Published online Jun 26, 2026. doi: 10.4252/wjsc.119118
Revised: February 21, 2026
Accepted: April 2, 2026
Published online: June 26, 2026
Processing time: 157 Days and 2.2 Hours
Aging impairs dental pulp regeneration, but upstream mechanisms driving dental pulp stem cell (DPSC) vulnerability remain unclear. Selenium-dependent an
To determine whether SEPP1 and selenium supplementation protect DPSCs from oxidative senescence injury.
Single-cell RNA sequencing data from young and aged human dental pulp were integrated. Human DPSCs were exposed to hydrogen peroxide (100 μmol/L, 4 hours), then treated for 48 hours with recombinant SEPP1 (50 ng/mL), sodium selenite (100 nmol/L), or both. Senescence, proliferation, redox injury, and fer
Aged pulp showed reduced DPSC abundance and weakened selenium-related antioxidant signatures. Combined SEPP1 and sodium selenite treatment reduced senescence-associated changes, improved proliferation, and preserved stemness-related markers after oxidative injury. The combined treatment produced stronger viability rescue than either treatment alone. It also reduced ferrous iron accumulation, glutathione imbalance, glutathione peroxidase 4 downregulation, and lipid peroxidation. Ferrostatin-1 improved viability after oxidative injury, supporting ferroptosis-associated injury involvement. Transcriptome analysis identified FOXM1-centered cell cycle and antioxidant programs, and FOXM1 knockdown attenuated the protective effects. Aged human pulp tissues showed reduced SEPP1 and FOXM1 expression.
SEPP1 and selenium co-treatment mitigates oxidative senescence and ferroptosis-associated lipid peroxidation in DPSCs through a FOXM1-dependent protective program.
Core Tip: Aging-associated decline of dental pulp stem cells is linked to impaired selenium-dependent antioxidant defense. This study integrates single-cell transcriptomics, oxidative stress modeling, and human tissue validation to show that selenoprotein P and sodium selenite co-treatment reduces senescence and ferroptosis-associated lipid peroxidation in dental pulp stem cells. The combined treatment is more effective than either agent alone, and ferrostatin-1 rescue supports ferroptosis-related injury involvement. Transcriptomic and knockdown analyses identify a forkhead box protein M1-dependent protective program downstream of selenoprotein P/selenium support, highlighting a candidate strategy for preserving dental pulp vitality during aging.
- Citation: Zhang RQ, Chu WH, Bai JB, Lei YH, Zhou D. Selenoprotein P attenuates oxidative senescence and ferroptosis-associated lipid peroxidation in dental pulp stem cells through a FOXM1-dependent program. World J Stem Cells 2026; 18(6): 119118
- URL: https://www.wjgnet.com/1948-0210/full/v18/i6/119118.htm
- DOI: https://dx.doi.org/10.4252/wjsc.119118
Dental pulp is a specialized connective tissue enclosed within the rigid dentin chamber and is essential for maintaining tooth vitality by providing nutrition, innervation, and immune defense[1]. However, during aging, the pulp undergoes progressive struc
Among the cellular components of dental pulp, dental pulp stem cells (DPSCs) play a central role in tissue homeostasis and regeneration because of their self-renewal capacity and multilineage differentiation potential[4,5]. Accumulating evidence indicates that DPSC function declines with aging, as reflected by reduced proliferation, impaired odontogenic differentiation, and increased senescence-associated phenotypes[6-8]. These changes are accompanied by elevated oxidative stress and persistent inflammatory signaling, which further exacerbate stem cell dysfunction and limit regenerative outcomes[9,10]. Accordingly, identifying upstream regulators of DPSC redox balance and senescence is an important step toward delaying pulp aging and improving regenerative therapy in older patients.
The molecular basis of DPSC aging is increasingly understood as a coordinated process involving cell-cycle arrest, oxidative injury, and stress-response remodeling[11]. Dysregulation of core cell-cycle checkpoint pathways, including p16, p21, and p53, is consistently implicated in this process[12-14]. Oxidative stress is a major driver of DPSC senescence because the enclosed pulp chamber is particularly vulnerable to fluctuations in oxygen and nutrient supply[15,16]. Excessive reactive oxygen species (ROS) damage cellular macromolecules and activate stress signaling pathways that promote senescence-associated secretory phenotypes and chronic inflammation[17,18]. In addition, recent studies suggest that ferroptosis, an iron-dependent form of lipid peroxidation-driven cell death, may also contribute to pulp degeneration under stress conditions[19,20]. However, current antioxidant-based interventions remain only partially effective, highlighting the need to identify upstream regulators that coordinate redox homeostasis, senescence, and ferroptosis-associated injury in DPSCs.
