, the anti-atrophic effects of ginsenosides Rh1, Rg2, and Rg3 on Dex-induced C2C12 myotube atrophy were assessed by XTT, myotube diameter, fusion index, and western blot analysis. The XTT assay results showed that treatment with Rh1, Rg2, and Rg3 enhanced cell viability in Dex-injured C2C12 myotubes. Compared with the control group, the myotube diameter and fusion index were both reduced in Dex-treated cells, but treatment with Rh1, Rg2, and Rg3 increased these parameters. Furthermore, Rh1, Rg2, and Rg3 significantly downregulated the protein expression of FoxO3a, MuRF1, and Fbx32, while also upregulating mitochondrial biogenesis through the SIRT1/PGC-1α pathway. It also prevents myotube atrophy by regulating the IGF-1/Akt/ mTOR signaling pathway. These findings indicate that Rh1, Rg2, and Rg3 have great potential as useful agents for the prevention and treatment of muscle atrophy.

Keywords: Ginsenosides Rh1, Rg2, Rg3; C2C12; Dexamethasone; Muscle atrophy; Myoblast differentiation

Introduction

Muscle atrophy primarily refers to the dystrophy of skeletal muscle, which is characterized by the thinning or even disappearance of muscle fibers, resulting in a reduction in muscle volume, loss of muscle strength, and muscle dysfunction (Yeon et al., 2022). Normally, the metabolism of muscle protein is in a state of dynamic equilibrium. However, when the rate of protein degradation exceeds that of protein synthesis, there is a net loss of muscle mass, resulting in a decrease in muscle strength and function, and the onset of muscle atrophy (Kim et al., 2020; Oh et al., 2022). The causes of muscle atrophy are complex. Prolonged immobilization or lack of physical activity can result in muscle atrophy, such as spinal cord injury, prolonged bed rest, and leg casts due to broken bones (Rudrappa et al., 2016). Insufficient protein intake or an imbalance of essential nutrients can also result in gradual muscle loss and muscle atrophy (Rennie et al., 2010). Additionally, conditions such as heart failure, cancer, and kidney failure can also lead to muscle atrophy (Wijaya et al., 2022). According to reports, glucocorticoids (GCs) are a crucial class of regulatory molecules in the body. They can enter cells directly and bind to the glucocorticoid receptor within the cell. Once the hormone binds to the receptor, the newly formed complex translocates into the nucleus and binds to the glucocorticoid response element located in the promoter region of the target gene, thereby regulating gene expression (Kadmiel and Cidlowski, 2013; Ricketson et al., 2007). For the muscular system, GCs can cause muscle wasting by inhibiting muscle protein synthesis and accelerating muscle protein degradation (Schakman et al., 2013). The degradation process of intracellular muscle proteins is mainly attributed to the ubiquitin-proteasome system (Singh et al., 2021). GCs induce an increase in proteolysis by upregulating two specific ubiquitin ligases, F-box protein (Fbx32) and muscle ring-finger 1 (MuRF1), which are associated with muscle atrophy (Liu et al., 2016). Concurrently, GCs promote the expression of the forkhead box O3a (FoxO3a) transcription factor, a major mediator of muscle atrophy, which can activate MuRF1 and Fbx32 to further increase muscle proteolysis (Peris-Moreno et al., 2020). In contrast to the degradation process of muscle protein, the insulin-like growth factor 1 (IGF-1)/protein kinase B (Akt)/mechanistic target of rapamycin (mTOR) muscle protein synthesis signaling pathway mitigates muscle atrophy caused by the ubiquitin-proteasome system. IGF-1 regulates muscle cell differentiation through the PI3K-Akt signaling pathway. Akt activates mTOR to increase muscle protein synthesis and inhibits the expression of FoxO3a to prevent muscle atrophy (Rommel et al., 2001). The IGF-1/Akt/mTOR signaling pathway plays a crucial role in regulating muscle protein synthesis and hypertrophy (Glass, 2003; Santo et al., 2013). Mitochondria are crucial organelles that supply energy for muscle function. When these organelles become dysfunctional, less energy is produced and an excess of reactive oxygen species are generated, leading to muscle atrophy, weakness, and reduced endurance (Yin et al., 2022). Silent information regulator 1 (SIRT1) can sense the state of energy metabolism at the cellular level and play an important role in regulating the energy homeostasis of the body. During the transcription process, SIRT1 can deacetylate the downstream gene peroxisome proliferator-activated receptor gamma coactivator-1α (PGC-1α) involved in energy metabolism. PGC-1α is the main regulator of mitochondrial biogenesis and activity, and the activation of PGC-1α can promote the biosynthesis of mitochondria (Jäger et al., 2007; Yuan et al., 2016). Therefore, the regulatory mechanism of the SIRT1/PGC-1α signaling pathway can be regarded as a potential target for the treatment of muscle atrophy (Li et al., 2022).

