The experimental protocol was approved by the Animal Ethics Committee of Jilin Agricultural University (Approval ID: 20220614005). Sixteen healthy, four-year-old male sika deer (Cervus nippon, average body weight: 108 $\pm$ 4.5 kg), with the same antler casting date (approximately 15 May 2023). The animals were randomly allocated into two experimental groups (n = 8 per group). All animals were fed a total mixed ration (TMR) consisting of 45% roughage and 55% concentrate on a dry matter basis (for detailed composition, see Supplementary Table 1). The control group received the basal diet, while the treatment group was given the same basal diet supplemented with 3.0 g/day of rumen-protected arginine. Each deer was housed individually and fed twice daily at 07:00 and 16:00, with free access to clean water. The experiment was conducted over an 8-week period following hard antler removal, which included a one-week dietary adaptation phase and a seven-week formal treatment phase. Antler mesenchymal tissue was subsequently collected. Antler mesenchymal tissue was then collected, with a portion immediately fixed for Immunofluorescence and the remaining tissue was placed in PBS for the isolation of RMCs. Animal health and welfare were monitored throughout the experiment by trained personnel under veterinary supervision. Appetite/feed intake, water intake, demeanor, locomotor activity, body condition, wound condition, bleeding, signs of infection, and pain-related behaviors were assessed daily, with particular attention during the perioperative period after hard antler removal and antler tissue collection. Predefined humane endpoints included severe body-weight loss ($>$15–20% of initial body weight), marked reduction in food intake for more than 24–48 h, inability to stand or move normally, persistent bleeding, severe wound infection, dehydration, or severe or unrelieved pain/distress. Animals reaching these criteria would be examined by a veterinarian and removed from the experiment for treatment or humane euthanasia if necessary. No animals reached the predefined humane endpoints, no severe adverse events or procedure-related mortality occurred, and no animals were euthanized in this study. Hard antler removal and subsequent antler mesenchymal tissue collection were performed by experienced personnel under aseptic conditions. Before the procedure, anesthesia and perioperative analgesia were provided with xylazine hydrochloride (working concentration: 100 mg/mL; 1.0–1.5 mg/kg, intramuscular [IM]) followed by ketamine hydrochloride (working concentration: 50 mg/mL; 2–4 mg/kg, IM). After clipping and aseptic preparation of the antler base, a tourniquet was applied around the pedicle to minimize blood loss. The antler was transected just above the pedicle coronet (approximately 2 cm above the pedicle) using a sterile fine-toothed saw, taking care to avoid pedicle injury. Hemostasis was confirmed before gradual release of the tourniquet, and the stump was treated with topical antiseptic and protected according to animal-welfare guidelines. Antler mesenchymal tissue was dissected under sterile conditions; part of the tissue was fixed for immunofluorescence, and the remaining tissue was placed in PBS for RMC isolation. Following sample collection, atipamezole hydrochloride (working concentration: 5 mg/mL; 0.1–0.2 mg/kg, IM) was administered to reverse the effects of xylazine, and animals were monitored until full recovery.

RMCs were isolated following an established protocol [20], and antler reserve mesenchymal layer (Fig. 1A) was minced and digested with 0.2% type II collagenase (093186, Gibco, Grand Island, NY, USA) at 37 °C for 2 h. After removing collagenase, digested complexes were cultured in growth medium consisting of Dulbecco’s Modified Eagle Medium (DMEM, 6125246, Gibco, Grand Island, NY, USA) supplemented with 10% fetal bovine serum (FBS, F8318, Sigma-Aldrich, St. Louis, MO, USA) and 1% penicillin-streptomycin (15140-122, Gibco, Grand Island, NY, USA) at 37 °C in a humidified incubator with 5% CO₂. Cells from passages 3–5 were used for all experiments. The identity of the primary cells was validated by flow cytometric analysis of surface markers. Mycoplasma contamination was tested using the Myco-Lum™ Luminescent Mycoplasma Detection Kit for Low Sensitivity Instrument (Beyotime, C0297S). The B/A ratio was 0.76, which is below the negative threshold of 0.9 according to the kit instructions; therefore, the cells were considered mycoplasma-negative.