Selenoprotein P (SEPP1, encoded by SELENOP) is a secreted glycoprotein that functions as the principal selenium transport protein and an important systemic antioxidant carrier[21]. By delivering selenium to peripheral tissues, SEPP1 supports the biosynthesis of key antioxidant selenoenzymes, including glutathione peroxidases (GPXs) and thioredoxin reductases, and thereby plays a central role in maintaining cellular redox homeostasis[21,22]. Emerging evidence further indicates that SEPP1 is critical for stem cell maintenance. In neural stem cells and muscle satellite cells, SEPP1-dependent selenium supply supports self-renewal, regenerative function, and resistance to oxidative exhaustion[23-25]. In parallel, SEPP1 has been increasingly linked to ferroptosis regulation, as SEPP1 deficiency can impair GPX4 synthesis and increase susceptibility to lipid peroxidation-driven injury[26,27]. Despite these findings in other tissues, whether SEPP1 plays a similar protective role in aging dental pulp remains unknown.
In this study, we investigated whether age-associated loss of SEPP1 contributes to oxidative vulnerability in DPSCs and whether restoring selenium support can mitigate senescence-associated injury. By integrating single-cell transcriptomic analysis of human dental pulp, an H2O2-induced oxidative stress model in DPSCs, bulk RNA sequencing, and human tissue validation, we examined the protective effects of SEPP1/selenite co-treatment on senescence, redox imbalance, and ferroptosis-associated lipid peroxidation. We further evaluated the downstream role of forkhead box protein M1 (FOXM1) using transcriptomic pathway analysis and small interfering RNA (siRNA)-mediated knockdown. Our findings identify a SEPP1/selenium-dependent, FOXM1-associated protective program in DPSCs and provide a mechanistic framework for targeting oxidative pulp aging.
This study was approved by the Medical Ethics Review Committee of Xiangya Hospital Central South University (Approval No. 2025122275), and written informed consent was obtained from all participants or their legal guardians. Human dental pulp tissues were collected from clinically indicated tooth extractions, including exfoliated deciduous teeth and permanent teeth (e.g., impacted third molars or premolars), and were grouped as young (9-25 years) or aged (50-67 years) for histological and molecular validation analyses. Only teeth with vital pulp tissue and without acute pulpitis, periapical lesions, or prior endodontic treatment were included. After extraction, pulp tissues were immediately placed in cold DMEM/F12 medium containing 1% penicillin-streptomycin and transported to the laboratory for processing. Donor metadata, including age, sex, tooth type, oral/pulp status, general health status, and assay allocation, are summarized in Supplementary Table 1.
Human DPSCs (Cyagen Biosciences, Catalog No. HUXDP-01001, CA, United States) were cultured in DMEM/F12 supplemented with 10% fetal bovine serum, 1% penicillin-streptomycin, and 1% L-glutamine in a humidified incubator (37 °C, 5% CO2). The culture medium was changed every 2-3 days. Cells were passaged with 0.25% trypsin-EDTA once they reached 80%-90% confluence.
For the acute oxidative stress/senescence-like injury model, DPSCs were exposed to 100 μM H2O2 for 4 hours. After H2O2 exposure, the medium was replaced, and cells were cultured for an additional 48 hours under the indicated treatment conditions. Recombinant human SEPP1 was used at 50 ng/mL, and sodium selenite was used at 100 nM. In the main treatment group (Trt), cells received SEPP1 + sodium selenite immediately after H2O2 challenge and throughout the 48 hours recovery period.
To clarify treatment attribution, additional control groups were included in CCK-8 optimization experiments: H2O2 + SEPP1 alone (50 ng/mL), H2O2 + sodium selenite alone (100 nM), and H2O2 + SEPP1 + sodium selenite (Supplementary Figure 1). For ferroptosis inhibitor rescue experiments, ferrostatin-1 (Fer-1, 10 μM) was added after H2O2 exposure and maintained for 48 hours before viability assessment (Supplementary Figure 2). SEPP1 and Fer-1 concentrations were determined based on preliminary CCK-8 optimization experiments.
Publicly available single-cell RNA-sequencing (scRNA-seq) datasets of young (GSM4365610 and GSM4998458) and aged (GSM8452933 and GSM8452934) human dental pulp were downloaded from the Gene Expression Omnibus database. Data processing and integration were performed using the Seurat package (v4.4) in R. Cells expressing fewer than 500 genes or more than 8000 genes, or with mitochondrial gene content > 20%, were excluded. Ribosomal genes were removed before downstream analysis.
