C2C12 myoblasts (Cat# ZQ0092, Zhong Qiao Xin Zhou Biotechnology, Shanghai, China) were cultured in high-glucose DMEM (Cat# 610502, Biopico, San Diego, CA, USA) supplemented with 10% fetal bovine serum (Cat# 086-150, Wisent, Saint-Jean-Baptiste, QC, Canada). All cell lines were authenticated by short tandem repeat (STR) profiling and tested negative for mycoplasma contamination prior to use. Cells were maintained at 37 °C in a humidified incubator with 5% CO₂ until they reached approximately 80% confluence. The culture medium was then replaced with differentiation medium comprising high-glucose DMEM and 2% horse serum (Cat# 16050130, Gibco, Grand Island, NY, USA), with media changes every 24 h. Overexpression or knock-down of XLOC_015548 was achieved via lentiviral vectors targeting mouse LncRNA XLOC_015548 (Hanbio, Shanghai, China). The full-length transcript for overexpression was cloned into the pHBLV-CMV-MCS-EF1-puro lentiviral vector under the control of the CMV promoter for constitutive expression. The shRNA sequence for XLOC_015548 knock-down (sense strand: 5^′-GATCCGCAGGGAGCTGTTGTCACCTTTGAAACTCGAGTTTCAAAGGTGACAACAGCTCCCTGTTTTTTG-3^′; antisense strand: 5^′-AATTCAAAAAACAGGGAGCTGTTGTCACCTTTGAAACTCGAGTTTCAAAGGTGACAACAGCTCCCTGCG-3^′) was subcloned into the pHBLV-U6-MCS-PGK-PURO interference vector (Hanbio, Shanghai, China). Overexpression of full-length Gadd45g was achieved using the PCDH-CMV-EF1-Puro lentiviral vector (ShuTong Biotech, Shanghai, China). In dual lentiviral infection experiments, C2C12 cells were subjected to four distinct conditions: the control cohort was co-infected with lentiviruses carrying the XLOC overexpression control (OE-Ctrl) and the Gadd45g overexpression control (OE-Ctrl); single-gene overexpression was achieved by infecting cells with OE-XLOC_015548 in combination with the Gadd45g OE-Ctrl; dual-gene co-overexpression involved simultaneous infection with OE-XLOC_015548 (MOI = 20) and OE-Gadd45g (MOI = 10); and for mechanistic validation, cells dually infected with OE-XLOC_015548 and OE-Gadd45g were treated with 10 µM MEK inhibitor U0126 for 48 hours prior to analysis. Stable cell lines were established through selection with 2 µg/mL puromycin (Cat# A1113803, Thermo Fisher Scientific, Waltham, MA, USA), following lentiviral transduction, which was enhanced using 8 µg/mL polybrene (Cat# TR-1003, Sigma-Aldrich, St. Louis, MO, USA). Infection efficiency was confirmed via real-time RT-PCR and Western blotting. For inhibitor experiments, the MEK inhibitor U0126 (Cat# U0126-EtO, Selleck, Shanghai, China) was dissolved in DMSO to prepare a 10 mM stock solution, then diluted in medium to a final concentration of 10 µM. The cells were treated with U0126 for 48 h prior to analysis. In therapeutic experiments involving XLOC_015548, the control and Dexamethasone (DEX) groups used OE-Ctrl lentiviral cell lines, while the DEX+OE-XLOC and DEX+OE-XLOC+U0126 groups used OE-XLOC cell lines. DEX and treatment groups were cultured in differentiation medium containing 10 µM dexamethasone (Dex, Cat# ST1254, Beyotime, Shanghai, China) or 10 µM U0126, while the control group was cultured in medium containing equivalent volumes of ethanol and DMSO.

Fluorescence in situ hybridization (FISH) was performed using biotin-labeled RNA probes specifically targeting XLOC_015548 mRNA. The probe sequence (5^′-GCTTCTTAATCTTCTGCTTAGGTGATGTATCCAGGATGGACCAATGGTACCCAAGGGATCAACTCTGCAAACAATCAGGTGGGCAGCGTCTTGTCATTATGGACCATGCTTTAAACTTTCTCAT-3^′) was synthesized and biotin-labeled by SaiCheng Biotech (Guangzhou, China). Hybridization was conducted at 42 °C for 2 h, followed by washes with 5X SSC and 1X SSC at room temperature. Fluorescent detection was performed with a Leica STELLARIS 5 laser scanning confocal microscope (Leica, Wetzlar, Germany). DAPI (Cat# D9542, Sigma, St. Louis, MO, USA) was used for nuclear staining (blue, Ex/Em = 405/420–566 nm), while FISH signals for XLOC_015548 were visualized as red fluorescence (Ex/Em = 561/566–716 nm). Three-dimensional (3D) imaging was captured over a 20 µm range, and 10 consecutive 2 µm slices were compiled into a composite 3D image. Co-localization analysis was performed using the Co-localization Finder plugin (version 1.8) in ImageJ (version 1.53k, National Institutes of Health, Bethesda, MD, USA). Three rounds of gating were performed for different groups, with consistent gating strategies across groups to obtain Pearson’s correlation coefficients for co-localized signals. In scatter plots and fluorescence merge images, co-localized signals from each replicate were marked in red, green, and blue, respectively. Further analysis was conducted using the Plot Profile function in ImageJ to examine pixel intensity distribution along characteristic lines, enabling the evaluation of FISH signal localization and intensity relative to DAPI staining.

During differentiation, images of myotubes were captured on days 1, 3, 5, and 7 using confocal immunofluorescence microscopy (STELLARIS 5, Leica, Wetzlar, Germany) and phase-contrast microscopy (AXIO Observer 3, Carl Zeiss, Oberkochen, Germany). The average myotube width was calculated from randomly selected fields within each group of cells. To ensure representative data, at least five fields per sample were analyzed to capture variations across the sample and minimize potential bias.

Cell cycle analysis was performed using the DNA Content Quantitation Assay Kit (Cell Cycle) (Cat# CA1510, Solarbio, Wuhan, China). After treatment, cells were collected, washed once with PBS (Cat# G4202, Servicebio, Wuhan, China), and centrifuged at 1500 rpm for 5 minutes. The cell concentration was adjusted to 1 $\times$ 10⁶ cells/mL and a 1 mL aliquot of the single-cell suspension was centrifuged, the supernatant discarded, and the cell pellet fixed in 500 µL of pre-cooled 70% ethanol at 4 °C for 2 h. Prior to staining, the fixation solution was removed by washing the cells with PBS. If necessary, a 200-mesh cell strainer was used to ensure a uniform cell suspension. The cell pellet was resuspended in 100 µL RNase A solution and incubated at 37 °C in a water bath for 30 minutes. Subsequently, 400 µL of PI staining solution was added, mixed thoroughly, and incubated at 4 °C in the dark for 30 minutes. Finally, the samples were analyzed with a CytoFLEX S Flow Cytometer (Beckman Coulter Life Sciences, Indianapolis, IN, USA), with red fluorescence signals recorded at an excitation wavelength of 488 nm.

Apoptosis was detected using an APC Annexin V Apoptosis Detection Kit with Propidium Iodide (PI) (Cat# 640932, BioLegend, San Diego, CA, USA). Cells were washed twice with cell staining buffer (Cat# 420201, BioLegend), resuspended in Annexin V binding buffer (Cat# 422201, BioLegend) at a concentration of 0.25–1.0 $\times$ 10⁶ cells/mL, followed by the addition of 5 µL APC-Annexin V and 10 µL PI solution. Samples were incubated at room temperature (25 °C) for 15 minutes in the dark, followed by the addition of 400 µL binding buffer and subsequent analysis using a NovoCyte Flow Cytometer (Agilent Technologies, Santa Clara, CA, USA).