Ginsenosides are the main pharmacological active ingredients in ginseng, which are mainly divided into protopanaxatriol of Rh1, Rg2, Rg1, and Re, and protopanaxadiol of Rh2, Rg3, Rd, Rc, Rb3, Rb2, and Rb1 (Kim et al., 2017). Modern pharmacological studies have demonstrated that ginsenosides possess various pharmacological effects, including anti-tumor, anti-aging, and regulatory effects on the central nervous system (Choi, 2008). There have been reports indicating that ginsenoside Rg2 exerts an antidepressant effect by activating the brain-derived neurotrophic factor signaling pathway in the hippocampus (Ren et al., 2017). Ginsenoside Rh1 can inhibit the invasion and migration of glioma cells by blocking PI3K/Akt and MAPK signaling pathways (Jung et al., 2013). Ginsenoside Rg3 can inhibit adipocyte differentiation and adipogenesis (Zhang et al., 2017). Wang et al. demonstrated through in vitro experiments that ginsenoside Rg3 exhibited a significant inhibitory effect on the proliferation of breast cancer cells (Wang et al., 2008). In addition, ginsenoside Rg3 can treat TNF-α-induced muscle atrophy by activating the Akt/mTOR signaling pathway (Lee et al., 2019). However, the therapeutic effects of ginsenosides Rh1, Rg2, and Rg3 on GCs-induced myotube atrophy and their underlying mechanisms have not been thoroughly studied. Therefore, in this study, we constructed a muscle atrophy model by inducing C2C12 myotube cells with dexamethasone (Dex) to investigate the effects of each ginsenoside (Rh1, Rg2, and Rg3) on myocyte differentiation, myotube growth, and mitochondrial biogenesis in the state of myotube atrophy, and analyzed the molecular mechanism of Rh1, Rg2, and Rg3 against Dex-induced muscle atrophy. Such a study design allows for the assessment of the individual activity characteristics of each ginsenoside and provides an in-depth exploration of their potential mechanisms of action during muscle atrophy.

Materials and methods

Chemicals and reagents

Ginsenoside Rg3 and Dex were purchased from Sigma-Aldrich Co. (St. Louis, MO, USA). Ginsenoside Rh1 and Rg2 were obtained from Med Chem Express (MCE, NJ, USA). Fetal bovine serum (FBS), phosphate-buffered saline (PBS), Dulbecco’s modified Eagle’s medium (DMEM), horse serum (HS), penicillin-streptomycin (P/S), and trypsin-ethylenediaminetetraacetic acid (EDTA) were obtained from Gibco (Grand Is-land, NY, USA).

Cell culture and differentiation

The myoblast C2C12 cells were purchased from the American Type Culture Collection (ATCC, Manassas, VA, USA). Cells were seeded into clear flat bottom tissue culture-treated 24-well plates, 96-well plates, and 100-mm Petri dishes (Corning) in DMEM containing 1% P/S, 10% FBS, and 3.7 g/L sodium bicarbonate at 37 °C in humidified atmosphere 5% CO2 until the cells reached 80–90% confluence. After the cells reached confluence, the medium was replaced with DMEM containing 2% HS and 1% P/S to induce myoblasts to differentiate into myotubes. During the induction of cell differentiation, the medium was changed every 2 days, Myoblasts were successfully induced to differentiate into C2C12 myotubes usually on day 5–6. For the study of the therapeutic effect of ginsenoside Rh1, Rg2, and Rg3 on Dex-induced muscle atrophy model. After the cells were fully differentiated, the cells were treated with 0.5, 1, 2 µM of ginsenosides Rh1, Rg2, and Rg3 and Dex (200 µM) for 24 h, and then the cells were then harvested for subsequent experiments.

Cell viability assay

The effect of Dex and/or ginsenosides Rh1, Rg2, and Rg3 on cell viability was detected using 2,3-bis(2-methoxy-4-nitro-5-sulfophenyl)-2 H-tetrazolium-5-carboxanilide (XTT) assay. Briefly, myoblasts were seeded in 96-well plates (1 × 104 cell/well) and cultured until the cells were induced to differentiate into myotubes. Cells were then treated with different concentrations (0.5, 1, 2 µM) of ginsenoside Rh1, Rg2, and Rg3 for 24 h. Subsequently, XTT and N-methyl dibenzopyrazine methyl sulfate (PMS) mixed solution (XTT:PMS = 50:1 (V:V)) was added to each well and incubated at 37 °C for 4 h. The absorbance was measured at 450 and 690 nm (Molecular Devices, Sunnyvale, CA, USA).

