The VSE (cat. no. KPM012-022) used in the present study was obtained from the Natural Product Central Bank at the Korea Research Institute of Bioscience and Biotechnology. The plant was collected in 2001 from Baegam-myeon, Cheoin, Yongin Gyeonggi, Korea. To create the extract, 74 g dried and powdered seeds were added to 1 liter of 99.9% methyl alcohol (high-performance liquid chromatography grade) and extracted at room temperature through 30 cycles of ultrasonication (40 KHz; 1,500 W) for 15 min using an ultrasonic extractor (SDN-900H; Sungdong Ultrasonic Co., Ltd.), then left to stand for 120 min. The extract was filtered (Qualitative Filter No. 100; HYUNDAI MICRO Co., Ltd.) and dried under reduced pressure. A total of 6.53 g VSE was obtained.

Murine C2C12 myoblast cells were obtained from American Type Culture Collection. The cells were grown at 37°C with 5% CO₂ in DMEM (Gibco; Thermo Fisher Scientific, Inc.) with fetal bovine serum (Gibco; Thermo Fisher Scientific, Inc.) at 10% (v/v) and penicillin-streptomycin (Gibco; Thermo Fisher Scientific, Inc.) at 1% (v/v). To induce myoblast differentiation into C2C12 myotubes, once cells reached 90% confluency, the medium was switched to DMEM containing horse serum (Gibco; Thermo Fisher Scientific, Inc.) at 2% (v/v). The differentiation media were changed every 2 days and the cells were cultivated for 4 days.

C2C12 myoblasts were seeded at a concentration of 1×10⁶ cells/ml in 6-well plates and differentiated into myotubes over 4 days. Subsequently, the existing medium was removed by aspiration, and the myotubes were rinsed twice with PBS and fixed in 4% paraformaldehyde for 10 min at room temperature. Giemsa solution, diluted 1:10 in distilled water and filtered through a 0.2-µm filter, was applied to fixed cells and left for 40 min at room temperature. After removing the Giemsa solution by aspiration, cells were rinsed three times with PBS, and then imaged under a bright-field microscope (Leica Microsystems GmbH) and analyzed with ImageJ (National Institutes of Health).

Cell viability was measured using a Cell Counting Kit-8 (CCK-8; Dojindo Laboratories, Inc.) according to the manufacturer's instructions. After differentiating C2C12 myoblasts into myotubes for 4 days in 96-well plates, the cells were treated with VSE at concentrations of 0 (control), 1.25, 2.5, 5 and 10 µg/ml for 48 h at 37°C. Given the lack of previous studies on VSE in muscle models, cytotoxicity testing was conducted to determine appropriate concentrations for subsequent experiments. After treatment, 10 µl CCK-8 solution was added to each well, and the cells were incubated for an additional 2 h. Absorbance was measured at 450 nm using a SPECTROstar microplate reader (BMG Labtech GmbH).

Total RNA was obtained from cells using RNAiso Plus Reagent (Takara Bio, Inc.). After measuring the mRNA yield, cDNA synthesis was carried out using ReverTra Ace™ qPCR RT Master Mix (cat. no. FSQ-201; Toyobo Co., Ltd.) according to the manufacturer's instructions. The reaction conditions were as follows: 37°C for 5 min for genomic DNA removal, followed by 37°C for 15 min for reverse transcription and 98°C for 5 min for enzyme inactivation. RT-qPCR was performed using a Cycler iQ™ Real-Time PCR Detection System (Bio-Rad Laboratories, Inc.) and SYBR Green master mix (Toyobo Co., Ltd.) with the appropriate primers (Table I) and the template cDNA. The thermocycling conditions were as follows: Initial denaturation at 95°C for 1 min, followed by 40 cycles of 95°C for 15 sec and 60°C for 60 sec. A melt curve analysis was conducted to verify amplification specificity. Analysis of relative gene expression data using RT-qPCR and the 2^−ΔΔCq method (14).

Differentiated myotubes were pretreated with VSE at concentrations of 0 (control), 1.25, 2.5 and 5 µg/ml for 3 h at room temperature and then co-treated with LPS (500 ng/ml) for 24 h at 37°C. Supernatants were acquired, and secretion of the proinflammatory cytokines TNF-α (cat. no. MTA00B; R&D systems, Inc.) and IL-1β was analyzed using ELISA kits (cat. no. DY401; R&D Systems, Inc.) according to the manufacturer's guidelines.