Total RNA was extracted from C2C12 cells using TRIzol reagent (Cat# R0016, Beyotime, Shanghai, China) following the manufacturer’s protocol. RNA integrity and concentration were assessed using a NanoDrop Spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA). cDNA synthesis was performed using the PrimeScript RT reagent kit with gDNA Eraser (Cat# RR047A, TaKaRa, Shiga, Japan). Quantitative real-time PCR (qPCR) was conducted using the TB Green Premix DimerEraser (Cat# RR091A, TaKaRa, Kyoto, Japan) on a LightCycler 480 qPCR system (Roche, Basel, Switzerland). Gene expression was normalized to GAPDH, and relative expression levels were calculated using the 2^{-Δ⁢Δ⁢Ct} method. Primer sequences used were: XLOC_015548: Forward, 5^′-CCTCAGCAGACCCTGACTGTAG-3^′; Reverse, 5^′-CAGTGGCTGTCTTAGTCCATCTCA-3^′. For GAPDH: Forward, 5^′-AGGTCGGTGTGAACGGATTTG-3^′; Reverse, 5^′-TGTAGACCATGTAGTTGAGGTCA-3^′. The 20 µL qPCR reaction included 2 µL diluted cDNA, 10 µL TB Green Premix DimerEraser, 0.4 µL of each primer (10 µM), and 7.2 µL nuclease-free water. Reactions were conducted as follows: 95 °C for 30 seconds (initial denaturation), followed by 40 cycles of 95 °C for 5 seconds (denaturation) and 60 °C for 34 seconds (annealing/extension). All reactions were performed in triplicate.

Proteins were extracted using RIPA lysis buffer (Cat# P0013B, Beyotime, Shanghai, China) supplemented with protease and phosphatase inhibitors (Cat# P1045, Beyotime, Shanghai, China). Protein concentrations were determined using a BCA Protein Assay Kit (Cat# P0398S, Beyotime, Shanghai, China). Equal amounts of protein were mixed with 6X SDS-PAGE sample loading buffer (Cat# P0015F, Beyotime, Shanghai, China) and separated via 10% SDS-PAGE (Cat# WB2102, Biotyscience, Shanghai, China). Proteins were transferred onto PVDF membranes (Cat# 0000191584, Millipore, Burlington, MA, USA) using a BIO-RAD wet transfer system. Membranes were blocked with QuickBlock™ Blocking Buffer (Cat# P0240, Beyotime, Shanghai, China) for 1 h at room temperature and then incubated overnight at 4 °C with the following primary antibodies: GADD45g (Cat# FNab03295, 1:500, FineTest, Wuhan, China); Phospho-MEK1/2 (Ser218/Ser222) (Cat#bsm-52176R, 1:500, Bioss, Beijing, China); MEK1 + MEK2 (Cat# ab178876, 1:20,000, Abcam, Cambridge, UK); Phospho-ERK1/2 (Thr202/Tyr204) (Cat# bs-3016R, 1:2000, Bioss, Beijing, China); ERK1 + ERK2 (Cat# bs-0022R, 1:1000, Bioss, Beijing, China); Myosin Heavy Chain (MHC) (Cat# MF20, 1:500, DSHB, Iowa City, IA, USA); Anti-iNOS Rabbit pAb (Cat# GB115703, 1:1000, Servicebio, Wuhan, China); Beta-tubulin (Cat# GB11017, 1:2000, Servicebio, Wuhan, China); Lamin B1 Antibody (Cat# FNab04682, 1:1000, Wuhan, China). Blots were washed with TBST and incubated with HRP-conjugated goat anti-rabbit IgG (Cat# GB23303, 1:10,000, Servicebio, Wuhan, China) or goat anti-mouse IgG (Cat# GB23301, 1:10,000, Servicebio, Wuhan, China) at room temperature for 1 h. Chemiluminescent detection was performed using the BeyoECL Moon Kit (Cat# P0018FM, Beyotime, Shanghai, China). Band intensities were quantified using ImageJ and normalized to beta-tubulin.

To investigate the regulatory effect of XLOC_015548 on the subcellular localization of Gadd45g, we employed nuclear-cytoplasmic fractionation combined with Western blotting for quantitative analysis. Nuclear and cytoplasmic proteins were extracted from C2C12 cells using a nuclear-cytoplasmic extraction kit (Cat# P0097, Beyotime, Shanghai, China). First, the mock-virus groups were verified to have no interference effect, with the following groupings: Untransfected group (Untransfected): normal cultured C2C12 cells; Mock-KD group: transfected with pHBLV-U6 empty vector (KD-NC); Mock-OE group: transfected with pHBLV-CMV empty vector (OE-Ctrl).To investigate the effect of XLOC_015548 on the subcellular localization of Gadd45g, the cell groups were as follows: Control group (Mock-KD, KD-NC): transfected with pHBLV-U6 empty vector (KD-NC); XLOC_015548 knockdown group (KD-XLOC) and overexpression group (OE-XLOC). Each group was performed in triplicate biological repeats.Western blot analysis was performed, and nuclear Gadd45g was normalized to Lamin B1, cytoplasmic Gadd45g was normalized to $\beta$-tubulin, and total Gadd45g was normalized to whole-cell $\beta$-tubulin. The nuclear-cytoplasmic ratio was calculated using the formula: nuclear Gadd45g (normalized to Lamin B1)/cytoplasmic Gadd45g (normalized to $\beta$-tubulin). Other western blot procedures followed the previously described methodology.

Cells were cultured on glass-bottom culture dishes (Cat#801001, NEST, Wuxi, China) until they adhered and proliferated normally. At the designated time points, cells were fixed with cell fixation solution (Cat# P0098, Beyotime, Shanghai, China) and washed three times with wash buffer (Cat# P0106, Beyotime, Shanghai, China) for 5 minutes each. Cells were permeabilized with permeabilization buffer (Cat# P0097, Beyotime, Shanghai, China) for 40 minutes and blocked with blocking solution (Cat# P0102, Beyotime, Shanghai, China) for 100 minutes to minimize nonspecific binding. Primary antibodies were diluted in primary antibody dilution buffer (Cat# P0103, Beyotime, Shanghai, China) and included rabbit anti-phospho-MEK1/2 (Ser218 + Ser222) (bsm-52176R, 1:100, Bioss, Beijing, China), rabbit anti-phospho-ERK1/2 (Thr202 + Tyr204) (Cat# bs-3016R, 1:100, Bioss, Beijing, China), rabbit anti-GADD45G (Cat# bs-7069R, 1:200, Bioss, Beijing, China), myosin heavy chain, sarcomere (MHC) (Cat# MF 20, 1:150, DSHB, Iowa City, IA, USA), MyoD (Cat# sc-71629, 1:50, Santa Cruz Biotechnology, Dallas, TX, USA), MYOD1 polyclonal antibody (Cat# 18943-1-AP, 1:150, Proteintech, Rosemont, IL, USA), and Mki67 rabbit polyclonal antibody (Cat# A25399, 1:150, ABclonal, Woburn, MA, USA). After overnight incubation at 4 °C with gentle shaking, the primary antibodies were recovered and the cells washed three times with wash buffer for 5 minutes each. Secondary antibodies, including Alexa Fluor® 647-conjugated goat anti-mouse IgG H&L (Cat# ab150115, 1:400, Abcam, Cambridge, UK) and Alexa Fluor® 488-conjugated goat anti-rabbit IgG H&L (Cat# ab150077, 1:400, Abcam, Cambridge, UK) were diluted in secondary antibody dilution buffer (Cat# P0108, Beyotime, Shanghai, China) and added to the samples. After 1 h incubation at room temperature, the secondary antibodies were recovered and the cells washed three times. Actin-Tracker Red-Rhodamine (Cat# C2207S, 1:2000, Beyotime, Shanghai, China) was then added for F-actin staining. Finally, after three washes with wash buffer, cells were mounted using anti-fluorescence quenching mounting medium with DAPI (Cat# P0131, Beyotime, Shanghai, China). Fluorescence detection was performed with a laser scanning confocal microscope (Leica, STELLARIS 5, Germany). DAPI staining was used to label nuclei in blue fluorescence (Ex/Em = 405/493–558 nm). P-MEK1/2, P-ERK1/2, GADD45G, Ki-67, and MYOD1 were labeled with rabbit primary antibodies and detected with green fluorescence (Ex/Em = 488/493–558 nm). F-actin, representing the structural remodeling of the cytoskeleton during C2C12 differentiation, was labeled in yellow fluorescence (Ex/Em = 561/566–643 nm). MHC and MyoD were detected with mouse primary antibodies and labeled with red fluorescence (Ex/Em = 638/644–818 nm). Co-localization analysis was conducted using the Co-localization Finder plugin (version 1.8) in ImageJ. Three rounds of gating were performed to obtain three Pearson correlation coefficients for co-localized signals. These coefficients were visualized in scatter plots and fluorescence dual-color merge images, with the co-localized signals from the three replicates marked in red, green, and blue, respectively.

