Oxysophoridine promotes osteoarthritis repair via GSH system activation and ROS suppression

Abstract

Osteoarthritis (OA) is a prevalent joint degenerative disease involving inflammation and oxidative stress, with reactive oxygen species (ROS) driving progression. Restoring joint redox balance mitigates cartilage damage. Osilyhizidine (OSR), from Sophora alopecuroides, has anti-inflammatory/antioxidant properties, but its OA-specific effects and mechanisms were unclear. In vitro experiments assessed OSR’s impact on OA chondrocyte proliferation, repair, and inflammation, focusing on Glutathione peroxidase 4 (GPX4) and solute carrier family 7 member 11 (SLC7A11) regulation. A murine OA model validated findings in vivo. OSR showed anti-inflammatory, antioxidant effects and promoted cartilage repair, enhancing chondrocyte functions under inflammation and suppressing pro-inflammation. It upregulated GPX4 (improving ROS detoxification) and SLC7A11 (facilitating glutathione synthesis for redox balance) at transcriptional and protein levels. These were confirmed in mice. OSR alleviates OA by activating GPX4/SLC7A11 to regulate ROS and oxidative stress, emerging as a promising OA therapeutic candidate, offering insights into redox-targeted interventions.

Supplementary Information

The online version contains supplementary material available at 10.1038/s41598-026-37912-1.

Introduction

Osteoarthritis (OA) develops as a degenerative musculoskeletal disorder in which articular cartilage gradually deteriorates while abnormal ossification occurs within the joint structures^1–3. Osteoarthritis represents the leading form of joint pathology across the globe, predominantly occurring in the ageing population⁴. World Health Organization (WHO) estimates indicate that among individuals older than 60 years, osteoarthritis prevalence reaches approximately 10% in men and 18% in women. Driven by demographic aging and the expanding prevalence of obesity, the burden of osteoarthritis has shown a continuous upward trend, emerging as an important issue within global public health⁵. Currently, osteoarthritis treatment primarily focuses on symptom relief and slowing disease progression, as no cure exists. Current therapeutic approaches to osteoarthritis integrate medication-based regimens (notably nonsteroidal anti-inflammatory agents), physiotherapy, lifestyle optimization, and various surgical techniques. However, these methods have limitations: pharmacological treatments mainly target pain relief with limited therapeutic efficacy; physical therapy and lifestyle changes have modest effects and are unlikely to reverse the disease; and surgical treatments carry higher risks, are expensive, and require lengthy recovery periods⁶. In summary, osteoarthritis is a complex and widespread chronic disease, highlighting the urgent need to develop novel, safe, and highly effective therapeutic drugs⁷.

Persistent inflammation is a key driver of articular cartilage degeneration and a hallmark of OA⁸. Among inflammatory cytokines, IL-1β (Interleukin-1β) is one of the most extensively studied in OA.Exposure of chondrocytes to IL‑1β results in pronounced up‑regulation of matrix metalloproteinases—particularly MMP‑1, MMP‑3, and MMP‑13—which catalyze the degradation of collagen and proteoglycans within the cartilage extracellular matrix, ultimately compromising its structural integrity. Concurrently, IL‑1β exposure diminishes the production of type II collagen and proteoglycans in chondrocytes, consequently aggravating the degenerative loss of cartilage matrix⁹. The interaction between OA and inflammatory cytokines is complex. These cytokines promote cartilage degradation, synovial inflammation, abnormal bone remodeling, and neuropathic sensitization, collectively driving the initiation, progression, and pain associated with OA¹⁰.Targeted anti-inflammatory therapies represent a critical direction in OA treatment. Ongoing and future studies should aim to delineate the detailed inflammatory pathways involved in OA, providing a foundation for the development of safer, more efficacious anti‑inflammatory therapeutics and consequently improving patients’ overall quality of life¹¹.

An imbalance between the generation of reactive oxygen species (ROS) and the neutralizing efficacy of endogenous antioxidants defines oxidative stress, resulting in oxidative modification of biomolecules and subsequent tissue impairment¹². In OA, oxidative stress is identified as a key pathological mechanism. Excessive ROS directly damages chondrocytes, leading to apoptosis or functional abnormalities. By activating pro-apoptotic signaling pathways (e.g., caspase, p53), ROS induces chondrocyte death, diminishing cell survival and repair capacity. ROS concurrently trigger intracellular signaling networks, including the NF‑κB pathway, thereby enhancing matrix metalloproteinase (MMP) expression and accelerating degradation of the cartilage matrix. ROS further compromise the biochemical and structural stability of the extracellular matrix by inducing oxidative alterations in collagen and proteoglycans¹³. Notably, a positive feedback loop exists between ROS and inflammatory cytokines.A reciprocal relationship exists between ROS and pro‑inflammatory mediators, creating a continuous amplification cycle in which ROS provoke cytokine secretion—including IL‑1β and TNF‑α—thereby stimulating additional ROS formation and driving progressive inflammation and joint matrix degeneration. Consequently, therapeutic strategies targeting oxidative stress—such as antioxidant therapies or the development of ROS scavengers—may provide novel approaches for treating osteoarthritis.

