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. 2025 Jun 18;26(4):120. doi: 10.1007/s10522-025-10262-7

β-hydroxy-β-methylbutyrate supplementation mitigates muscle atrophy induced by inactivity and protein deprivation

Xu He 1,2,3, Yan Li 1,2,3,✉, Jun Chen 1,2, Yan Huang 1,2, Ying Zhou 1,2, Yang Li 1,2, Jing Quan 1,2
PMCID: PMC12176917  PMID: 40531264

Abstract

Muscle atrophy, resulting from physical inactivity or protein deficiency, is a significant health concern. β-hydroxy-β-methylbutyrate (HMB) has potential in preserving muscle mass, but its mechanisms in various atrophy-inducing conditions are not fully understood. This study aimed to investigate HMB’s effects on muscle atrophy induced by inactivity and protein deprivation, and to elucidate the underlying molecular mechanisms. Rats were subjected to inactivity or protein-deficient diets with or without HMB supplementation. Muscle morphology, strength, and biochemical parameters were assessed. In vitro studies using C2C12 myoblasts and mouse skeletal muscle satellite cells exposed to interleukin-6 (IL-6) explored molecular pathways involved in HMB’s protective effects. Inactivity and protein deprivation led to muscle atrophy, reduced strength, and altered biochemical markers. HMB supplementation partially mitigated these effects, preserving muscle mass and function. HMB attenuated atrophy markers (Muscle Atrophy F-box and Muscle RING Finger 1 (MuRF1)) and maintained myogenic factor (Myogenin (MyoG)) levels. In vitro studies revealed that HMB’s protective effects were mediated through the AKT/mTOR pathway, with concurrent regulation of autophagy pathways and preservation of mitochondrial function in both myoblasts and satellite cells. HMB specifically protected satellite cell viability and function through AKT-dependent mechanisms, maintaining protein synthesis and reducing apoptosis under IL-6-induced stress conditions. HMB supplementation shows protective effects against muscle atrophy induced by inactivity and protein deprivation, through multiple mechanisms including AKT/mTOR pathway activation, autophagy regulation, and maintenance of mitochondrial function in both myoblasts and satellite cells. These findings suggest HMB as a potential therapeutic strategy for preventing muscle atrophy in various clinical scenarios.

Keywords: Muscle atrophy, HMB supplementation, Protein deprivation, Physical inactivity, AKT/mTOR pathway, Myogenesis

Introduction

Muscle atrophy, characterized by the loss of muscle mass and function, is a significant health concern with diverse etiologies (Bodine and Baehr 2014). This condition can result from various factors, including aging, disuse, malnutrition, and chronic diseases (Nunes et al. 2022). Muscle atrophy features reduced muscle fiber size, decreased protein content, and diminished force-generating capacity (Du et al. 2021). Risk factors encompass prolonged bed rest, immobilization, protein-deficient diets, and inflammatory conditions (Zhou et al. 2023). Current treatment interventions primarily focus on resistance exercise and nutritional supplementation (Huang et al. 2022a). However, these approaches have limitations, particularly in situations where physical activity is restricted or appetite is compromised.

Physical activity, protein intake, and chronic inflammation play crucial roles in muscle homeostasis and atrophy development (Ziaaldini et al. 2017). Inactivity leads to rapid muscle loss through reduced protein synthesis and increased proteolysis (Breen et al. 2013; Powers et al. 2020). Adequate protein consumption is essential for maintaining muscle mass, with deficiency resulting in accelerated atrophy (Carbone and Pasiakos 2019). Chronic inflammation, often associated with diseases such as cancer and rheumatoid arthritis, can exacerbate muscle wasting through pro-inflammatory cytokine signaling (Londhe and Guttridge 2015). For instance, IL-6 has been shown to induce muscle atrophy through activation of the JAK2/STAT3 pathway and subsequent suppression of protein synthesis via mTOR inhibition (Haddad et al. 2005). These factors modulate key pathways involved in muscle atrophy, including the ubiquitin–proteasome system, autophagy-lysosome pathway, and myostatin signaling (Vainshtein and Sandri 2020). For instance, Muscle Atrophy F-box (MAFbx) and Muscle RING Finger 1 (MuRF1) are well-established markers of muscle atrophy and essential components of the E3 ubiquitin ligase system responsible for protein degradation through ubiquitination. Their increased expression is consistently observed during muscle atrophy conditions and serves as a reliable indicator of muscle wasting (Bodine and Baehr 2014; Nunes et al. 2022).

The phosphatidylinositol 3-kinase (PI3K)/protein kinase B (AKT)/mammalian target of rapamycin (mTOR) signaling pathway plays a pivotal role in de novo protein synthesis and muscle cell homeostasis (Jaiswal et al. 2022). Activation of this pathway promotes protein synthesis through phosphorylation of downstream targets such as p70S6 kinase and 4E-BP1 (Huang et al. 2022b). For instance, insulin-like growth factor 1 (IGF-1) stimulates this pathway, leading to increased muscle protein synthesis and hypertrophy (Russell and Schneider 2020). Conversely, inhibition of PI3K/AKT/mTOR signaling, as observed during muscle atrophy, results in reduced protein synthesis and increased proteolysis (Castets et al. 2019). β-hydroxy-β-methylbutyrate (HMB), a metabolite of leucine, has gained attention for its potential to mitigate muscle atrophy (Phillips et al. 2022). HMB is synthesized primarily in the liver and muscle tissues, with approximately 5% of leucine converted to HMB in the body (Kaczka et al. 2019). Physiologically, HMB promotes protein synthesis, possibly through activation of the mTOR pathway (Eley et al. 2007). Additionally, HMB may inhibit protein degradation by reducing proteasome activity (Smith et al. 2005). These properties make HMB a promising candidate for preventing and treating muscle atrophy in various clinical scenarios (Holecek 2017).

