Introduction
Metabolic dysfunction-associated steatotic liver disease (MASLD) represents a chronic hepatic disorder defined by ≥5% steatosis concurrent with at least one cardiometabolic risk factor.1 With a global prevalence exceeding 30%, MASLD represents a disease spectrum ranging from simple steatosis to metabolic dysfunction-associated steatohepatitis (MASH), which may progress to advanced fibrosis, cirrhosis, and hepatocellular carcinoma, and is closely linked to obesity, diabetes, and other metabolic disorders.2 Beyond the liver, MASLD has systemic consequences, with strong associations with cardiovascular disease, chronic kidney disease, and extrahepatic metabolic complications.3
Skeletal muscle, the largest insulin-sensitive organ, is central to energy balance.4 Progressive decline in muscle mass and function, known as sarcopenia, is common in older adults and patients with chronic diseases.5 It is reported to affect 10% to 27% of individuals aged 60 and above worldwide, substantially increasing risks of frailty, disability, and all-cause mortality.6 Sarcopenia has been increasingly recognized as a disease rather than simply an aging phenotype, and its risk can be exacerbated by factors such as physical inactivity, inadequate nutrition, and metabolic disorders.7
Growing epidemiological evidence underscores a clinically significant bidirectional relationship between sarcopenia and MASLD. Several cross-sectional studies have identified that sarcopenia, independent of insulin resistance and obesity, is associated with MASLD and MASH-related advanced fibrosis.8,9 This association carries important prognostic implications, as demonstrated by meta-analytic data showing that sarcopenic individuals face over twofold greater odds of MASLD development, alongside a 59% increased all-cause mortality risk among MASLD patients.10 Emerging evidence also positions MASLD as an independent risk factor for sarcopenia. A multicenter prospective study demonstrated that baseline hepatic steatosis predicted incident sarcopenia even after comprehensive adjustment for age, sex, lifestyle factors, comorbidities, and metabolic confounders.11 While these observational studies establish epidemiological associations, they cannot delineate causal directionality between sarcopenia and MASLD.
The pathogenesis of sarcopenia and MASLD demonstrates substantial mechanistic overlap, including insulin resistance, chronic low-grade inflammation, and imbalances in hormones and cytokines.12 The liver and skeletal muscle function as endocrine organs capable of secreting a variety of proteins and metabolites, including hepatokines and myokines, that facilitate inter-organ communication.13,14 Some of these molecules have been implicated in metabolic homeostasis. For example, muscle-derived follistatin-like protein 1 promotes the progression of MASH.15 In contrast, increased secretion of the myokine irisin enhances lipid catabolism and effectively ameliorates hepatic injury.16 Fetuin-A, primarily expressed and secreted by the liver, is elevated in the hepatic tissue and serum of MASLD patients and may induce insulin resistance in muscle.17 Although certain myokines and hepatokines have been identified as key regulators in muscle atrophy, hepatic steatosis, and fibrosis, the functional roles of other secreted factors in the crosstalk between skeletal muscle and liver remain to be fully elucidated.
Currently, pharmacological interventions for coexisting MASLD and sarcopenia remain limited, partly due to a lack of adequate preclinical animal data addressing the co-occurrence and mechanistic interplay between these two conditions.18 To address this gap, we employed an integrative approach combining human genetic analyses with complementary mouse models. First, we applied Mendelian randomization (MR) to evaluate the causal relationship between sarcopenia-related traits and MASLD risk. To experimentally dissect the liver-muscle axis, we established muscle atrophy-combined MASLD models to examine how sarcopenia influences MASLD progression. In parallel, we utilized well-established MASLD models to investigate how MASLD affects muscle function. By integrating these approaches with multi-tissue transcriptomic profiling, we aimed to construct a mechanistic framework for this inter-organ crosstalk and identify potential therapeutic strategies targeting hepatokine-myokine signaling for patients with MASLD and sarcopenia.
Methods
MR analysis
To investigate the causal relationship between sarcopenia and MASLD, we employed a two-sample MR approach. In this study, sarcopenia-related traits were defined as exposures, while MASLD was designated as the outcome. Genetic instruments for MASLD were derived from a publicly available genome-wide association study comprising 778,614 European individuals (8,434 cases and 770,180 controls).19 Appendicular lean mass data were obtained from a genome-wide association study of 450,243 participants.20 Furthermore, we acquired instrumental variables (IVs) associated with walking speed and leg impedance from the MRC-IEU consortium (https://gwas.mrcieu.ac.uk/ ). These variables involved 454,857 participants for left-leg impedance, 454,863 for right-leg impedance, and 335,349 for walking pace.
