Short-chain fatty acids alleviate sarcopenia in rats via modulation of the NF-κB signaling pathway

Abstract

Sarcopenia, characterized by progressive impairment of muscle quantity and performance, is closely linked to chronic inflammatory responses and cell death pathways. Short-chain fatty acids (SCFAs), metabolites originating from the gut microbial community, have shown immunomodulatory effects. This investigation evaluated the protective role and mechanistic basis of SCFAs in an induced sarcopenia rat model, focusing on the NF-κB signaling cascade. A pharmacologically induced sarcopenia model was established in adult female SD rats (6 months old) through bilateral ovariectomy followed by dexamethasone injections. Animals were grouped as follows: Sham, Model, and SCFAs (administered 150 mM SCFAs in drinking water for 4 weeks). Muscle function (forelimb grip strength, gastrocnemius muscle index), histological changes (H&E), reactive oxygen species, apoptosis (TUNEL), inflammatory cytokines (ELISA), gene expression (RT-qPCR), and protein expression (immunoblotting) were evaluated to assess the impact of SCFAs on sarcopenia and its underlying mechanistic basis. Relative to the Model group, SCFAs significantly increased circulating SCFA levels, improved grip strength and gastrocnemius index, restored muscle fiber structure, and reduced reactive oxygen species levels and apoptosis. SCFAs were associated with reduced NF-κB pathway activation (p-IKKβ and p-p65), downregulated NLRP3 inflammasome components (NLRP3, ASC, cleaved-Caspase-1), and reduced pyroptosis (GSDMD-N expression) and inflammatory cytokines (IL-1β and IL-18). SCFAs ameliorated sarcopenia in a pharmacologically induced rat model by suppressing NF-κB/NLRP3-mediated inflammation and pyroptosis, highlighting their potential as a candidate nutritional intervention for muscle atrophy associated with inflammation, hormonal imbalance, or metabolic stress.

Supplementary Information

The online version contains supplementary material available at 10.1038/s41598-026-47873-0.

Introduction

As a prevalent clinical syndrome, sarcopenia involves a progressive reduction in skeletal muscle mass accompanied by diminished muscle performance and is commonly associated with aging as well as other pathological conditions such as hormonal imbalance and metabolic stress. Patients with sarcopenia are more susceptible to falls, fracture events, functional disability, and hospitalizations, often leading to loss of independence and elevated all-cause mortality rates¹. Epidemiological evidence shows that the prevalence of sarcopenia varies between approximately 10% and 27% among older adults aged 60 years and above, although it may also occur under conditions such as hormonal imbalance and metabolic disorders². The pathogenesis of sarcopenia is multifactorial, involving imbalances in protein turnover, mitochondrial dysfunction, hormonal decline, chronic low-grade inflammation, fat infiltration, and degeneration of neuromuscular junctions³.

In the past few years, increasing scientific interest has focused on the “gut–muscle axis” as a novel regulatory mechanism in sarcopenia. Conditions such as aging, hormonal imbalance, and metabolic stress are frequently accompanied by shifts in gut microbial composition, particularly a decline in beneficial bacteria such as Faecalibacterium and Lactobacillus. This dysbiosis can impair immune homeostasis and metabolic functions, while promoting systemic inflammation and oxidative stress, ultimately contributing to muscle atrophy and dysfunction⁴. Short-chain fatty acids (SCFAs)—for example, acetate, propionate, and butyrate—represent crucial metabolites derived via gut microbial fermentation of dietary fiber⁵. SCFAs exhibit multiple biological functions, including maintaining intestinal barrier integrity, modulating immune responses, reducing oxidative stress, and regulating energy metabolism⁶. These functions suggest that SCFAs may exert protective effects on skeletal muscle health, especially through their involvement in the gut–immune–muscle regulatory network^7,8. However, how SCFAs mechanistically influence muscle wasting, particularly in the context of sarcopenia, remains incompletely understood.

The nuclear factor kappa-light-chain-enhancer of activated B cells (NF-κB) signaling pathway plays a central role in regulating inflammation, oxidative stress, and apoptosis in various tissues, including skeletal muscle⁹. Sustained activation of NF-κB has been linked to upregulation of pro-inflammatory mediators, elevated proteolysis, impaired muscle regeneration, and myofiber apoptosis¹⁰. Furthermore, NF-κB is an upstream activator of the nucleotide-binding oligomerization domain-like receptor protein 3 (NLRP3) inflammasome, a central mediator of pyroptosis—a pro-inflammatory mode of programmed cell death¹¹. Upon NLRP3 activation, pro-caspase-1 and gasdermin D (GSDMD) undergo cleavage, facilitating pore formation in the plasma membrane via GSDMD-N fragments, which induces release of pro-inflammatory cytokines (IL-1β and IL-18), thereby exacerbating tissue damage and inflammation¹².