The present study aimed to investigate the effects of β-hydroxy-β-methylbutyrate (HMB) supplementation on muscle atrophy induced by inactivity, protein deprivation and inflammation. We employed a combination of in vivo experiments using rat models subjected to inactivity or protein-deficient diets, and in vitro studies with C2C12 myoblasts and skeletal muscle satellite cells exposed to interleukin-6. Our findings demonstrated that HMB supplementation partially mitigates muscle atrophy through multiple mechanisms, including activation of the AKT/mTOR pathway, regulation of autophagy, and preservation of mitochondrial function. In both myoblasts and satellite cells, HMB maintained protein synthesis and reduced apoptosis under stress conditions through AKT-dependent mechanisms. These results suggest that HMB supplementation could be a promising therapeutic strategy for preventing muscle atrophy in various clinical scenarios.

Methods

Animals and experimental design

Male Sprague–Dawley rats (6 months old, 700–800 g) were obtained from Shanghai Laboratory Animal Center (Shanghai, China). Animals were housed in a temperature-controlled room (22 ± 2 °C) with a 12 h light/dark cycle and free access to water. All animal procedures were approved by the Institutional Animal Care and Use Committee of Medical Ethics Committee of The First People’s Hospital of Yunnan Province (NO.2020KYS074). Rats were randomly divided into five groups (n = 6 per group): Control, Inactive, Inactive + HMB, No-protein diet, and No-protein diet + HMB. Animals in the inactivity model group were maintained by suspending rats in specially designed cages that prevented hindlimb weight bearing while allowing free movement of the forelimbs for 30 days. Control and No-protein diet groups were housed in standard cages allowing free movement.

Diet composition and HMB supplementation

Control and Inactive groups received a standard diet (18% protein, 58% carbohydrate, 7.5% fat; Research Diets, Inc., New Brunswick, NJ, USA). The No-protein diet group received a protein-deficient diet (0.5% protein, 75.5% carbohydrate, 7.5% fat; Research Diets, Inc.). HMB supplementation groups received 340 mg/kg/day of calcium HMB (Ca-HMB, purity ≥ 98%, TSI Group Ltd., Shanghai, China) mixed into their respective diets. The Ca-HMB powder was thoroughly mixed with the diet at a concentration of 3.4 g/kg of feed, assuming an average daily food intake of 100 g/kg body weight for rats. Food intake was monitored daily, and all groups received similar amounts of food by weight (approximately 20–22 g/day/rat). After 30 days of experimental treatment, all animals underwent performance tests, followed by sample collection. The experimental treatments included standard diet (control), inactivity, inactivity with HMB supplementation, protein-deficient diet, and protein-deficient diet with HMB supplementation.

Grip strength test

Hindlimb grip strength was measured using a grip strength meter (Columbus Instruments, Columbus, OH, USA). Rats were gently pulled backward by the tail while grasping a metal grid attached to the force gauge. The maximum force exerted before releasing the grid was recorded. Five trials were performed for each rat with a 1 min rest between trials. The highest three values were averaged for analysis.

Exercise capacity test

Exercise capacity was assessed using a motorized treadmill (Columbus Instruments) set at a 35° incline. Rats were acclimatized to the treadmill for three days before testing. On the test day, rats ran at an initial speed of 10 m/min, with speed increased by 1 m/min every 2 min until exhaustion. Exhaustion was defined as the inability to run for 10 s despite mechanical prodding. Exercise time was recorded and normalized to the control group’s average time.

Sample collection and preparation

After 30 days of dietary intervention or 30 days of inactivity, rats were euthanized by CO2 inhalation. Blood was collected via cardiac puncture, allowed to clot for 30 min at room temperature, and centrifuged at 3000×g for 15 min at 4 °C. Serum was aliquoted and stored at − 80 °C until analysis. The gastrocnemius muscles were quickly dissected, weighed, and either snap-frozen in liquid nitrogen for protein analysis or fixed in 4% paraformaldehyde for histological examination.

Histological analysis

Fixed gastrocnemius muscles from hindlimb were embedded in paraffin, sectioned at 5 μm thickness, and stained with hematoxylin and eosin (H&E) using a commercial H&E Staining Kit (Abcam, Cambridge, UK) according to the manufacturer’s instructions. Images were captured using a light microscope (Olympus BX51, Tokyo, Japan) equipped with a digital camera (Olympus DP72).

Biochemical assays

Serum levels of alanine aminotransferase (ALT), aspartate aminotransferase (AST), albumin, creatinine (CREA), creatine kinase (CK), and testosterone were measured using commercial kits (Nanjing Jiancheng Bioengineering Institute, Nanjing, China) according to the manufacturer’s instructions. These markers were selected to assess liver function (ALT, AST), protein metabolism (albumin), kidney function (CREA), muscle damage (CK), and anabolic status (testosterone). IL-6 levels in serum and muscle homogenates were quantified using a rat IL-6 ELISA kit (R&D Systems, Minneapolis, MN, USA). TNF-α and CRP levels in muscle homogenates were measured using rat TNF-α ELISA kit and rat CRP (C-Reactive Protein) ELISA kit (both from R&D Systems, Minneapolis, MN, USA) to evaluate the inflammatory status during muscle atrophy and the anti-inflammatory effects of HMB supplementation.