Single-nucleotide polymorphisms (SNPs) significantly associated with the exposure were selected as IVs at a genome-wide significance threshold of p < 5 × 10−8. The strength of the IVs was evaluated using the F-statistic, with a value greater than 10 considered indicative of a robust instrument. The F-statistic was calculated as F = R2 × (N−2) / (1−R2), where R2 represents the proportion of variance in the exposure explained by the SNPs, and N is the sample size.21 To ensure independence among the IVs, linkage disequilibrium clumping was applied with parameters set to R2 = 0.001 and a window size of 10,000 kb. SNPs associated with potential confounders or the outcome were systematically removed using Phenoscanner V2. After harmonizing the exposure and outcome datasets, the remaining eligible SNPs were retained for subsequent analysis. Five MR methods were employed, with inverse variance weighting serving as the primary approach.22 Pleiotropy was evaluated using the MR-Egger intercept test, while heterogeneity was assessed with Cochran’s Q test. All analyses were conducted using the TwoSampleMR and MR-PRESSO packages in R.
Animals
Specific pathogen-free C57BL/6J mice, aged six to eight weeks, were procured from HuaFukang BioScience Company (Beijing, China) and maintained on a 12-h light cycle with ad libitum access to food and water.
For the botulinum toxin A (BTX-A)-induced sarcopenia model,23 eight-week-old C57BL/6J mice were randomized into two groups: (i) Saline; (ii) BTX-A. Lyophilized BTX-A (Botox, Allergan Pharmaceuticals, Ireland) was reconstituted in sterile saline. Following anesthesia, the BTX-A group received intramuscular injections of BTX-A at a dose of 20 IU/kg in the right hindlimb.24 The Saline group received equivalent volumes of saline at the same site. After two weeks, both groups were subjected to a high-fat methionine–choline-deficient diet (60 kcal% fat with 0.1% methionine and no added choline; A06071302, Research Diets, New Brunswick, USA) for three weeks.25
For the stelic animal model (STAM),26 male C57BL/6J mice were randomized into two groups: (i) Ctrl; (ii) STAM. The STAM group received a single subcutaneous injection of 200 µg streptozotocin (STZ; Sigma-Aldrich, MO, USA) at postnatal day 2, followed by feeding with a high-fat diet (HFD; 60 kcal% fat; D12492, Research Diets, New Brunswick, USA) from four to seven weeks of age. Control mice were administered vehicle solution subcutaneously at postnatal day 2 and maintained on the same HFD from four weeks of age.
To establish a diet-induced MASLD model, male C57BL/6J mice (six to eight weeks old) were fed either a low-fat diet (LFD; 10 kcal% fat; D17112301R, Research Diets, New Brunswick, USA) or a Gubra-Amylin NASH (GAN) diet (40 kcal% fat, 20 kcal% fructose, and 2% cholesterol; D09100310, Research Diets, New Brunswick, USA) for 20 weeks.25
To establish a spontaneous obesity model, eight-week-old male leptin-deficient (ob/ob) and control (ob/m) mice, obtained from HuaFukang BioScience Company (Beijing, China), were maintained on a normal chow diet (13.5 kcal% fat; LabDiet, MO, USA) for 16 weeks.27,28
All animal care and experimental protocols were in accordance with the ethical guidelines of the Shanghai Institute of Materia Medica, Chinese Academy of Sciences.
Grip test
Forelimb and whole-limb grip strength were assessed using a grip strength meter (XR501, XinRuan, Shanghai, China). Mice were gently pulled horizontally by the tail after grasping a metal grid (forelimbs or all limbs).16 Peak force from three trials was recorded and normalized to body weight.
Biochemical analysis
Serum levels of alanine aminotransferase (ALT) and aspartate aminotransferase (AST), and cellular total triglyceride (TG) levels were quantified using commercially available assay kits (Jiancheng, Nanjing, China). Hepatic total cholesterol (TC) and TG levels were measured using enzymatic colorimetric assays in accordance with the manufacturer’s protocols (Jiancheng, Nanjing, China). Frozen liver tissue was homogenized in ice-cold Tris-HCl buffer containing 1% (v/v) Triton X-100. After centrifugation, the lipid-containing supernatant was analyzed.
Cell culture and treatment
All cells were maintained at 37°C in a 5% CO2 atmosphere with saturated humidity. The C2C12 cell line (ATCC, VA, USA) was cultured in growth medium composed of high-glucose DMEM (Gibco, MA, USA), 10% fetal bovine serum (Gibco, MA, USA), and 1% penicillin–streptomycin (Meilunbio, Liaoning, China). At 90% confluence, differentiation was initiated by replacing the growth medium with differentiation medium (high-glucose DMEM supplemented with 2% horse serum and 1% penicillin–streptomycin). To induce muscle atrophy in vitro, myotubes were treated five days post-differentiation with 100 ng/mL recombinant murine tumor necrosis factor-alpha (TNF-α) (RP01071, ABclonal, Hubei, China) in differentiation medium for 24 h.29
AML12 cells (Fenghui Biotechnology, Hunan, China) were cultured in DMEM supplemented with 10% fetal bovine serum, 1% insulin–transferrin–selenium (Gibco, MA, USA), 40 ng/mL dexamethasone (MedChemExpress, Shanghai, China), and 1% penicillin–streptomycin (Meilunbio, Liaoning, China). To induce intracellular lipid accumulation, AML12 cells were treated with 0.2 mM palmitic acid (PA; Sigma-Aldrich, MO, USA) for 24 h.30
Following TNF-α treatment, C2C12 cells were washed with phosphate-buffered saline (PBS) and then incubated in differentiation medium. After 24 h, the conditioned medium (CM) was collected. AML12 cells were treated for 24 h with a mixture of CM and fresh intrinsic medium (3:1 ratio) along with 0.2 mM PA, followed by cellular TG quantification.