Oxysophoridine (OSR), an alkaloid obtained from Sophora alopecuroides, demonstrates significant pharmacological activities, notably anti‑inflammatory actions, regulation of oxidative stress, and inhibition of apoptosis. Research indicates that OSR exhibits anti-inflammatory and antioxidant effects in various diseases¹⁴. According to Chen Jian‑Yu et al., OSR counteracts oxidative injury and inflammatory activation during liver fibrosis progression by influencing the Nrf2 and NF‑κB signaling cascades. Mechanistically, OSR inhibits the induction of iNOS and COX‑2, thereby dampening inflammatory signaling. Furthermore, by lowering malondialdehyde (MDA) levels and increasing glutathione (GSH) levels, OSR protects cell membranes from oxidative damage and decreases lactate dehydrogenase (LDH) release, thus exerting antioxidant effects. Evidence from Wang Hongbo et al. indicates that OSR confers neuroprotection in murine cerebral ischemia/reperfusion models, primarily by attenuating oxidative stress and repressing the expression of the NMDA receptor subunit NR1¹⁵. Consistent with these findings, Wang Yong‑Sheng and co‑workers revealed that OSR confers neuroprotection against ischemic damage in murine models through modulation of inflammatory mediators and suppression of the NF‑κB signaling cascade. These findings highlight OSR’s potential for treating diseases caused by inflammation and oxidative stress¹⁶. Although OSR exhibits promising pharmacological effects, its specific molecular mechanisms in osteoarthritis pathogenesis have yet to be comprehensively clarified, underscoring the need for further in‑depth research.

Our research focused on evaluating how OSR modulates IL‑1β‑induced responses in osteoarthritic chondrocytes to elucidate its protective potential for cartilage cells. In addition, we clarified the mechanisms through which OSR counteracts inflammatory signaling and oxidative stress triggered by IL‑1β stimulation. Furthermore, we evaluated OSR’s chondroprotective effects in vivo using a DMM-induced OA mouse model, confirming that OSR reduces inflammation and oxidative stress, resulting in a therapeutic benefit for osteoarthritis.

Materials and methods

Reagents

OSR was acquired from TopScience (Shanghai, China), and recombinant human IL‑1β was obtained from PeproTech (Rocky Mount, NJ, USA). Dimethyl sulfoxide (DMSO) was provided by Sigma‑Aldrich (St. Louis, MO, USA). Primary antibodies recognizing GPX4 (ab125066), SLC7A11 (ab307601), and goat anti‑rabbit IgG‑HRP (ab150077) were obtained from Abcam (Cambridge, UK). The GAPDH antibody (R24404) was supplied by ZenBio (Chengdu, China), while antibodies against IL‑6 (21,865‑1‑AP), TNF‑α (26,405‑1‑AP), COX2 (27,308‑1‑AP), and MMP13 (18,165‑1‑AP) were purchased from Proteintech (Wuhan, China).

Ethical Statement

Human articular cartilage samples were collected from individuals diagnosed with osteoarthritis who underwent total knee arthroplasty (TKA) at Xuzhou Mining Group General Hospital. Written informed consent was obtained from all participants prior to sample collection, and all procedures were conducted in accordance with the ethical principles outlined in the Declaration of Helsinki. Approval for the use of human tissues was granted by the Biomedical Research Ethics Committee of Xuzhou Mining Group General Hospital (Approval No. [2024]100901).

All animal experimental protocols were reviewed and approved by the Laboratory Animal Ethics Committee of Anhui Medical University (Approval No. LLSC20252136). All animal studies were performed in accordance with the institutional guidelines for the care and use of laboratory animals, national regulations, and the ARRIVE guidelines (https://arriveguidelines.org).

Osteoarthritis was induced in six-week-old male C57BL/6 mice through destabilization of the medial meniscus (DMM) surgery. Animals were randomly divided into five groups (n = 6 per group): Sham, DMM, DMM + DMSO, DMM + DMSO + OSR-down (25 mg/kg), and DMM + DMSO + OSR-up (50 mg/kg). The Sham and DMM groups received intraperitoneal injections of normal saline, the DMM + DMSO group received equivalent volumes of DMSO solvent, and the OSR-treated groups received intraperitoneal injections of OSR solution once per week for eight weeks.

At the end of the experimental period, mice were deeply anesthetized with sodium pentobarbital (50 mg/kg, intraperitoneal) prior to euthanasia. After confirming complete loss of reflexes, cervical dislocation was performed to ensure death. This method of euthanasia complies with the American Veterinary Medical Association (AVMA) Guidelines for the Euthanasia of Animals (2020) and was chosen to minimize animal pain and distress.All possible efforts were made to reduce the number of animals used and to alleviate suffering.

Cell isolation, identification, and culture

Cartilage specimens obtained from osteoarthritis patients during total knee arthroplasty were subjected to enzymatic digestion for chondrocyte isolation. Following recovery, cells were maintained in DMEM/F-12 supplemented with fetal bovine serum, L-glutamine, and antibiotics under standard incubator conditions (37 °C, 5% CO₂, humidified). Their morphology and identity were characterized through methylene blue cytochemical staining and ultrastructural evaluation using electron microscopy. The C28/I2 chondrocyte line was maintained in parallel under identical culture parameters.

