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OGA promotes human dental pulp stem cell senescence and inhibits mitophagy by inhibition of O-GlcNAcylation of KLF2

BMC Oral Health volume 25, Article number: 595 (2025) Cite this article

Abstract

Background

Dental pulp stem cells (DPSCs) aging impedes its application in tooth regeneration techniques, involving abnormal mitophagy. O-GlcNAcylation is a post-translational modification that regulates various cellular processes. Here, we aimed to investigate the role of O-GlcNAcylation in mitophagy and senescence.

Methods

DPSCs were cultured and passaged in vitro, and the 7th (p7) and 15th (p15) generation cells were collected. OGA and KLF2 were knocked down in p15 cells. Cell senescence was evaluated using senescence associated β-galactosidase staining, enzyme-linked immunosorbent assay, and western blotting; mitophagy was evaluated using western blotting. The regulation of OGA on the O-GlcNAcylation of KLF2 was analyzed using immunoprecipitation and western blotting.

Results

The results showed that p15 cells were more senescent than p7 cells and had poor mitophagy, with the higher expression of OGA. Knockdown of OGA inhibited senescence and promoted mitophagy in DPSCs. Moreover, silencing of KLF2 reversed the effects on senescence and mitophagy mediated by OGA knockdown. Additionally, OGA suppressed the O-GlcNAcylation of KLF2 at S177 site and thus reduced its stability.

Conclusion

Silencing of OGA promotes mitophagy and inhibits DPSC senescence by promoting the O-GlcNAcylation of KLF2, suggesting a novel mechanism underlying DPSC senescence.

Peer Review reports

Introduction

Dental pulp stem cells (DPSCs), a type of mesenchymal stem cell residing within the pulp cavity of teeth, play a critical role in pulp homeostasis and injury repair [1]. They have shown immense promise in tissue engineering and regenerative dentistry due to their multipotent differentiation potential [2, 3]. Clinical application of DPSCs requires a large number of cells, and it is necessary to expand the population through in vitro culture. However, the number of cell divisions is restricted due to cell senescence in this process [4]. Senescence in DPSCs is a complex process, which inhibits their proliferative and differentiation capabilities, leading to a reduction in the regenerative potential of DPSCs [5]. With the prevalence of dental diseases, understanding the mechanisms of DPSC aging has become a critical area of research in regenerative dentistry.

Mitophagy is a selective degradation process crucial for maintaining cellular homeostasis [6]. It is responsible for the removal of damaged mitochondria, thereby preventing the accumulation of reactive oxygen species and the induction of apoptosis [7]. Aging cells usually accumulate many dysfunctional mitochondria. Emerging evidence suggests that the impairment of mitophagy is associated with cellular aging [8]. Nevertheless, whether mitophagy is involved in delaying the senescence of DPSCs remains not understood.

O-GlcNAcylation is a dynamic and reversible post-translational modification involving the addition of O-linked N-acetylglucosamine (O-GlcNAc) to serine or threonine residues of proteins [9, 10]. This process is regulated by two enzymes: O-GlcNAc transferase (OGT) and O-GlcNAcase (OGA). It has been reported that O-GlcNAcylation is a key regulator of cell physiological processes by modulating protein stability and location, mitochondrial function, chromatin remodeling, and signal transduction [11]. Notably, alterations in O-GlcNAcylation have been associated with cellular aging [12]. However, whether O-GlcNAcylation is related to DPSC senescence needs further study.

Kruppel-like factor 2 (KLF2) is a zinc-finger transcription factor that has been implicated in the regulation of endothelial function and vascular homeostasis [13]. KLF2 has been shown to regulate several cellular processes, such as inflammation, proliferation, apoptosis, and senescence [14,15,16]. Notably, KLF2 has the ability to modulate autophagy and mitophagy in DPSCs [17, 18]. However, the role of KLF2-mediated mitophagy in the aging of DPSCs and its potential regulation by O-GlcNAcylation remains largely unexplored.

