C2C12 cells were obtained from ATCC and maintained in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 15% fetal bovine serum, 1% penicillin-streptomycin, and 0.1% tetracycline. The cells were cultured at 37°C with 5% CO₂ until reached a density of approximately 70%–80%. Subsequently, C2C12 cell differentiation was induced by treating the cells with 2% horse serum and 1% penicillin-streptomycin. The horse serum was refreshed daily. On the fourth day, the cells were treated with 100 μM entacapone (MCE, dissolved in DMSO) or the corresponding vehicle control. On the sixth day, C2C12 myotubes were cultured with DMEM with 2% horse serum and 50 μM dexamethasone (Sigma, D4902) for 24 h. Each group underwent four distinct experiments in vitro to ensure reliability and reproducibility of the results.

Male C57BL/6J mice and APOE^−/− mice (C57BL/6J background), aged 6–8 weeks, were purchased from GemPharmatech Co., Ltd. The mice were housed under controlled conditions (22°C–25°C, 12-h light/dark cycle, 45%–55% humidity) with ad libitum access to food and water. All animal experiments adhered to the National Institutes of Health guide for the care and use of laboratory animals and were approved by the Bio-Safety Level III Animal Laboratory of Wuhan University (Wuhan, China).

At the conclusion of the experiments, specific tissues and blood samples were collected for various analyses: Muscle Tissue Collection: The diaphragm and gastrocnemius muscle samples were excised immediately following sacrifice and rinsed in cold PBS. For biochemical analysis, muscle tissues were snap-frozen in liquid nitrogen and stored at −80°C. For histological analysis, muscle samples were fixed in 4% paraformaldehyde and then processed for paraffin embedding.

After weighing, the mice were intraperitoneally anesthetized with 1% pentobarbital sodium. The shaved area was disinfected, and the mice were then connected to a small animal ventilator (VentElite, Harvard Apparatus, United States). The tidal volume was set at 10 mL/kg, with a positive end-expiratory pressure (PEEP) of 2 cm H₂O. The respiratory rate (RR) was maintained at 150 breaths per minute (Li et al., 2022b; Mrozek et al., 2012). To maintain their body temperature at 37°C, the mice were placed on a temperature-controlled blanket. Anesthesia was continuously administered to the mice using a micropump throughout the duration of the experiment.

After collection, the diaphragm was promptly immersed in a Krebs-Henseleit solution (pH 7.4, 25°C) that had been equilibrated with a gas mixture of 95% O₂ and 5% CO₂. The diaphragm was allowed to equilibrate in this solution for 15 min to ensure uniform adaptation to the environment. Subsequently, the muscles were secured onto the equipment and connected to an experimental instrument system (Aurora Scientific Inc, 1200A, Canada) for the measurement of diaphragm muscle strength (Qaisar et al., 2019; Ham et al., 2022). Muscle contractility was assessed by delivering electrical stimulations at varying frequencies (10 Hz, 20 Hz, 40 Hz, 60 Hz, 80 Hz, 100 Hz, 120 Hz, and 140 Hz) using a stimulation voltage of 15V, a pulse duration of 2 milliseconds, and a stimulation interval of 2 min. Each stimulation lasted consistently across frequencies, with a 1-min rest interval between stimuli to allow for recovery. The isometric contraction force was recorded in Newtons (N) during each stimulation. To calculate the muscle’s CSA, the diaphragm’s mass was measured, and CSA was derived using the formula: muscle mass divided by the product of its optimal initial length (Lo) and an assumed muscle density of 1.056 g/cm³. The specific force (in Newtons per square centimeter, N/cm²) was calculated by dividing the recorded muscle force by the CSA (in cm²). This comprehensive methodology provided an accurate assessment of the diaphragm’s force output under varying stimulation conditions, yielding critical data for further physiological research.

The levels of total cholesterol (TC), high-density lipoprotein cholesterol (HDL-C), triglycerides, and low-density lipoprotein cholesterol (LDL-C) in the mice were quantified using an automated chemistry analyzer (Cobas, MiraPlus, Roche Diagnostics) according to the manufacturer’s instructions.