After PA treatment, AML12 cells were washed with PBS and returned to growth medium. CM was collected 24 h later. Following five days of differentiation, C2C12 myotubes were then treated for 24 h with a mixture of this CM and intrinsic medium at a 3:1 ratio.
To investigate the role of secretory factors in vitro, AML12 hepatocytes were treated with 200 ng/mL recombinant C-C motif chemokine ligand 2 (CCL2) protein (RP01626, ABclonal, Hubei, China), and five-day differentiated C2C12 myotubes were treated with 100 nM recombinant adrenomedullin (ADM) protein (TP09904, TenZoBio, Hubei, China) for 24 h.
ELISA
Serum and cell-CM levels of CCL2 and ADM were quantified using commercial ELISA kits (CCL2: ml037533; ADM: ml002194; both from Enzyme-Linked Biotechnology, Shanghai, China) in accordance with the manufacturer’s protocols.
RNA sequencing (RNA-seq) and analysis
Total RNA was isolated from the livers and muscles of mice and subsequently subjected to high-throughput sequencing using the Illumina platform. RNA libraries were prepared using the Illumina TruSeq Stranded Total RNA Library Prep Kit (Illumina, San Diego, CA) following the manufacturer’s protocol. All subsequent library processing and paired-end sequencing were conducted by Majorbio Bio-Pharm Technology (Shanghai, China) on an Illumina NovaSeq platform. Quality-filtered reads were then aligned to the murine reference genome.
Transcriptomic data were analyzed with the edgeR package to identify differentially expressed genes (DEGs). Functional annotation of DEGs was conducted using Gene Ontology (GO) and Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway enrichment analyses, alongside gene set enrichment analysis, with the clusterProfiler package. Building on prior secretome profiling,31 we identified DEGs encoding hepatokines and myokines. To systematically map liver-muscle crosstalk, we employed the NicheNetR package to analyze bulk RNA-seq data.32
Histological assessment
Murine muscle and liver samples were fixed in paraformaldehyde, dehydrated, and embedded in paraffin. Tissue morphology was assessed via hematoxylin and eosin (H&E) staining (Servicebio, Hubei, China), while intramuscular lipid deposition was quantified in cryosectioned samples using Oil Red O staining (Servicebio, Hubei, China). Hepatic collagen deposition was evaluated by Sirius red staining (Servicebio, Hubei, China).30 The NAFLD activity score (NAS) was evaluated according to the histopathological characteristics observed in liver biopsy specimens, encompassing steatosis (scored 0–3), hepatocellular ballooning (scored 0–2), and lobular inflammation (scored 0–3).33 Muscle cross-sectional area (CSA) and minimum Feret’s diameter were quantified using ImageJ.34
Immunofluorescence staining
C2C12 myotubes were fixed with 4% paraformaldehyde for 30 min, permeabilized with 0.1% Triton X-100, and blocked with 3% bovine serum albumin in PBS for 30 min at room temperature. Subsequently, the cells were incubated overnight at 4°C with an anti-myosin heavy chain antibody (MF 20, DSHB, Iowa, USA) diluted 1:100 in blocking buffer.35 Following primary antibody incubation, the cells were washed and then incubated with an Alexa Fluor 488-conjugated anti-mouse secondary antibody (4408S, CST, MA, USA; 1:500 dilution) and counterstained with DAPI (Beyotime, Shanghai, China). Fluorescence images were acquired using a Zeiss LSM 900 fluorescence microscope, and myotube diameter was quantified using ImageJ.
Quantitative real-time PCR
Total RNA was extracted from frozen muscle, liver, and cells using TRIzol (Yeasen Biotechnology, Shanghai, China). cDNA was synthesized with HiScript® III RT SuperMix (Vazyme, Jiangsu, China). Real-time PCR amplification was conducted using Hieff qPCR SYBR Green Master Mix (Yeasen Biotechnology, Shanghai, China) and a CFX384 system. Gene expression data were normalized to their respective housekeeping genes, using ACTIN for liver tissue and AML12 cells and CYPA for C2C12 cells. Primer sequences are detailed in Supplementary Table 1.
Statistical analysis
All statistical analyses were conducted using GraphPad Prism (GraphPad Software, CA, USA). Continuous variables are presented as mean ± standard deviation. Between-group comparisons were performed using an unpaired two-tailed Student’s t test. The threshold for statistical significance was set at *p < 0.05, with more stringent levels indicated as **p < 0.01 and ***p < 0.001.