Cell viability assay (CCK-8)

Stock preparations of OSR (10, 20, and 40 mM) were generated in DMSO, and recombinant human IL‑1β was adjusted to a final concentration of 10 ng/mL. Chondrocytes (3 × 10³ per well) were cultured in 96‑well plates for 24, 48, or 72 h under standard conditions. OSR was administered at 10–40 μM concentrations in DMEM/F‑12 medium. To induce OA‑like phenotypes, cells were preincubated with IL‑1β for 2 h prior to OSR exposure. After the designated treatments, 10 μL of CCK‑8 reagent was added, and absorbance was quantified spectrophotometrically at 450 nm.

EdU proliferation assay

To simulate inflammatory conditions, chondrocytes were stimulated with IL‑1β (10 ng/mL) for 2 h prior to exposure to OSR at graded concentrations of 10, 20, and 40 μM for a 24‑h incubation period. EdU assays (Beyotime) followed the manufacturer’s protocol. 2 × EdU was mixed 1:1 with medium and applied to cells.After the 2‑h pretreatment period, cells underwent fixation (15 min), washing, and permeabilization steps, followed by incubation with EdU solution for 30 min protected from ligh. After final washes, DAPI staining was done for 10 min before microscopy.

Western blotting

To assess protein expression following exposure to OSR and IL‑1β, chondrocytes were lysed using RIPA lysis buffer enriched with protease inhibitors, and total protein concentration was measured via the BCA method (Beyotime, China). Equivalent protein samples were subjected to SDS‑polyacrylamide gel electrophoresis and subsequently transferred onto nitrocellulose membranes. Membranes were blocked in 5% skim milk for 2 h before incubation overnight at 4 °C with the primary antibodies targeting GPX4, SLC7A11, and GAPDH (1:1000). After washing, HRP‑linked secondary antibodies were applied for 2 h at room temperature. Protein signals were detected by BeyoECL Plus chemiluminescence (Beyotime) and quantified using ImageJ analysis software.

RT-qPCR

Total RNA from chondrocyte lysates was extracted using the TRIzol Reagent (Thermo Fisher Scientific, USA) according to the manufacturer’s protocol. Complementary DNA (cDNA) was generated via reverse transcription following the supplier’s instructions. Quantitative real‑time PCR (qPCR) was carried out on an Agilent Mx3000P system using the cycling conditions of 95 °C for 5 min for initial denaturation, followed by 40 cycles of 95 °C for 10 s and 60 °C for 30 s. Gene‑specific primer pairs for MMP‑13, TNF‑α, COX‑2, and IL‑6 were designed using the NCBI Primer‑BLAST tool. The primer sequences were as follows:

GAPDH (human): Forward 5′‑GTCTCCTCTGACTTCAACAGCG‑3′; Reverse 5′‑ACCACCCTGTTGCTGTAGCCAA‑3′.

MMP‑13 (human): Forward 5′‑CCTTGATGCCATTACCAGTCTCC‑3′; Reverse 5′‑AAACAGCTCCGCATCAACCTGC‑3′.

TNF‑α (human): Forward 5′‑CCTCTCTCTAATCAGCCCTCTG‑3′; Reverse 5′‑GAGGACCTGGGAGTAGATGAG‑3′.

COX‑2 (human): Forward 5′‑TGAAACCCACTCCAAACACAG‑3′; Reverse 5′‑TCTCCTATGAGTATGAGTCTGCTGG‑3′.

IL‑6 (human): Forward 5′‑ACTCACCTCTTCAGAACGAATTG‑3′; Reverse 5′‑CCATCTTTGGAAGGTTCAGGTTG‑3′.

Lipid peroxidation assay

Lipid peroxidation levels in IL‑1β‑exposed chondrocytes subjected to graded OSR treatments were determined using the oxidation‑sensitive fluorescent probe C11‑BODIPY581/591 (Thermo Fisher Scientific). After 24 h of incubation, cells were labeled with 10 μM C11‑BODIPY at 37 °C for 30 min in darkness, subsequently rinsed with pre‑chilled PBS, harvested, and analyzed for fluorescence intensity via CytoFLEX flow cytometry (Beckman Coulter).

For the oxidative level of cells, we performed relative quantitative analysis using the FITC detection channel (excitation/emission: 488/530 nm) of a flow cytometer, and conducted relative comparisons of cellular oxidation degrees based on the fluorescence shift of oxidative probes and standardized percentages.

Determination of GSH, MDA, and 4‑HNE levels in chondrocytes.