The lung tissues underwent fixation, dehydration, and subsequent paraffin embedding. Afterward, the tissues were sectioned and subjected to routine Hematoxylin and Eosin (H&E) staining. H&E-stained sections were used primarily for qualitative assessment of histopathological changes. The focus of this analysis was to visually identify structural alterations rather than to quantify specific parameters.

To evaluate the influence of lipids on muscle, frozen sections were fixed with 4% paraformaldehyde for 10 min. Following fixation, the sections were washed with 60% isopropanol and stained with Oil Red O at 37°C for 15 min to visualize lipid droplets within the cytoplasm. Subsequently, nuclear counterstaining was carried out using hematoxylin. Lipid Accumulation Measurement: Lipid accumulation was assessed through visual observation of Oil Red O-stained areas.

C2C12 cells were seeded at a density of 2 × 10³ cells per well in 96-well plates. After inducing differentiation for 4 days to form myotubes, the cells were treated with different concentrations of entacapone (50, 100, 150, 200, 250 μM) for 48 h. Additionally, varying concentrations of dexamethasone (50, 100, 150, 200, 250 μM) were added during the last 24 h of treatment. Following the manufacturer’s instructions, the CCK-8 reagent was mixed with the cell culture medium. After a 2-h incubation, the absorbance at 450 nm was measured using a microplate reader. Cell viability was calculated according to the provided instructions.

C2C12 cells and mouse muscle sections were fixed in 4% paraformaldehyde for 25 min. Subsequently, permeabilization was performed using 0.5% Triton X-100 for 15 min, followed by blocking with 5% BSA for 1 h. For C2C12 cells, the primary antibody used was myosin heavy chain (Abclonal; A4963, 1:200). For mouse muscle sections, specific primary antibodies included anti-slow-twitch myosin heavy chain (NOQ7.5.4D, Abcam; ab11083, 1:200), anti-fast-twitch myosin heavy chain (MY-32, Abcam; ab51263, 1:200), and anti-laminin (Abcam; ab11575, 1:500). The samples were incubated overnight at 4°C with the respective antibodies. Afterward, the samples were incubated with the appropriate secondary antibody for 1 h at room temperature. Following PBS washing steps, the nuclei were stained with DAPI. Finally, the samples were sealed, and images were acquired using a fluorescence microscope (Olympus, IX73, Japan).

Protein samples were extracted by lysing muscle tissue or cells using RIPA buffer (Solarbio, China) supplemented with protease and phosphatase inhibitors. For each sample type (cells, diaphragm, gastrocnemius), 30 µg of total protein was loaded into the gel. Following determination of protein sample concentrations, equal amounts of protein samples were subjected to SDS-polyacrylamide gel electrophoresis and subsequently transferred to an NC membrane at 300 mA for 1 h. The membranes were then blocked with 5% nonfat milk. Specific primary antibodies, including Atrogin-1 (Abcam; ab168372, 1:1,000), Murf-1 (Abcam; ab172479, 1:1,000), 4HNE (Abcam; ab46545, 1:1,000), GAPDH (Abcam; ab181602, 1:1,000), superoxide dismutase 1 (SOD1) (Abclonal; A0274, 1:2000), and superoxide dismutase 2 (SOD2) (Abclonal; A19576, 1:2000) were incubated with the membranes overnight at 4°C. After washing with TBST, the membranes were incubated with corresponding secondary antibodies (goat anti-rabbit IgG, Abcam; ab6721, 1:2000) for 1 h at room temperature, followed by further washing with TBST. Chemiluminescent detection was performed using the ECL reagent (Thermo Scientific; 32,106) according to the manufacturer’s instructions. Blot quantification was carried out using Image Lab Software (Bio-Rad), normalizing band intensity to GAPDH as a loading control.