Jenner–Giemsa staining

To evaluate the effect of ginsenosides Rh1, Rg2, and Rg3 on Dex-induced atrophy of C2C12 myotube cells, cells were stained with Jenner–Giemsa to allow quantitative measurements of myotube structure and nuclei. Briefly, 24 h after cells were treated with experimental samples, the medium was aspirated and washed twice with 1×PBS. The cells were then fixed with 100% methanol for 5 min and dried at room temperature for 10 min. The dried cells were stained with Jenner staining solution for 5 min and washed twice with deionized water (DW). The cells were then stained with Giemsa staining solution for 10 min, washed twice with DW again, and observed and photographed under an inverted microscope (CKX41SF, Tokyo, Japan). Jenner staining solution is to dissolve 1 g Jenner’s stain (Sigma–Aldrich) in 400 mL methanol, and mix it with DW at a volume ratio of 1:1 (V:V), and then mixing the mixed solution with 1mM sodium phosphate buffer (pH 5.6) at a ratio of 1:3 (V:V). Giemsa staining solution was prepared by mixing Giemsa stain (Sigma–Aldrich) with 1mM sodium phosphate buffer (pH 5.6) at a volume ratio of 1:20 (V:V). The Jenner and Giemsa staining solutions used in the experiments were prepared from the same batch.

Measurement of myotube diameters

Myotube diameters were measured by using ImageJ software. After the myotube cells were stained, use an inverted microscope (200 × magnification) to take pictures of five random areas in each well of the 24-well plate, and measure the diameter of the myotube cells containing at least three nuclei in the picture.

Measurement of myotube fusion index

To determine the fusion index, an inverted microscope was used to take pictures of 5 random areas in each well of a 24-well plate. The number of nuclei in each myotube cell containing at least three nuclei and total number of nuclei were calculated using the ImageJ software, and the fusion index of the cell was calculated using the following formula:

Fusionindex(%)=numberofnucleiinmyotubes/totalnumberofnuclei×100

Western blot analysis

After washing the cells twice with 1×PBS, the cells were lysed using a protein lysis buffer and centrifuged at 4 °C and 12,000×g for 15 min. After centrifugation, the supernatant was collected, and the protein concentration was determined using a Bio-Rad protein assay kit. Samples were then separated into equal amounts of protein on 10% sodium dodecyl sulfate polyacrylamide gel electrophoresis. After electrophoresis, the proteins were transferred to a polyvinylidene difluoride membrane by wet transfer method. The membrane was blocked with 5% skim milk for 1 h at room temperature. After blocking, the membrane was washed three times with 1× TBST, and then incubated with primary antibodies [IGF-1 and Fbx32 (1:1000; Abcam, Cambridge, UK), β-actin, SIRT1, Akt, p- Akt, mTOR, p- mTOR, and FoxO3a (1:1000; Cell Signaling Technology, Danvers, MA, USA), PGC-1α and MuRF1 (1:1000; Santa Cruz Biotechnology, Santa Cruz, CA, USA)] overnight at 4 °C. After the primary antibody incubation, the membrane was washed three times with 1×TBST for 10 min each, and then incubated with anti-rabbit or anti-mouse IgG, HRP-linked secondary antibodies (1:2000; Cell Signaling Technology, Danvers, MA, USA) for 1 h at room temperature. The membrane was then washed three times with 1×TBST for 10 min each. Then, after developing with the ECL reagent, the images of protein bands were detected using ChemiDoc imaging system (Bio-Rad Laboratories, Inc., Hercules, CA, USA).

Statistical analysis

All experimental results are expressed as the mean ± standard deviation (SD). Statistical analysis was performed on each test group by one-way analysis of variance and Duncan’s multiple tests using SPSS software (version 24.0; SPSS Inc., Chicago, IL, USA). A p-value < 0.05 was considered significant.

Results and discussion

Effect of ginsenosides Rh1, Rg2, and Rg3 on cell viability in Dex-induced C2C12 myotube cells

Firstly, we assessed the effects of different concentrations of Dex (25, 50, 100, and 200 µM) and ginsenosides Rh1, Rg2, and Rg3 (0.5, 1, and 2 µM) on the viability of C2C12 myotubes by XTT assay (Fig. 1A–D). As shown in Fig. 1A, the Dex treatment group significantly reduced the cell activity of C2C12 myotubes in a dose-dependent manner compared to the control group (Con). At a concentration of 200 µM, the cell activity decreased to 75.52 ± 9.27%. All concentrations of Rh1, Rg2, and Rg3 in the experimental group showed no cytotoxic effects on C2C12 cells. (Fig. 1B–D). Based on the inhibitory effect of Dex (200 µM) on the viability of C2C12 myotube cells, we further evaluated the effects of Rh1, Rg2, and Rg3 on Dex-induced cytotoxicity. As shown in Fig. 1E–G, co-treatment of Rh1, Rg2, and Rg3 with Dex significantly increased the cell viability of C2C12 myotube cells compared to the Dex group. These results suggest that Rh1, Rg2, and Rg3 can effectively prevent the reduction of myotube cell viability induced by Dex.

