Polydeoxyribonucleotide attenuates dexamethasone-induced skeletal muscle atrophy by suppressing oxidative stress and regulating muscle protein turnover

Abstract

Glucocorticoid-induced muscle atrophy is characterized by excessive oxidative stress and disrupted muscle protein homeostasis. Polydeoxyribonucleotide (PDRN) exerts tissue-protective and regenerative effects; however, its therapeutic potential in muscle atrophy remains unclear. This study investigated the protective effects of PDRN against dexamethasone (DEX)-induced muscle atrophy in vitro and in vivo. Differentiated C2C12 myotubes were treated with PDRN in the presence or absence of DEX (100 µM). PDRN showed no cytotoxicity, and concentrations of 10, 25, and 50 µg/mL were selected for subsequent experiments. PDRN significantly attenuated DEX-induced myotube thinning and reduced intracellular reactive oxygen species. For in vivo evaluation, C57BL/6 mice received DEX (25 mg/kg/day) for 14 days with or without intramuscular PDRN administration. PDRN treatment improved treadmill performance and preserved GCM mass compared with DEX-treated mice. Histological analyses demonstrated significant increases in muscle fiber area and minimum Feret diameter following PDRN treatment. Furthermore, PDRN reduced oxidative damage, as evidenced by decreased 4-hydroxynonenal staining and malondialdehyde levels while restoring glutathione content. PDRN also upregulated nuclear factor erythroid 2-related factor 2 signaling and normalized antioxidant-related gene expression. In addition, PDRN enhanced expression of anabolic markers, including insulin-like growth factor-1, myogenic differentiation 1, and myogenin while suppressing the atrophy-related factors muscle RING finger-1 and Atrogin-1. These changes were accompanied by increased MHC expression and activation of the mTOR/S6K pathway, together with suppression of the myostatin/Smad/FoxO3a signaling pathway. Collectively, PDRN attenuated DEX-induced skeletal muscle atrophy by suppressing oxidative stress and regulating muscle protein turnover, suggesting its potential as a therapeutic strategy for glucocorticoid-induced muscle wasting.

Introduction

Skeletal muscle atrophy is a debilitating condition characterized by the progressive loss of muscle mass, strength, and function. It is associated with aging, immobilization, chronic disease, and prolonged glucocorticoid therapy, leading to impaired physical performance and reduced quality of life (Yin et al. 2021). Among these factors, glucocorticoids are widely prescribed for inflammatory and autoimmune disorders; however, long-term use frequently results in muscle wasting and weakness (Gensler 2013). Therefore, the development of effective therapeutic strategies to prevent or attenuate glucocorticoid-induced muscle atrophy remains an important clinical challenge.

Dexamethasone (DEX), a synthetic glucocorticoid, is commonly used to establish experimental models of skeletal muscle atrophy (Chang and Kong 2020). DEX disrupts muscle protein homeostasis by suppressing anabolic pathways while activating proteolytic systems. In particular, DEX downregulates insulin-like growth factor-1 (IGF-1)/Akt/mechanistic target of rapamycin (mTOR) signaling and simultaneously promotes the expression of muscle-specific ubiquitin ligases, including muscle RING finger-1 (MuRF1) and muscle atrophy F-box (Atrogin-1), resulting in accelerated protein degradation and muscle loss (Stitt et al. 2004). Furthermore, glucocorticoid exposure induces excessive oxidative stress, which contributes to mitochondrial dysfunction, muscle fiber degeneration, and activation of catabolic signaling pathways (Chen et al. 2023).

Oxidative stress is increasingly recognized as a key contributor to the pathogenesis of skeletal muscle atrophy. Excessive production of reactive oxygen species (ROS) disrupts redox homeostasis, leading to lipid peroxidation, mitochondrial dysfunction, and activation of proteolytic pathways that ultimately accelerate muscle wasting (Agrawal et al. 2023). Accordingly, reducing oxidative stress and enhancing endogenous antioxidant defense systems have emerged as promising therapeutic strategies for preserving skeletal muscle integrity. Among these defense mechanisms, nuclear factor erythroid 2-related factor 2 (Nrf2) regulates the transcription of antioxidant enzymes, including superoxide dismutase (SOD) and catalase (CAT), thereby contributing to cellular protection against oxidative damage (Huang et al. 2023).

Polydeoxyribonucleotide (PDRN) is a mixture of deoxyribonucleotide polymers derived from salmonid DNA and is widely used for tissue repair and regeneration (Squadrito et al. 2017). PDRN exerts anti-inflammatory, antioxidant, angiogenic, and wound-healing effects, primarily through activation of adenosine A2A receptors (Squadrito et al. 2017, Irrera et al. 2018, Galeano et al. 2008). In addition, PDRN has shown protective effects in various models of tissue injury by reducing oxidative stress and promoting cellular recovery. Recently, PDRN has been increasingly utilized as a pharmacopuncture agent in Korean medicine. Emerging clinical evidence suggests that PDRN pharmacopuncture may be a safe and promising therapeutic option for various musculoskeletal disorders, including tendinopathy, osteoarthritis, ligament injury, plantar fasciitis, radiculopathy, and spinal disorders, by alleviating pain, improving functional recovery, and promoting tissue regeneration (Nam et al. 2025). Beyond musculoskeletal diseases, the clinical application of PDRN has expanded to dermatology and aesthetic medicine. PDRN has been reported to stimulate fibroblast proliferation, enhance collagen and elastin synthesis, restore the skin barrier, and accelerate wound healing. Moreover, a recent case report demonstrated that integrated treatment including PDRN pharmacopuncture successfully improved chronic recurrent dyshidrotic eczema, suggesting that PDRN pharmacopuncture may represent a safe and clinically applicable regenerative therapy beyond musculoskeletal indications (Lee and Lee 2026).