Batch correction and integration were performed using the FindIntegrationAnchors and IntegrateData functions with 2000 variable features. Dimensionality reduction and clustering were conducted using uniform manifold approximation and projection. Cell clusters were annotated according to canonical marker genes. Visualization was generated using the plot1cell and scRNAtoolVis packages. Differentially expressed genes (DEGs) and pathway enrichment analyses were performed using Metascape.
Total RNA was extracted using TRIzol reagent (Invitrogen, CA, United States), and RNA integrity was assessed with an Agilent 2100 Bioanalyzer. Poly(A)+ mRNA was enriched using oligo(dT) magnetic beads, fragmented, and reverse-transcribed into cDNA. Sequencing libraries were constructed and sequenced on an Illumina HiSeq 2500 platform. Raw reads were filtered using Fastp (v0.18.0), and ribosomal RNA reads were removed by Bowtie2 (v2.2.8) alignment. Clean reads were aligned to the reference genome using StringTie (v1.3.1). DEGs were identified using DESeq2, and P values were adjusted for multiple testing using the Benjamini-Hochberg method. Genes with FDR-adjusted P values (Padj) < 0.05, FPKM ≥ 1, and |log2 fold change| ≥ 1 were considered significantly differentially expressed. The full bulk RNA-seq gene expression matrix (FPKM values) is provided in Supplementary Table 2. Gene Ontology and Kyoto Encyclopedia of Genes and Genomes enrichment analyses were performed with the ClusterProfiler package in R.
Total RNA was isolated using RNAiso Plus (Takara, Japan), and RNA concentration was measured using a NanoDrop spectrophotometer (Thermo Fisher Scientific, MA, United States). cDNA was synthesized using a reverse transcription kit (Roche, Switzerland). Quantitative real-time polymerase chain reaction (qPCR) was performed on an ABI Prism 7700 system (Applied Biosystems, CA, United States) using SYBR Green chemistry. Relative mRNA expression levels were calculated by the 2-ΔΔCt method, with ACTB acting as the internal control. Primer sequences are provided in Supple
Cells or tissue samples were lysed in RIPA buffer supplemented with protease and phosphatase inhibitors (Thermo Fisher Scientific, MA, United States). Protein concentration was measured using a BCA protein assay. Equal amounts of protein (20-40 μg) were separated by sodium-dodecyl sulfate gel electrophoresis and transferred to polyvinylidene fluoride membranes. Membranes were blocked with 5% non-fat milk in Tris-buffered saline with Tween for 1 hour at room temperature and incubated overnight at 4 °C with primary antibodies (Supplementary Table 4). After Tris-buffered saline with Tween washes, membranes were incubated with HRP-conjugated secondary antibodies for 1 hour at room temperature. Protein bands were visualized using an enhanced chemiluminescence detection system.
Human dental pulp tissues were fixed, paraffin-embedded, and sectioned. After deparaffinization and rehydration, antigen retrieval was performed in 0.01 mol/L sodium citrate buffer (98 °C, 18 minutes). Sections were treated with 3% H2O2 to block endogenous peroxidase activity, permeabilized with 0.25% Triton X-100, and blocked with 5% bovine serum albumin. Sections were incubated with primary antibodies (Supplementary Table 4) overnight at 4 °C. For immunohistochemistry, staining was developed using a DAB detection kit (Zsbio, China). For immunofluorescence, Alexa Fluor-conjugated secondary antibodies were used, and nuclei were counterstained with DAPI. Images were acquired using a Zeiss microscope under identical exposure settings within each comparison group.
siRNAs targeting FOXM1 and a negative control siRNA were purchased from RiboBio (Guangzhou, Guangdong Province, China). DPSCs were transfected using Lipofectamine 3000 (Thermo Fisher Scientific, MA, United States) at 50%-70% confluence. siRNA-lipid complexes were prepared in Opti-MEM and added to cells at a final siRNA concentration of 20-50 nM. After 6 hours, the transfection medium was replaced with complete culture medium. Cells were collected 48-72 hours later for qPCR, western blotting, and functional assays.
For the CCK-8 assay, DPSCs were seeded into 96-well plates and treated as indicated. At each time point, CCK-8 reagent (10% v/v; Dojindo, Japan) was added and incubated for 3 hours at 37 °C. Absorbance was measured at 450 nm using a microplate reader. For the EdU assay, cells were incubated with 50 μM EdU (RiboBio, Guangzhou, Guangdong Province, China) for 12 hours, fixed with 4% paraformaldehyde, permeabilized with 0.5% Triton X-100, and stained using the Apollo reaction cocktail according to the manufacturer’s instructions. Nuclei were counterstained with DAPI, and images were captured by fluorescence microscopy. EdU-positive nuclei were quantified as a percentage of total nuclei.
