Targeting the SARM1–cADPR–Ca²⁺ pathway attenuates mitochondrial fragmentation and osteoarthritis progression

Abstract

Introduction

Osteoarthritis (OA) is a prevalent degenerative joint disease accompanied by increased number of senescent chondrocytes. Mitochondrial dysfunction is a well-established hallmark of chondrocyte senescence in OA pathogenesis. Sterile α and Toll/Interleukin-1 Receptor motif–containing 1 (SARM1), known to drive mitochondrial impairment in various cell types, has not been thoroughly investigated in the context of chondrocyte aging or OA.

Methods

We established a doxorubicin (DOX)-induced senescence model in primary mouse chondrocytes. Gain- and loss-of-function approaches were employed using siRNA-mediated knockdown and lentiviral overexpression of SARM1, followed by assessment of senescence markers, mitochondrial function, and morphology. To investigate the mechanistic pathway, exogenous cyclic ADP-ribose (cADPR) and its specific inhibitor 8-Br-cADPR were applied, with subsequent evaluation of intracellular calcium dynamics and Drp1 translocation to mitochondria. We next tested the efficacy of blocking this SARM1/cADPR axis both ex vivo on human femoral head tissue as well as in an experimental OA mouse model.

Results

Chondrocytes isolated from human OA cartilage and aged murine cartilage showed increased expression of SARM1. Knockdown of SARM1 reduced DOX-induced chondrocyte senescence and mitochondrial dysfunction, while overexpression of wild-type but not catalytic-inactive SARM1-TIR domain mutant (TIR-E642A) induced intrinsic apoptosis and mitochondrial fragmentation. Exogenous cADPR recapitulated senescence and mitochondrial fragmentation, whereas treatment with 8-Br-cADPR abolished SARM1-dependent effects. Mechanistically, SARM1-generated cADPR increased intracellular calcium levels, triggering Drp1 phosphorylation at Ser616 and dephosphorylation at Ser637, thereby resulting in Drp1-FIS1 interaction and mitochondrial fission. Interestingly, pharmacological or genetic inhibition of the SARM1/cADPR pathway ameliorated cartilage degradation in the experimental OA model.

Conclusion

We show that SARM1 mediates mitochondrial fragmentation by activating cADPR-dependent calcium signaling, which in turn promotes Drp1 binding to FIS1. This suggests an unappreciated role for the SARM1–cADPR pathway in OA etiology and presents this pathway as an attractive candidate to be targeted therapeutically.

Supplementary Information

The online version contains supplementary material available at 10.1186/s13075-026-03837-3.

Introduction

Osteoarthritis (OA) is the most common chronic joint disease, typically characterized by cartilage degeneration, subchondral bone sclerosis, osteophyte formation, and synovitis [1]. Due to the global aging population, OA is placing a heavy socioeconomic burden on societies worldwide [2–4]. Currently, no disease-modifying therapies exist for OA [5], which underscores the critical need to elucidate its pathogenesis and find new therapeutic targets.

Mitochondria play a central role in chondrocyte metabolism, supplying key energy in the low-oxygen environment of avascular cartilage. Disruptions in mitochondrial homeostasis are now recognized as a major contributor to the development and progression of osteoarthritis (OA) [6, 7]. These organelles are highly dynamic, constantly undergoing fusion and fission—processes collectively referred to as mitochondrial dynamics—that help regulate their shape and function [8]. This balance is essential for mitochondrial quality control: it allows removal of damaged parts, supports efficient energy production, and helps cells cope with stress [9]. Dysregulation of this balance has been associated with chondrocyte dysfunction and apoptosis throughout OA pathogenesis [10, 11]. In fact, studies have observed fragmented mitochondria in articular cartilage from both human OA patients and mouse models of the disease [12, 13]. Despite these observations, the precise molecular mechanisms driving this pathological shift in mitochondrial dynamics are not yet fully clear.

Mitochondrial-interacting proteins are proteins that physically associate with or functionally engage mitochondria to regulate their morphology, metabolic functions, and stress responses. Research indicates that these proteins play critical roles in various mitochondria-related disorders [14, 15]. Among them, SARM1 (Sterile Alpha and TIR Motif Containing 1) has attracted attention as an important regulator. It localizes to the mitochondrial outer membrane, a position that puts it in direct contact with the machinery governing mitochondrial dynamics. When activated, SARM1 throws off this balance by pushing excessive fission, generating a surplus of small, dysfunctional mitochondrial fragments. These impaired organelles struggle to produce energy efficiently, ultimately driving mitochondrial dysfunction. In neurodegenerative conditions, SARM1 disrupts axonal energy metabolism severely [16]. Similar SARM1-mediated mitochondrial damage has been reported in other non-dividing (post-mitotic) cell types, such as cardiomyocytes and hepatic stellate cells [17–20]. This evidence prompts the hypothesis that SARM1 may play an important role in driving chondrocyte senescence and cartilage degeneration.

At baseline, SARM1 is autoinhibited. Cellular stress or injury can relieve this inhibition by triggering a decline in NAD⁺. In this way, SARM1 acts as a key metabolic sensor, converting the energetic state of the cell into a catalytic signal [21]. Once activated, the SARM1 octameric complex disassembles and relocates to the mitochondria, where dimerization of the TIR domains drives its NADase activity [16, 22]. Activated SARM1 hydrolyzes NAD⁺ to produce Nam (Nicotinamide) and cADPR, rapidly triggering a cascade of pathological changes: collapse of the mitochondrial membrane potential, intracellular calcium overload, and, ultimately, cell death [23]. But for now, the function of SARM1 on promoting chondrocyte senescence and OA has not been reported yet.

In this work, we discovered increased SARM1 expression in OA patient’s articular cartilage and aged mouse joints. Subsequent loss-of-function experiments demonstrated that SARM1 plays a role in inducing mitochondrial dysfunction and chondrocyte senescence. Through comparison between chondrocytes overexpressing wild-type SARM1 and TIR-domain catalytically inactive mutants, we showed that SARM1 activation induces mitochondrial dysregulation. Fu rthermore, we validated that SARM1, via its enzymatic product—the calcium-mobilizing agent cADPR—induces calcium overload, disrupts mitochondrial dynamics homeostasis, and promotes mitochondrial fragmentation, ultimately driving mitochondrial dysfunction in OA Progression.

Materials and methods

Human cartilage samples

Osteoarthritic (OA) cartilage samples were obtained from six patients (n = 6; age: 61.33 ± 2.88 years; 5 females, 1 male) undergoing total knee arthroplasty for end-stage OA. Non-OA controls were collected from six trauma or severe diabetic foot amputation patients (n = 6; age: 56.5 ± 7.69 years; 3 females, 3 males). Patients with rheumatoid arthritis, metabolic disorders, or other inflammatory conditions were excluded.

