METTL3-mediated fibroblast-like synoviocytes senescence promotes temporomandibular joint osteoarthritis progression

Kun Tian; Qin Du; Jun Guo; Wen Liao; Xianrui Yang; Shuang Lai; Juan Liao; Yandong Mu; Li Xiao

doi:10.1038/s42003-026-09773-x

. 2026 Mar 3;9:510. doi: 10.1038/s42003-026-09773-x

METTL3-mediated fibroblast-like synoviocytes senescence promotes temporomandibular joint osteoarthritis progression

Kun Tian ^1,², Qin Du ¹, Jun Guo ¹, Wen Liao ³, Xianrui Yang ⁴, Shuang Lai ¹, Juan Liao ^1,^✉, Yandong Mu ^1,^2,^✉, Li Xiao ^1,^2,^✉

PMCID: PMC13066444 PMID: 41776281

Abstract

Temporomandibular joint osteoarthritis (TMJOA) is a disease that affects the TMJ and is associated with degenerative changes in the articular cartilage. Fibroblast-like synoviocytes (FLSs) have been found to contribute to osteoarthritis. Here, we aim to investigate the role of METTL3-mediated FLS senescence in the TMJOA process. TMJOA model rats were successfully generated, displaying typical structural and inflammatory alterations, and primary FLSs were isolated from monosodium iodoacetate (MIA)-induced TMJOA rats; these FLSs were accompanied by increased senescence, attenuated mitophagy, and upregulated METTL3. FLSs from TMJOA rats also induced cartilage degradation. Mechanistically, METTL3 silencing can increase PINK1 expression by increasing its RNA stability through m⁶A modification. In addition, we found that METTL3 silencing could delay cellular senescence and promote mitophagy by upregulating PINK1 in bleomycin (BLM)-induced hFLSs. Senescent FLSs can also accelerate pathological progression and cartilage degradation in Sprague–Dawley (SD) rats. This study revealed that METTL3 silencing could suppress the senescence of FLSs and promote mitophagy by mediating m⁶A modification to upregulate PINK1 during TMJOA progression, which might provide a theoretical basis for TMJOA therapy.

Subject terms: Epigenetics, Senescence, Mitophagy, Oral diseases, Cell signalling

METTL3 can regulate mitophagy and fibroblast-like synoviocyte senescence through m6A modification, affecting the stability of PINK1 mRNA and thereby promoting the progression of TMJOA

Introduction

The temporomandibular joint (TMJ) is a unique synovial joint that enables essential functions such as mastication and facial expression through its complex anatomical structure and bilateral coordination¹. Temporomandibular joint osteoarthritis (TMJOA) is a common degenerative condition, with a prevalence affecting 8% to 16% of the population worldwide². Its onset is often linked to factors such as trauma, articular disc displacement, joint overload, and developmental abnormalities³. Additionally, advanced age, occlusal disorders, joint injuries, and certain systemic conditions contribute to its progression⁴. Current treatment strategies primarily include symptomatic therapies and surgical interventions. While drug-based symptom management can provide relief, it frequently leads to adverse effects. Surgical options, on the other hand, are invasive and carry substantial risks^2,5. Despite growing research, the underlying mechanisms driving TMJOA remain elusive, underscoring the need for further exploration.

Fibroblast-like synoviocytes (FLSs) are key cellular components of synovial tissue and play a central role in maintaining joint homeostasis as well as driving pathological changes during TMJOA progression⁶. Increasing evidence indicates that aberrant activation of FLSs contributes to TMJOA development through enhanced inflammatory responses, excessive secretion of cartilage-degrading enzymes, and heightened sensitivity to mechanical stress^7,8. Within the TMJ microenvironment, activated FLSs secrete a broad spectrum of cytokines and chemokines that amplify local inflammation and accelerate cartilage destruction⁹. In parallel, aging-related cellular alterations further aggravate joint degeneration. Senescence of joint-resident cells not only impairs normal tissue function but also disrupts intracellular quality-control mechanisms, thereby promoting disease progression¹⁰.

Notably, cellular senescence is closely associated with mitochondrial dysfunction, particularly impaired mitophagy, which is essential for maintaining mitochondrial integrity. Senescent cells exhibit reduced mitophagic activity, leading to the accumulation of dysfunctional mitochondria and further reinforcement of the senescent phenotype¹¹. In TMJOA, insufficient mitophagy has been linked to increased cell death and cartilage degradation, highlighting its protective role in joint tissues¹². However, the molecular mechanisms that coordinate mitochondrial quality control and senescence in FLSs during TMJOA progression remain largely unclear.

RNA methylation has emerged as an important regulatory mechanism governing gene expression¹³. Among various RNA modifications, N6-methyladenosine (m6A) is the most abundant internal modification in eukaryotic mRNAs and influences RNA transcription, processing, translation, and stability^14,15. Accumulating studies suggest that dysregulated m6A modification is involved in osteoarthritis progression^16,17. METTL3, a core component of the m6A methyltransferase complex, catalyzes m6A deposition on target RNAs and has been implicated in the regulation of cellular senescence and mitochondrial function¹⁸. Previous work has shown that inhibition of m6A modification alleviates chondrocyte senescence and attenuates TMJOA progression in animal models¹⁰. Based on these observations, we hypothesized that METTL3-mediated m6A modification may regulate mitophagy and senescence in FLSs, thereby contributing to TMJOA development. In the present study, we investigated the role of senescent FLSs in TMJOA progression and further explored whether METTL3 modulates mitophagy and senescence in FLSs through m6A-dependent regulation of PINK1 mRNA.

Results

MIA induces pathological progression in the TMJ of Sprague–Dawley (SD) rats

To investigate the possible mechanism of TMJOA progression, this study first constructed a rat model of TMJOA with MIA (Fig. 1A). Micro-CT images revealed that, compared with that in the sham group, the trabecular bone microstructure of the subchondral bone deteriorated in the TMJOA group (Fig. 1B). The statistics revealed that BV/TV and Tb.Th were lower and that Tb.Sp was greater in the TMJOA model group than in the sham group (Fig. 1C–F). The data also revealed that the percentage of HWT decreased most significantly on the first day of MIA treatment and gradually increased with time; the degree of pain in the rats significantly differed during pain assessment (Fig. 1G). The pathological staining results revealed cartilage degeneration with a proteoglycan area and chondrocyte reduction in the TMJOA group, and the OARSI score was also greater in the TMJOA group than in the sham group (Fig. 1H–K). Moreover, the IHC results revealed that MIA induction increased the number of MMP-13⁺ cells and decreased the number of Col2A1⁺ cells in the TMJs of SD rats (Fig. 1L–N). Furthermore, inflammatory cytokine expression in synovial tissues was examined. The mRNA levels of IL-1β, TNF-α, and IL-6 were higher in the TMJOA group than in the sham group (Fig. 1O–Q). ELISA analysis also showed increased protein levels of IL-1β, TNF-α, and IL-6 in synovial tissue homogenates from MIA-treated rats (Fig. 1R–T). These findings indicate that MIA injection induces structural alterations of the TMJ accompanied by synovial inflammatory changes in SD rats.

Insufficient mitophagy and increased METTL3 in synovial tissues of TMJOA rats

Compared with those in the sham group, the levels of senescence-related marker p16INK4a were markedly increased, while vimentin was significantly decreased in MIA-induced TMJOA model rats (Supplementary Figs. 1A–E and 23). We also observed that the protein level of p62 in synovial tissue mitochondria was greater in the TMJOA model group than in the sham group (Supplementary Figs. 1F, G and 24). The results of mitochondrial-related protein detection revealed that Mfn2 expression was notably decreased and that Drp1 expression was elevated in the synovial tissue of the TMJOA group, suggesting mitochondrial dysfunction (Supplementary Figs. 1H–J, 25). Besides, double IF staining revealed that mitophagy-related proteins (LC3B and TOMM20) were decreased in the synovial tissue of MIA-induced TMJOA rats (Supplementary Fig. 1K). The protein level of METTL3 was elevated in the TMJOA rats (Supplementary Figs. 1L and M, 26). We further measured the content of m⁶A in synovial tissue and detected increased levels of m⁶A in MIA-induced TMJOA rats (Supplementary Fig. 1N). β-Galactosidase activity was also greater in the TMJOA group than in the sham group (Supplementary Fig. 1O).Through correlation analysis, the OARSI score was positively correlated with the proportion of p16^INK4a-positive FLSs (r = 0.8978, p = 0.0025), METTL3 expression (r = 0.8756, p = 0.0044), and β-GAL activity (r = 0.8939, p = 0.0028), while it was negatively correlated with LC3B expression (r = −0.6234, p = 0.0025). In addition, TOMM20 expression was positively correlated with LC3B expression (r = 0.8053, p = 0.0159) (Supplementary Fig. 1P–T). Overall, cellular senescence, mitophagy, and METTL3 are relevant to TMJOA in rats.