In this research, we developed a pharmacologically induced sarcopenia model in rats through ovariectomy combined with dexamethasone administration to investigate the influence of SCFAs on muscle mass, strength, tissue morphology, apoptosis, and pyroptosis. We further explored the regulatory role of SCFAs in modulating the NF-κB/NLRP3 signaling axis. Through functional, histological, and molecular analyses, this work seeks to define the protective mechanistic basis of SCFAs in sarcopenia and offer a theoretical and experimental foundation for developing SCFA-based nutritional interventions against sarcopenia and muscle atrophy related to hormonal deficiency or metabolic stress.

Materials and methods

Experimental animals and housing

Twenty healthy female Sprague-Dawley (SD) rats (6 months old, 285.5 ± 35.2 g) were obtained from the same institution and housed under specific pathogen-free (SPF) conditions (temperature: 20–25 °C; humidity: 50%–60%; 12-hour light/dark cycle, 08:00–20:00). All procedures were approved by the Animal Ethics Committee of Bestcell Model Biological Center(BSMS-2024-07-31 A). The study was conducted in compliance with the ARRIVE guidelines (Animal Research: Reporting of In Vivo Experiments). Rats were acclimated for 7 days before experimentation.

Sarcopenia was induced by bilateral ovariectomy followed by dexamethasone injection. Fifteen rats were assigned at random to three groups (n = 5 per group): Sham, Model, and short-chain fatty acids (SCFAs). Rats were anesthetized prior to surgery according to standard laboratory procedures, and aseptic techniques were strictly followed. Both ovaries were exposed through a midline abdominal incision, ligated, and excised. Sham-operated rats received the identical surgical procedure without ovary removal. All animals were provided a 7-day recovery interval with ad libitum food and water. On postoperative day 7, Model and SCFAs groups received subcutaneous injections of dexamethasone (5 mg/kg, Sigma-Aldrich, Cat. No. D4902) once daily for 7 days. The Sham group received no treatment. After dexamethasone administration, rats in the SCFAs group were provided with drinking water containing 150 mM SCFAs once per week for four weeks. The SCFA mix included sodium acetate (600 mg/kg, Solarbio, Cat. No. A1070), sodium propionate (200 mg/kg, YuanYe Bio, Cat. No. C1434005), and sodium butyrate (200 mg/kg, Macklin, Cat. No. C14314005).

Successful model establishment was confirmed when the gastrocnemius muscle index (GMI), calculated as the bilateral gastrocnemius weight-to-body weight ratio, was over two standard deviations (SDs) lower than the mean value of the Sham group.

Grip strength and body weight measurement

After confirming successful model induction, forelimb grip strength was assessed with a digital grip strength meter (Model 47200, UGO Basile, Italy). All animals were positioned on the grip grid and permitted to grasp it with only the forepaws while maintaining a horizontal body position. The tail was gently pulled backward until the animal released its grip, and the maximal force value was documented. This procedure was repeated five times per rat, and the average of the five readings was used as the final grip strength value.

Body weight was measured using an electronic scale before sacrifice. The GMI was calculated as the total weight of bilateral gastrocnemius muscles divided by the body weight. These assessments were performed to assess alterations in muscle function as well as mass.

Gastrocnemius muscle acquisition and histological assessment

At the end of the experiment, rats were deeply anesthetized and euthanized by intraperitoneal administration of sodium pentobarbital (150 mg/kg). Blood samples were collected via cardiac puncture. Bilateral gastrocnemius muscles were carefully dissected, rinsed in PBS to remove blood and adipose tissue, and weighed using an electronic balance. The GMI was calculated as the combined weight of the bilateral gastrocnemius muscles divided by the body weight.