Animals and establishment of OA model

Male C57BL/6J mice (wild-type, 12–96 weeks old) were purchased from Bestest Biotechnology (Zhuhai, China) and housed under SPF conditions (22–26 °C, 40–60% humidity, 12-h light/dark cycle). To induce OA, 12-week-old mice underwent destabilization of the medial meniscus (DMM): the meniscotibial ligament was exposed and transected via a medial parapatellar approach. 10 µL AAV (5 × 10¹¹ vg/ml) containing Sarm1 shRNA (AAV5-sarm1shRNA) was injected into the knee joint of surgically treated C57BL/6J mice. Mice were euthanized at 8 weeks post-injection for histological assessment. The control groups were all treated with negative control (AAV-control) for the same indicated time. All shRNA sequences used in this study are provided in Table 1.

Table 1.

shRNA sequences

shRNA	Sense Strand (5’ → 3’)	Antisense Strand (5’ → 3’)
shSarm1-1	CAAGGTCTATGCGATGCTAT	ATAGCATCGCATAGACCTTG
shSarm1-2	CAGCCAGAGAAATGCTACAT	ATGTAGCATTTCTCTGGCTG
shSarm1-3	AGAACATTGTGCCCATCATT	AATGATGGGCACAATGTTCT
shControl	ATACAGCCATCGCTGTTATTC	GAATAACAGCGATGGCTGTAT

RNA sequencing (RNA-seq) experimental methods

All operations were performed in an RNase-free ultra-clean workbench with RNase-free consumables to avoid RNA degradation, with all tissue handling conducted on ice. Immediately after mouse euthanasia, knee joints were dissected, the superficial cartilage layer was excised on ice, snap-frozen in liquid nitrogen, and ground into powder in a pre-cooled mortar. TRIzol lysis solution (Invitrogen, USA) was added per the manufacturer’s instructions, with repeated pipetting for complete tissue lysis.

Total RNA was isolated via chloroform extraction, isopropanol precipitation, and two pre-cooled 75% ethanol washes, then dissolved in RNase-free water. RNA concentration, purity and integrity were assessed via NanoDrop spectrophotometer and Agilent 2100 Bioanalyzer; only samples with RIN ≥ 8 were eligible for library construction. Poly(A) mRNA was enriched with oligo(dT) magnetic beads, fragmented, and reverse-transcribed into double-stranded cDNA. Libraries were constructed via end repair, A-tailing, adapter ligation and PCR amplification. After quality verification, high-throughput sequencing was performed on the Illumina NovaSeq platform with 150 bp paired-end reads, and differentially expressed genes were identified using DESeq2.

Isolation and culture of primary chondrocytes

Primary chondrocytes were isolated from neonatal C57BL/6J mouse knee cartilage. Briefly, neonatal mice were euthanized, and the knee cartilage tissue was carefully dissected. Under sterile conditions, the cartilage tissue was minced into small fragments and thoroughly washed with PBS. Cartilage samples were initially digested with 3% Collagenase D for 1 h, followed by further digestion with a diluted concentration of 0.5% Collagenase D overnight. After digestion, the cell suspension was filtered and centrifuged at 100 × g for 5 min.

The purified cells were cultured in DMEM/F12 medium (Gibco) supplemented with 10% fetal bovine serum (FBS; Gibco, Cat. # A5670701), 1% Penicillin-Streptomycin at 37 °C under 5% CO₂. Experiments used first-passage cells at 70–80% confluency. Senescence was induced with 100 nM doxorubicin (24 h) and confirmed by SA-β-gal staining.

Transfection of siRNA

siRNA targeting Sarm1 and negative control siRNA were purchased from Tsingke Biotechnology (Beijing, China) and are listed in Table 2. Primary chondrocytes were transfected with 50 nM Sarm1 siRNA or negative control siRNA using Lipofectamine 3000 reagent (Sigma-Aldrich), according to the manufacturer’s protocol.

Table 2.

siRNA sequences

siRNA	Sense Strand (5’ → 3’)	Antisense Strand (5’ → 3’)
siSarm1-1	GCUUAUCCAAAGCGUCAUA	UAUGACGCUUUGGAUAAGC
siSarm1-2	GGUUUCUUACUCUACGAAU	AUUCGUAGAGUAAGAAACC
siSarm1-3	GAUGGUGACCUGCUUCUAA	UUAGAAGCAGGUCACCAUC
Negative Control	GGUUAAUCGCGUAUAAUACGCG	CGCGUAUUAUACGCGAUUACC

Transduction of lentiviral

Lentiviruses overexpressing wild-type SARM1 or the TIR domain mutant E642A (BankIt accession: KY584388–KY584401) were obtained from OBiO Technology (Shanghai, China). Chondrocytes were seeded in 6-well plates and transduced at 30–50% confluency with MOI = 50. After 24 h, virus-containing medium was replaced with fresh complete medium. Cells were harvested 48 h post-transduction, and overexpression was validated by western blot.

RNA isolation and qRT-PCR

Total RNA was extracted using the Omega Bio-tek Total RNA Kit (Cat. # R6834). cDNA was synthesized with SuperScript IV UniPrime Kit (Invitrogen). qPCR was performed using Taq Pro Universal SYBR Master Mix (Vazyme, Cat. # Q712-02) on an Applied Biosystems real-time PCR system. All steps followed manufacturer protocols. The primer sequences used in this study are listed in Table 3.

Table 3.

Primers for qPCR detection

Gene name	Forward primer(Human)	Reverse primer (Human)
SARM1	AGCAACAAGCACTCTGAATGG	AGATCACGCCTAGACCGATG
COL2A1	TGGACGATCAGGCGAAACC	GCTGCGGATGCTCTCAATCT
MMP13	ACTGAGAGGCTCCGAGAAATG	GAACCCCGCATCTTGGCTT
CDKN1A	TGTCCGTCAGAACCCATGC	AAAGTCGAAGTTCCATCGCTC
CDKN2A	GATCCAGGTGGGTAGAAGGTC	CCCCTGCAAACTTCGTCCT
GAPDH	GGAGCGAGATCCCTCCAAAAT	GGCTGTTGTCATACTTCTCATGG

Histological assessment

Tissues were fixed in 4% paraformaldehyde (24 h, 4 °C), decalcified in 10% EDTA (pH 7.4, 4 weeks), embedded in paraffin, and sectioned sagittally (3 μm). Sections were stained with H&E or Safranin O/Fast Green. Cartilage damage was scored using the OARSI system; synovitis was graded 0–3 as previously described [24, 25].

β-Galactosidase activity assay

Senescence was assessed by:

(1).SA-β-Gal Staining: Cellular senescence in chondrocytes was evaluated using a SA-β-Gal Staining Kit (Solarbio, Cat. # G1580) strictly following the manufacturer’s instructions.

(2).C12FDG Fluorescence Assay: Briefly, adherent cells were first pretreated with 100 nM Bafilomycin A1 (MCE, Cat. # HY-100558) for 1 h. Subsequently, the cells were stained with 6.5 µM C12FDG (MCE, Cat. # HY-126839) and incubated at 37 °C under 5% CO₂ for 1 h. Fluorescent signals were captured using a confocal microscope (Zeiss, LSM 880).