Abnormal cell senescence, mitophagy, and METTL3 expression in primary FLSs

Senescence-related proteins were first examined, and p21, p16INK4a, and DcR2 expression in primary FLSs from TMJOA rats was greater than that in primary FLSs from normal rats (Fig. 2A–D and Supplementary Fig. 19). P62 expression in primary FLSs was also elevated in the TMJOA group compared with the normal group (Fig. 2E, F, and Supplementary Fig. 20). In the mitochondrial protein assay, Drp1 was upregulated, and Mfn2 was downregulated in TMJOA-FLSs compared with normal FLSs (Fig. 2G–I and Supplementary Fig. 21). Notably, the accumulation of p62 together with increased Drp1 and reduced Mfn2 reflects an imbalance in mitochondrial dynamics and insufficient mitochondrial clearance, rather than enhanced completion of mitophagy. Besides, METTL3 expression, m⁶A levels, and β-galactosidase activity were prominently increased in TMJOA-FLSs (Fig. 2J–N and Supplementary Fig. 22). TMRM staining revealed that the mitochondrial membrane potential (MMP) decreased in TMJOA-FLSs (Fig. 2O). Compared with those in normal FLSs, the expression of LC3B and TOMM20 was reduced (Fig. 2Q), accompanied by the production of many mitochondrial fragments in TMJOA-FLSs (Fig. 2R) and enhanced secretion of IL-1β and TNF-α (Supplementary Fig. 2). These data support the importance of cellular senescence, mitophagy, and METTL3 in TMJOA in vitro.

Fig. 2 — Primary FLSs were isolated from the synovial tissues of normal and MIA-induced TMJOA rats. A–D Western blot assays were used to assess the expression of p21, p16INK4a, and DcR2. E, F Western blotting analysis of P62 expression in the mitochondrial fraction. TOMM20 was used as a mitochondrial loading control, and β-actin was included as a cytosolic marker to assess the purity of mitochondrial isolation. G–I Western blotting analysis of Drp1 and Mfn2 expression. J, K Western blotting analysis of METTL3 expression. L The m⁶A level was verified via an ELISA kit. M, N SA-β-Gal staining revealed the senescence of FLSs, and the activity of β-Gal was determined. O, P The MMP was confirmed by TMRM staining. Q LC3B and TOMM20 expression was analyzed via double IF staining. R TEM was used to observe the mitochondrial structure. *P < 0.05, **P < 0.01, ***P < 0.001.

FLSs from TMJOA rats induce cartilage degradation

Next, human articular chondrocytes were cocultured with FLSs isolated from normal and TMJOA rats using a Transwell chamber system to establish an in vitro model that mimics the OA microenvironment (Supplementary Fig. 3A). By examining cartilage degradation-related proteins, we found that ADAMTS5 and MMP13 were upregulated and that Collagen II was downregulated in human articular chondrocytes after coculture with TMJOA-FLSs (Supplementary Figs. 3B–F, 4 and 27). These findings suggested that FLSs from TMJOA rats promoted cartilage degradation.

Bleomycin (BLM) accelerates cell senescence, reduces mitophagy, and upregulates METTL3 in hFLSs

BLM markedly increased β-galactosidase production in hFLSs (Supplementary Fig. 5A, B), as did p16^INK4a, p21, and DcR2 expression (Supplementary Fig. 5C–F and Supplementary Fig. 28), and increased the secretion of IL-1β and TNF-α (Supplementary Fig. 6). BLM treatment upregulated P62 in the mitochondria and downregulated Drp1 and Mfn2 in hFLSs (Supplementary Figs. 5G, H and 29, 7 and 32). Moreover, BLM treatment upregulated METTL3 (Supplementary Figs. 5I and J and 30), increased the m⁶A level (Supplementary Fig. 5K), and reduced the MMP (Supplementary Fig. 5L and M) in hFLSs. Besides, BLM reduced LC3B and TOMM20 expression (Supplementary Fig. 5N) and increased the number of mitochondrial fragments (Supplementary Fig. 5O) in hFLSs. Through the detection of mitophagy-related proteins, we found that BLM downregulated Parkin and PINK1 but did not affect DFCP1, WIPI1, optineurin, NDP52, or ULK1 expression in hFLSs (Supplementary Figs. 5P–U, 31, 8, 33), suggesting that Parkin and PINK1 are key factors involved in attenuating mitophagy. Thus, we demonstrated that cell senescence and METTL3 expression were increased and that mitophagy was decreased in BLM-induced senescent hFLSs.

Torin1 inhibits cell senescence and promotes mitophagy in BLM-induced senescent hFLSs

The data indicated that BLM stimulation enhanced cellular senescence and that the mitophagy agonist Torin1 weakened the BLM-induced increase in hFLSs (Supplementary Fig. 9A, B). Torin1 reduced the number of mitochondrial fragments (Supplementary Fig. 9C), decreased p62 expression in the mitochondria (Supplementary Figs. 9D, E and 34), downregulated Drp1, upregulated Mfn2 in BLM-induced hFLSs (Supplementary Figs. 9F–H and 35), and suppressed BLM-induced IL-1β and TNF-α secretion (Supplementary Fig. 10). Moreover, Torin1 downregulated p16I^NK4a, p21, and DcR2 in BLM-induced hFLSs (Supplementary Figs. 9I–L and 36). The decreases in LC3B, TOMM20, and MMP mediated by BLM could also be partially reversed by Torin1 in hFLSs (Supplementary Fig. 9M–O). Overall, insufficient mitophagy is essential in senescent hFLSs.

METTL3 silencing weakens cell senescence and promotes mitophagy in BLM-treated hFLSs

The results indicated that METTL3 knockdown notably reduced METTL3 expression in BLM-induced hFLSs, suggesting the high transfection efficiency of sh-METTL3 in BLM-induced hFLSs (Fig. 3A, 3B and Supplementary Fig. 37). METTL3 knockdown also partially reversed the BLM-mediated increase in β-galactosidase activity in hFLSs (Fig.3C, D). Besides, METTL3 knockdown notably reduced the number of mitochondrial fragments (Fig. 3E), increased the MMP (Fig. 3F, G), and downregulated p62 in the mitochondria (Fig. 3H, I and Supplementary Fig. 38) of hFLSs. METTL3 knockdown also downregulated Drp1 and upregulated Mfn2 in BLM-induced hFLSs (Supplementary Figs. 11 and 41). Additionally, METTL3 knockdown downregulated p16^INK4a, p21, and DcR2 in BLM-induced hFLSs (Fig. 3J–M and Supplementary Fig. 39) and suppressed BLM-induced IL-1β and TNF-α secretion (Supplementary Fig. 12). Our results revealed that METTL3 silencing upregulated PINK1 and Parkin but did not affect the expression of optineurin, DFCP1, WIPI1, NDP52, or ULK1 in BLM-induced hFLSs (Fig. 3N–P, Supplementary Figs. 13 and 40). These results suggest that METTL3 may affect mitophagy mainly by regulating PINK1 and Parkin. Besides, METTL3 silencing elevated LC3B and TOMM20 expression in BLM-induced hFLSs (Fig. 3Q). These data revealed that METTL3 knockdown inhibited cell senescence and accelerated mitophagy in senescent hFLSs.

Fig. 3 — BLM-treated hFLSs were transfected with shMETTL3-1 (IME4-1) or shMETTL3-2 (IME4-2). A, B The transfection efficiency of shMETTL3-1 and shMETTL3-2 was verified by Western blotting. C, D Verification of cellular senescence with a β-GAL kit. E TEM for observing mitochondrial autophagosomes. F, G TMRM staining. H, I Western blotting data of P62 expression in the extracted mitochondria. J–P Western blotting analysis of p16^INK4a, p21, DcR2, PINK1, and Parkin. Q Double IF staining results for LC3B and TOMM20. *** P < 0.001 vs. the CTRL group; ## P < 0.01, ### P < 0.001 vs. the BLM+shCTRL group.

METTL3 silencing stabilizes PINK1 mRNA via m⁶A modification

The results revealed that the level of PINK1 m⁶A mRNA was increased in the synovial tissues of the MIA-induced TMJOA rats and in the primary FLSs from the MIA-induced TMJOA rats (Fig. 4A, B). Besides, the PINK1 m⁶A mRNA level was also increased in BLM-treated hFLSs relative to hFLSs (Fig. 4C), which was also decreased by METTL3 knockdown (Fig. 4D). Based on publicly available m⁶A modification data from m⁶A Atlas, several putative m⁶A sites were identified within the coding region of PINK1 mRNA. The sites at positions 2028, 2174, 2483, and 2074 were selected for subsequent analysis. The corresponding methylation profiles are shown in Fig. 4E. Through m⁶A RIP validation, we found that m⁶A was notably enriched at the 2028, 2074, and 2174 sites and that m⁶A enrichment at the 2028, 2174, 2483, and 2074 sites was increased in BLM-induced hFLSs relative to hFLSs, which could also be partially reversed by METTL3 knockdown (Fig. 4F–I). After treatment with 2.5 μM actinomycin D, the mRNA level of PINK1 gradually decreased as the processing time increased, and BLM decreased PINK1 expression in hFLSs. METTL3 (IME4) knockdown increased PINK1 expression, which was reduced in BLM-induced hFLSs. Consistently, METTL3 knockdown prolonged the estimated half-life of PINK1 mRNA (Fig. 4J). These data indicate that METTL3 knockdown reduces PINK1 m⁶A modification and enhances its mRNA stability.