Total RNA was extracted from cells and muscle tissue using Trizol Reagent following the manufacturer’s instructions. The concentration and purity of RNA were determined using a NanoDrop spectrometer (Thermo Fisher Scientific, Inc.). Total RNA was prepared at a concentration of 1 μg/μL for cDNA synthesis using the ABScript II cDNA First-Strand Synthesis Kit (RK20400). Quantitative Real-time PCR was conducted on a QuantStudio 1 Plus system (Applied Biosystems) using the Universal SYBR Green Fast qPCR Mix (2X) (Abclonal, RK21203) and analyzed with QuantStudio TM Design and Analysis Software v1.0.0 (United States). The relative gene expression levels were calculated by the 2^−ΔΔCT method using Glyceraldehyde 3-phosphate dehydrogenase (GAPDH) as the internal control. The primer sequences used in the study are reported below:

Atrogin-1 (F-TTCAGCAGCCTGAACTACGA; R-AGTATCCATGGCGCTCCTTC), Murf-1 (F-GCGTGACCACAGAGGGTAAA; R-CTCTGCGTCCAGAGCGTG) and GAPDH (F-AGGTCGGTGTGAACGGATTTG; R-TGTAGACCATGTAGTTGAGGTCA).

The levels of MDA and GSH-PX activity in cells were determined using assay kits (MDA assay kit, Nanjing Jiancheng Bioengineering Institute; Cat. No. A003-1-1; GSH-PX assay kit, Nanjing Jiancheng Bioengineering Institute; Cat. No. A005-1-1) following the manufacturer’s instructions.

Data were presented as mean ± standard error of the mean (SEM). The normality of the data was assessed using the Shapiro-Wilk test. We performed statistical analyses using one-way analysis of variance (ANOVA), acknowledging that the unbalanced sample sizes could affect the type I and type II error rates. We recommend cautious interpretation of the results, particularly where group sizes differ significantly. Statistical comparisons between groups were conducted using ANOVA. When significant differences were found, post hoc analysis was performed using Tukey’s test for multiple comparisons between groups. For comparisons between two groups, unpaired two-tailed Student’s t-test was employed. All statistical analyses were performed using Prism 8.0.1 software (GraphPad Software, Inc., CA, United States). A two-tailed p-value below 0.05 was deemed statistically significant.

To investigate the impact of entacapone on cellular myotube atrophy in vitro, various concentrations of ENT (50, 100, 150, 200, and 250 μM) were added to myotubes to prevent dexamethasone (Dex)-induced cell death at concentrations of 50 and 150 μM (Figures 1A, B).

Furthermore, the addition of 100 μM ENT was found to effectively protect against the upregulation of Atrogin-1 and Murf-1 protein levels induced by 50 μM Dex (Figure 1C). Specifically, Atrogin-1 levels significantly increased by 660.6% (p < 0.05) in the Dex-treated group compared to the control, while entacapone treatment reduced Atrogin-1 levels by 84.4% (p < 0.05) compared to the Dex group. Similarly, Murf-1 levels rose by 365% (p < 0.05) in the Dex group compared to control and were decreased by 89.5% (p < 0.05) with entacapone treatment. Similar observations were made regarding the attenuation by 100 μM ENT of the upregulation of Atrogin-1 and Murf-1 mRNA levels induced by 50 μM dexamethasone (Figures 2A, B). Collectively, these findings indicate that ENT demonstrates potential in ameliorating Dex-induced muscle atrophy in C2C12 myotubes. To investigate the potential protective effect of ENT against myotube atrophy in vitro, we examined the impact of ENT on C2C12 myotube diameter, as shown in Figure 2C. Dex treatment significantly reduced the diameter of C2C12 myotubes, which was subsequently restored by the addition of ENT. The microscopic images were consistent with the immunofluorescence results. Notably, ENT effectively protected against Dex-induced myotube atrophy.