Cellular senescence was assessed using an SA-β-Gal Staining Kit (Beyotime, Shanghai, China). Cells were fixed with 4% paraformaldehyde for 15 minutes and incubated with staining solution overnight at 37 °C (without CO2). SA-β-gal-positive cells were identified by blue staining and imaged using a light microscope (Zeiss). The percentage of positive cells was quantified from randomly selected fields.
DNA strand breaks were detected using an In Situ Cell Death Detection Kit (Roche, Switzerland). Cells were permeabilized with proteinase K (20 μg/mL) for 15 minutes, rinsed with phosphate buffered saline, and incubated with TdT enzyme/dUTP reaction mixture for 1 hour at 37 °C in the dark. After DAPI counterstaining, fluorescence images were obtained using a fluorescence microscope.
To evaluate ferroptosis-associated oxidative injury, iron homeostasis, lipid peroxidation, and glutathione (GSH) metabolism were assessed as follows.
Lipid ROS (C11-BODIPY 581/591): Cells were incubated with 5 μM C11-BODIPY 581/591 (Invitrogen, CA, United States) for 30 minutes at 37 °C. Oxidized (green) and reduced (red) fluorescence signals were captured by fluorescence microscopy, and the oxidized/reduced fluorescence ratio was used as an index of lipid peroxide accumulation.
Labile Fe2+ (FerroOrange): Cells were stained with 1 μM FerroOrange (Dojindo, Japan) for 30 minutes at 37 °C in the dark and imaged immediately (excitation 561 nm, emission 580 nm). Fluorescence intensity was quantified to estimate intracellular labile Fe2+ levels.
GSH/oxidized GSH assay: Intracellular reduced GSH and oxidized GSH (GSSG) were measured using a commercial assay kit (Beyotime, Shanghai, China). Total GSH and GSSG were quantified via spectrophotometry at 412 nm, and GSH levels were calculated by subtraction. The GSH/GSSG ratio was used to evaluate GSH redox balance.
Malondialdehyde: Malondialdehyde (MDA) levels were measured using an MDA assay kit (Solarbio, China). Cell lysates were incubated with thiobarbituric acid at 100 °C for 60 minutes, and absorbance was read at 532 nm and 600 nm.
Data were analyzed using R (including the dplyr package) and GraphPad Prism. Results are presented as mean ± SD from at least three independent experiments. Differences between two groups were analyzed using a Student’s t test, and comparisons among multiple groups were analyzed using one-way ANOVA followed by an appropriate post hoc test. A two-sided P value < 0.05 was considered statistically significant.
The statistical methods of this study were reviewed by a researcher with biomedical statistics training, and all analyses were performed in accordance with the study design and data distribution characteristics.
To systematically characterize the cellular landscape of dental pulp aging, we integrated public scRNA-seq datasets from young and aged human pulp tissues (GSM4365610 and GSM4998458 for young pulp; GSM8452933 and GSM8452934 for aged pulp). After quality control and batch integration, a total of 22915 cells were retained for downstream analysis. Uniform manifold approximation and projection clustering identified the major pulp cell populations, including DPSCs, fibroblasts, endothelial cells, immune cells, and other stromal populations (Figure 1A and B).
Comparative compositional analysis revealed an age-associated contraction of the DPSC compartment in aged pulp relative to young pulp (approximately 6% vs 10%), accompanied by broader shifts in stromal and immune cell com
To dissect the molecular basis of the decreased DPSC population, we next analyzed DEGs and pathway signatures within the DPSC cluster. Aged DPSCs exhibited downregulation of genes involved in redox regulation and selenium metabolism, including SEPP1/SELENOP-associated pathways and downstream selenoprotein antioxidant programs (Figure 1D and E). Enrichment analysis further indicated suppression of selenium-dependent antioxidant defense and oxidative stress buffering pathways in aged DPSCs (Figure 1F). Together, these data suggest that erosion of selenium-related antioxidant capacity is a prominent molecular feature of DPSC aging.
Based on the transcriptomic signature of selenium deficiency in aged DPSCs, we next tested whether restoring SEPP1/selenite support could protect DPSCs from oxidative injury. We established an acute oxidative stress/senescence-like model by exposing DPSCs to 100 μM H2O2 for 4 hours (Figure 2A). In this experimental workflow, untreated cells served as the blank group, H2O2-treated cells were designated as the senescence control (Ctrl), and cells receiving recombinant SEPP1 (50 ng/mL) plus sodium selenite (100 nM) after H2O2 exposure were defined as Trt.