Despite these beneficial properties, the therapeutic potential of PDRN in glucocorticoid-induced skeletal muscle atrophy has not yet been fully elucidated. Therefore, the present study investigated the protective effects of PDRN in DEX-induced muscle atrophy using both differentiated C2C12 myotubes and a mouse model of glucocorticoid-induced muscle wasting. We further explored whether the effects of PDRN are associated with attenuation of oxidative stress, enhancement of endogenous antioxidant defense, and regulation of muscle protein synthesis and degradation pathways.

Materials and methods

PDRN preparation

PDRN was prepared at the Jaseng Hospital of Korean Medicine (Seoul, Republic of Korea). PDRN was supplied by GenoCure Co., Ltd. (Seoul, Republic of Korea) and used as the active pharmaceutical ingredient. The PDRN solution was formulated with sterile physiological saline and prepared under aseptic conditions. The formulated solution underwent sequential sterilization through a 0.2 μm membrane filtration system using compressed medical-grade nitrogen. Primary and secondary sterilizing filtration processes were performed to ensure sterility. The integrity of the sterilizing filters was verified before and after filtration using a filter integrity test. Following sterile filtration, the final product was aseptically filled into 10 mL sterile vials and stored under controlled conditions until use. This final formulation contained PDRN at a concentration of 1.875 mg/mL. All preparation procedures were conducted in accordance with Good Manufacturing Practice guidelines to ensure product quality, sterility, and consistency.

C2C12 myoblast culture and myotube differentiation

Mouse C2C12 myoblasts (ATCC, Manassas, VA, USA) were cultured in Dulbecco’s Modified Eagle Medium (DMEM; Gibco, Grand Island, NY, USA) supplemented with 10% fetal bovine serum (Gibco) and 1% penicillin–streptomycin (Gibco) at 37 °C in a humidified atmosphere containing 5% CO₂. Cells were seeded at 8 × 10⁴ cells/well in 24-well culture plates or 1.5 × 10⁴ cells/well in 96-well culture plates, according to the experimental purpose. On the day following seeding, when the cells had reached approximately 80–90% confluence, the growth medium was replaced with differentiation medium consisting of DMEM supplemented with 2% horse serum (HS; Gibco) and 1% penicillin–streptomycin. The differentiation medium was replaced every 2 days, and cells were differentiated for 4 days to form mature multinucleated myotubes. Fully differentiated myotubes were subsequently used for further analyses.

DEX and PDRN treatment

Differentiated C2C12 myotubes were treated with DEX and PDRN on day 4 of differentiation for 24 h and subsequently harvested or fixed according to the requirements of each analysis. For the cell viability assessment, differentiated myotubes cultured in 96-well plates were treated with DEX in the presence or absence of PDRN at concentrations of 1, 5, 10, 25, 50, 75, and 100 µg/mL. Based on the results of the cell viability assay, PDRN concentrations of 10, 25, and 50 µg/mL were selected for subsequent experiments. To evaluate the effects of PDRN on DEX-induced myotube atrophy and oxidative stress, C2C12 myotubes used for MyHC immunocytochemistry and acquisition of representative DCFDA fluorescence images were cultured directly in 24-well µ-plate (µ-Plate 24 Well ibiTreat; ibidi, Gräfelfing, Germany), without the use of separate coverslips. For MyHC immunocytochemistry, the cells were fixed and immunolabeled directly in the wells and subsequently imaged using confocal microscopy. Myotubes cultured in 96-well plates were used for quantitative analysis of DCFDA fluorescence.

Cell viability assay

Cell viability was evaluated using a Cell Counting Kit-8 assay (CCK-8; Dojindo, Kumamoto, Japan). Briefly, C2C12 myoblasts were seeded into 96-well plates and differentiated as described above. On day 4 of differentiation, the cells were co-treated with DEX and PDRN at concentrations of 1, 5, 10, 25, 50, 75, and 100 µg/mL for 24 h. Following treatment, CCK-8 reagent was added to each well at 10% of the culture medium volume and incubated for 4 h at 37 °C. Absorbance was measured at 450 nm using a microplate reader (Synergy H1 Hybrid Multi-Mode Reader, BioTek Instruments, Winooski, VT, USA). Cell viability was expressed as a percentage relative to the untreated control group using the following formula: Cell viability (%) = (OD of sample / Mean OD of control group) × 100.

MyHC immunocytochemistry and morphometric analysis

MyHC immunocytochemistry was performed 24 h after treatment. Differentiated C2C12 myotubes were fixed with 4% paraformaldehyde at room temperature and then washed with phosphate-buffered saline (PBS). The cells were permeabilized with 0.2% Triton X-100 and washed twice with PBS. To block nonspecific binding, the cells were incubated with 2% normal goat serum for 1 h at room temperature. The cells were then incubated overnight at 4 °C with an anti-MyHC primary antibody (1:200; Novus Biologicals, Centennial, CO, USA). Following primary antibody incubation, the cells were washed with PBS and incubated with an FITC-conjugated secondary antibody (1:300; Jackson ImmunoResearch, West Grove, PA, USA) for 2 h at room temperature in the dark. Nuclei were counterstained with 4′,6-diamidino-2-phenylindole (DAPI). Fluorescence images were acquired using a confocal microscope (Nikon, Tokyo, Japan). Quantitative analysis was performed using ImageJ software (National Institutes of Health, Bethesda, MD, USA). MyHC-positive cells containing two or more nuclei were defined as multinucleated myotubes. For myotube thickness analysis, 10 randomly selected images were acquired from each well at 100× magnification. The experiment was independently performed seven times on separate occasions using independently cultured and differentiated C2C12 cells. Within each independent experiment, measurements obtained from multiple wells and images were treated as technical measurements and averaged to obtain a single value for each treatment group. Each independently performed experiment was considered a biological replicate (n = 7), and quantitative data are presented as the mean ± SD of the seven independent experiments. The fusion index was calculated as the percentage of nuclei within MyHC-positive multinucleated myotubes relative to the total number of nuclei.