Results
MR analysis revealed a causal relationship between sarcopenia and MASLD
Given the established clinical overlap between MASLD and sarcopenia, along with prior MR evidence indicating a causal effect of MASLD on sarcopenia risk,36 we first investigated whether sarcopenia-related traits conversely exerted a causal influence on MASLD risk in humans. To address this, we performed two-sample MR analysis using large-scale genome-wide association studies of muscle mass, muscle function, and MASLD. Our MR analysis revealed that higher appendicular lean mass was significantly associated with reduced MASLD risk (OR 0.91, 95% CI 0.84–0.98) (Fig. 1). Walking pace, reflecting physical function, showed an even stronger protective relationship (OR 0.34, 95% CI 0.15–0.80), with consistent results observed in weighted median and simple mode analyses. Leg impedance, an indicator of muscle quality, was also inversely associated with MASLD (right leg: OR 0.85, 95% CI 0.76–0.95; left leg: OR 0.87, 95% CI 0.77–0.97). MR-Egger intercept tests indicated an absence of horizontal pleiotropy, suggesting that the IVs were not influenced by unobserved confounding factors (Fig. 1). These findings demonstrate that reduced muscle mass and impaired function increase MASLD risk, supporting a causal role of sarcopenia in MASLD pathogenesis.
BTX-A–induced muscle atrophy promoted MASLD in high-fat methionine–choline-deficient diet-fed mice
To validate this causal link experimentally, we next investigated whether sarcopenia exacerbated MASLD progression in vivo. We established a localized muscle atrophy model by a single intramuscular injection of BTX-A (Fig. 2A), a neurotoxin that induces muscle atrophy by blocking neuromuscular transmission.37,38 The injection was administered into the hindlimb muscle to better recapitulate the clinical progression of sarcopenia, which affects the lower limbs more rapidly and severely than the upper limbs in humans.39
Administration of BTX-A resulted in a modest reduction in body weight (Fig. 2B). Compared with the saline-treated cohort, mice receiving BTX-A exhibited significantly diminished grip strength, indicative of impaired muscle function (Fig. 2C). Furthermore, BTX-A resulted in profound muscle atrophy, with significant reductions in both quadriceps femoris and gastrocnemius mass (Fig. 2D). H&E staining of muscle confirmed a decreased myofiber CSA, a diminished minimum Feret’s diameter, and a leftward shift in myofiber size distribution toward smaller CSA (Fig. 2E and F).
Subsequently, we assessed whether muscle atrophy induced by BTX-A could exacerbate liver dysfunction. The ratio of liver weight to body weight was slightly higher in the BTX-A group (Fig. 2G). Serum ALT, AST, and hepatic TC levels also increased, but the differences were not statistically significant (Fig. 2H and I). In contrast, hepatic TG content was significantly increased, accompanied by marked hepatic steatosis in BTX-A-treated mice (Fig. 2I and J). Histological assessment further revealed enhanced inflammatory infiltration and fibrosis in the BTX-A group (Fig. 2J and K). RT-qPCR analysis indicated upregulation of the fatty acid transporter gene CD36, whereas the expression of genes involved in lipogenesis and fatty acid oxidation remained unaltered (Fig. 2L). Notably, mRNA levels of pro-inflammatory genes such as IL1b and CASP1, along with profibrotic genes, were significantly elevated in the livers of BTX-A-treated mice (Fig. 2L). Taken together, these results demonstrate that BTX-A–induced muscle atrophy aggravated the progression of MASLD.
Coexistence of diabetic sarcopenia and MASLD in STAM mice
To further assess the association between systemic sarcopenia and MASLD, we next employed a STAM model.26,40 In the STAM group, neonatal mice received a low, non-hepatotoxic dose of STZ via subcutaneous injection, which effectively induced pancreatic islet inflammation and dysfunction, constituting a high-risk factor for diabetic sarcopenia.26,40 Subsequently, at four weeks of age, both the control group and the STZ-treated group were maintained on an HFD to develop MASLD (Fig. 3A).
Compared with control mice, STAM mice demonstrated a significant reduction in body weight (Fig. 3B). Notably, STAM mice exhibited severe muscle weakness, as evidenced by impaired grip strength, along with a substantial decline in muscle mass (Fig. 3C and D). Additionally, STZ administration induced a notable decrease in myofiber CSA and fiber diameter, and a shift toward smaller myofiber size distribution (Fig. 3E and F).
STAM mice exhibited significant liver dysfunction, as evidenced by a marked increase in the ratio of liver weight to body weight and elevated serum ALT and AST levels (Fig. 3G and H). Hepatic TC and TG levels were also significantly increased in STAM mice compared with controls (Fig. 3I). Histological assessment revealed severe hepatic steatosis, inflammatory infiltration, and fibrosis in STAM mice (Fig. 3J and K). Molecular analysis showed upregulated expression of CD36 alongside downregulated lipogenic genes in the livers of STAM mice (Fig. 3L). Additionally, pro-inflammatory cytokines and profibrotic markers were significantly elevated in STAM mice relative to controls (Fig. 3L). Previous studies have shown that administration of an appropriate dose of STZ is generally sufficient to induce sarcopenia within two weeks,34,40 whereas STZ-induced diabetes alone has a limited effect on hepatic lipid deposition, requiring a “second hit” from HFD to precipitate MASLD.26 Together, these data demonstrate that, in the STAM model, the presence of severe sarcopenia was associated with exacerbated MASLD progression.