Fig. 1.

Arginine promotes the proliferation of RMCs in vivo and in vitro. (A) Schematic diagram of the antler sampling site. (B) Sections were co-stained for the proliferation marker Ki67 (green) and the mesenchymal marker CD90 (red). Nuclei are counterstained with DAPI (blue). Scale bar, 100 µm. (C) Quantitative analysis of the percentage of Ki67-positive cells from images in (A,B). Data are presented as mean $\pm$ SEM (n = 3). *p 0.05 vs. CON group (Student’s t-test). (D) Immunofluorescence characterization of isolated and cultured RMCs, showing positive expression of mesenchymal stem cell surface markers CD44 and CD105. Nuclei are stained with DAPI (blue). Scale bar, 200 µm. (E) CCK-8 assay showing the viability of RMCs treated with 0 (CON), 200, 400, or 800 µM arginine for 48 h. **p 0.01, ****p 0.0001 compared to the CON group (one-way ANOVA with Tukey’s post-hoc test). (F) Representative fluorescence images of EdU (red) incorporation in RMCs treated with 0 (CON), 200, 400, or 800 µM arginine for 48 h. Nuclei are stained with DAPI (blue). Scale bar, 100 µm. (G) Quantitative analysis of the percentage of EdU-positive cells from images in (F). Data are presented as mean $\pm$ SEM (n = 3). *p 0.05, ***p 0.001 compared to the CON group (one-way ANOVA with Tukey’s post-hoc test). RMCs, reserve mesenchymal cells; DAPI, 4^′,6-diamidino-2-phenylindole; SEM, Standard Error of the Mean; EdU, EdU (5-Ethynyl-2^′-deoxyuridine) incorporation assay.

Antler tissue was fixed in 4% paraformaldehyde at 4 °C for 24 h, decalcified in decalcification solution at room temperature with solution changes every 3 days, then dehydrated through graded ethanol, cleared in xylene and embedded in paraffin. Sections (4 µm) were cut with a rotary microtome, mounted on poly-L-lysine–coated slides and dried. For immunofluorescence, sections were deparaffinized in xylene, rehydrated through descending ethanol to water, and subjected to antigen retrieval in 10 mM citrate buffer (pH 6.0) by microwave heating for about 10 min. After cooling, sections were washed with PBS, permeabilized with 0.3% Triton X-100 (P0096, Beyotime, Shanghai, China) and blocked with 10% normal goat serum for 30 min at room temperature. Slides were incubated with CD90 (27178-1-AP, dilution 1:500; Proteintech, Wuhan, China) and Ki67 (27309-1-AP, dilution 1:500; Proteintech, Wuhan, China) antibody at 4 °C overnight, washed, and then incubated with a fluorophore-conjugated secondary antibody (A0453, dilution 1:500; Beyotime, Shanghai, China) for 1 h in the dark. Nuclei were counterstained with DAPI (ZE0815, Vector Laboratories, Newark, CA, USA), and sections were mounted with anti-fade medium. Images were captured using a fluorescence microscope under identical settings, and fluorescence signals were analyzed with ImageJ software (Version 1.53t, National Institutes of Health, Bethesda, MD, USA). All animal samples were coded by a third party who was not involved in the histological quantification, and the histological quantification was performed in a blinded manner to avoid subjective bias.

RMCs were cultured in a 24-well cell culture dish for immunofluorescence. Cells were incubated with the indicated primary antibodies CD44 (5675-1-AP, dilution 1:500; Proteintech, Wuhan, China), CD105 (10862-1-AP, dilution 1:500; Proteintech, Wuhan, China) or $\beta$-catenin (P35222, dilution 1:1000; Cell Signaling Technology, Danvers, MA, USA) overnight at 4 °C. Samples were incubated with appropriate Alexa Fluor-conjugated secondary antibodies (A0453, dilution 1:500; Beyotime, Shanghai, China) and counterstained with DAPI. Images were captured using an inverted fluorescence microscope (Nikon, Tokyo, Japan). For quantification of $\beta$-catenin nuclear localization, 5 random fields of view were captured per group from three independent experiments. Nuclear $\beta$-catenin positive cells were defined as cells showing clear overlap between $\beta$-catenin fluorescence and the DAPI stained nucleus. The total number of cells and the number of nuclear $\beta$-catenin positive cells were counted in each field, and the percentage of nuclear $\beta$-catenin positive cells was calculated as nuclear positive cells divided by total cells multiplied by 100. Quantification was performed in a blinded manner using ImageJ software.