Fig. 1.

Fig. 1

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Effect of ginsenosides Rh1, Rg2, and Rg3 on cell viability in Dex-induced C2C12 myotubes. A Cell viability of C2C12 myotubes at different concentrations (25, 50, 100, and 200 µM) of Dex. The cell viability of C2C12 myotubes was assessed in the presence of ginsenoside Rh1 (B), Rg2 (C), and Rg3 (D) at concentrations of 0, 0.5, 1, and 2 µM. Cell viability of C2C12 myotubes was co-treated with 0.5, 1, and 2 µM Rh1 (E), Rg2 (F), and Rg3 (G) along with 200 µM Dex. Data values are presented as mean ± SD. Different letters on the graphs indicate significant differences (p < 0.05)

Effects of ginsenosides Rh1, Rg2, and Rg3 on myotube diameter and fusion index in Dex-induced C2C12 myotubes

To determine the effects of Rh1, Rg2, and Rg3 on Dex-induced C2C12 myotube atrophy, we measured changes in fusion index and myotube diameter. The C2C12 myotubes of fully differentiated cells stained with Jenner-Giemsa after 24 h treatment with Rh1, Rg2, Rg3, and Dex, respectively, are shown in Fig. 2A. In the Dex treatment group, myotube diameter was significantly reduced to 28.30% compared to the Con. However, when Rh1, Rg2, and Rg3 were co-treated with Dex, the Dex-induced reduction in C2C12 myotube diameter was significantly inhibited. Especially in the 2 µM Rh1, Rg2, and Rg3 treatment groups, the diameter of atrophied myotubes induced by Dex was significantly increased to 83.40%, 74.97%, and 87.44% respectively (Fig. 2B–D). The fusion index was calculated as the percentage of the number of nuclei incorporated into the myotubes to the total number of nuclei (Veliça and Bunce, 2011). The experimental results showed that compared to the Con, treatment of myotubes with 200 µM Dex for 24 h significantly decreased the fusion index. However, treatment with 0.5, 1, and 2 µM Rh1, Rg2, or Rg3 significantly prevented the Dex-induced reduction in fusion index (Fig. 2E–G). In particular, the fusion index of myotubes was significantly increased to 65.41 ± 6.77%, 63.50 ± 7.06%, and 62.33 ± 5.49% with the treatment of 2 µM Rh1, Rg2, and Rg3, respectively. These results indicate at the morphological level that Rh1, Rg2, and Rg3 can significantly promote the differentiation of C2C12 myoblasts in the Dex-induced muscle atrophy model and have a significant therapeutic effect on Dex-induced cell atrophy.

Fig. 2.

Fig. 2

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Effect of Rh1, Rg2, and Rg3 on diameter and fusion index in Dex-induced C2C12 myotubes. A After Jenner-Giemsa staining, representative photographs of cells in different experimental groups (Con, Dex, Dex + Rh1/Rg2/Rg3) observed under the microscope (× 200); red arrows indicate nuclei. The diameter of C2C12 myotubes was evaluated upon treatment with ginsenoside Rh1 (B), Rg2 (C), and Rg3 (D) at concentrations of 0, 0.5, 1, and 2 µM in combination with 200 µM Dex. The fusion index of C2C12 myotubes was evaluated upon treatment with ginsenoside Rh1 (E), Rg2 (F), and Rg3 (G) at concentrations of 0, 0.5, 1, and 2 µM in combination with 200 µM Dex. Quantization was performed using the Image J program. Data values are presented as mean ± SD. Different letters on the graphs indicate significant differences (p < 0.05)

Ginsenosides Rh1, Rg2, and Rg3 attenuated Dex-induced mitochondrial energy metabolic function disorder by activating the SIRT1/PGC-1α signaling pathway

