γ-Tocotrienol attenuates oxidative stress and preserves mitochondrial function in inflammation-induced muscle atrophy

Abstract

Muscle atrophy, marked by the loss of skeletal muscle mass and strength, presents a major health concern with diverse etiologies, including chronic inflammation. Effective interventions are urgently needed for its prevention and treatment. Although α-tocopherol, the most abundant form of vitamin E, is known for its antioxidant benefits in muscle health, γ-tocotrienol exhibits superior antioxidant and anti-inflammatory properties. This study investigates the protective effects of γ-tocotrienol against muscle atrophy and compares its efficacy with α-tocopherol. Muscle atrophy was induced in differentiated C2C12 myotubes using lipopolysaccharide (LPS), with vitamin E pre-treatment applied prior to LPS challenge. Myotube morphology, expression of atrophy-related markers, and underlying molecular pathways were examined through immunofluorescence, western blotting, and quantitative proteomics. LPS treatment induced significant myotube atrophy without affecting cell viability. Notably, γ-tocotrienol pre-treatment preserved myotube size and suppressed key atrophy markers, including the E3 ubiquitin ligases MuRF-1 and Fbxo32/Atrogin-1. Proteomic analysis quantified 5,371 proteins and revealed that γ-tocotrienol alleviated atrophy by enhancing extracellular matrix organization and attenuating oxidative stress and mitochondrial dysfunction. These protective effects were further confirmed in vivo, where γ-tocotrienol administration preserved muscle strength, suppressed pro-inflammatory signaling, and restored mitochondrial biogenesis in LPS-treated mice. Collectively, these findings demonstrate that γ-tocotrienol offers superior protection against muscle atrophy compared to α-tocopherol, highlighting its therapeutic potential for individuals at risk of muscle wasting.

Graphical abstract

Highlights

• •γ-Tocotrienol exhibits superior muscle-protective effects than α-tocopherol in LPS-induced atrophy.
• •Pre-treatment of γ-tocotrienol preserves myotube morphology and suppresses MuRF-1 and Atrogin-1 expression.
• •Proteome analysis reveals γ-tocotrienol enhances ECM organization, reduce oxidative stress, and protects mitochondria.
• •In vivo administration of γ-tocotrienol restores muscle strength, promotes mitochondrial biogenesis, and reduces inflammation.
• •γ-Tocotrienol as a promising therapy for inflammation-induced muscle loss, offering advantages over α-tocopherol.

1. Introduction

Muscle atrophy, characterized by a reduction in muscle mass and strength, represents a significant health challenge with diverse etiologies [1]. Various pathological conditions—including metabolic syndromes such as diabetes, cancer, and sclerosis—can disrupt skeletal muscle homeostasis and trigger atrophic processes [2,3]. These disorders are frequently associated with chronic inflammation, mitochondrial dysfunction, apoptosis, and dysregulated protein turnover. The inflammatory mechanisms underlying skeletal muscle atrophy are highly complex, necessitating molecular-level insights to develop effective preventive and therapeutic strategies. Although exercise remains the most effective intervention for both preventing and reversing muscle atrophy [4], its applicability is often limited in patients with physical impairments or chronic illnesses [2]. Therefore, identifying alternative pharmacological or nutritional approaches is essential to address the multifactorial nature of muscle wasting.

Lipopolysaccharide (LPS) is a widely used agent for inducing muscle atrophy in vitro and mimics the molecular and physiological features of muscle loss observed in inflammation-related diseases [[5], [6], [7], [8]]. Elevated circulating endotoxins, including LPS, have been reported in various conditions such as obesity [9], insulin resistance [10], type 2 diabetes [11], non-alcoholic fatty liver disease [12], cancer [13], and other chronic inflammatory disorders [14], all of which are associated with skeletal muscle wasting.

As a component of the outer membrane of Gram-negative bacteria, LPS acts as a potent pro-inflammatory stimulus by binding to toll-like receptor 4 (TLR4), thereby triggering the production of pro-inflammatory cytokines such as tumor necrosis factor-alpha (TNF-α), interleukin-6 (IL-6), and interleukin-1β (IL-1β), as well as generating reactive oxygen species (ROS). LPS activates NADPH oxidases and impairs mitochondrial function, leading to excessive ROS production. These ROS further amplify inflammation and activate transcription factors such as NF-κB and FOXO. The FOXO family (including FOXO1, FOXO3, and FOXO4) directly induces the expression of the muscle-specific E3 ubiquitin ligases MuRF1 and Atrogin-1 (Fbxo32), key mediators of proteasome-dependent protein degradation during muscle atrophy [15].

This study aims to investigate the protective effects of γ-tocotrienol, a potent isoform of vitamin E, against LPS-induced muscle atrophy using C2C12 murine myotubes. While α-tocopherol, the predominant form of vitamin E, is well-known for its antioxidant benefits in muscle health [16], γ-tocotrienol has demonstrated superior antioxidant and anti-inflammatory activities [17,18]. Structurally, tocotrienols possess an unsaturated isoprenoid side chain that facilitates better membrane penetration and distribution within lipid bilayers. This unique property contributes to their wide-ranging health benefits, including improved glycemic control and insulin sensitivity in diabetic rats [19], favorable modulation of lipid profiles [20], and cardioprotective effects [21]. Although several studies have reported the muscle-preserving potential of tocotrienols, most employed tocotrienol-rich fractions (TRFs) derived from palm oil [22], without directly comparing the effects of γ-tocotrienol to α-tocopherol. For example, recent findings showed that TRF supplementation mitigated hyperglycemia-induced muscle atrophy and insulin resistance in diabetic mice [19], and enhances myogenic differentiation in myoblasts [23]. These collective findings suggest that γ-tocotrienol may exert more potent protective effects on skeletal muscle than α-tocopherol.

In the present study, we systematically compared the effects of γ-tocotrienol and α-tocopherol on LPS-induced muscle atrophy using mass spectrometry-based quantitative proteomics. We further examined the molecular pathways involved in muscle protein turnover in vivo and in vitro to elucidate the mechanisms by which γ-tocotrienol exerts its protective actions against inflammation-driven muscle wasting.

2. Materials and methods

2.1. Cell culture and treatment

C2C12 murine myoblast cell line was obtained from Bioresource Collection and Research Center (BCRC), Hsinchu, Taiwan). Cells was cultured in high-glucose DMEM (Gibco, Grand Island, NY, USA) supplemented with 10 % FBS (Gibco) at 37 °C with 5 % CO₂. Culture medium were changed to differentiation medium (DM, DMEM supplemented with 2 % HS (Gibco)) upon reaching 80 % confluency, and replaced every 2 days throughout the differentiation process. After 6 days of differentiation, multinuclear myotubes were formed. C2C12 myotubes were pretreated with different doses of γ-tocotrienol (0.2, 1, 5 μM) (Cayman Chemical, Ann Arbor, MI, USA) or α-tocopherol (Cayman Chemical) for 6 h before treated with LPS (Chemscene, Monmouth Junction, NJ, USA) for 48 h to induce muscle atrophy. All experiments were conducted using C2C12 cells at passages between 5 and 10.