Fusion index (%) = (Number of nuclei within MyHC-positive multinucleated myotubes / Total number of nuclei) × 100.

DCFDA fluorescence assay

Intracellular reactive oxygen species (ROS) levels were assessed using a 2′,7′-dichlorofluorescin diacetate (DCFDA) assay kit (Abcam, Cambridge, UK). Cells were incubated with 10 µM DCFDA prepared in Live Cell Imaging Solution (LCIS; Gibco) at 37 °C for 30 min in the dark. After incubation, cells were washed twice with LCIS, and representative fluorescence images were captured using a confocal laser scanning microscope. For quantitative analysis, 100 µL of LCIS was added to each well, and fluorescence intensity was measured using an Epoch microplate reader (BioTek, Winooski, VT, USA) with excitation and emission wavelengths of 485 and 535 nm, respectively. DCFDA fluorescence intensity was normalized to cell viability.

DEX-induced muscle atrophy mouse model and PDRN treatment

A total of 60 eight-week-old male C57BL/6 mice maintained under specific pathogen-free conditions were used in this study (Samtako Bio, Osan, Republic of Korea). The animals were allocated to four experimental groups (n = 15 per group). All animal experiments were approved by the Jaseng Animal Care and Use Committee (Approval No. JSR-2024-12-001-A-003). Mice were randomly divided into four groups. The sham group received an intraperitoneal injection of physiological saline containing 20% sulfobutyl ether-β-cyclodextrin and 5% dimethyl sulfoxide (Sigma-Aldrich, St. Louis, MO, USA). The control group received DEX (Sigma-Aldrich) at a dose of 25 mg/kg by intraperitoneal injection. The PDRN-treated groups received DEX (25 mg/kg, intraperitoneal), along with PDRN into the gastrocnemius muscle (GCM). PDRN was administered at 50 µL per injection as either a two-fold diluted solution containing 0.047 mg PDRN (PDRN-L, low-dose PDRN) or the original solution containing 0.094 mg PDRN (PDRN-H, high-dose PDRN). DEX and PDRN were administered once daily for 14 consecutive days. Body weight was monitored every other day throughout the experimental period. At the experimental endpoint, mice designated for transcardial perfusion were deeply anesthetized with 2–3% isoflurane (JW Pharmaceutical, Gwacheon, Republic of Korea). After confirmation of an adequate depth of anesthesia, transcardial perfusion was performed as a terminal procedure with 0.9% saline followed by 4% paraformaldehyde (Biosesang, Yongin, Republic of Korea). The animals remained under deep anesthesia throughout the perfusion procedure. Following completion of perfusion, death was confirmed before tissue harvesting. The GCM was then harvested and weighed using an electronic balance for subsequent analyses.

Treadmill exhaustion test

The treadmill exhaustion test was performed as previously described with slight modifications. Before the exhaustion test, mice underwent a 3-day acclimation period on a motorized treadmill (Rodent Treadmill NG, Ugo Basile, Varese, Italy). During acclimation, mice ran at a speed of 10 m/min on a 0° incline for 5 min daily. The electric stimulus grid located at the rear of the treadmill was activated to encourage continuous running.

For the exhaustion test, the treadmill was set at a 10° incline with an initial running speed of 10 m/min. Running speed was increased by 2 m/min every 2 min until a maximum speed of 46 m/min was reached. Mice were placed individually in separate lanes, and the test was initiated by activating both the treadmill belt and the electric stimulus grid. Exhaustion was defined as the inability of a mouse to resume running despite repeated mechanical and electrical stimulation and remaining continuously on the shock grid for more than 10 s. Total running time and distance to exhaustion were recorded and used as indicators of exercise performance and fatigue resistance.

Histological and morphometric analysis of GCM

Following tissue collection, the GCMs were fixed in 4% paraformaldehyde (Biosesang, Yongin, Republic of Korea) at room temperature for 48 h. Fixed tissues were dehydrated through a graded ethanol series, cleared in xylene, and embedded in paraffin. Paraffin blocks were sectioned at a thickness of 5 μm using a HistoCore MULTICUT microtome (Leica, Wetzlar, Germany). Sections were floated on a 40 °C water bath, mounted onto coated slides, and dried at 37 °C.

For histological evaluation, tissue sections were stained with hematoxylin and eosin (H&E). Briefly, sections were stained with hematoxylin for 2 min 30 s, rinsed under running tap water for 2 min, and counterstained with eosin for 50 s. After staining, the slides were dehydrated through graded ethanol solutions and cleared in xylene, and coverslipped.

Histological images were acquired using a bright-field microscope (Nikon, Tokyo, Japan). For morphometric analysis, three non-overlapping fields from the central region of each transverse GCM section were randomly selected and imaged at ×400 magnification. Muscle fiber cross-sectional area (CSA) and minimum Feret diameter were quantified using ImageJ software after image scale calibration. Approximately 200 muscle fibers per group were analyzed. The mean CSA (µm²) and minimum Feret diameter (µm) were calculated for each animal and used for subsequent statistical analyses.

