Wild-type C57BL/6 male mice were purchased from (Orient-Bio, Seongnam, Korea). Fetal bovine serum (FBS), horse serum (HS), and Dulbecco modified Eagle's medium (DMEM) were purchased from Thermo Scientific (Waltham, MA, USA). 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT), DEX, and all other chemicals were from Sigma-Aldrich (St. Louis, MO, USA). Antibodies were purchased as follows: myosin heavy chain (MHC, Developmental Studies Hybridoma Bank (DSHB), Iowa, IA, USA) Myogenin, Myoglobin, total-OxPHOS (Abcam, Cambridge, MA, USA), total-Akt, phospho-Akt, phospho-mTOR, mTOR, phospho-S6K, S6K (Cell Signaling Technology, Beverly, MA, USA), MuRF1, Atrogin-1, HSP90 (Santa Cruz Biotechnology, Santa Cruz, CA, USA), and β-tubulin (Zymed, South San Francisco, CA, USA).

The fresh fruits of A. melanocarpa were collected at Danyang-gun, Chungcheongbuk-do, Korea, in August 2016 and were identified by one of the authors (K. H. Kim). A voucher specimen (SKKU AR-2016-08) has been deposited in the herbarium of the School of Pharmacy, Sungkyunkwan University, Suwon, Korea. Dried fruits (350 g) of A. melanocarpa were extracted with 80% MeOH for 3 days twice at room temperature. The extracts were then filtered, and the filtrate was concentrated under vacuum pressure, generating an AR (21.8 g). The AR was stored at −20°C until use.

All animal experiments were approved by the Institutional Animal Care and Research Advisory Committee at Sungkyunkwan University School of Medicine (SUSM) and complied with the regulations of the institutional ethics committee. All mice were maintained at 23°C with a 12:12 light-dark cycle and free access to food and water. To examine the effect of AR, these mice were orally administered a daily dose of 3.3 mg/kg AR for 8 weeks (8-month-old mice). Control mice were administrated the same amount of vehicle (dimethylsulfoxide, DMSO) dissolved in saline. All animals were sacrificed after fasting for 6 h with ad libitum to water.

C2C12 myoblasts were cultured as previously described (36). They were grown in Dulbecco's Modified Eagle Medium high glucose (DMEM; Thermo Scientific, Waltham, MA, USA) containing 15% FBS (growth medium, GM), 10 units/ml penicillin, and 10 μg/ml streptomycin (Welgene, Daegu, Korea) at 37°C, 5% CO₂. To induce differentiation of C2C12 myoblasts, cells at near confluence were changed growth medium into DMEM containing 2% HS (differentiation medium, DM), and myotube formation was observed at 2 or 3 days after differentiation. For the DEX-induced atrophy study, C2C12 cells were induced to differentiate in DM for 3 days (D3), followed by the treatment with 100 μM DEX along with vehicle DMSO or AR for an additional 1 day (D4) (37).

Western blot analysis was performed as previously described (38). Briefly, cells were lysed in cell extraction buffer (10 mM Tris-HCl, pH 8.0, 150 mM NaCl, 1 mM ethylenediaminetetraacetic acid (EDTA), and 1% Triton X-100) containing a complete protease inhibitor cocktail (Roche Diagnostics, Basel, Switzerland), followed by sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) and incubation with primary and secondary antibodies.

Immunostaining for MHC expression was carried out as previously described (39). Briefly, the differentiated cultures were then immunostained for MHC antibodies and Alexa 568-conjugated secondary antibodies (Molecular Probes, Eugene, OR, USA). Images were captured and processed with a Nikon ECLIPSE TE-2000U microscope and NIS-Elements F software (Nikon, Tokyo, Japan). To analyze the efficiency of myotube formation, the MHC-positive myotubes containing two to five, or six or more nuclei, were quantified at least three times and measured using ImageJ software.

Muscle tissue was embedded in Tissue-Tek OCT Compound (Sakura Finetek, Nagano, Japan), and 7 mm thick serial sections for staining were cut using a cryomicrotome. To analyze the nicotinamide adenine dinucleotide (NADH) dehydrogenase activity, we dried the sectioned tissues for 10 min at room temperature and incubated them in 0.9 mM NADH and 1.5 mM nitro blue tetrazolium (NBT; Sigma-Aldrich) in 3.5 mM phosphate buffer (pH 7.4) for 30 min. To analyze the succinate dehydrogenase (SDH) activity, we incubated the sections for 30 min in 50 mM sodium succinate and 0.3 mM nitro blue tetrazolium in 114 mM phosphate buffer containing K-EGTA (Sigma-Aldrich). To analyze the glycerol-3-phosphate dehydrogenase (GPDH) activity, we used the dried tissue samples and incubated them in 1.2 mM NBT, 2 mM phenazine methosulfate, and 1.86 mM glycerol phosphate in sodium phosphate buffer (pH 7.4) for 40 min.