For tissue histology, muscle specimens were formalin-fixed, alcohol-dehydrated, paraffin-embedded, and sectioned. Paraffin sections were deparaffinized using xylene (15 min ×2), rehydrated through graded ethanol (100%, 90%, and 75%, 5 min per step), and washed with tap water. Hematoxylin and eosin (H&E) staining was then performed. Nuclei received hematoxylin staining (3 min), were differentiated in 1% acid alcohol, and subsequently blued in 1% ammonia water. After rinsing, sections received eosin counterstaining for 1–3 min, ethanol-dehydrated, xylene-cleared, and mounted with neutral balsam.

Stained sections were observed and imaged using a light microscope (Olympus Corporation, Japan) under identical acquisition settings. For quantitative analysis, at least five randomly selected non-overlapping fields per section were analyzed. The degree of inflammatory cell infiltration was evaluated using a semi-quantitative scoring system based on the number of infiltrating inflammatory cells as follows: 0, no or minimal inflammatory cell infiltration; 1, mild infiltration with a small number of scattered inflammatory cells; 2, moderate infiltration with increased inflammatory cell accumulation; and 3, severe infiltration with extensive inflammatory cell aggregation. Image analysis was performed using ImageJ software (NIH, USA).

Terminal deoxynucleotidyl transferase dUTP nick end labeling (TUNEL) staining

To evaluate apoptotic cell death in gastrocnemius muscle tissue, paraffin-embedded sections were processed with TUNEL staining. Once deparaffinized and rehydrated, sections were treated with proteinase K (20 µg/mL; Solarbio Life Sciences, Beijing, China) at 37 °C for 30 min to enhance tissue permeability. TUNEL staining was conducted using a One Step TUNEL Apoptosis Assay Kit (Beyotime Biotechnology, China; Cat. No. C1088) in accordance with the manufacturer’s protocol. Sections were incubated with the TUNEL reaction solution containing terminal deoxynucleotidyl transferase (TdT) and fluorescein-dUTP at 37 °C for 1 h under humid conditions. Following washing, nuclei underwent DAPI counterstaining, and slides were mounted in anti-fade medium.

Stained sections were observed and imaged using a fluorescence microscope (Olympus, Japan) under identical acquisition settings. For quantitative analysis, at least five randomly selected non-overlapping fields per section were analyzed using ImageJ software (NIH, USA). The apoptotic index was calculated as the ratio of TUNEL-positive nuclei to the total number of nuclei.

MitoSOX Red staining for mitochondrial ROS detection

Frozen gastrocnemius muscle sections were incubated with MitoSOX Red (Servicebio, China; Cat. No. GMS10124) at a final concentration of 5 µM for 45 min at 37 °C in the dark after PBS washing. Sections were then rinsed with PBS and counterstained with Hoechst 33,342 for 10 min. After mounting with anti-fade medium, images were captured using a fluorescence microscope (Olympus, Japan) under identical acquisition settings. For quantitative analysis, at least five randomly selected non-overlapping fields per section were analyzed using ImageJ software (NIH, USA), and the mean fluorescence intensity was calculated to represent mitochondrial ROS levels.

Cell culture and treatment

Rat skeletal muscle L6 myoblasts were cultured in DMEM supplemented with 10% fetal bovine serum at 37 °C with 5% CO₂. For differentiation, cells were grown to 90% confluence and switched to DMEM containing 2% horse serum.

Differentiated myotubes were treated as follows: control (vehicle), dexamethasone (DEX, 1 µM, 24 h), DEX combined with short-chain fatty acids (SCFAs; acetate, propionate, and butyrate, 0.5 mM each) following 1 h pretreatment, or with pathway modulators including BAY 11-7082 (10 µM), MCC950 (1 µM), or nigericin (5 µM). Samples were collected for subsequent analyses.

Cell viability assay (CCK-8)

Cell viability was assessed using a Cell Counting Kit-8 (CCK-8, Beyotime, China). Cells were seeded into 96-well plates at a density of 5 × 10⁴ cells/mL (100 µL per well) and incubated for 24 h at 37 °C with 5% CO₂. After treatment, 10 µL of CCK-8 solution was added to each well, followed by incubation for 2 h. The absorbance at 450 nm was measured using a microplate reader. Cell viability was calculated relative to the control group. All experiments were performed in triplicate.

Enzyme-linked immunosorbent assay (ELISA) for Inflammatory Cytokines

Interleukin-1β (IL-1β) and interleukin-18 (IL-18) levels were quantified using commercial ELISA kits according to the manufacturer’s instructions.