PINK1 overexpression affects cell senescence and mitophagy in BLM-induced hFLSs

As presented in Supplementary Fig. 14A and B and Supplementary Fig. 42, BLM treatment reduced PINK1 expression in hFLSs, and PINK1 expression was markedly elevated after OE-PINK1 transfection in BLM-treated hFLSs, indicating that PINK1 was successfully overexpressed. PINK1 overexpression attenuated β-galactosidase activity, reduced the number of mitochondrial fragments, and increased the MMP in BLM-treated hFLSs (Supplementary Fig. 14C–14G). Moreover, PINK1 overexpression reduced p62 expression in the mitochondria of BLM-induced hFLSs, likely reflecting enhanced mitophagy flux and clearance of damaged mitochondria rather than impaired p62 recruitment (Supplementary Figs. 14H and I and 43). With respect to mitochondrial function, PINK1 overexpression downregulated Drp1 and upregulated Mfn2 in BLM-induced hFLSs (Supplementary Figs. 14J, L and 44). In mitophagy, PINK1 overexpression downregulated p16^INK4a, p21, and DcR2 in BLM-induced hFLSs (Supplementary Figs. 14M and 45, 15) and suppressed BLM-induced IL-1β and TNF-α secretion (Supplementary Fig. 16). The IF results revealed that PINK1 overexpression increased the fluorescence intensity of LC3B and TOMM20 in BLM-induced hFLSs (Supplementary Fig. 14N). Thus, PINK1 overexpression can also induce mitophagy in senescent hFLSs.

METTL3 silencing delays cellular senescence and enhances mitophagy by upregulating PINK1 in BLM-induced hFLSs

As verified by Western blotting, we found that METTL3 silencing upregulated PINK1 in BLM-treated hFLSs, which was reversed by PINK1 silencing (Fig. 5A, B and Supplementary Fig. 46). PINK1 silencing also attenuated the reduction in β-galactosidase activity and the number of mitochondrial fragments and the increase in the MMP in BLM-treated hFLSs (Fig. 5C–G). The downregulation of mitochondrial p62, Drp1, p16^INK4a, p21, and DcR2 and the upregulation of Mfn2 could also be partially reversed by PINK1 silencing in BLM-induced hFLSs (Fig. 5H–M and Supplementary Figs. 47–49, 17). Moreover, PINK1 silencing weakened the increase in LC3B and TOMM20 expression mediated by METTL3 silencing in BLM-stimulated hFLSs (Fig. 5N). These results suggested that the delay of senescence and induction of mitophagy mediated by METTL3 silencing could be achieved in part by regulating PINK1 in BLM-induced hFLSs.

Fig. 5 — BLM-induced hFLSs were transfected with shMETTL3 or shPINK1. A, B Validation of PINK1 protein expression. C, D Assessment of cellular senescence with a β-GAL kit. E TEM images. F, G TMRM staining. H, I P62 expression in mitochondria. J–M Western blotting analysis of Drp1, Mfn2, p16^INK4a, p21, and DcR2 expression. N Double IF staining of LC3B and TOMM20 expression. *** P < 0.001.

Senescent FLSs induced cartilage degradation in vivo

A total of 2.5×10⁵ normal FLSs or senescent FLSs induced by bleomycin treatment were intra-articularly injected into the rats (Fig. 6A). The micro-CT results revealed that senescent FLSs induced by BLM could result in deterioration of the trabecular microstructure in the subchondral bone (Fig. 6B). Senescent FLSs induced by BLM decreased the BV/TV, Tb.Th, and HWT, and increased the Tb.Sp and OARSI scores of SD rats (Fig. 6C–H). The staining results revealed that in the sham and N-FLS groups, the surface of the cartilage was smooth, whereas in the S-FLS group, the chondrocytes were disorganized and even contained vacuoles and acellular zones. The proteoglycans and chondrocytes in the S-FLS group were prominently lower than those in the other two groups (Fig. 6I–K). The IF staining results also revealed that p16INK4a and MMP-13 expression was dramatically greater and that Col2A1, LC3B, and TOMM20 expression was lower than that in the sham and N-FLS groups (Fig. 6L–O, Supplementary Fig. 18). The in vivo results further confirmed that senescent FLSs could accelerate OA progression.

Discussion

In TMJOA, FLSs can activate and secrete inflammatory mediators and cytokines that promote joint destruction and fibrosis of the synovium¹⁹. Therefore, exploration of the mechanism of FLS is crucial for the development of treatment strategies for TMJOA. In this study, we also isolated primary FLSs from the synovial tissues of normal and MIA-induced TMJOA rats. Besides, senescent FLSs from TMJOA rats accelerated cartilage degradation.

Senescence is a crucial factor in TMJ degeneration²⁰. Senescent cells have a unique secretory phenotype, namely, the senescence-associated secretory phenotype (SASP)²¹. The common SASPs of chondrocytes include interleukin (IL), matrix metalloproteinase, ADAMTS, and VEGF²². Other important senescence phenotypes include the accumulation of SA-β-gal activity and elevated p16, p21, and DcR2 expression^23–26. Research has shown that Fyn can accelerate the degradation of articular cartilage and the development of age-dependent OA²⁷. Based on our data, we demonstrated that cellular senescence was enhanced in MIA-induced TMJOA rats and primary FLSs. We also found that BLM-induced senescent FLSs can also induce cartilage degradation in rats, suggesting a potential feedback loop where aging synovial cells perpetuate the disease progression.

Mitophagy is essential for the maintenance of cellular homeostasis²⁸. Excessive activation of mitophagy can cause a decrease in mitochondrial number and mitochondrial dysfunction, which affects cellular metabolic processes and leads to the development of diseases such as cancer, aging, diabetes, and OA²⁹. Our results first revealed that mitophagy was decreased during the process of TMJOA. Besides, we demonstrated that a mitophagy agonist (Torin1) could delay senescence and accelerate mitophagy in senescent hFLSs. A study also reported that mitochondrial damage in FLSs is associated with rheumatoid arthritis³⁰. Through correlation analysis, we further revealed that TMJOA progression was associated with mitophagy and senescence. A study also demonstrated that mitophagy could participate in senescence³¹. Our study also revealed that mitophagy was reduced in BLM-induced senescent hFLSs and senescent FLSs from SD rats. These findings underscore the importance of mitophagy in maintaining mitochondrial integrity and delaying cellular senescence, particularly in the context of TMJOA.

Mitophagy involves two main pathways: the PTEN-induced PINK1/Parkin pathway and the related receptor-mediated pathway³². Upon MMP depolarization, PINK1 accumulates on the mitochondrial outer membrane and activates Parkin^33,34. Activated Parkin functions as an E3 ubiquitin ligase to promote the ubiquitination of mitochondrial outer membrane proteins³². These ubiquitinated proteins are recognized by autophagy-associated proteins and guide the encapsulation of damaged mitochondria into the autophagosome for eventual degradation. Besides, PINK1 can recruit autophagy receptors to mitochondria through ubiquitin phosphorylation to accelerate autophagy. p62 also acts as an autophagic substrate in addition to its role as a mitophagy receptor³⁵. Accordingly, the decreased p62 level detected in mitochondrial fractions following PINK1 overexpression likely reflects enhanced mitophagy flux and more efficient clearance of damaged mitochondria, during which p62 is degraded together with the cargo, rather than reduced recruitment. ULK1 is a key kinase of the autophagy initiation complex³⁶. DFCP1 is involved in autophagosome formation³⁷. WIPI1 interacts with autophagy-related proteins³⁸. Our research suggested that BLM could downregulate Parkin and PINK1 but did not change the expression of DFCP1, WIPI1, optineurin, NDP52, or ULK1 in hFLSs. These data suggested that only PINK1/Parkin pathway-mediated mitophagy was associated with hFLS senescence. Functionally, our data revealed that PINK1 overexpression slowed cellular senescence and induced mitophagy in senescent hFLSs, suggesting the importance of the PINK1/Parkin pathway in senescent hFLSs. Although the PINK1/Parkin pathway was strongly implicated in mitophagy regulation in this study, mitophagy activation was mainly evaluated through LC3B/TOMM20 co-localization and changes in p62 expression. However, the ubiquitination of mitochondrial membrane proteins, a key downstream event of the PINK1/Parkin pathway, was not directly examined. Future studies assessing mitochondrial ubiquitination may further strengthen these findings. m⁶A methylation is the most important posttranscriptional mRNA modification of genes and plays a crucial role in physiological processes such as cellular senescence, differentiation, apoptosis, and autophagy^39,40. Many studies have confirmed that METTL3 can increase m6A levels^39,41. A recent study revealed that METTL3 is highly expressed in OA-isolated FLSs and that METTL3 can mediate ATG7 mRNA m6A modification to regulate autophagy to increase the senescence of FLSs to accelerate the OA process⁴². It has also been demonstrated that abnormal mechanical stress can enhance chondrocyte senescence by upregulating METTL3, which is mediated by the loss of YAP¹⁰. Our data revealed that METTL3 levels were markedly increased in MIA-induced TMJOA rats, primary FLSs, and BLM-induced senescent hFLSs. The results of functional experiments also revealed that METTL3 silencing inhibited senescence and induced mitophagy in senescent hFLSs. Besides, we found that the PINK1 m⁶A mRNA level was increased in TMJOA rats, FLSs from MIA-induced TMJOA rats, and senescent hFLSs. Moreover, METTL3 knockdown reduced PINK1 m6A modification and enhanced its mRNA stability. METTL3 silencing also delays cellular senescence and induces mitophagy by upregulating PINK1 in BLM-induced hFLSs.