To examine the differentiation ability of RMCs, osteogenic and chondrogenic differentiation was induced in vitro. Briefly, cells were seeded at 5 $\times$ 10⁵ cells per well in 6-well plates, and differentiation was started after they attained 80% confluence. For osteogenic differentiation, cells were cultured in osteogenic induction medium. The medium included DMEM, 10% FBS, 1% P/S, 100 nM dexamethasone, 10 mM $\beta$-glycerophosphate, 50 µM ascorbic acid. For chondrogenic differentiation, cells were cultured in chondrogenic differentiation medium supplemented with DMEM, 5% FBS, 1% P/S, 10 ng/mL transforming growth factor-beta 1 (TGF-$\beta$1, R&D Systems, Minneapolis, MN, USA), 50 µM ascorbic acid, 1% insulin-transferrin-selenium (ITS, OriCell, Guangzhou, China) for 1 week. For adipogenic differentiation, cells were cultured in adipogenic induction medium supplemented with DMEM, 10% FBS, 1% P/S, 1 µM dexamethasone (D4902, Sigma-Aldrich, St. Louis, MO, USA), 200 µM indomethacin (I7378, Sigma-Aldrich, St. Louis, MO, USA), 10 µg/mL insulin (I6634, Sigma-Aldrich, St. Louis, MO, USA), and 0.5 mM 3-Isobutyl-1-methylxanthine (IBMX, I5879, Sigma-Aldrich, St. Louis, MO, USA) for two days, followed by incubation in adipogenic maintenance medium (DMEM, 10% FBS, 1% P/S, 10 µg/mL insulin) for one day. This cycle was repeated 7–8 times. Subsequently, staining was performed using alizarin red (C0138, Beyotime, Shanghai, China), Alcian blue (ALCB-10001, Cyagen, Suzhou, Jiangsu, China) and Oil red O (C0158S, Beyotime, Shanghai, China) solutions, respectively. Images were captured using a fluorescence microscope (Nikon Corporation, Tokyo, Japan).

Flow cytometry was performed to assess the expression of CD44 and CD105 in isolated RMCs. Cells were harvested, washed with PBS, and incubated with the corresponding primary antibodies or isotype controls, followed by incubation with the appropriate fluorescent secondary antibody. After washing, the cells were analyzed using a flow cytometer (BD Biosciences, San Jose, CA, USA), and the data were processed with FlowJo software (Version 10.8, BD Biosciences, Ashland, OR, USA). Positive gates were set according to the isotype control.

To assess the effect of Arg on RMCs proliferation, cells were treated with various L-Arg concentrations in an L-Arg-deficient basal medium (MA0545, Meilunbio, Dalian, Liaoning, China). L-Arg (A8094, Sigma-Aldrich, St. Louis, MO, USA) was dissolved in PBS to prepare a stock solution and then diluted into the Arg-deficient basal medium to the indicated final concentrations (0, 200, 400, and 800 µM). Control cells received the same volume of vehicle. RMCs were seeded in 96-well plates at a density of 3 $\times$ 10³ cells per well. Cell viability was evaluated at 48 h using the Cell Counting Kit-8 (CCK8, HY-K0301, MedChemExpress (MCE), Monmouth Junction, NJ, USA) according to the manufacturer’s instructions. Absorbance was measured at 450 nm with a microplate reader. Proliferating cells were labeled with the BeyoClick™ 5-Ethynyl-2^′-deoxyuridine (EdU) Cell Proliferation Kit with AF555 (C0076S, Beyotime, Shanghai, China). Briefly, RMCs treated with Arg for 48 h were incubated with 10 µM EdU for 2 h. After fixation and permeabilization, cells were stained with the Click iTRAQ reagent at room temperature in the dark for 30 min, then counterstained with Hoechst 33342 for nuclear staining. Images were acquired using a fluorescence microscope, and the percentage of EdU-positive cells was calculated from five randomly selected fields per well. EdU and CCK-8 assays were performed independently, using different batches of RMC cultures (3 independent batches), and each assay was repeated 3 times independently to ensure the reliability of the results. The cell seeding density and treatment conditions were consistent between the two assays.