In this study, we investigated the role of O-GlcNAcylation in the regulation of DPSC senescence, focusing on the interplay between OGA, KLF2, and mitophagy. Our findings suggest that OGA can influence the aging process of DPSCs by affecting mitophagy through the modulation of O-GlcNAcylation of KLF2. This study will provide novel insights into the molecular mechanisms underlying DPSC senescence.

Materials and methods

Primary DPSC isolation and culture

This study was approved by the Ethics Committee of Ningbo Stomatology Hospital (approval number: NBKQYY2025LS-01). Human dental pulp tissues were collected from healthy donors (age 18–26 years old) who received the third molar extraction in our hospital. All participants provided written informed consent. DPSCs were isolated as previously described [19, 20]. In brief, pulp tissues were cut into small pieces and digested with 0.3% type I collagenase (Sigma-Aldrich, St. Louis, MO, USA) and 0.4% dispase (Sigma-Aldrich) for 30 min at 37 °C. The single-cell suspension was collected and centrifuged at 112 ×g for 5 min. Primary DPSCs were cultured at alpha modification of minimal essential medium (α-MEM; Gibco, Grand Island, NY, USA) supplemented with 10% fetal bovine serum (FBS; Gibco) and 1% penicillin/streptomycin (Invitrogen, Carlsbad, CA, USA) at an incubator containing 5% CO2 at 37 °C.

Cell identification

To induce multilineage differentiation, DPSCs were seeded into six-well plates one night before induction, and the basal medium was replaced with low glucose- Dulbecco’s modification of Eagle’s medium (LG-DMEM; Gibco) supplemented with 10% FBS. The induction methods were performed as previously described [21]. To induce osteogenic differentiation, the cells were cultured in LG-DMEM supplemented with 15% FBS, 1% penicillin/streptomycin, 0.1 mM L-ascorbic acid phosphate, 0.01 μM dexamethasone, 0.01 mM β-glycerophosphate, and 2 mM L-glutamine. After 14 days, alkaline phosphatase (ALP) staining was conducted to evaluate osteogenic differentiation. After 21 days, Alizarin Red S (ARS) staining was also conducted to evaluate osteogenic differentiation. To induce adipogenic differentiation, the cells were cultured in LG-DMEM supplemented with 10% FBS, 1 μM dexamethasone, 10 μg/mL insulin, 200 μM indomethacin, and 0.5 mM 3-Isobutyl-1-methylxanthine (IBMX). After 21 days, Oil Red O staining was performed to observe lipid droplets. Undifferentiated DPSCs were also stained by ALP, ARS, and Oil Red O dyes. All media additives were acquired from Sigma-Aldrich. ALP, ARS, and Oil Red O stain kits were acquired from Beyotime (Shanghai, China). The cell culture medium was replaced every 2 or 3 days.

To detect the cell surface markers, DPSCs after osteogenic differentiation or not were incubated with monoclonal antibodies against CD45, CD90, and CD105 (BioLegend, San Diego, CA, USA) on ice in the dark for 30 min. A flow cytometer (Beckman Coulter, Miami, FL, USA) was used to detect their levels. Data were analyzed using the FlowJo software.

Cell passage

To analyze cellular senescence, DPSCs underwent passage when they were grown until about 80% confluence. The cells at passage 7 (p7) and 15 (p15) were collected.

Senescence associated β-galactosidase (SA‐β‐gal) staining

DPSC senescence was evaluated using the lysosomal β-gal staining kit (Beyotime). Briefly, the cells in the six-well plates were washed with PBS and fixed with 1 mL β-gal staining fixative at room temperature for 15 min. Next, the cells were incubated with 1 mL staining working solution at 37 °C overnight. The results were observed under a light microscope.