Myh immunostaining of muscle tissue sections was performed in the sequence of fixation, permeation, and incubation with primary antibodies against MyhIIa and MyhIIb (DSHB) and laminin (Abcam). Images were captured with a Nikon ECLIPSE TE-2000U using NIS-Elements F software. Myofiber area was measured with ImageJ software. For muscle histology, the cryosections were stained with Mayer's hematoxylin and eosin (BBC Biomedical, McKinney, TX, USA). The images were captured using a Nikon ECLIPS TE-2000U.

Total RNA extraction and quantitative RT-PCR analysis were performed as previously described (40). Tissues were homogenized by FastPrepR-24 (MP Biomedicals, Santa Ana, CA, USA) and extracted using the easy-spin Total RNA Extract kit (iNtRON, Seongnam, Korea). Gene expression fold change was normalized against the expression of 18S ribosomal RNA. The sequences of the primers used in this study are provided in Table 1.

Grip strength was measured using a grip strength meter (Bioseb, Pinellar Park, FL, USA). The animal was allowed to grab the grid with a fore and back limb through a blind test. The limbs are pulled gently with consistent force until the forelimb was detached from the grid. The maximal strength was recorded when the grid was detached. Each animal was tested in three trials.

Values are expressed as mean ± SD for in vitro systems or ± SEM for in vivo systems, as indicated in the figure legends. The statistical significance was calculated using either Student's t-test (unpaired, two-tailed) or ANOVA by post-hoc Tukey's tests for multiple comparisons. Differences were considered statistically significant at or under values of P < 0.05.

To examine the effect of AR on myogenic differentiation, C2C12 myoblasts were induced to differentiate for 3 days (D3) in the presence of vehicle DMSO or AR with the indicated concentration ranging from 0.1 to 1 μg/ml in DM and subjected to the assessment of myoblast differentiation by immunoblotting analysis. The treatment with AR increased the expression levels of MHC and Myogenin in a dose-sensitive fashion and peaked at the concentration between 0.2 and 0.4 μg/ml (Figure 1A). This slight decrease at higher AR concentration was not due to any cytotoxicity as assessed by MTT assay (Figure 1B). To explore the molecular regulatory pathways of AR-mediated myogenic promotion, C2C12 myoblasts were treated with AR at the concentration of 0.1, 0.3, and 1.0 μg/ml for 48 h. Cell lysates were subjected to immunoblotting analysis for the active phosphorylated Akt (pAkt) to assess for the activation status of a promyogenic kinase Akt. The treatment of AR, especially 0.3 μg/ml concentration, increased the level of pAkt without altering the total Akt protein level (Figures 1C,D). In addition, AR treatment induced the phosphorylation of mTOR, a downstream target of Akt, in a dose-dependent manner (Figures 1C,D). These results indicate that AR promotes myogenic differentiation with enhanced Akt activation.

To confirm the promyogenic effects of AR, myotube formation was assessed by MHC immunostaining. As a result, AR treatment in C2C12 myoblasts elicited the formation of larger MHC-positive multinucleated myotubes, compared to control cells (Figure 1E). To quantify the myotube formation, MHC-positive myocytes were counted as mononucleate, myotubes containing two to five nuclei or containing six or more nuclei and plotted as a percentile (Figure 1F). The treatment with AR decreased the proportion of mononucleate myocytes, while it substantially elevated the proportion of larger myotubes containing six or more nuclei in a dose-sensitive manner. Our current data further support the positive effect of AR on myoblast differentiation at a morphological and at a biochemical level. AR can exert its promyogenic effect through Akt activation without overt cytotoxicity.