For in vivo experiments, gastrocnemius muscle tissues were collected, rinsed with cold PBS, homogenized in PBS (1:9, w/v), and centrifuged at 5,000 × g for 10 min at 4 °C. The supernatants were collected for ELISA measurement to assess tissue inflammatory cytokine levels. For in vitro experiments, culture supernatants from L6 myotubes were collected and centrifuged to remove debris prior to analysis, and were used for ELISA detection of cytokine secretion.

ELISA kits for IL-1β (Cat. No. PI301) and IL-18 (Cat. No. PI518) were obtained from Beyotime Biotechnology (China). Absorbance was measured at 450 nm using a microplate reader (Cmax Plus, Molecular Devices, USA), and cytokine concentrations were calculated based on standard curves.

Determination of SCFAs by GC–MS

Serum levels of short-chain fatty acids (SCFAs), including acetate, propionate, and butyrate, were measured using gas chromatography–mass spectrometry (GC–MS). Blood samples were collected from the abdominal aorta after fasting and centrifuged at 3,000 rpm for 15 min to obtain serum, which was stored at − 80 °C until analysis.

For sample preparation, serum samples were subjected to acidification and derivatization using propanol/pyridine and propyl chloroformate (PCF), followed by extraction with n-hexane. After centrifugation, the organic phase containing SCFA derivatives was collected and analyzed using a GC–MS system equipped with an autosampler and capillary column.

Quantification was performed using external standard curves prepared with acetate, propionate, and butyrate standards. The total SCFA concentration was calculated as the sum of individual components.

Quantitative real-time PCR (RT-qPCR)

Extraction of total RNA from gastrocnemius muscle tissues was achieved with TRIzol (Invitrogen, USA), using the standard phenol–chloroform procedure. RNA quality and quantity were determined via spectrophotometry, and 1 µg of total RNA was employed as the template for each reaction.

Quantitative real-time PCR was carried out with the UniPeak U + One Step RT-qPCR SYBR Green Kit (Yujia Biotechnology, Guangzhou, China) on a StepOnePlus System (Thermo Fisher Scientific, USA). Each 20 µL reaction mixture consisted of 10 µL 2× SYBR Mix, 0.4 µL of each primer (10 µM), 2 µL RNA, and 7.2 µL RNase-free water. Cycling parameters were set as 55 °C for reverse transcription, 95 °C for pre-denaturation, then 40 cycles of denaturation (95 °C) and annealing/extension (60 °C). Specificity was confirmed by performing a melting-curve analysis after each run.

The expression levels of ASC, NLRP3, and GAPDH were analyzed through the 2^–ΔΔCt method (Table 1). GAPDH functioned as the internal standard. Each assay was performed three times.

Table 1.

Primer sequences were as follows:

Gene	Primer	Sequence (5’–3’)
ASC	F	TCGTCAGCTACTATCTGGAG
	R	TCCAGCAAACCATCAACTTCTG
NLRP3	F	CCAACCATTCATACGCACTC
	R	TCCAGCATCTGTAGCTTACAG
GAPDH	F	ACTCCCTCAAGATTGTCAGC
	R	AGTTGCTGTTGAAGTCACAGG

Western blot analysis

Gastrocnemius muscle tissues were lysed in RIPA buffer containing protease and phosphatase inhibitors. After centrifugation (12,000 ×g, 15 min, 4 °C), the supernatants were collected, and protein concentrations were determined using a BCA Kit (Beyotime, China). Equal amounts of protein were separated by 12% SDS-PAGE and transferred onto PVDF membranes (Millipore, USA).

Each membrane was blocked with 5% non-fat milk in TBST for 1 h at room temperature and then incubated overnight at 4 °C with primary antibodies against p-IKKβ (1:1000, Cat. No. AF3009), IKKβ (1:1000, Cat. No. AF6009) (both from Affbiotech, China); p-NF-κB p65 (1:1000, Cat. No. #3033), NF-κB p65 (1:3000, Cat. No. #8242) (both from CST, USA); NLRP3 (1:1000, Cat. No. ab263899, Abcam, UK), ASC (1:1000, Cat. No. ab155970, Abcam, UK), cleaved Caspase-1 (1:2000, Cat. No. AF4005, Affinity, China), GSDMD (1:2000, Cat. No. ab219800, Abcam, UK), GSDMD-N (1:70000, Cat. No. A24059, Abclonal, China), and GAPDH (1:10000, Cat. No. 81640-5-RR, Proteintech, China) as the internal control.