2.2. Cell viability assay

Cells were plated in a 96-well culture plate at a density of 1 × 10⁴ cells per well, induced to differentiation for 6 days, and pre-treated with various concentrations of γ-tocotrienol for 6 h prior to 48-h LPS administration. Cell viability was determined by CCK8 assay (Dojindo Laboratories, Mashiki, Japan). 10 μL of the CCK-8 reagent was added directly into the wells and incubated in the dark for 30 min. Absorbance at 450 nm wavelength was read with an ELISA microplate reader (Molecular Devices, San Jose, CA, USA).

2.3. Immunofluorescence staining

Differentiated myotubes were fixed by 4 % paraformaldehyde (Sigma‐Aldrich, Burlington, MA, USA), for 10 min at room temperature, permeabilized by 0.2 % Triton X-100 (Sigma-Aldrich) in PBS for 5 min, and then blocked with blocking buffer (BlockPRO, Taiwan) for 1 h at room temperature. Myotubes were incubated with Recombinant Alexa Fluor® 488 Anti-TOMM20 antibody (Abcam, Cambridge, UK), anti-NFκB p65 (Cell signalling Technology, Danvers, MA, USA), anti-Myh1 (Abclonal) or anti-Myh4 (DHSB, Iowa City, IA, USA) diluted in blocking buffer overnight at 4 °C, followed by incubation with Alexa Fluor 562-goat anti-mouse IgG and Alexa Fluor 488-goat anti-rabbit IgG antibody (Thermo Fisher Scientific (Boston, MA, USA)). Cells were visualized with a fluorescence microscope (Axio Observer D1, Carl Zeiss AG, Oberkochen, Germany), and the diameter of myotubes was measured by ImageJ software (NIH, Bethesda, MD, USA). The nuclear fusion index (NFI) were measured using MyoCount software version 1.3 [24].

2.4. Nuclear protein fractionation

Nuclear fraction isolation was performed using the REAP method as described previously [25]. Briefly, collected cells were centrifuged down and resuspended in 900 μL of ice-cold 0.1 % NP40 in PBS, and triturated 5 times using a P1000 pipette tip. The lysate was then centrifuged on a tabletop microcentrifuge (GeneReach, Taiwan), and the supernatant was collected as the cytosol fraction. The remaining pellet was then wash with 0.1 % NP40 in PBS twice and the nuclear pellet was resuspended in RIPA buffer.

2.5. Proteomics analysis

Cells lysis and protein extraction were performed using PTS buffer [26]. The proteins were treated with 10 mM TCEP (Sigma-Aldrich) at 37 °C for 30 min to disrupt disulfide bonds, and alkylated by 25 mM CAA (Sigma-Aldrich) at 37 °C for 30 min. Protein digestion into peptides was achieved by incubating with ABC (Sigma-Aldrich), Lys-C endopeptidase (FUJIFILM Wako Chemicals USA, Richmond, VA, USA), and trypsin (w:w = 1:100) (Promega) for 3 h and 16 h respectively. After digestion, the reaction was quenched with 10 % TFA (Thermo Fisher Scientific) and EtOAc (Sigma-Aldrich) was added for detergent extraction. The mixture was vigorously shaken for 2 min, followed by centrifugation at 17000×g for 2 min to remove the organic layer containing EtOAc and detergent. The remaining EtOAc was removed using SpinVac (Thermo Fisher Scientific) and the sample was desalted with SDB-XC StageTip [27]. Peptide concentration was quantified according to the absorbance at 214 nm with a SpectraMax ABS Plus Absorbance ELISA Microplate Reader. For TMT labeling and sample fractionation, equal amount of samples were dissolved in 200 mM HEPES pH 8.5 (Thermo Fisher Scientific) and labeled with TMTpro™ 18 plex Label Reagent Set (Thermo Fisher Scientific) according to the manual. Pooled sample was fractionated under alkaline condition, followed by desalted with C18 StageTip.

Samples were reconstituted 2 % ACN (Thermo Fisher Scientific) with 0.1 % FA (Thermo Fisher Scientific). Analysis was performed using an Ultimate system 3000 nanoLC system (Thermo Fisher Scientific, Bremen, Germany) coupled with an Orbitrap Fusion Lumos Tribrid quadrupole-ion trap-Orbitrap mass spectrometer (Thermo Fisher Scientific, San Jose, CA). The column used was a 75 μm ID, 25 cm length C18 Acclaim PepMap NanoLC column (Thermo Fisher Scientific, San Jose, CA) packed with 2 μm particles with a pore of 100 Å. Mobile phase A (0.1 % FA in Milli-Q water) and B (0.1 % FA in 80 % ACN) were used as the LC buffer. The initial mobile phase composition was set to 98 % A and 2 % B, with a gradient increase to 40 % B over 100 min, separating the peptides at a flow rate of 300 nL/min in 50 min. Mass spectrometer parameters were set as follows: Data-dependent mode with Full-MS (mass accuracy of <5 ppm and a resolution of 120,000 at m/z 200, AGC target 5e5, maximum injection time of 50 ms); HCD-MS/MS of the most intense ions in 3 s. MS/MS (resolution of 15,000) within a 1.4 Da isolation window at a normalized collision energy of 32. AGC value 5e4 was set for MS/MS analysis with previously selected ions dynamically excluded for 60 s. Max injection time of MS/MS was set at 50 ms.

2.6. Peptide identification and analysis of differential expressed proteins

The raw MS data was processed using MaxQuant software version 2.0.3.0 for peptide and protein identification and quantification. The Andromeda search engine [28] was employed to search MS2 data against the mouse reference protein database UniProt (downloaded on December 24, 2023). Subsequent processing and analysis of the identified proteins file were performed using Perseus 1.6.15.0 [29]. Decoy hits and contaminations were removed, and protein data was log2 transformed and normalized. Differentially expressed proteins (DEPs) were identified using Significance Analysis of Microarrays (SAM) with an S0 factor applied to a two-sample t-test and false discovery rate (FDR) of less than 0.05. DEPs were visualized in heatmaps and volcano plots. Subsequently, STRING and Metascape was utilized to obtain protein-protein interaction networks [30], and functional pathway enrichment analysis was performed using Enrichr [31] and GSEA [32].