TUNEL staining combined with dual immunofluorescence for Gadd45g and muscle markers (Myog or MHC) was performed to assess the expression level of Gadd45g during myotube maturation. Cells cultured on glass-bottom dishes were fixed with cell fixation solution (Cat# P0098, Beyotime, Shanghai, China) and washed twice with wash buffer for 5 minutes each. They were then permeabilized with permeabilization buffer (Cat# P0097, Beyotime, Shanghai, China) for 5 minutes and washed twice. A TUNEL reaction mixture (5 µL TdT enzyme + 45 µL fluorescent labeling solution) was prepared using the One-Step TUNEL Cell Apoptosis Detection Kit (Cat# KGA1408-50, KeyGEN, Nanjing, China) and incubated at 37 °C for 60 minutes in the dark. After TUNEL staining, cells were washed three times with PBS for 5 minutes each. Primary antibodies were diluted in primary antibody dilution buffer and two staining schemes were used: Scheme 1 with rabbit anti-GADD45G (Cat# bs-7069R, 1:200, Bioss, Beijing, China) and mouse anti-Myogenin (Cat# ab1835, 1:800, Abcam, Cambridge, UK), and Scheme 2 with rabbit anti-GADD45G and mouse anti-MHC (Cat# MF 20, 1:150, DSHB, Iowa City, IA, USA). After overnight incubation at 4 °C, cells were washed and stained with secondary antibodies (Alexa Fluor® 647 and Alexa Fluor® 488) for 1 h at room temperature. Finally, cells were washed and mounted with DAPI-containing mounting medium. Fluorescent signals were visualized with a confocal microscope (Leica, Germany). TUNEL-positive cells were marked as yellow (Ex/Em = 561/566–643 nm), Gadd45g as green (Ex/Em = 488/494–588 nm), and the markers Myogenin and MHC as red (Ex/Em = 638/643–817 nm).

Giemsa staining was used to observe myotube morphology. Cells cultured on glass-bottom dishes were fixed with methanol for 5 minutes at room temperature. A working solution was prepared by mixing 1 mL Giemsa concentrate with 9 mL Giemsa diluent (Cat# CU0001, Solarbio, Wuhan, China). The working solution was applied to cover the cells for 30 minutes at room temperature. After staining, cells were gently rinsed with distilled water to remove excess stain and then observed under an inverted light microscope (DMi8, Leica, Germany).

Cellular senescence was assessed using the Senescence $\beta$-Galactosidase Staining Kit (Cat#C0602, Beyotime, Shanghai, China). Cells were fixed with 1 mL $\beta$-galactosidase fixative for 15 minutes at room temperature after PBS washing. Fixed cells were then washed three times with PBS, and 1 mL staining solution (10 µL each of solutions A, B, and C diluted with 930 µL of distilled water) was added to each well. Cells were incubated overnight at 37 °C and observed under a light microscope (DMi8, Leica, Germany).

Cell proliferation was analyzed using the BeyoClick™ EdU-488 Cell Proliferation Assay Kit (Cat# C0078S, Beyotime, Shanghai, China). Cells were cultured on glass-bottom culture dishes (Cat# 801001, NEST, Wuxi, China) until fully adhered and proliferating. At the designated time points, 500 µL of 20 µM EdU working solution prewarmed to 37 °C was added to each well and incubated for 5 h to label proliferating cells. After incubation, the medium was discarded and cells were fixed with 1 mL of fixation solution (Cat# P0098, Beyotime, Shanghai, China) for 15 minutes at room temperature. Cells were then washed three times with 1 mL of wash buffer for 5 minutes each. Permeabilization was performed with 1 mL of strong permeabilization solution (Cat# P0097, Beyotime, Shanghai, China) at room temperature for 20 minutes, followed by three additional washes. Staining solution (1 µL Azide 488 + 20 µL CuSO₄ + 50 µL Click Additive Solution + 430 µL Click Reaction Buffer) prepared according to the manufacturer’s protocol was added to each well (500 µL per well) and incubated at room temperature in the dark for 30 minutes. After staining, the solution was removed and the cells washed three times with wash buffer, mounted with DAPI-containing antifade mounting medium (Cat# P0131, Beyotime, Shanghai, China), and observed using a laser scanning confocal microscope (STELLARIS 5, Leica, Wetzlar, Germany). Fluorescence signals were detected as blue for DAPI (Ex/Detectors = 405/420–503 nm) and green for EdU-positive nuclei (Ex/Em = 488/503–709 nm).

Intracellular ROS levels were detected using the Reactive Oxygen Species Assay Kit (Cat# C13000-2, Applygen, Beijing, China). C2C12 cells were cultured on glass-bottom dishes (Cat# 801001, NEST, Wuxi, China). After removing the medium, 500 µL of serum-free DMEM containing 10 µM dihydroethidium (DHE) probe was added and the cells were incubated at 37 °C for 60 minutes. They were then washed thoroughly with serum-free medium to remove excess probe, and Hoechst 33342 live-cell staining solution (Cat# C1027, Beyotime, Shanghai, China) was applied for 30 minutes, followed by three PBS washes. Fluorescence signals were observed using a laser scanning confocal microscope (STELLARIS 5, Leica, Wetzlar, Germany). Hoechst 33342 emitted blue fluorescence (Ex/Detectors = 405/420–566 nm), while oxidized DHE resulting from intracellular ROS activity emitted bright red fluorescence (Ex/Em = 561/566–716 nm).

Lipid peroxidation was assessed using the Lipid Peroxidation Probe BDP 581/591 C11 (Cat#L267, Tocris, Tokyo, Japan). C2C12 cells were cultured on glass-bottom dishes (Cat# 801001, NEST, Wuxi, China) until fully adhered. After removing the medium, cells were washed three times with serum-free DMEM and incubated with 500 µL of BDP 581/591 C11 working solution for 30 minutes at 37 °C in a 5% CO₂ incubator. Cells were washed three times with serum-free medium and then stained for 30 minutes with Hoechst 33342 live-cell staining solution (Cat#C1027, Beyotime, Shanghai, China). After three washes with PBS, fluorescence was detected with a laser scanning confocal microscope (STELLARIS 5, Leica, Wetzlar, Germany). Lipid peroxidation-associated radicals converted the probe’s fluorescence to green (Ex/Em = 488/493–553 nm), while unreacted probe emitted red fluorescence (Ex/Em = 561/566–712 nm).

Mitochondrial membrane potential (MMP) was assessed using the Rhodamine 123-based MMP Assay Kit (Cat# C2008S, Beyotime, Shanghai, China). C2C12 cells were cultured on glass-bottom dishes (Cat# 801001, NEST, Wuxi, China). After removing the medium, cells were washed three times with serum-free DMEM. Subsequently, 500 µL of Rhodamine 123 staining working solution (0.5 µL Rhodamine 123 + 500 µL detection buffer) was added and the cells were incubated at 37 °C for 60 minutes. After staining, cells were washed three times with prewarmed DMEM and stained with Hoechst 33342 (Cat# C1027, Beyotime, Shanghai, China) for 30 minutes. Cells were then washed three times with PBS and fluorescence detected with a laser scanning confocal microscope (STELLARIS 5, Leica, Wetzlar, Germany). Hoechst 33342 emitted blue fluorescence (Ex/Detectors = 405/420–503 nm), and Rhodamine 123 emitted green fluorescence dependent on the MMP (Ex/Detectors = 488/503–709 nm).

Cell viability and cytotoxicity were evaluated using the Calcein/PI Cell Viability and Cytotoxicity Assay Kit (Cat# C2015, Beyotime, Shanghai, China). Both adherent and trypsinized cells were assessed. For adherent cells, C2C12 cells were cultured on glass-bottom dishes (Cat# 801001, NEST, Wuxi, China) until fully adhered. After removing the medium, 500 µL of Calcein AM/PI working solution was added and the cells incubated at 37 °C in the dark for 30 minutes. After three washes with prewarmed PBS, fluorescence was observed with a laser scanning confocal microscope (STELLARIS 5, Leica, Wetzlar, Germany). For trypsinized cells, the cells were first detached using trypsin, centrifuged at 250–1000 $\times$g for 5 minutes, washed with PBS, and centrifuged again. Cells were then resuspended in 500 µL of Calcein AM/PI working solution and incubated at 37 °C in the dark for 30 minutes. After staining, the cells were observed by placing the suspension in glass-bottom dishes. Calcein AM stains viable cells green (Ex/Em = 488/493–553 nm), while PI stains dead cells red (Ex/Em = 561/566–712 nm). Fluorescence was detected with a confocal microscope (STELLARIS 5, Leica, Wetzlar, Germany).