Enzyme-linked immunosorbent assay (ELISA)

The activity of telomerase reverse transcriptase (TERT) was detected using the human TERT ELISA kit (Yanqi, Shanghai, China). DPSCs were suspended in PBS to adjust cell density to 106 cells/mL. Then, the cells were broken through repeated freezing and thawing. After centrifuging at 3000 rpm for 20 min, the supernatant was collected for analysis. The sample (50 μL) was incubated with 100 μL horseradish peroxidase (HRP)-labeled antibody working solution at 37 °C for 60 min, followed by washing five times. After incubating with the substrate in the dark for 15 min at 37 °C, the reaction was stopped, and the absorbance was measured at 450 nm.

5-ethynyl-2’-deoxyuridine (EdU) assay

The proliferation of DPSCs was evaluated using the kFluor647-EdU detection kit (KeyGen, Nanjing, China). DPSCs (2 × 105 cells) in six-well plates were grown overnight and labeled with 20 μM EdU solution. Subsequently, the cells were fixed with 4% paraformaldehyde protecting the light and permeated permeable solution for 15 min. Click-iT reaction mixture was added to each well to incubate with the cells for 30 min. The cell nucleus was stained with DAPI for 15 min. After washing with PBS, the cells were visualized using a fluorescence microscope.

Western blotting

Cell lysate was collected after lysing the DPSCs or HEK293T cells using the radio-immunoprecipitation assay buffer. Protein concentration was detected using a BCA protein assay kit (Beyotime). Equal amounts of proteins were loaded for sodium dodecyl sulfate-polyacrylamide gel electrophoresis and then transferred to polyvinylidene fluoride membranes. The membranes were blocked with 5% non-fat milk for 1 h, incubated with primary antibodies overnight at 4 °C, and continued to incubate with HRP-labeled secondary antibodies for 1 h. Enhanced chemiluminescent (ECL) kit (KeyGen) was used to develop the protein bands.

The antibodies (Abcam, Cambridge, MA, USA) used were as follows: anti-p53 (ab32389), anti-p21 (ab109520), anti-p16 (ab51243), anti-β-actin (ab8227), anti-Pink1 (ab216144), anti-LC3B (ab48394), anti-Beclin 1 (ab207612), anti-O-GlcNac/RL2 (ab93858), anti-OGT (ab177941), anti-OGA (ab124807), anti-KLF2 (ab236507), goat anti-rabbit (ab6721), and goat anti-mouse (ab205719).

Cell transfection

OGA short hairpin RNA (shOGA, targeting sequences: 5’-GTGTCTCAGTCTCCATATTTA-3’), KLF2 short hairpin RNA (shKLF2, targeting sequences: 5’-TTGTGATGCCTTGTGAGAAAT-3’), and negative control short hairpin RNA (shNC, 5’-CCTAAGGTTAAGTCGCCCTCG-3’) were synthesized by Invitrogen. In p7 and p15 cells, transfection was performed 48 h after passage. DPSCs were plated into six-well plates and transfected with shRNAs using Lipofectamine 2000 (Invitrogen) for 48 h following the manufacturer’s protocols.

Quantitative PCR (qPCR)

Total RNA was extracted from DPSCs using the RNA easy fast tissue/cell kit (Tiangen, Beijing, China). Reverse transcription and qPCR were performed using the FastKing one step RT-qPCR kit (SYBR) (Tiangen). The expression levels of RNAs were calculated using the 2−ΔΔCt method by normalizing to β-actin. The specific primer sequences were as follows. OGA forward: 5’-CATAGGATGTTTTGGCGAGAGAT-3’; reverse 5’-GGTGAGATCGCATAGATGAACTC-3’, KLF2 forward: 5’-CTACACCAAGAGTTCGCATCTG-3’; reverse 5’-CCGTGTGCTTTCGGTAGTG-3’, β-actin forward: 5’-CATGTACGTTGCTATCCAGGC-3’; reverse 5’-CTCCTTAATGTCACGCACGAT-3’.