2.7. Measurement of intracellular reactive oxygen species (ROS)

C2C12 cells were seeded and differentiated in 6-well plates containing coverslips, treated with γ-tocotrienol for 6 h and then stimulated with LPS (1 μg/mL) for 48 h. To detect intracellular ROS, H₂-DCFDA fluorescence staining (Thermo Fisher Scientific) was performed on live C2C12 myotubes. To monitor the generation of mitochondrial ROS, cells were stained with 50 nM MitoTracker™ Red CMXRos (Thermo Fisher Scientific, Boston, MA, USA) for 30 min in the dark. After washing three times with PBS, cells were fixed with 4 % paraformaldehyde for 15 min and proceed to fluorescence immunostaining. Coverslips were mounted using a mounting medium containing 4,6-diamidino-2-phenylindole (DAPI, Biotium, CA, USA). Fluorescent images were captured by a Axio Observer D1 microscope (ZEISS, Oberkochen, Germany) and processed in ZEN Imaging Software (ZEISS).

2.8. Mitochondrial membrane potential (ΔΨm) measurement

C2C12 cells were seeded and differentiated in 6-well plates containing coverslips, treated with γ-tocotrienol for 6 h and then stimulated with LPS (1 μg/mL) for 48 h. The fluorescent cationic dye JC-1 (Thermo Fisher Scientific) was used to assess mitochondrial membrane potential. Fluorescent images were captured by a Axio Observer D1 microscope (ZEISS) and processed in ZEN Imaging Software (ZEISS). Changes in the JC-1 fluorescence intensity ratio (red: 540/570 nm to green: 495/520 nm) were calculated based on signals recorded within a fixed exposure time.

2.9. Mitochondrial oxygen consumption rate analysis

Seahorse XFe24 analyzer (XFe24, Agilent Technologies, CA) was employed to evaluate the mitochondrial oxygen consumption rate (OCR) of C2C12 myotubes. C2C12 myoblasts were seeded at equal density in Seahorse XFe24 microplates, cultured to full confluence, and subsequently differentiated into myotubes to ensure consistent differentiation and minimize variability from post-assay normalization. Myotubes were treated with either γ-tocotrienol or α-tocopherol (2 μM) for 6 h, followed by exposure to LPS (1 μg/mL) for 48 h. Before conducting the assay, the sensors were hydrated overnight at 37 °C in a non-CO2 incubator using Agilent Seahorse XF calibrant. The culture medium was then replaced with Agilent Seahorse XF basal medium, and the plates were incubated at 37 °C in an incubator without CO₂ for 1 h. Mitochondrial function was assessed using the Seahorse XF Cell Mito Stress Test, with measurements taken at baseline and after the sequential addition of oligomycin (1 μM), FCCP (3 μM), and a combination of antimycin A and rotenone (1 μM).

2.10. Animal model of LPS-induced muscle atrophy

All animal experiments were conducted in accordance with the guidelines and regulations approved by the Institutional Animal Care and Use Committee (IACUC) of the Laboratory Animal Center at National Defense Medical University (IACUC-25-076). Six-week-old male C57BL/6 mice were purchased from the National Laboratory Animal Center (Taipei, Taiwan) and housed under controlled conditions with a 12-h light/dark cycle, at 25 °C and 65 % relative humidity. Male mice were used to ensure consistent experimental conditions and to minimize potential confounding effects of sex steroids on metabolism and atrophy progression [33,34].

After one-week acclimation, supplementation was administered daily by oral gavage for six weeks, starting four weeks before LPS exposure. Mice were randomly divided into six experimental groups (n = 8 per group): Control, LPS-only, and four treatment groups receiving LPS with prior supplementation of either γ-tocotrienol or α-tocopherol at 20 mg/kg or 200 mg/kg body weight. The dosages were selected based on previous studies [35].

Beginning at week 11, muscle atrophy was induced by intraperitoneal injection of LPS (1 mg/kg body weight), administered once every five days for a total of three injections. Body weight was monitored twice weekly throughout the study. Mice were sacrificed 14 days after the initial LPS injection, and the gastrocnemius muscles were collected for subsequent histological and biochemical analyses.

2.11. Hematoxylin-eosin (H&E) staining and muscle fiber cross-sectional area analysis

Gastrocnemius muscle samples were fixed, sectioned, and stained with hematoxylin and eosin (H&E) by Catching Micro-Tech Co., Ltd. (Taipei, Taiwan). Stained tissue sections were imaged at 40× magnification using a ZEISS Axioscan 7 Slide Scanner (Zeiss). The cross-sectional area (CSA) of individual muscle fibers was quantified using ImageJ software. At least 100 fibers per sample were measured to ensure statistical robustness.

2.12. Immunohistochemical staining

Paraffin-embedded tissue sections were deparaffinized, rehydrated, and subjected to blocking using a blocking buffer for 1 h at room temperature. The sections were then incubated overnight at 4 °C with primary antibodies against PGC-1α and SIRT1 (1:500 dilution). After washing, the sections were incubated for 1 h at room temperature with fluorescently labeled secondary antibodies: Alexa Fluor 562-conjugated goat anti-mouse IgG and Alexa Fluor 488-conjugated goat anti-rabbit IgG. Immunofluorescent signals were visualized using a multiphoton laser scanning confocal microscope (LSM980, Zeiss).

2.13. Grip strength test

Muscle strength was evaluated using a grip strength meter (SH‐III‐20, China). Mice were gently placed on a stainless-steel grid and allowed to grasp it with all four limbs while being held by the tail. The grid was then pulled horizontally at a constant speed until the mouse released its grip, and the peak force was recorded. Each mouse underwent four consecutive trials, and the average force from these trials was calculated as the representative grip strength. Measurements were performed both before and after LPS treatment to assess treatment-related changes in muscle strength.

2.14. Enzyme-linked immunosorbent assay (ELISA)

Serum interleukin-6 (IL-6) levels were measured using a commercial ELISA kit (BioLegend, CA, USA; Cat. No. 431304), following the manufacturer's instructions. Each sample was analyzed in at least two technical replicates to ensure accuracy. Absorbance was read at 450 nm using a microplate reader (Molecular Devices, USA), and cytokine concentrations were calculated based on standard curves generated from known IL-6 concentrations.