Western blot analysis

Frozen muscle samples or cell samples were homogenized in RIPA buffer (Beyotime Biotechnology, Shanghai, China) containing protease and phosphatase inhibitors. Protein concentrations were determined using the BCA method (Thermo Fisher Scientific, Waltham, MA, USA). Equal amounts of protein (30 μg) were separated by SDS-PAGE and transferred to PVDF membranes. Membranes were blocked with 5% non-fat milk and incubated overnight at 4 °C with primary antibodies against Muscle Atrophy F-box (FBXO32/MAFbx, PA5-19056, Thermo Fisher Scientific), Muscle RING Finger 1 (MuRF1, ab313852, Abcam), Myogenin (MyoG, sc-12732, Santa Cruz), phospho-AKT (Ser473, #4060), total AKT (#4691), phospho-mTOR (Ser2448, #5536), total mTOR (#2983), β-actin (#4970), total ULK1 (#8054), cleaved LC3B (LC3B-II, SAB1305635, Sigma-Aldrich), Beclin1 (#3495), ATG7 (#8558), p62 (#5114), phospho-AMPKα (Thr172, #2535), total AMPKα (#5831), phospho-FoxO3a (Ser253, #9466), total FoxO3a (#2497), phospho-JAK2 (Tyr1007/1008, #3776), total JAK2 (#3230), phospho-STAT3 (Tyr705, #9145), total STAT3 (#4904), and IL-6 (#12912) (unless otherwise noted, all from Cell Signaling Technology, Danvers, MA, USA). After washing, membranes were incubated with HRP-conjugated secondary antibodies. Protein bands were visualized using an ECL detection system (Millipore, Billerica, MA, USA) and quantified using Image J software.

Cell culture and treatments

C2C12 myoblasts (ATCC, Manassas, VA, USA) were cultured in DMEM supplemented with 10% fetal bovine serum and 1% penicillin/streptomycin at 37 °C in a 5% CO2 incubator. Mouse skeletal muscle satellite cells (SunCell Biotech, Wuhan, China) were maintained in specific complete culture medium for mouse skeletal muscle satellite cells (SNPM-M084, SunCell Biotech) at 37 °C in a 5% CO2 incubator. Both cell types were seeded at a density of 1 × 104 cells/cm2 for all experiments unless otherwise stated. Cells were treated with 20 ng/ml recombinant mouse IL-6 (R&D Systems) for 72 h, with or without 50 μM HMB (Sigma-Aldrich, St. Louis, MO, USA). For AKT inhibition experiments, 1 μM MK-2206 (Selleck Chemicals, Houston, TX, USA) was added simultaneously with HMB treatment.

Protein synthesis assay

De novo protein synthesis was measured using the Click-iT Plus OPP Protein Synthesis Assay Kit (Thermo Fisher Scientific) according to the manufacturer’s instructions. Briefly, C2C12 cells were seeded in 96-well plates at a density of 5 × 103 cells/well. After treatments, cells were incubated with O-propargyl-puromycin (OPP) for 30 min, fixed with 4% paraformaldehyde, and permeabilized with 0.5% Triton X-100. The Click-iT reaction cocktail was then added to detect OPP incorporation. Nuclei were counterstained with Hoechst 33,342. Fluorescence images were captured using an EVOS 5000 Auto Imaging System (Thermo Fisher Scientific) with appropriate filter sets for OPP (GFP channel, excitation/emission: 470/525 nm) and Hoechst (DAPI channel, excitation/emission: 357/447 nm). At least five random fields per well were imaged using a 20 × objective.

Cell proliferation assay

Cell proliferation was assessed using the Cell Counting Kit-8 (CCK-8) (Dojindo Molecular Technologies, Kumamoto, Japan). C2C12 cells were seeded in 96-well plates at a density of 2 × 103 cells/well. After treatments, 10 μL of CCK-8 solution was added to each well and incubated for 2 h at 37 °C. Absorbance was measured at 450 nm using a microplate reader. Measurements were taken at 24, 48, and 72 h after treatment initiation.

Apoptosis assay

Apoptosis was evaluated using the Annexin V-FITC Apoptosis Detection Kit (BD Biosciences, San Jose, CA, USA). C2C12 cells were seeded in 6-well plates at a density of 2 × 105 cells/well. After treatments, cells were harvested by trypsinization, washed twice with cold PBS, and resuspended in 1X binding buffer. Cells were then stained with 5 μL Annexin V-FITC and 5 μL propidium iodide (PI) for 15 min at room temperature in the dark. Stained cells were analyzed within 1 h using a FACSCalibur flow cytometer (BD Biosciences). A minimum of 10,000 events were recorded for each sample. The percentage of early apoptotic (Annexin V-FITC positive, PI negative) and late apoptotic (Annexin V-FITC positive, PI positive) cells was calculated using FlowJo software (Tree Star, Ashland, OR, USA).

Mitochondrial analysis

For mitochondrial morphology assessment, cells were stained with 500 nM MitoTracker Green FM (Thermo Fisher Scientific, Waltham, MA, USA) for 30 min at 37 °C in the dark. Nuclei were counterstained with Hoechst 33,342 (5 μg/mL, Thermo Fisher Scientific) for 10 min at room temperature. Images were captured using an EVOS 5000 Auto Imaging System (Thermo Fisher Scientific) with appropriate filter sets for MitoTracker Green (excitation/emission: 490/516 nm) and Hoechst (excitation/emission: 350/461 nm).

For flow cytometric analysis of mitochondrial mass and membrane potential, cells were harvested by trypsinization and stained with 200 nM MitoTracker Green FM and 50 nM MitoTracker Deep Red FM (Thermo Fisher Scientific) for 30 min at 37 °C in the dark. After washing twice with PBS, cells were analyzed using a FACSCalibur flow cytometer (BD Biosciences, San Jose, CA, USA). MitoTracker Green FM signal was detected in the FL1 channel (488 nm excitation, 530/30 nm emission) for mitochondrial mass measurement, while MitoTracker Deep Red FM signal was detected in the FL4 channel (635 nm excitation, 661/16 nm emission) for membrane potential assessment. Data were analyzed using FlowJo software (Tree Star, Ashland, OR, USA).