Morphologically, H2O2-treated DPSCs displayed enlarged, flattened, and irregular cell shapes, consistent with a senescent phenotype, whereas Trt cells retained a more spindle-like morphology resembling the blank group (Figure 2B). SA-β-gal staining further confirmed a marked increase in senescent cells in the Ctrl group, which was significantly reduced by SEPP1/selenite co-treatment (Figure 2B and C).
At the molecular level, western blotting showed that H2O2 exposure significantly increased the senescence-associated proteins P53, P21, and P16, while decreasing SEPP1 and the proliferation marker proliferating cell nuclear antigen (Figure 2D and E). SEPP1/selenite co-treatment effectively reversed these changes, indicating attenuation of stress-induced senescence signaling. Consistent with these findings, EdU staining demonstrated that DNA synthesis was strongly suppressed in Ctrl cells but was significantly restored in Trt cells (Figure 2F and G). Likewise, CCK-8 assays showed that SEPP1/selenite co-treatment improved cell viability and growth after oxidative injury (Figure 2H).
To further clarify treatment attribution, we compared SEPP1 alone (50 ng/mL), sodium selenite alone (100 nM), and SEPP1 + sodium selenite co-treatment under the same H2O2 injury conditions. CCK-8 assays showed that the co-treatment group exhibited the strongest viability rescue, whereas single-agent treatment produced partial protection (Supplementary Figure 1). These results support the use of SEPP1/selenite co-treatment in subsequent experiments.
Finally, we evaluated stemness-related markers and DNA damage responses. H2O2 exposure reduced the expression of OCT4 and THY1 and increased the DNA damage marker γH2AX, whereas Trt treatment partially restored OCT4 and THY1 and significantly reduced γH2AX levels (Figure 2I and J). Collectively, these data demonstrate that SEPP1/selenite co-treatment protects DPSCs from oxidative stress-induced senescence and preserves functional competence.
Given that GPX4 is a key selenium-dependent enzyme that suppresses ferroptosis-associated lipid peroxidation, we next examined whether SEPP1/selenite co-treatment affects iron-redox homeostasis in H2O2-injured DPSCs. FerroOrange staining showed that H2O2 exposure markedly increased intracellular labile Fe2+ levels, whereas Trt treatment significantly reduced ferrous iron accumulation (Figure 3A and B), indicating partial restoration of iron homeostasis.
We then assessed GSH metabolism and lipid peroxide burden. H2O2-treated DPSCs showed a pronounced disruption of GSH redox balance, characterized by decreased GSH levels, increased GSSG levels, and a collapse of the GSH/GSSG ratio. SEPP1/selenite co-treatment significantly restored GSH homeostasis and re-established a more favorable intracellular redox state (Figure 3C-E).
At the molecular level, western blot analysis demonstrated that oxidative stress increased TFRC expression and reduced GPX4 expression, consistent with ferroptosis-associated molecular changes. SEPP1/selenite co-treatment reversed these alterations by downregulating TFRC and restoring GPX4 (Figure 3F and G), supporting recovery of GPX4-dependent lipid peroxide detoxification. Consequently, H2O2 stress increased lipid peroxidation, while Trt treatment significantly reduced lipid peroxide accumulation, as reflected by C11-BODIPY/MDA readouts (Figure 3H-J).
To further evaluate ferroptosis involvement, we performed a Fer-1 rescue experiment. Fer-1 (10 μM) significantly improved cell viability in H2O2-treated DPSCs (CCK-8; Supplementary Figure 2), supporting the interpretation that ferroptosis-associated injury contributes to oxidative damage in this model. Taken together, these findings indicate that acute oxidative stress induces ferroptosis-associated lipid peroxidation features in DPSCs, and that SEPP1/selenite co-treatment mitigates these changes by improving iron-redox balance and preserving selenium-dependent antioxidant defense.
To elucidate the downstream mechanisms underlying the protective effects of SEPP1/selenite co-treatment, we performed bulk RNA sequencing comparing Ctrl and Trt groups. Differential expression analysis was conducted using DESeq2 with Benjamini-Hochberg multiple-testing correction, and genes with Padj < 0.05, FPKM ≥ 1, and |log2 fold change| ≥ 1 were considered significantly differentially expressed. Using these criteria, we identified a broad transcriptional response to treatment, including 395 DEGs (128 upregulated and 267 downregulated in Trt vs Ctrl) (Figure 4A-C). The full bulk RNA-seq gene expression matrix (FPKM values) is provided in Supplementary Table 2.