Immunohistochemistry

Paraffin-embedded GCM sections with a thickness of 5 μm were used for 4-hydroxynonenal (4-HNE) immunohistochemistry. Tissue sections were baked at 60 °C for 1 h, deparaffinized in xylene, and rehydrated through a graded ethanol series. After rinsing with distilled water and PBS, antigen retrieval was performed by heating the sections in 10 mM sodium citrate buffer (pH 6.0) at 95 °C for 20 min. The sections were then allowed to cool to room temperature in the same buffer and washed with PBS. The sections were permeabilized with 0.2% Triton X-100 for 10 min and blocked with 2% normal goat serum to reduce nonspecific binding. Subsequently, the sections were incubated overnight with an anti-4-HNE primary antibody (1:100; Abcam, Cambridge, UK) at 4 °C. After washing with PBS, the sections were incubated with an appropriate FITC-conjugated goat anti-mouse IgG (1:300; Jackson ImmunoResearch) at room temperature in the dark. Nuclei were counterstained with DAPI, and the slides were mounted using fluorescence mounting medium.

Fluorescent images were acquired using a confocal microscope (Eclipse C2 Plus, Nikon, Tokyo, Japan). Quantitative analysis of 4-HNE fluorescence intensity was performed using ImageJ software. The mean fluorescence intensity of 4-HNE-positive staining was measured in randomly selected fields from each section and used for statistical analysis.

Malondialdehyde and glutathione analysis

Malondialdehyde (MDA) and glutathione (GSH) levels were measured using commercially available colorimetric assay kits. MDA levels were determined using a Lipid Peroxidation (MDA) Assay Kit (Abcam), and total GSH levels were measured using a GSH/GSSG Assay Kit (Abcam). Briefly, GCM tissues were homogenized, and the resulting supernatants were used for analysis. All procedures were performed according to the manufacturers’ instructions, and absorbance was measured using a microplate reader (Synergy H1 Hybrid Multi-Mode Reader, BioTek Instruments).

Reverse transcription quantitative PCR analysis

Total RNA was isolated from GCM tissues using the RNeasy Mini Kit (Qiagen, Hilden, Germany) according to the manufacturer’s instructions. The extracted RNA was reverse-transcribed into complementary DNA (cDNA) using Oligo-dT primers and AccuPower RT PreMix (Bioneer, Daejeon, Republic of Korea). Reverse transcription quantitative PCR (RT-qPCR) was performed using the CFX Connect Real-Time PCR Detection System (Bio-Rad, Hercules, CA, USA). The expression levels of antioxidant-related genes, including nuclear factor erythroid 2-related factor 2 (Nrf2), superoxide dismutase 1 (Sod1), and catalase (Cat), as well as muscle growth- and atrophy-related genes, including insulin-like growth factor-1 (Igf1), myogenin (MyoG), myogenic differentiation 1 (MyoD1), muscle RING-finger protein-1 (Murf1), and atrogin-1 (Fbxo32), were evaluated. The primer sequences used in this study are listed in Table 1. Relative gene expression levels were normalized to glyceraldehyde-3-phosphate dehydrogenase (Gapdh) and calculated using the 2^-ΔΔCt method. The results are presented as fold changes relative to the sham group.

Table 1.

Primer sequences used for real-time PCR analysis

Gene	5’-3’	Primer sequence
Nrf2	Forward	GATCCGCCAGCTACTCCCAGGTTG
	Reverse	CAGGGCAAGCGACTCATGGTCATC
Sod1	Forward	GTACCAGTGCAGGACCTCAT
	Reverse	AGTCACATTGCCCAGGTCTC
Cat	Forward	CACTGACGAGATGGCACACT
	Reverse	TGTGGAGAATCGAACGGCAA
Igf1	Forward	ACCTCAGACAGGCATTGTGG
	Reverse	GTTTGTCGATAGGGACGGGG
Myog	Forward	GCAGGCTCAAGAAAGTGAATGA
	Reverse	TAGGCGCTCAATGTACTGGAT
Myod1	Forward	TGAATGAGGCCTTCGAGACG
	Reverse	GCCTGCAGACCTTCGATGTA
Murf1	Forward	AGGGCCATTGACTTTGGGAC
	Reverse	TTTACCCTCTGTGGTCACGC
Atrogin-1	Forward	ACCTGCCTGTGTGCTTACAA
	Reverse	CTCTTCCACAGTAGCCGGTC
Gapdh	Forward	ACTGAGCAAGAGAGGCCCTA
	Reverse	TATGGGGGTCTGGGATGGAA

Western blotting

GCM tissues were homogenized in radioimmunoprecipitation assay buffer (GenDEPOT, Barker, TX, USA) supplemented with protease and phosphatase inhibitor cocktails (Millipore, Burlington, VT, USA). The homogenates were centrifuged, and the supernatants were collected for protein analysis. Total protein concentrations were determined using a Pierce™ BCA Protein Assay Kit (Thermo Fisher Scientific, Waltham, MA, USA) according to the manufacturer’s instructions. Equal amounts of protein were separated by electrophoresis on 10–12% sodium dodecyl sulfate–polyacrylamide gels and subsequently transferred onto polyvinylidene difluoride membranes (Millipore). The membranes were blocked with 5% Difco™ Skim Milk (BD Biosciences, Franklin Lakes, NJ, USA) prepared in Tris-buffered saline containing 0.1% Tween-20 (TBST) and incubated overnight at 4 °C with the primary antibodies listed in Table 2. Following incubation, the membranes were washed with TBST and incubated with horseradish peroxidase-conjugated secondary antibodies for 2 h at room temperature. Protein bands were visualized using an enhanced chemiluminescence detection system (Bio-Rad, Hercules, CA, USA) and imaged using an Amersham Imager 600 (GE Healthcare Life Sciences, Uppsala, Sweden).

Table 2.