After washing with TBST, membranes were incubated with HRP-conjugated goat anti-rabbit IgG secondary antibody (1:3000, Cat. No. bs-0295G-HRP, BIOSS, China) for 1 h at room temperature. Protein bands were visualized using an enhanced chemiluminescence kit (Beyotime, China) and captured using a chemiluminescence imaging system (Olympus, Japan). Band intensities were quantified using AlphaEase FC software (Alpha Innotech, USA).

Statistical analysis

Analyses of statistical data were conducted through SPSS 25.0 (IBM, USA) and summarized as mean ± SD. Differences among groups were evaluated using one-way analysis of variance (ANOVA), followed by Tukey’s post hoc multiple comparisons test for pairwise comparisons. P < 0.05 was regarded as significant.

Results

SCFAs supplementation increases circulating SCFA levels

As shown in Fig. 1, serum levels of major SCFA components, including acetic acid, propionic acid, and butyric acid, were measured to verify the effectiveness of SCFAs administration. No significant differences were observed between the Sham and Model groups. In contrast, SCFAs supplementation significantly increased the concentrations of acetic acid, propionic acid, and butyric acid compared with the Model group (all P < 0.05). These findings indicate that oral SCFAs supplementation effectively elevates circulating SCFA levels in vivo.

Fig. 1.

SCFAs supplementation increases circulating SCFA levels in rats. Serum concentrations of acetic acid, propionic acid, and butyric acid in the Sham, Model, and SCFAs groups. Data are presented as mean ± SD (n = 5 per group). *P < 0.05, **P < 0.01 vs. Sham;#P < 0.05, ##P < 0.01 vs. Model.

SCFAs improve muscle strength, mass, and histological integrity in sarcopenic rats

Following confirmation of successful SCFAs supplementation, we next evaluated its effects on muscle function and morphology in sarcopenic rats.

As shown in Fig. 2A, forelimb grip strength was markedly reduced in the Model group compared to the Sham group (P < 0.001), indicating significant functional impairment. SCFAs treatment significantly increased grip strength in contrast to the Model rats (P < 0.01), suggesting improved muscle performance.

Fig. 2.

SCFAs improve muscle function, mass, and morphology in sarcopenic rats. (A) Forelimb grip strength; (B) Gastrocnemius muscle index (GMI); (C) Representative H&E staining images of gastrocnemius muscle (upper: 50 μm; lower: 20 μm) and quantification of inflammatory cell infiltration score. Data are presented as mean ± SD (n = 5 per group). *P < 0.05, ^**P < 0.01, ^***P < 0.001 vs. Sham; ^#P < 0.05, ^##P < 0.01 vs. Model.

Similarly, the GMI was substantially lower among Model rats relative to the Sham controls (P < 0.01), while the SCFAs group revealed a notable elevation in GMI relative to the Model rats (P < 0.05), reflecting recovery of muscle mass (Fig. 2B).

H&E staining revealed clear histological differences among the groups (Fig. 2C). Sham rats displayed well-organized, intact muscle fibers without inflammatory infiltration. In contrast, the Model group showed irregular and atrophic fibers with evident cellular infiltration and partial necrosis. Notably, SCFA-treated rats exhibited improved fiber alignment, reduced muscle atrophy, and a marked decrease in inflammatory cell infiltration score relative to the Model rats (all P < 0.05).

These findings suggest that SCFAs alleviate muscle atrophy, restore muscle strength, and improve tissue structure in rats with sarcopenia.

SCFAs attenuate apoptosis and mitochondrial ROS accumulation in skeletal muscle

To further explore the mechanisms underlying the protective effects of SCFAs on skeletal muscle, apoptosis and mitochondrial reactive oxygen species (ROS) levels were evaluated using TUNEL staining and MitoSOX fluorescence, respectively. As shown in Fig. 3A, the Model group exhibited a marked increase in TUNEL-positive cells compared with the Sham group (P < 0.001), indicating enhanced apoptosis in skeletal muscle. Notably, SCFAs treatment significantly reduced the apoptosis rate relative to the Model group (P < 0.01).

Fig. 3.