Immunohistochemistry (IHC) and Immunofluorescence (IF)

For IHC, sections were deparaffinized, rehydrated, quenched with 3% H₂O₂, subjected to antigen retrieval (0.1% trypsin, 37 °C, 20 min), blocked with 5% goat serum, and incubated overnight at 4 °C with primary antibodies. Detection used HRP-polymer secondary antibody and DAB (Proteintech PK10006).

For IF, samples were incubated with Alexa Fluor™ 488 (1:500, A-11001) or Alexa Fluor™ 568 (1:500, A-11011)-conjugated secondary antibodies (1 h, RT, dark), counterstained with DAPI, and imaged using Zeiss LSM 880.

Western blot

Proteins were extracted from chondrocytes using RIPA buffer (Beyotime, China) supplemented with protease and phosphatase inhibitors (Roche). Protein concentration was determined by the BCA assay (Beyotime, China). Equal amounts of protein were separated by 10–12.5% SDS-PAGE and transferred to PVDF membranes (Millipore). Membranes were blocked with 5% non-fat milk in TBST, incubated with primary antibodies overnight at 4 °C, and then with HRP-conjugated secondary antibodies at room temperature for 1 h. Immunoreactive bands were detected using enhanced chemiluminescence (ECL) reagents (Thermo Fisher Scientific). A complete list of the antibodies used in this study is provided in Table 4.

Table 4.

Anbibody for western blots

Anbibody	Manufacturer	Catalogue number	Source of species	Dilution
SARM1	Abcam	ab309195	Rabbit	1:1000
COL2A1	Proteintech	28459-1-AP	Rabbit	1:1000
MMP13	Proteintech	18165-1-AP	Rabbit	1:1000
p21	Abcam	ab109199	Rabbit	1:1000
p16	Abcam	ab211542	Rabbit	1:1000
γ- H2A.X	Abcam	ab26350	Mouse	1:500
COXIV	Proteintech	11242-1-AP	Rabbit	1:10000
Caspase 3	Proteintech	19677-1-AP	Rabbit	1:1000
Cleaved Caspase 3	Proteintech	25128-1-AP	Rabbit	1:500
Caspase 9	CST	9504	Rabbit	1:1000
Cleaved Caspase 9	CST	9509	Rabbit	1:1000
OPA1	Proteintech	27733-1-AP	Rabbit	1:5000
MFN1	Proteintech	13798-1-AP	Rabbit	1:5000
MFN2	Proteintech	12186-1-AP	Rabbit	1:5000
p-Drp1 s616	CST	4494	Rabbit	1:500
p-Drp1 s637	CST	20,990	Rabbit	1:500
Drp1	CST	8570	Rabbit	1:1000
MFF	Proteintech	17090-1-AP	Rabbit	1:5000
FIS1	Proteintech	10956-1-AP	Rabbit	1:5000
p-Erk1/2	Proteintech	28733-1-AP	Rabbit	1:1000
Erk1/2	Proteintech	11257-1-AP	Rabbit	1:5000
CnA(PPP3CA)	Proteintech	13422-1-AP	Rabbit	1:5000
Flag	Proteintech	66008-4-Ig	Mouse	1:5000
Gadph	Abcam	ab8245	Rabbit	1:5000
β-actin	Proteintech	66009-1-Ig	Mouse	1:20000

4D label-free proteomics analysis of cultured chondrocytes

For detailed total protein extraction procedures from cultured chondrocytes, refer to the Western blot section above.

Equal amounts of protein from each sample were reduced with dithiothreitol, alkylated with iodoacetamide, and digested overnight with trypsin at 37 °C at an enzyme-to-protein ratio of 1:50. The resulting peptide mixtures were desalted with C18 spin columns, vacuum-dried, and reconstituted in 0.1% formic acid for subsequent mass spectrometry analysis. 4D label-free proteomic profiling was performed on a timsTOF platform coupled with nano-flow liquid chromatography. This technology adds a 4th dimension of ion mobility separation (based on peptide collision cross section) to boost proteome coverage, sensitivity and throughput. Protein identification and relative quantification were completed by comparing MS signal intensity of corresponding peptides across samples, without stable isotope internal standards.

Co-immunoprecipitation assay

ATDC5 cell lysates were prepared in Co-IP Lysis Buffer (Proteintech, Cat. # PR20037), centrifuged (12,000 × g, 15 min, 4 °C), and incubated with Anti-Flag Nanobody Magnetic Agarose Beads (AlpalifeBio, KTSM1361) at 4 °C for 8 h with rotation. Beads were washed three times, resuspended in SDS loading buffer, boiled (95 °C, 10 min), and analyzed by immunoblotting.

Seahorse assay

Mitochondrial oxygen consumption rate (OCR) was measured using the Seahorse XFe24 Analyzer (Agilent) and XF Cell Mito Stress Test Kit (Cat. # 103015-100). Chondrocytes were seeded in 24-well plates, cultured overnight, washed, and switched to assay medium (DMEM/F12 with 20 mM glucose, 2 mM glutamine, 1 mM sodium pyruvate). OCR was measured following sequential injections of oligomycin, FCCP, and rotenone/antimycin A.

Assessment of mitochondrial network fragmentation in chondrocytes

Mitochondrial network fragmentation was assessed using MitoTracker fluorescent probes. Briefly, chondrocytes were plated uniformly on 35 mm glass-bottom dishes and stained with either MitoTracker Deep Red or MitoTracker Green (100 nM) by incubating for 30 min at 37 °C under 5% CO₂ in the dark. Images were acquired using a confocal microscope (Zeiss, LSM 880).

Measurement of Intracellular Ca²⁺

1. Fluo-8 AM and Rhod-2 AM.

Fluo-8 AM / Rhod-2 AM: Cells were loaded with 4 µM Fluo-8 AM (Abcam) or Rhod-2 AM (AAT Bioquest) for 20 min at 37 °C, washed, and incubated in HBSS + 1% FBS (40 min, dark). Images were acquired using Zeiss LSM 880.

• 2.Fura-2 AM.

Cells in black 96-well plates were loaded with 1 µM Fura-2 AM (MCE) for 30 min, washed, incubated (40 min, dark), and analyzed on an EnVision 2105 reader (excitation: 340/380 nm; emission: 510 nm).

Measurement of NAD⁺ and cADPR Concentrations

Cells from 6-well plates were scraped, homogenized on ice, and centrifuged (12,000 × g, 15 min, 4 °C). Supernatants were used to measure:

1. NAD⁺: Enhanced NAD⁺/NADH Assay Kit (Beyotime, S0176S).

2. cADPR: Amplite^® Fluorimetric cADP-Ribose Assay Kit (AAT Bioquest, #20305).

Absorbance/fluorescence was read on an EnVision 2105 plate reader (PerkinElmer).