Statistical analysis

Data are presented as mean ± standard deviation (SD). Statistical analyses were performed using GraphPad Prism 8.0 (GraphPad Software, San Diego, CA, USA). One-way ANOVA followed by Tukey’s post hoc test was used for multiple group comparisons. P < 0.05 was considered statistically significant.

Results

Muscle atrophy and altered molecular signaling in response to inactivity

In order to assess the effects of physical inactivity on muscle morphology and related molecular pathways, rats were divided into control and inactive groups, with the inactive group subjected to restricted movement for 30 days. Analysis of gastrocnemius muscle diameter revealed significant differences between the control and inactive groups (Fig. 1A), with the inactive group showing a marked reduction. This finding was corroborated by the gastrocnemius muscle-to-body weight ratio data (Fig. 1B), which decreased from 0.56% in the control group to 0.35% in the inactive group, representing a 37.5% reduction. Histological examination of muscle fibers using H&E staining (Fig. 1C) further supported these findings. The control group exhibited larger, uniformly sized, and closely packed polygonal muscle fibers. In contrast, the inactive group showed smaller fiber diameters with greater size variability and more pronounced spaces between fibers, indicative of muscle atrophy. Western blot analysis (Fig. 1D) revealed changes of several key proteins related to muscle homeostasis in the inactive group compared to the control. Notably, MyoG (a myogenesis marker) expression significantly decreased, indicating reduced muscle growth signaling. Conversely, MAFbx and MuRF1, both markers of muscle atrophy, showed significant increases. The phosphorylation levels of both AKT and mTOR were significantly decreased. Moreover, the phosphorylation levels of both AKT and mTOR were significantly decreased (Fig. 1D). Additionally, measurement of interleukin-6 (IL-6) levels showed a substantial increase in the inactive group compared to the control group (Fig. 1E). These molecular changes suggest alterations in muscle protein synthesis, degradation pathways, and inflammatory responses due to inactivity, indicating of muscle atrophy.

Fig. 1.

Fig. 1

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Muscle atrophy and altered molecular signaling in response to inactivity. Rats were divided into control and inactive groups, with the inactive group subjected to restricted movement for 30 days. A Gastrocnemius muscle diameter comparison between control and inactive groups. B Gastrocnemius muscle-to-body weight ratio in control and inactive groups. C H&E staining of muscle fibers from control and inactive groups. Quantification is from 250 muscle fibers for each individual. D Western blot analysis of MyoG, MAFbx, MuRF1, phosphorylated AKT, and phosphorylated mTOR in control and inactive groups. E Interleukin-6 (IL-6) levels in control and inactive groups. (n = 6 animals in each group; *p < 0.05; **p < 0.01; ***p < 0.001; ****p < 0.0001)

Protein deprivation induces muscle atrophy and alters molecular signaling pathways despite maintained physical activity

Next, we examined the impact of protein deficiency on muscle health and related signaling pathways. Rats were fed with normal diet or protein-deficient diet for 30 days, with all animals being allowed to move and exercise freely. Protein deprivation significantly impacted muscle characteristics and molecular signaling pathways, as evidenced by our comprehensive analysis of muscle morphology, mass, and gene expression. Examination of gastrocnemius muscle diameter (Fig. 2A) revealed a marked reduction in the no-protein diet group compared to the normal diet group. This finding was further corroborated by the gastrocnemius muscle-to-body weight ratio (Fig. 2B), which decreased from 0.58% in the normal diet group to 0.31% in the no-protein diet group, indicating substantial muscle mass loss due to protein deficiency. Histological analysis provided visual confirmation of these changes (Fig. 2C), with the muscle tissues in no-protein diet group exhibiting smaller fiber diameters, increased size variability, and more pronounced interfibrillar spaces. At the molecular level, MyoG expression was significantly reduced, while the muscle atrophy markers (MAFbx and MuRF1) showed significant increases after protein deprivation (Fig. 2D). Additionally, the phosphorylation levels of both AKT and mTOR were significantly reduced in the no-protein diet group, indicating disruption of protein synthesis pathways. Furthermore, IL-6 levels were substantially elevated in the no-protein diet group, pointing to an increased inflammatory response (Fig. 2E). Collectively, these findings highlight the crucial role of dietary protein in maintaining muscle health and function, even when animals are allowed to move and exercise freely.

Fig. 2.

Fig. 2

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Protein deprivation induces muscle atrophy and alters molecular signaling pathways despite maintained physical activity. Rats were fed with normal diet or protein-deficient diet for 30 days, with all animals allowed to move and exercise freely. A Gastrocnemius muscle diameter comparison between normal diet and no-protein diet groups. B Gastrocnemius muscle-to-body weight ratio in normal diet and no-protein diet groups. C Histological analysis of muscle tissues from normal diet and no-protein diet groups. Quantification is from 250 muscle fibers for each individual. D Western blot analysis of MyoG, MAFbx, MuRF1, phosphorylated AKT, and phosphorylated mTOR in normal diet and no-protein diet groups. E IL-6 levels in normal diet and no-protein diet groups. (n = 6 animals in each group; *p < 0.05; **p < 0.01; ***p < 0.001; ****p < 0.0001)

HMB supplementation improves muscle atrophy and physiological alterations induced by inactivity and protein deprivation