It is known that SIRT1 can sense the energy metabolic status at the cellular level (Yang et al., 2022). The activation of SIRT1 can regulate the activity of the PGC-1α protein, which is involved in energy metabolism, thereby improving the biosynthesis level of mitochondria and the oxidation capacity of muscle fibers. This plays an important role in improving exercise ability and preventing muscle atrophy (Gouspillou et al., 2014; Yeon et al., 2022). Therefore, we further investigated the effect of Rh1, Rg2, and Rg3 on the SIRT1/PGC-1α signaling pathway in Dex-induced myotube cells. Our results showed that the protein expressions of SIRT1 and PGC-1α were significantly decreased in all Dex-treated groups compared to Con. Rh1, Rg2, and Rg3 experimental groups all dose-dependently increased the expression of SIRT1 and PGC-1α protein. Among them, in the 2 µM Rh1 and 2 µM Rg2 experimental groups, the expressions of SIRT1 and PGC-1α proteins were significantly restored to normal levels (Fig. 3A, B, D, and E). In addition, the protein expressions of SIRT1 and PGC-1α were significantly enhanced in the 2 µM Rg3 experimental group, and their protein expression levels were even higher than the Con (Fig. 3C and F). These results indicated that Rh1, Rg2, and Rg3 can increase the expression levels of SIRT1 and PGC-1α proteins to ameliorate Dex-induced C2C12 myotube atrophy.

Fig. 3.

Fig. 3

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Effects of Rh1, Rg2, and Rg3 on SIRT1/PGC-1α protein expression in Dex-induced c2c12 myotube cells. Rh1 (A), Rg2 (B), and Rg3 (C) promoted the expression of SIRT1 protein. Rh1 (D), Rg2 (E), and Rg3 (F) promoted the expression of PGC-1α protein. SIRT1 and PGC-1α proteins were identified by Western blot analysis and normalized to β-actin. Data values are presented as mean ± SD. Different letters on the graphs indicate significant differences (p < 0.05)

Effects of ginsenosides Rh1, Rg2, and Rg3 on muscle protein synthesis-related pathways in Dex-induced C2C12 myotube cells

IGF-1 plays a crucial role in the synthesis of skeletal muscle protein and muscle formation. It activates the mTOR/Akt signaling pathway, promoting myoblast differentiation, protein synthesis, and enhancing muscle hypertrophy and growth. As a result, upregulating the IGF-1/Akt/mTOR signaling pathway is important in preventing and treating muscle atrophy (Yoshida and Delafontaine, 2020). To clarify the potential mechanism of Rh1, Rg2, and Rg3 on the muscle protein synthesis pathway in Dex-induced c2c12 myotube cells, we measured the expression levels of IGF-1, Akt (p-Akt), and mTOR (p-mTOR) proteins (Fig. 4). The protein expressions of phosphorylated Akt and mTOR (p-Akt and p-mTOR), as well as IGF, were significantly decreased in all Dex-treated groups compared to the Con. However, Rh1, Rg2, and Rg3 showed a concentration-dependent increase in the expression of IGF-1, p-Akt, and p-mTOR proteins, and these protein expression levels returned to normal significantly in the 2 µM Rh1 and Rg2 treatment group. Surprisingly, in the 2 µM Rg3 treatment group, the protein expression of IGF-1 and p-Akt returned to normal, while the expression level of p-mTOR was higher than that of the Con (Fig. 4A–C). Taken together, these results suggest that Rh1, Rg2, and Rg3 can effectively treat Dex-induced muscle atrophy by increasing the expression of related muscle differentiation proteins through activation of the IGF-1/Akt/mTOR signaling pathway.

Fig. 4.

Fig. 4

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Effects of Rh1, Rg2, and Rg3 on IGF-1/Akt/mTOR protein expression in Dex-induced c2c12 myotube cells. A Rh1, Rg2, and Rg3 promoted the expression of IGF-1 protein. B Rh1, Rg2, and Rg3 promoted the expression of p-Akt/Akt protein. C Rh1, Rg2, and Rg3 promoted the expression of p-mTOR/mTOR protein. IGF-1, p-Akt, Akt, p-mTOR, and mTOR proteins were identified by Western blot analysis and normalized to β-actin. Data values are presented as mean ± SD. Different letters on the graphs indicate significant differences (p < 0.05)

Effects of ginsenosides Rh1, Rg2, and Rg3 on muscle atrophy and muscle protein degradation related pathways in Dex-induced C2C12 muscle cells