Primary antibodies used for western blot analysis

Antibody	Source	Cat.no	Dilution ratio
FOXO3a	CST	2497	1:1000
p-FOXO3a	Affinity	AF3020	1:1000
SMAD	CST	3103	1:1000
p-SMAD	Invitrogen	PA5-110155	1:1000
S6K	Abclonal	A1290	1:1000
p-S6K	Abclonal	AP0564	1:1000
mTOR	CST	2972	1:1000
p-mTOR	CST	2971	1:1000
MHC	DSHB	MF20	1:300
Myostatin	Proteintech	19142-1-AP	1:1000
GAPDH	Santa cruz	SC-32,233	1:2000

Band intensities were quantified using ImageJ software and normalized to the corresponding loading control. Relative protein expression levels were expressed as fold changes relative to the sham group.

Statistical analysis

All data are presented as the mean ± standard deviation (SD). Statistical analyses were performed using GraphPad Prism version 8.0 (GraphPad Software, Boston, MA, USA). Comparisons among multiple groups were conducted using one-way analysis of variance, followed by Tukey’s post hoc test for multiple comparisons. Differences were considered statistically significant at p < 0.05.

For graphical presentation, the symbol (#) indicates a significant difference compared to the sham group in the in vivo experiments or the blank group in the in vitro experiments, whereas the symbol (*) indicates a significant difference compared to the DEX-treated group. Statistical significance was denoted as follows: # or *, p < 0.05; ## or **, p < 0.01; ### or ***, p < 0.001; and #### or ****, p < 0.0001.

Results

PDRN attenuates DEX-induced myotube atrophy and oxidative stress in C2C12 myotubes

To determine the cytotoxicity of PDRN, differentiated C2C12 myotubes were treated with increasing concentrations of PDRN (1–100 µg/mL) for 24 h, and cell viability was assessed using a CCK-8 assay (Fig. 1A). PDRN treatment did not induce significant cytotoxicity at any tested concentration. Although cell viability gradually increased up to 50 µg/mL, it showed a slight decline at concentrations of 75 µg/mL and above; however, these changes were not statistically significant. Next, the protective effects of PDRN against DEX-induced myotube injury were evaluated (Fig. 1B). Treatment with DEX significantly reduced cell viability compared with that in the blank group. Co-treatment with PDRN did not significantly restore cell viability at any tested concentration up to 100 µg/mL. Based on the cytotoxicity and viability results, concentrations of 10, 25, and 50 µg/mL were selected for subsequent experiments to investigate the anti-atrophic effects of PDRN.

Fig. 1.

PDRN attenuates DEX-induced myotube atrophy and oxidative stress in differentiated C2C12 myotubes. (A) Cell viability of differentiated C2C12 myotubes treated with increasing concentrations of PDRN (1–100 µg/mL) for 24 h, as determined by the CCK-8 assay. (B) Cell viability of differentiated C2C12 myotubes co-treated with 100 µM dexamethasone (DEX) and PDRN (10–100 µg/mL) for 24 h. (C) Quantitative analysis of the fusion index. (D) Representative immunofluorescence images of MyHC (green) and DAPI (blue) staining following treatment with DEX (100 µM) in the presence or absence of PDRN (10, 25, and 50 µg/mL). White scale bar = 200 μm and yellow scale bar = 100 μm. (E) Quantitative analysis of myotube thickness. (F) Representative fluorescence images of intracellular reactive oxygen species (ROS) detected by DCFDA staining. White scale bar = 200 μm and yellow scale bar = 100 μm. (G) Quantification of DCFDA fluorescence intensity. Data are expressed as mean ± SD. Significant differences are indicated as ####p < 0.0001 vs. the blank group or *p < 0.01, **p < 0.01, ***p < 0.001, and ****p < 0.0001 vs. the DEX group according to one-way analysis of variance with Tukey’s post-hoc test

To evaluate myotube morphology, MyHC immunocytochemistry was performed following co-treatment with DEX and PDRN (Fig. 1C, D). DEX treatment markedly impaired myotube formation and significantly reduced the fusion index compared with that of the blank group. In contrast, PDRN treatment increased the fusion index in a concentration-dependent manner, indicating improved myogenic differentiation and myotube formation. Quantitative analysis of myotube thickness further demonstrated the protective effects of PDRN (Fig. 1E). DEX treatment significantly decreased myotube thickness, whereas co-treatment with PDRN significantly attenuated this reduction. The protective effect showed a concentration-dependent tendency, with the highest mean value observed in the 50 µg/mL PDRN-treated group.

To investigate whether the anti-atrophic effects of PDRN were associated with oxidative stress regulation, intracellular ROS production was evaluated using DCFDA staining (Fig. 1F, G). DEX treatment significantly increased DCFDA fluorescence intensity compared with that in the blank group, indicating enhanced ROS generation. PDRN co-treatment reduced ROS levels, with a decreasing trend observed from 25 µg/mL onward. Notably, only the 50 µg/mL PDRN group showed a statistically significant reduction in DCFDA fluorescence intensity compared with that of the DEX-treated group, suggesting that high-dose PDRN effectively suppresses DEX-induced oxidative stress in C2C12 myotubes.

PDRN improves physical performance and preserves muscle mass in DEX-treated mice

Body weight was measured every 2 days throughout the 14-day experimental period (Fig. 2A). Compared with the Sham group, the DEX group exhibited significantly lower body weights from Day 2 through Day 14. In contrast, both the low-dose PDRN (PDRN-L) and high-dose PDRN (PDRN-H) groups generally exhibited higher mean body weights than the DEX group, with significant differences observed from Day 4 onward, except on Day 8. Although the mean body weight tended to be higher in the PDRN-H group than in the PDRN-L group at several time points, no definitive dose-dependent effect of PDRN on body weight was established. To evaluate exercise capacity and endurance, a treadmill exhaustion test was performed after the 14-day treatment period (Fig. 2B, C). DEX administration significantly decreased both running distance and exhaustion time compared with those in the sham group, indicating impaired physical performance. In contrast, PDRN significantly increased both parameters relative to those in the DEX group. These findings suggest that PDRN alleviates DEX-induced impairment of exercise performance and endurance capacity.