For RNA Sequencing (RNA-seq), cells cultured under proliferative conditions were treated with 400 µM Arg, which showed the strongest pro-proliferative effect. Total RNA was extracted from control and 400 µM Arg-treated RMCs (n = 3 biological replicates per group) using TRIzol reagent (RE34142520, Invitrogen, Carlsbad, California, USA). RNA integrity was confirmed with an Agilent 2100 Bioanalyzer (RIN $>$8.0). Library construction and 150-bp paired-end sequencing were performed on an Illumina HiSeq 4000 by Novogene Co., Ltd. Raw reads were processed with fastp for quality control. Clean reads were aligned to the Sika deer reference genome (Cervus nippon, PRJCA001220) using HISAT2 (Version 2.2.1, Daehwan Kim, University of Texas Southwestern Medical Center, Dallas, TX, USA) [21]. Gene expression levels were quantified from the alignment files and used for differential expression analysis. Differential expression analysis was performed using the DESeq2 package in R (Version 1.53.0, Michael Love, Simon Anders, Wolfgang Huber, Bioconductor, Boston, MA, USA) [22], with genes satisfying $|$log₂FoldChange$|$ $>$0.5 and p-value 0.05 considered significantly differentially expressed. Principal component analysis (PCA) was employed to assess the overall transcriptional variation between the Arg and Con groups. Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway enrichment was performed using the clusterProfiler package (Version 4.21.0, Guangchuang Yu, Bioconductor, Boston, MA, USA) [23], applying an false discovery rate (FDR) threshold of 0.05 for significance.

Micromass cultures were used to assess Arg’s effect on RMCs chondrogenic differentiation [24]. Briefly, 1.5 $\times$ 10⁵ cells in 10 µL medium were seeded as micromass droplets per well. After 1 h of adherence at 37 °C, 500 µL of chondrogenic differentiation medium containing L-Arg was added. The groups received L-Arg at final concentrations of 0, 200, 400, or 800 µM. After 4 d, samples were fixed for Alcian blue staining or collected for gene and protein analyses. Since 800 µM Arg showed the strongest promotive effect on chondrogenic differentiation, this concentration was used in the subsequent pathway perturbation experiments with rapamycin (RAPA) and a Wnt/$\beta$-catenin signaling pathway activator (CHIR99021).

Total RNA was reverse transcribed using the PrimeScript RT reagent Kit (RR037A, Takara, Kusatsu, Shiga, Japan). Quantitative Real-Time PCR (qRT-PCR) was performed on a CFX96 Touch system (CFX96 Touch System, Bio-Rad, Hercules, CA, USA) using TB Green Premix Ex Taq II (RR820A, Takara, Kusatsu, Shiga, Japan). Gene expression was normalized to glyceraldehyde-3-phosphate dehydrogenase (GAPDH) and quantified using the 2^{-Δ⁢Δ⁢Ct} method. The primer sequences are listed as follows: SOX9 forward, 5^′-CAAGAACAAGCCGCACGTCAAG-3^′, and SOX9 reverse, 5^′-TCTCGCTCTCGTTCAGCAGTCT-3^′; COL2A1 forward, 5^′-GTGGAGCAGCAAGAGCAAGGA-3^′, and COL2A1 reverse, 5^′-AGCAGGCGGAGGAAGGTCAT-3^′; LEF1 forward, 5^′-CGGGTGGTGTTGGACAGATTAC-3^′, and LEF1 reverse, 5^′-GCGTTACGATGGCTGGATGAG-3^′; AXIN2 forward, 5^′-GGCGATCAGGACGGTGCTTA-3^′, and AXIN2 reverse, 5^′-GCTTGGAGACGATGCTGTTGTT-3^′; GAPDH forward, 5^′-AGATGGTGAAGGTCGGAGTG-3^′, and GAPDH reverse, 5^′-CCTTTCCATTGATGACGAGC-3^′.