XLOC_015548 is located on chromosome 16 and spans the region 86,044,931–86,047,144. CRISPR/Cas9 gene editing technology was employed to generate XLOC_015548 mutant mouse models. Myf5 is a well-established mouse-specific promoter widely used in skeletal muscle-specific conditional knockout mouse studies [46, 47, 48, 49]. The present study utilized conditionally overexpressed XLOC_015548 mice (XLOC_015548cKI/+; iCre⁺), overexpression control mice (XLOC_015548cKI/+; iCre⁻), conditionally deleted XLOC_015548 mice (XLOC_015548fl/fl; Cre⁺), and knock-out control mice (XLOC_015548fl/fl; Cre⁻). The F₀ and breeding mice were obtained from Jiangsu GemPharmatech Co., Ltd. (Nanjing, China).

Conditional Knockout Mouse Model: To generate the conditional knockout mice for XLOC_015548, we used CRISPR/Cas9 technology. The target CRISPR site was located in the XLOC_015548 gene on chromosome 16 (Chr16:86044931-86047144), with LoxP sites inserted approximately 5 kb upstream and 2 kb downstream of the coding region. Cas9 mRNA, sgRNA, and a donor vector with LoxP sequences were co-injected into C57BL/6J zygotes, producing chimeric F0 mice. These were then bred with Myf5-Cre mice for skeletal muscle-specific knockout (genotype: XLOC_015548fl/fl; Cre⁺), confirmed by PCR and sequencing.

Overexpression Mouse Model: For the overexpression model, we targeted the Myf5 gene, specifically the Myf5-201 transcript (ENSMUST00000000445.1). We inserted the iCre-P2A sequence upstream of the ATG start codon of Myf5, using the Myf5 promoter to drive expression. Cas9 and sgRNA were used to create a double-strand break, and homologous recombination facilitated the insertion. The insertion was confirmed by PCR and sequencing. The iCre expression was specific to skeletal muscle due to the Myf5 promoter.

Genotypes were confirmed using PCR and agarose gel electrophoresis. All animal-related procedures were approved by the Ethics Committee of Shenzhen Top-Biotech Co., Ltd., and carried out in accordance with the Guide for the Care and Use of Laboratory Animals established by the National Research Council.

Ten-week-old C57BL/6 mice of four genotypes (22–25 g): XLOC_015548cKI/+; iCre⁺, XLOC_015548cKI/+; iCre⁻, XLOC_015548fl/fl; iCre⁺, and XLOC_015548fl/fl; iCre⁻ (n = 12 per genotype). Each genotype group was equally divided into sciatic nerve transection-induced atrophy (n = 6) and sham-operated (n = 6) subgroups under standardized isoflurane anesthesia (3% induction, 1.5%–2% maintenance). Each group of 6 mice included 3 reserve mice to mitigate the risk of attrition due to potential mortality or loss during the experiment.Post-surgery, the mice were housed individually with free access to food and water. At two weeks post-denervation, the success of the sciatic nerve transection model was confirmed under isoflurane anesthesia. Mice were euthanized with excess isoflurane, and the sciatic nerve transection site was inspected to ensure successful establishment of the model. The right gastrocnemius muscle was harvested and weighed to evaluate muscle wet weight. Samples were processed for paraffin and frozen embedding for subsequent histological analyses, including measurement of the cross-sectional area (CSA) and dihydroethidium (DHE) staining to assess oxidative stress. Molecular-level studies were also performed by immunohistochemistry (IHC). Additionally, tissues from the heart, liver, spleen, lungs, and kidneys were collected for hematoxylin and eosin (HE) staining and analysis.

Gastrocnemius muscle samples (~0.5 cm $\times$ 0.5 cm $\times$ 0.25 cm) were fixed in GD muscle fixative (Cat#G1111, Servicebio, Wuhan, China), dehydrated, paraffin-embedded, and sectioned at ~7 µm thickness. Sections were stained with HE and immunohistochemically labeled for ERK, phospho-ERK (Thr202 + Tyr204), MEK, phospho-MEK (Ser218 + Ser222), Gadd45g, and MyoD. Additional muscle samples (~0.5 cm $\times$ 0.5 cm $\times$ 0.25 cm) were processed for frozen embedding using OCT compound (Cat# OCT4583SA, KURA, Osaka, Japan) and stored at –80 °C.

Paraffin sections were deparaffinized with xylene and rehydrated through graded ethanol to distilled water. Sections were stained with hematoxylin for 3 minutes, differentiated with 1% acid alcohol, washed, and counterstained with eosin for 30 seconds. After dehydration, clearing in xylene, and mounting with neutral resin, the sections were examined by light microscopy. The muscle cross-sectional area (CSA) was measured using Image-Pro Plus software (version 6.0, Media Cybernetics, Rockville, MD, USA). The polygon tool was used to outline typical muscle fibers in HE-stained sections. Measurements were performed using ImageJ . For each section, at least five randomly selected fields were analyzed to calculate the average myotube area. The selection of fields was randomized to minimize bias, and the final average was calculated from all fields across each experimental group.

Paraffin sections were deparaffinized, rehydrated, and subjected to antigen retrieval using pepsin-based antigen retrieval solution (Cat# DIG-3009, Maixin, Fuzhou, China). Endogenous peroxidase was blocked with peroxidase blocking solution (Cat# KIT-9710, MXB, Fuzhou, China). After blocking non-specific binding sites, sections were incubated overnight at 4 °C with the following primary antibodies: rabbit anti-MEK1/2 (Cat# bs-1041R, 1:100, Bioss, Beijing, China), rabbit anti-phospho-MEK1/2 (Ser218 + Ser222) (Cat# bsm-52176R, 1:200, Bioss, Beijing, China), rabbit anti-ERK1/2 (Cat# bs-0022R, 1:200, Bioss, Beijing, China), rabbit anti-phospho-ERK1/2 (Thr202 + Tyr204) (Cat# bs-3016R, 1:200, Bioss, Beijing, China), rabbit anti-MyoD1 (Cat# 18943-1-AP, 1:150, Proteintech, Rosemont, IL, USA), and rabbit anti-GADD45G (Cat# GB113573, 1:1000, Servicebio, Wuhan, China). After three washes with PBS, sections were incubated with biotinylated secondary antibodies and streptavidin-HRP complex (Cat# KIT-9710, MXB, Fuzhou, China). DAB substrate (Cat# G1212, Servicebio, Wuhan, China) was applied for color development. Sections were counterstained with hematoxylin, dehydrated, cleared, and mounted with neutral resin. DAB-positive signals appeared brown, while nuclei were stained blue.

Frozen sections were warmed to room temperature and treated with autofluorescence quencher (Cat# G1221, Servicebio, Wuhan, China) for 5 minutes, followed by washing. Sections were stained with dihydroethidium (DHE) solution (Cat# D7008, Sigma-Aldrich, Shanghai, China) at 37 °C for 40 minutes in the dark. After washing with PBS, sections were mounted with antifade medium (Cat# 1401, Servicebio, Wuhan, China). DHE-positive signals were visualized under a fluorescence microscope (DMi8, Leica, Germany) as red fluorescence (Ex/Em = 535/550 nm).

Quantification of fluorescence intensity and Western blot band density was performed using ImageJ software. The remaining image analyses, including histological evaluations were performed using Image-Pro Plus software (version 6.0, Media Cybernetics, Rockville, MD, USA). Statistical analyses were performed using GraphPad Prism (version 10.1.1, GraphPad Software, San Diego, CA, USA). Data were obtained from at least three independent experiments. Time-course comparisons were conducted using two-way analysis of variance (two-way ANOVA) with Šídák’s multiple comparisons test, while multi-group comparisons were evaluated using one-way analysis of variance (one-way ANOVA) followed by Tukey’s post hoc test. Pairwise comparisons were performed using Student’s t-test. For experiments involving the comparison of means across different genotypes, statistical analysis was performed using multiple unpaired t-tests. The False Discovery Rate (FDR) correction was applied using the two-stage, step-up method proposed by Benjamini, Krieger, and Yekutieli to ensure reliability in multiple comparisons. Results are presented as the mean $\pm$ standard deviation (SD), with statistical significance defined as p 0.05, or FDR-adjusted q 0.05. A complete list of abbreviations is provided in Supplementary Table 1.

