Danshensu sodium salt alleviates muscle atrophy via CaMKII‐PGC1α‐FoxO3a signaling pathway in D‐galactose‐induced models

Abstract

Sarcopenia is an age‐related muscle atrophy syndrome characterized by the loss of muscle strength and mass. Although many agents have been used to treat sarcopenia, there are no successful treatments to date. In this study, we identified Danshensu sodium salt (DSS) as a substantial suppressive agent of muscle atrophy. We used a D‐galactose (DG)‐induced aging‐acceleration model, both in vivo and in vitro, to confirm the effect of DSS on sarcopenia. DSS inhibits the expression of muscle atrophy‐related factors (MuRF1, MAFbx, myostatin, and FoxO3a) in DG‐induced mouse C2C12 and human skeletal muscle cells. Additionally, DSS restored the diameter of reduced C2C12 myotubes. Next, we demonstrated that DSS stimulates AMPK and PGC1α through CaMKII. DSS inhibits the translocation of FoxO3a into the nucleus, thus inhibiting muscle atrophy in a calcium‐dependent manner. DSS initiated the protein–protein interaction between FoxO3a and PGC1α. The reduction of the PGC1α‐FoxO3a interaction by DG was restored by DSS. Also, DSS suppressed increased intracellular reactive oxygen species (ROS) by DG. In animal models, DSS administration improved mouse muscle mass and physical performance (grip strength and hanging test) under DG‐induced accelerated aging conditions. These findings demonstrated that DSS attenuates muscle atrophy by inhibiting the expression of muscle atrophy‐related factors. Therefore, DSS may be a potential therapeutic agent for the treatment of sarcopenia.

Danshensu sodium salt, an analogue of Danshensu, downregulates the expression of muscle atrophy‐related genes via the CaMKII‐PGCα‐FoxO3a signaling pathway, thereby enhancing muscle function.

Abbreviations

AMPK

AMP‐activated protein kinase

CaMKII

Ca²⁺/calmodulin‐dependent protein kinase II

DG

D‐galactose

DM

differentiation medium

DS

Danshensu

DSS

Danshensu sodium salt

FoxO3a

Forkhead box O3

GCM

gastrocnemius

GTT

glucose tolerance test

LKB1

liver kinase B1

MAFbx

muscle atrophy F‐box

MuRF1

muscle RING‐finger protein 1

PGC1α

peroxisome proliferator‐activated receptor gamma coactivator 1‐alpha

QA

quadriceps

TA

tibialis anterior

1. INTRODUCTION

Danshensu (DS), also known as salvianic acid A ((R)‐3‐(3,4‐Dihydroxyphenyl)‐2‐hydroxypropanoic acid), is a water‐soluble phenolic compound derived from Danshen (Salvia miltiorrhiza Bunge), a standard herbal medicine. ¹ Danshen is widely used to treat cardiovascular diseases, such as atherosclerosis, angina, and myocardial infarction, and enhance immunity. ² , ³ DS can improve inflammatory cytokine secretion and histopathological changes by inhibiting the TLR4/NF‐κB pathway in rats with high‐fat diet‐induced atherosclerosis and acute pulmonary inflammation. ⁴ , ⁵ However, DS exhibits limited absorption because of its low hydrophilicity and stability. A newly derived compound of DS, Danshensu sodium salt (DSS) or (R)‐α,3,4‐trihydroxy‐benzenepropanoic acid sodium salt, has shown equivalent efficacy, improved oral absorption, ability to cross the blood–brain barrier, and greater clinical applicability. ⁶ , ⁷ DSS inhibits cardiomyocyte apoptosis through Akt and ERK1/2 phosphorylation and suppresses oxidative stress‐induced autophagy damage via the AMP‐activated protein kinase (AMPK)‐mTOR‐Ulk1 signaling pathway. ⁸ , ⁹ Despite these findings, the effects of DSS on skeletal muscles remain underexplored. Therefore, this study focused on the potential benefits of DSS in sarcopenia.

Skeletal muscle, which constitutes a substantial portion of the human body, accounts for ~40% of body weight. ¹⁰ Sarcopenia, characterized by age‐related loss of muscle mass and strength, involves the upregulation of atrogenes, such as E3 ubiquitin ligases (MuRF1 and MAFbx) within the ubiquitin‐proteasome system and myostatin, a negative regulator of skeletal muscle growth. ¹¹ , ¹² The transcription factor, forkhead box O3 (FoxO3a), which regulates atrogenes in skeletal muscles, is activated and translocates to the nucleus, inducing muscle atrophy. ¹³