In summary, both the BTX-A and STAM models consistently exhibited impaired muscle function and structural atrophy, suggesting a link between sarcopenia and MASLD. In the STAM model, beyond sarcopenia, other STZ-induced systemic alterations, such as lipodystrophy and oxidative stress, may act as confounding factors that could influence the development of MASLD.41,42 In contrast, the BTX-A model more directly illustrated the contribution of sarcopenia induced by neuromuscular inhibition.
Sarcopenia was observed in both diet-induced and genetically obese mice
Evidence from a prior MR study has indicated a causal role of MASLD in sarcopenia pathogenesis.36 To experimentally validate this causal link, we employed a GAN diet-induced nutritional model and genetically obese leptin-deficient (ob/ob) mice (Fig. 4A and H).
Compared with ob/m controls, ob/ob mice exhibited marked obesity (Supplementary Fig. 1A and B). The ob/ob group developed severe hepatic steatosis with significantly elevated hepatic TG content (Fig. 4B and C). As expected in this genetic model,43 no apparent fibrosis was observed (Fig. 4C). These animals also demonstrated prominent sarcopenic characteristics, including decreased grip strength, reduced muscle mass, and diminished myofiber CSA (Fig. 4D–F). Such muscular impairment was associated with ectopic lipid accumulation in skeletal muscle (Fig. 4F). Notably, all components of the NAS score were strongly inversely correlated with both muscle mass and muscle function (Fig. 4G).
To further investigate the relationship between the severity of MASLD and sarcopenia, we subjected mice to a MASH-inducing GAN diet (Fig. 4H). After 20 weeks, GAN diet-fed mice developed significant obesity (Supplementary Fig. 1C and D). Histological and biochemical assessments confirmed the onset of MASH, evidenced by pronounced hepatic steatosis, inflammation, and collagen deposition in the liver (Fig. 4I and J). Additionally, these mice displayed significant muscle weakness and atrophy compared with LFD-fed controls, as indicated by reduced grip strength, muscle mass, myofiber CSA, and increased ectopic lipid accumulation (Fig. 4K–M). Notably, all components of the NAS score and Sirius red–positive area were strongly inversely correlated with both muscle mass and muscle function, with these associations becoming more marked as MASLD severity increased (Fig. 4N).
Collectively, these results demonstrate that sarcopenia is a consistent phenotype in both dietary and genetic models of MASLD, supporting the notion that hepatic metabolic disturbances serve as a direct driver of sarcopenia progression, with the severity of muscle loss correlating with the advancement of MASLD pathology.
Transcriptomic profiling revealed sarcopenia-driven liver dysfunction via myokine signaling
Given that STZ-induced metabolic disturbances exhibit broad systemic effects due to pancreatic β-cell destruction,26,40 whereas BTX-A–mediated muscle atrophy is more localized with minimal extra-muscular impact,38 the BTX-A model enabled investigation of the direct impact of sarcopenia on MASLD pathogenesis. To elucidate how sarcopenia remodels liver metabolism and mediates muscle-to-liver communication, we performed RNA-seq on liver and skeletal muscle from BTX-A–treated mice and their controls.
We first identified significant transcriptomic alterations in the livers of BTX-A–induced sarcopenia models. GO analysis of upregulated DEGs revealed that sarcopenia perturbed multiple hepatic processes, particularly nutrient sensing, RNA metabolism, and transcriptional regulation (Fig. 5A). KEGG pathway analysis further demonstrated activation of mTOR signaling, insulin signaling, and PPAR signaling pathways (Fig. 5B). Conversely, GO analysis showed sarcopenia-induced downregulation of mitochondrial electron transport chain components and mitochondrial function, suggesting compromised cellular energy production and metabolic dysfunction (Fig. 5C). This was corroborated by KEGG analysis revealing broad disturbances in metabolic pathways, including oxidative phosphorylation, detoxification capacity, and cholesterol homeostasis (Fig. 5D). Additionally, gene set enrichment analysis identified significant upregulation of lipid transport pathways concurrent with suppression of lipid catabolic pathways in livers of the BTX-A group, providing a mechanistic basis for the observed exacerbation of hepatic steatosis (Fig. 5E). These findings demonstrate that sarcopenia caused broad transcriptional reprogramming in the liver, characterized by mitochondrial dysfunction and dysregulated lipid homeostasis.