GSH (glutathione) is the most important and abundant non-enzymatic antioxidant in cells, directly scavenging reactive oxygen species (ROS) and serving as the first line of cellular defense. Its level directly reflects the antioxidant reserve and reducing capacity of cells. MDA (malondialdehyde), as one of the most classic and stable end products of lipid peroxidation, is generated when ROS attack unsaturated fatty acids in cell membranes. Elevated MDA content directly indicates oxidative damage to cells. 4-HNE (4-hydroxynonenal), another major end product of lipid peroxidation, differs fundamentally from MDA. It reflects the total extent of damage and acts as a macroscopic indicator of structural injury caused by oxidative stress. By simultaneously analyzing GSH, MDA, and 4-HNE, researchers can comprehensively evaluate oxidative stress status across three dimensions: defense capacity (via GSH), damage intensity (via MDA), and toxic consequences (via 4-HNE). This multi-angle approach provides a holistic assessment of oxidative stress, justifying the combined use of these three assays.

GSH, MDA, and 4‑hydroxynonenal (4-HNE) levels in chondrocytes and DMM mouse cartilage were measured using commercial kits. GSH and MDA kits were from Nanjing Jiancheng, and 4-HNE ELISA kits from Sinobest Bio. The specific steps for GSH include: Cell lysis: Collect cells, wash them with pre-cooled PBS, and add an appropriate amount of reagent 1 based on the cell count. Perform repeated freeze–thaw cycles or ultrasonic disruption. Take the resulting lysate and mix it with reagent 2 at a 1:1 volume ratio. Vortex to mix thoroughly and allow it to stand on ice for 5–10 min. Centrifuge at 4 °C and 3500 rpm for 10 min. Carefully collect the supernatant, which serves as the sample solution for testing.Mix the sample solution with the working solution (prepared according to the instructions). Gently shake the microplate to ensure thorough mixing and allow it to stand for 5 min. Measure the absorbance of each well at 405 nm using a microplate reader. The specific steps for MDA are as follows: Discard the cell culture supernatant, collect the cells into a centrifuge tube, add 0.3 mL of reagent five extraction solution, mix well for 2 min, then disrupt the cells (via ultrasonication) to form a suspension. Take a 0.1 mL sample in a 1.5 mL centrifuge tube (pre-puncture a small hole in the tube cap using a needle heated over an alcohol lamp). Vortex the mixture thoroughly, place it in a water bath at 95 °C or above for 40 min, remove and cool under running water, centrifuge at 4000 rpm for 10 min, and measure the absorbance at 530 nm. First scan the empty microplate, then accurately pipette 0.25 mL of the supernatant from each tube into a 96-well plate, and measure the absorbance of each well using a microplate reader. Specific steps for 4-HNE:Discard the culture medium and wash the cells with pre-cooled PBS. Add an appropriate amount of lysis buffer and lyse the cells on ice for 15 min. Collect the lysate and centrifuge at 4 °C and 12,000 rpm for 15 min. Use the supernatant as the test sample. Add standard solutions and samples to the antibody-precoated microplate wells (50 μL per well is recommended). Gently shake to mix, cover with a plate sealer, and incubate at 37 °C for 30–60 min. Discard the liquid from the wells and add 300 μL of wash buffer to each well. Let stand for 30 s, then discard. Repeat this wash step 3–5 times, and after the final wash, blot the plate dry on absorbent paper.Add 50 μL of biotin-labeled antibody working solution to each well, cover with a plate sealer, and incubate at 37 °C for 30 min. Wash the plate again 3 times. Add 50 μL of HRP-Streptavidin working solution to each well, cover with a plate sealer, and incubate at 37 °C for 30 min. Add 50 μL of TMB substrate to each well, cover, and incubate at 37 °C in the dark for 15 min. Wash the plate 3 times. Immediately measure the absorbance (OD value) of each well at a wavelength of 450 nm using a microplate reader.

Molecular markers and flow cytometry

Human articular chondrocytes and C28/I2 cells were plated in 6‑well plates and cultured for 24 h prior to treatment.The cells were then stimulated with IL‑1β (10 ng/mL) for 2 h and subsequently incubated with graded concentrations of OSR (10, 20, or 40 μM) for a further 24 h. After the treatment period, cells were fixed in 4% paraformaldehyde, permeabilized using 90% methanol, and incubated at room temperature for 1 h with primary antibodies against GPX4 and SLC7A11 (1:2000). Following extensive washing, samples were stained with Alexa‑fluor‑conjugated secondary goat anti‑rabbit IgG (1:2000) for 30 min in the dark. Labeled cells were resuspended in 1 × PBS and analyzed for fluorescence intensity using a flow cytometer (CytoFLEX, Beckman Coulter, USA). Normal rabbit IgG (Abcam, ab172730) served as the negative control.

Molecular docking

The ligand structure of OSR (PubChem CID 114,850) was acquired from the PubChem compound repository in SDF file format for subsequent computational analysis. Hydrogens were added and energy minimized (Amber10-EHT, rigid water solvent, RMS gradient 0.1 kcal/mol/Å, no constraints). The optimized structure underwent conformation searching (MOE, LowModeMD). Docking to Protein.pdb was performed in MOE using TriangleMatcher and London δG scoring. PLIF clustering was applied, and optimal docking conformations were selected based on score and binding mode. The S value indicates protein–compound affinity (lower S = stronger binding).