Fig. 2.

PDRN improves physical performance and attenuates DEX-induced skeletal muscle atrophy in mice. (A) Changes in body weight during the 14-day experimental period. (B, C) Running distance and exhaustion time measured using a treadmill exhaustion test at the end of the experimental period. (D) Representative gross images of the gastrocnemius muscle (GCM). (E, F) Absolute GCM weight and GCM weight normalized to body weight (mg/g). (G) Representative hematoxylin and eosin (H&E)-stained cross-sections of the GCM. Black scale bar = 500 μm and red scale bar = 200 μm. (H) Quantitative analysis of muscle fiber cross-sectional area (CSA). (I) Quantitative analysis of the minimum Feret diameter. Data are presented as the mean ± SD. Significant differences are indicated as ####p < 0.0001 vs. the Sham group or *p < 0.01, **p < 0.01, ***p < 0.001, and ****p < 0.0001 vs. the DEX group according to one-way analysis of variance with Tukey’s post-hoc test

Representative images of the GCM collected at the end of the experimental period are shown in Fig. 2D. Visual inspection revealed a marked reduction in GCM size in the DEX-treated group compared with the sham group, whereas both PDRN-treated groups exhibited partial preservation of muscle mass. Consistent with the gross morphological findings, DEX administration significantly reduced GCM weight compared with that in the sham group (Fig. 2E). In contrast, PDRN significantly increased GCM weight relative to that in the DEX group. Similar protective effects were observed in both PDRN-treated groups, indicating that PDRN attenuated DEX-induced skeletal muscle loss. To account for differences in body weight, GCM mass was normalized to body weight and expressed as mg/g (Fig. 2F). DEX treatment significantly decreased the GCM mass-to-body weight ratio compared with that in the sham group. Both PDRN-treated groups exhibited significantly higher normalized GCM mass values than those in the DEX group, indicating that PDRN attenuated DEX-induced muscle loss in the GCM.

Histological analysis of H&E-stained GCM sections revealed marked muscle atrophy following DEX administration (Fig. 2G). Representative images from the central (mid-belly) region of the GCM showed reduced muscle fiber size in the DEX-treated group compared with the sham group. In contrast, PDRN treatment partially preserved muscle morphology, with the PDRN_H group exhibiting a histological appearance comparable to that of the sham group. Consistent with these observations, quantitative analysis demonstrated that DEX administration significantly reduced muscle fiber CSA compared with that in the sham group (Fig. 2H). Although PDRN_L did not significantly affect CSA, PDRN_H significantly increased muscle fiber area relative to the DEX group. Similarly, minimum Feret diameter was significantly decreased in the DEX group compared with the sham group (Fig. 2I). The PDRN_H group, but not the PDRN_L group, showed significantly restored minimum Feret diameter compared with that in the DEX group. Collectively, these findings demonstrate that PDRN mitigates DEX-induced skeletal muscle atrophy by preserving GCM mass and maintaining muscle fiber morphology, with more pronounced effects observed at the higher dose.

PDRN Attenuated Oxidative Stress and Restored Antioxidant Defense in DEX-Induced Skeletal Muscle Atrophy

Oxidative stress and antioxidant defense in the gastrocnemius muscle were assessed by immunohistochemical, biochemical, and gene expression analyses. Immunohistochemical analysis demonstrated that the expression of 4-HNE, a marker of lipid peroxidation, was significantly increased in the DEX group compared with the Sham group (Fig. 3A, B). Although no significant difference in 4-HNE expression was observed between the DEX and low-dose PDRN (PDRN-L) groups, treatment with high-dose PDRN (PDRN-H) significantly reduced 4-HNE expression compared with the DEX group, indicating an attenuation of oxidative damage. In contrast, the expression of SOD1, an endogenous antioxidant enzyme, was markedly decreased following DEX administration (Fig. 3C, D). Low-dose PDRN did not significantly alter SOD1 expression compared with the DEX group, whereas PDRN_H significantly restored SOD1 immunoreactivity.

Fig. 3.

PDRN attenuates oxidative stress and restores antioxidant defense in DEX-induced skeletal muscle atrophy. (A) Representative immunohistochemical images of 4-hydroxynonenal (4-HNE) staining in gastrocnemius muscle sections. White scale bar = 200 μm and yellow scale bar = 50 μm. (B) Quantitative analysis of 4-HNE-positive staining. (C) Representative immunohistochemical images of superoxide dismutase 1 (SOD1) staining in gastrocnemius muscle sections. White scale bar = 200 μm and yellow scale bar = 50 μm. (D) Quantitative analysis of SOD1-positive staining. (E) Glutathione (GSH) levels in gastrocnemius muscle. (F) Malondialdehyde (MDA) levels in gastrocnemius muscle. (G) Relative mRNA expression levels of antioxidant defense-related genes (Nrf2, Sod1, and Cat) determined by quantitative real-time PCR. Data are presented as the mean ± SD. Significant differences are indicated as #p < 0.05, ##p < 0.01, and ####p < 0.0001 vs. the Sham group or *p < 0.01, **p < 0.01, ***p < 0.001, and ****p < 0.0001 vs. the DEX group according to one-way analysis of variance with Tukey’s post-hoc test

Consistent with these findings, biochemical analysis revealed that DEX treatment significantly reduced GSH levels and increased MDA levels compared with the Sham group (Fig. 3E, F). High-dose PDRN significantly increased GSH levels while simultaneously reducing MDA accumulation compared with the DEX group, whereas low-dose PDRN did not produce significant changes. To further investigate the molecular antioxidant response, the mRNA expression levels of antioxidant-related genes were evaluated by real-time PCR (Fig. 3G). DEX administration significantly decreased the expression of Nrf2, Sod1, and Cat compared with the Sham group. PDRN treatment restored the expression of these antioxidant genes, with the high-dose group showing significant upregulation of Nrf2, Sod1, and Cat compared with the DEX group.