Functional enrichment analysis of the filtered DEGs revealed that upregulated genes were strongly associated with mitotic progression, DNA replication, and cell-cycle regulation, whereas downregulated genes were enriched in inflammatory and senescence-associated pathways (Figure 4D). Notably, FOXM1 emerged as a central candidate regulator within the SEPP1/selenite-responsive transcriptional program, together with multiple FOXM1-associated mitotic genes (e.g., CCNB1, CDC20, BUB1B, and PLK1) (Figure 4D and E).
Gene Set Enrichment Analysis further supported these findings, showing enrichment of cell-cycle and mitotic spindle programs in Trt cells and suppression of inflammatory stress and senescence-associated signatures (Figure 4E). To validate the RNA-seq results, we performed qPCR for representative target genes and confirmed the treatment-associated induction of FOXM1-centered cell-cycle genes and repression of stress-related genes (Figure 4F). These data indicate that SEPP1/selenite co-treatment activates a FOXM1-centered protective transcriptional program in DPSCs under oxidative stress.
To determine whether FOXM1 is required for the protective effects of SEPP1/selenite co-treatment, we first validated the knockdown efficiency of two siRNAs targeting FOXM1 (siF1 and siF2). Both siRNAs significantly reduced FOXM1 mRNA and protein expression compared with the negative control, and siF1 showed stronger knockdown efficiency; therefore, siF1 was used for subsequent experiments (Figure 5A-C).
We then examined the impact of FOXM1 depletion on the anti-senescence effect of SEPP1/selenite co-treatment under oxidative stress. Although SEPP1/selenite co-Trt markedly reduced senescence compared with the H2O2-treated Ctrl group, FOXM1 knockdown (Trt + siF1) largely abolished this protection. The Trt + siF1 group showed a clear rebound in SA-β-gal-positive cells and reacquired a flattened, senescent morphology (Figure 5D and E). Consistently, western blotting demonstrated that FOXM1 knockdown reversed the molecular benefits of treatment: The suppression of P53, P21, and P16 was lost, and the restoration of proliferating cell nuclear antigen was significantly blunted in the Trt + siF1 group (Figure 5F and G).
The proliferative rescue conferred by SEPP1/selenite co-treatment was also FOXM1-dependent. CCK-8 assays showed that FOXM1 depletion significantly compromised the growth advantage of the Trt group, with viability curves shifting back toward Ctrl levels (Figure 5H). These data indicate that FOXM1 is required for the anti-senescence and pro-proliferative effects of SEPP1/selenite co-treatment in oxidatively stressed DPSCs.
We next assessed whether FOXM1 is also required for SEPP1/selenite-mediated redox recovery. In the Trt group, SEPP1/selenite co-treatment restored GSH homeostasis, as shown by increased reduced GSH, reduced GSSG, and recovery of the GSH/GSSG ratio. However, these protective changes were largely nullified after FOXM1 knockdown, and the Trt + siF1 group exhibited a redox profile resembling stressed Ctrl cells (Figure 5I-K).
At the molecular level, FOXM1 depletion also disrupted the ferroptosis-associated protective program induced by SEPP1/selenite co-treatment. The treatment-dependent downregulation of TFRC and restoration of GPX4 were both attenuated in the Trt + siF1 group (Figure 5L and M). This molecular failure was accompanied by impaired iron handling: FerroOrange staining showed significant re-accumulation of intracellular labile Fe2+ in FOXM1-knockdown cells despite SEPP1/selenite co-treatment (Figure 5N and O).
Finally, qPCR analysis demonstrated that the induction of downstream FOXM1-associated targets by SEPP1/selenite co-treatment, including MYBL2, SESN3, and CDC20, was abolished in the Trt + siF1 group (Figure 5P). Taken together, these results demonstrate that FOXM1 is a required downstream effector for the anti-senescence, antioxidant, and anti-ferroptotic effects of SEPP1/selenite co-treatment in DPSCs.
To validate the clinical relevance of our transcriptomic and in vitro findings, we analyzed histological and molecular changes in human dental pulp tissues from young and aged donors. Donor metadata and sample allocation for each assay are summarized in Supplementary Table 1. Histological examination by hematoxylin and eosin staining showed clear age-related structural differences: Young pulp tissues displayed relatively high cellular density and organized stromal architecture, whereas aged pulp tissues exhibited reduced cellularity and degenerative changes (Figure 6A).