Total protein was extracted using RIPA lysis buffer. Equal amounts of protein (20 µg) were separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), transferred to polyvinylidene difluoride (PVDF, IPVH00010, MilliporeSigma, Burlington, MA, USA) membranes, and blocked. Membranes were incubated overnight at 4 °C with primary antibodies. The following primary antibodies were used: rabbit COL2A1 (ab307674, dilution 1:1000; Abcam, UK), rabbit SOX9 (P48436, dilution 1:1000; Cell Signaling Technology, Danvers, MA, USA), rabbit mTOR (28273-1-AP, dilution 1:5000; Proteintech, Wu Han, China), rabbit p-mTOR (5536T, dilution 1:1000; Cell Signaling Technology, Danvers, MA, USA), rabbit p70 S6 Kinase (2708T, dilution 1:1000; Cell Signaling Technology, Danvers, MA, USA); rabbit p-p70 S6 Kinase (Thr389) (9234T, dilution 1:1000; Cell Signaling Technology, Danvers, MA, USA), mouse $\beta$-catenin (8480T, dilution 1:1000; Cell Signaling Technology, Danvers, MA, USA), mouse GAPDH Antibody (MAB374, dilution 1:20,000; Sigma-Aldrich, St. Louis, MO, USA), followed by incubation with anti-rabbit IgG horseradish peroxidase (HRP, P0948; dilution 1:500; Beyotime, Shanghai, China) or anti-mouse IgG HRP (P0946, dilution 1:500; Beyotime, Shanghai, China). Finally, blot signals were detected using a chemiluminescence kit (26619, Thermo Fisher Scientific, Altrincham, Cheshire, UK) and imaged with a multifunctional imaging system (Tanon, Shanghai, China). Relative protein levels were normalized to the GAPDH level on the same blot. Proteins were analyzed in grayscale using ImageJ software to obtain quantitative data.

All quantitative data are presented as mean $\pm$ standard error of the mean (SEM) from at least three independent experiments. Statistical comparisons among multiple groups were analyzed using one-way analysis of variance (ANOVA) followed by Tukey’s post hoc test in GraphPad Prism (Version 9.0, GraphPad Software, San Diego, CA, USA). p-value 0.05 was considered statistically significant.

The antler reserve mesenchymal layer in the antler growth center was selected for analysis (Fig. 1A). CD90 and Ki67 double immunofluorescence staining were performed in this region (Fig. 1B). CD90 (red) labels the mesenchymal stem cells, Ki67 (green) labels proliferating cells, and DAPI (blue) stains the nuclei. Compared with the control group, the Arg group showed a marked increase in the proportion of Ki67-positive cells among CD90-positive cells, with stronger green fluorescence signals overall. Quantitative analysis (Fig. 1C) revealed that Arg treatment significantly increased the proportion of Ki67⁺ cells (p 0.05). These results indicate that dietary Arg supplementation markedly enhances cell proliferation in the antler mesenchymal cell region.

The primary cultured cells displayed a typical spindle-shaped morphology (Supplementary Fig. 1A). Immunofluorescence (Fig. 1D) and flow cytometric analysis (Supplementary Fig. 1B) showed high expression of CD44 and CD105, and multilineage differentiation assays confirmed their osteogenic (Supplementary Fig. 1C), and chondrogenic (Supplementary Fig. 1D) and adipogenic (Supplementary Fig. 1E) differentiation potential. Together, these findings indicate that the isolated cells possess mesenchymal stem cell characteristics. The CCK-8 assay demonstrated that Arg treatment increased cell viability compared with the control group, with the 400 µM group exhibiting the strongest effect (Fig. 1E, p 0.05). EdU staining further showed that the percentage of EdU-positive cells was significantly higher in the Arg-treated groups than in the CON group, indicating that Arg promotes DNA synthesis in RMCs (Fig. 1F,G, p 0.05). Overall, these results suggest that Arg enhances the proliferation of RMCs.