PDRN regulates muscle protein synthesis and degradation pathways in DEX-induced muscle atrophy

To investigate the molecular mechanisms underlying the anti-atrophic effects of PDRN, the expression of genes associated with muscle protein synthesis and degradation was evaluated in GCM tissues. DEX administration significantly reduced the mRNA expression levels of the anabolic factors Igf1, Myog, and Myod1 compared with those in the sham group (Fig. 4A). PDRN treatment partially restored the expression of these genes. Specifically, Igf1 expression was significantly increased only in the PDRN_H group, whereas both PDRN_L and PDRN_H groups showed significantly increased expression of MyoG and MyoD1 relative to that in the DEX group. Although both PDRN-treated groups exhibited significant improvements, the increases in Myog and Myod1 expression did not follow a clear dose-dependent pattern.

Fig. 4.

PDRN restores muscle protein homeostasis in DEX-induced skeletal muscle atrophy. (A) Relative mRNA expression levels of muscle protein synthesis-related genes (Igf1, Myog, and Myod1) in gastrocnemius muscle determined by quantitative real-time PCR. (B) Relative mRNA expression levels of muscle atrophy-related genes (Murf1 and Atrogin-1) determined by quantitative real-time PCR. (C) Representative western blot band images of total mTOR, phospho-mTOR (p-mTOR), total S6, phospho-S6 (p-S6), MHC, and GAPDH. (D) Quantitative analysis of phosphorylation levels of mTOR and S6K expressed as the p-mTOR/total mTOR and p-S6K/total S6K ratios, and MHC protein expression normalized to GAPDH. (E) Representative western blot band images of myostatin, total Smad2/3, phospho-Smad2/3 (p-Smad2/3), total FoxO3a, phospho-FoxO3a (p-FoxO3a), and GAPDH. (F) Quantitative analysis of Myostatin protein expression normalized to GAPDH, and phosphorylation levels of SMAD and FOXO3a expressed as the p-SMAD/total SMAD and p-FOXO3a/total FOXO3a ratios. Data are presented as the mean ± SD. Significant differences are indicated as #p < 0.05, ##p < 0.01, ##p < 0.001, and ####p < 0.0001 vs. the Sham group or *p < 0.01, **p < 0.01, ***p < 0.001, and ****p < 0.0001 vs. the DEX group according to one-way analysis of variance with Tukey’s post-hoc test

In contrast, DEX treatment significantly increased the expression of the muscle atrophy-related genes Murf1 and Atrogin-1 compared with those in the sham group (Fig. 4B). PDRN treatment significantly suppressed the expression of both genes. No significant difference in Murf1 expression was observed between the two PDRN-treated groups. Similarly, although Atrogin-1 expression tended to decrease as the PDRN concentration increased, the difference between the PDRN_L and PDRN_H groups did not reach statistical significance.

To further investigate the effects of PDRN on muscle protein homeostasis, protein expression levels associated with anabolic and catabolic signaling pathways were evaluated using western blot analysis (Fig. 4C–F). DEX administration significantly reduced MHC expression and decreased the phosphorylation levels of mTOR and S6, indicating suppression of anabolic signaling and protein synthesis (Fig. 4D). In contrast, high-dose PDRN significantly restored MHC expression and increased the phosphorylation levels of mTOR and S6 compared with those in the DEX group. In parallel, DEX treatment increased myostatin expression and enhanced Smad2/3 phosphorylation, indicating activation of catabolic signaling pathways. PDRN treatment attenuated these changes by reducing myostatin expression and suppressing Smad2/3 activation. Furthermore, PDRN increased FoxO3a phosphorylation compared with that in the DEX group.

Collectively, these results indicate that PDRN attenuates DEX-induced skeletal muscle atrophy by restoring anabolic signaling through the mTOR/S6 pathway while simultaneously suppressing myostatin/Smad2/3-mediated catabolic signaling and inhibiting FoxO3a-dependent muscle protein degradation.

Discussion

Glucocorticoid-induced muscle atrophy is a major complication associated with long-term corticosteroid therapy and is characterized by excessive oxidative stress, impaired protein synthesis, and accelerated protein degradation (Permpoon et al. 2025). To the best of our knowledge, we demonstrate for the first time that PDRN attenuates DEX-induced skeletal muscle atrophy in both differentiated C2C12 myotubes and a murine model of glucocorticoid-induced muscle wasting. Our findings indicate that PDRN mitigates glucocorticoid-induced muscle loss by suppressing oxidative stress and restoring the balance between anabolic and catabolic signaling pathways. Collectively, these results suggest that PDRN may represent a promising therapeutic strategy for preventing and treating glucocorticoid-induced muscle atrophy.

Oxidative stress is a key contributor to glucocorticoid-induced muscle atrophy. Excessive glucocorticoid exposure promotes mitochondrial dysfunction and ROS generation, leading to oxidative damage, activation of proteolytic pathways, and progressive muscle loss. Oxidative stress directly damages myofibers and amplifies catabolic signaling through FoxO-dependent pathways, thereby accelerating muscle protein degradation (Agrawal et al. 2023, Permpoon et al. 2025). In the present study, PDRN markedly attenuated oxidative stress in both in vitro and in vivo models, as evidenced by reduced ROS accumulation and decreased levels of lipid peroxidation markers. These findings are consistent with previous reports describing the antioxidant properties of PDRN in various tissues. Although the precise mechanism was not directly investigated in the current study, PDRN exerts its biological effects primarily through activation of adenosine A2A receptors. Activation of A2A receptor signaling enhances intracellular cAMP production and promotes antioxidant responses, including Nrf2-mediated transcriptional regulation. Consistent with this mechanism, PDRN treatment increased the expression of Nrf2 and its downstream antioxidant genes, suggesting that reinforcement of endogenous antioxidant defense systems may contribute to the observed protection against glucocorticoid-induced muscle damage (Mari et al. 2025).