To evaluate the effect of HMB on muscle performance under conditions of inactivity or protein deprivation, we conducted a comprehensive analysis across five groups: Control, Inactive, Inactive + HMB, No-protein diet, and No-protein diet + HMB (Fig. 3). Gastrocnemius muscle diameter measurements revealed significant reductions in all experimental groups compared to the control, with the most pronounced decrease observed in the no-protein diet group (Fig. 3A). HMB supplementation partially mitigated this reduction in both inactive and protein-deprived conditions. Hindlimb grip strength tests showed a similar pattern, with the control group exhibiting the highest strength, followed by progressive decreases in both the no-protein diet and inactive groups. Notably, HMB supplementation significantly attenuated strength loss in both inactive and protein-deprived conditions (Fig. 3B). Exercise capacity, as measured by relative exercise duration, decreased across all experimental groups, with the no-protein diet group showing the most severe reduction. HMB supplementation improved exercise capacity in both experimental groups (Fig. 3C). Biochemical analysis in the blood samples revealed alterations in various parameters across groups. ALT and AST levels were elevated in all experimental groups compared to control, suggesting potential hepatic stress. Serum albumin levels decreased in protein-deprived and inactive conditions, with HMB supplementation showing a protective effect. CREA, an indicator of muscle mass, showed significant decreases in both no-protein diet and inactive groups, while HMB supplementation tended to prevent its decrease. Furthermore, CK levels, indicative of muscle damage, were elevated in inactive and protein-deprived groups, with HMB supplementation appearing to attenuate this increase. Testosterone levels were reduced in all experimental groups, particularly in the no-protein diet condition, with HMB supplementation showing a modest protective effect (Fig. 3D). These results collectively demonstrate the detrimental effects of inactivity and protein deprivation on muscle mass, strength, and related physiological parameters, while also suggesting a potential protective role for HMB supplementation under these conditions.

Fig. 3.

Fig. 3

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HMB supplementation improves muscle atrophy and physiological alterations induced by inactivity and protein deprivation. Five experimental groups were examined: Control, Inactive, Inactive + HMB, No-protein diet, and No-protein diet + HMB. A Gastrocnemius muscle diameter measurements across all groups. B Hindlimb grip strength test results. C Relative exercise duration as a measure of exercise capacity. D Biochemical analysis of blood samples, including ALT, AST, serum albumin, CREA, CK, and testosterone levels. (n = 6 animals in each group; *p < 0.05; **p < 0.01; ***p < 0.001; ****p < 0.0001)

Molecular and histological analysis reveals protective effects of HMB supplementation on muscle homeostasis

To further elucidate the molecular mechanisms underlying the effects of HMB supplementation on muscle homeostasis during inactivity and protein deprivation, we analyzed various muscle-related parameters across our experimental groups (Fig. 4). The gastrocnemius muscle to body weight ratio decreased in all experimental groups, with the most severe reduction in the no-protein diet group and partial mitigation observed with HMB supplementation (Fig. 4A). Histological analysis revealed similar trends, with reduced muscle fiber size and increased atrophy in the inactive and protein-deprived groups. Notably, HMB supplementation demonstrated a rescuing effect, partially preserving muscle fiber morphology and size in both conditions (Fig. 4B).

Fig. 4.

Fig. 4

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Molecular and histological analysis reveals protective effects of HMB supplementation on muscle homeostasis. Analysis was performed across five groups: Control, Inactive, Inactive + HMB, No-protein diet, and No-protein diet + HMB. A Gastrocnemius muscle to body weight ratio. B Histological analysis of muscle fiber morphology and size. Quantification is from 250 muscle fibers for each individual. C Western blot analysis of MyoG, MAFbx, MuRF1, phosphorylated AKT, and phosphorylated mTOR. D ELISA analysis of IL-6 levels in muscle tissue and blood samples. E Analysis of IL-6 protein expression in muscle tissue. F Measurement of inflammatory markers TNF-α and CRP by ELISA in muscle tissues. G Western blot analysis of autophagy-related proteins (ULK1, LC3BII, Beclin1, ATG7, p62) and the activation status of energy sensors (phospho-AMPKα, phospho-FoxO3a). (n = 6 animals in each group; *p < 0.05; **p < 0.01; ***p < 0.001; ****p < 0.0001)

Western blot analysis revealed decreased MyoG levels and increased MAFbx and MuRF1 expression in experimental groups, particularly in protein-deprived conditions, with HMB showing a protective effect (Fig. 4C). Phosphorylation levels of AKT and mTOR were also reduced in experimental groups, and HMB supplementation partially restored these levels. ELISA analysis demonstrated elevated IL-6 levels in both skeletal muscle and blood samples across all experimental groups, with HMB moderating this increase (Fig. 4D). Further molecular analysis revealed increased IL-6 expression in muscle tissue (Fig. 4E), accompanied by elevated levels of inflammatory markers TNF-α and CRP (Fig. 4F). These elevated inflammatory markers indicate a pronounced inflammatory state during inactivity and protein deprivation, which was effectively attenuated by HMB supplementation. The examination of autophagy-related proteins showed significant changes, with increased levels of ULK1, LC3BII, Beclin1, and ATG7 in experimental groups, while p62 levels decreased. HMB supplementation partially normalized these autophagy markers (Fig. 4G). Additionally, analysis of energy sensors showed increased phospho-AMPKα levels in experimental groups, while phospho-FoxO3a levels decreased, with HMB supplementation moderating these changes. These alterations in energy sensors and autophagy markers suggest a metabolic shift towards catabolic processes during muscle stress, which was partially reversed by HMB intervention. Our findings collectively indicate that HMB supplementation may help preserve muscle mass and structure, attenuate atrophy signaling, and moderate inflammatory responses during inactivity and protein deprivation.