Co-immunoprecipitation (co-IP)

The interaction between OGA and KLF2 proteins was evaluated using the co-IP kit (GeneSeed, Guangzhou, China). HEK293T cells (ATCC, Manassas, VA, USA) were used here due to their high transfection efficiency, stable properties, and ease of culture. They were maintained in DMEM supplemented with 10% FBS at 37 °C with 5% CO2. Plasmids with HA-tagged OGA and His-tagged KLF2 were transfected into HEK293T cells. Afterward, the cells were lysed using the binding buffer containing the protease inhibitor on ice. The supernatant was collected after centrifugation. Protein A + G magnetic beads were incubated with 5 μg anti-HA, anti-His, or anti-IgG at 4 °C for 2 h. The supernatant was incubated with pre-treated magnetic beads at 4 °C overnight. After washing and elucidating the mixture, the levels of HA-OGA and His-KLF2 were measured using western blotting.

O-GlcNAcylation analysis

To detect the O-GlcNAcylation levels of KLF2 (KLF2-RL2), the immunoprecipitation (IP) kit (Beyotime) was used. DPSCs transfected with shNC or shOGA were lysed using the IP buffer containing protease inhibitor cocktail on ice, and the cell lysate was collected by centrifuging. Protein A + G magnetic beads were pre-coated with 5 μg anti-KLF2 for 1 h, and incubated with cell lysate for 2 h. The magnetic beads were washed with IP buffer, and the proteins were elucidated and collected. Western blotting was conducted to measure KLF2-RL2 levels with the anti-RL2 primary antibody.

The O-GlcNAcylation sites in KLf2 were predicted using the DictyOGlyc-1.1 database (https://services.healthtech.dtu.dk/services/DictyOGlyc-1.1/).

To analyze the modifying sites, wild-type (WT) KLF2 plasmids were constructed. KLF2 mutated plasmids (Thr or Ser to Ala) were constructed using the QuickMutation site-directed mutagenesis kit (Beyotime). These plasmids were transfected into the HEK293T cells using Lipofectamine 2000. The levels of KLF2 and KLF2-RL2 were measured using western blotting.

Cycloheximide (CHX) treatment

DPSCs after shNC and shOGA transfection were treated with 100 μM CHX (Sigma-Aldrich) for 0, 8, 16, and 24 h. The levels of KLF2 were examined using western blotting.

Statistical analysis

Data were analyzed using the GraphPad Prism 7 software and were represented as means ± standard deviation (SD). Differences were analyzed using the Student’s t-test (two groups) and one-way ANOVA followed by Tukey’s post hoc test (more than two groups). P < 0.05 was considered statistically significant.

Results

Multilineage differentiation of DPSCs

To identify the human DPSCs, we measured the osteogenic and adipogenic differentiation capabilities. We then measured the osteogenic and adipogenic differentiation capabilities. After the cells were cultured in osteogenic differentiation induction medium, ALP staining assay showed that the cells had ALP activity (Fig. 1A), and ARS staining results showed that the cells had obvious mineralized nodule deposition (Fig. 1B). In addition, DPSCs were cultured in adipogenic differentiation induction medium, and Oil Red O staining was performed. The results showed that the cells had obvious lipid droplets (Fig. 1C). However, DPSCs without osteogenic induction could not be stained by ALP, ARS and Oil Red O (Fig. 1A-C). Besides, the surface markers of stem cells were measured using flow cytometry. As shown in Fig. 1D, DPSCs expressed CD90 and CD105, but rarely expressed CD45. Additionally, DPSCs after osteogenic differentiation did not express CD90, CD105, and CD45. Taken together, the cells can differentiate into osteoblasts and lipoblasts, and express stem markers, suggesting they are DPSCs.