To acquire lysates from myotubes in vitro, cells were treated with 1% protease and 1% phosphatase inhibitors in RIPA buffer (Biosesang) at room temperature for 30 min. To acquire lysates in vivo, gastrocnemius (GAS) samples were lysed using PRO-PREP^TM (Intron Biotechnology, Inc.) at room temperature for 30 min. After Bradford assay quantification of protein concentrations, 30 µg of protein were separated by SDS-PAGE using 10% polyacrylamide gels. Subsequently, the proteins were transferred to a 0.2-µm nitrocellulose membrane and the membrane was blocked in 5% skimmed milk in Tris-buffered saline containing 0.1% Tween 20 (TBST) for 1 h at room temperature. Each membrane was incubated with one of 18 primary antibodies (Table II) on a shaker overnight at 4°C to facilitate primary antibody binding. The membrane was then rinsed with TBST three times, and the membrane was incubated with secondary antibody (rabbit and mouse) at room temperature for 1 h. After rinsing three times with TBST, protein bands were detected using an enhanced chemiluminescence detection kit (Cytiva). For semi-quantitative protein expression analysis, membrane images were processed using ImageJ version 1.54d (National Institutes of Health).

Myotubes were treated with either 500 ng/ml LPS alone or co-treated with 1.25, 2.5 or 5 µg/ml VSE for 24 h at 37°C and then stained with MitoTracker Orange (Invitrogen; Thermo Fisher Scientific, Inc.) for 30 min at 37°C. To target mitochondrial ROS (mtROS), cells were stained with MitoSOX (Invitrogen; Thermo Fisher Scientific, Inc.) dissolved in anhydrous N,N-dimethylformamide (MilliporeSigma) for 30 min at 37°C. The mitochondria were observed at 554/576 nm, and the generation of mtROS was observed at 488/510 nm, under a fluorescence microscope. Images were recorded using the LAS X version 3.7.2.22383 software (Leica Microsystems GmbH).

To assess the mitochondrial membrane potential (Δψm), cells were incubated with JC-1 dye (Invitrogen; Thermo Fisher Scientific, Inc.) at a final concentration of 5 µM at 37°C in the dark for 30 min according to the manufacturer's protocol. JC-1 fluorescence was detected at 514/529 nm for green fluorescence and 514/590 nm for red fluorescence using a fluorescence microscope (Leica Microsystems GmbH). All fluorescence intensities were quantified using ImageJ version 1.54d software (National Institutes of Health).

A total of 6 male C57BL/6J mice aged 5 weeks (average body weight, 23 g) were purchased from Hyochang Science and underwent an acclimation period of 1 week. The mice were housed in an appropriately controlled environment at 25–30°C with 50–60% relative humidity and a 12-h light/dark cycle. No restrictions were applied on access to food and water. To validate the effects of VSE in the sepsis-induced sarcopenia model, mice were divided into four groups to ensure comparable average body weights across groups with six mice per group: i) Control (saline); ii) LPS + saline (1 mg/kg LPS for 24 h); iii) LPS + low VSE (1 mg/kg LPS and 2.5 mg/kg/day VSE); and iv) LPS + high VSE (1 mg/kg LPS and 5 mg/kg/day VSE) for 7 days. LPS was dissolved in sterile water and VSE was dissolved in saline solution; all groups, including the control group, were orally administered the same volume of saline (100 µl/mouse) either alone or containing VSE daily for 7 days, followed by administration of 1 mg/kg LPS in 100 µl sterile water by intraperitoneal injection on day 8 to induce an inflammatory response in vivo. Body weight was measured on days 0, 2, 4, 6, 8 (before LPS treatment) and 9 (before sacrifice) to monitor the effects of VSE and LPS over time. Daily body weight measurement was avoided to minimize animal handling stress. The control group was administered sterile saline (100 µl; intraperitoneally), equivalent in volume and route to the LPS-treated group.