HMB attenuates IL-6-induced myoblast defects via AKT/mTOR pathway modulation and regulation of multiple cellular processes

We next sought to elucidate the molecular mechanisms underlying HMB’s protective effects against IL-6-induced muscle atrophy using C2C12 myoblasts stimulated with 20 ng/ml IL-6 for 72 h under various conditions (Fig. 5). Western blot analysis revealed that IL-6 stimulation decreased phosphorylation of AKT and mTOR, which was partially restored by HMB supplementation (Fig. 5A). MyoG levels decreased with IL-6 stimulation but were partially recovered with HMB, while atrophy markers MAFbx and MuRF1 increased with IL-6 treatment and their upregulations were attenuated by HMB (Fig. 5A). De novo protein synthesis significantly decreased under IL-6 stimulation, with HMB partially restoring synthesis levels (Fig. 5B). The protective effects of HMB were blocked by MK-2206 (an AKT inhibitor), confirming the critical role of AKT signaling in HMB’s protective mechanism (Fig. 5B). Cell proliferation showed reduced rates in IL-6-stimulated cells over 72 h, which were improved by HMB supplementation and suppressed after MK-2206 treatment (Fig. 5C). Flow cytometry analysis indicated increased apoptotic rates in IL-6-stimulated cells, significantly reduced by HMB, with MK-2206 reversing this protection (Fig. 5D).

Fig. 5.

Fig. 5

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HMB protects myoblast viability and function from IL-6-induced impairment via AKT/mTOR pathway. C2C12 myoblasts were stimulated with 20 ng/ml IL-6 for 72 h under various conditions, including HMB supplementation and MK-2206 treatment. A Western blot analysis of phosphorylated AKT, phosphorylated mTOR, MyoG, MAFbx, and MuRF1. B De novo protein synthesis measured by Click-IT assay. C Cell proliferation assessed by CCK-8 assay over 72 h. D Flow cytometry analysis of apoptotic rates. E Western blot analysis of autophagy markers (ULK1, LC3BII, p62, Beclin1, ATG7) and the activation status of energy sensors (phospho-AMPKα, phospho-FoxO3a). F Western blot analysis of JAK2/STAT3 signaling pathway components. G Analysis of mitochondrial morphology by fluorescence microscopy. H Flow cytometric analysis of mitochondrial mass (MitoTracker Green) and membrane potential (MitoTracker Deep Red). (n = 3 independent experiments; *p < 0.05; **p < 0.01; ***p < 0.001; ****p < 0.0001)

Further molecular analysis revealed that IL-6 treatment activated autophagy pathways, evidenced by increased ULK1, LC3BII, Beclin1, and ATG7 levels, along with decreased p62. Additionally, IL-6 increased AMPKα phosphorylation and decreased FoxO3a phosphorylation, suggesting activation of catabolic signaling. HMB supplementation moderated all these changes, reducing autophagy markers and AMPKα activation while partially restoring FoxO3a phosphorylation. The addition of MK-2206 abolished HMB’s protective effects, returning all markers to levels similar to IL-6 treatment alone. This suggests HMB alleviates IL-6-induced excessive autophagy through AKT-dependent mechanisms, which involve modulation of both AMPK and FoxO3a signaling pathways (Fig. 5E). Analysis of IL-6 signaling showed increased phosphorylation of JAK2 and STAT3 with IL-6 treatment, which was partially attenuated by HMB but restored with AKT inhibition (Fig. 5F), indicating the interplay between PI3K/AKT signaling and JAK2/STAT3 pathways in mediating muscle atrophy response, where HMB’s protective effects appear to require intact AKT signaling to suppress IL-6-induced inflammatory cascade. Furthermore, mitochondrial morphology assessment demonstrated significant disruption of mitochondrial network under IL-6 treatment, while HMB supplementation helped maintain mitochondrial integrity (Fig. 5G). Flow cytometric analysis using MitoTracker dyes revealed that IL-6 treatment reduced both mitochondrial mass (MitoTracker Green) and membrane potential (MitoTracker Deep Red), while HMB supplementation preserved mitochondrial activity and abundance (Fig. 5H). The protective effects of HMB on mitochondria were also abolished by AKT inhibitor, suggesting that AKT-dependent signaling is essential for HMB’s mitochondrial protective effects, possibly through regulation of mitochondrial quality control and biogenesis pathways.

Taken together, these results suggest that HMB exerts its protective effects against IL-6-induced muscle atrophy through multiple mechanisms: maintaining protein synthesis through the AKT/mTOR pathway, reducing atrophy-related gene expression, moderating autophagy activation, preserving mitochondrial function and integrity, and reducing apoptosis. The dependency of these protective effects on AKT signaling highlights the central role of this pathway in HMB’s therapeutic mechanism.

HMB preserves satellite cell function and viability under IL-6-induced stress via AKT-dependent regulation of protein synthesis and mitochondrial homeostasis

Skeletal muscle satellite cells, as essential stem cells responsible for muscle repair and regeneration, play a crucial role in maintaining muscle homeostasis (Haddad et al. 2005; Londhe and Guttridge 2015; Carbone and Pasiakos 2019). Therefore, we investigated whether HMB could protect satellite cells from IL-6-induced stress using a mouse skeletal satellite cell line. Similar to our observations in myoblasts, IL-6 treatment of satellite cells led to decreased phosphorylation of AKT/mTOR, reduced MyoG expression, and increased atrophy markers MAFbx and MuRF1. HMB supplementation effectively counteracted these changes, but its effects were abolished by AKT inhibitor MK-2206 (Fig. 6A). HMB also restored IL-6-suppressed protein synthesis (Fig. 6B), maintained cell proliferation (Fig. 6C), and prevented apoptosis (Fig. 6D), with all these protective effects being dependent on AKT signaling. Further molecular analysis showed IL-6 activated autophagy, increased AMPKα phosphorylation and decreased FoxO3a phosphorylation, with HMB moderating these changes in an AKT-dependent manner (Fig. 6E). IL-6 signaling analysis demonstrated increased JAK2/STAT3 pathway activation, which was attenuated by HMB but restored with AKT inhibition (Fig. 6F). Mitochondrial analysis revealed that IL-6 disrupted mitochondrial network structure (Fig. 6G) and reduced both mitochondrial mass and membrane potential (Fig. 6H). HMB preserved mitochondrial integrity and function under IL-6-induced stress through an AKT-dependent mechanism.