AMPK, a serine/threonine kinase family member, acts as a cellular energy sensor by monitoring the AMP:ATP ratio to maintain cellular homeostasis. Phosphorylation at the Thr172 site on the α subunit of AMPK activates the kinase through various upstream kinases, including Ca²⁺/calmodulin‐dependent protein kinase II (CaMKII) and liver kinase B1 (LKB1). ¹⁴ In skeletal muscles, AMPK activation leads to PGC‐1α regulation, which influences myofiber‐type transformation via mitochondrial biogenesis in myotubes. ¹⁵ The inhibition of FoxO3a activity can improve muscle atrophy. For example, the VDR agonist, 1,25‐dihydroxyvitamin D3, activates SIRT1 and AMPK, leading to FoxO3a inactivation and improved muscle atrophy. ¹⁶ Resistance exercise also ameliorates sarcopenia by downregulating FoxO3a and modulating AMPK‐mediated mitochondrial quality control. ¹⁷

Several studies have used D‐galactose (DG)‐induced mouse models to represent accelerated aging. ¹⁸ , ¹⁹ Based on previous studies showing that DSS activates AMPK, we hypothesized that DSS improves sarcopenia symptoms in rapidly aging DG‐induced mice. ⁹ , ²⁰ In this study, we aimed to identify the significant activating signal transduction factors exerted by DSS on skeletal muscles and determine their effects on sarcopenia.

2. MATERIALS AND METHODS

2.1. Cell culture

Mouse C2C12 myoblasts (ATCC) and human skeletal muscle cells (Thermo Fisher Scientific) were cultured under standard conditions (37°C, 5% CO₂) in high‐glucose DMEM (Welgen) supplemented with 10% fetal bovine serum (Gibco) and 100 mg/mL of a penicillin–streptomycin solution (Gibco). When C2C12 cells reached 60–70% confluency, they were subcultured and used for no more than 10 passages. For myogenic differentiation, when C2C12 myoblasts reached 90–100% confluence, they were washed with PBS and replaced with DMEM supplemented with 2% horse serum (Gibco) as the differentiation medium (DM). The DM was changed daily for 5 days until the myotubes were fully differentiated. D‐galactose (Sigma) was added to the DM for 24 h to induce the fully differentiated C2C12 myotubes atrophy. Images of the C2C12 myotubes were captured and processed using a CKX53 microscope (Olympus). All diameters were measured in 10 random fields from each section using the ImageJ software (NIH). Hanks' Balanced Salt Solution (HBSS, Welgen) without calcium chloride was used for calcium signaling imaging.

2.2. Cell viability assay

C2C12 and human skeletal muscle cells were seeded into 96 well‐clear plates and treated with 100 μL of fresh medium containing various doses of DS, DSS, and DG after cell adherence. MTT was added to the medium and incubated at 37°C for 4 h. The medium was discarded, and the formazan crystals were eluted with DMSO and measured at a wavelength of 540 nm.

2.3. Real‐time polymerase chained reaction (RT‐PCR)

Total RNA was extracted from C2C12 myotubes and human skeletal muscle cells using QIAzol reagent (Qiagen, Hilden, Germany). The RNA concentration was then measured using a NanoDrop spectrophotometer (Thermo Fisher Scientific). Complementary DNA was synthesized from 5 μg RNA using a reverse transcription system (Promega). The mRNA expression was assessed using RT‐PCR (Thermo Fisher Scientific) and a SYBR Green system (Enzynomics). The primer sequences used are listed in Table S1. Data were normalized to GAPDH for each sample and fold‐change values were calculated using the ΔΔCt method.

2.4. Immunoblotting (IB)

Protein extraction was performed using C2C12 myotube lysates. Lysates were prepared using RIPA buffer (Thermo Fisher Scientific). RIPA buffer was supplemented with freshly added 1 mM sodium orthovanadate (Na₃VO₄), 1 mM phenylmethylsulfonyl fluoride (PMSF), and 5 mM sodium fluoride (NaF). The extracted protein (20 μg) was subjected to 10% SDS‐PAGE and transferred to a 0.45‐μm nitrocellulose membrane (Millipore). The membranes were blocked in Tris‐buffered saline with 0.1% Tween 20 (TBST) containing 5% bovine serum albumin (BSA) for 1 h, and then washed with TBST. Next, the cells were incubated with primary antibodies in blocking buffer (5% BSA in TBST) overnight at 4°C. The following primary antibodies and dilutions were used: anti‐myostatin (GENTEX, 1:1000), anti‐MuRF1 (Enogene, 1:1000), anti‐MAFbx (Abcam, 1:1000), anti‐FoxO3a (CellSignaling, 1:1000), anti‐p‐AMPKα^(T172) (CellSignaling, 1:1000), anti‐AMPKα (CellSignaling, 1:1000), anti‐p‐CaMKII^(T286) (CellSignaling, 1:1000), anti‐CaMKII (Abcam, 1:1000), anti‐PGC1α (Abclonal, 1:1000), and anti‐β‐actin (Enogene, 1:5000). The secondary antibodies and dilution factors used were HRP‐conjugated anti‐mouse (ENZO, 1:5000) and HRP‐conjugated anti‐rabbit (ENZO, 1:3000). The membranes were washed with TBST to remove excess primary antibodies and then incubated for 2 h with the appropriate HRP‐conjugated secondary antibodies in a blocking buffer before detection. Enhanced chemiluminescence IB substrates (Thermo Fisher Scientific) were used to visualize protein expression using ChemiDoc (Solo6S Edge).