This study uncovers the critical role of METTL3-mediated m6A modification of PINK1 in regulating mitophagy and cellular senescence, shedding light on the molecular mechanisms driving TMJOA progression (Fig. 6P). These findings suggest that the interplay between METTL3, PINK1, and mitochondrial function may represent a broader biological phenomenon, potentially relevant to other inflammatory or degenerative conditions. The regulation of senescence and mitochondrial homeostasis by these pathways could play a significant role in diseases characterized by chronic inflammation or tissue degeneration. Although this research identifies potential molecular targets for intervention, further validation using human clinical samples is necessary to establish their relevance beyond experimental models. Future investigations should also examine the dynamic changes in METTL3 and PINK1 during different stages of TMJOA and explore their involvement in other joint disorders or systemic inflammatory processes. Such studies will help to clarify their broader impact on human health and may provide a foundation for developing novel therapeutic strategies.

Methods

Experimental animals

Male SPF-grade SD rats (8 weeks old, weighing 210 ~ 250 g) were provided by Sun Yat-sen University Laboratory Animal Center (SCXK (Yue) 2024-0072). The Guiding Opinions on Kindness to Laboratory Animals were strictly followed during the experiments. All the rats were fed for 1 week before the experiment in a warm room (room temperature, 21–25 °C; light and darkness, 12 h each; and free access to food and water). We have complied with all relevant ethical regulations for animal use. The experiment was approved by the Animal Ethics Committee of Sichuan Provincial People’s Hospital, University of Electronic Science and Technology of China (Lunshen(Yan)2022-233).

Construction of the rat model of TMJOA

To establish the TMJOA model, all procedures were conducted under aseptic conditions. Rats were randomly allocated and deeply anesthetized using pentobarbital sodium (Merck, Germany; 40 mg/kg, intraperitoneally). The hair around the TMJ region was shaved, and the skin was sterilized with iodophor. For the TMJOA model, 50 μL of monosodium iodoacetate (MIA; Sigma-Aldrich, USA; 4 mg/kg) was injected into the upper joint cavity of the TMJ on both sides using a microsyringe. In another experimental group, 2 × 10⁵ normal or senescent FLSs suspended in PBS were injected into the TMJ upper cavity using the same method. The sham group, which served as the negative control, received an injection of 50 μL of sterile PBS into the TMJ cavity without MIA or FLSs. Postoperative monitoring was conducted to ensure the rats’ well-being, and efforts were made to minimize stress and discomfort throughout the experiment. Each group contained 8 rats.

Micro-CT detection

All the samples were fixed, washed, and scanned with a SkyScan 1172 Micro-CT system (SkyScan, Bruker, Belgium)⁴³. After scanning, the two-dimensional image sequence was automatically reconstructed via the SkyScanNR econ software and the Feldkamp cone-beam algorithm. Bone microstructural parameters, such as the bone volume/total volume (BV/TV), trabecular number (Tb. N), trabecular separation (Tb. Sp), and trabecular thickness (Tb. Th), were analyzed with the custom analysis program (CTAn, SkyScan).

Detection of the head withdrawal threshold (HWT)

Pain behavior was analyzed by measuring the HWT of the rats in each group as previously reported⁴⁴. Briefly, the TMJ region of each rat was contacted via a Von Frey tenderness filament, and pressure was gradually applied until the rat’s head swung, at which time the final pressure value was recorded as the HWT, and each side was tested 5 times.

H&E staining

Fresh tissues from each group were fixed in 4% paraformaldehyde at 4 °C for 24 h. The samples were then decalcified in 10% EDTA solution (pH 7.4) at 4 °C until complete decalcification, with the solution refreshed regularly, followed by dehydration, paraffin embedding, and sagittal sectioning at a thickness of 5 μm. The sections were dewaxed, hydrated, and stained with hematoxylin–eosin. Histopathological changes were observed under a light microscope (Olympus BX53, Olympus, Tokyo, Japan).

Safranin O staining

The samples were processed using the same fixation, decalcification, and paraffin-embedding procedures described above and sectioned at a thickness of 5 μm. The sections were stained with freshly prepared Weigert’s iron hematoxylin, counterstained with Fast Green (Solarbio), and followed by Safranin O (Solarbio). After dehydration and sealing, the morphology of the articular cartilage was observed under a light microscope (Olympus BX53, Olympus, Tokyo, Japan). Images were acquired at a magnification of 100×. The severity of the TMJ-OA-like phenotype was assessed using the OARSI scoring system, as previously described^5,45,46. Scoring was performed on sagittal sections by observers blinded to the experimental groups.

Alcian blue (AB) staining

Paraffin sections of the cartilage tissues (5 μm thickness) were dewaxed, rehydrated, and immersed in 1% Alcian blue solution for 1 h. After washing, the sections were dehydrated, cleared with xylene for 5 min, and sealed with neutral gum. Images were obtained using a light microscope (Olympus BX53, Olympus, Tokyo, Japan).

Immunohistochemistry (IHC)

Paraffin-embedded sections prepared using the same fixation, decalcification, and sectioning procedures described above were deparaffinized with xylene and hydrated with graded alcohols. Antigen retrieval was performed by microwave heating for 10 min. The sections were treated with 3% H₂O₂ for 15 min and blocked with 5% BSA for 30 min. After washing, the sections were incubated with anti-MMP-13 (1:200) or anti-Col2A1 (1:150), followed by secondary antibodies (1:1000; Abcam). The results were visualized with diaminobenzidine (DAB, Sigma). Images were acquired using an upright light microscope (BX53, Olympus, Tokyo, Japan). Five non-overlapping microscopic fields were randomly selected from each section. Cells with distinct DAB staining above background were counted as positive, and the average number of positive cells per field was calculated per animal for statistical analysis.

Isolation of primary FLSs

The synovial tissues of normal and MIA-induced TMJOA model rats were cut into 1 mm pieces, washed with saline, supplemented with 1% collagenase I, and digested at 37 °C for 2 h. After centrifugation, the cells were cultured with RPMI 1640 (Gibco) supplemented with 20% FBS, 1% nonessential amino acids, 1% sodium pyruvate, and 1% glutamine at 37 °C with 5% CO₂. When the cell confluence reached 95%, the cells were digested and passaged. After the 3rd generation, the remaining cells were mainly FLSs and were cultured for subsequent experiments.

Cell culture

Human articular chondrocytes were obtained from Procell Life Science & Technology (CP-H096, Wuhan, China) and cultured in DMEM/F12 medium supplemented with 10% fetal bovine serum (FBS; Sigma, USA) at 37 °C in a humidified atmosphere with 5% CO₂. Human fibroblast-like synoviocytes (hFLSs) were obtained from Cell Applications, Inc. (No. 408-05a, San Diego, CA, USA) and maintained in HFLS medium (No. 415–500, Cell Applications) at 37 °C with 5% CO₂, as previously described⁴⁷. Rat fibroblast-like synoviocytes (FLSs) were isolated from the synovial tissues of normal and MIA-induced TMJOA Sprague-Dawley rats (6–8 weeks old, 220–270 g) following standard enzymatic digestion protocols. The isolated rat FLSs were cultured in DMEM supplemented with 10% FBS at 37 °C in 5% CO₂ until they reached confluence. Cells were used for experiments after 2–3 passages to ensure uniformity and viability.

Cell treatment

PINK1 overexpression plasmids (OE-PINK1), empty vectors (OE-CTRL), METTL3 shRNAs (shMETTL3-1 and shMETTL3-2, also named shIME4-1 and shIME4-2), PINK1 shRNAs (shPINK1), and sh-control (shCTRL) were obtained from GenePharma (Shanghai, China). hFLSs (5 × 10⁴ cells/well) were placed in a 12-well plate, cultured overnight, and transfected with the plasmids or shRNAs via Lipofectamine 3000 for 48 h according to the instructions. Besides, hFLSs were treated with 10 μM bleomycin (BLM) or 100 nmol/L mitophagy agonist (Torin1; Selleck, S2827) for 6 hours.