Statistical analyses

Data are presented as mean ± SD. Two-group comparisons were performed using Student’s t-test. For multi-group analyses, parametric one-way or two-way ANOVA followed by Fisher’s LSD post hoc test was applied for normally distributed data; for animal experiments with small sample sizes or non-normally distributed data, the nonparametric Kruskal‑Wallis H test followed by Dunn’s multiple comparisons test was used. Significance was set at p < 0.05. All statistical analyses were performed using GraphPad Prism 9.0.

Results

SARM1 is upregulated and activated in senescent chondrocytes

To identify key genes associated with both cellular senescence and mitochondrial dysfunction, we first performed RNA sequencing (RNA-seq) on primary chondrocytes isolated from C57BL/6J mice of different ages: 12-week-old (young adult), 48-week-old (middle-aged), and 96-week-old (aged) animals (Fig. 1A). Cellular senescence was evaluated by senescence-associated β-galactosidase (SA-β-gal) staining (Fig. 1B-C). Meanwhile, we established a replicative senescence model in mouse primary chondrocytes followed by RNA-seq (P6 vs. P0 controls; Fig. S1A, n = 3 per group). RNA-seq analysis revealed 4,865 upregulated and 4,828 downregulated genes in cartilage tissues from aged versus young mice (adjusted p-value < 0.05). Similarly, in replicatively senescent chondrocytes (P6 vs. P0), 3,389 genes were upregulated and 3,590 were downregulated (adjusted p-value < 0.05). Venn analysis was used to intersect our senescence-associated DEGs with the MitoCarta 3.0 database (Fig. 1D). This step yielded 28 overlapping candidates, among which SARM1 exhibited the greatest fold change (Fig. 1E), leading to its prioritization for further study.

Fig. 1.

SARM1 is highly expressed in chondrocytes from OA patients and aged mice.A Schematic overview of the RNA-seq workflow performed on chondrocytes isolated from 12-, 48-, and 96-week-old mice (n = 3 per group). B Representative images of SA-β-Gal staining in chondrocytes from mice of different ages. Scale bars: 50 μm; n = 3. C Quantification of SA-β-Gal–positive cells shown in (B). Six fields of view per group were analyzed. ***p < 0.001. D Venn diagram illustrating the overlap of differentially expressed genes (DEGs) identified in young vs. old and P0 vs. P6 comparisons, with mitochondria-interacting genes. E Heatmap showing expression levels of the overlapping genes identified in (D). F Representative Safranin O/Fast Green and immunohistochemical (IHC) staining for SARM1, p16, and p21 in human cartilage tissues from OA and non-OA donors. Scale bars: 500 μm; n = 6. G Quantitative RT-PCR analysis of SARM1, COL2A1, MMP13, CDKN1A, and CDKN2A mRNA levels in human cartilage tissues (OA vs. non-OA). n = 6; *p < 0.05, ***p < 0.001. H Representative Safranin O/Fast Green and IHC staining of SARM1 in mouse cartilage tissues following destabilization of the medial meniscus (DMM) or sham surgery. Scale bars: 50 μm; n = 6. I Representative immunoblots showing protein expression of SARM1, Col2a1, MMP13, p21, and p16 in primary mouse chondrocytes treated with or without 100 nM doxycycline (DOX) for 24 h. n = 3. J Subcellular fractionation followed by immunoblotting to detect SARM1 in cytosolic and mitochondrial fractions of chondrocytes. COX IV and GAPDH served as mitochondrial and cytosolic loading controls, respectively. n = 3. K-L Confocal microscopy images of Flag-SARM1 (green) and the mitochondrial marker Tomm20 (red) in chondrocytes overexpressing Flag-SARM1, with or without DOX treatment. Intensity line profiles along the white dashed lines are shown in (L). Scale bars: 5 μm; n = 3

We next examined SARM1 expression in human OA cartilage samples. IHC and qRT-PCR analyses revealed that OA patient cartilage (n = 6) expressed significantly higher levels of SARM1 and the senescence markers p16 and p21 compared to non-OA controls (n = 6) (Fig. 1F–G). In the murine DMM model, we similarly observed markedly increased SARM1 expression in OA-induced joints (n = 6), which paralleled the development of severe cartilage degradation when compared to sham controls (n = 6) (Fig. 1H).

The recruitment of SARM1 to mitochondria has been reported to be coupled with its activation [16, 26]. A cellular senescence model was generated in primary mouse chondrocytes using 100 nM doxorubicin (DOX) for 24 h. We confirmed the upregulation of SARM1 alongside key senescence markers at the protein level in this model (Fig. 1I, Fig. S1B). Furthermore, SARM1 localization from the cytosol to mitochondria was observed by western blot in senescence chondrocytes (Fig. 1J, Fig. S1C). For direct visualization, immunofluorescence analysis of chondrocytes expressing Flag-tagged SARM1 showed that DOX treatment significantly increased the mitochondrial localization of SARM1 relative to controls (Fig. 1K–L).

Taken together, our data establish that SARM1 is both upregulated and mitochondrially redistributed in senescent chondrocytes, events that are consistent with its activation and a direct role in driving mitochondrial dysfunction. This positions SARM1 as a potential nexus coordinating cellular senescence with metabolic failure in the pathogenesis of osteoarthritis.

SARM1 plays a role in Cellular senescence and mitochondrial dysfunction in primary chondrocytes

To define the functional role of SARM1 in cellular senescence, we therefore performed loss-of-function studies by transfecting senescent chondrocytes with SARM1 siRNA. SARM1 knockdown successfully reduced the expression of key senescence markers p16 and p21 (Fig. 2A, Fig. S1D-F). Flow cytometry also showed that silencing SARM1 partially rescued the DOX-induced cell cycle arrest (Fig. 2B). Furthermore, SARM1 depletion diminished senescence-associated β-galactosidase activity (measured by C12FDG fluorescence) and reduced DNA damage, as evidenced by fewer γ-H2AX foci (Fig. 2C, Fig. S1G-H). Finally, EdU assays revealed a higher proportion of proliferating cells in the SARM1-deficient group compared to controls (Fig. 2D–E).

Fig. 2.