Fig. 6.

Fig. 6

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HMB maintains muscle satellite cell function during IL-6-induced stress via AKT signaling. Mouse skeletal muscle satellite cells were treated with 20 ng/ml IL-6 for 72 h under various conditions, including HMB supplementation and MK-2206 treatment. A Western blot analysis of phosphorylated AKT, phosphorylated mTOR, MyoG, MAFbx, and MuRF1. B De novo protein synthesis assessment. C Cell proliferation analysis over 72 h. D Flow cytometric analysis of apoptosis. E Western blot analysis of autophagy markers (ULK1, LC3BII, p62, Beclin1, ATG7) and the activation status of energy sensors (phospho-AMPKα, phospho-FoxO3a). F Analysis of JAK2/STAT3 signaling components. G Mitochondrial morphology assessment. H Flow cytometric analysis of mitochondrial mass and membrane potential. (n = 3 independent experiments; *p < 0.05; **p < 0.01; ***p < 0.001; ****p < 0.0001)

These findings demonstrate that HMB protects satellite cell function through multiple AKT-dependent mechanisms, including maintenance of protein synthesis, reduction of atrophic signaling, preservation of mitochondrial function, and regulation of autophagy, suggesting its potential therapeutic value in maintaining satellite cell pool and function under inflammatory conditions.

Discussion

Our study demonstrated that HMB supplementation significantly mitigated muscle atrophy induced by inactivity and protein deprivation. Our findings revealed that HMB partially preserved muscle mass and function, attenuated the expression of atrophy markers (MAFbx, MuRF1), and maintained levels of myogenic factors (MyoG) in both in vivo and in vitro models. Mechanistically, we showed that HMB’s protective effects were mediated through the AKT/mTOR pathway, enhancing protein synthesis and reducing apoptosis in IL-6 stimulated C2C12 myoblasts and skeletal muscle satellite cells. HMB also demonstrated protective effects through regulation of autophagy pathways and preservation of mitochondrial function, while maintaining satellite cell viability through AKT-dependent mechanisms. These results have important implications for the prevention and management of muscle atrophy in various clinical scenarios, including prolonged bed rest, immobilization, and malnutrition. Our findings suggested that HMB supplementation could be a valuable therapeutic strategy to maintain muscle health and improve outcomes in patients at risk of muscle wasting.

Impaired AKT/mTOR signaling and reduced protein synthesis are hallmarks of muscle atrophy induced by inactivity, protein deprivation, or inflammation. Recent studies have elucidated the molecular mechanisms underlying these processes. For instance, lack of exercise or prolonged sedentary behavior could impair the AKT/mTOR signaling and undermine protein synthesis in muscle fibers (Agostini et al. 2018; Zhang et al. 2019). In contrast, resistance exercise-induced mTORC1 signaling mediates muscle regeneration by enhancing de novo protein synthesis (Ge et al. 2009; D’Hulst et al. 2022). Furthermore, amino acid or protein deprivation has been shown to downregulate the AKT/mTOR signaling pathway, leading to decreased protein synthesis and increased protein breakdown in skeletal muscle (Laplante and Sabatini 2012). This nutrient deficiency activates the amino acid sensor General Control Nonderepressible 2 (GCN2), which phosphorylates eukaryotic Initiation Factor 2 alpha (eIF2α) and consequently inhibits global protein synthesis while selectively upregulating the translation of Activating Transcription Factor 4 (ATF4), a transcription factor that induces the expression of genes involved in amino acid synthesis and transport (Zhang et al. 2002; Rasmussen and Adams 2020; Brüggenthies et al. 2022; Misra et al. 2024). Moreover, prolonged amino acid starvation can induce autophagy in muscle cells through the inhibition of mTORC1, further contributing to muscle protein degradation and potentially leading to muscle atrophy (Sandri 2013). Our findings align with these observations, as we noted decreased AKT and mTOR phosphorylation in muscle tissues exposed to inactivity or protein deprivation. Collectively, these molecular mechanisms highlight the intricate interplay between nutrient availability, physical activity, and signaling pathways in maintaining muscle mass, underscoring the importance of balanced nutrition and regular exercise in preventing muscle atrophy and preserving overall musculoskeletal health.

Inflammatory conditions also play a crucial role in muscle atrophy through their effects on the AKT/mTOR pathway and protein synthesis. Chronic inflammation, often associated with diseases such as cancer, diabetes, and rheumatoid arthritis, can significantly impair muscle protein homeostasis (Bonaldo and Sandri 2013). Pro-inflammatory cytokines, particularly tumor necrosis factor-alpha (TNF-α) and interleukin-6 (IL-6), have been shown to interfere with insulin/IGF-1 signaling, leading to reduced activation of the AKT/mTOR pathway (Londhe and Guttridge 2015; Ji et al. 2022; Yang et al. 2022). This interference results in decreased protein synthesis and increased protein degradation, contributing to muscle wasting (Londhe and Guttridge 2015; Ji et al. 2022). Additionally, inflammation-induced activation of nuclear factor-kappa B (NF-κB) can upregulate the expression of muscle-specific E3 ubiquitin ligases, such as MuRF1 and MAFbx/atrogin-1, further promoting protein breakdown (Li and Reid 2000). Our findings corroborate these observations, as we observed decreased phosphorylation of AKT and mTOR in muscle tissues exposed to IL-6 stimulation, accompanied by elevated levels of pro-inflammatory cytokines and increased expression of ubiquitin ligases. Moreover, novel evidence has highlighted the role of inflammasomes, particularly the NLRP3 inflammasome, in exacerbating muscle atrophy through the production of IL-1β and IL-18, which can further suppress the AKT/mTOR signaling pathway (Huang et al. 2017). The complex interplay between inflammation, protein synthesis, and degradation underscores the importance of managing inflammatory conditions to prevent muscle atrophy and maintain muscle mass in various pathological states.