Parallel RNA-seq analysis of skeletal muscle also revealed distinct changes. Pathways associated with muscle development and extracellular matrix remodeling were significantly upregulated, potentially reflecting compensatory tissue repair mechanisms in response to neuromuscular inhibition (Supplementary Fig. 2A and B). Conversely, marked downregulation of carbohydrate metabolism and energy production pathways was observed (Supplementary Fig. 2C and D). Notably, contractile apparatus-related pathways (including myofibril, sarcomere, and calcium signaling) were substantially suppressed, consistent with the expected muscle relaxation effects of BTX-A (Supplementary Fig. 2C). These transcriptional alterations demonstrate that BTX-A induced muscle dysfunction through impaired energy metabolism and contractile dysfunction.
As the primary effector of movement and a key metabolic regulator, skeletal muscle secretes myokines that orchestrate systemic metabolic homeostasis.13 In BTX-A–induced sarcopenia, we identified 97 upregulated and 50 downregulated myokine-encoding genes through cross-referencing muscle DEGs with a secreted protein database (Supplementary Fig. 2E).31 To delineate muscle-liver crosstalk mechanisms, we applied NicheNet analysis using the upregulated myokines as putative ligands. This computational prediction demonstrated robust sarcopenia-associated muscle-to-liver signaling, with CCL2, CCL8, AGRP, TIMP1, and SERPINE1 emerging as the top five muscle-derived ligands exhibiting the strongest predicted regulatory activity in liver and high-affinity receptor interaction potential (Fig. 5F and Supplementary Fig. 2F). Downstream analysis identified twelve hepatic target genes regulated by the myokine signaling network (Fig. 5G and H). These results demonstrate that muscle-derived ligands in sarcopenia exerted a substantial influence on hepatic gene expression.
Collectively, these findings reveal that sarcopenia promoted hepatic steatosis through impaired mitochondrial function and disrupted lipid homeostasis and established a mechanistic link to MASLD progression via myokine-mediated inter-organ communication.
CCL2 secretion induced by muscle atrophy promoted lipid accumulation in hepatocytes
To investigate whether sarcopenia directly altered hepatocyte lipid metabolism, we induced muscle atrophy in vitro by treating C2C12 myotubes with TNF-α, which resulted in a significant reduction in myotube diameter (Fig. 6A and B). Following removal of TNF-α–containing medium, CM from these C2C12 cultures was collected and used to treat AML12 hepatocytes in combination with PA administration (Fig. 6A). Notably, CM from atrophic C2C12 myotubes markedly increased TG levels in AML12 cells (Fig. 6C). Based on RNA-seq data from sarcopenic mice, we identified CCL2, CCL8, TIMP1, AGRP, and SERPINE1 as potential myokines that may mediate the effects of muscle atrophy on MASLD (Fig. 5F). Consistent with this, TNF-α–induced muscle atrophy significantly elevated the expression of CCL2, CCL8, TIMP1, and SERPINE1 in C2C12 cells, with CCL2 exhibiting the most pronounced upregulation (Fig. 6D). Increased secretion of CCL2 was further confirmed in the CM of atrophic C2C12 cells (Fig. 6E). Additionally, serum levels of CCL2 were significantly elevated in both BTX-A–treated mice and STAM mice (Fig. 6F). To functionally validate the role of CCL2, AML12 cells were treated with recombinant CCL2 protein along with PA, which resulted in a significant increase in intracellular TG accumulation (Fig. 6G). These findings demonstrate that muscle atrophy directly promoted lipid accumulation in hepatocytes and identify CCL2 as a key muscle-derived mediator linking sarcopenia to MASLD progression.
Transcriptomic profiling revealed MASLD-driven muscle dysfunction via hepatokine signaling
To investigate how MASLD contributed to skeletal muscle dysfunction and inter-organ communication, we performed RNA-seq on the liver and muscle from mice fed either an LFD or a GAN diet. This nutritional model better reflects the metabolic disturbances that initiate and drive MASLD in humans.
Significant transcriptomic alterations were first identified in skeletal muscle of MASLD models. GO analysis of upregulated DEGs indicated that MASLD affected multiple aspects of muscle biology, primarily involving immune system activation, inflammatory response, and hormone- and cytokine-related processes (Fig. 7A). Furthermore, KEGG pathway analysis demonstrated that cytokine–cytokine receptor interaction, muscle cytoskeleton, and hormone signaling pathways were upregulated, suggesting that MASLD induced muscle inflammation, remodeling, and metabolic regulation (Fig. 7B). In contrast, GO analysis revealed that the pathways most downregulated by MASLD were related to polyamine metabolism (Fig. 7C). Additionally, MASLD disrupted pathways involved in muscle secretory function and cellular polarity, potentially affecting inter-tissue communication, while KEGG analysis further demonstrated broad changes in metabolic processes, including carbohydrate, lipid, and amino acid metabolism (Fig. 7C and D). These findings demonstrate that MASLD drove inflammatory activation and metabolic dysfunction in skeletal muscle, which together may underlie the observed sarcopenic phenotype.