In addition to its antioxidant effects, maintenance of muscle protein homeostasis appears to be another important mechanism underlying the protective actions of PDRN. Skeletal muscle mass is maintained through a dynamic balance between protein synthesis and protein degradation, and glucocorticoid-induced muscle atrophy disrupts this balance by suppressing anabolic signaling while activating catabolic pathways (Salehian and Kejriwal 1999). In the present study, PDRN treatment significantly increased the mRNA expression of the myogenic regulators Igf1, Myod1, and Myog, suggesting enhanced myogenic and anabolic potential. These transcriptional changes were accompanied by restoration of MHC expression and increased phosphorylation of mTOR and S6K, indicating recovery of protein synthesis-related signaling.

Conversely, PDRN suppressed multiple mediators of muscle protein degradation. The expression of the muscle atrophy-related genes Trim63 (MuRF1) and Fbxo32 (Atrogin-1) was significantly reduced, together with decreased myostatin expression and reduced phosphorylation of Smad2/3 and FoxO3a. Since myostatin signaling promotes muscle wasting through activation of Smad and FoxO transcription factors, leading to increased expression of muscle-specific ubiquitin ligases, the coordinated suppression of these pathways suggests that PDRN effectively attenuates glucocorticoid-induced proteolytic signaling. Collectively, these findings indicate that PDRN preserves skeletal muscle homeostasis by simultaneously enhancing protein synthesis and suppressing protein degradation, thereby mitigating DEX-induced muscle atrophy.

Although both PDRN doses produced favorable trends in several functional and molecular parameters, the therapeutic effects of the low-dose treatment were generally modest and inconsistent. In contrast, the high-dose group consistently demonstrated significant improvements across histological, biochemical, and oxidative stress-related outcomes. While several markers in the low-dose group showed changes in the same direction as those observed in the high-dose group, these changes frequently failed to reach statistical significance and were not consistently observed across all outcome measures, particularly those associated with oxidative stress. Therefore, the therapeutic efficacy of low-dose PDRN remains inconclusive under the present experimental conditions, whereas high-dose PDRN produced reproducible protective effects against DEX-induced muscle atrophy. Future studies examining additional dose levels and treatment durations are warranted to better define the therapeutic window and dose–response relationship of PDRN.

Some limitations should be considered when interpreting the present findings. First, although several signaling pathways were modulated by PDRN treatment, the present study cannot establish direct causal relationships between these molecular changes and the observed anti-atrophic effects. While our findings suggest the involvement of the Nrf2 pathway, adenosine A2A receptor signaling, and muscle protein turnover pathways, direct mechanistic validation was not performed. Future studies employing A2A receptor antagonists, Nrf2 inhibitors, or gene-silencing approaches are required to confirm the causal contribution of these pathways. Second, because PDRN was administered via intramuscular injection, the relative contribution of local versus systemic effects could not be determined. Third, the present study used only a glucocorticoid-induced muscle atrophy model. Additional investigations in aging-related sarcopenia, disuse atrophy, cancer cachexia, and neuromuscular disease models are needed to determine the broader applicability of PDRN. Finally, only a 14-day treatment period was evaluated, and the long-term efficacy and safety of repeated PDRN administration remain unclear. Moreover, muscle function was assessed using a treadmill exhaustion test, which reflects overall exercise performance rather than direct muscle contractile force. Future studies incorporating grip strength measurements, ex vivo muscle contractility testing, and comprehensive metabolic analyses would provide a more detailed understanding of the therapeutic effects of PDRN.

In conclusion, PDRN effectively attenuated DEX-induced skeletal muscle atrophy by reducing oxidative stress and preserving muscle protein homeostasis. These protective effects were accompanied by attenuation of oxidative damage, enhancement of endogenous antioxidant defenses, restoration of MHC expression and mTOR/S6K signaling, and suppression of myostatin/FoxO3-associated protein degradation pathways. Collectively, our findings suggest that PDRN represents a promising therapeutic candidate for the prevention and treatment of glucocorticoid-induced muscle atrophy.

Author contributions

Jin Young Hong: Formal analysis, Writing – original draft, Validation, Data Curation. Hyunseong Kim: Data curation, Writing - Review & Editing. Changhwan Yeo: Formal analysis, Visualization. Junseon Lee: Formal Analysis, Visualization. Wan-Jin Jeon: Formal analysis, Validation. Hyun Kim: Formal analysis, Software. In-Hyuk Ha: Formal analysis, Writing—review and editing. Seung Ho Baek: Writing—review and editing. Yoon Jae Lee: Conceptualization, Supervision, Project administration, Funding acquisition, Writing—review and editing.

Funding

This research was supported by a grant of the Korea Health Technology R&D Project through the Korea Health Industry Development Institute (KHIDI), funded by the Ministry of Health & Welfare, Republic of Korea (grant number: RS-2025-02310152).

Data availability

No datasets were generated or analysed during the current study.

Declarations

Consent for publication

Not applicable.

Ethics approval

All animal experiments were approved by the Institutional Animal Care and Use Committee (IACUC) of Jaseng Spine and Joint Research Institute (Approval No. R-2024-12-001-A-003) and were performed in accordance with the institutional guidelines for animal care and use.

Competing interests

The authors declare no competing interests.