2.15. Western blot

Cell pellets or mice tissues were sonicated in RIPA buffer with phosphatase/protease inhibitor (MedChemexpress LLC, Middlesex, NJ, USA). The lysates were centrifuged at 17,000×g for 20 min at 4 °C. Protein concentration was determined by BCA protein assay (T-Pro biotechnology, New Taipei City, Taiwan) using different concentrations of BSA (BioShop Canada Inc., Burlington, ON, Canada) as standard, and equal amounts of protein samples were separated by 10–15 % SDS-PAGE and transferred to a PVDF membrane (Millipore, Burlington, MA, USA). The PVDF membranes were blocked in 5 % skimmed milk in TBS containing 0.1 % Tween 20 for 1 h at room temperature and then incubated with primary antibodies diluted in blocking buffer at 4 °C overnight with the following concentration: MuRF-1 (1:1000), Fbxo32 (1:2000), Myh1 (1:2000), Sod2 (1:1000) (Abclonal, Woburn, MA, USA), NFκB p65 (1:1000), Sirt1 (1:1000), p-Akt (1:1000), Akt (1:1000) (Cell signalling Technology), Lamin B1 (1:1000), Pgc-1α (1:1000) (Genetex, Irvine, CA, USA), OXPHOS (1:2000) and α-Tubulin (1:5000) (Abcam). Membranes were then washed with TBST, and incubated with anti-mouse (Abcam) and anti-rabbit IgG horseradish peroxidase-conjugated secondary antibody (Abcam) for 1h in TBST. An ECL chemiluminescent kit (T-Pro biotechnology) was used to visualize the protein bands, and the chemical luminescence of membranes was detected using an eBlot Touch Imager (eBlot Photoelectric Technology, Shanghai, China). The relative band intensity was quantified using ImageJ software. The data were expressed as fold change compared with the untreated group (n > 3).

2.16. Statistical analysis

All data are performed with at least 3 independent experiment of biological replicates. Data are presented as mean ± standard deviation (SD). Statistical significance between treatments was analyzed using one-way analysis of variance (ANOVA) followed by Tukey's multiple comparisons test, unless specified otherwise. All analyses were conducted using GraphPad Prism 8.0. A p-value of less than 0.05 was considered statistically significant.

3. Results

3.1. LPS effectively induced myotube atrophy in differentiated C2C12 cells

To model inflammation-induced muscle atrophy, C2C12 cells were differentiated in 96-well plates and treated with various concentrations of LPS for 48 h. Cell viability analysis indicated that only the highest dose of LPS exhibited cytotoxic effects (Fig. 1A). In LPS-treated groups, significant reductions in myotube diameter and size were observed, accompanied by marked morphological changes. Even at non-cytotoxic doses, LPS-treated myotubes exhibited fragmented and shrunken morphologies, indicating that LPS-induced atrophy occurs independently of cell death (Fig. 1B).

Fig. 1.

LPS-induced myotube atrophy and the protective effects of γ-tocotrienol in C2C12 cells. (A) Cell viability of C2C12 myotubes treated with various concentrations of LPS for 48 h, as measured by the CCK-8 assay. (B) Phase-contrast images of myotubes treated with LPS, showing reductions in diameter and size. Scale bar = 200 μm. (C) Cell viability of C2C12 myotubes treated with γ-tocotrienol at different concentrations for 48 h, measured by the CCK-8 assay. (D) Fluorescent images of myotubes pretreated with γ-tocotrienol and exposed to LPS, stained for nuclei (blue, DAPI) and Myh1 (green). Scale bar = 100 μm. (E) Western blot analysis of Myh1, MuRF-1, and Atrogin-1 protein expression in LPS-treated myotubes, with α-tubulin as the internal control. Data are presented as mean ± SD from three independent experiments.

To evaluate the protective effects of γ-tocotrienol against LPS-induced muscle atrophy, myotubes differentiated for 5 days were pretreated with γ-tocotrienol at various concentrations before LPS exposure. γ-Tocotrienol demonstrated excellent tolerance in myotubes, even at high concentrations (Fig. 1C). Pretreatment with γ-tocotrienol significantly preserved the morphological integrity of myotubes and prevented LPS-induced atrophy (Fig. 1D). To confirm the occurrence of myotube atrophy in LPS-treated C2C12 cells, we analyzed the expression of the differentiation marker Myh1 (MyHC-IIx/d) and the atrophy markers MuRF-1 and Atrogin-1. LPS treatment led to a significant reduction in Myh1 expression, coupled with an upregulation of MuRF-1 and Atrogin-1, corroborating the observed morphological changes (Fig. 1E). These findings validate that LPS effectively induces muscle atrophy in differentiated C2C12 cells and demonstrate the protective effects of γ-tocotrienol against this process.

3.2. γ-tocotrienol alleviates LPS-induced muscle atrophy

To further assess the protective effects of γ-tocotrienol on LPS-induced myotube atrophy, differentiated C2C12 myotubes were pretreated with varying concentrations of γ-tocotrienol prior to LPS exposure. Specifically, 0.2 μM and 2 μM of γ-tocotrienol were added to the culture medium 6 h before a 48-h LPS treatment. Myotubes pretreated with γ-tocotrienol showed a dose-dependent reduction in susceptibility to LPS-induced damage, with the 2 μM concentration demonstrating a more pronounced protective effect compared to 0.2 μM. This was evidenced by improved cell morphology and the preservation of myotube diameter (Fig. 2A and B). In contrast, myotubes exposed to LPS alone exhibited significant disruptions in myogenesis, characterized by reduced myotube diameter and structural disintegration (Fig. 2B).

Fig. 2.

Protective effects of γ-tocotrienol against LPS-induced myotube atrophy in C2C12 cells. C2C12 cells (5000 cells/well) were differentiated in DMEM supplemented with 2 % horse serum (HS) and pretreated with various concentrations of γ-tocotrienol or vehicle on day 5 post-differentiation for 6 h, followed by treatment with 1 μg/mL LPS or vehicle for 48 h. (A) Phase-contrast images of C2C12 myotubes under different treatments. Scale bar = 100 μm. (B) Quantification of myotube diameter using ImageJ software. (C) Western blot analysis of the differentiation marker (Myh1) and atrophy markers (MuRF-1 and Fbxo32/Atrogin-1) under different treatment conditions, with α-tubulin as the loading control. (D) Statistical analysis of protein expression. Data are presented as mean ± SD from three independent experiments. ∗p < 0.05, ∗∗p < 0.01 compared to the control group; #p < 0.05, ##p < 0.01 compared to the LPS-treated group.

Western blot analysis supported these morphological findings, revealing increased Myh1 expression in γ-tocotrienol-treated groups and a lack of significant upregulation of atrophy markers MuRF-1 and Atrogin-1 compared to the LPS-only group (Fig. 2C and D). These results suggest that γ-tocotrienol effectively mitigates LPS-induced myotube atrophy in C2C12 cells, potentially by modulating proteasome-dependent protein degradation pathways.

3.3. γ-tocotrienol inhibits LPS-induced nuclear translocation of NF-κB

Lipopolysaccharide (LPS) triggers an inflammatory response by activating nuclear factor kappa-light-chain-enhancer of activated B cells (NF-κB) and facilitating its translocation into the nucleus [33]. To determine whether γ-tocotrienol mitigates myotube atrophy via NF-κB pathway modulation, differentiated myotubes were pretreated with γ-tocotrienol prior to LPS exposure. NF-κB p65 levels were assessed using immunoblotting and immunocytochemistry (ICC). LPS treatment significantly increased NF-κB p65 protein expression, while γ-tocotrienol pretreatment restored p65 levels in a dose-dependent manner (Fig. 3A and B). ICC analysis revealed that LPS-treated myotubes exhibited increased NF-κB p65 expression with pronounced nuclear localization. In contrast, γ-tocotrienol pretreatment significantly inhibited NF-κB p65 nuclear translocation (Fig. 3C). Additionally, γ-tocotrienol preserved myotube diameter and nuclear fusion index (NFI), calculated as the ratio of nuclei in multinucleated myotubes to total nuclei [34] (Fig. 3D).