The maintenance of muscle mass is dependent on the balance between two processes: the rate of protein synthesis and the rate of protein degradation (Romanello and Sandri, 2021). FoxO3a plays a central mediating role in the process of muscle synthesis and degradation, and is a key protein involved in muscle protein degradation. It can promote the expression of Fbx32 and MuRF1, which in turn block muscle synthesis and increase muscle protein breakdown (Ghafouri-Fard et al., 2021). Next, to further analyze the effects of Rh1, Rg2, and Rg3 on key proteins involved in Dex-induced myotube atrophy, we investigated the protein expression levels of FoxO3a, Fbx32, and MuRF1. As shown in Fig. 5A–C, the protein expression levels of FoxO3a, Fbx32, and MuRF1 were significantly increased in Dex-treated C2C12 myotubes compared to the Con. When Dex-induced cells were treated with Rh1, Rg2, and Rg3, the results demonstrated a significant reduction in the protein expression of FoxO3a, Fbx32, and MuRF1. Furthermore, the MuRF1 protein expression level was observed to be lower than the Con in both the 2 µM Rh1 and Rg2 treatment groups. In the 2 µM Rg3 treatment group, the protein expression levels of FoxO3a and MuRF1 returned to normal levels. Taken together, these results suggest that Rh1, Rg2, and Rg3 are effective in inhibiting the Dex-induced increase in muscular dystrophy protein expression. Several studies have demonstrated that the expression of Fbx32 and MuRF1 proteins increases with age, thereby contributing to the development of sarcopenia (Mankhong et al., 2020). From another perspective, these findings suggest that these ginsenosides may have potential applications in indirectly influencing muscle loss conditions associated with aging.

Fig. 5.

Fig. 5

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Effects of Rh1, Rg2, and Rg3 on FoxO3a, Fbx32, and MuRF1 protein expression in Dex-induced C2C12 myotube cells. A Rh1, Rg2, and Rg3 promoted the expression of FoxO3a protein. B Rh1, Rg2, and Rg3 promoted the expression of Fbx32 protein. C Rh1, Rg2, and Rg3 promoted the expression of MuRF1 protein. FoxO3a, Fbx32, and MuRF1 proteins were identified by Western blot analysis and normalized to β-actin. Data values are presented as mean ± SD. Different letters on the graphs indicate significant differences (p < 0.05)

In conclusion, our results suggest that Rh1, Rg2, and Rg3 have the potential to prevent Dex-induced mitochondrial dysfunction by promoting the expression of energy metabolism-related proteins (SIRT1 and PGC-1α) and by increasing the expression of myoblast differentiation and myogenesis-related proteins (IGF-1/Akt/mTOR). Furthermore, Rh1, Rg2, and Rg3 inhibit ubiquitin ligases (FoxO3a, Fbx32, and MuRF1) in myotube cells, which effectively treats and prevents Dex-induced muscle atrophy (Fig. 6). This study provides a foundation for the development of therapeutic drugs to prevent and treat muscle atrophy in the future. Moreover, it suggests the potential application of ginsenosides in indirectly addressing muscle loss conditions associated with aging or nutrition, thus providing valuable insights into the therapeutic prospects of ginsenosides for muscle atrophy.

Fig. 6.

Fig. 6

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Schematic diagram of the effects of ginsenosides Rh1, Rg2, and Rg3 in Dex-induced C2C12 myotube atrophy

Acknowledgements

This research was funded by the Ministry of SMEs and Startups (Grant No. S3241314), the Basic Science Research Program (NRF-2021R1A6A1A03044242) through the National Research Foundation of Korea (NRF) funded by the Ministry of Education, the BK21 FOUR (Fostering Outstanding Universities for Research) funded by the Ministry of Education (MOE, Korea) and National Research Foundation of Korea (NRF) (Grant No. 4299990913942).

Declarations

Conflict of interest

None of the authors of this study has any financial interest or conflict with industries or parties.

Footnotes

Publisher’s Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Contributor Information

Sun-Il Choi, Email: docgotack89@hanmail.net.

Ok-Hwan Lee, Email: loh99@kangwon.ac.kr.