Fig. 1
figure 1

Multilineage differentiation of DPSCs. (A) ALP and (B) ARS staining assay were performed to analyze osteogenic differentiation capability of osteogenic-induced or undifferentiated DPSCs. (C) The adipogenic differentiation of adipogenic-induced or undifferentiated DPSCs was evaluated using Oil Red O staining. (D) Stem cell surface markers of osteogenic-induced or undifferentiated DPSCs were detected using flow cytometry

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Mitophagy is involved in DPSC aging in vitro

To identify whether the DPSCs senescence occurs during passage, we conducted continuous natural passage of the cells and collected p7 and p15 cells. The results of SA-β‐gal staining showed that the β‐gal-positive cells were more in the p15 group than that in the p7 group (Fig. 2A). The TERT activity was lower in the p15 group than that in the p7 group (Fig. 2B). The senescence markers including p53, p21, and p16 were measured by western blotting, and the results showed that their levels were increased in the p15 cells, compared with p7 cells (Fig. 2C). Aging cells inhibit their own proliferation [22]. DPSC proliferation was evaluated using EdU assay. We found that the proliferation capability was suppressed in the p15 cells, compared with the p7 cells (Fig. 2D). Mitophagy disorder occurs in senescent cells. To investigate whether mitophagy is involved in DPSC senescence, the levels of related markers were measured using western blotting. It was observed that the levels of Pink1, parkin, Beclin-1, and LC3B-II/LC3B-1 ratio were lower in the p15 cells than that in the p7 cells (Fig. 2E). The results demonstrated that p15 cells are more senescent than p7 cells, and mitophagy is inhibited in senescent DPSCs.

Fig. 2
figure 2

Mitophagy is involved in DPSC aging in vitro. (A) DPSC senescence was evaluated using SA-β‐gal staining. Scale bar = 100 μm. (B) TERT activity was measured using ELISA. (C) Western blotting was carried out to examine the protein levels of p53, p21, and p16. (D) The proliferation of DPSCs was assessed using EdU assay. EdU-positive cells (red) were quantified. The nucleus (blue) was stained by DAPI. Scale bar = 100 μm. (E) Western blotting was carried out to examine the protein levels of mitophagy markers (Pink1, parkin) and autophagy markers (LC3B, Beclin-1). Data were analyzed using Student’s t-test. **P < 0.01

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OGA expression was increased in p15 cells

O-GlcNAcylation can regulate several cellular processes. Nevertheless, whether O-GlcNAcylation is involved in DPSC behaviors remains unclear. We measured total O-GlcNAc levels in DPSCs and found that its levels were lower in the p15 group than that in the p7 group (Fig. 3). Next, the two enzymes OGA and OGT that regulate O-GlcNAcylation were measured. OGA expression was elevated in p15 cells, compared with p7 cells, while no difference in OGT levels was found in DPSCs in these two groups (Fig. 3). Thus, we considered that the aging of DPSCs involves the inhibition of O-GlcNAcylation, which is modulated by high expression of OGA.

Fig. 3
figure 3

OGA expression was increased in p15 cells. The levels of O-GlcNAc, OGT, and OGA were measured by western blotting. Data were analyzed using Student’s t-test. **P < 0.01. ns: no significance

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Knockdown of OGA inhibits DPSC senescence and promotes mitophagy

Next, the effect of OGA on cellular phenotypes was investigated after its knocking down in p15 cells. OGA expression was downregulated in shOGA-transfected DPSCs, compared with shNC-transfected DPSCs (Fig. 4A). β-gal-positive cells in the p15 group were reduced after OGA silence (Fig. 4B). The activity of TERT was enhanced after OGA knockdown (Fig. 4C). Moreover, interfering with OGA downregulated the levels of p53, p21, and p16 (Fig. 4D). Additionally, the inhibition of DPSC proliferation in p15 cells was counteracted by knockdown of OGA (Fig. 4E). Besides, silencing of OGA in p15 cells upregulated Pink1, parkin, and Beclin-1 levels, as well as LC3BII/LC3BI ratio (Fig. 4F). The results indicates that knockdown of OGA contributes to attenuating senescence of DPSCs and facilitating mitophagy.