β-hydroxy-β-methylbutyrate (HMB), a leucine metabolite, shows promise in maintaining nutrient balance and muscle homeostasis in sarcopenia (Wu et al. 2015). HMB enhances protein synthesis by activating the mTOR pathway and reduces protein degradation by inhibiting the ubiquitin–proteasome system (Wilkinson et al. 2013). In sarcopenia, HMB supplementation improves muscle mass and strength (Deutz et al. 2013). HMB also modulates inflammatory responses, enhances mitochondrial function, and promotes muscle satellite cell activity (Nunes et al. 2008; Alway et al. 2013; He et al. 2016; Zhong et al. 2019). Our findings further showed that HMB supplementation increased mTOR phosphorylation and decreased E3 ubiquitin ligase expression (MAFbx and MuRF1) in both C2C12 myoblasts and skeletal muscle satellite cells under IL-6 stimulation, as well as in the muscle tissues from inactive and protein-deprived rat models. Notably, our study revealed novel mechanisms of HMB’s protective effects through regulation of autophagy-related proteins (ULK1, LC3B, Beclin1, ATG7, p62) and energy sensors (AMPK, FoxO3a) in both cell types, suggesting its role in maintaining cellular energy homeostasis and protein quality control. Furthermore, we demonstrated that HMB’s beneficial effects extend to preserving mitochondrial function in both myoblasts and satellite cells, as evidenced by maintained mitochondrial mass and membrane potential under stress conditions. The protective effects of HMB were particularly pronounced in satellite cells, where it maintained their viability and function through AKT-dependent mechanisms, providing new insights into its potential role in muscle regeneration and repair. In both myoblasts and satellite cells, HMB attenuated JAK2/STAT3 signaling in response to IL-6 stimulation, suggesting its anti-inflammatory properties may contribute to muscle preservation across different cellular contexts. These multifaceted effects highlight HMB’s potential as a therapeutic agent for sarcopenia and muscle homeostasis in aging and pathological conditions, particularly through its comprehensive impact on protein synthesis, mitochondrial function, and satellite cell maintenance in multiple cell types critical for muscle health. However, the effect of HMB on differentiated myotube cells remains to be studied in future research. This limitation is important to note, as myotubes represent a more terminal differentiation state that may respond differently to HMB treatment compared to myoblasts or satellite cells.

Of note, the alterations in biochemical markers provide important insights into the systemic effects of muscle atrophy and HMB supplementation. The elevated ALT and AST levels suggest that both inactivity and protein deprivation induce not only muscle stress but also hepatic stress. These findings are consistent with a previous report in which hindlimb suspension induces hepatic injury in a rodent animal model (Rivera et al. 2003). The decrease in serum albumin in atrophic conditions reflects compromised protein synthesis capacity, while HMB’s ability to partially maintain albumin levels suggests its role in preserving protein synthesis machinery. The reduction in CREA levels confirms muscle mass loss, while elevated CK levels indicate muscle damage in both atrophic conditions. Notably, the decrease in testosterone levels, particularly in protein-deprived conditions, suggests impaired anabolic signaling, which could contribute to muscle loss. HMB supplementation’s ability to partially normalize these parameters, especially CK and albumin levels, supports its protective role through multiple mechanisms, including maintenance of protein synthesis and reduction of muscle damage. These systemic effects complement our findings of HMB’s direct actions on muscle tissue and inflammatory pathways.

Conclusion

HMB supplementation demonstrates protective effects against muscle atrophy induced by inactivity, protein deprivation and inflammation, through mechanisms involving AKT/mTOR pathway activation, autophagy regulation, and preservation of mitochondrial function in both myoblasts and satellite cells. These findings suggest HMB as a potential therapeutic strategy for preventing muscle atrophy in various clinical scenarios. Future research should focus on optimizing HMB dosage and timing for different populations, exploring potential synergistic effects with other nutrients or exercise protocols, and conducting long-term clinical trials to assess its efficacy and safety in various age groups and health conditions.

Acknowledgements

Not applicable.

Author contributions

The authors confirm contribution to the paper as follows: study conception and design: Xu He, Yan Li; data collection: Jun Chen, Yan Huang, Yang Li; analysis and interpretation of results: Ying Zhou, Yang Li, Jing Quan; draft manuscript preparation: Xu He, Yan Li. All authors reviewed the results and approved the final version of the manuscript.

Funding

This work was supported by Kunming Medical Joint Program [202101AY070001-240]; Project of Clinical Medical Research Center of Geriatric Diseases in Yunnan Province [202102AA310002] and Open Project of Clinical Medical Research Center for Geriatric Diseases in Yunnan Province [2023YJZX-LN-14].

Data availability

No datasets were generated or analysed during the current study.

Declarations

The manuscript has been read and approved by all the authors, that the requirements for authorship as stated earlier in this document have been met, and that each author believes that the manuscript represents honest work.

Conflict of interest

The authors declare no competing interests.

Ethical approval

All the sample handling and data processing steps were following the Declaration of Helsinki. The animal protocols gained the approval of Ethical Review Committee for The First People’s Hospital of Yunnan Province (NO.2020KYS074).

Footnotes

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