This study explored the protective role of XLOC_015548 in promoting myogenic differentiation and mitigating muscle atrophy by regulating Gadd45g, activating the MEK/ERK signaling pathway, and alleviating oxidative stress (Fig. 1A). Confocal microscopy-based Fluorescence In Situ Hybridization (FISH) 3D imaging revealed that XLOC_015548 was predominantly localized in the cytoplasm. Stably transfected C2C12 cell lines were prepared using lentiviral constructs for XLOC_015548 knock-down (KD-XLOC) and its control (KD-NC), or XLOC_015548 overexpression (OE-XLOC) and its control (OE-Ctrl). Analysis with Co-localization Finder revealed negative Pearson correlation coefficients, further confirming the cytoplasmic localization of XLOC_015548. Moreover, Plot Profile analysis revealed there was no overlap between nuclear DAPI signals and FISH signals, thus confirming the cytoplasmic localization of XLOC_015548. Both 3D imaging and Plot Profile analysis showed that overexpression or knock-down of XLOC_015548 altered its expression level in cells (Fig. 1B, Supplementary Fig. 1A). Additionally, RT-qPCR analysis revealed that XLOC_015548 mRNA levels were decreased in the knock-down group, and increased in the overexpression group compared to their respective controls (Supplementary Fig. 1B). These results confirm successful transfection of the target gene into C2C12 myogenic cells and its correct localization in the cytoplasm (Fig. 1B, Supplementary Fig. 1B).

Fig. 1.

Mechanism of XLOC_015548 in promoting myogenic differentiation and alleviating muscle atrophy through its regulation of Gadd45g expression, activation of the MEK/ERK pathway, and mitigation of oxidative stress, along with analysis of its cellular localization and co-localization. (A) Schematic diagram illustrating the regulatory mechanism of XLOC_015548 in promoting myogenesis under denervation-induced or dexamethasone (DEX)-induced muscle atrophy conditions. Overexpression of XLOC_015548 inhibited mitochondrial dysfunction caused by reactive oxygen species (ROS), activated the MEK/ERK signaling pathway, and increased the expression of myogenesis-related molecules. Solid arrows indicate promotion or inhibition effects, while red and blue arrows represent upregulation and downregulation, respectively. (B) FISH images showing the localization and co-localization of XLOC_015548 in control cells (KD-NC) and XLOC_015548 in knockdown cells (KD-XLOC_015548). FISH signals are shown in red, and nuclei are labeled in blue. White arrows indicate the decreased XLOC_015548 expression in KD-XLOC_015548 cells. Pearson co-localization coefficients for three independent measurements were recorded in scatter plots, with merged fluorescent signals annotated in red, green, and blue. Line plots represent pixel intensity distributions for DAPI (blue fluorescence) and FISH (red fluorescence) signals. Scale bars: 20 µm (3D image), 40 µm (full image), and 15 µm (zoomed image).

To investigate the effects of XLOC_015548 on cell proliferation and survival, stable lentivirus-transfected C2C12 cells were analyzed on differentiation day 0 and day 7 under control conditions, or after 48 h treatment with the MEK/ERK inhibitor U0126. The EdU assay was used to quantify cell proliferation under various experimental conditions (Supplementary Fig. 2A). XLOC_015548 overexpression (OE-XLOC) significantly increased cell proliferation compared to the control group (OE-Ctrl), while knock-down (KD-XLOC) markedly suppressed proliferation (Supplementary Fig. 2B,C). Notably, the addition of U0126 completely abolished the pro-proliferative effects of OE-XLOC, indicating its dependence on the MEK/ERK pathway.

Calcein-AM/PI staining was used to evaluate cell survival under different conditions (Supplementary Fig. 2B,D; Supplementary Fig. 3A,B). Knockdown of XLOC_015548 showed a trend toward increased PI-positive (dead) cells, whereas OE-XLOC exhibited a trend toward reduced cell death, suggesting a potential protective role in undifferentiated states. This protective effect was partially reversed by U0126 treatment, further confirming its reliance on the MEK/ERK pathway. This phenomenon was not observed in differentiated cells.

Stable KD-XLOC and OE-XLOC cell lines were established to investigate the role of XLOC_015548 in C2C12 myogenic differentiation, with or without intervention from the MEK inhibitor U0126. Immunofluorescence (IF) and Western blot analyses were performed to evaluate the expression of related proteins and myotube formation during differentiation (Fig. 2A). On day 1, U0126 treatment significantly increased the proportion of Myosin heavy chain (MHC)-positive nuclei, as well as the myotube diameter (Fig. 2B,C). However, from day 3 onward, KD-XLOC showed significant inhibition of myotube formation and disorganization, whereas OE-XLOC promoted myotube formation and enhanced cytoplasmic MyoD distribution. These effects were evident on days 3, 5, and 7. Addition of U0126 reversed these effects, impairing myotube morphology in both the KD-XLOC and OE-XLOC groups (Fig. 2B). Further analysis on days 3, 5, and 7 revealed that KD-XLOC significantly reduced MHC-positive nuclei and myotube diameter, whereas OE-XLOC significantly increased these metrics. U0126 intervention suppressed the pro-differentiation effects of OE-XLOC, further demonstrating the dependence of XLOC on the MEK/ERK pathway (Fig. 2C). Western blot analysis corroborated these findings for day 7. KD-XLOC significantly reduced MHC expression and MEK activation, while OE-XLOC showed an promoting trend in these metrics. U0126 treatment suppressed OE-XLOC-induced pathway activation and protein expression. Furthermore, Gadd45g, a key regulatory protein, was negatively regulated by XLOC (Fig. 2D–F). These findings suggest that XLOC_015548 promotes myotube formation and regulates myogenic protein expression by modulating Gadd45g or activating the MEK/ERK signaling pathway, thus providing critical insights into the mechanisms underlying XLOC_015548-mediated myogenic differentiation.

Fig. 2.

Regulation of myotube formation and myogenic protein expression by XLOC via the MEK/ERK pathway. (A) Experimental timeline showing the differentiation of C2C12 myoblasts under conditions of XLOC knock-down (KD) or overexpression (OE), and with or without the MEK inhibitor U0126. (B) Representative immunofluorescence images at days 1, 3, 5, and 7 of differentiation. Green: MyoD; Red: MHC; Yellow: F-actin; Blue: nuclei (DAPI). Scale bar = 100 µm. (C) Quantification of the proportion of Myosin heavy chain (MHC)-positive nuclei and myotube diameter under different experimental conditions. Data are presented as the mean $\pm$ SD (n = 3). Statistical analyses: One-way ANOVA with Tukey’s post hoc test for day 1 (**p 0.01, *p 0.05, ns = not significant); Two-way ANOVA with Šídák’s multiple comparisons test for days 3, 5, and 7 (****p 0.0001, ***p 0.001, **p 0.01, *p 0.05, ns = not significant). Intra-group temporal comparisons were denoted by lowercase letters (a-g) indicating significant differences (*p 0.05) relative to Day 3 in the following groups: KD-NC, KD-NC+U0126, KD-XLOC, KD-XLOC+U0126, OE-Ctrl, OE-XLOC, and OE-XLOC+U0126. (D,E) Western blot analysis of MHC, p-MEK1/2, MEK1/2, p-ERK1/2, ERK1/2, and Gadd45g expression under KD and OE conditions. (F) Quantitative analysis of Western blot results normalized to $\beta$-Tubulin. Data are presented as the mean $\pm$ SD (n = 3). Statistical analysis: One-way ANOVA with Tukey’s post hoc test (***p 0.001, **p 0.01, *p 0.05, ns = not significant).

During the proliferation phase (DM 0d), XLOC_015548 increased cell proliferation primarily by activating the MEK/ERK pathway. KD-XLOC reduced the proportion of Ki67-positive nuclei, whereas OE-XLOC increased the proportion of Ki67-positive nuclei (Supplementary Fig. 4A–D). Treatment with U0126 reversed the pro-proliferative effects of OE-XLOC, confirming the critical role of the MEK/ERK pathway in XLOC_015548-mediated regulation of proliferation.