We next examined the expression of the key proteins SEPP1 and FOXM1 in pulp tissues. Western blotting of tissue lysates showed that both SEPP1 and FOXM1 protein levels were significantly lower in the aged group compared with the young group (Figure 6B and C; n = 3 for western blotting). To assess whether these reductions were associated with the putative DPSC compartment, we performed immunofluorescence co-staining with THY1 as a DPSC-associated marker. In young pulp, SEPP1 showed strong fluorescence intensity and clear co-localization with THY1-positive cells (Figure 6D). In contrast, aged pulp exhibited markedly reduced SEPP1 and THY1 fluorescence signals (Figure 6E and F). Notably, Pearson’s correlation coefficient analysis showed that the spatial co-localization between SEPP1 and THY1 remained detectable and was not markedly disrupted despite the reduction in signal intensity (Figure 6G), suggesting that SEPP1 remains associated with residual THY1-positive cells while overall abundance declines with age.
A similar pattern was observed for FOXM1. FOXM1 immunofluorescence was enriched in THY1-positive cells in young pulp (Figure 6H), whereas aged tissues showed substantially weaker FOXM1 staining (Figure 6I and J). Pearson’s correlation coefficient analysis again indicated preserved spatial association between FOXM1 and THY1 signals despite reduced overall expression (Figure 6K). To further assess the selenium-dependent antioxidant network identified by transcriptomic analysis, we measured downstream selenoprotein-related transcripts in human pulp tissues. qPCR showed significant downregulation of GPX4 (Figure 6L), thioredoxin reductase 1 (Figure 6M), and DIO2 (Figure 6N) in aged pulp compared with young pulp.
Taken together, these human tissue data support an age-associated reduction in SEPP1 and FOXM1 expression, accompanied by decreased expression of selenium-dependent antioxidant genes in the dental pulp microenvironment. Given the low human sample numbers, these findings should be interpreted as supportive validation of the SEPP1/FOXM1-associated protective program rather than definitive proof of causality in vivo.
The progressive functional decline of DPSCs is a central event in pulp aging and contributes to reduced pulp vitality and impaired repair capacity in older individuals. In the present study, we integrated scRNA-seq analysis, in vitro oxidative stress experiments, bulk RNA-seq, and human tissue validation to show that aging-associated contraction of the DPSC compartment is accompanied by suppression of a selenium-dependent antioxidant program, including reduced SELENOP/SEPP1 expression and downstream selenoprotein-related genes. Functionally, SEPP1/selenite co-treatment alleviated oxidative senescence phenotypes and reduced ferroptosis-associated lipid peroxidation features in DPSCs, whereas FOXM1 knockdown attenuated these protective effects. Together, these findings support a model in which a SEPP1/selenium-associated redox program is functionally linked to FOXM1-dependent maintenance of DPSC homeostasis, rather than establishing a fully resolved direct signaling axis.
Our findings are consistent with the established view that oxidative stress is a major driver of DPSC senescence. Previous studies have shown that aged DPSCs exhibit increased ROS accumulation, persistent DNA damage signaling, and activation of the p53-p21/p16 senescence pathway[28,29]. Building on this framework, our scRNA-seq analysis highlights a more specific vulnerability in aged pulp, namely erosion of the selenium-dependent antioxidant network within the mesenchymal compartment. In contrast to prior studies primarily focused on canonical senescence regulators[30] or epigenetic mechanisms[31], our data suggest that impaired selenium handling, particularly reduced SEPP1 availability, may contribute to weakened antioxidant capacity in DPSCs. This interpretation is also consistent with reports in other stem/progenitor systems showing that selenium insufficiency accelerates functional decline[32]. Importantly, our human pulp validation further supports the clinical relevance of this framework by showing reduced SEPP1/FOXM1 expression, together with lower expression of selenium-dependent antioxidant genes in aged tissues.
A key implication of this study is that ferroptosis-associated lipid peroxidation appears to be involved in oxidative DPSC injury under aging-like stress conditions. In our model, H2O2 challenge induced a coordinated pattern of labile Fe2+ accumulation, GSH imbalance, GPX4 reduction, and increased lipid peroxidation, all of which were mitigated by SEPP1/selenite co-treatment[33,34]. These data support the presence of ferroptosis-associated injury features, but they do not by themselves establish definitive ferroptotic cell death. To address this point, we performed Fer-1 rescue experiments, which showed improved cell viability after oxidative injury and strengthened the interpretation that ferroptosis-related mechanisms contribute to the observed damage. Accordingly, we use the more conservative term “ferroptosis-associated lipid peroxidation/injury” rather than making an overly strong claim that ferroptosis is the sole death mechanism[35]. Future studies using genetic perturbation of core ferroptosis regulators (for example, GPX4 or SLC7A11) will be important to further refine this conclusion.