Cellular thermal transfer assay (CETSA)

Human primary chondrocytes and C28/I2 cell lines were cultured in complete medium under typical conditions (37 °C, 5% CO₂, humidified atmosphere) until reaching a cell density of approximately 90%. Following lysis, the cell lysates were combined and portioned (1 mL per sample) into DMSO and OSR experimental groups. Each aliquot was incubated for 2 h at room temperature with either 1 μL DMSO or 1 μL of 500 mM DDZ. Subsequently, 150 μL supernatant from each treatment was collected, exposed to incremental heating at 50, 55, 60, 65, 70, and 75 °C for 5 min, and immediately cooled on ice to halt denaturation. Protein samples were then prepared for Western blot analysis to assess GPX4 expression and thermal stability.

Animal experiments

Six‑week‑old male C57BL/6 mice were randomly assigned to five treatment groups (n = 6 per group): sham‑operated, DMM model, DMM + DMSO vehicle, DMM + OSR 25 mg/kg, and DMM + OSR 50 mg/kg. Osteoarthritis was established by surgical destabilization of the medial meniscus (DMM), and OSR was delivered to the knee joint via intra‑articular injection once per week. Body weights were tracked every three days during the study period. After 90 days of treatment, animals were humanely euthanized, and knee joint tissues were harvested, fixed, decalcified, paraffin‑embedded, sectioned, and examined using hematoxylin‑eosin staining, immunohistochemical analysis, and immunofluorescence microscopy.

Immunohistochemical and immunofluorescence staining methodology

Immunohistochemistry (IHC) and immunofluorescence (IF) staining were performed on mouse bone tissue. The specific procedures are as follows:

First, fresh bone tissue was immediately fixed in 4% paraformaldehyde. After fixation, the samples were transferred to 10% EDTA decalcification solution for decalcification, followed by embedding and sectioning. The sections were dewaxed in xylene and then rehydrated through a graded series of ethanol. Antigen retrieval was performed using citrate buffer (pH 6.0) at 95 °C for 20 min. Subsequently, blocking was carried out with 5% BSA + 0.3% Triton X-100. For IHC staining, primary antibodies (GPX4 (ab125066), SLC7A11 (ab307601), IL-6 (21,865–1-AP), TNF-α (26,405–1-AP), COX2 (27,308–1-AP), and MMP13 (18,165–1-AP)) were applied at a dilution of 1:200 and incubated overnight at 4 °C, followed by incubation with an HRP-conjugated secondary antibody and DAB color development. For TUNEL staining, sections were placed flat in a humidity chamber (protected from light) and incubated in a 37 °C constant temperature incubator for 60 min. After incubation, the sections were transferred to PBS and gently agitated for three washes (5 min each) to terminate the reaction and remove unbound fluorescent dyes. Nuclei were counterstained with hematoxylin (for IHC) or DAPI (for Tunel). Imaging was performed using a slide scanning microscope.

Statistical analysis

Quantitative data are expressed as mean values ± standard deviation (SD). Group differences were evaluated by one‑way analysis of variance (ANOVA) using GraphPad Prism (version 8.0; GraphPad Software, USA), and p values below 0.05 were interpreted as statistically significant.

Results

OSR promotes chondrocyte activity under inflammatory conditions

The structural formula of OSR is shown in Fig. 1A. Human chondrocytes were isolated from total knee replacement samples and identified by morphology (Fig. 1B1) and toluidine blue staining, which revealed elliptical or spindle-shaped cells with purple-stained cytoplasmic proteoglycans (Fig. 1B). To mimic inflammation-induced injury, IL‑1β‑treated chondrocytes (including C28/I2 cells) were exposed to graded concentrations of OSR. CCK‑8 and EdU assays demonstrated that OSR dose‑dependently restored chondrocyte viability impaired by IL‑1β (Fig. 1C–E). Meanwhile, OSR itself exhibited no toxicity toward chondrocytes (Supplementary Data 1). Collectively, OSR protects chondrocytes from inflammation‑induced impairment.

Fig. 1.

Oxysophoridine (OSR) mitigates IL‑1β‑induced inflammatory responses in chondrocytes.(A) Structural representation of OSR.(B) Morphological features of human primary chondrocytes observed under (1) light microscopy and (2) toluidine blue staining (scale bar = 50 µm).(C) CCK‑8 assay results showing the effects of different OSR concentrations on the viability of human primary chondrocytes and C28/I2 cells after 24, 48, and 72 h (n = 6).(D) EdU live/dead staining images of both cell types following 24 h of OSR exposure (scale bar = 200 µm).(E) Quantitative evaluation of EdU‑positive cells using ImageJ (n = 3).Values are presented as the mean ± SD from three or six independent experiments. ****p < 0.0001, ***p < 0.001, **p < 0.01, ns > 0.05.

OSR inhibits inflammation and reduces oxidative stress in chondrocytes

Inflammation is central to osteoarthritis, making anti‑inflammatory actions critical for potential therapeutics. IL‑1β stimulation markedly increased IL‑6, TNF‑α, COX‑2, and MMP13 expression in chondrocytes, whereas OSR treatment suppressed these markers in a dose‑dependent manner (Fig. 2A), indicating notable anti‑inflammatory activity. Because oxidative stress contributes to osteoarthritis progression, the lipid peroxidation probe C11‑BODIPY581/591 was used to assess lipid ROS, and OSR significantly decreased lipid ROS levels in IL‑1β‑treated chondrocytes (Fig. 2B–C). OSR also elevated GSH concentrations while reducing MDA and 4‑HNE levels (Fig. 2D–F). These data demonstrate that OSR mitigates both inflammatory and oxidative‑stress responses in chondrocytes.