SARM1 contributes to cellular senescence and mitochondrial dysfunction in mouse primary chondrocytes. A Immunoblots show SARM1, p16, and p21 levels in doxycycline (DOX)-induced senescent chondrocytes transfected with control or two SARM1-targeting siRNAs (si#1, si#2); untreated cells serve as negative control. n = 3. B Flow cytometry reveals that SARM1 knockdown reduces DOX-induced G1/S arrest. n = 3; **p < 0.01. C Representative images of senescence markers: (Top) Detection of SA-β-Gal activity via C12FDG fluorescence staining. (Bottom) Immunofluorescence staining of γH2A.X (red), a DNA damage marker, co-stained with phalloidin (green) to label F-actin. Nuclei were counterstained with DAPI. D–E EdU assay shows SARM1 siRNA partially restores proliferation in DOX-treated chondrocytes. DAPI labels nuclei. Quantification shows the percentage of EdU-positive cells. Scale bars: 500 μm; n = 6; ***p < 0.001. F JC-1 staining reflects mitochondrial membrane potential: red (aggregates, polarized), green (monomers, depolarized). SARM1 knockdown attenuates DOX-induced loss of red/green ratio. Scale bars: 50 μm; n = 3. G Oxygen consumption rate (OCR) measured via Seahorse XF24 Analyzer to assess mitochondrial respiration. H Bar graph summarizes basal, maximal, and spare respiratory capacity from (G). n = 3; *p < 0.05, **p < 0.01, ***p < 0.001. I TEM images show mitochondrial ultrastructure; SARM1 knockdown reduces DOX-induced swelling and cristae disruption. Scale bars: 500 nm; n = 3. J Mitochondrial networks labeled with MitoTracker Deep Red; insets show magnified regions. Scale bars: 5 μm; n = 6. K–L Morphometric analysis using MINA plugin: SARM1 knockdown increases branch number (K) and length (L). n = 6; *p < 0.05, ***p < 0.001

Based on the established mitochondrial role of SARM1, we next examined whether its knockdown could rescue the functional and structural deficits in mitochondria of senescent chondrocytes. Depletion of SARM1 restored the loss of mitochondrial membrane potential (ΔΨm, JC-1 assay, Fig. 2F). Seahorse metabolic analysis further demonstrated that SARM1 silencing significantly improved mitochondrial respiration, enhancing basal, maximal, and spare respiratory capacity (Fig. 2G–H). Ultrastructurally, transmission electron microscopy (TEM) showed that cells lacking SARM1 had better-preserved mitochondria, with neatly arranged cristae and much less swelling, in contrast to the severely disrupted appearance seen in control cells (Fig. 2I). Visualization of the mitochondrial network revealed that knocking down SARM1 shifted the organelles from a fragmented state to longer, more interconnected structures (Fig. 2J). This change was quantified by measuring the average number of branching points per network and the average branch length (Fig. 2K–L), pointing to a preference for mitochondrial fusion over fission.

Overall, these results demonstrate that increased SARM1 activity drives mitochondrial fragmentation and impairs function, while blocking or reducing SARM1 helps restore a healthy, interconnected mitochondrial network and improves respiratory performance.

Activated SARM1 mediates mitochondrial dysfunction and fragmentation in chondrocytes

SARM1 needs to be activated to carry out its NAD⁺ hydrolase (NADase) function. We hypothesized that a similar activation occurs in senescent chondrocytes, allowing SARM1 to perform its role there. To investigate this, we used the E642A point mutation in the TIR domain, a change known to completely eliminate SARM1’s enzymatic activity [21].

Primary mouse chondrocytes were transduced with lentiviruses expressing either wild-type SARM1 (SARM1 WT) or the catalytically inactive mutant (SARM1 E642A). After treating cells with DOX for 24 h, TUNEL staining showed markedly higher levels of apoptosis in cells expressing SARM1 WT than in those with the E642A mutant. This difference disappeared in the absence of DOX treatment (Fig. 3A, Fig. S2A). Flow cytometry with Annexin V/PI staining confirmed these findings (Fig. 3B, Fig. S2B). SARM1’s enzymatic activity appears to contribute to DOX-induced apoptosis. Since the intrinsic apoptotic pathway typically begins with mitochondrial damage, we next examined signs of its activation. Treatment with the SARM1 activator CZ-48 led to progressive, time-dependent cleavage of caspase-9 and caspase-3 (Fig. 3C, Fig. S2C-D), further supporting that SARM1 activation contributes to apoptotic signaling in this context. Immunofluorescence further showed prominent release of cytochrome c (Cyt c) from mitochondria in DOX-treated cells expressing SARM1 WT, whereas release was barely detectable in E642A-expressing cells. Importantly, this Cyt c release in WT cells was entirely prevented by the SARM1-specific inhibitor dehydronitrosonisoldipine (DeNSDP).

Fig. 3.

Activation of SARM1 is essential for mitochondrial dysfunction in senescent chondrocytes. A TUNEL staining of chondrocytes overexpressing wild-type (WT) or catalytically inactive E642A SARM1, with or without DOX treatment. Yellow arrowheads: TUNEL-positive apoptotic cells. Scale bars: 100 μm; n = 3. B Annexin V/PI flow cytometry shows apoptosis dynamics; WT SARM1 enhances DOX-induced apoptosis, attenuated by E642A mutant. n = 3. C Immunoblots show time-dependent cleavage of caspase-3 and caspase-9 in CZ-48-treated chondrocytes. n = 3. D Co-immunofluorescence shows cytochrome c (green) release from mitochondria (red, MitoTracker); overexpression of WT SARM1 promotes cytochrome c leakage after DOX, reduced by E642A mutant or dehydronitrosonisoldipine (4 μM). n = 3. E JC-1 staining reveals mitochondrial membrane potential: red (polarized), green (depolarized). WT SARM1 exacerbates DOX-induced depolarization, partially rescued by E642A or inhibitor. Scale bars: 50 μm; n = 3. F Mitochondrial networks labeled by MitoTracker; insets show magnified regions. WT SARM1 induces fragmentation, mitigated by E642A or inhibitor. Scale bars: 5 μm; n = 6. G–H MINA analysis shows SARM1 overexpression reduces branch number (G) and length (H), reversed by E642A or inhibitor. n = 6; *p < 0.05, ***p < 0.001. I Seahorse metabolic assay assessing mitochondrial respiration. J Quantification shows basal, maximal respiration, and spare capacity; WT SARM1 impairs mitochondrial function, effects blunted by E642A or dehydronitrosonisoldipine. n = 3; **p < 0.01, ***p < 0.001

We then asked whether SARM1’s enzymatic activity is necessary for the mitochondrial damage itself. JC-1 dye measurements demonstrated a sharp drop in mitochondrial membrane potential in DOX-stressed cells expressing active (WT) SARM1; this depolarization was largely avoided when cells expressed the inactive E642A mutant or were treated with DeNSDP (Fig. 3E). Seahorse extracellular flux analysis showed that only catalytically active SARM1 impaired mitochondrial respiration under these stress conditions (Fig. 3I–J). Finally, MitoTracker imaging revealed extensive mitochondrial fragmentation specifically in cells with active SARM1, while both the E642A mutant and DeNSDP treatment maintained elongated, interconnected mitochondrial networks (Fig. 3F–H).

Taken together, these experiments indicate that SARM1’s NADase activity is indispensable for causing the mitochondrial dysfunction and fragmentation that occur during chondrocyte stress responses.