Parallel RNA-seq of liver tissue showed that pathways related to leukocyte activation and adhesion, collagen-containing extracellular matrix, and cytokine–cytokine receptor interaction were predominantly upregulated in MASLD livers (Supplementary Fig. 3A and B). Conversely, downregulated DEGs induced by MASLD were primarily enriched in sterol biosynthesis, protein synthesis and transport, and protease inhibitor activity (Supplementary Fig. 3C and D). These findings suggest that coordinated immune-inflammatory and fibrotic pathway activation drove hepatocyte injury in MASLD, while suppressed metabolic pathways exacerbated lipid dysregulation and protein metabolism impairment.
Emerging evidence indicates that steatotic livers exhibit altered endocrine functions compared with healthy livers, releasing differentially expressed secretory proteins (hepatokines) into circulation that may disrupt systemic metabolic homeostasis.14 To characterize hepatokine dysregulation in MASLD, we cross-referenced hepatic DEGs with a database of secreted proteins,31 identifying 250 upregulated and 66 downregulated hepatokine-encoding genes (Supplementary Fig. 3E). To investigate the molecular basis of liver–muscle crosstalk in MASLD, we performed NicheNet analysis using the upregulated hepatokines as putative ligands. The top five liver ligands exhibiting the highest predicted activity in MASLD were LAMA2, HBEGF, CFD, ADM, and CCL19, showing strong receptor interaction potential in muscle (Fig. 7E and Supplementary Fig. 3F). Downstream analysis identified 15 muscle target genes regulated by these hepatokines (Fig. 7F and G). These results indicate that hepatokine dysregulation in MASLD can directly reshape skeletal muscle gene expression.
Collectively, these findings establish that MASLD contributed to muscle wasting by triggering inflammatory and metabolic disruption in muscle and by altering hepatokine-mediated liver-muscle crosstalk.
ADM secretion induced by MASLD promoted myotube atrophy in vitro
To investigate the potential direct influence of MASLD on muscle atrophy, we induced hepatic lipid accumulation in vitro by treating AML12 hepatocytes with PA. In AML12 cells, PA exposure markedly increased intracellular TG content (Fig. 8A and B). After PA removal, CM from these lipid-laden hepatocytes was collected and applied to differentiated C2C12 myotubes (Fig. 8A). Notably, CM from PA-treated hepatocytes significantly reduced C2C12 myotube diameter, indicating a direct pro-atrophic effect on muscle cells (Fig. 8C). Based on RNA-seq data from MASLD model mice, we identified HBEGF, CFD, ADM, CCL19, and LAMA2 as potential hepatokines that might mediate the effects of MASLD on sarcopenia (Fig. 7E). Consistent with this, PA-induced lipid accumulation significantly upregulated the expression of CFD, ADM, CCL19, and LAMA2 in AML12 cells, with ADM showing the most pronounced increase (Fig. 8D). Enhanced secretion of ADM was further confirmed in the CM of PA-treated AML12 cells (Fig. 8E). Additionally, serum ADM levels were significantly elevated in both diet-induced MASLD mice and ob/ob mice (Fig. 8F). To functionally validate the role of ADM, C2C12 myotubes were treated with recombinant ADM protein, resulting in a significant reduction in myotube diameter (Fig. 8G). Collectively, these findings demonstrate that MASLD directly promoted myotube atrophy and that ADM secreted by steatotic hepatocytes acted as a pathogenic signal to impair muscle integrity, providing evidence for hepatokine-mediated liver–muscle crosstalk in MASLD-driven sarcopenia.
Discussion
This study provides integrated evidence that MASLD and sarcopenia are not only epidemiologically associated but also causally linked through inter-organ signaling. Using MR analysis, we demonstrated that reduced muscle mass and function increase the risk of MASLD. By employing complementary murine models, we confirmed that muscle atrophy exacerbates the progression of MASLD, while steatotic liver injury accelerates muscle dysfunction. These findings establish that sarcopenia and MASLD form a reciprocal pathogenic loop rather than independent complications of metabolic disease. Our transcriptomic and functional analyses further revealed the molecular basis of this crosstalk. Sarcopenia disrupts hepatic lipid homeostasis by upregulating fatty acid uptake and suppressing oxidative phosphorylation, whereas MASLD impairs muscle integrity by potentiating inflammatory responses and metabolic dysfunction. Among the secreted mediators, CCL2 emerged as a key myokine promoting hepatic lipid accumulation, while ADM was identified as a hepatokine that directly induces muscle atrophy. These results highlight a bidirectional signaling axis that links the liver and skeletal muscle and explain how MASLD and sarcopenia reinforce each other.