2.5. Calcium and reactive oxygen species (ROS) assay

The C2C12 cells were seeded into confocal dishes (SPL) and incubated. Cells were labeled with Fluo3/AM (Invitrogen) to monitor the intracellular calcium concentration. C2C12 cells were seeded onto confocal dishes and black plates (SPL) to assess and quantify intracellular ROS generation. The differentiation of C2C12 myoblasts was initiated using DM. After a 5‐day differentiation, the cells were cultured for an additional 24 h with or without DG and DSS treatment, washed once with PBS, and stained with DCF‐DA (Invitrogen) for 30 min. Images of ROS generation were captured using a confocal microscope, and quantification was performed using a fluorescence plate reader. The fluorescence intensity of Fluo‐3 AM in living cells was measured using a confocal microscope (Carl Zeiss, LSM800), and that of DCF‐DA was observed using a fluorescence plate reader (BioTek) and a confocal microscope.

2.6. Immunocytochemistry

C2C12 cells were seeded in chamber slides (SPL), and C2C12 myoblasts were differentiated using DM. After a 5‐day differentiation, the cells were cultured for 24 h with or without DG and DSS treatment. Subsequently, the cells were fixed with 4% paraformaldehyde in PBS, permeabilized with 0.1% Triton X‐100 for 5 min, and blocked with 5% BSA for 30 min. Then, the cells were incubated overnight with an anti‐FoxO3a antibody (1:500) in PBS containing 0.1% BSA at 4°C. After multiple washes with PBS containing 0.1% Tween 20, the cells were treated with R‐phycoerythrin goat anti‐rabbit IgG (H + L) (Invitrogen, 1:500) for 30 min at 25°C. Then, the samples were mounted onto slides using PBS with DAPI (Biomeda) and visualized under a confocal microscope.

2.7. Cytosolic and nuclear protein fractions

According to the manufacturer's instructions, C2C12 myotubes were extracted for cytoplasmic and nuclear proteins using the CelLytic NuCLEAR extraction kit (Sigma).

2.8. Co‐immunoprecipitation (Co‐IP) assay

C2C12 myotubes lysate proteins (1 mg) were incubated with rabbit anti‐FoxO3a (1:100) or rabbit anti‐IgG (1:100) for 24 h at 4°C. The immune complexes were captured using protein A‐sepharose (Amersham, UK) after 1 h of incubation. The precipitated immune complexes were washed five times with a wash buffer (25 mM HEPES, 5 mM EDTA, 1% Triton X‐100, 50 mM NaF, 150 mM NaCl, 10 mM PMSF) and denatured in an SDS sample buffer (125 mM Tris–HCl (pH 6.8), 20% (v/v) glycerol, 4% (w/v) SDS, 100 mM DTT, and 0.1% (w/v) bromophenol blue) by boiling at 95°C for 5 min.

2.9. Animal experiment

The Institutional Animal Care and Use Committee of Korea University approved all animal experiments, which were conducted according to relevant guidelines and regulations (KOREA‐2024‐0023). Male C57BL/6 mice aged 8 weeks were obtained from Japan SLC (Hamamatsu) and housed in a highly controlled environment (temperature, 21–23°C; relative moisture, 50–60%; 12 h light:12 h dark cycle). Animals were randomly divided into four groups (n = 6 per group): sham control, DG administration, DG + DSS50 administration, and DG + DSS100 administration. The experiment was performed when the mice reached 9 weeks of age (after acclimation). DG (300 mg/kg) was administered daily via intraperitoneal injection from weeks 0 to 9. After 5 weeks of DG administration, the mice were intraperitoneally injected daily with DSS (50 mg/kg) or DSS (100 mg/kg) along with DG (300 mg/kg) for 4 weeks. Nine weeks after DG administration, the mice were euthanized, and their muscles were removed.

2.10. Glucose tolerance test (GTT)

The GTT was performed after a 12‐h fast. Blood glucose levels were assessed by tail incision at week 9. Blood glucose levels were measured using glucose strips inserted into an Accu‐Check Performa II blood glucose meter (Roche).

2.11. In vivo physical performance test

The grip strength test was performed using a calibrated grip strength tester (BIOSEB) to assess the grip strength of the fore and hind limbs (four paws of the mice). For the hanging test, mice were placed on a wire mesh that was subsequently inverted, forcing them to grip the wire. We recorded drop latencies for up to 15 min before falling. The results of the grip strength and hanging analyses are represented as the means of at least three repetitions.