To identify the molecular pathways by which Arg promotes RMCs proliferation, we performed RNA-seq analysis on cells from both the control group and the 400 µM Arg-treated group. PCA revealed a clear separation between the two groups, indicating distinct global transcriptional profiles, with PC1 and PC2 together accounting for 67.78% of the total variance (Fig. 2A). Differential expression analysis using $|$log₂FC$|$ $>$0.5 and p 0.05 identified 2907 differentially expressed genes (DEGs) after Arg treatment, including 1388 upregulated genes and 1519 downregulated genes. Among the upregulated DEGs, SST, UBE2C, PLK1, PCLAF, and FAM83D were markedly increased, while DES, TGFB1, CSPG4, and KIFC1 showed notable decreases among the downregulated DEGs (Fig. 2B). KEGG pathway enrichment analysis revealed the upregulated DEGs were significantly enriched in multiple canonical signaling pathways, including ECM-receptor interactions, focal adhesion, axon guidance, Hippo signaling pathways, mTOR signaling pathways, osteoclast differentiation, PI3K-Akt signaling pathways, and Wnt signaling pathways (Fig. 2C). Downregulated DEGs were mainly enriched for pathways related to DNA replication, cellular senescence, nucleocytoplasmic transport, the proteasome, oxidative phosphorylation, p53 signaling pathways, nucleotide metabolism, and the cell cycle (Fig. 2C). Next, to intuitively illustrate how DEGs are distributed across pathways and to explore potential inter-pathway connections, we built a gene-pathway conceptual network. Using pathways significantly enriched by upregulated genes and those already linked to antler growth and chondrogenesis, we visualized DEGs mapped to ECM–receptor interaction, focal adhesion, Wnt signaling, mTOR signaling, and osteoclast differentiation (Fig. 2D). The network topology showed that ECM–receptor interaction and focal adhesion occupied central hubs, connecting many DEGs and forming a closely linked cluster with Wnt signaling, mTOR signaling, and osteoclast differentiation, highlighting extracellular matrix-adhesion signaling as a core part of the Arg-induced transcriptional response. Within this cluster, numerous ECM/adhesion-related components (e.g., FN1, LAMA1/LAMA3, COL1A2/COL4A3/COL6A1, and various integrins ITGA/ITGB) were densely represented, supporting extensive remodeling of cell-matrix interactions following Arg treatment. Additionally, key Wnt receptors and ligands, like FZD1, FZD2, WNT5A, WNT10B, and WNT16, showed overall upregulation, indicating activation of Wnt signaling. In addition, several nutrient-sensing and translation-related nodes associated with mTOR signaling (e.g., CASTOR2, RRAGA, EIF4E, EIF4EBP1, EIF4B, SLC7A5/SLC3A2) displayed significant differential expression, pointing to mTOR-regulated processes involved in the Arg response. Overall, the gene-pathway network analysis indicates that Arg-driven transcriptional changes are organized around an ECM-adhesion hub and are linked with Wnt/mTOR growth-regulating pathways, providing systems-level support for Arg’s pro-proliferative effects in RMCs (Fig. 2D). Overall, the results suggest that Arg treatment significantly remodels the RMCs transcriptome. These changes are linked to the coordinated modulation of ECM/adhesion-related signaling and key growth-regulatory pathways, which may play a role in Arg-driven RMC proliferation and chondrogenic biological processes.

Fig. 2.

Transcriptomic profiling of RMCs in response to arginine treatment. (A) PCA plot showing distinct transcriptional profiles between the CON and Arg groups. (B) Volcano plot of differentially expressed genes (DEGs). Significantly upregulated (red) and downregulated (blue) genes are highlighted. (C) KEGG pathway enrichment analysis of upregulated and downregulated DEGs (based on $|$log₂FC$|$ $>$0.5 and p 0.05). (D) Gene-pathway interaction network. Nodes represent DEGs, colored by their expression change (red: up, blue: down), and are mapped to their significantly enriched KEGG pathways (grey circles). PCA, Principal component analysis; KEGG, Kyoto Encyclopedia of Genes and Genomes.