Fig. 3.

Effects of γ-tocotrienol on NF- κB activation. (A) Western blot analysis of NF-κB p65 protein levels in myotubes pretreated with γ-tocotrienol at different concentrations prior to LPS exposure. α-Tubulin was used as the loading control. (B) Quantification of NF-κB p65 protein expression relative to α-tubulin. Data are presented as mean ± SD from three independent experiments. (C) Immunocytochemistry (ICC) analysis of NF-κB p65 (green) and Myh4 (red) in myotubes. DAPI (blue) was used to stain nuclei. Scale bar = 100 μm. (D) Quantification of myotube diameter and nuclear fusion index (NFI), calculated as the ratio of nuclei in myocytes with two or more nuclei to the total number of nuclei. Data are presented as mean ± SD. (E) Western blot analysis of NF-κB p65 protein levels in nuclear and cytosolic fractions of myotubes. Lamin B1 was used as the nuclear loading control, and α-tubulin as the cytosolic loading control. ∗p < 0.05, ∗∗p < 0.01 compared to the control group; #p < 0.05; ##p < 0.01; ###p < 0.001 compared to the LPS-treated group.

Nuclear and cytosolic fractionation corroborated these findings, demonstrating elevated nuclear NF-κB p65 levels following LPS treatment, which were attenuated in γ-tocotrienol-pretreated cells (Fig. 3E). These results indicate that γ-tocotrienol inhibits NF-κB activation and nuclear translocation, contributing to its protective effects against LPS-induced myotube atrophy.

3.4. Comparative protective effects of γ-tocotrienol and α-tocopherol against LPS-induced myotube atrophy

The protective effects of γ-tocotrienol and α-tocopherol on LPS-induced myotube atrophy were compared by pretreating C2C12 myotubes with each compound before LPS exposure. Morphological analysis revealed that both compounds mitigated LPS-induced atrophy, but γ-tocotrienol exhibited superior preservation of structural integrity (Fig. 4A). Western blot analysis revealed that γ-tocotrienol significantly reduced the upregulation of atrophy markers, including Fbxo32/Atrogin-1, and maintained higher levels of Myh1, a differentiation marker, compared to α-tocopherol (Fig. 4B and C). These findings highlight γ-tocotrienol's greater efficacy in protecting against both morphological and molecular hallmarks of LPS-induced myotube atrophy, suggesting its potential as a more effective therapeutic agent.

Fig. 4.

Differential effects of γ-tocotrienol and α-tocopherol on LPS-induced myotube atrophy (A) Effects of γ-tocotrienol and α-tocopherol on LPS-induced morphological changes in C2C12 myotubes. Scale bar = 100 μm. (B) Western blot analysis of atrophy-related protein expression, with α-tubulin as the loading control. (C) Quantification of protein expression relative to α-tubulin. Data are presented as mean ± SD from three independent experiments. ∗p < 0.05, ∗∗p < 0.01 compared to the control group; #p < 0.05, ##p < 0.01 compared to the LPS-treated group.

3.5. Proteomic insights into γ-Tocotrienol's protective mechanisms

To elucidate the molecular mechanisms underlying the protective effects of γ-tocotrienol and α-tocopherol against LPS-induced myotube atrophy, an isobaric label-based quantitative proteomics analysis was conducted. A total of 4,003 quantifiable protein groups were identified from 23,902 peptides. Principal component analysis (PCA) revealed four distinct clusters, with γ-tocotrienol-treated myotubes exhibiting a protein expression profile more closely resembling the control group compared to α-tocopherol-treated myotubes (Fig. 5A).

Fig. 5.

Proteomic and functional enrichment analysis of differentially expressed proteins in C2C12 myotubes treated with LPS and γ-tocotrienol. (A) Principal component analysis (PCA) of proteomic profiles under different treatments. (B) Heatmap and volcano plot of differentially expressed proteins identified by ANOVA (FDR = 0.01). (C) Overlap analysis of DEPs between the LPS-only group and the LPS + γ-tocotrienol group. (D) Protein-protein interaction (PPI) network of overlapping DEPs generated using STRING. Proteins downregulated in the LPS + γ-tocotrienol group are marked in blue, upregulated proteins in red, and mitochondria-related proteins are highlighted in a green box. (E) Functional enrichment analysis of upregulated (orange) and downregulated (purple-blue) proteins in the γ-tocotrienol vs. LPS group, using Enrichr with the Wikipathway 2023 database.

LPS treatment induced significant changes in protein expression, with 172 proteins upregulated and 133 proteins downregulated compared to the vehicle control (Table S1). Pathway enrichment analysis revealed that upregulated proteins were predominantly associated with pro-inflammatory responses, including C3, P2RX4, HMOX1, and NFKB2 (Table S3). In contrast, downregulated proteins were linked to extracellular matrix (ECM) organization, such as COL1A1 and ITGB1 (Table S4).

γ-Tocotrienol pretreatment mitigated these effects, with 23 proteins upregulated and 49 downregulated compared to the LPS-only group. Notably, 27 proteins overlapped with those upregulated by LPS (Fig. 5B, Table S2, Table S6). γ-Tocotrienol reversed LPS-induced proteomic changes by reducing the expression of oxidative stress-related proteins and enhancing those associated with ECM organization and energy metabolism (Fig. 5E, Table S5). Approximately one-third of the proteins upregulated by LPS were restored to normal levels following γ-tocotrienol pretreatment (Fig. 5B).

Protein-protein interaction and functional enrichment analyses revealed that γ-tocotrienol primarily restored proteins involved in mitochondrial functions, metal ion homeostasis, structural molecule activity, and myofibril assembly (Fig. 5D). Notably, the downregulation of superoxide dismutase 2 (Sod2) expression in γ-tocotrienol-treated myotubes suggested reduced oxidative stress (Fig. 5D, Table S2).

These findings highlight γ-tocotrienol's ability to modulate critical cellular pathways, particularly those related to oxidative stress and ECM integrity, thereby protecting against LPS-induced myotube atrophy.