Co-culture setup

To simulate the osteoarthritic microenvironment in vitro, a Transwell co-culture system (Corning, USA) was employed. An insert with a 0.4 μm pore membrane was used to allow the exchange of soluble mediators while preventing direct cell–cell contact, thereby specifically focusing on paracrine interactions between FLSs and chondrocytes. Human articular chondrocytes were selected because commercially available human cells exhibit relatively stable phenotypic characteristics and good batch-to-batch consistency compared with primary rat chondrocytes, which helps reduce variability associated with animal-derived samples. Under this non-contact condition, cross-species co-culture is suitable for assessing soluble factor–mediated cellular responses. Human chondrocytes were seeded into the lower chamber of the Transwell plates, while rat FLSs were plated in the upper chamber separated by a 0.4 μm pore membrane, which allowed the exchange of soluble factors between the cell types without direct cell-cell contact. Two experimental groups were established to assess the effects of FLSs on chondrocytes: Normal-FLS group (Human chondrocytes co-cultured with FLSs isolated from normal rats) and TMJOA-FLS group (Human chondrocytes co-cultured with FLSs isolated from MIA-induced TMJOA rats). All co-cultures were maintained under identical conditions, including equal cell densities, medium volumes, and incubation times, to minimize experimental bias. Medium changes were performed carefully to avoid contamination, and each experiment was repeated independently to confirm reproducibility.

RT-qPCR

Total RNA from the treated cells and tissues was extracted with TRIzol (Invitrogen). cDNA was reverse-transcribed with the PrimeScript RT Master Mix Kit (Takara). PCR was conducted using SYBR Green Real-Time PCR Master Mix (Thermo Fisher Scientific). The relative expression was calculated via the 2^−∆∆Ct method. The primer sequences are shown in Supplementary Table1.

RNA stability assay

The treated hFLSs were treated with 5 μg/mL actinomycin D (MCE, Cat. no. HY-17559) for 0, 4, 8, or 12 h. PINK1 expression was confirmed via RT-qPCR.

Western blot

Mitochondria were extracted via a mitochondrial extraction kit (LMAl Bio, Shanghai, China) following the manufacturer’s instructions. For western blotting analysis of mitochondrial proteins, equal amounts of isolated mitochondrial protein were loaded. TOMM20 was used as a mitochondrial loading control. β-actin was used as a cytosolic marker to verify the purity of the mitochondrial fraction. Total protein was extracted from RIPA lysis buffer (Beyotime). The protein concentration was quantified with a BCA kit (Thermo Fisher Scientific, USA). The protein (30 μg) was separated via 10% SDS-PAGE and transferred to a PVDF membrane (Bio-Rad, USA). The PVDF membrane was blocked with 5% skim milk powder at 37 °C for 1 h and incubated with specific primary antibodies at 4 °C overnight. Then, the membranes were washed and incubated with a secondary antibody (Proteintech) for 2 hours. Visualization of the protein bands was conducted via an enhanced ECL detection kit (Pierce, Rockford, IL, USA). The gray values of the protein bands were quantitatively analyzed via ImageJ software (NIH, USA). The primary antibodies included p16INK4a (1:1000, Santa Cruz Biotechnology Cat# sc-81156, RRID:AB_1126947), Vimentin (1:1000, Santa Cruz Biotechnology Cat# sc-6260, RRID:AB_628437), p62 (1:1200, Santa Cruz Biotechnology Cat# sc-48402, RRID:AB_2255371), Mfn2 (1:1000, Santa Cruz Biotechnology Cat# sc-100560, RRID:AB_2235195), Drp1 (1:1000, Santa Cruz Biotechnology Cat# sc-101270, RRID:AB_2093545), p21 (1:1500, Santa Cruz Biotechnology Cat# sc-166630, RRID:AB_2059940), DcR2 (also designated TNFRSF10B or DR5, 1:1000, Abcam Cat# ab8416, RRID:AB_306551), METTL3 (1:20000, Proteintech Cat# 15073-1-AP, RRID:AB_2142033), TOMM20 (1:30000, Proteintech Cat# 11802-1-AP, RRID:AB_2207530), ADAMTS5 (1:1000, Abcam Cat# ab41037, RRID:AB_2222327), MMP13 (1:2000, Proteintech Cat# 18165-1-AP, RRID:AB_2144858), Col2A1 (1:2000, Proteintech Cat# 28459-1-AP, RRID:AB_2881147), Parkin (1:2000, Proteintech Cat# 14060-1-AP, RRID:AB_2878005), PINK1 (1:1000, Proteintech Cat# 23274-1-AP, RRID:AB_2879244), DFCP1 (1:1000, Abcam Cat# ab102599, RRID:AB_10711773), WIPI1 (1:1000, Proteintech Cat# 25204-1-AP, RRID:AB_2879958), and Optineurin (1:5000, Proteintech Cat# 10837-1-AP, RRID:AB_2156665).

Immunofluorescence (IF) staining

Tissues and cell samples from each group were fixed, treated with 0.5% Triton-X for membrane disruption, blocked with 10% BSA at room temperature, diluted with a specific primary antibody, and incubated in a refrigerator at 4 °C overnight. Finally, FITC-labeled goat anti-rabbit IgG (Life Technologies, USA) (1:1000) and DAPI were added. After cleaning and sealing, the images were observed using a Nikon A1 confocal laser scanning microscope (Nikon, Tokyo, Japan). The primary antibodies used included p16INK4a (1:100), vimentin (1:100), LC3B (1:100, Proteintech Cat# 18725-1-AP, RRID: AB_2137745), TOMM20 (1:100), ADAMTS5 (1:100), collagen II (1:150), and MMP-13 (1:100) antibodies.

ELISA

Synovial tissues were collected and homogenized on ice, and the supernatants were used for ELISA. The levels of m6A were measured using an m6A ELISA kit (EpiQuik). The protein levels of IL-1β, TNF-α, and IL-6 were determined using ELISA kits (Solarbio) according to the manufacturer’s instructions.

SA-β-galactosidase staining

The collected cells were fixed for 15 min, washed, and incubated with SA-β-gal staining solution (Beyotime, C0602) at 37 °C for 16–20 h. After washing, the senescent cells were observed under an inverted light microscope (IX73, Olympus, Tokyo, Japan). For frozen sections, the sections were dried at 37 °C for 20–30 min and fixed for 15 minutes. After washing, the sections were treated with SA-β-gal (Beyotime) overnight, stained with eosin for 1–2 min, rinsed for 1 min, differentiated with 1% acidic ethanol for 10–20 s, rinsed for 1 min, dehydrated with gradient ethanol and washed with xylene. After drying, the samples were observed under a light microscope (Olympus BX53, Olympus, Tokyo, Japan).

Tetramethylrhodamine methyl ester (TMRM) staining

The processed cells were collected and made into a single-cell suspension of 1 × 10⁵ cells/ml and seeded in six-well plates. After culture and adherence, the cells were stained with 100 μmol/L TMRM reagent (Invitrogen) for 30 min. The fluorescence was imaged using an inverted microscope (IX73, Olympus, Tokyo, Japan) equipped for fluorescence imaging.

Transmission electron microscopy (TEM)

After washing, the collected cells were fixed with 2.5% glutaraldehyde for 90 min. After being fixed with 1% osmium tetroxide, the cells were dehydrated in gradient ethanol. The cells were subsequently embedded in Epon812 resin. The blocks were cut into ultrathin sections (60–80 nm) via a slicer. The sections were stained with saturated hydrogen peroxide acetate, and lead citrate. The mitochondrial structure was observed via a JEM-1200EX transmission electron microscope (JEOL, Japan).

RNA methylation immunoprecipitation (MeRIP-qPCR)

Putative m⁶A modification sites on PINK1 mRNA were identified using publicly available m⁶A modification data from m⁶A Atlas (https://rnamd.org/m6a/). Candidate regions at positions 2028, 2074, 2174, and 2483 were selected based on their localization within predicted m⁶A-enriched regions and typical m⁶A consensus motifs. Then, the RNA was fragmented via a m⁶A-MeRIP Kit (GS-T-001, Gen Seq) following the manufacturer’s instructions, and a small amount of RNA was used as the input group, while the remainder was incubated with IgG or a m⁶A antibody and purified via qRT-PCR.

Statistics and Reproducibility

Each experiment was repeated at least three times independently. The experimental data were analyzed via SPSS 23.0 software (SPSS Inc., Chicago, IL, USA). If the data were normally distributed and chi-square tests were performed, one-way analysis of variance was applied for comparisons among multiple groups. If the data did not conform to a normal distribution or chi-square test, nonparametric tests were applied. The test level was α = 0.05. The correlation between the data was analyzed via Spearman correlation.