References

  1. Choi KT. Botanical characteristics, pharmacological effects and medicinal components of Korean Panax ginseng CA Meyer. Acta Pharmacologica Sinica. 2008;29:1109–1118. doi: 10.1111/j.1745-7254.2008.00869.x. [DOI] [PubMed] [Google Scholar]
  2. Ghafouri-Fard S, Abak A, Khademi S, Shoorei H, Bahroudi Z, Taheri M, Dilmaghani NA. Functional roles of non-coding RNAs in atrophy. Biomedicine & Pharmacotherapy. 2021;141:111820. doi: 10.1016/j.biopha.2021.111820. [DOI] [PubMed] [Google Scholar]
  3. Glass DJ. Signalling pathways that mediate skeletal muscle hypertrophy and atrophy. Nature Cell Biology. 2003;5:87–90. doi: 10.1038/ncb0203-87. [DOI] [PubMed] [Google Scholar]
  4. Gouspillou G, Sgarioto N, Norris B, Barbat-Artigas S, Aubertin-Leheudre M, Morais JA, Burelle Y, Taivassalo T, Hepple RT. The relationship between muscle fiber type-specific PGC-1α content and mitochondrial content varies between rodent models and humans. PloS ONE. 2014;9:e103044. doi: 10.1371/journal.pone.0103044. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Jäger S, Handschin C, St.-Pierre J, Spiegelman BM (2007)AMP-activated protein kinase (AMPK) action in skeletal muscle via direct phosphorylation of PGC-1α. Proceedings of the national academy of sciences. 104: 12017-12022  [DOI] [PMC free article] [PubMed]
  6. Jung JS, Ahn JH, Le TK, Kim DH, Kim HS. Protopanaxatriol ginsenoside Rh1 inhibits the expression of matrix metalloproteinases and the in vitro invasion/migration of human astroglioma cells. Neurochemistry International. 2013;63:80–86. doi: 10.1016/j.neuint.2013.05.002. [DOI] [PubMed] [Google Scholar]
  7. Kadmiel M, Cidlowski JA. Glucocorticoid receptor signaling in health and disease. Trends in Pharmacological Sciences. 2013;34:518–530. doi: 10.1016/j.tips.2013.07.003. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Kim JH, Yi YS, Kim MY, Cho JY. Role of ginsenosides, the main active components of Panax ginseng, in inflammatory responses and diseases. Journal of Ginseng Research. 2017;41:435–443. doi: 10.1016/j.jgr.2016.08.004. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Kim IY, Park S, Jang J, Wolfe RR. Understanding muscle protein dynamics: technical considerations for advancing sarcopenia research. Annals of Geriatric Medicine and Research. 2020;24:157. doi: 10.4235/agmr.20.0041. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Lee SJ, Bae JH, Lee H, Lee H, Park J, Kang JS, Bae GU. Ginsenoside Rg3 upregulates myotube formation and mitochondrial function, thereby protecting myotube atrophy induced by tumor necrosis factor-alpha. Journal of Ethnopharmacology. 2019;242:112054. doi: 10.1016/j.jep.2019.112054. [DOI] [PubMed] [Google Scholar]
  11. Li M, Wu H, Huang KH, Hu JH, Liu HS. Paeoniflorin ameliorates skeletal muscle atrophy in chronic kidney disease via AMPK/SIRT1/PGC-1α-mediated oxidative stress and mitochondrial dysfunction. Frontiers in pharmacology. 2022;13:662. doi: 10.3389/fphar.2022.859723. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Liu J, Peng Y, Wang X, Fan Y, Qin C, Shi LE, Tang Y, Li H, Long J, Liu J. Mitochondrial dysfunction launches dexamethasone-induced skeletal muscle atrophy via AMPK/FOXO3 signaling. Molecular Pharmaceutics. 2016;13:73–84. doi: 10.1021/acs.molpharmaceut.5b00516. [DOI] [PubMed] [Google Scholar]
  13. Mankhong S, Kim S, Moon S, Kwak HB, Park DH, Kang JH. Experimental models of sarcopenia: bridging molecular mechanism and therapeutic strategy. Cells. 2020;9:1385. doi: 10.3390/cells9061385. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Oh HJ, Jin H, Kim BY, Lee OH, Lee BY. A Combined Angelica gigas and Artemisia dracunculus Extract Prevents Dexamethasone-Induced Muscle Atrophy in Mice through the Akt/mTOR/FoxO3a Signaling Pathway. Cells. 2022;11:3245. doi: 10.3390/cells11203245. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Peris-Moreno D, Taillandier D, Polge C. MuRF1/TRIM63, master regulator of muscle mass. International Journal of Molecular Sciences. 2020;21:6663. doi: 10.3390/ijms21186663. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Ren Y, Wang JL, Zhang X, Wang H, Ye Y, Song L, Wang YJ, Tu MJ, Wang WW, Yang L, Jiang B. Antidepressant-like effects of ginsenoside Rg2 in a chronic mild stress model of depression. Brain Research Bulletin. 2017;134:211–219. doi: 10.1016/j.brainresbull.2017.08.009. [DOI] [PubMed] [Google Scholar]
  17. Rennie MJ, Selby A, Atherton P, Smith K, Kumar V, Glover EL, Philips SM. Facts, noise and wishful thinking: muscle protein turnover in aging and human disuse atrophy. Scandinavian Journal of Medicine & Science in Sports. 2010;20:5–9. doi: 10.1111/j.1600-0838.2009.00967.x. [DOI] [PubMed] [Google Scholar]