3.6. γ-tocotrienol protects against LPS-induced mitochondrial dysfunction and oxidative stress

Gene set enrichment analysis (GSEA) revealed that pathways associated with mitochondrial fusion and apoptotic mitochondrial changes were significantly enriched among downregulated proteins in γ-tocotrienol-pretreated C2C12 myotubes, indicating a protective effect on mitochondrial integrity in response to LPS exposure (Fig. 6A). JC-1 and MitoTracker staining further confirmed this protective effect, as γ-tocotrienol-pretreated myotubes maintained mitochondrial integrity and membrane potential despite LPS-induced damage (Fig. 6B, Fig. S1A). Immunostaining for translocase of outer mitochondrial membrane 20 (TOMM20) corroborated these findings, demonstrating that γ-tocotrienol mitigated LPS-induced reductions in mitochondrial mass and metabolic activity, thereby preventing mitochondrial damage-related myotube atrophy (Fig. 6C).

Fig. 6.

γ-Tocotrienol protects C2C12 myotubes from atrophy by maintaining mitochondrial integrity. (A) Gene Set Enrichment Analysis (GSEA) of mitochondria-related pathways and the corresponding protein expression levels in each gene set. (B) Representative fluorescence images of JC-1 dye staining for mitochondrial membrane potential. (C) Representative fluorescence images of TOMM20 staining for mitochondrial mass. Scale bar = 100 μm. Data are presented as mean ± SD from three independent experiments. ∗∗p < 0.01 compared to the control group; ##p < 0.05 compared to the LPS-treated group.

Given the established antioxidant properties of vitamin E isoforms, mitochondrial reactive oxygen species (mtROS) and total ROS levels were assessed. LPS treatment significantly elevated mtROS in C2C12 myotubes; however, γ-tocotrienol pretreatment effectively prevented mtROS accumulation, with effects comparable to those observed in α-tocopherol-treated cells (Fig. 7A). A similar trend was observed for total ROS levels, where γ-tocotrienol significantly attenuated LPS-induced ROS accumulation (Fig. S1B).

Fig. 7.

γ-Tocotrienol prevents LPS-induced mitochondrial oxidative stress and preserves mitochondrial respiration in C2C12 myotubes. (A) Mitochondrial superoxide levels were measured using MitoSOX staining and fluorescence microscopy. Representative images show (top) nuclear staining with DAPI, (middle) MitoSOX fluorescence, and (bottom) merged images of MitoSOX and DAPI. (B) Oxygen consumption rate (OCR) was measured using a Seahorse XFe24 analyzer. Sequential injections of oligomycin (1 μM), FCCP (3 μM), and rotenone/antimycin-A (1 μM) were added at the indicated time points (dashed lines). (C) Quantification of basal OCR, maximal respiration, non-mitochondrial oxygen consumption, ATP production OCR, coupling efficiency, and spare respiratory capacity. Data are presented as mean ± SEM from three independent experiments (n = 3). ∗p < 0.05, ∗∗p < 0.01 compared to the control group; #p < 0.05, ##p < 0.01 compared to the LPS-treated group.

To further evaluate the impact of γ-tocotrienol on mitochondrial respiratory function, the oxygen consumption rate (OCR) was analyzed. LPS exposure led to a marked reduction in basal OCR, maximal respiration, and ATP production in C2C12 myotubes. Pretreatment with γ-tocotrienol or α-tocopherol effectively mitigated these LPS-induced reductions in OCR parameters (Fig. 7B and C). Interestingly, while coupling efficiency and spare respiratory capacity were unaffected, non-mitochondrial oxygen consumption increased in myotubes pretreated with γ-tocotrienol or α-tocopherol.

3.7. γ-tocotrienol inhibits LPS‐induced muscle loss and muscle strength in vivo

To evaluate whether the in vitro findings could be replicated in an in vivo context, an animal study was performed using male C57BL/6 mice treated with LPS (Fig. 8A). Mice were pretreated with different doses of γ-tocotrienol and α-tocopherol for four weeks prior to LPS administration. Following LPS injection, all groups except the untreated control exhibited a significant drop in body weight. However, body weight gradually recovered over the subsequent two weeks, with the LPS-only group showing the greatest weight loss at the study endpoint (Fig. 8B). Pretreatment with either γ-tocotrienol or α-tocopherol effectively prevented LPS-induced muscle strength loss, as assessed by grip strength normalized to body weight (Fig. 8C). Serum IL-6 levels were also significantly reduced in the pretreated groups compared to the LPS-only group, displaying a dose-dependent effect (Fig. 8D). Histological analysis revealed a reduction in muscle fiber cross-sectional area in the LPS group, whereas γ-tocotrienol pretreatment displayed slightly better protection than α-tocopherol, particularly in the high dose treatment (Fig. 8E and F).

Fig. 8.

γ-tocotrienol effectively prevent LPS-induced myotube atrophy in vivo. (A) Experiment setting of the in vivo study using LPS-induced atrophy model (n = 8). (B) Percentage body weight changes across the 6-week study. (C) Left: Measurement of grip strength test before and after LPS administration; Right: Grip strength normalized to (D) Comparison of serum IL-6 levels between groups. (E) Raw figure and H&E-stained tissue section of the gastrocnemius muscle. Scale Bar: 100 μm. (F) Quantification of cross sectional area of muscle fibre using imageJ. (G) Relative expression of atrophy-related protein was quantified using Western Blot. Data represented the mean ± SD (n = 5–8). ∗p < 0.05, ∗∗p < 0.01 vs. control group (vehicle only); #p < 0.05, ##p < 0.01 vs. LPS-treated group.

3.8. γ-tocotrienol protects muscle degradation via Sirt1/Pgc-1α axis

To further characterize LPS-induced muscle atrophy, the expression of the E3 ubiquitin ligases MuRF-1 and Atrogin-1 was analyzed in gastrocnemius muscle tissue. Consistent with in vitro results, LPS significantly upregulated both proteins, while pretreatment with γ-tocotrienol or α-tocopherol attenuated this increase (Fig. 8G). The expression of NF-κB p65 and Foxo3a was also examined to explore the inflammatory response. LPS treatment significantly increased NF-κB p65 expression, which was reduced by both forms of vitamin E, with γ-tocotrienol showing a slightly better inhibitory effect than α-tocopherol (Fig. 9A). A similar result was also observed with Foxo3a, where LPS induced a significant upregulation, while pretreatment with γ-tocotrienol suppressed Foxo3a expression more effectively compared to α-tocopherol. As proteomics study found the protection effect of γ-tocotrienol were primarily mediated through mitochondrial maintenance, the expressions of the mitochondrial biogenesis marker Pgc-1α and its upstream regulator Sirt1 were investigated. Both proteins were notably upregulated only in the γ-tocotrienol-treated groups, with minimal increases observed in mice treated with α-tocopherol (Fig. 9B). While Sirt1 expression was elevated in both γ-tocotrienol treatment groups, increased Pgc-1α expression was observed only in the higher-dose group.

Fig. 9.