To validate the role of Arg in promoting commitment of RMCs to the chondrogenic lineage, an in vitro chondrogenic induction system was supplemented with different concentrations of Arg and cell differentiation was subsequently induced for 4 d. Alcian blue staining revealed that cells treated with 200 µM, 400 µM, or 800 µM Arg showed a significantly larger staining area compared to the CON group, indicating increased cartilage matrix deposition and a higher level of chondrogenic differentiation (Fig. 3A). Similarly, qPCR (Fig. 3B) and Western blot (Fig. 3C,D) analyses confirmed that Arg treatment markedly increased both the mRNA and protein levels of the key chondrogenic markers SOX9 and COL2A1. Overall, these findings demonstrate that Arg boosts the chondrogenic differentiation potential of RMCs.

Fig. 3.

Arginine enhances chondrogenic differentiation of RMCs. (A) Alcian Blue staining of micromass cultures after 4 d chondrogenic induction with the indicated concentrations of arginine, showing increased proteoglycan deposition (n = 3). Scale bar, 800 µm. (B) Relative mRNA expression levels of chondrogenic marker genes COL2A1 and SOX9 as determined by qRT-PCR. Data are presented as mean $\pm$ SEM (n = 3). *p 0.05; ****p 0.0001 compared to CON group (one-way ANOVA with Tukey’s test). (C,D) Western blot analysis of COL2A1 and SOX9 protein expression after arginine treatment (n = 3). **p 0.01; ****p 0.0001 compared to CON group (one-way ANOVA with Tukey’s test). GAPDH served as the loading control. qRT-PCR, Quantitative Real-Time PCR; COL2A1, Collagen type II alpha 1 chain; SOX9, SRY-box transcription factor 9; GAPDH, Glyceraldehyde-3-phosphate dehydrogenase.

To determine whether mTOR signaling contributes to Arg-induced chondrogenesis, we performed pharmacological perturbation experiments during chondrogenic induction. The phosphorylation levels of mTOR and its downstream target S6K were analyzed after Arg treatment. Western blot analysis showed that Arg increased the p-mTOR/mTOR and p-S6K/S6K ratios during chondrogenic differentiation, indicating enhanced mTOR signaling under Arg treatment (Fig. 4A,B).

Fig. 4.

Arg activates mTOR signaling during chondrogenic differentiation of RMCs. (A) Western blot analysis of p-mTOR, mTOR, p-S6K, and S6K in RMCs treated with different concentrations of Arg. (B) Quantification of the p-mTOR/mTOR and p-S6K/S6K ratios (n = 3). (C) Alcian blue staining of chondrogenic differentiation in the CON, Arg, Arg+RAPA, and RAPA groups (n = 3). Scale bar, 500 µm. (D) Relative mRNA expression of SOX9 and COL2A1 in the CON, Arg, Arg+RAPA, and RAPA groups (n = 3). Western blot analysis (E) and quantification (F) of p-S6K/S6K, SOX9, and COL2A1 protein expression in the CON, Arg, Arg+RAPA, and RAPA groups (n = 3). Data are presented as mean $\pm$ SEM. For panel (B), statistical significance was determined relative to the control group. For panels (D,F), ^# indicates comparison with the Arg group. *p 0.05, **p 0.01; ^#p 0.05, ^#⁢#p 0.01, ^#⁢#⁢#p 0.001.

To further assess the role of mTOR activity, RAPA (an inhibitor of mTOR signaling) was applied during chondrogenic induction. Alcian blue staining revealed that Arg promoted the deposition of cartilage matrix, whereas this effect was attenuated by RAPA (Fig. 4C). Consistent with this observation, the mRNA expression levels of SOX9 and COL2A1 were increased in the Arg group, but reduced in the Arg+RAPA group (Fig. 4D). Western blot analysis further showed that RAPA decreased the p-S6K/S6K ratio and attenuated Arg-induced upregulation of SOX9 and COL2A1 (Fig. 4E,F). Together, these results suggest that mTOR activity is involved in the Arg-enhanced chondrogenic differentiation of RMCs.