SCFAs attenuate apoptosis and mitochondrial ROS accumulation in skeletal muscle. (A) TUNEL staining showing apoptotic cells in skeletal muscle tissues from the Sham, Model, and SCFAs groups, along with quantitative analysis of apoptosis rate. (B) MitoSOX staining showing mitochondrial ROS levels in skeletal muscle, with corresponding fluorescence intensity quantification. Data are presented as mean ± SD (n = 5 per group). **P < 0.01, ***P < 0.001 vs. Sham; ###P < 0.001 vs. Model.

Consistently, MitoSOX staining revealed a substantial elevation of mitochondrial ROS levels in the Model group compared with the Sham group (P < 0.001) (Fig. 3B). In contrast, SCFAs supplementation markedly decreased mitochondrial ROS accumulation compared with the Model group (P < 0.001).

Together, these findings suggest that SCFAs may reduce skeletal muscle apoptosis and oxidative stress, indicating a potential role in mitigating mitochondrial dysfunction in sarcopenic rats.

SCFAs Inhibit NF-κB–mediated NLRP3 inflammasome activation in skeletal muscle

To clarify how SCFAs ameliorate sarcopenia, we examined the NF-κB signaling cascade and NLRP3 inflammasome signaling activity in gastrocnemius muscle specimens.

Immunoblotting results indicated that phosphorylated IKKβ and NF-κB p65 levels were notably enhanced in Model rats relative to the Sham rats (P < 0.01 ), indicating NF-κB pathway activation. SCFA treatment significantly decreased p-IKKβ/IKKβ and p-NF-κB p65/p65 ratios versus the Model rats (P < 0.05) (Fig. 4A).

Fig. 4.

SCFAs inhibit NF-κB/NLRP3 inflammasome pathway activation in skeletal muscle. (A) Immunoblot detection and densitometric analysis of p-IKKβ, IKKβ, p-p65 and total p65. (B) Transcript levels of NLRP3 and ASC. (C) Immunoblot evaluation of NLRP3, ASC, and cleaved Caspase-1 proteins. (D) Caspase-1 enzymatic activity in gastrocnemius muscle. Results are reported as mean ± SD (n = 5 per group). ^*P < 0.05, ^**P < 0.01, ^***P < 0.001 vs. Sham; ^#P < 0.05, ^##P < 0.01, ^###P < 0.001 vs. Model.

RT-qPCR results markedly elevated mRNA levels of NLRP3 and ASC in Model rats relative to Sham rats (P < 0.001), whereas SCFAs treatment markedly downregulated both transcripts (P < 0.001) (Fig. 4B).

Consistent with the transcriptional changes, Western blotting demonstrated an elevated abundance of NLRP3, ASC, and cleaved Caspase-1 among Model rats (P < 0.01). SCFA administration notably suppressed NLRP3 and ASC levels (P < 0.05) and markedly reduced cleaved Caspase-1 levels (P < 0.05) (Fig. 4C).

Furthermore, Caspase-1 activity was markedly higher in Model rats than in Sham rats (P < 0.01), whereas SCFAs notably suppressed Caspase-1 activity (P < 0.05) (Fig. 4D).

Observations above suggest that SCFAs inhibit the NF-κB/NLRP3/Caspase-1 signaling axis, thereby suppressing inflammation and pyroptosis in skeletal muscle of sarcopenic rats.

SCFAs alleviate pyroptosis and inflammation-related cytokine secretion

To determine the influence of SCFAs on pyroptosis and associated inflammation, we assessed the expression of GSDMD and its cleaved form GSDMD-N, along with the levels of IL-1β and IL-18 in gastrocnemius muscle homogenates.

As shown in Fig. 5, the protein level of full-length GSDMD remained comparable among groups, whereas cleaved GSDMD-N was significantly increased in the Model group compared with the Sham group (P < 0.05), indicating enhanced pyroptotic activity. SCFAs treatment markedly reduced GSDMD-N expression relative to the Model group (P < 0.05) (Fig. 5A, B).

Fig. 5.

SCFAs treatment attenuates pyroptosis and pro-inflammatory cytokine secretion in sarcopenic rats. (A) Western blotting of GSDMD and GSDMD-N. (B) Densitometric analysis of GSDMD and GSDMD-N proteins. (C) ELISA-based measurement of IL-1β and IL-18 in gastrocnemius muscle homogenates. Results are reported as mean ± SD (n = 5 per group). ^*P < 0.05, ^***P < 0.001 vs. Sham; ^#P < 0.05, ^##P < 0.01 vs. Model.