γ-tocotrienol treatment protects LPS‐induced inflammatory response and mitochondrial dysregulation in muscle. Western blot analysis showing the protein expression of (A) SIRT1 and PGC-1α, (B) FOXO3a and NF-κB, (C) SOD2 and mitochondrial complex protein expression in response to γ-tocotrienol or α-tocopherol treatment in LPS-treated mice gastrocnemius muscle. α-tubulin was used as an internal control. (D) Immunohistological staining using Pgc-1α (green) and Sirt1 (red) antibodies on the harvested gastrocnemius muscle tissue section, with DAPI (blue) counterstaining. Scale Bar: 50 μm. Values are expressed as the mean ± SD (n = 5–8). ∗p < 0.05, ∗∗p < 0.01 vs. control group (vehicle only); #p < 0.05, ##p < 0.01 vs. LPS-treated group.

3.9. Mitochondrial damage protection by inhibition of mitochondrial ROS

Additionally, the expression of Sod2 and mitochondrial oxidative phosphorylation (OXPHOS) proteins was also assessed. Western blot analysis found increased γ-tocotrienol pre-treated mice expressed a higher level of Sod2 expression compared to other groups. Additionally, the dysregulation in mitochondria caused by LPS was observed with diminishing levels of the OXPHOS respiratory complexes, while both treatment group protects the muscle from the impaired mitochondrial function, γ-tocotrienol has shown a more significant protection compared to α-tocopherol (Fig. 9C). Immunohistological staining was also performed on harvested gastrocnemius muscle tissue sections. Both Pgc-1α and Sirt1 expression were reduced in the LPS-treated group. This decrease was prevented by pre-supplementation with either γ-tocotrienol or α-tocopherol. However, only the γ-tocotrienol group showed a significant increase in muscle fiber cross-sectional area. These results indicated that γ-tocotrienol is better in protecting cells from damaged mitochondrial function and muscle atrophy induced by LPS.

These findings highlight γ-tocotrienol's ability to preserve mitochondrial function, reduce oxidative stress, and sustain cellular energy metabolism under LPS-induced stress, further supporting its role as a protective agent against inflammation-associated mitochondrial dysfunction and myotube atrophy.

4. Discussion

Lipopolysaccharide (LPS) induces muscle atrophy through multiple mechanisms, primarily by triggering pro-inflammatory responses. LPS binds to toll-like receptors (TLRs), activating downstream signaling pathways such as NF-κB and MAPK, which drive the transcription of pro-inflammatory cytokines [36]. These inflammatory responses impair muscle function and contribute to muscle atrophy [37]. In this study, well-differentiated myotubes were treated with LPS to induce muscle atrophy, providing a model to examine the protective effects of γ-tocotrienol.

Previous research has demonstrated that different isoforms of vitamin E, such as α-tocopherol and γ-tocotrienol, exhibit distinct effects on muscle health. While both α-tocopherol and γ-tocotrienol are potent antioxidants, γ-tocotrienol uniquely suppresses COX- and 5-LOX-mediated eicosanoids to reduce inflammation—an effect not observed with α-tocopherol [18]. Tocotrienols, including γ-tocotrienol, have shown superior efficacy in reducing oxidative stress and inflammation in muscular dystrophy models, enhancing muscle stem cell function, and decreasing fibro-adipogenic progenitors [38]. Furthermore, γ-tocotrienol has demonstrated greater potency than tocopherols in lowering cholesterol levels, reducing inflammatory cytokines, and mitigating oxidative stress markers, suggesting its potential to improve muscle health and overall well-being [20].

Proteomic analysis in this study revealed that γ-tocotrienol modulates key molecular pathways associated with muscle atrophy. It suppressed the expression of pro-atrophy markers such as E3 ubiquitin ligases MuRF-1 (Trim63) and Fbxo32 (Atrogin-1), while enhancing muscle fiber markers like myosin heavy chain 1 (Myh1). γ-Tocotrienol also downregulated oxidative stress response pathways and upregulated proteins involved in calcium transport regulation and extracellular matrix organization, further contributing to its protective effects. These findings highlight γ-tocotrienol as a promising therapeutic strategy for muscle atrophy, necessitating further research to elucidate the distinct mechanisms of α-tocopherol and γ-tocotrienol in muscle health.

The measurement of oxygen consumption rate (OCR) revealed that γ-tocotrienol effectively mitigates LPS-induced mitochondrial dysfunction, with results comparable to the well-established antioxidant α-tocopherol. Interestingly, non-mitochondrial oxygen consumption was elevated in both γ-tocotrienol- and α-tocopherol-treated groups, potentially compensating for mitochondrial deficits. Although ATP production OCR levels were preserved, the reduced maximal OCR observed in γ-tocotrienol-treated groups suggests a protective metabolic shift that prioritizes ATP synthesis over maximal respiratory capacity [39]. This higher non-mitochondrial OCR may reflect a broader shift in metabolic pathways, highlighting the need for further studies on the interaction between mitochondrial and non-mitochondrial respiration in γ-tocotrienol-mediated protection.

LPS is known to disrupt zinc homeostasis by upregulating the expression of the metal transporter Slc39a14 (Zip14) through acute inflammatory responses [40]. Prior studies have linked increased Slc39a14 expression to muscle wasting [41], and our findings are consistent with this observation. Proteomic analysis revealed significant upregulation of Slc39a14 in LPS-treated C2C12 myotubes, a phenomenon that was partially prevented by γ-tocotrienol pretreatment. qPCR analysis corroborated these findings, showing concurrent changes in metallothionein 1 (Mt1) expression across treatments (Fig. S2). The disruption of zinc homeostasis is strongly associated with muscle atrophy, as evidenced by studies reporting Slc39a14 upregulation in cachectic muscles of mice with metastatic cancer [42]. Notably, the upregulation of Slc39a14 in muscle cells has been proposed as a direct driver of muscle atrophy, rather than a secondary effect of muscle wasting.

The animal study supports the protective effect of γ-tocotrienol against LPS-induced muscle atrophy. While both γ-tocotrienol and α-tocopherol were effective in preserving muscle strength and mitigating systemic inflammation, as evidenced by improved grip strength and reduced serum IL-6 levels, γ-tocotrienol demonstrated a more robust protective effect compared to α-tocopherol, particularly in preserving muscle fiber morphology and suppressing the expression of atrophy-related proteins MuRF-1 and Atrogin-1. While Western blot analysis showed that Akt activation did not significantly differ between groups (Fig. S6), Akt phosphorylation is known to be transient, and previous studies have suggested that LPS-induced muscle atrophy in vivo may occur independently of the Akt pathway [37]. Therefore, the enhanced protective effect of γ-tocotrienol against muscle atrophy was examined through alternative mechanisms, particularly the key regulators of mitochondrial biogenesis, Sirt1 and Pgc-1α. Indeed, γ-tocotrienol significantly upregulated these two proteins, suggesting that its protective role is closely linked to mitochondrial preservation These results are consistent with the in vitro findings and proteomic data, suggesting that γ-tocotrienol confers muscle-protective effects in part through maintenance of mitochondrial function.