  18. Ricketson D, Hostick U, Fang L, Yamamoto KR, Darimont BD. A conformational switch in the ligand-binding domain regulates the dependence of the glucocorticoid receptor on Hsp90. Journal of Molecular Biology. 2007;368:729–741. doi: 10.1016/j.jmb.2007.02.057. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Romanello V, Sandri M. The connection between the dynamic remodeling of the mitochondrial network and the regulation of muscle mass. Cellular and Molecular Life Sciences. 2021;78:1305–1328. doi: 10.1007/s00018-020-03662-0. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Rommel C, Bodine SC, Clarke BA, Rossman R, Nunez L, Stitt TN, Yancopoulos GD, Glass DJ. Mediation of IGF-1-induced skeletal myotube hypertrophy by PI (3) K/Akt/mTOR and PI (3) K/Akt/GSK3 pathways. Nature Cell Biology. 2001;3:1009–1013. doi: 10.1038/ncb1101-1009. [DOI] [PubMed] [Google Scholar]
  21. Rudrappa SS, Wilkinson DJ, Greenhaff PL, Smith K, Idris I, Atherton PJ. Human skeletal muscle disuse atrophy: effects on muscle protein synthesis, breakdown, and insulin resistance—a qualitative review. Frontiers in Physiology. 2016;7:361. doi: 10.3389/fphys.2016.00361. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Santo EE, Stroeken P, Sluis PV, Koster J, Versteeg R, Westerhout EM. FOXO3a Is a major target of inactivation by PI3K/AKT signaling in aggressive neuroblastomatumor suppressive role of FOXO3a in neuroblastoma. Cancer Research. 2013;73:2189–2198. doi: 10.1158/0008-5472.CAN-12-3767. [DOI] [PubMed] [Google Scholar]
  23. Schakman O, Kalista S, Barbé C, Loumaye A, Thissen JP. Glucocorticoid-induced skeletal muscle atrophy. The International Journal of Biochemistry & Cell Biology. 2013;45:2163–2172. doi: 10.1016/j.biocel.2013.05.036. [DOI] [PubMed] [Google Scholar]
  24. Singh A, Phogat J, Yadav A, Dabur R. The dependency of autophagy and ubiquitin proteasome system during skeletal muscle atrophy. Biophysical Reviews. 2021;13:203–219. doi: 10.1007/s12551-021-00789-7. [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Veliça P, Bunce CM. A quick, simple and unbiased method to quantify C2C12 myogenic differentiation. Muscle & Nerve. 2011;44:366–370. doi: 10.1002/mus.22056. [DOI] [PubMed] [Google Scholar]
  26. Wang CZ, Aung HH, Zhang B, Sun S, Li XL, He H, Xie JT, He TC, Du W, Yuan CS. Chemopreventive effects of heat-processed Panax quinquefolius root on human breast cancer cells. Anticancer Research. 2008;28:2545–2551. [PMC free article] [PubMed] [Google Scholar]
  27. Wijaya YT, Setiawan T, Sari IN, Park K, Lee CH, Cho KW, Lee YK, Kim JY, Yoon JK, Lee SH, Kwon HY. Ginsenoside Rd ameliorates muscle wasting by suppressing the signal transducer and activator of transcription 3 pathway. Journal of Cachexia, Sarcopenia and Muscle. 2022;13:3149–3162. doi: 10.1002/jcsm.13084. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Yang M, Yang H, Yu KH, Gao JH, Zheng HS. Regulation of yak longissimus lumborum energy metabolism and tenderness by the AMPK/SIRT1 signaling pathways during postmortem storage. Plos one. 2022;17:e0277410. doi: 10.1371/journal.pone.0277410. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Yeon M, Choi H, Chun KH, Lee JH, Jun HS. Gomisin G improves muscle strength by enhancing mitochondrial biogenesis and function in disuse muscle atrophic mice. Biomedicine & Pharmacotherapy. 2022;153:113406. doi: 10.1016/j.biopha.2022.113406. [DOI] [PubMed] [Google Scholar]
  30. Yin Z, Tian Y, Cai Y, Qi L, Yuan C, Liu J, Xu T. Mitochondrial activity as an indicator of scallop (Mizuhopecten yessoensis) adductor muscle in early cold storage. Journal of Food Science. 2022;87:206–215. doi: 10.1111/1750-3841.16007. [DOI] [PubMed] [Google Scholar]
  31. Yoshida T, Delafontaine P. Mechanisms of IGF-1-mediated regulation of skeletal muscle hypertrophy and atrophy. Cells. 2020;9:1970. doi: 10.3390/cells9091970. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Yuan Y, Cruzat VF, Newsholme P, Cheng J, Chen Y, Lu Y. Regulation of SIRT1 in aging: roles in mitochondrial function and biogenesis. Mechanisms of Ageing and Development. 2016;155:10–21. doi: 10.1016/j.mad.2016.02.003. [DOI] [PubMed] [Google Scholar]
  33. Zhang L, Zhang L, Wang X, Si H. Anti-adipogenic effects and mechanisms of ginsenoside Rg3 in pre-adipocytes and obese mice. Frontiers in Pharmacology. 2017;8:113. doi: 10.3389/fphar.2017.00113. [DOI] [PMC free article] [PubMed] [Google Scholar]

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