Ethics approval and consent to participate

We have complied with all relevant ethical regulations for animal use. The experiment was approved by the Animal Ethics Committee of Sichuan Provincial People’s Hospital, University of Electronic Science and Technology of China (Lunshen (Yan) 2022-233).

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Supplementary information

Supplementary Information^{(6.7MB, pdf)}

42003_2026_9773_MOESM2_ESM.pdf^{(27.6KB, pdf)}

Description of Additional Supplementary Files

Supplementary Data^{(51.1KB, xlsx)}

Reporting Summary^{(2.5MB, pdf)}

Acknowledgements

The present study was supported in part by the National Natural Science Foundation of China (82470953) and the Natural Science Foundation of Sichuan Province (2025ZNSFSC0750).

Author contributions

Conceptualization: L.X. and Y.M. Data curation: J.L., K.T., and Q.D. Formal analysis: L.X., J.L., and J.G. Funding acquisition: Y.M. and L.X. Investigation: J.L., W.L., X.Y., and S.L. Methodology: L.X. and K.T. Project administration: L. X. and J.L. Visualization: L.X. and K.T. Writing - original draft: L.X. and Y.M. Writing - review & editing: All authors.

Peer review

Peer review information

Communications Biology thanks Wacharapol Saengsiwaritt and Ruina Kong for their contribution to the peer review of this work. Primary Handling Editors: Carmen Hueasa and Joao Valente.

Data availability

The datasets used and/or analyzed during the current study are available from the corresponding author on reasonable request. The numerical source data underlying graphs in the manuscript can be found in the supplementary data file.

Competing interests

The authors declare no competing interests.

Footnotes

Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Contributor Information

Juan Liao, Email: 109497731@qq.com.

Yandong Mu, Email: muyd@uestc.edu.cn.

Li Xiao, Email: xiaolikq@med.uestc.edu.cn.

Supplementary information

The online version contains supplementary material available at 10.1038/s42003-026-09773-x.

References

1.Roberts, W. E. & Goodacre, C. J. The Temporomandibular joint: a critical review of life-support functions, development, articular surfaces, biomechanics and degeneration. J. Prosthodont.29, 772–779 (2020). [DOI] [PubMed] [Google Scholar]
2.Wang, D. et al. Recent advances in animal models, diagnosis, and treatment of temporomandibular joint Osteoarthritis. Tissue Eng. B Rev.29, 62–77 (2023). [DOI] [PubMed] [Google Scholar]
3.Liu, G. et al. Insights into the Notch signaling pathway in degenerative musculoskeletal disorders: Mechanisms and perspectives. Biomed. Pharmacother.169, 115884 (2023). [DOI] [PubMed] [Google Scholar]
4.Zhao, X. et al. Symptoms, disc position, occluding pairs, and facial skeletal characteristics of older patients with temporomandibular disorders. J. Int. Med. Res.49, 300060521990530 (2021). [DOI] [PMC free article] [PubMed] [Google Scholar]
5.Zhou, J., Ren, R., Li, Z., Zhu, S. & Jiang, N. Temporomandibular joint osteoarthritis: A review of animal models induced by surgical interventions. Oral. Dis.29, 2521–2528 (2023). [DOI] [PubMed] [Google Scholar]
6.Nygaard, G. & Firestein, G. S. Restoring synovial homeostasis in rheumatoid arthritis by targeting fibroblast-like synoviocytes. Nat. Rev. Rheumatol.16, 316–333 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
7.Tsaltskan, V. & Firestein, G. S. Targeting fibroblast-like synoviocytes in rheumatoid arthritis. Curr. Opin. Pharm.67, 102304 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]
8.Yang, J. et al. Targeting YAP1-regulated glycolysis in fibroblast-like synoviocytes impairs macrophage infiltration to ameliorate diabetic osteoarthritis progression. Adv. Sci.11, e2304617 (2024). [DOI] [PMC free article] [PubMed] [Google Scholar]
9.Liu, B. et al. Inflammatory fibroblast-like synoviocyte-derived exosomes aggravate osteoarthritis via enhancing macrophage glycolysis. Adv. Sci.11, e2307338 (2024). [DOI] [PMC free article] [PubMed] [Google Scholar]
10.Yang, F. et al. Abnormal mechanical stress induced chondrocyte senescence by YAP loss-mediated METTL3 upregulation. Oral. Dis.30, 3308–3320 (2024). [DOI] [PubMed] [Google Scholar]
11.Miwa, S., Kashyap, S., Chini, E. & von Zglinicki, T. Mitochondrial dysfunction in cell senescence and aging. J. Clin. Investig.132 (2022). [DOI] [PMC free article] [PubMed]
12.Jiang, W. et al. Mechanisms linking mitochondrial mechanotransduction and chondrocyte biology in the pathogenesis of osteoarthritis. Ageing Res. Rev.67, 101315 (2021). [DOI] [PubMed] [Google Scholar]
13.Bai, Y. et al. RNA methylation, homologous recombination repair and therapeutic resistance. Biomed. Pharmacother.166, 115409 (2023). [DOI] [PubMed] [Google Scholar]
14.Zhao, X. et al. Alterations of the m(6)A Methylation Induced by TGF-β2 in ARPE-19 Cells. Front Biosci.28, 148 (2023). [DOI] [PubMed] [Google Scholar]
15.Wei, F., Zhang, J. N., Zhao, Y. Q., Lyu, H. & Chen, F. Expression of m6A RNA Methylation Regulators and Their Clinical Predictive Value in Intrahepatic Cholangiocarcinoma. Front. Biosci.28, 120 (2023). [DOI] [PubMed] [Google Scholar]
16.Ye, G. et al. ALKBH5 facilitates CYP1B1 mRNA degradation via m6A demethylation to alleviate MSC senescence and osteoarthritis progression. Exp. Mol. Med.55, 1743–1756 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]
17.Lu, Y. et al. Expression pattern analysis of m6A regulators reveals IGF2BP3 as a key modulator in osteoarthritis synovial macrophages. J. Transl. Med.21, 339 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]
18.Jiang, X. et al. The role of m6A modification in the biological functions and diseases. Signal Transduct. Target Ther.6, 74 (2021). [DOI] [PMC free article] [PubMed] [Google Scholar]
19.Han, D. et al. The emerging role of fibroblast-like synoviocytes-mediated synovitis in osteoarthritis: An update. J. Cell Mol. Med.24, 9518–9532 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
20.Zhou, Y. et al. Senolytics alleviate the degenerative disorders of temporomandibular joint in old age. Aging Cell20, e13394 (2021). [DOI] [PMC free article] [PubMed] [Google Scholar]
21.Saul, D. et al. A new gene set identifies senescent cells and predicts senescence-associated pathways across tissues. Nat. Commun.13, 4827 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]
22.Coryell, P. R., Diekman, B. O. & Loeser, R. F. Mechanisms and therapeutic implications of cellular senescence in osteoarthritis. Nat. Rev. Rheumatol.17, 47–57 (2021). [DOI] [PMC free article] [PubMed] [Google Scholar]
23.Yagi, M., Endo, K., Komori, K. & Sekiya, I. Comparison of the effects of oxidative and inflammatory stresses on rat chondrocyte senescence. Sci. Rep.13, 7697 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]
24.Huang, W., Hickson, L. J., Eirin, A., Kirkland, J. L. & Lerman, L. O. Cellular senescence: the good, the bad and the unknown. Nat. Rev. Nephrol.18, 611–627 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]
25.O’Sullivan, E. A., Wallis, R., Mossa, F. & Bishop, C. L. The paradox of senescent-marker positive cancer cells: challenges and opportunities. NPJ Aging10, 41 (2024). [DOI] [PMC free article] [PubMed] [Google Scholar]
26.Chen, J. et al. DCR2, a cellular senescent molecule, is a novel marker for assessing tubulointerstitial fibrosis in patients with immunoglobulin A nephropathy. Kidney Blood Press Res44, 1063–1074 (2019). [DOI] [PubMed] [Google Scholar]
27.Li, K. et al. Tyrosine kinase Fyn promotes osteoarthritis by activating the β-catenin pathway. Ann. Rheum. Dis.77, 935–943 (2018). [DOI] [PubMed] [Google Scholar]
28.Sun, K., Jing, X., Guo, J., Yao, X. & Guo, F. Mitophagy in degenerative joint diseases. Autophagy17, 2082–2092 (2021). [DOI] [PMC free article] [PubMed] [Google Scholar]
29.Bernabei, I., So, A., Busso, N. & Nasi, S. Cartilage calcification in osteoarthritis: mechanisms and clinical relevance. Nat. Rev. Rheumatol.19, 10–27 (2023). [DOI] [PubMed] [Google Scholar]
30.Wang, Y. et al. Mst1 promotes mitochondrial dysfunction and apoptosis in oxidative stress-induced rheumatoid arthritis synoviocytes. Aging12, 16211–16223 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
31.Luo, Y., Zhang, L., Su, N., Liu, L. & Zhao, T. YME1L-mediated mitophagy protects renal tubular cells against cellular senescence under diabetic conditions. Biol. Res.57, 10 (2024). [DOI] [PMC free article] [PubMed] [Google Scholar]
32.Li, J. et al. PINK1/Parkin-mediated mitophagy in neurodegenerative diseases. Ageing Res. Rev.84, 101817 (2023). [DOI] [PubMed] [Google Scholar]
33.Imberechts, D. et al. DJ-1 is an essential downstream mediator in PINK1/parkin-dependent mitophagy. Brain145, 4368–4384 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]
34.Yan, C. et al. PHB2 (prohibitin 2) promotes PINK1-PRKN/Parkin-dependent mitophagy by the PARL-PGAM5-PINK1 axis. Autophagy16, 419–434 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
35.Lippai, M. & Lőw, P. The role of the selective adaptor p62 and ubiquitin-like proteins in autophagy. Biomed. Res. Int.2014, 832704 (2014). [DOI] [PMC free article] [PubMed] [Google Scholar]
36.Zhang, S. et al. Negative pressure wound therapy improves bone regeneration by promoting osteogenic differentiation via the AMPK-ULK1-autophagy axis. Autophagy18, 2229–2245 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]
37.Nähse, V. et al. ATPase activity of DFCP1 controls selective autophagy. Nat. Commun.14, 4051 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]
38.Sirko, A., Wawrzyńska, A., Brzywczy, J. & Sieńko, M. Control of ABA signaling and crosstalk with other hormones by the selective degradation of pathway components. Int. J. Mol. Sci.22. 10.3390/ijms22094638 (2021). [DOI] [PMC free article] [PubMed]
39.Luo, S. et al. METTL3-mediated m6A mRNA methylation regulates neutrophil activation through targeting TLR4 signaling. Cell Rep.42, 112259 (2023). [DOI] [PubMed] [Google Scholar]
40.Kumari, R. et al. mRNA modifications in cardiovascular biology and disease: with a focus on m6A modification. Cardiovasc. Res.118, 1680–1692 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]
41.Chen, L. et al. METTL3-mediated m6A modification stabilizes TERRA and maintains telomere stability. Nucleic Acids Res.50, 11619–11634 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]
42.Chen, X. et al. METTL3-mediated m(6)A modification of ATG7 regulates autophagy-GATA4 axis to promote cellular senescence and osteoarthritis progression. Ann. Rheum. Dis.81, 87–99 (2022). [DOI] [PubMed] [Google Scholar]
43.Chen, B. et al. Metformin suppresses oxidative stress induced by high glucose via activation of the Nrf2/HO-1 signaling pathway in Type 2 diabetic osteoporosis. Life Sci.312, 121092 (2023). [DOI] [PubMed] [Google Scholar]
44.Jiang, H. et al. Chronic pain causes peripheral and central responses in MIA-induced TMJOA Rats. Cell. Mol. Neurobiol.42, 1441–1451 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]
45.Feng, S. Y. et al. Increased joint loading induces subchondral bone loss of the temporomandibular joint via the RANTES-CCRs-Akt2 axis. JCI Insight7. 10.1172/jci.insight.158874 (2022). [DOI] [PMC free article] [PubMed]
46.Yuan, W. et al. Comparison and applicability of three induction methods of temporomandibular joint osteoarthritis in murine models. J. Oral. Rehabil.49, 430–441 (2022). [DOI] [PubMed] [Google Scholar]
47.Sukkaew, A. et al. Heterogeneity of clinical isolates of chikungunya virus and its impact on the responses of primary human fibroblast-like synoviocytes. J. Gen. Virol.99, 525–535 (2018). [DOI] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supplementary Information^{(6.7MB, pdf)}