Sirt1 and Pgc-1α are known to regulate mitochondrial biogenesis and overall muscle function. In skeletal muscle, the Sirt1/Pgc-1α axis contributes to the maintenance of mitochondrial content and function and also protects muscle fibers from oxidative stress and atrophy [43]. One previous study has shown that a tocotrienol-rich fraction can indeed upregulate this pathway to ameliorate skeletal muscle damage in diabetic mice [19]. Vitamin E has also been shown to upregulate Pgc-1α under normal conditions [44]. Although α-tocopherol did restore the affected Pgc-1α expression to some extent following LPS treatment, the present study demonstrated that γ-tocotrienol induced a significantly higher upregulation. Immunohistostaining also observed that LPS treatment reduced the expression of Sirt1 and Pgc-1α in gastrocnemius muscle, while pre-supplementation with either γ-tocotrienol or α-tocopherol was able to prevent this decline, suggesting that both compounds exert protective effects against LPS-induced downregulation of mitochondrial regulatory proteins, which is consistent with the in vitro findings. However, only γ-tocotrienol supplementation resulted in a significant increase in muscle fiber cross-sectional area. This might indicate that while both compounds can preserve molecular markers of mitochondrial function, γ-tocotrienol can provide better myoprotective or anabolic effects which is weaker in α-tocopherol treatment. The improvement in muscle fiber size observed with γ-tocotrienol may be related to enhanced mitochondrial biogenesis mediated by the enhanced Sirt1/Pgc-1α axis, leading to improved energy metabolism and muscle maintenance under inflammatory stress.

In the animal study, Sod2 was markedly downregulated by LPS, while γ-tocotrienol pretreatment significantly prevented this suppression. Interestingly, proteomic data in differentiated C2C12 myotubes shown a reduction in Sod2 following γ-tocotrienol. This may due to the differences of cell culture and animal model, where in systemic inflammation plays an important role in producing excessive ROS [45]. The elevated Sod2 expression suggested that γ-tocotrienol supplementation could respond to mitochondrial ROS in vivo more effectively compared to the relatively homogenous cell culture environment. This has shown the importance to consider the physiological complexity of whole organisms when studying antioxidant responses of γ-tocotrienol.

One of the limitations of the present study is the lack of discussion on the deacetylation activity of Sirt1. While the study focused on the protein expression levels of Sirt1 and Pgc-1α, it is well established that Sirt1 modulates cellular function through the deacetylation of key transcriptional regulators, including Pgc-1α and NF-κB [46]. Deacetylation of Pgc-1α enhances its transcriptional activity and promotes mitochondrial biogenesis, while deacetylation of the p65 subunit of NF-κB reduces its pro-inflammatory effect [47]. Future work measuring acetylation levels to study the post-translational modifications of these proteins may provide a more complete understanding of how γ-tocotrienol modulates these signaling cascades to protect against muscle atrophy.

Another important consideration in the application of γ-tocotrienol is its bioavailability, which is known to be affected by α-tocopherol. Studies have shown that α-tocopherol, commonly included in supplements, can reduce plasma concentrations of γ-tocotrienol, thereby impairing its efficacy [48]. When supplemented without tocopherols, δ-tocotrienol demonstrates superior bioavailability and can be metabolically converted into other tocotrienols and tocopherols, enhancing its potential to combat age-related and chronic disorders [49]. Besides, other research indicates that α-tocopherol exhibits significantly higher oral bioavailability than γ-tocotrienol due to differences in intestinal permeability [50], whereas tocotrienols are more readily taken up into cells compared to tocopherols [51]. In the present study, γ-tocotrienol demonstrated potent biological activity despite its faster clearance in cell culture studies, suggesting that its efficacy is not solely dependent on intracellular concentration (Fig. S5). The in vivo study clearly demonstrated biological efficacy, even though LC-MS/MS analysis found lower intracellular concentrations of γ-tocotrienol in C2C12 cells after 48 h of treatment. This may be due to its preferential accumulation in specific tissues or its ability to modulate key cellular pathways at lower concentrations [52]. These observations underscore the importance of considering both bioavailability and tissue-specific efficacy when evaluating tocotrienol-based interventions and highlight the need for optimized formulations of vitamin E isoforms. Lastly, this study was limited to a male-only cohort, which restricts the generalizability of the findings. Previous research has demonstrated that sex-specific differences influence vitamin E metabolism [33,53] and susceptibility to muscle atrophy [34,54]. Therefore, while the present results highlight the therapeutic potential of γ-tocotrienol, future studies incorporating both male and female subjects are warranted to account for sex-specific biological differences and to ensure broader translational relevance.

5. Conclusion

Overall, this study demonstrated that γ-tocotrienol effectively prevents the onset of myotube atrophy induced by LPS in vitro and in vivo. LPS treatment of C2C12 myotubes and male mice led to characteristic atrophy symptoms, including reduced myofiber diameter and muscle strength. Notably, pre-treatment with γ-tocotrienol, in contrast to α-tocopherol, significantly inhibited the LPS-induced upregulation of the inflammatory cytokine NF-κB and atrophy-associated proteins MuRF-1 and Atrogin-1, while increasing the expression of mitochondrial biogenesis markers Sirt1 and Pgc-1α. Additionally, γ-tocotrienol effectively reduced ROS accumulation and the expression of Slc39a14, mitigating oxidative stress and protecting against mitochondrial damage. These findings provide evidence that γ-tocotrienol is a promising bioactive compound with muscle-protective properties, supporting its potential use as a dietary intervention to prevent inflammation-induced muscle wasting.

CRediT authorship contribution statement

Jun Yi Chong: Writing – original draft, Methodology, Investigation. Tsui-Chin Huang: Methodology, Investigation, Formal analysis. Sheng-Ming Chueh: Methodology, Formal analysis. Cheng-Yi Ma: Methodology. Tzu-Ting Kuo: Methodology. Jia-Jun He: Formal analysis. Yii-Jwu Lo: Formal analysis. Kuan-Chieh Peng: Formal analysis. Mohamed Ali: Resources. Hsin-Yi Chang: Writing – review & editing, Supervision, Resources, Project administration, Funding acquisition, Conceptualization. Shih-Min Hsia: Writing – review & editing, Supervision, Resources, Project administration, Conceptualization.

Ethical approval

Not applicable.

Declaration of competing interest

The authors have declared no conflict of interest.

Appendix A. Supplementary data

The following is the Supplementary data to this article:

Data availability

The datasets generated and analyzed in this study are available in the Japan ProteOme STandard Repository Database (jPOST, https://jpostdb.org/) [55], with the provided accession numbers JPST003208. Other data will be made available on request.