2.12. Statistical analysis

Data are expressed as mean ± standard error of the mean. Statistical differences between and among groups were evaluated using Student's t‐test and ANOVA, followed by Tukey's post hoc test. GraphPad Prism 5 (GraphPad) software was used for statistical analyses, and p‐values <.05 significance levels were defined as *p < .05, **p < .01, and ***p < .001.

3. RESULTS

3.1. Effect of DSS in skeletal muscle cells

We evaluated cytotoxicity and muscle atrophy factors in mouse and human skeletal muscle cells to determine the effects of DS and DSS. The structures of DS and DSS (Figure 1A). Cytotoxicity was investigated using MTT in C2C12 and human skeletal muscle cells treated with various doses of DS and DSS. After 72 h, cell viability in C2C12 cells was 82.0% for DS and 96.8% for DSS up to 100 μM (Figure 1B). In human skeletal muscle cells, cell viability was 74.5% for DS and 89.6% for DSS at 100 μM (Figure 1C). Both DS and DSS exhibited greater cytotoxicity against human skeletal muscle cells than against C2C12 cells. The maximum concentration that maintained 80% cell viability in both cell types was set at 30 μM for further comparison of their effects. To investigate the effects of DS and DSS on skeletal muscles, we examined the transcriptional expression of skeletal muscle atrophy‐related factors (MuRF1, MAFbx, Myostatin, and FoxO3a). Both DS and DSS significantly reduced the mRNA levels in C2C12 and human skeletal muscle cells (Figure 1D,E). In particular, DSS effectively suppressed atrophy‐related factors with significantly lower cytotoxicity than that with DS. DG‐treated skeletal muscles exhibit age‐related phenotypes. ²¹ , ²² We used DG‐treated C2C12 myotubes as an in vitro model. The cytotoxicity of DG was tested using the MTT assay, and cell viability was ~80% at 30 g/L for up to 72 h (Figure 2A). We used RT‐PCR to determine whether DG affected the transcriptional expression of atrophy‐related factors at a concentration range that was not confirmed to be toxic to C2C12 cells (30 g/L). We observed a significant increase in atrophy‐related factors in the DG treatment (30 g/L); therefore, this was used as the inducing concentration (Figure 2B). We evaluated whether DSS affects the expression of atrophy‐related factors in DG‐treated C2C12 myotubes using RT‐PCR and IB. DSS significantly decreased mRNA and protein expression (Figure 2C,D) and increased the diameter of C2C12 myotubes, which was reduced by DG induction (Figure 2E). These results suggest that DSS alleviates muscle atrophy in DG‐induced C2C12 myotubes.

FIGURE 1.

DSS inhibits the expression of the muscle atrophy‐related factors in skeletal muscle cells. (A) The structural characteristics of DS and DSS. Cell viability of (B) mouse C2C12 cells and (C) human skeletal muscle cells treated with DS and DSS at indicated doses (1–100 μM) for 72 h. Cell viability was evaluated using the MTT assay (n = 3). Comparison of the relative mRNA expression of atrophy‐related genes (MuRF1, MAFbx, Myostatin, and FoxO3a) in (D) C2C12 myotubes and (E) human skeletal muscle cells were treated with the 30 μM of DS and DSS for 24 h. mRNA expression was assessed using PCR and normalized to GAPDH (n = 3). Results are expressed as the mean ± SEM, analyzed using one‐way ANOVA with Tukey's posttest. *p < .05, **p < .01, versus control. DS, Danshensu; DSS, Danshensu sodium salt.

FIGURE 2.

DSS inhibits the myotube atrophy in D‐galactose‐induced skeletal muscle cells. (A) Cell viability of mouse C2C12 cells treated with DG at various doses (5–60 g/L) for 24–72 h, evaluated using the MTT assay (n = 3). (B) Relative mRNA expression of atrophy‐related genes (MuRF1, MAFbx, Myostatin, and FoxO3a) in C2C12 myotubes treated with DG (10–30 g/L) for 24 h. (C) Relative mRNA and (D) protein expression of atrophy‐related genes (MuRF1, MAFbx, Myostatin, and FoxO3a), and (E) morphology images in C2C12 myotubes pretreated with DSS (10–30 μM) for 30 min, then cultured with or without DG (30 g/L) for 24 h. mRNA expression was assessed using PCR and normalized to GAPDH (n = 3). Myotube diameters were calculated using ImageJ (n = 10). Results are expressed as mean ± SEM, analyzed using one‐way ANOVA with Tukey's posttest (*p < .05, **p < .01, ***p < .001, versus control). Scale bar: 100 μm. DG, D‐galactose; DSS, danshensu sodium salt.