SARM1 promotes mitochondrial fragmentation by regulating Drp1 phosphorylation and translocation

To figure out how SARM1 drives mitochondrial fragmentation, we started by screening key proteins involved in mitochondrial dynamics. Activating SARM1 didn’t change the overall levels of these proteins, but it did specifically affect the phosphorylation of Drp1 (dynamin-related protein 1). In particular, it increased phosphorylation at Ser616 and decreased it at Ser637 (as shown in Fig. 4A). As the primary regulator of mitochondrial fission, Drp1 sits in the cytosol and moves to the mitochondrial outer membrane to help pinch the membrane and split the organelle. Its activity is controlled by opposing phosphorylation events: Ser616 phosphorylation promotes this recruitment and oligomerization, while Ser637 phosphorylation inhibits it. When we activated SARM1, we saw much stronger overlap between Drp1 and mitochondria (Fig. 4B, Fig. S2E). Direct visualization confirmed robust translocation to mitochondrial in SARM1-overexpressing cells treated with CZ-48 (Fig. 4C-D). Importantly, loss-of-function experiments cemented this regulatory relationship: SARM1 knockdown reversed the phosphorylation pattern (Fig. 4E) and reduced Drp1’s mitochondrial localization (Fig. 4F-H, Fig. S2F). These data demonstrate that SARM1 promotes fragmentation by activating Drp1 and facilitating its recruitment to mitochondria.

Fig. 4.

SARM1 promotes mitochondrial fragmentation by activating Drp1. A Representative immunoblot images showing the levels of key mitochondrial fusion (OPA1, MFN1, MFN2) and fission (Drp1, MFF, FIS1) proteins, as well as Drp1 phosphorylation at Serine 616 and Serine 637, in primary chondrocytes overexpressing SARM1 treated with or without DOX or CZ-48. Relative phosphorylation levels of Drp1 at Ser616 and Ser637 (normalized to total Drp1) are shown on the right. n = 3. B Primary chondrocytes transduced with lentivirus overexpressing SARM1 were treated with or without DOX or CZ-48. Representative immunoblot images of Drp1 in Chondrocyte cytosolic and mitochondrial fractions. COX IV and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) were used as loading controls of mitochondrial and cytosolic fractions, respectively. n = 3. C-D Representative confocal fluorescence microscopy of Drp1(green) and mitochondria (red, labeled with MitoTracker) in SARM1 overexpressing chondrocytes treated with or without CZ-48 at indicated time points. Intensity traces (offset white lines) are plotted at Fig(D). n = 3, Scale bars: 5 μm. E Representative immunoblot images showing the levels of Drp1 phosphorylation at Serine 616 and Serine 637 in primary chondrocytes transfected with control or SARM1 siRNA and treated with or without DOX. Relative phosphorylation levels of Drp1 at Ser616 and Ser637 (normalized to total Drp1) are shown below. n = 3. F primary chondrocytes transfected with control or SARM1 siRNA were treated with or without DOX. Representative immunoblot images of Drp1 in Chondrocyte cytosolic and mitochondrial fractions. COX IV and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) were used as loading controls of mitochondrial and cytosolic fractions, respectively. n = 3. G-H Representative confocal fluorescence microscopy of Drp1(green) and mitochondria (red, labeled with MitoTracker) in primary chondrocytes transfected with control or SARM1 siRNA and treated with or without 100nM DOX for 24 h. Intensity traces (offset white lines) are plotted at Fig(H). n = 3, Scale bars: 5 μm

Activated SARM1 promotes Drp1 mitochondrial translocation by cADPR-Ca²⁺signaling

To determine whether SARM1 loss reshapes the expression of proteins in senescent chondrocytes, we performed quantitative proteomics. Knocking down SARM1 produced only very minor changes in the proteome (Fig. 5A), with just a handful of collagen subunits showing differences. This suggests that SARM1 does not drive massive protein-level reprogramming. So, we shifted focus to its known metabolic outputs.

Fig. 5.

Activated SARM1 promotes Drp1 mitochondrial translocation by cADPR-Ca²⁺ signaling. A Volcano plot showing transcriptomic changes in DOX-induced senescent chondrocytes transfected with control or SARM1 siRNA (siSARM1 vs. siCtrl); n = 5. B–C NAD⁺ and cADPR levels in chondrocytes after SARM1 knockdown with or without DOX treatment, normalized to protein; n = 6. D Fluo-8AM staining shows cytosolic Ca²⁺ levels; SARM1 knockdown reduces DOX-induced Ca²⁺ elevation; n = 3. E Co-staining with Rhod-2 AM (mitochondrial Ca²⁺, red) and MitoTracker (green) reveals that SARM1 silencing attenuates mitochondrial Ca²⁺ overload; DAPI labels nuclei; n = 3. F Cytosolic Ca²⁺ measured by Fura-2 AM in chondrocytes treated with cADPR (0–4 μM), with BAPTA-AM as chelator; n = 6. G Time course of cytosolic Ca²⁺ in SARM1-overexpressing (CZ-48-activated) chondrocytes, suppressed by 8-Br-cADPR; n = 6. H Drp1 translocation to mitochondria induced by cADPR is reduced by BAPTA-AM; COX IV and GAPDH are mitochondrial/cytosolic markers; n = 3. I cADPR increases Drp1 phosphorylation at Ser616 (pro-fission) and decreases Ser637 (anti-fission); quantified relative to total Drp1; n = 3. J MitoTracker staining shows cADPR-induced mitochondrial fragmentation, prevented by BAPTA-AM; insets: magnified views; scale bars: 5 μm; n = 6. K–L MINA analysis confirms reduced branch number (K) and length (L) with cADPR, reversed by Ca²⁺ chelation; ***p < 0.001; n = 6. M–N cADPR activates Erk1/2 and upregulates calcineurin A (CnA), effects blocked by BAPTA-AM or 8-Br-cADPR; n = 3.O Proposed model: SARM1 promotes cADPR production, elevating cytosolic and mitochondrial Ca²⁺, activating CnA/Erk pathways, driving Drp1-mediated fission and mitochondrial dysfunction

SARM1 knockdown didn’t bring NAD⁺ levels back up, but it did strongly cut down the buildup of cADPR (Fig. 5B–C). Since cADPR is a powerful signal that releases calcium, we looked at calcium handling next. Activating SARM1 raised calcium levels in both the cytosol and mitochondria, while knocking it down prevented this overload (Fig. 5D–E, Fig. S3A-D). Adding cADPR from outside the cell increased intracellular calcium in a dose-dependent way (Fig. 5F), and the SARM1 activator CZ-48 triggered a calcium rise that could be fully blocked by a cADPR antagonist (Fig. 5G)—showing the pathway is specific.

This calcium increase turned out to be functionally important. Treating cells with cADPR caused mitochondrial fragmentation (Fig. 5J–L) and boosted the activating phosphorylation of Drp1 at Ser616 along with its recruitment to mitochondria (Fig. 5H–I, Fig. S2G). All of these effects disappeared when we used the calcium chelator BAPTA-AM to buffer the calcium surge. On the molecular side, we saw increased levels of the calmodulin-binding catalytic subunit A (CnA) and phosphorylated ERK1/2 (p-ERK1/2), positioning them downstream of the cADPR/calcium signal and upstream of Drp1 (Fig. 5M-N, Fig. S2H-K). Finally, adding cADPR by itself was enough to trigger senescence markers like p16 and p21, whereas blocking cADPR signaling stopped SARM1 from driving senescence (Fig. S3E–J).