At the mid-differentiation phase (DM 3d), XLOC_015548 optimized the differentiation microenvironment by modulating the cell cycle and early apoptosis (Fig. 3A). Cell cycle analysis showed that OE-XLOC significantly increased the proportion of G1-phase cells while reducing the proportion of S-phase cells, suggesting that XLOC_015548 facilitates transition of C2C12 cells from the proliferation phase to the differentiation phase by regulating the cell cycle at the intracellular level (Fig. 3B,C). Additionally, Annexin V/PI staining revealed that OE-XLOC significantly increased the proportion of early apoptotic cells (Fig. 3D,E), indicating that it may clear undifferentiated cells, thereby creating a more favorable microenvironment for differentiation.

Fig. 3.

Regulation of myogenic differentiation by XLOC through activation of the MEK/ERK pathway and inhibition of Gadd45g nuclear translocation. (A) Experimental timeline for differentiation experiments under KD or OE conditions, with or without the MEK inhibitor U0126. (B) Flow cytometry analysis of cell cycle distribution on day 3 of differentiation, comparing the OE-Ctrl and OE-XLOC groups. (C) Quantification of G1, S, and G2 phase cell cycle distributions, comparing the OE-Ctrl and OE-XLOC groups. Statistical analysis: Student’s t-test (***p 0.001). (D) Annexin V/PI staining flow cytometry analysis, comparing apoptotic rates between the OE-Ctrl and OE-XLOC groups. (E) Quantification of total, early, and late apoptotic cells. Statistical analysis: Student’s t-test (***p 0.001, *p 0.05). (F) Representative immunofluorescence images on day 7 of differentiation under different experimental conditions. Green: Gadd45g; Yellow: F-actin; Red: MyoD; Blue: nuclei (DAPI). Scale bar = 100 µm. (G,H) Quantification of normalized fluorescence intensity for Gadd45g and p-MEK1/2. Data are presented as the mean $\pm$ SD (n = 3). Statistical analysis: One-way ANOVA with Tukey’s post hoc test (***p 0.001, ns = not significant). (I) Immunofluorescence images of p-MEK1/2 (green), F-actin (yellow), MyoD (red), and nuclei (blue, DAPI) on differentiation days 0 and 7. Scale bar = 100 µm. (J) Schematic diagram summarizing the potential mechanism by which XLOC regulates myogenesis through the MEK/ERK pathway by influencing Gadd45g nuclear translocation and myogenesis-related molecules.

During the late differentiation phase (DM 7d), XLOC_015548 maintained myotube formation by regulating the cytoplasmic localization of Gadd45g and the activity of the MEK/ERK pathway. IF analysis showed that KD-XLOC increased Gadd45g expression and nuclear translocation, leading to disorganized myotube alignment and impaired differentiation. In contrast, OE-XLOC reduced Gadd45g expression and nuclear localization while improving myotube structure (Fig. 3F,G). Nuclear-cytoplasmic fractionation combined with Western blot analysis demonstrated that XLOC_015548 bidirectionally regulates both protein stability and subcellular distribution of Gadd45g during myogenic differentiation (Fig. 2D–F, Supplementary Fig. 5A–C). Control experiments with empty vectors (OE-Control and KD-NC) showed no significant differences in total Gadd45g levels or nuclear-cytoplasmic partitioning compared to untransfected cells (Supplementary Fig. 5A), ruling out nonspecific vector effects. Specifically, XLOC_015548 overexpression (OE-XLOC) significantly reduced total Gadd45g levels (vs. OE-Ctrl), whereas knockdown (KD-XLOC) induced its accumulation (vs. KD-NC) (Fig. 2D–F). Subcellular localization analysis further revealed that OE-XLOC markedly decreased nuclear Gadd45g with a concomitant cytoplasmic reduction trend, while KD-XLOC enhanced nuclear enrichment alongside cytoplasmic elevation (Supplementary Fig. 5B,C). Notably, dynamic changes in nuclear-to-cytoplasmic ratios correlated strongly with functional outcomes: reduced ratios in OE-XLOC group coincided with MEK/ERK pathway activation and enhanced myotube fusion, whereas elevated ratios in KD-XLOC group were associated with differentiation blockade (Fig. 3F). These findings collectively indicate that XLOC_015548 orchestrates the pro-differentiation function of Gadd45g primarily through its dominant regulatory role in subcellular partitioning. Notably, these differences in Gadd45g distribution were not observed in undifferentiated cells (DM 0d), suggesting that Gadd45g localization plays a functional role primarily during differentiation (Supplementary Fig. 5D,E). Furthermore, p-MEK1 was predominantly localized in mature myotubes (Fig. 3I), while KD-XLOC decreased p-MEK1/2 expression (Fig. 3H). Although the positive nuclear rate for p-ERK1/2 was not significantly different in the KD-XLOC group, the co-localization of p-ERK1/2 with MyoD-positive nuclei in partially differentiated cells was reduced (Supplementary Fig. 5F,G). This indicates that XLOC_015548 optimizes myogenic differentiation by regulating the nuclear localization of p-ERK.

Histological analysis of major organs from gene-edited mice revealed no structural abnormalities (Supplementary Fig. 6A,B). However, conditional knockout mice (XLOC_015548fl/fl; Cre⁺) exhibited significantly elevated Gadd45g expression, predominantly in the nucleus. Additionally, the Pearson correlation coefficients for nuclear localization of p-ERK1/2 were reduced, indicating impaired ERK nuclear translocation (Supplementary Fig. 7E–G). After two weeks of denervation, Cre⁺ mice displayed disorganized muscle fiber structure, reduced muscle cross-sectional area (CSA), increased oxidative stress (Supplementary Fig. 7A–D), and significantly decreased MyoD expression (Supplementary Fig. 7E,G). These findings suggest that XLOC_015548 alleviates denervation-induced muscle atrophy by inhibiting the nuclear translocation of Gadd45g.

In summary, XLOC_015548 plays a crucial role in regulating cell proliferation, differentiation, and myotube formation during different stages of myogenesis. In the early phase (DM 0d), it enhances proliferation through the MEK/ERK pathway. In the mid-differentiation phase (DM 3d), it optimizes the differentiation microenvironment by regulating the cell cycle and early apoptosis. In the late phase (DM 7d), it maintains myotube formation by modulating the cytoplasmic localization of Gadd45g and the activity of the MEK/ERK pathway. These findings highlight the importance of cytoplasmic localization of Gadd45g and nuclear distribution of p-ERK in myogenic differentiation, with Gadd45g serving as a key upstream regulator of the MEK/ERK pathway in myogenesis and stress regulation (Fig. 3J).

To further investigate the regulatory role of Gadd45g in myogenic differentiation, we established C2C12 stable cell lines that overexpress Gadd45g (OE-Gadd45g) and then systematically analyzed the impact on the MEK/ERK signaling pathway, myotube formation, and functional development (Fig. 4B). Western blot analysis revealed that OE-Gadd45g significantly upregulated Gadd45g protein expression, activated the MEK/ERK signaling pathway, and increased the expression of the myogenic marker myosin heavy chain (MHC) (Fig. 4A,C). On the basis of OE-XLOC_015548 cells, further overexpression of Gadd45g (OE-Gadd45g) reversed the downregulation of Gadd45g caused by XLOC_015548 overexpression and enhanced p-MEK and MEK/ERK activation, as well as MHC expression (Supplementary Fig. 8A,B). Upon treatment with the MEK inhibitor U0126, these effects were suppressed, accompanied by a reduction in MHC expression (Fig. 4A,C; Supplementary Fig. 8A,B).

Fig. 4.