3.2. DSS activates AMPK‐PGC1α via CaMKII

DSS activates AMPK. ⁹ , ²⁰ Therefore, we treated C2C12 myotubes at various time points to determine whether DSS activated AMPK. As a result, we observed that AMPK was activated 30 min after DSS treatment, and the expression of PGC‐1α increased after 3 h (Figure 3A). Accordingly, we used Compound C, an AMPK inhibitor, to verify the increased expression of p‐AMPKα and PGC‐1α due to DSS treatment. As a result, Compound C blocked the increase in p‐AMPKα and PGC‐1α induced by DSS. These results indicated that DSS activated AMPK in C2C12 myotubes (Figure 3B). Next, we investigated upstream factors involved in AMPK activation. AMPK can be activated by calcium, therefore, we evaluated whether DSS treatment affected calcium secretion at the cellular level using Fluo‐3; DSS treatment increased calcium secretion. Next, we identified the source of calcium secretion by blocking inside or outside the cell. Thapsigargin (an endoplasmic reticulum stress inducer) and BAPTA (a calcium chelator) blocked calcium secretion, confirming that intracellular calcium was involved (Figure 3C). Based on these results, we evaluated the phosphorylation of CaMKII, an upstream regulator of AMPK, to confirm whether DSS treatment activates calcium‐dependent AMPK. CaMKII phosphorylation was significantly increased after 30 min (Figure 3D). We used STO609 (a CaMKII inhibitor) to verify whether DSS is involved in AMPK and PGC1α expression through CaMKII in skeletal muscle cells. As a result, STO609 effectively blocked the increase of p‐AMPKα and PGC‐1α induced by DSS (Figure 3E). These results suggest that DSS can enhance PGC‐1α expression by activating AMPK through a calcium‐dependent mechanism in C2C12 myotubes.

FIGURE 3.

DSS activates the AMPKα‐PGC1α via CaMKII. (A) Expression of AMPKα and PGC1α in C2C12 myotubes treated with DSS (30 μM) for various times (0.5–24 h). (B) Expression of AMPKα and PGC1α in C2C12 myotubes pretreated with compound C (30 μM) for 30 min, then cultured with or without DSS (30 μM) for 24 h. (C) C2C12 myoblasts were stained with Fluo‐3 (1 μM) for 30 min, and fluorescence intensity was analyzed using confocal microscopy. Cells were exposed to DSS in calcium‐free HBSS or a combination of Thapsigargin (1 μM) and BAPTA‐AM (10 μM). Fluorescence images were captured every second after DSS treatment (n = 4). (D) Expression of CaMKII in C2C12 myotubes treated with DSS (30 μM) for various times (0.5–24 h). (E) Expression of AMPKα, CaMKII, and PGC1α in C2C12 myotubes pretreated with STO609 (30 μM) for 30 min, then cultured with or without DSS (30 μM) for 24 h. Scale bar, 20 μm. DSS, danshensu sodium salt; HBSS, Hanks' balanced salt solution.

3.3. DSS attenuates muscle atrophy via CaMKII‐AMPK‐PGC1α in DG‐induced C2C12 myotubes

DG‐induced cells exhibit decreased AMPK activation. ²³ , ²⁴ We investigated whether DG reduced AMPK activation in DG‐treated C2C12 myotubes. AMPK activation was reduced after 6 h, and the expression of PGC1α was significantly reduced at 24 h (Figure 4A). We used DG‐treated C2C12 myotubes to determine whether DSS counteracts the DG‐induced decrease in AMPK activation. DSS restored the decreased p‐AMPKα and PGC1α expression (Figure 4B). DSS ameliorated the decreased AMPK activation induced by DG, therefore, we used STO609 to inhibit the upstream factors of AMPK to verify its effect. DSS treatment ameliorated the increased expression of muscle atrophy‐related factors and decreased expression of AMPK‐PGC1a in DG‐induced C2C12 myotubes, but this effect was blocked by STO609 (Figure 4C). Additionally, DSS treatment reversed the decrease in the diameter of DG‐treated C2C12 myotubes; however, this effect was inhibited by STO609 (Figure 4D). Next, we explored whether DSS, which inhibits muscle atrophy‐related factors, affected FoxO3a, a transcription factor involved in regulating atrophy factors. To determine whether DSS inhibits the nuclear translocation and activation of FoxO3a in DG‐treated C2C12 myotubes, we used confocal microscopy to localize FoxO3a. DSS treatment reduced FoxO3a translocation to the nucleus (Figure 5A). Cell lysates were fractionated into nuclear and cytoplasmic fractions to quantitatively evaluate both the localization and distribution of FoxO3a. DSS treatment of C2C12 myotubes without DG induction, increased the expression of FoxO3a in the cytoplasm (Figure 5B). In contrast, DSS treatment of DG‐treated C2C12 myotubes resulted in a decrease in nuclear FoxO3a levels, and this effect was blocked by STO609 (Figure 5C). FoxO3a interacts with PGC‐1α. This interaction suppresses oxidative stress and reduces the expression of target genes. ²⁵ , ²⁶ We performed Co‐IP to determine whether DSS enhances the interaction between PGC‐1α and FoxO3a. We observed a strong interaction 3 h after DSS treatment (Figure 5D). We then investigated whether this interaction was also present in DG‐induced C2C12 myotubes. Our results showed that DSS treatment reversed the decreased interaction between PGC‐1α and FoxO3a in DG‐induced C2C12 myotubes, and this effect was blocked by STO609 (Figure 5E). Therefore, we investigated whether DSS protects skeletal muscle from oxidative stress. Treatment with various concentrations of DG increased intracellular ROS levels in skeletal muscle cells; however, DSS effectively inhibited this increase. Specifically, C2C12 myotubes induced with DG (30 g/L) exhibited significantly reduced ROS levels after DSS treatment (Figure 5F). These findings suggest that DSS may help prevent skeletal muscle atrophy by inhibiting FoxO3a by activating CaMKII‐PGC‐1α.