Fig. 4
figure 4

Knockdown of OGA inhibits DPSC senescence and promotes mitophagy. (A) OGA expression in DPSCs at p15 was detected using qPCR after transfection. (B) DPSC senescence was evaluated using SA-β‐gal staining. Scale bar = 100 μm. (C) TERT activity was measured using ELISA. (D) The protein levels of p53, p21, and p16 were examined by western blotting. (E) EdU assay was performed to assess DPSC proliferation. Scale bar = 100 μm. (F) The protein levels of mitophagy markers (Pink1, parkin) and autophagy markers (LC3B, Beclin-1) were examined by western blotting. Data were analyzed using Student’s t-test in (A) and one-way ANOVA in (B-F). **P < 0.01

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Knockdown of OGA stabilizes KLF2 protein by promoting O-GlcNAcylation

We chose KLF2 for molecular mechanism study since KLF2 is an autophagy and mitophagy regulator in DPSCs [17]. Due to the inhibition effect of OGA on O-GlcNAcylation, we analyzed the regulation of OGA on the O-GlcNAcylation of KLF2. We found that knockdown of OGA promoted the O-GlcNAcylation of KLF2, and concurrently elevated the protein levels of KLF2 (Fig. 5A). Next, we identified that OGA protein interacted with KLF2 protein (Fig. 5B). Potential O-GlcNAcylation sites in KLF2 were predicted, with three of the most likely sites being T141, T173, and S177 (Fig. 5C and D). Then, the modifying sites were confirmed. As compared to the WT group, S177A reduced the protein levels of KLF2 and its O-GlcNAcylation, while T141A and T173A did not affect their levels (Fig. 5E), suggesting that S177 site is the O-GlcNAcylation site in KLF2. Moreover, we found that silencing of OGA enhanced the protein stability of KLF2 (Fig. 5F). In summary, OGA knockdown promotes the O-GlcNAcylation of KLF2 at S177 site, enhances KLF2 stability, and elevates KLF2 protein levels.

Fig. 5
figure 5

Knockdown of OGA stabilizes KLF2 protein by promoting O-GlcNAcylation. (A) After OGA knocking down, KLF2 protein and O-GlcNAcylation levels were measured by western blotting. (B) The interaction between OGA and KLF2 was verified by exogenous co-IP in HEK293T cells. (C) Possible O-GlcNAcylation sites in KLF2 were predicted using the DictyOGlyc-1.1 database. (D) Three most likely modification sites in KLF2. (E) O-GlcNAcylation modification sites were confirmed by IP and western blotting in HEK293T cells. (F) KLF2 protein stability was measured using western blotting after CHX treatment for indicated times. Data were analyzed using Student’s t-test in (A), one-way ANOVA in (E), and two-way ANOVA in (F). **P < 0.01. ns: no significance

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OGA affects DPSC senescence and mitophagy by modulating KLF2

To explore the impact of KLF2 on cellular behaviors, shKLF2 was transfected into p15 cells, and KLF2 expression was decreased (Fig. 6A). The reduction of β-gal-positive cells and the enhancement of TERT activity caused by OGA knockdown were counteracted by KLF2 knockdown (Fig. 6B and C). Silencing of KLF2 reversed the downregulation of p53, p21, and p16 protein levels induced by knockdown of OGA (Fig. 6D). Additionally, interfering of OGA promoted DPSC proliferation, which was abrogated by KLF2 knockdown (Fig. 6E). Moreover, silencing of KLF2 reduced Pink1, parkin, and Beclin-1 levels and LC3BII/LC3BI ratio in p15 cells after OGA knockdown (Fig. 6F). Taken together, knockdown of OGA inhibits DPSC senescence and promotes mitophagy by elevating KLF2 expression.

Fig. 6
figure 6

OGA affects DPSC senescence and mitophagy by modulating KLF2. (A) KLF2 expression in DPSCs at p15 was detected using qPCR after transfection. (B) DPSC senescence was evaluated using SA-β‐gal staining. Scale bar = 100 μm. (C) TERT activity was measured using ELISA. (D) The protein levels of p53, p21, and p16 were examined by western blotting. (E) EdU assay was performed to assess DPSC proliferation. Scale bar = 100 μm. (F) The protein levels of mitophagy markers (Pink1, parkin) and autophagy markers (LC3B, Beclin-1) were examined by western blotting. Data were analyzed using Student’s t-test in (A) and one-way ANOVA in (B-F). **P < 0.01