Gadd45g regulates myogenic differentiation and functionality of C2C12 myoblasts through the MEK/ERK pathway. (A) Western blot analysis of Gadd45g, p-MEK1/2, p-ERK1/2, and the myogenic differentiation marker MHC in OE-Ctrl, OE-Gadd45g, and OE-Gadd45g + U0126 groups. Overexpression of Gadd45g (OE-Gadd45g) significantly upregulated Gadd45g and MHC levels, while U0126 treatment reduced p-MEK/MEK, p-ERK/ERK, and MHC expression. (B) Experimental design outlining the infection of C2C12 cells with OE-Gadd45g or OE-Ctrl lentivirus and subsequent differentiation with or without U0126 treatment. (C) Quantification of Western blot results normalized to $\beta$-tubulin (**p 0.01, *p 0.05, ns = not significant). (D) Immunofluorescence images on day 7 of differentiation. Gadd45g (green) exhibited high expression in mature myotubes in the OE-Gadd45g group. p-MEK1/2 (green) showed no significant changes, whereas the nuclear localization and co-localization of p-ERK1/2 (green) with MyoD (red) were significantly increased (white arrows). U0126 treatment reduced nuclear localization of p-ERK1/2 and resulted in disordered myotube morphology (scale bar = 100 µm). (E) Immunofluorescence images on day 7 showing MHC (red), F-actin (yellow), and nuclei (blue, DAPI). OE-Gadd45g increased the differentiation signals, whereas U0126 treatment suppressed differentiation (scale bar = 100 µm). (F) Quantification of Gadd45g and p-MEK1/2 fluorescence intensity. OE-Gadd45g significantly upregulated Gadd45g (**p 0.01, ns = not significant), while U0126 did not affect Gadd45g expression. (G) Pearson co-localization coefficients for nuclear p-ERK1/2 and MyoD signals in undifferentiated cells. OE-Gadd45g showed increased co-localization (white arrows), which was reversed by U0126. A large number of p-ERK nuclear-localized undifferentiated cells were observed around mature myotubes, while MyoD typically translocates from the nucleus to the cytoplasm during differentiation. Co-localization analysis suggests that cells with co-localized p-ERK and MyoD may represent incompletely differentiated cells that are potentially cleared via the apoptotic pathway. (H) Quantification of multinucleated myotubes ($>$5 nuclei) revealed significant increases in OE-Gadd45g, with a reduction following U0126 treatment (**p 0.01). (I) Quantification of nuclear p-ERK1/2 positivity showed a significant increase in OE-Gadd45g, which was reversed by U0126 treatment (**p 0.01, *p 0.05).

Treatment with the MEK inhibitor U0126 resulted in decreased p-MEK/MEK and p-ERK/ERK ratios, and reduced MHC expression (Fig. 4C). These results highlight the critical role of the MEK/ERK signaling pathway in Gadd45g-mediated regulation of myogenic differentiation. IF analysis further confirmed the role of Gadd45g in myogenesis. On differentiation day 7 (DM 7d), OE-Gadd45g promoted the localization of Gadd45g to the cytoplasm and distribution in mature myotubes (Fig. 4D,F). Notably, although XLOC_015548 overexpression decreased total Gadd45g protein levels, its cytoplasmic localization was largely maintained. Similarly, OE-Gadd45g enhanced Gadd45g distribution in the cytoplasm and its enrichment in mature myotubes (Fig. 4D,F). These findings further support the notion that cytoplasmic Gadd45g helps activate MEK/ERK signaling and promotes the expression of the myogenic marker MHC. Although p-MEK levels remained unchanged (Fig. 4F), the nuclear localization of p-ERK1/2 and its co-localization with MyoD-marked nuclei (indicative of undifferentiated cells) were significantly increased in the OE-Gadd45g group. This effect was reversed by U0126 treatment (Fig. 4D,G,I). Together, these findings suggest that Gadd45g optimizes the myogenic differentiation environment by facilitating nuclear p-ERK signaling. In addition, the OE-Gadd45g group had an increased number of multinucleated myotubes ($>$5 nuclei), with a more orderly myotube alignment. In contrast, U0126 treatment inhibited myotube fusion, resulting in fewer, and more disorganized myotubes (Fig. 4E,H). Quantitative analysis of myotube width further supported these observations (Supplementary Fig. 8C,D).

Of note, in a DEX-induced muscle atrophy model, the key oxidative stress marker iNOS was suppressed under both OE-Gadd45g and OE-XLOC_015548 overexpression conditions (Supplementary Fig. 8A,B). This observation suggests that XLOC_015548 may play a significant role in alleviating DEX-induced oxidative stress. However, in the absence of DEX treatment, no significant differences in iNOS expression were observed.

In conclusion, this study revealed that cytoplasmic Gadd45g positively regulates myogenic differentiation, myotube formation, and modulates oxidative stress through activation of the MEK/ERK signaling pathway and nuclear translocation of p-ERK (Supplementary Fig. 8E). These findings provide critical insights into the mechanistic role of Gadd45g in myogenic regulation and highlight its potential as a target for enhancing muscle differentiation and function.

This study used XLOC_015548-stable transfected C2C12 cell lines (OE-Ctrl and OE-XLOC), combined with DEX treatment and MEK inhibitor U0126 intervention, to systematically investigate the role of XLOC_015548 in cell viability, senescence, proliferation, and myogenic differentiation (Fig. 5A). OE-XLOC significantly reversed DEX-induced inhibition of myotube formation and improved myotube morphology and function by activating the MEK/ERK signaling pathway, suppressing cellular senescence, and promoting cell proliferation. Brightfield microscopy revealed that DEX treatment markedly reduced the number and diameter of myotubes, while OE-XLOC restored these parameters. However, the restorative effects were partially inhibited by U0126 treatment (Fig. 5B,C).

Fig. 5.

XLOC_015548 reverses DEX-induced muscle atrophy by regulating myogenic differentiation. (A) Schematic showing the design of experiments involving OE-XLOC and control C2C12 stable lines treated with or without DEX, and with the MEK inhibitor U0126. (B) Top: Bright-field images of myotube morphology on differentiation day 0 (DM 0d) and day 7 (DM 7d) under different conditions (scale bar = 100 µm). Middle: $\beta$-galactosidase staining showing senescence levels (scale bar = 100 µm). Bottom: Representative immunofluorescence images of Ki67 (green), MyoD (red), MHC (red), and F-actin (yellow), with DAPI-stained nuclei (blue) (scale bar = 50 µm). (C) Quantification of myotube width, showing that DEX significantly reduced myotube diameter, which was reversed by OE-XLOC. U0126 partially inhibited this recovery effect. (D) Quantification of $\beta$-galactosidase-positive cells showing DEX-induced senescence. This was significantly reduced by OE-XLOC. U0126 treatment partially restored senescence levels. (E) Quantification of Ki67-positive nuclei showed that OE-XLOC enhanced proliferation, which was suppressed by U0126 treatment. (F) Western blot analysis showing protein levels for MHC, iNOS, p-MEK1/2, p-ERK1/2, and Gadd45g under different conditions. OE-XLOC downregulated Gadd45g expression, activated the MEK/ERK pathway, and increased MHC expression. These effects were reversed by U0126. (G) Quantitative analysis of Western blot results normalized to $\beta$-tubulin (n = 3). Statistical data are presented as the mean $\pm$ SD. Statistical analysis: One-way ANOVA with Tukey’s post hoc test (***p 0.001, **p 0.01, *p 0.05, ns = not significant). Scale bars = 50 µm or 100 µm.

$\beta$-Galactosidase ($\beta$-gal) staining confirmed that DEX significantly increased cellular senescence levels, while OE-XLOC substantially reduced the proportion of senescent cells. U0126 treatment diminished the anti-senescence effect of OE-XLOC (Fig. 5B,D). Additionally, staining for live/dead cells revealed improved cell viability with OE-XLOC (Supplementary Fig. 9A–C), while Ki67 IF and EdU assays revealed enhanced proliferation. These effects were partially reversed by U0126 (Fig. 5B,E; Supplementary Fig. 9B,D). Western blot analysis at day 7 of differentiation revealed that OE-XLOC downregulated Gadd45g expression, activated the MEK/ERK signaling pathway, and further enhanced MHC expression. Additionally, OE-XLOC significantly reduced iNOS expression, a key oxidative stress marker, suggesting its role in mitigating oxidative damage. However, U0126 treatment markedly suppressed MEK/ERK pathway activity and reduced MHC expression, and partially reversed the reduction of iNOS expression (Fig. 5F,G).

TUNEL staining and multiplex fluorescence analysis were used to further elucidate the role of Gadd45g in DEX-induced myogenic inhibition. DEX was shown to significantly increase the apoptosis rate on day 7 of differentiation, while OE-XLOC markedly reduced the proportion of TUNEL-positive cells and restored the proportions of MHC- and Myog-positive nuclei in myotubes (Supplementary Fig. 10A–G), thereby promoting myotube formation. These effects were partially reversed by U0126 treatment. Mechanistically, OE-XLOC may function by suppressing Gadd45g expression and enhancing its cytoplasmic localization, leading to the activation of MEK/ERK signaling and optimization of the differentiation environment (Supplementary Fig. 10H). Residual apoptosis observed on day 3 in the DEX + OE-XLOC group may reflect the natural process of selective apoptosis during myogenesis.