Fig. 2.

OSR suppresses inflammation by inhibiting ROS production in chondrocytes. IL-1β-stimulated human primary chondrocytes and C28/I2 cells were cultured with OSR for 24 h. (A) RT-qPCR analysis of IL-6, TNF-α, and IL-1β mRNA expression. (B) Intracellular lipid ROS levels measured by flow cytometry after C11-BODIPY581/591 staining. (C) Quantification of lipid ROS. (D) GSH levels assessed with a glutathione assay kit. (E) MDA content measured by MDA assay kit. (F) 4-HNE expression determined by ELISA. Data are mean ± SD from three independent experiments. ****P<0.0001, ***P<0.001, **P<0.01, *P<0.05, ns>P 0.05.

OSR inhibits oxidative stress in chondrocytes by upregulating GPX4 expression

Lipid peroxidation reflects oxidative stress, and GPX4 serves as a key enzyme that reduces lipid hydroperoxides via GSH to limit ROS damage. OSR treatment increased GPX4 mRNA and protein levels in both human chondrocytes and C28/I2 cells (Fig. 3A–C), as further confirmed by flow cytometry (Fig. 3D–E). Molecular docking (PLIF clustering) revealed that OSR directly binds GPX4 with a binding energy of − 5.21 kcal/mol, suggesting enhanced protein stability (Fig. 3F). The CETSA assay demonstrated that OSR protected GPX4 from heat‑induced degradation (Fig. 3G–H). Together, these results indicate that OSR alleviates oxidative stress through dual regulation of GPX4—enhancing its expression and stabilizing the protein.

Fig. 3.

OSR reduces oxidative stress in chondrocytes by upregulating GPX4. (A–E) IL-1β-stimulated chondrocytes (primary and C28/I2) were treated with OSR (24 h), and GPX4 levels were analyzed by qPCR (A), Western blot (B–C), and flow cytometry (D–E), quantified with ImageJ. (F) Binding conformation of OSR and GPX4 shown in 3D/2D views, highlighting hydrogen bonds. (G–H) CETSA/Western blot analysis showed OSR stabilizes GPX4 at increasing temperatures, quantified by ImageJ. Data: mean ± SD (n=3); significance: ****P<0.0001, ***P<0.001, **P<0.01, *P<0.05, ns > 0.05.

OSR suppresses oxidative stress in osteoarthritis by activating the GSH synthesis-related pathways.

GSH, a major intracellular antioxidant, maintains redox balance through regulation of genes such as SLC7A11, GCLC/GCLM, GSS, and SLC3A2. IL‑1β‑induced chondrocytes (primary and C28/I2) displayed decreased SLC7A11 and GCLC/GCLM expression, whereas GSS and SLC3A2 levels remained unchanged (Fig. 4A). Under oxidative stress, OSR treatment significantly upregulated SLC7A11 expression at both mRNA and protein levels, thus enhancing antioxidant capacity (Fig. 4A–C). Flow cytometry confirmed that OSR elevated SLC7A11 expression, attenuated oxidative stress, and alleviated inflammation (Fig. 4D–E). Therefore, OSR promotes GSH biosynthesis via SLC7A11 activation, sustaining redox equilibrium under osteoarthritic conditions.

Fig. 4.

OSR suppresses osteoarthritis oxidative stress and inflammation by activating GSH synthesis genes. (A) RT-qPCR heatmap shows mRNA levels of SLC7A11, GCLC/GCLM, GSS, and SLC3A2. (B–C) Western blot and ImageJ quantification of SLC7A11 protein. (D–E) Flow cytometry of SLC7A11 in IL-1β-induced chondrocytes treated with OSR; data shown as histograms. Results: mean ± SD (n=3); ****P<0.0001, ***P<0.001, **P<0.01, *P<0.05, ns>P 0.05.

OSR efficiently and safely restores chondrocyte activity in DMM model mice

To evaluate OSR’s chondroprotective and anti‑osteoarthritic effects in vivo, a DMM model was established in C57BL/6 mice with five groups: Sham, DMM, DMSO, OSR 25 mg/kg, and OSR 50 mg/kg (Fig. 5A). TUNEL staining confirmed higher cell viability in the OSR groups compared with the DMM group (Fig. 5B–C). OSR treatment did not significantly affect body weight (Fig. 5D), and H&E staining showed no evident organ toxicity (Fig. 5E). These findings demonstrate that OSR effectively restores chondrocyte activity and exhibits good in vivo safety.

Fig. 5.