Putting it all together, the data outline a straightforward pathway in these cells: SARM1 activation leads to cADPR accumulation → calcium overload → activation of CnA and p-ERK1/2 → Drp1-dependent mitochondrial fission → chondrocyte senescence.

SARM1-cADPR-Ca²⁺signaling enhances mitochondrial fission by promoting the interaction between Drp1 and FIS1

Drp1 carries out mitochondrial fission by interacting with specific receptor proteins on the mitochondrial outer membrane, including Mid49/Mid51 (also called MIEF1/2), MFF, and FIS1. To investigate which of these receptors Drp1 binds to under SARM1 activation, we created stable ATDC5 cell lines that stably express Flag-tagged versions of Mid51, MFF, or FIS1. We then performed co-immunoprecipitation experiments to check for Drp1 binding to each receptor.

The results showed that activating SARM1 (using DOX) selectively strengthened the interaction between Drp1 and FIS1, while binding to Mid51 or MFF remained unchanged (Fig. 6A-C). This enhanced Drp1–FIS1 association was lost when SARM1 was knocked down. Exogenous cADPR externally increased Drp1 binding to FIS1 in a dose-dependent way, and this effect was completely blocked by the calcium chelator BAPTA-AM—pointing again to the cADPR–calcium pathway as the key driver (Fig. 6D-F). Immunofluorescence in primary chondrocytes further confirmed that inducing SARM1 with DOX led to stronger colocalization of Drp1 with FIS1 on mitochondria, but knocking down SARM1 reduced this overlap (Fig. 6G-H). Thus, SARM1 fine-tunes fission by specifically promoting cADPR/Ca²⁺-dependent recruitment of Drp1 to the FIS1 receptor.

Fig. 6.

SARM1-cADPR-Ca2 + signaling enhances mitochondrial fission by promoting the interaction between Drp1 and Fis1. A-C ATDC5 cell lines stably overexpressing Flag-MiD51, Flag-MFF, and Flag-Fis1 were transfected with control or SARM1 siRNA and treated with or without 100 nM DOX for 24 h. Immunoprecipitation analysis of the MiD51-Drp1 interaction 、MFF-Drp1 interaction and the Fis1-Drp1 interaction. D-G ATDC5 cell lines stably overexpressing Flag-MiD51, Flag-MFF, and Flag-Fis1 were treated with cADPR (0, 2, and 4 µM) or 4 µM cADPR followed by BAPTA-AM. Immunoprecipitation analysis of the MiD51-Drp1 interaction 、MFF-Drp1 interaction and the Fis1-Drp1 interaction. H-I Representative confocal fluorescence microscopy of Drp1(green) and Fis1 (red) in primary chondrocytes transfected with control or SARM1 siRNA and treated with or without 100nM DOX for 24 h. Intensity traces (offset white lines) are plotted at Fig(I). n = 3, Scale bars: 5 μm

Targeting SARM1 and cADPR attenuates chondrocyte senescence and OA progression

To minimize potential biases from monolayer culture systems, we validated our findings in mouse femoral head cartilage explants, which preserve the native extracellular matrix. In this model, DOX treatment increased Drp1 phosphorylation at Ser616 and decreased it at Ser637, confirming enhanced mitochondrial fission. These changes were attenuated by both the SARM1 inhibitor DeNSDP and the cADPR antagonist 8-Br-cADPR (Fig. 7A, Fig. S4B-C).

Fig. 7.

Targeting SARM1 and cADPR attenuates chondrocyte senescence and OA progression.A Representative immunofluorescence images showing the expression of p-Drp1 (Ser616) and p-Drp1 (Ser637) in primary chondrocytes treated for 24 h with 100 nM DOX alone, or 100 nM DOX co-treated with dehydronitrosonisoldipine or 8-Br-cADPR. Leftmost lane: NC (Negative Control). Nuclei were visualized with DAPI staining. n = 6, Scale bars: 200 μm. B-C Mouse femoral head explants were harvested and cultured as described in Methods, and then treated as described in Fig(A). Z-stack images were acquired at 2-µm intervals through a tissue depth of 200–300 μm using a ZEISS LSM 880 confocal microscope, followed by 3D reconstruction. Representative images of Fluo-8 AM staining showing intracellular ca.2+ levels in mouse femoral head cartilage (Fig B). Representative images of JC-1 staining showing changes in mitochondrial membrane potential (ΔΨm) (Fig C). Leftmost lane: NC (Negative Control). n = 3, Scale bars: 200 μm. D overview of the experimental setup with injections of aaV5- control or aaV5- SARM1 shrna. E Representative images of Safranin O/Fast Green staining in mouse knee joints subjected to the following treatments: sham operation, DMM surgery with AAV5-control or AAV5-shSARM1 treatment, or intra-articular injection of PBS or 8-Br-Cadpr. The inset is shown in the lower right corner. n = 6 mice per group. Scale bar:50 μm. The corresponding OARSI scores (0–6) are shown in (Fig G). Statistical analysis was performed using Kruskal‑Wallis H test followed by Dunn’s multiple comparisons test. F Representative H&E staining of knee joints from mice treated as described previously (see Fig E). The inset is shown in the lower right corner. n = 6 mice per group. Scale bar:50 μm. The corresponding synovitis grade (0 to 3) are shown in (Fig H) (same statistical test as Fig G). I Representative IHC staining for SARM1, Col2a1, MMP13, p16, and p21 in knee joint cartilage from mice treated as described previously (see Fig E)

We next assessed intracellular Ca²⁺ and mitochondrial membrane potential (ΔΨm) within the explants. Using Fluo-8 AM and JC-1 probes that efficiently penetrated the tissue, we acquired z-stack images at 2-µm intervals (200–300 μm depth) on a Zeiss LSM 880 confocal microscope for 3D reconstruction. DOX treatment elevated intracellular Ca²⁺ levels (Fig. 7B) and increased JC-1 green fluorescence (indicating ΔΨm loss) (Fig. 7C). Both DeNSDP and 8-Br-cADPR significantly reduced Ca²⁺ overload and restored the JC-1 red/green fluorescence ratio, demonstrating preserved ΔΨm.

Finally, we employed a DMM-induced OA model in C57BL/6J mice (6–8 weeks old). Mice received weekly intra-articular (IA) injections of AAV5-shSarm1 or control vector for four weeks post-surgery (viral transduction was confirmed by IVIS imaging, Fig.S4A), with tissues harvested at eight weeks. A parallel cohort received weekly IA injections of 8-Br-cADPR or PBS vehicle.

DMM surgery induced severe cartilage erosion (Fig. 7E, G) and synovitis (Fig. 7F, H). Both AAV5-shSarm1 and 8-Br-cADPR treatments markedly preserved cartilage structure and reduced synovitis. IHC analysis showed that both interventions suppressed senescence markers (p16, p21), enhanced the anabolic marker COL2A1, and reduced the catabolic enzyme MMP13 in cartilage (Fig. 7I, Fig. S4D-H).