The causal relationship uncovered by MR analysis is in line with epidemiological evidence showing inverse associations between sarcopenia-related traits, such as appendicular lean mass, leg bioimpedance, and walking pace, and MASLD risk, suggesting potential protective effects of muscle preservation against hepatic steatosis. Currently, dietary and lifestyle modifications remain the primary therapeutic strategies for MASLD.1 In patients with MASLD-related cirrhosis, it is particularly crucial to improve lifestyle to achieve weight reduction while preventing muscle mass loss, as sarcopenia is an independent predictor of survival, especially in patients with decompensated cirrhosis.44
Mechanistically, both BTX-A-treated mice and STAM mice exhibited increased hepatic lipid accumulation, partly through increased CD36 expression, implicating enhanced fatty acid uptake as a driver of steatosis. This aligns with studies showing increased CD36 localization on hepatocyte membranes in MASH patients, where CD36 palmitoylation promotes free fatty acid uptake and exacerbates intracellular lipid deposition.45 Additionally, our RNA-seq data revealed an upregulation of pathways associated with hepatic lipid transport in the BTX-A group. Moreover, mitochondrial dysfunction likely further disrupts liver lipid metabolic homeostasis and promotes MASH progression.46
In this study, we identified CCL2 as a critical myokine that mediates the pathogenic crosstalk between skeletal muscle and liver. As a key chemokine, CCL2 is known for its role in immune cell chemotaxis and inflammatory responses.47 Recent studies have shown that CCL2 expression is upregulated in specific subpopulations of muscle stem cells within senescent human skeletal muscle, as well as in various cell types in the muscle fiber microenvironment, where it contributes to disrupted immune homeostasis.48 Our data suggest that muscle-derived CCL2 may enter the circulation and exert effects on the liver. Previous studies have demonstrated that inhibition of CCL2 during chronic liver injury attenuates steatohepatitis by reducing hepatic monocyte and macrophage infiltration.49
This study demonstrates that obesity induces not only hepatic steatosis but also profound sarcopenic alterations in mice, including diminished grip strength, reduced myofiber CSA, and increased intramuscular lipid deposition. These findings are consistent with clinical observations of the high prevalence of sarcopenic obesity in MASLD patients.50 Notably, the pathological significance of myosteatosis extends beyond muscle dysfunction, as emerging evidence indicates that it independently predicts MASH development, fibrosis progression, and adverse outcomes in cirrhosis, underscoring the pathological importance of altered muscle lipid metabolism.51,52
Our transcriptomic analysis demonstrated concurrent activation of inflammatory pathways in both liver and skeletal muscle under MASLD conditions. In the obese state, numerous cytokines released by immune cells and adipocytes, such as interleukin-6, C-reactive protein, and TNF-α, induce chronic low-grade inflammation.53 The progression of MASLD is markedly accelerated through a multistage inflammatory cascade, comprising immune cell activation, proinflammatory cytokine release, and establishment of chronic inflammation.54 This inflammatory milieu disrupts insulin and IGF-1 signaling, promotes endoplasmic reticulum stress and mitochondrial dysfunction in skeletal muscle, and collectively drives insulin resistance, protein metabolism imbalance, and muscle atrophy.7
In the reverse direction, ADM was identified as a critical hepatokine linking steatotic liver to muscle atrophy. As a peptide hormone, ADM is known primarily for its roles in regulating vascular tone, vascular remodeling, and endothelial barrier function and integrity.55 A recent study highlighted its involvement in type 2 diabetes, reporting elevated plasma levels of ADM in both obese humans and mice.56 Specifically, ADM was shown to activate protein tyrosine phosphatase 1B, which inhibits phosphorylation of the insulin receptor in vascular endothelial cells, thereby impairing insulin signaling.56 Treatment of mice with an ADM mimetic recapitulated the effects of obesity, inducing both endothelial and systemic insulin resistance.56
Building upon the direct atrophic effects of ADM on myocytes in our study, an intriguing question arises regarding its potential interplay with muscle-derived CCL2. We hypothesize that ADM and CCL2 may reinforce each other by mutually exacerbating insulin resistance and chronic inflammation. On one hand, ADM-mediated insulin resistance may diminish the anti-inflammatory effects of insulin, thereby potentially augmenting CCL2 expression.56,57 Conversely, elevated levels of CCL2 may exacerbate hepatic inflammation,49 which in turn can further enhance the secretion of hepatic ADM via cytokines such as TNF-α.58,59 The subsequently increased ADM may then circulate back to muscle, worsening insulin resistance and muscle atrophy. Therefore, beyond their direct roles, ADM and CCL2 likely form a self-amplifying axis that disrupts muscle-liver crosstalk, a mechanism that warrants further investigation.
Several limitations warrant attention. The MR analysis was conducted exclusively in European populations, potentially limiting the extrapolation of our findings to other ethnic groups. Regarding animal models, while BTX-A-induced muscle atrophy replicates core sarcopenia features, it does not fully emulate the chronic and systemic progression of age-related muscle decline observed in humans.39 Similarly, the STAM model induces insulin-deficient diabetes rather than the insulin-resistant state typically observed in MASLD-associated sarcopenia patients.7,60 It is necessary to design longitudinal studies to clarify the temporal relationship between the onset of sarcopenia and MASLD, especially in STAM mice, GAN diet-induced MASLD mice, and ob/ob mice. While the roles of CCL2 and ADM have been established in vitro, their in vivo functions and the potential interplay between their downstream signaling pathways warrant further investigation. Furthermore, the clinical relevance and translational potential of these mechanisms identified in murine models to human pathophysiology remain to be elucidated.