42003_2026_9773_MOESM2_ESM.pdf^{(27.6KB, pdf)}

Description of Additional Supplementary Files

Supplementary Data^{(51.1KB, xlsx)}

Reporting Summary^{(2.5MB, pdf)}

Data Availability Statement

[CR1] 1.Roberts, W. E. & Goodacre, C. J. The Temporomandibular joint: a critical review of life-support functions, development, articular surfaces, biomechanics and degeneration. J. Prosthodont.29, 772–779 (2020). [DOI] [PubMed] [Google Scholar]

[CR2] 2.Wang, D. et al. Recent advances in animal models, diagnosis, and treatment of temporomandibular joint Osteoarthritis. Tissue Eng. B Rev.29, 62–77 (2023). [DOI] [PubMed] [Google Scholar]

[CR3] 3.Liu, G. et al. Insights into the Notch signaling pathway in degenerative musculoskeletal disorders: Mechanisms and perspectives. Biomed. Pharmacother.169, 115884 (2023). [DOI] [PubMed] [Google Scholar]

[CR4] 4.Zhao, X. et al. Symptoms, disc position, occluding pairs, and facial skeletal characteristics of older patients with temporomandibular disorders. J. Int. Med. Res.49, 300060521990530 (2021). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR5] 5.Zhou, J., Ren, R., Li, Z., Zhu, S. & Jiang, N. Temporomandibular joint osteoarthritis: A review of animal models induced by surgical interventions. Oral. Dis.29, 2521–2528 (2023). [DOI] [PubMed] [Google Scholar]

[CR6] 6.Nygaard, G. & Firestein, G. S. Restoring synovial homeostasis in rheumatoid arthritis by targeting fibroblast-like synoviocytes. Nat. Rev. Rheumatol.16, 316–333 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR7] 7.Tsaltskan, V. & Firestein, G. S. Targeting fibroblast-like synoviocytes in rheumatoid arthritis. Curr. Opin. Pharm.67, 102304 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR8] 8.Yang, J. et al. Targeting YAP1-regulated glycolysis in fibroblast-like synoviocytes impairs macrophage infiltration to ameliorate diabetic osteoarthritis progression. Adv. Sci.11, e2304617 (2024). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR9] 9.Liu, B. et al. Inflammatory fibroblast-like synoviocyte-derived exosomes aggravate osteoarthritis via enhancing macrophage glycolysis. Adv. Sci.11, e2307338 (2024). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR10] 10.Yang, F. et al. Abnormal mechanical stress induced chondrocyte senescence by YAP loss-mediated METTL3 upregulation. Oral. Dis.30, 3308–3320 (2024). [DOI] [PubMed] [Google Scholar]

[CR11] 11.Miwa, S., Kashyap, S., Chini, E. & von Zglinicki, T. Mitochondrial dysfunction in cell senescence and aging. J. Clin. Investig.132 (2022). [DOI] [PMC free article] [PubMed]

[CR12] 12.Jiang, W. et al. Mechanisms linking mitochondrial mechanotransduction and chondrocyte biology in the pathogenesis of osteoarthritis. Ageing Res. Rev.67, 101315 (2021). [DOI] [PubMed] [Google Scholar]

[CR13] 13.Bai, Y. et al. RNA methylation, homologous recombination repair and therapeutic resistance. Biomed. Pharmacother.166, 115409 (2023). [DOI] [PubMed] [Google Scholar]

[CR14] 14.Zhao, X. et al. Alterations of the m(6)A Methylation Induced by TGF-β2 in ARPE-19 Cells. Front Biosci.28, 148 (2023). [DOI] [PubMed] [Google Scholar]

[CR15] 15.Wei, F., Zhang, J. N., Zhao, Y. Q., Lyu, H. & Chen, F. Expression of m6A RNA Methylation Regulators and Their Clinical Predictive Value in Intrahepatic Cholangiocarcinoma. Front. Biosci.28, 120 (2023). [DOI] [PubMed] [Google Scholar]

[CR16] 16.Ye, G. et al. ALKBH5 facilitates CYP1B1 mRNA degradation via m6A demethylation to alleviate MSC senescence and osteoarthritis progression. Exp. Mol. Med.55, 1743–1756 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR17] 17.Lu, Y. et al. Expression pattern analysis of m6A regulators reveals IGF2BP3 as a key modulator in osteoarthritis synovial macrophages. J. Transl. Med.21, 339 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR18] 18.Jiang, X. et al. The role of m6A modification in the biological functions and diseases. Signal Transduct. Target Ther.6, 74 (2021). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR19] 19.Han, D. et al. The emerging role of fibroblast-like synoviocytes-mediated synovitis in osteoarthritis: An update. J. Cell Mol. Med.24, 9518–9532 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR20] 20.Zhou, Y. et al. Senolytics alleviate the degenerative disorders of temporomandibular joint in old age. Aging Cell20, e13394 (2021). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR21] 21.Saul, D. et al. A new gene set identifies senescent cells and predicts senescence-associated pathways across tissues. Nat. Commun.13, 4827 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR22] 22.Coryell, P. R., Diekman, B. O. & Loeser, R. F. Mechanisms and therapeutic implications of cellular senescence in osteoarthritis. Nat. Rev. Rheumatol.17, 47–57 (2021). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR23] 23.Yagi, M., Endo, K., Komori, K. & Sekiya, I. Comparison of the effects of oxidative and inflammatory stresses on rat chondrocyte senescence. Sci. Rep.13, 7697 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR24] 24.Huang, W., Hickson, L. J., Eirin, A., Kirkland, J. L. & Lerman, L. O. Cellular senescence: the good, the bad and the unknown. Nat. Rev. Nephrol.18, 611–627 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR25] 25.O’Sullivan, E. A., Wallis, R., Mossa, F. & Bishop, C. L. The paradox of senescent-marker positive cancer cells: challenges and opportunities. NPJ Aging10, 41 (2024). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR26] 26.Chen, J. et al. DCR2, a cellular senescent molecule, is a novel marker for assessing tubulointerstitial fibrosis in patients with immunoglobulin A nephropathy. Kidney Blood Press Res44, 1063–1074 (2019). [DOI] [PubMed] [Google Scholar]