Furthermore, ELISA analysis demonstrated that IL-1β and IL-18 levels in gastrocnemius muscle homogenates were significantly elevated in the Model group compared with the Sham group (P < 0.001), reflecting a pronounced inflammatory response. Notably, SCFAs treatment significantly decreased the levels of both cytokines, although they remained higher than those in the Sham group (P < 0.01), indicating partial attenuation of inflammation (Fig. 5C).

Collectively, these results suggest that SCFAs suppress pyroptosis by inhibiting GSDMD cleavage and reducing the release of key inflammatory mediators, thereby alleviating inflammatory damage in skeletal muscle of sarcopenic rats.

SCFAs attenuate DEX-induced pyroptosis via the NF-κB/NLRP3 axis in L6 myotubes

To further elucidate the underlying mechanism, in vitro experiments were performed in differentiated L6 myotubes using dexamethasone (DEX) stimulation combined with SCFAs and pathway modulators.

As shown in Fig. 6A, ELISA results demonstrated that DEX treatment significantly increased the secretion of IL-1β and IL-18 compared with the Control group (P < 0.001). SCFAs markedly reduced the levels of both cytokines (P < 0.001). Similar reductions were observed in the BAY 11-7082 and MCC950 groups, whereas the addition of nigericin partially reversed the inhibitory effects of SCFAs (P < 0.001).

Fig. 6.

SCFAs attenuate DEX-induced pyroptosis and inflammatory responses via the NF-κB/NLRP3 signaling pathway in L6 myotubes. (A) ELISA analysis of IL-1β and IL-18 levels in the culture supernatants of L6 myotubes under different treatments. (B) Cell viability assessed by CCK-8 assay. (C) Western blot analysis and corresponding densitometric quantification of p-p65, NLRP3, cleaved-Caspase-1, and GSDMD-N protein expression. Cells were treated as follows: Control, DEX (1 µM), DEX + SCFAs, DEX + BAY 11-7082, DEX + MCC950, and DEX + SCFAs + nigericin. Data are presented as mean ± SD (n = 3 independent experiments). ***P < 0.001 vs. Control; ###P < 0.001 vs. DEX; &&&P < 0.001 vs. DEX + SCFAs.

As shown in Fig. 6B, CCK-8 analysis revealed no significant differences in cell viability among all groups, indicating that the treatments did not induce cytotoxicity.

Consistent with these findings, Western blot analysis (Fig. 6C) showed that DEX markedly upregulated the expression of p-p65, NLRP3, cleaved-Caspase-1, and GSDMD-N compared with the Control group (P < 0.001). SCFAs significantly suppressed the expression of these proteins. BAY 11-7082 reduced p-p65 levels and concurrently decreased downstream NLRP3 inflammasome activation, whereas MCC950 inhibited NLRP3, cleaved-Caspase-1, and GSDMD-N expression without affecting p-p65 levels (P < 0.001). Importantly, nigericin treatment partially restored the expression of pyroptosis-related proteins in SCFAs-treated cells (P < 0.001).

Collectively, these results indicate that SCFAs attenuate DEX-induced inflammation and pyroptosis in skeletal muscle cells through inhibition of the NF-κB/NLRP3 signaling axis.

Discussion

In this study, a pharmacologically induced sarcopenia model was established, and SCFAs supplementation was observed to significantly elevate circulating SCFA levels, accompanied by improvements in muscle mass and grip strength, alleviation of histological muscle damage, reduction of oxidative stress and apoptosis, and suppression of inflammatory and pyroptosis-related markers. These findings suggest that SCFAs may exert protective effects against sarcopenia through multiple mechanisms.

Prolonged low-grade inflammatory activity represents a hallmark abnormality in sarcopenia pathophysiology¹³. The NF-κB signaling cascade serves a critical function in mediating the transcription of inflammatory mediators, and its overactivation has been linked to enhanced muscle protein degradation and atrophy^14,15. In our experiments, enhanced abundance of phosphorylated IKKβ and NF-κB p65 among the model rats indicated NF-κB pathway activation, while SCFAs treatment markedly suppressed this phosphorylation. These results, together with pharmacological modulation using BAY 11-7082, MCC950, and nigericin, suggest that SCFAs exert anti-inflammatory effects in skeletal muscle at least in part through inhibition of the NF-κB/NLRP3 signaling axis. Previous research has also demonstrated that SCFAs can inhibit NF-κB activity through G-protein–linked receptors (e.g., GPR41/GPR43) or through histone deacetylase (HDAC) inhibition^16,17, which may partially explain the inhibitory effects on NF-κB signaling observed in this study.