To examine Wnt/$\beta$-catenin signaling during chondrogenic differentiation, $\beta$-catenin immunofluorescence and gene expression analyses were performed after Arg treatment. Immunofluorescence staining showed that nuclear $\beta$-catenin fluorescence was reduced after Arg treatment and was lowest at a concentration of 800 µM (Fig. 5A and Supplementary Fig. 2A). Western blot analysis revealed the effect of Arg on total $\beta$-catenin expression was non-monotonic, with increased $\beta$-catenin expression at 200 and 400 µM, but decreased expression at 800 µM (Fig. 5B,C). In addition, the canonical Wnt target genes Axin2 and LEF1 were modulated by Arg treatment and showed lower expression at 800 µM (Fig. 5D). Together, these results suggest that canonical Wnt/$\beta$-catenin signaling was attenuated by 800 µM Arg during chondrogenic differentiation.

Fig. 5.

Effects of Arg and CHIR on the Wnt/$\beta$-catenin signaling during chondrogenic differentiation of RMCs. (A) Immunofluorescence staining of nuclear β-catenin in RMCs treated with different concentrations of Arg (n = 3). Scale bar, 50 µm. (B) Western blot analysis and quantification (C) of total β-catenin protein expression after Arg treatment (n = 3). (D) Relative mRNA expression of Axin2 and LEF1 in RMCs treated with different concentrations of Arg (n =3 ). (E) Alcian blue staining of chondrogenic differentiation in the CON, Arg, CHIR, and Arg+CHIR groups (n = 3). Scale bar, 500 µm. (F) Relative mRNA expression of SOX9 and COL2A1 in the CON, Arg, CHIR, and Arg+CHIR groups (n = 3). (G) Relative mRNA expression of Axin2 and LEF1 in the CON, Arg, CHIR, and Arg+CHIR groups (n = 3). (H) Immunofluorescence staining and quantification of nuclear $\beta$-catenin in the CON, Arg, CHIR, and Arg+CHIR groups (n = 3). Scale bar, 50 µm. Western blot analysis (I) and quantification (J) of $\beta$-catenin, SOX9, and COL2A1 protein expression in the CON, Arg, CHIR, and Arg+CHIR groups (n = 3). Data are presented as mean $\pm$ SEM. The white arrows in panels A and H indicate representative RMCs showing different levels of nuclear $\beta$-catenin staining. For panels (C,D), **p 0.01, ***p 0.001, ****p 0.0001 indicates comparison with the CON group. For panels (F–J), *p 0.05, **p 0.01, ***p 0.001 indicates comparison with the CON group. ^#p 0.05, ^#⁢#p 0.01 vs. the Arg group.

To further investigate the role of Wnt/$\beta$-catenin signaling in Arg-mediated chondrogenic differentiation, CHIR99021 (CHIR, a Wnt/$\beta$-catenin signaling pathway activator) was applied during chondrogenic induction. Alcian blue staining showed that 800 µM Arg promoted cartilage matrix deposition, with the effect attenuated by CHIR (Fig. 5E). Consistent with this finding, the mRNA expression levels of SOX9 and COL2A1 were increased in the Arg group but reduced in the Arg+CHIR group (Fig. 5F). Moreover, CHIR increased the expression levels of LEF1 and Axin2 (Fig. 5G), which showed a similar trend to nuclear $\beta$-catenin accumulation (Fig. 5H and Supplementary Fig. 2B). Western blot analysis further showed that CHIR increased $\beta$-catenin protein expression and reduced the Arg-induced upregulation of SOX9 and COL2A1 (Fig. 5I,J). Together, these results suggest that activation of Wnt/$\beta$-catenin signaling attenuates the promotive effect of Arg on the chondrogenic differentiation of RMCs.