FIGURE 4.

DSS attenuates the myotube atrophy via CaMKII. (A) Expression of AMPKα and PGC1α in C2C12 myotubes treated with DG (30 g/L) for various times (0.5–24 h). (B) Expression of AMPKα and PGC1α in C2C12 myotubes pretreated with DSS (30 μM) for 30 min, then cultured with or without DG (30 g/L) for 24 h. (C) Expression of MuRF1, MAFbx, Myostatin, FoxO3a, AMPKα, CaMKII, and PGC1α, and (D) morphological images in C2C12 myotubes pretreated with DSS (30 μM) for 30 min, then cultured with or without DG (30 g/L) or STO609 (30 μM) for 24 h. Myotube diameters were calculated using ImageJ (n = 10). Results are expressed as mean ± SEM, analyzed using one‐way ANOVA with Tukey's posttest (*p < .05, **p < .01, ***p < .001, versus control). Scale bar: 100 μm. DG, D‐galactose.

FIGURE 5.

DSS inactivates FoxO3a through interaction between FoxO3a and PGC1α via CaMKII. (A) C2C12 myotubes were cultured in DG (30 g/L) with or without DSS (30 μM) for 24 h. Protein expression and localization of FoxO3a were assessed using anti‐FoxO3a antibody (green) and DAPI (blue) for nuclear staining, visualized by confocal microscopy. Subcellular fractions were prepared to detect FoxO3a localization. (B) C2C12 myotubes were treated with DSS (30 μM) for varying times (0.5–24 h). (C) C2C12 myotubes were pretreated with DSS (30 μM) for 30 min, then cultured with or without DG (30 g/L) or STO609 (30 μM) for 24 h. FoxO3a protein expression was evaluated by immunoblotting, normalized to GAPDH (cytoplasmic) and Lamin B (nuclear). (D) Protein–protein interaction between PGC1α and FoxO3a in C2C12 myotubes treated with DSS (30 μM) for different durations (1–24 h). (E) Interaction between PGC1α and FoxO3a in C2C12 myotubes pretreated with DSS (30 μM) for 30 min, then cultured with or without DG (30 g/L) or STO609 (30 μM) for 24 h. Anti‐rabbit IgG served as a negative control. (F) Intracellular ROS generation in C2C12 cells pretreated with DSS (30 μM) for 30 min, then cultured with or without DG (5–40 g/L) for 24 h. Intracellular ROS measurements were assessed using DCF‐DA. Statistical analysis was performed using one‐way ANOVA with Tukey's posttest (**p < .01, ***p < .001 versus control). Scale bar: 10 μm. DG, D‐galactose; DSS, Danshensu sodium salt.