In conclusion, XLOC_015548 exerts significant protective effects against DEX-induced inhibition of myogenesis through multiple mechanisms, including the suppression of senescence, promotion of proliferation, regulation of Gadd45g localization, modulation of oxidative stress via iNOS downregulation, and activation of the MEK/ERK signaling pathway. This study provides new insights into the molecular mechanisms underlying DEX-induced muscle atrophy, and establishes a theoretical foundation for therapeutic interventions that target muscle wasting.

We next investigated the role of XLOC_015548 in regulating oxidative stress and mitochondrial function during myogenic differentiation by utilizing XLOC_015548-stably transfected C2C12 cell lines (OE-Ctrl and OE-XLOC) in combination with dexamethasone (DEX) treatment and intervention with the MEK inhibitor U0126 (Fig. 6A). Analysis of redox homeostasis, mitochondrial function, and cell viability revealed that DEX treatment significantly increased the green fluorescence intensity of the oxidative probe (BDP 581/591 C11), indicating an increase in oxidative stress. In contrast, OE-XLOC cells exhibited a marked reduction in oxidative signals and increased red fluorescence intensity of unreacted probe, which was partially reversed by U0126 treatment (Fig. 6B,D). These findings suggest that XLOC_015548 may regulate redox balance through activation of the MEK/ERK signaling pathway.

Fig. 6.

XLOC protects myogenic cells by regulating oxidative stress and mitochondrial function, reversing DEX-induced oxidative damage. (A) Experimental schematic for the oxidative stress and mitochondrial function assays in OE-XLOC and control C2C12 stable lines with or without dexamethasone (DEX) and MEK inhibitor U0126 treatments and at days 0, 3 and 7 of differentiation. (B) Oxidative-reductive fluorescent marker (BDP 581/591 C11) images showing the distribution of oxidized probe (green) and unreacted probe (red) within cells. DEX significantly increased oxidative signals, while OE-XLOC enhanced reductive signals. U0126 partially reversed the protective effects of OE-XLOC (scale bar = 50 µm). (C) Fluorescent images of DHE staining (red indicates reactive oxygen species), live/dead cell staining (Calcein-AM/PI; green for live cells, red for dead cells), and Rhodamine 123 (green indicates MMP). DEX significantly increased DHE signals, impaired MMP, and enhanced cell death. OE-XLOC reduced the DHE signal, partially restored Rhodamine 123 fluorescence, and improved viability. U0126 partially reversed these effects (scale bar = 50 µm). (D) Quantification of the redox fluorescence intensity ratios (green-to-red) shows DEX-induced oxidative signals. OE-XLOC reduced oxidative signals and enhanced reductive signals, which was partially counteracted by U0126. (E) Quantification of DHE-positive fluorescence intensity indicates that DEX increased oxidative stress levels, while OE-XLOC significantly reduced DHE signals. U0126 treatment partially reversed the antioxidant effects of OE-XLOC. (F) Quantification of Rhodamine 123 fluorescence intensity showed that DEX impaired mitochondrial function, while OE-XLOC improved the stability of MMP. U0126 partially inhibited the protective effects of OE-XLOC. Data are presented as the mean $\pm$ SD (n = 3). Statistical analysis was performed using one-way ANOVA with Tukey’s post hoc test (***p 0.001, **p 0.01, *p 0.05, ns = not significant). Scale bar = 50 µm. MMP, mitochondrial membrane potential.

Further analysis using DHE staining, Calcein-AM/PI double staining, and MMP (Rhodamine 123) assays allowed in-depth evaluation of the mechanisms by which XLOC_015548 modulates myogenic differentiation. DEX treatment was found to significantly increase DHE fluorescence intensity, impair MMP, and increase the cell death rate. In contrast, OE-XLOC cells showed a marked reduction in DHE-positive signals, partial recovery of MMP, and decreased cell death. However, the protective effects of OE-XLOC were attenuated by U0126 treatment, further confirming the critical role of the MEK/ERK signaling pathway in XLOC_015548-mediated regulation of oxidative stress and mitochondrial function (Fig. 6C,E,F; Supplementary Fig. 11A,B).

In summary, XLOC_015548 mitigates oxidative stress and maintains the stability of MMP, thus exerting significant protective effects during myogenic differentiation. It effectively counteracts DEX-induced oxidative damage, providing strong theoretical support for the therapeutic potential of XLOC_015548 in muscle atrophy-related diseases and highlighting its potential as a novel therapeutic target.

This study found that XLOC_015548 mitigates denervation-induced skeletal muscle atrophy through multiple mechanisms, including regulation of Gadd45g expression, enhancement of p-ERK nuclear localization, reduction of oxidative stress, and promotion of myogenic differentiation. A skeletal muscle-specific conditional overexpression mouse model (XLOC_015548cKI/+; iCre⁺) was observed to retain muscle mass and CSA compared to control mice (XLOC_015548cKI/+; iCre⁻) on day 14 post-sciatic nerve transection (Fig. 7A–D). IHC analysis confirmed that XLOC_015548 significantly downregulated Gadd45g expression under denervation conditions. Although the overall levels of p-MEK/MEK and p-ERK/ERK remained the same on day 14 post-denervation (Fig. 7E,G), p-ERK nuclear localization was enhanced in XLOC_015548cKI/+; iCre⁺ mice, suggesting a role in optimizing the myogenic environment during later stages of denervation (Fig. 7F). Additionally, XLOC_015548 overexpression upregulated the expression of the myogenic regulator MyoD in denervated skeletal muscle (Fig. 7E,G), further highlighting its critical role in promoting myogenic differentiation. Collectively, these findings indicate that XLOC_015548 can alleviate denervation-induced muscle atrophy through regulation of Gadd45g/ERK signaling, and promotion of myogenic differentiation.

Fig. 7.

XLOC_015548 overexpression alleviates denervation-induced muscle atrophy in conditional overexpression mice. (A) Representative images of gastrocnemius muscles from conditional XLOC_015548 overexpression mice (XLOC_015548cKI/+; iCre⁺ ) and control mice (XLOC_015548cKI/+; iCre⁻) under sham and denervation conditions (scale bar = 4 mm). (B) H&E staining highlights muscle fiber morphology and structural differences. Denervated mice in the Cre⁻ group exhibit fiber atrophy, whereas the Cre⁺ group demonstrate tighter fiber alignment and larger cross-sectional areas (CSA) (scale bar = 50 µm). (C) Schematic illustration of the potential mechanism for XLOC_015548 in mitigating denervation-induced muscle atrophy and promoting regeneration by regulating Gadd45g expression and MEK/ERK pathway activity. (D) Quantitative analyses of muscle mass (g) and average muscle fiber CSA (µm²) show that Cre⁺ mice exhibit recovery of gastrocnemius muscle mass and CSA under denervation conditions. Data are presented as the mean $\pm$ SD. Statistical analysis was performed using multiple unpaired t-tests and two-stage step-up corrections (*q 0.05, ns = not significant). (E) Immunohistochemistry (IHC) showing the expression of ERK, MEK, p-ERK1/2 (T202/T204), p-MEK1/2 (S218/S222), and Gadd45g under sham and denervation conditions (scale bar = 50 µm). (F) IHC analysis shows co-localization of p-ERK1/2 DAB-positive signals (red) with hematoxylin-stained nuclei (blue). Pearson correlation coefficients for co-localization are shown below the co-localization plots. H refers to hematoxylin, which stains cell nuclei blue, while DAB refers to 3,3^′-diaminobenzidine, which stains protein expression as a brown signal in IHC (scale bar = 50 µm). (G) Quantification of IHC results for MyoD, p-ERK/ERK, p-MEK/ERK, and Gadd45g protein expression. All results are normalized to the control group (XLOC_015548cKI/+; iCre⁻). Data are presented as the mean $\pm$ SD (n = 3). Statistical analysis was performed using multiple unpaired t-tests with two-stage step-up corrections (*q 0.05, ns = not significant).