Pyroptosis, an inflammasome-dependent programmed cell death modality, has recently been implicated in various degenerative diseases^18,19. In the present study, NLRP3, ASC, cleaved caspase-1, and GSDMD-N were notably upregulated in the model rats, along with elevated IL-1β and IL-18 levels in muscle tissues and cell culture supernatants. These changes were notably reversed following SCFAs intervention, suggesting that SCFAs may suppress pyroptosis by inhibiting inflammasome assembly and downstream gasdermin D cleavage. Although direct evidence for this regulatory mechanism in skeletal muscle remains limited, existing literature supports that SCFAs modulate inflammasome activity²⁰, providing indirect support for our observations.

Oxidative stress is another major contributor to muscle degeneration and may activate NF-κB signaling and mitochondrial damage, ultimately leading to apoptosis^21,22. In our study, increased ROS accumulation and TUNEL-positive cells were observed in sarcopenic rats, whereas SCFAs markedly reduced ROS levels and apoptosis. These findings indicate that SCFAs may confer antioxidant and cytoprotective effects in skeletal muscle tissue.

It should be noted that several limitations exist in this study. First, the in vivo sample size was relatively small, and only female rats were included; thus, potential sex-related differences and broader population variability remain to be explored. Second, whether SCFAs act directly on skeletal muscle or indirectly via modulation of the gut microbiota requires further clarification, and alternative interpretations cannot be excluded, as the observed effects may be secondary to broader metabolic or oxidative stress changes rather than direct regulation of the NF-κB/NLRP3 pathway. Third, although multiple molecular indicators were used to assess pyroptosis, direct morphological evidence was not provided. Finally, although pharmacological modulators supported pathway involvement in vitro, causal relationships have not been fully validated in vivo. In addition, the translational relevance remains limited due to differences in dosage, administration route, and species-specific metabolism.

In conclusion, SCFAs were associated with multiple protective effects in sarcopenic rats, potentially through suppression of the NF-κB/NLRP3 inflammasome pathway, reduction of oxidative stress, and suppression of pyroptosis. These results offer early support for the potential use of SCFAs in sarcopenia; however, further mechanistic studies and clinical validation are necessary to fully elucidate their roles and application value.

Conclusion

This study demonstrated that SCFAs supplementation significantly elevated circulating SCFA levels and improved skeletal muscle mass, morphology, and strength in sarcopenic rats. Mechanistically, SCFAs alleviated muscle atrophy, at least in part, by suppressing NF-κB activation and inhibiting NLRP3 inflammasome assembly, thereby reducing Caspase-1 activation, GSDMD-N–mediated pyroptosis, and the release of inflammatory cytokines (IL-1β and IL-18). In addition, SCFAs decreased ROS accumulation and apoptosis in skeletal muscle, indicating antioxidative and cytoprotective effects. Collectively, these findings suggest that SCFAs mitigate skeletal muscle injury under sarcopenic conditions primarily through modulation of the NF-κB/NLRP3/GSDMD axis, supporting their potential as a metabolic intervention strategy. However, the direct targets of SCFAs and their interactions with gut microbiota and other regulatory pathways require further investigation.

Supplementary Information

Below is the link to the electronic supplementary material.

Author contributions

Conceptualization: HX; Methodology: ZHY, NC; Formal Analysis: ZHY, HX; Data curation: NC, HX; Writing - Original Draft: HX; Writing - Review and Editing: ZHY, NC; Funding acquisition: HX; Investigation: NC, HX; Supervision: ZHY, HX; Software: ZHY, HX; Visualization: ZHY, NC; Validation: ZHY, HX; All authors have agreed on the journal to which the article has been submitted.

Funding

This study is supported by Research Project of Xinjiang Uygur Autonomous People’s Hospital (20220205).

Data availability

The data used to support the findings of this study are available from the corresponding author upon request.

Declarations

Competing interests

The authors declare no competing interests.

Ethics approval and consent to participate

All procedures were approved by the Animal Ethics Committee of Bestcell Model Biological Center(BSMS-2024-07-31 A). The study was conducted in compliance with the ARRIVE guidelines (Animal Research: Reporting of In Vivo Experiments) and conducted in compliance with the institutional guidelines (Directive 2010/63/EU in Europe) for the care and use of animals.