3.4. DSS administration ameliorates symptoms of sarcopenia

Next, we evaluated the effects of DSS in a DG‐induced accelerated aging model at the animal level. DSS (50–100 mg/kg) was administered to mice exposed to DG (300 mg/kg) for 9 weeks. These effects were assessed using grip strength and hanging tests. Behavioral assessments were performed weekly throughout the 9 weeks of DG treatment to monitor accelerated aging. DSS was administered from week 5 to 9 after DG induction (Figure 6A). We monitored the weights of the mice throughout the 9‐week experiment, and no significant differences were found (Figure 6B). However, when the GTT was performed at 9 weeks, the GTT of DG‐treated mice significantly increased, and this increase was significantly reduced by DSS administration (Figure 6C). DG‐induced mice exhibit age‐related phenotypes characterized by an increase in GTT, although there is no difference in weight. ²⁷ , ²⁸ We evaluated whether DSS improved sarcopenia symptoms by performing grip strength and hanging tests to assess muscle strength. The results of grip strength measurements showed that the DG treatment group experienced a significant decrease in grip strength compared with the sham group from the second week of the experiment. Additionally, DSS administration (50–100 mg/kg) significantly recovered the decrease in grip strength induced by DG treatment (Figure 6D). The hanging test was performed to evaluate skeletal muscle endurance performance. The results showed a decline in endurance in DG‐administered mice from week 5. After 9 weeks, DSS (100 mg/kg) significantly improved endurance (Figure 6E). At the end of the 9 weeks, the mice were euthanized, and their skeletal muscles were extracted and weighed. The results showed that the weights of the tibialis anterior (TA), gastrocnemius (GCM), and quadricep (QA) muscles in DG‐treated mice were significantly reduced compared to those in the sham group, and significantly increased after DSS administration (Figure 6F,G). DSS can increase PGC1a‐FoxO3a interaction via CaMKII by increasing calcium inside skeletal muscle cells. This protected against oxidative stress, improved muscle atrophy, and enhanced muscle function (Figure 7). These results suggested that DSS improved sarcopenia symptoms in mice with accelerated DG‐induced aging.

FIGURE 6.

DSS administration improves the muscle atrophy in D‐galactose‐induced mice. (A) Schema of the experimental schedule for the animal study. (B) Weekly body weight measurement. (C) GTT measurement at 9 weeks. (D) Grip strength testing and (E) hanging test during weeks 0–9. (F) Skeletal muscle (tibialis anterior; TA, gastrocnemius; GCM, quadriceps; QA, soleus; SOL, and extensor digitorum longus; EDL) morphology image and (G) skeletal muscle weight of mice were evaluated at the time of sacrifice. One‐way ANOVA was used to determine statistical significance. Each result was expressed as mean ± SEM of 6 ice in each group. *p < .05, **p < .01, and ***p < .001 versus sham; ^# p < .05, ^## p < .01, and ^### p < .001 versus DG. Sham: Sham mice; DG: D‐galactose (300 mg/kg) intraperitoneal administrated mice; DG + DSS50: D‐galactose (300 mg/kg) intraperitoneal administrated mice treated with DSS (50 mg/kg) intraperitoneally; DG + DSS100: D‐galactose (300 mg/kg) intraperitoneal administrated mice treated with DSS (100 mg/kg) intraperitoneally. DSS, Danshensu sodium salt; GTT, glucose tolerance test.

FIGURE 7.

Schematic of DSS working model. DSS increases CaMKII‐AMPK via intracellular calcium in skeletal muscle cells. DSS inhibits FoxO3a, whose nuclear translocation is mediated by increased protein interaction between PGC1α and FoxO3a. DSS‐induced FoxO3a inhibition inhibits transcription of muscle atrophy factors. DSS, Danshensu sodium salt.

4. DISCUSSION

We investigated the effects of DSS on sarcopenia, focusing on its impact on muscle atrophy and signaling pathways in skeletal muscle cells, using a DG‐induced accelerated aging mouse model. Currently, pathological mechanisms, such as muscle protein reduction, increased inflammation, oxidative stress, and hormonal changes are involved in sarcopenia treatment research. Treatments, such as myostatin inhibitors and exercise, are recommended for sarcopenia; however, these have limitations. ²⁹ For example, myostatin inhibitors may not be appropriate for patients with cardiovascular disease, and exercise therapy may have unintended side effects in patients with poor joint function. Therefore, we investigated DSS as an AMPK activator for treating sarcopenia to overcome these limitations. ⁹ , ²⁰

Our study confirmed that DSS reduced muscle atrophy by increasing AMPK‐PGC‐1α expression in C2C12 myotubes. AMPK activation by DSS was calcium‐dependent and reversed the decrease in AMPK activation and PGC‐1α expression in DG‐induced C2C12 myotube models. The increase in CaMKII‐AMPK phosphorylation induced by DSS, possibly via intracellular calcium, may involve the ER and mitochondria. ³⁰ DSS inhibited the nuclear translocation of FoxO3a, a key regulator of muscle atrophy, promoted its interaction with PGC‐1α, and reduced intracellular ROS levels in skeletal muscle. These results suggest that DSS may not only increase PGC‐1α expression in a calcium‐dependent manner in skeletal muscle cells, but also prevent skeletal muscle atrophy by protecting against oxidative stress. ²⁵ , ²⁶ We confirmed the mechanism by which DSS reduces muscle atrophy‐related factors at the cellular level, as well as the reduction of symptoms of sarcopenia at the animal level. ³¹ DSS administration restores decreased GTT and skeletal muscle function in DG‐induced accelerated aging mice. ²⁷ , ²⁸ In particular, our results confirmed an improvement in grip strength, which represents the weight and strength of the skeletal muscle, and hanging test, which represents endurance. Additionally, DSS treatment improved the weights of the TA, GCM, and QA muscles in DG‐induced accelerated aging mice. These results suggest that DSS improves skeletal muscle mass and function, which are currently known diagnostic criteria for sarcopenia, through AMPK activation.