OSR enables effective and safe cartilage repair in DMM mice. Five groups: Sham, DMM, DMSO, OSR 25 mg/kg, and OSR 50 mg/kg. (A) Experimental schematic. (B–C) TUNEL immunofluorescence of cartilage, quantified by ImageJ. (D) Mouse body weights recorded every 30 days (n=6). (E) H&E staining of heart, liver, spleen, lung, and kidney. Scale bar: 100 μm. B–E: mean ± SD (n=3); F: mean ± SD (n=6). Significance: ****P<0.0001, ***P<0.001, **P<0.01, ns>P 0.05.

OSR alleviates inflammation and oxidative stress in DMM model mice

Immunohistochemical analysis showed that DMM and DMSO groups displayed elevated IL‑6, TNF‑α, COX‑2, and MMP13 expression in mouse knee cartilage. OSR administration—especially at 50 mg/kg—significantly reduced these inflammatory markers, approaching levels observed in the Sham group (Fig. 6A–B), thereby confirming strong in vivo anti‑inflammatory activity. Consistent with the in vitro findings, OSR increased GPX4 and SLC7A11 expression in cartilage (Fig. 7A–B) and improved oxidative‑stress parameters, including elevated GSH and reduced MDA and 4‑HNE levels (Fig. 7C–G). Collectively, OSR exerts potent anti‑inflammatory and antioxidant effects by upregulating GPX4 and SLC7A11, thereby decelerating osteoarthritis progression (Fig. 8).

Fig. 6.

OSR reduces inflammation in DMM model mice. Five groups: Sham, DMM, DMSO, OSR 25 mg/kg, and OSR 50 mg/kg. (A–B) Immunohistochemical analysis of IL-6, TNF-α, COX-2, and MMP13 in cartilage, quantified by ImageJ. Scale bar: 100 μm. Data are mean ± SD from three independent experiments. Significance: ****P<0.0001, ***P<0.001, **P<0.01, *P<0.05, ns>P 0.05.

Fig. 7.

OSR reduces oxidative stress in DMM model mice. Five groups: Sham, DMM, DMSO, OSR 25 mg/kg, and OSR 50 mg/kg. (A–B) Immunohistochemistry of GPX4 and SLC7A11 in cartilage chondrocytes, quantified by ImageJ. Scale bar: 100 μm. (C–E) Levels of GSH, MDA, and 4-HNE in chondrocytes. Data are mean ± SD from three independent experiments. Significance: ****P<0.0001, ***P<0.001, **P<0.01, *P<0.05, ns>P 0.05.

Fig. 8.

Schematic illustration of OSR’s protective mechanism in osteoarthritis. OSR mitigates cartilage damage from inflammatory factors in vivo and in vitro by suppressing ROS-induced oxidative stress and regulating GPX4 activity, thereby preserving cartilage integrity.

Discussion

The present study demonstrates that OSR exerts protective effects on osteoarthritic chondrocytes by restoring cellular activity under inflammatory stress conditions. Exposure to OSR significantly attenuated the IL‑1β‑induced reduction in cell viability in both human primary chondrocytes and C28/I2 cells, reflecting a robust cytoprotective capacity of OSR under inflammatory conditions. Persistent inflammatory signaling is a hallmark of OA progression, mediated by elevated production of IL‑6, TNF‑α, COX‑2, and MMP13, which together accelerate cartilage degradation and joint deterioration^17–19.OSR substantially downregulated pro‑inflammatory factor expression in IL-1β-stimulated chondrocytes, underscoring its potential as an emerging candidate for osteoarthritis therapy.

Excessive oxidative stress is another central driver of OA pathogenesis, amplifying inflammation and accelerating chondrocyte apoptosis through ROS‑induced cellular injury and activation of degenerative signaling cascades^20–22.Mutual reinforcement between oxidative stress and inflammation establishes a pathological feedback loop that aggravates cartilage damage and joint deterioration^23,24.In this study, OSR efficiently reduced intracellular ROS accumulation and enhanced intrinsic antioxidant defenses, thereby breaking this pathological feedback loop and improving cellular redox homeostasis.

Mechanistically, the antioxidant capacity of OSR is largely attributed to its ability to induce and stabilize GPX4, a selenium‑dependent enzyme that neutralizes lipid peroxides and serves as a central safeguard against ferroptotic cell death^25,26.Mechanistic analyses revealed that OSR promotes GPX4 expression at both the transcriptional and translational stages and engages in direct binding with GPX4, validated through computational docking and CETSA experiments. Such interaction reinforces GPX4 structural stability and sustains its peroxidase activity during oxidative stress. This dual mode of GPX4 regulation is pivotal for preserving intracellular redox equilibrium and preventing oxidative cartilage degeneration²⁷.

Simultaneously, OSR upregulated SLC7A11, a critical subunit of system Xc⁻, which mediates cystine–glutamate exchange and regulates cystine uptake, the rate‑limiting step in GSH synthesis^28,29. GSH is the primary intracellular antioxidant and one of the key regulatory factors in GPX4 expression^30,31.Through restoration of SLC7A11 expression, OSR facilitated greater cystine uptake and GSH accumulation, while simultaneously lowering the levels of lipid peroxidation by-products such as MDA and 4-HNE. These findings highlight OSR’s dual ability to augment antioxidant capacity and mitigate oxidative stress-induced chondrocyte damage.