Grip strength tests were carried out 18 h after LPS administration. The mice were then fasted from food only, with free access to water, for 24 h in order to reduce biological variability and ensure consistency in downstream measurements. Grip strength was measured 18 h after LPS administration, based on the results of a pilot study (Fig. S1). In this study, C57BL/6J mice were intraperitoneally injected with LPS (1 mg/kg), and physiological and molecular parameters were assessed at 18 and 24 h post-injection. Body and skeletal muscle weights (gastrocnemius, tibialis anterior, extensor digitorum longus and soleus) were slightly decreased at both time points, although this was not significant. Serum TNF-α levels were significantly elevated at 18 h, and mRNA expression levels of MuRF1 and Atrogin-1 were markedly increased at 18 h compared with those in the control group (Fig. S1). Based on these results, 18 h was selected as the optimal time point for evaluating early-stage systemic inflammation and muscle atrophy. These findings support the appropriateness of the 18 h time point for grip strength measurement. Blood samples were collected and mice were sacrificed 6 h after the grip strength test. The animal experiments were approved by the Animal Ethics Committee of Kyungpook National University (approval no. KNU 2024-0511; Daegu, South Korea) and were carried out in accordance with the institution's ethical guidelines. The sample size (n=6 per group) was selected based on previous studies using murine models of inflammation-induced muscle atrophy, where similar group sizes yielded statistically valid and biologically relevant results (15–17). Additionally, the group size was determined in consideration of ethical guidelines, statistical feasibility and animal welfare. At the end of the experiment, mice were deeply anesthetized with isoflurane (4–5% in oxygen), followed by cervical dislocation. Death was confirmed by the absence of a heartbeat and corneal reflex. Immediately after sacrifice, the GAS muscles were carefully dissected, rinsed in ice-cold PBS to remove residual blood, blotted dry, snap-frozen in liquid nitrogen, and stored at −80°C until analysis. For protein extraction, tissues were lysed in PRO-PREP™ protein extraction solution (Intron Biotechnology, Inc.) on ice, homogenized using a tissue grinder, and centrifuged at 13,000 × g for 20 min at 4°C. The supernatants were collected and used for subsequent western blotting.

Grip strengths of the mice were measured using a grip strength test device (Grip Strength Meter for Mice and Rats; Ugo Basile SRL), with five repetitions per group. After acquiring the results, the data were standardized to body weight and expressed as the mean.

GAS muscle tissues were obtained from each group. The tissues were fixed in 10% neutral buffered formalin at room temperature for 24 h, dehydrated and embedded in paraffin. Paraffin-embedded tissues were sectioned at a thickness of 5 µm, deparaffinized in xylene and rehydrated through a graded ethanol series followed by distilled water. The sections were then stained with hematoxylin for 5–10 min at room temperature, rinsed in water and counterstained with eosin for 1–3 min at room temperature. All staining procedures were performed using a standard protocol by a commercial pathology service (18). After staining, muscle tissue sections were imaged using a bright-light microscope, and muscle fiber organization was analyzed using ImageJ 1.54d (National Institutes of Health).

In vitro experiments were conducted with three independent biological replicates, and in vivo experiments were performed with six mice per group (n=6), and all data are presented as the mean ± SD. Statistical analyses were carried out using GraphPad Prism 9.4.1 (Dotmatics). One-way ANOVA was used to compare differences among multiple groups, followed by Tukey's post hoc test for pairwise comparisons. P<0.05 was considered to indicate a statistically significant difference.

Inflammation is one of the main elements influencing skeletal muscle atrophy. Inflammation was induced in muscles using LPS, an effective proinflammatory agent (6,19). Muscle atrophy results from activation of the ubiquitin system, which is associated with muscle metabolism (20). Controlling skeletal muscle atrophy specifically requires regulation of the expression of the muscle-specific E3 ubiquitin ligases MuRF1 and muscle atrophy F-box protein (MaFbx) and their upstream mechanism, the Akt/Foxo3a pathway (21). Fig. 1 shows the effects of VSE on cell viability and muscle atrophy-related factors. As Fig. 1A illustrates, VSE exerted no toxic effects on C2C12 myotubes at concentrations ≤5 µg/ml. Meanwhile, cell viability significantly increased with VSE treatment (114–131%) compared with the non-treatment group (100%). Cell viability slightly decreased between 5 and 10 µg/ml (Fig. 1A); however, the viability at 10 µg/ml was still increased compared with the control, suggesting no cytotoxicity at concentrations >5 µg/ml. The treatment concentrations were set as 1.25, 2.5 and 5 µg/ml in subsequent experiments.