Discussion

Our findings establish a pathogenic pathway in which SARM1 upregulation drives cADPR-dependent Ca²⁺ overload, promotes Drp1-mediated mitochondrial fission, and ultimately leads to chondrocyte senescence and OA. Inhibition of this axis, both in explants and in a mouse OA model, attenuated disease phenotypes, pointing to mitochondrial stabilization as a viable therapeutic strategy.

SARM1 can flip between inactive and active forms depending on metabolic stress, connecting different cellular processes to cell death and playing a role in many diseases [17–19, 21, 27]. A recent study by Lina Wang et al. further showed that SARM1 can also detect cytosolic double-stranded DNA (dsDNA) and trigger cell death [28]. Since senescent cells often accumulate cytosolic dsDNA, SARM1 probably acts through several different mechanisms during senescence. More work is needed to fully understand its additional functions.

Although SARM1’s harmful effects are often blamed on NAD⁺ breakdown, emerging data highlight separate roles for its product cADPR [29]. As a second messenger that triggers calcium-induced calcium release, cADPR has been well studied across systems—from its original discovery in sea urchin eggs to many mammalian cell types, including T cells, heart muscle cells, and neurons [30–33]. Our work now identifies cADPR as a key driver of chondrocyte senescence. When we activate SARM1 (using CZ-48), we see calcium overload, mitochondrial fragmentation, and higher levels of senescence markers; all of these are strongly reduced by the specific cADPR blocker 8-Br-cADPR. On the flip side, adding cADPR directly to chondrocytes causes calcium release and Drp1-dependent mitochondrial fragmentation. Together, these findings describe a new pathway that does not depend on NAD⁺ loss: SARM1 produces excess cADPR, which causes abnormal calcium signaling, mitochondrial fission, and chondrocyte senescence.

We identified FIS1—a protein on the mitochondrial outer membrane—as the main receptor that recruits Drp1 to carry out fission after SARM1 activation. While the Drp1–FIS1 interaction is already known to drive mitochondrial division, its importance in aging-related diseases is becoming clearer. Kleele et al. showed that mitochondrial fission happens in two ways: one through MFF at the midzone (for normal turnover) and another through FIS1 at the periphery (often tied to stress, damage, and pathological conditions like mitophagy) [34]. This peripheral, FIS1-driven fission is triggered by problems such as loss of mitochondrial membrane potential (ΔΨm), diminished proton motive force, increased reactive oxygen species (ROS), and notably, by intracellular Ca²⁺ overload. In diabetic heart disease, Wu et al. found that calcium overload activates Drp1 through either ERK1/2-dependent Ser616 phosphorylation or calcineurin (CnA)-mediated Ser637 dephosphorylation, leading to fragmentation [35]. Similarly, Ansari et al. reported that ERK1/2-driven Drp1 activation disrupts mitochondrial balance and speeds up cell death in OA chondrocytes [10] —a result that aligns closely with what we observed.

OA is tightly coupled to chondrocyte mitochondrial dysfunction, a long-standing focus of intensive mechanistic research. Cumulative studies have identified several core drivers of mitochondrial impairment in OA chondrocytes: reduced activity of the aging-associated mitochondrial deacetylase SIRT3 disrupts mitochondrial homeostasis by altering the acetylation status of cytochrome c oxidase subunit 4 isoform 2 (COX4I2) [36]; sustained IL-1β stimulation in the OA microenvironment inhibits Parkin-dependent mitophagy, leading to accumulation of damaged mitochondria and subsequent chondrocyte apoptosis [37]; and TRPV4-mediated calcium overload drives hyperactivation of Drp1, triggering mitochondrial network fragmentation and irreversible organelle damage [38]. While multiple small molecules targeting these pathways (including dihydromyricetin and the clinically approved metformin) exert robust preclinical chondroprotective effects in OA models, their clinical translation is severely hampered by inherent adverse effect risks, given that their targets are core proteins governing essential mitochondrial and cellular physiology [39, 40].

Here, we advance the mechanistic understanding of OA-related mitochondrial dysfunction via a novel SARM1-mediated pathway. Specifically, SARM1 drives mitochondrial fragmentation through its enzymatic product cyclic ADP-ribose (cADPR), which triggers calcium overload to promote Drp1 phosphorylation and subsequent Drp1-Fis1 interaction, in a manner independent of global cellular NAD+ depletion. Critically, unlike the aforementioned proteins (including SIRT3 and the PINK1-Parkin mitophagy axis), SARM1 is a stress-responsive executioner protein that is specifically activated only under cellular stress. Thus, SARM1 inhibition exerts negligible off-target effects on normal resting cells, rendering it a promising, high-safety disease-modifying therapeutic target for OA [41].

This study has some limitations. We did not have access to SARM1-knockout mice, so we could not fully evaluate how complete SARM1 loss affects chondrocyte senescence and OA development in vivo. Also, while the destabilization of the medial meniscus (DMM) mouse model is widely used and informative, differences in joint anatomy, biomechanics, and walking patterns compared to humans mean it does not perfectly mimic human OA. Moving forward, using conditional SARM1-knockout mice and larger animal models that more closely resemble human disease will help confirm these findings and move them closer to clinical use.

In summary, this work shows how the SARM1–cADPR pathway throws off intracellular calcium balance, causing mitochondrial fragmentation and chondrocyte senescence. We further show the therapeutic relevance of this pathway, as inhibiting either SARM1 or cADPR signaling effectively mitigates mitochondrial dysfunction and senescence in mouse cartilage explants and in a surgical OA model. These results collectively identify SARM1 as a promising target for OA therapy.

Authors’ contributions

YG, XZ, BX conceived and designed the study. YG and XZ were responsible for data acquisition, analysis, and manuscript drafting. BW, RP, CL contributed to data collection and analysis. YG wrote the manuscript. BX, XZ, and HL reviewed, edited, and provided final approval of the manuscript. All authors reviewed the manuscript and provided critical feedback.

Funding

This work was supported by the Guangdong Basic and Applied Basic Research Foundation (No. 2022A1515010713, No. 2023A1515110860).

Data availability

The datasets used and analysed during the current study are available from the corresponding author on reasonable request.

Declarations

Ethics approval and content to participate

This study was performed in line with the principles of the Declaration of Helsinki. The use of discarded, deidentified human cartilage was approved by Medical Ethics Committee of the Fifth Affiliated Hospital of Sun Yat-sen University (Permit Number: No.2025K198-1), with a waiver of informed consent. All animal procedures were approved by the Institutional Animal Care and Use Committee of The Fifth Affiliated Hospital of Sun Yat-sen University (Permit Number: No. 00484) and in accordance with the Guide for the Care and Use of Laboratory Animals.

Consent for publication

Not applicable.

Competing interests

The authors declare no competing interests.