Dex treatment resulted in a significant decrease in GSH-PX content and an increase in MDA content, indicating elevated oxidative stress in C2C12 myotubes (Figures 3A, B). Specifically, treatment with 50 μM Dex led to a marked increase in MDA levels, a well-known marker of lipid peroxidation, which suggests enhanced oxidative damage to cell membranes. However, ENT effectively reduced this Dex-induced increase in MDA content by 63.98% (p < 0.05), demonstrating its potential protective role against lipid oxidative stress. Additionally, ENT significantly increased GSH-PX content compared to the Dex-treated group by 385.6% (p < 0.05). GSH-PX is a key antioxidant enzyme that plays a critical role in reducing oxidative damage by catalyzing the reduction of hydrogen peroxide and organic peroxides. This increase suggests that ENT not only mitigates oxidative damage but also enhances the antioxidant defense system in C2C12 myotubes. To assess the efficacy of ENT in countering Dex-induced oxidative stress, we further analyzed additional oxidative stress-related markers, including 4-HNE, SOD1, and SOD2. Dex treatment significantly increased the levels of 4-HNE-modified proteins, indicating heightened lipid peroxidation. ENT treatment effectively attenuated this increase, reinforcing its role in reducing oxidative damage. Additionally, we observed that ENT inhibited the Dex-induced upregulation of SOD1 and SOD2, which are important antioxidant enzymes that help mitigate ROS levels (Figures 3C–F). The downregulation of these enzymes suggests that while Dex induces oxidative stress, ENT may help restore the balance by modulating these protective pathways. Collectively, these findings indicate that Dex exposure significantly upregulates markers of oxidative stress, while ENT exerts a protective effect against Dex-induced oxidative stress and muscle atrophy in C2C12 myotubes.

After 12 h of continuous MV, diaphragm muscle strength exhibited a significant decrease compared to the control group. Treatment with ENT substantially mitigated this decrease (Figure 4A). Additionally, MV resulted in elevated expression levels of Atrogin-1 and Murf-1; however, ENT treatment effectively reduced the ventilation-induced upregulation of these markers (Figure 4B). H&E staining of lung tissue revealed alveolar septal thickening, alveolar hemorrhage, and alveolar collapse in the mechanically ventilated group. Notably, these pathological changes were significantly alleviated following the administration of ENT (Figure 4C). High doses of Dex led to a decline in diaphragm strength, but diaphragm contractility in mice was enhanced in the ENT + Dex group compared to the Dex group (Figure 4D). Mice subjected to Dex-induced muscle atrophy experienced substantial weight loss when compared to the control group. Mice in the ENT + Dex group also exhibited weight loss relative to both the control group and the Dex group alone (Figure 4E). The CSA of both the gastrocnemius and diaphragm displayed a reduction in the Dex group when compared with the control group. However, ENT effectively safeguarded against this reduction (Figure 4F).

To investigate the effects of LPS-induced muscle atrophy and the protective role of ENT against inflammation-related muscle atrophy, we examined the expression levels of muscle atrophy and oxidative stress markers: Atrogin-1, Murf-1, and 4-HNE. As anticipated, LPS-induced muscle atrophy led to a notable elevation in the levels of Atrogin-1, Murf-1, and 4-HNE (p < 0.05), indicating the activation of pathways associated with muscle degradation and oxidative stress. However, in the ENTLPS group, these levels exhibited a significant reduction compared to the LPS group (p < 0.05) (Figure 5). This reduction suggests that ENT effectively mitigates oxidative stress, thereby exerting a protective influence on the molecular mechanisms involved in muscle atrophy.

To evaluate the impact of hyperlipidemia on muscle atrophy and the potential effects of ENT on the muscles of mice with hyperlipidemia, we employed APOE^−/− mice and measured blood lipid levels. The results indicated that APOE^−/− mice exhibited higher levels of TC, LDL, and TC/HDL compared to C57 mice (Figure 6A). Furthermore, muscle strength was reduced in hyperlipidemic mice relative to C57 mice (Figure 6B). Concurrently, immunofluorescence analysis of the gastrocnemius muscle revealed a smaller CSA of gastrocnemius myofibers in APOE^−/− mice compared to C57 mice, suggesting myofiber atrophy. Notably, the administration of ENT mitigated this myofiber atrophy (Figure 6C). Lipid content was assessed through Oil Red O staining, which corroborated the reduction in gastrocnemius muscle CSA in APOE^−/− mice and demonstrated that ENT treatment protected against muscle fiber atrophy. Additionally, APOE^−/− mice exhibited more lipid droplet accumulation in muscle compared to C57 mice, while treatment with ENT significantly alleviated lipid aggregation in the gastrocnemius muscle of APOE^−/− mice (Figure 6D). Taken together, our findings demonstrate that ENT treatment confers a protective effect against muscle atrophy associated with hyperlipidemia.