VSE treatment at 5 µg/ml reduced the upregulation of MuRF1 and atrogin-1 mRNA expression caused by LPS Meanwhile, an analogous pattern in expression was observed for protein expression levels of MuRF1 and MaFbx. Moreover, VSE treatment significantly attenuated the LPS-induced decrease in MyHC protein expression (Fig. 1D). Giemsa staining revealed that VSE treatment restored the thinned myotube morphology caused by LPS exposure, indicating an alleviating effect on LPS-induced atrophy (Fig. 1C). Furthermore, LPS significantly reduced the phosphorylation of both Akt and Foxo3a, leading to the activation of muscle atrophy-related genes such as MuRF1 and MaFbx. By contrast, VSE treatment increased the protein levels of MyHC, phosphorylated Akt and Foxo3a (Fig. 1E). These findings indicated that VSE alleviated inflammation-induced skeletal muscle atrophy by modulating the Akt/Foxo3a signaling pathway and subsequently suppressing the expression of atrophy-related genes.

Pyroptosis, which is activated under various inflammatory states, including LPS-induced inflammation, is modulated by the NLRP inflammasome, with NLRP3 initiating inflammasome expression (22,23). Thus, the effect of VSE on NLRP inflammasome activation via the NLRP3 pathway was investigated (Fig. 2). When administered alone, LPS increased the mRNA levels of NLRP3, which were downregulated by VSE treatment to levels similar to those observed in the control group (Fig. 2A). Additionally, the protein expression levels of pyroptosis-related factors were investigated, including those of NLRP3, caspase-1, cleaved-caspase-1, GSDMD and cleaved-GSDMD. The ratios of cleaved to total caspase-1 and GSDMD were significantly increased in the LPS group, indicating activation of pyroptosis. VSE treatment effectively suppressed these ratios, suggesting inhibition of LPS-induced pyroptotic signaling. A significant reduction in cleaved caspase-1 levels was observed only for 5 µg/ml VSE (Fig. 2B). These results demonstrated that VSE may reduce NLRP3 inflammasome-mediated pyroptosis.

Release of IL-1β, a proinflammatory cytokine, is increased during pyroptosis, leading to the initiation of several cellular processes, including inflammation (24), and the release of IL-1β through cell membrane pores created by cleaved-GSDMD is considered to be a key factor in pyroptosis (25). To ascertain the effect of VSE on LPS-induced inflammatory responses, VSE-mediated release of IL-1β and TNF-α was observed. The secretion of both cytokines was evaluated at the mRNA and protein levels. When administered alone, LPS substantially increased the levels of IL-1β and TNF-α; however, VSE treatment decreased the LPS-induced upregulation at all concentrations (Fig. 3A and B). These findings suggested that the anti-inflammatory effects exerted by VSE include modifying the release of proinflammatory cytokines.

Pyroptosis and the generation of ROS are mutually dependent, functioning as cause and effect. Specifically, regulation of mtROS is a key process in pyroptosis (26). The expression levels of factors associated with the nuclear factor erythroid 2-related factor 2 (Nrf2)/heme oxygenase-1 (HO-1) signaling pathway were assessed to examine the potential alleviating effects of VSE on oxidative stress. VSE treatment at 5 µg/ml significantly upregulated the Nrf2/HO-1 signaling pathway (Fig. 4A). Although statistical significance was not assessed, treatment with 5 µg/ml VSE appeared to increase HO-1, Nrf2 and PGC-1α expression when compared with the untreated control. Furthermore, the impact of VSE on mtROS release was assessed. To quantify this impact using immunofluorescence staining, mitotracker Orange staining, which labels mitochondria, was combined with mitoSOX green staining, specifically targeting mtROS. The LPS-induced generation of mtROS was decreased by VSE treatment (Fig. 4B), suggesting that VSE may influence pyroptosis by regulating mtROS levels.

The NLPR3 protein translocates to the mitochondria in response to stimuli such as LPS, which induces mitochondrial fragmentation. Healthy mitochondria are required to suppress NLPR3 inflammasome activation (27,28). Additionally, muscle atrophy is largely influenced by the degradation of mitochondrial functions, including oxidative phosphorylation, aerobic respiration and Δψm (12).

The present study aimed to investigate the effects of VSE treatment on mitochondrial functions, which are known to be negatively impacted following LPS stimulation (29). Therefore, the expression levels of oxidative phosphorylation system (OXPHOS) complexes I–V were examined. While LPS stimulation significantly reduced the expression of most OXPHOS complexes, complex IV remained unchanged. VSE treatment, however, dose-dependently restored the expression of complexes I–III and V, with complexes IV and V showing particularly notable increases at higher VSE concentrations (Fig. 5A). These findings suggest that VSE helps maintain mitochondrial metabolism at near-normal levels.