In vivo experiments using the DMM mouse model corroborated the in vitro results, demonstrating that OSR mitigates joint degeneration in an experimental setting that closely mimics osteoarthritic progression. OSR administration significantly reduced chondrocyte apoptosis (TUNEL‑positive cells), collectively indicating functional repair of OA-damaged cartilage. Immunohistochemical analysis confirmed that OSR downregulated inflammatory mediators and oxidative stress markers while upregulating GPX4 and SLC7A11 expression in joint tissues.Furthermore, OSR demonstrated excellent systemic tolerability, with consistent body weight and preserved histoarchitecture across major organs, indicating minimal off‑target toxicity^32,33.Current pharmacological treatments for OA primarily provide symptomatic relief rather than disease modification.While NSAIDs and analgesics provide transient pain relief, they are ineffective in rebuilding cartilage or interrupting the chronic inflammatory and oxidative feedback loops that propel osteoarthritic progression³⁴. Moreover, long-term NSAID use carries notable safety concerns, including an increased risk of gastrointestinal damage and cardiovascular complications³⁵. These limitations underline the critical need for agents capable of targeting molecular mechanisms that drive OA development instead of providing transient relief.

Despite promising results from preclinical studies, translation of antioxidant or anti‑inflammatory therapies into clinical success has been hindered by ambiguous molecular mechanisms and inconsistent safety profiles^36,37. OSR, a quinolizidine alkaloid purified from Sophora alopecuroides, has been reported to exert potent anti‑inflammatory and antioxidative effects in experimental models of hepatic fibrosis, cerebral ischemia, and cancer, highlighting its potential as a multifunctional cytoprotective agent^38–40; however, its specific role in OA had not been characterized before this study. Our findings reveal that OSR addresses fundamental pathogenic mechanisms of OA by restoring redox homeostasis through the GPX4/SLC7A11-GSH axis, thereby simultaneously attenuating inflammation and oxidative stress while promoting cartilage regeneration.

Taken together, these results position OSR as a promising dual‑action candidate for OA therapy, providing complementary anti‑inflammatory and antioxidant properties superior to single‑target interventions. Future research should focus on pharmacokinetic profiling, dose optimization, and genetic model validation (e.g., GPX4 or SLC7A11 knockout mice) to fully define OSR’s regulatory network. Ultimately, well-designed clinical trials will be necessary to confirm its therapeutic efficacy and safety in human OA patients.

Conclusion

Taken together, our findings reveal that OSR restores cartilage homeostasis in osteoarthritis by dampening ROS‑driven oxidative stress and enhancing the glutathione antioxidant network through GPX4 stabilization and SLC7A11 activation. By coupling antioxidant reinforcement with anti‑inflammatory effects, OSR preserves chondrocyte survival and tissue architecture. These results underscore OSR as an innovative and safe therapeutic candidate with strong potential for clinical translation in osteoarthritis management.

Supplementary Information

Below is the link to the electronic supplementary material.

Abbreviations

CETSA

Cellular thermal shift assay

COX‑2

Cyclooxygenase‑2

DMSO

Dimethyl sulfoxide

GPX4

Glutathione peroxidase 4

GSH

Glutathione

H&E

Hematoxylin and eosin

IL‑6

Interleukin‑6

MDA

Malondialdehyde

MMP‑13

Matrix metallopeptidase 13

OA

Osteoarthritis

OSR

Oxysophoridine

ROS

Reactive oxygen species

SLC7A11

Solute carrier family 7 member 11

TNF‑α

Tumor necrosis factor‑α

4‑HNE

4‑Hydroxynonenal

Author contributions

Jun Tu (First Author): Conceptualization, Methodology, Software, Investigation, Formal Analysis, Writing—Original Draft, Writing—Review & Editing; Zhiwei Peng: Data Curation, Writing—Original Draft, Writing—Review & Editing; Haobo Liu: Visualization, Investigation, Writing—Review & Editing; Xiyang Sun :Visualization, Writing—Review & Editing; Jianqi Zhao: Resources, Supervision, Writing—Review & Editing; Bin Xu: Software, Validation, Writing—Review & Editing; XuZhu: Concepualization,Methodology, Writing—review Editing. Baofang Liu:(Corresponding Author): Conceptualization, Funding Acquisition, Resources, Supervision, Writing—Review Editing. All authors agreed on the journal for submission, reviewed and approved all versions of the manuscript, and accept responsibility for the published content.

Funding

This work was supported by the Health Research Program of Anhui (Grant No. AHWJ2024Aa20135).

Data availability

All data generated or analyzed during this study are available from the corresponding author upon reasonable request. The datasets include raw Western blot images, RT-qPCR Ct values, flow cytometry files, histological and immunohistochemical images, and molecular docking results. All data have been carefully validated, and the authors affirm their accuracy and reliability. Requests for access should be directed to the corresponding author.

Declarations

Competing interests

The authors declare no competing interests.

Consent for publication

Written informed consent for publication was obtained from all participants who provided human cartilage samples. All identifying information has been removed to ensure anonymity, and signed consent forms are available for review by the journal’s editorial office upon request.