AMPK, a well‐known signal that is generally activated during exercise, is activated in a calcium‐dependent manner. ³² , ³³ This activation of AMPK may affect mitochondria within the cell, which may affect PGC1α. Our study confirmed that DSS activates AMPK and increases PGC1α expression in a CaMKII‐dependent manner. ³² DSS requires further comparison with exercise performance. Aerobic and resistance exercises enhance bidirectional signaling involving AMPK and Akt. ³⁴ DSS also activates Akt, therefore, it has the potential to replace exercise; however, this requires further study. ³⁵ , ³⁶ The role of AMPK in skeletal muscle is complex and controversial, as its activation can either increase or decrease muscle mass depending on the pathological or physiological conditions. Our previous study indicated that metformin impaired muscle function by modulating myostatin in skeletal muscle cells via the AMPK‐FoxO3a‐HDAC6 axis. ³⁷ In contrast, other studies have demonstrated that myonectin can enhance muscle function via an AMPK/PGC1α‐dependent mechanism in mouse models of accelerated aging and Duchenne muscular dystrophy. ³⁸ Additionally, 20(S)‐ginsenoside Rg3 improves mitochondrial function and prevents muscle atrophy by regulating the AMPK‐FoxO3 signaling pathway in C2C12 myotubes. ³⁹ Furthermore, ampelopsin mitigates DG‐induced muscle atrophy in SD rats by upregulating the AMPK and SIRT1 signaling pathways. ⁴⁰ Decreased PGC1α expression affects FoxO3a regulation and exacerbates muscle atrophy. ⁴¹ Although controversial, our study suggests that DSS can increase PGC1a expression as well as AMPK activation, which may provide the rationales for the atrophy inhibition effect in DG‐induced accelerated aging mice.

Although our study provides evidence supporting the efficacy of DSS in alleviating muscle atrophy, several limitations must be addressed. First, the relatively small sample size (n = 6 per group) in the animal experiments, which was designed to balance exploratory objectives and ethical considerations, may limit the statistical power and generalizability of the results. ⁴² Second, while the DG model effectively induces age‐related muscle atrophy through oxidative stress, it lacks the natural aging and the ability to replicate the localized muscle atrophy induced by denervation. Natural aging models encompass systemic processes, such as inflammation and mitochondrial dysfunction, which closely align with human sarcopenia but require extended study durations. ⁴³ , ⁴⁴ In contrast, denervation models specifically target neuromuscular disruptions, providing insights into localized atrophy mechanisms while failing to recapitulate systemic aging effects. ⁴⁵ Third, although the effectiveness of DSS was confirmed in a standardized DG model, its long‐term efficacy has not yet been evaluated. ¹⁹ , ²⁸ Each sarcopenia model offers distinct advantages and can be combined to provide evidence for the long‐term effects of DSS treatment. Future studies should involve larger cohorts and integrate complementary models to comprehensively evaluate the potential of DSS for long‐term treatment of sarcopenia.

In summary, our findings demonstrated that DSS administration is required in sarcopenia to confirm the direct CaMKII‐PGC1α–FoxO3a relationship, which can trigger the inhibition of muscle atrophy‐related factors expression. The evidence presented here shows that DSS treatment effectively inhibits muscle atrophy‐causing protein degradation in both DG‐treated mice and C2C12 myotubes, suggesting that DSS is a potential therapeutic candidate for improving skeletal muscle protein degradation in sarcopenia.

AUTHOR CONTRIBUTIONS

P. R. Lim: Conceptualization, data curation, formal analysis, validation, investigation, visualization, writing–original draft, investigation, methodology, writing–review and editing. S. W. Woo: Data curation, formal analysis, validation, visualization, writing–review and editing. J. H. Han: Data curation. Y. L. Lee: Writing–review and editing. J. H. Shim: Conceptualization, supervision, funding acquisition, writing–original draft, project administration, writing–review and editing. H. S. Kim: Conceptualization, supervision, funding acquisition, validation, writing–original draft, project administration, writing–review, and editing.

DISCLOSURES

The authors declare no conflicts of interest.

Supporting information

Table S1.

Lim P, Woo SW, Han J, Lee YL, Shim JH, Kim HS. Danshensu sodium salt alleviates muscle atrophy via CaMKII‐PGC1α‐FoxO3a signaling pathway in D‐galactose‐induced models. The FASEB Journal. 2025;39:e70280. doi: 10.1096/fj.202402158R

DATA AVAILABILITY STATEMENT

Data sharing is not applicable to this article as no datasets were generated or analyzed during the current study.