The synthetic glucocorticoid DEX induces muscle-specific ubiquitin ligase expression contributing to muscle atrophic phenotypes with reduced myotube diameter (37). Akt activation is one of the key events to suppress the induction of muscle-specific ubiquitin ligase and muscle atrophy triggered by DEX (25). Since AR can activate Akt, the effects of AR at the concentration of 0.1 or 1.0 μg/ml on DEX-induced C2C12 myotube atrophy were examined. C2C12 cells were induced to differentiate in DM for 3 days (D3), followed by the treatment with DEX along with vehicle DMSO or AR for an additional 1 day (D4) (Figure 2A). Myotubes were then subjected to MHC immunostaining to access the thickness of myotubes. The treatment with DEX-elicited myotube atrophy, which was suppressed by AR treatment, was evident by the presence of larger multinucleated myotubes in AR-treated cultures (Figure 2B). The quantification of myotube diameter revealed that DEX treatment caused declines in myotube diameter, which was significantly recovered by AR treatment (Figure 2C). In addition, the qRT-PCR analysis showed that DEX treatment greatly elevated muscle-specific E3 ubiquitin ligases, MuRF1, and Atrogin-1 and AR treatment attenuated this induction (Figure 2D). To further define, the protein levels of muscle-specific proteins and ubiquitin ligases were determined by immunoblotting analysis. Consistent with qRT-PCR results, the treatment with AR in DEX-treated myotubes reduced the level of MuRF1 and Atrogin-1 proteins, compared to control (Figures 2E,F). The treatment of DEX elicited a reduction in MHC protein levels and the co-treatment of DEX with AR attenuated the decline in MHC proteins in myotubes. In addition, DEX-treated myotubes exhibited decreased levels of pAkt while AR treatment in DEX-treated myotubes abrogated this decrease. Taken together, these results suggest that AR protects DEX-induced myotube atrophy through inhibition of muscle-specific ubiquitin ligases mediated by Akt activation.

To assess the in vivo effects of AR on muscles, 8-month-old-mice were fed with control or AR for 8 weeks, followed by muscle functional analysis and biopsy. The hindlimb muscles of AR-ingested mice appeared darker than those of control-fed mice, which can be readily detected in gastrocnemius (GAS) muscles among five hindlimb muscle groups, such as soleus (SOL), tibialis anterior (TA), extensor digitorum longus (EDL), and quadriceps (QU) (Figure 3A). AR treatment mildly but significantly elevated body weights, compared with the vehicle-treated group (Figure 3B). Among four hindlimb muscles, the weights of SOL and EDL muscles were increased an approximately 20.0 and 15.3% in AR-fed mice, compared to vehicle-fed mice, respectively (Figure 3C). The heart mass was slightly reduced but the liver mass was not changed (Figure 3D). One week prior to harvesting muscles, muscle strength was examined by measuring the grip strength. Mice fed with AR displayed approximately 14.5% increase in grip strength relative to control mice (Figure 3E). These data collectively suggest that AR treatment improves muscle mass and function without overt hypertrophic effect on the cardiac muscle.

To examine the detailed histology of muscles, control and AR-treated TA muscles were cryosectioned and subjected to hematoxylin and eosin staining, followed by the measurement of the cross-sectional area of myofibers (Figures 4A,B). AR-treated muscles had more lager myofibers, compared to the control-treatment muscles. To examine the effect of AR on myofiber types, muscle sections were analyzed by immunostaining with antibodies to MyhIIb and MyhIIa (Figure 4C). AR treatment elicited a shift toward larger MyhIIa- and MyhIIb-positive myofibers, compared to the vehicle treatment (Figures 4D,E). Consistently, qRT-PCR analysis of AR-treated muscles had elevated expression of MyhIIa and MyhIIb, compared to the control muscles (Figure 4F). These data suggest that AR treatment enhances MyhIIa and MyhIIb muscles.

Myofibers have different metabolic characteristics. Myofiber type IIa has higher mitochondrial content, compared to glycolytic myofiber type IIb (41, 42). Since AR substantially elevated both MyhIIa and MyhIIb expressions, we next examined the muscle metabolism by measuring activities of SDH and nicotinamide adenine dinucleotide tetrazolium reductase (NADH-TR) for muscle oxidative metabolism and GPDH for glycolytic metabolism. In agreement with the myofiber type staining, AR treatment elevated the proportion of myofibers with strong (dark) activities for SDH, NADH-TR, and GPDH, compared to the control muscles (Figures 5A,B), indicating that AR enhances both oxidative and glycolytic muscle metabolism. Consistently, AR-treated muscles expressed higher levels of mitochondrial genes (MTCO1, Sdhb), myoglobin, and glucose metabolism-related genes [hexokinase (HK), pyruvate kinase (PK), phosphofructokinase (PFK)], compared to the control muscles (Figures 5C,D). To further confirm, we examined the protein levels of total OXPHOS [ATP5A (CV), MTCO1 (CIII), SDHB (CII) and NDUF88 (CI)] proteins, myoglobin and S6K in Qu muscle tissues. In consistent with the RNA expression data, AR-treated muscles had elevated levels of total OXPHOS, myoglobin, and phosphorylated S6K proteins (Figures 5E,F). These data suggest that AR enhances the expression of muscle-specific genes and muscle metabolic genes, such as mitochondrial components.