[CR27] 27.Li, K. et al. Tyrosine kinase Fyn promotes osteoarthritis by activating the β-catenin pathway. Ann. Rheum. Dis.77, 935–943 (2018). [DOI] [PubMed] [Google Scholar]

[CR28] 28.Sun, K., Jing, X., Guo, J., Yao, X. & Guo, F. Mitophagy in degenerative joint diseases. Autophagy17, 2082–2092 (2021). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR29] 29.Bernabei, I., So, A., Busso, N. & Nasi, S. Cartilage calcification in osteoarthritis: mechanisms and clinical relevance. Nat. Rev. Rheumatol.19, 10–27 (2023). [DOI] [PubMed] [Google Scholar]

[CR30] 30.Wang, Y. et al. Mst1 promotes mitochondrial dysfunction and apoptosis in oxidative stress-induced rheumatoid arthritis synoviocytes. Aging12, 16211–16223 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR31] 31.Luo, Y., Zhang, L., Su, N., Liu, L. & Zhao, T. YME1L-mediated mitophagy protects renal tubular cells against cellular senescence under diabetic conditions. Biol. Res.57, 10 (2024). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR32] 32.Li, J. et al. PINK1/Parkin-mediated mitophagy in neurodegenerative diseases. Ageing Res. Rev.84, 101817 (2023). [DOI] [PubMed] [Google Scholar]

[CR33] 33.Imberechts, D. et al. DJ-1 is an essential downstream mediator in PINK1/parkin-dependent mitophagy. Brain145, 4368–4384 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR34] 34.Yan, C. et al. PHB2 (prohibitin 2) promotes PINK1-PRKN/Parkin-dependent mitophagy by the PARL-PGAM5-PINK1 axis. Autophagy16, 419–434 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR35] 35.Lippai, M. & Lőw, P. The role of the selective adaptor p62 and ubiquitin-like proteins in autophagy. Biomed. Res. Int.2014, 832704 (2014). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR36] 36.Zhang, S. et al. Negative pressure wound therapy improves bone regeneration by promoting osteogenic differentiation via the AMPK-ULK1-autophagy axis. Autophagy18, 2229–2245 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR37] 37.Nähse, V. et al. ATPase activity of DFCP1 controls selective autophagy. Nat. Commun.14, 4051 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR38] 38.Sirko, A., Wawrzyńska, A., Brzywczy, J. & Sieńko, M. Control of ABA signaling and crosstalk with other hormones by the selective degradation of pathway components. Int. J. Mol. Sci.22. 10.3390/ijms22094638 (2021). [DOI] [PMC free article] [PubMed]

[CR39] 39.Luo, S. et al. METTL3-mediated m6A mRNA methylation regulates neutrophil activation through targeting TLR4 signaling. Cell Rep.42, 112259 (2023). [DOI] [PubMed] [Google Scholar]

[CR40] 40.Kumari, R. et al. mRNA modifications in cardiovascular biology and disease: with a focus on m6A modification. Cardiovasc. Res.118, 1680–1692 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR41] 41.Chen, L. et al. METTL3-mediated m6A modification stabilizes TERRA and maintains telomere stability. Nucleic Acids Res.50, 11619–11634 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR42] 42.Chen, X. et al. METTL3-mediated m(6)A modification of ATG7 regulates autophagy-GATA4 axis to promote cellular senescence and osteoarthritis progression. Ann. Rheum. Dis.81, 87–99 (2022). [DOI] [PubMed] [Google Scholar]

[CR43] 43.Chen, B. et al. Metformin suppresses oxidative stress induced by high glucose via activation of the Nrf2/HO-1 signaling pathway in Type 2 diabetic osteoporosis. Life Sci.312, 121092 (2023). [DOI] [PubMed] [Google Scholar]

[CR44] 44.Jiang, H. et al. Chronic pain causes peripheral and central responses in MIA-induced TMJOA Rats. Cell. Mol. Neurobiol.42, 1441–1451 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR45] 45.Feng, S. Y. et al. Increased joint loading induces subchondral bone loss of the temporomandibular joint via the RANTES-CCRs-Akt2 axis. JCI Insight7. 10.1172/jci.insight.158874 (2022). [DOI] [PMC free article] [PubMed]

[CR46] 46.Yuan, W. et al. Comparison and applicability of three induction methods of temporomandibular joint osteoarthritis in murine models. J. Oral. Rehabil.49, 430–441 (2022). [DOI] [PubMed] [Google Scholar]

[CR47] 47.Sukkaew, A. et al. Heterogeneity of clinical isolates of chikungunya virus and its impact on the responses of primary human fibroblast-like synoviocytes. J. Gen. Virol.99, 525–535 (2018). [DOI] [PubMed] [Google Scholar]

PERMALINK

METTL3-mediated fibroblast-like synoviocytes senescence promotes temporomandibular joint osteoarthritis progression

Kun Tian

Qin Du

Jun Guo

Wen Liao

Xianrui Yang

Shuang Lai

Juan Liao

Yandong Mu

Li Xiao

Abstract

Introduction

Results

MIA induces pathological progression in the TMJ of Sprague–Dawley (SD) rats

Fig. 1. MIA induces pathological progression in the TMJ of SD rats.

Insufficient mitophagy and increased METTL3 in synovial tissues of TMJOA rats

Abnormal cell senescence, mitophagy, and METTL3 expression in primary FLSs

Fig. 2. Mitophagy is attenuated, and cell senescence and METTL3 expression are enhanced in primary FLSs from MIA-induced TMJOA rats.

FLSs from TMJOA rats induce cartilage degradation

Bleomycin (BLM) accelerates cell senescence, reduces mitophagy, and upregulates METTL3 in hFLSs

Torin1 inhibits cell senescence and promotes mitophagy in BLM-induced senescent hFLSs

METTL3 silencing weakens cell senescence and promotes mitophagy in BLM-treated hFLSs

Fig. 3. METTL3 silencing weakens cell senescence and accelerates mitophagy in BLM-induced senescent hFLSs.

METTL3 silencing stabilizes PINK1 mRNA via m6A modification

Fig. 4. METTL3 silencing stabilizes PINK1 mRNA via m6A modification.

PINK1 overexpression affects cell senescence and mitophagy in BLM-induced hFLSs

METTL3 silencing delays cellular senescence and enhances mitophagy by upregulating PINK1 in BLM-induced hFLSs

Fig. 5. METTL3 silencing delays cellular senescence and enhances mitophagy by upregulating PINK1 in BLM-induced hFLSs.

Senescent FLSs induced cartilage degradation in vivo

Fig. 6. Senescent FLSs induce cartilage degradation in vivo.

Discussion

Methods

Experimental animals

Construction of the rat model of TMJOA

Micro-CT detection

Detection of the head withdrawal threshold (HWT)

H&E staining

Safranin O staining

Alcian blue (AB) staining

Immunohistochemistry (IHC)

Isolation of primary FLSs

Cell culture

Cell treatment

Co-culture setup

RT-qPCR

RNA stability assay

Western blot

Immunofluorescence (IF) staining

ELISA

SA-β-galactosidase staining

Tetramethylrhodamine methyl ester (TMRM) staining

Transmission electron microscopy (TEM)

RNA methylation immunoprecipitation (MeRIP-qPCR)

Statistics and Reproducibility

Ethics approval and consent to participate

Reporting summary

Supplementary information

Acknowledgements

Author contributions

Peer review

Peer review information

Data availability

Competing interests

Footnotes

Contributor Information

Supplementary information

References

Associated Data

Supplementary Materials

Data Availability Statement

ACTIONS

PERMALINK

RESOURCES

Similar articles

Cited by other articles

Links to NCBI Databases

METTL3 silencing stabilizes PINK1 mRNA via m⁶A modification

Fig. 4. METTL3 silencing stabilizes PINK1 mRNA via m⁶A modification.