Additionally, JC-1 staining was performed to assess changes in Δψm. In healthy mitochondria, JC-1 forms aggregates emitting red fluorescence, whereas in depolarized mitochondria it exists as monomers emitting green fluorescence. LPS treatment reduced red fluorescence and significantly decreased the red/green fluorescence ratio, indicating mitochondrial depolarization. By contrast, VSE treatment restored Δψm in a dose-dependent manner, as evidenced by the increasing red/green ratio (Fig. 5B).

Collectively, these results indicate that VSE preserves mitochondrial function at both the molecular (OXPHOS protein expression) and functional (Δψm) levels under inflammatory conditions, potentially alleviating muscle atrophy induced by mitochondrial dysfunction.

Sepsis-induced sarcopenia is a disease in which sepsis causes a loss of muscle mass and strength; this condition is considered an important model for studying the effects of systemic inflammatory responses on muscle (30). Therefore, the potential preventive impact of VSE treatment on muscle atrophy was investigated in a sepsis-induced sarcopenia mouse model, since in vitro analysis revealed that VSE treatment reduced pyroptosis and muscle atrophy under LPS-induced inflammatory conditions. To this end, 2.5 and 5 mg/kg/day VSE was orally administered daily to 5-week-old male C57BL/6J mice (n=6). On the 8th day, 1 mg/kg LPS was administered intraperitoneally. Body weight was measured at specific time points (days 0, 2, 4, 6, 8 and 9). The decrease in body weight observed in all groups after day 8 can be partially attributed to pre-sacrifice fasting, while the greater reduction in the LPS group may represent LPS-induced catabolic effects. Although not statistically analyzed, VSE treatment appeared to partially restore body weight compared with the LPS group (Fig. 6A). The decrease in body weight and TA tissue weight induced by LPS was slightly restored by VSE treatment. In particular, the reduction in tissue weight caused by LPS in the tibialis anterior muscle and the recovery caused by VSE were both significant (Fig. 6B). GAS muscle tissue in the LPS-treated group comprised very narrow fibers, and LPS treatment reduced the muscle surface area significantly. Tissue patterns were severely disrupted by LPS treatment, as evidenced by a significant reduction in muscle fiber surface area. However, VSE administration (2.5 and 5 mg/kg/day) markedly restored the tissue morphology, with the muscle fiber surface area returning to levels comparable to the control group. These improvements were confirmed by quantitative analysis using ImageJ (Fig. 6C). LPS treatment significantly weakened the grip strength of mice; however, the grip strength was significantly improved by 5 µg/ml VSE treatment (Fig. 6D). These results suggested that VSE treatment may prevent sepsis-induced sarcopenia by improving muscle strength.

NLRP3 inflammasome activation during sepsis induces muscle cell pyroptosis, a key factor in accelerating muscle fiber damage and muscle strength loss (31). Pyroptosis may also promote sarcopenia by modulating the inflammatory microenvironment within muscles (32). Similar to the in vitro experiments, the regulatory effects of VSE treatment on the NLRP3 inflammasome were investigated in vivo. The protein expression levels of the proinflammatory cytokine TNF-α were reduced in the serum of mice following VSE administration (Fig. 7A), which also reduced the mRNA expression levels of muscle degradation factor MuRF1 in GAS tissues at 5 µg/ml of VSE (Fig. 7B). Additionally, the protein expression levels of the muscle protein degradation factors MuRF1 and MaFbx and the NLPR3 pathway-related factors NLRP3, GSDMD, cleaved-GSDMD, caspase-1 and cleaved-caspase-1 in GAS tissues were analyzed. While LPS treatment decreased myosin heavy chain (MyHC) expression, the treatment generally increased the expression levels of muscle protein degradation factors and NLRP3 pathway-related factors. Furthermore, VSE treatment restored the expression of MyHC and MafBx, and reduced the expression of MuRF1, NLRP3, cleaved-GSDMD and cleaved-caspase-1 to levels comparable with or lower than those observed in the LPS group, suggesting a protective effect against muscle atrophy and pyroptotic cell death (Fig. 7C). These results indicate that VSE may effectively counteract inflammatory muscle damage in vivo.