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Discussion

DPSCs, similar to mesenchymal stem cells, have high differentiation, proliferation, and regeneration capabilities, and can synthesize a dentin-pulp complex in vivo and amplify into single colonies in vitro [23]. In recent years, DPSC senescence has received more and more attention. Age is a risk factor for DPSC senescence. With the increase of age, the proliferation and differentiation of DPSCs gradually decline, which is easy to cause diseases such as periodontitis, resulting in tooth loss [24]. In addition, when DPSCs need to be applied clinically, in vitro replicative senescence is the main limiting factor [4]. In this study, since there are currently no suitable culture methods to prevent cell senescence, we focused on the mechanism of DPSC senescence under long-term culture conditions.

Mitochondrial functions and dynamics are closely linked to the differentiation of DPSCs, which contributes to understanding their senescence and self-renewal [25]. Clearance of damaged mitochondria through mitophagy helps restore normal mitochondrial function. Moreover, intercellular reactive oxygen species (ROS), which are produced by mitochondria, is a main cause of senescence [26]. In our work, we found that p15 DPSCs were more senescent than p7 DPSCs, and their mitophagy ability was reduced, indicating a close relationship between inhibiting aging and activating mitophagy, consistent with several previous studies [27, 28].

Growing evidence has reported that O-GlcNacylation regulates cellular senescence [29, 30]. This regulatory effect involves abnormal expression of OGA and/or OGT. OGT deletion inhibits premature aging of human fibroblasts, while OGT overexpression accelerates aging [31]. OGA expression is elevated in the process of delaying brain aging caused by meclofenoxate [32]. However, whether they regulate DPSC senescence remains unknown. In this study, we found that OGA expression was increased in p15 cells, while OGT expression was not altered, suggesting that OGA may be involved in DPSC senescence. Interestingly, inhibition of OGA by Thiamet-G treatment can promote mitophagy [33]. Herein, we identified that knockdown of OGA suppressed the senescence of DPSCs and facilitated mitophagy, suggesting an important role of OGA in DPSC aging.

We further discovered that OGA knockdown stabilized KLF2 by promoting its O-GlcNAcylation at S177 site, upregulating its expression level. KLF2 has been demonstrated to induce mitophagy and autophagy of DPSCs [17, 18, 34]. Nevertheless, whether KLF2 affects DPSC senescence is still unclear. In this study, we found that the knockdown of KLF2 reverses the effects of OGA knockdown on mitophagy and cellular aging. These findings expand our understanding of the multifaceted regulation of KLF2 and suggest a critical role for the OGA/KLF2 axis in DPSC senescence.

In conclusion, we demonstrated that silencing of OGA inhibits DPSC senescence by facilitating mitophagy, which is achieved through the promotion of KLF2 O-GlcNAcylation. Our findings give evidence that implicates O-GlcNAcylation as a key regulator of cellular senescence and highlights its potential to preserve the regenerative capacity of DPSCs, which will provide a new strategy for delaying the replicative senescence of DPSC in vitro.

Data availability

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

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  1. Engineering Laboratory for Biomaterials and Tissue Regeneration, Ningbo Stomatology Hospital, No.435, Xinxing Road, Wangchun Street, Haishu District, Ningbo, 315000, Zhejiang, China

    Yinhao Ding & Yan Ran

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All authors participated in the design, interpretation of the studies and analysis of the data and review of the manuscript. Y D drafted the work and revised it critically for important intellectual content and was responsible for the acquisition, analysis and interpretation of data for the work; Y R made substantial contributions to the conception or design of the work. All authors read and approved the final manuscript.

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Ding, Y., Ran, Y. OGA promotes human dental pulp stem cell senescence and inhibits mitophagy by inhibition of O-GlcNAcylation of KLF2. BMC Oral Health 25, 595 (2025). https://doiorg.publicaciones.saludcastillayleon.es/10.1186/s12903-025-05927-1

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BMC Oral Health

ISSN: 1472-6831

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