Our transcriptomic and loss-of-function results further indicate that FOXM1 is a key downstream effector of the protective phenotype observed under SEPP1/selenite co-treatment conditions. FOXM1 is well recognized as a regulator of cell-cycle progression, DNA damage recovery, and stress adaptation[36]. In the present study, SEPP1/selenite co-treatment was associated with induction of a FOXM1-centered transcriptional program, including MYBL2, CDC20, and SESN3. FOXM1 knockdown reversed the anti-senescence, antioxidant, and ferroptosis-associated protective effects[37,38]. These findings extend FOXM1 biology to the context of DPSC oxidative injury and support a functional link between SEPP1/selenite-supported redox recovery and FOXM1 activity. However, our data do not establish whether FOXM1 is directly regulated by SEPP1 itself, by improved selenium bioavailability, by restored selenoprotein function (for example, GPX4-dependent redox buffering), or by secondary redox-sensitive signaling pathways. In addition, we did not evaluate FOXM1 post-translational regulation (such as phosphorylation or ubiquitination), nor did we perform ChIP or promoter-reporter assays to determine whether FOXM1 directly regulates downstream antioxidant or ferroptosis-related genes[39,40]. Therefore, the SEPP1-FOXM1 relationship in this study should be interpreted as a functionally linked pathway module rather than a fully resolved direct molecular axis.
An important strength of this study is the integration of multiple data layers to position selenium metabolism within DPSC aging biology. Compared with prior DPSC studies that primarily described senescence phenotypes or tested general antioxidant interventions[41-43], our work provides a transcriptome-informed rationale for focusing on selenium-dependent redox maintenance in the DPSC compartment. The scRNA-seq analysis suggests that age-related DPSC decline is accompanied by suppression of a specific antioxidant program, rather than a nonspecific loss of cellular function[44]. At the same time, we agree that reduced DPSC abundance in aged pulp is a recognized phenomenon, and our data do not prove that selenium pathway impairment is the sole driver of this process in vivo. Instead, our results support SEPP1/selenium insufficiency as one important contributor within a broader aging microenvironment, which likely also involves vascular, immune, and extracellular matrix alterations. This framing better reflects the scope of the current dataset and avoids overinterpretation of causality.
Several limitations of this study should be noted. First, the in vitro model is based on acute H2O2 exposure, which is useful for inducing oxidative stress/senescence-like injury but does not fully recapitulate chronic or replicative aging[29,31]. Future studies should evaluate the SEPP1/FOXM1-associated protective program in long-term passaging or selenium-restriction models to better mimic progressive aging of the pulp microenvironment. Second, because our intervention used a combined SEPP1 + sodium selenite regimen, the individual biochemical contributions of recombinant SEPP1 and inorganic selenium cannot yet be completely separated, although we added single-agent comparison groups and Fer-1 rescue validation to improve treatment attribution. A more systematic design, including dose-response and timing analyses, will be needed to clarify whether SEPP1 primarily acts through selenium delivery, synergizes with selenite, or also exerts additional effects. Third, although SEPP1 classically depends on receptor-mediated selenium uptake, we did not assess SEPP1 receptor expression or uptake mechanisms in DPSCs, and we did not perform any receptor-blocking experiments. Fourth, the human tissue validation supports clinical relevance but remains limited by sample size and assay-specific sample allocation, and therefore should be interpreted as supportive rather than definitive evidence for in vivo causality.
Despite these limitations, the present study provides a practical translational direction for regenerative endodontics. Rather than positioning SEPP1 as a stand-alone anti-aging factor, our data support SEPP1/selenium-based redox support as a potential adjunctive strategy to preserve DPSC viability and function in oxidative or aging-compromised pulp microenvironments. In practical terms, this approach may be explored through local delivery systems, such as pulp-capping biomaterials, injectable hydrogels, or bioactive scaffolds designed to support redox homeostasis during pulp repair[19,45,46]. Future in vivo studies should evaluate whether SEPP1/selenium-supportive formulations improve histologic repair quality, reduce inflammatory burden, and preserve long-term pulp vitality, particularly in aged or stress-compromised settings. These translational considerations remain preliminary but provide a clinically relevant direction for further investigation.
Our study identifies a selenium-dependent antioxidant vulnerability in aging dental pulp and shows that SEPP1/selenite co-treatment mitigates oxidative senescence and ferroptosis-associated lipid peroxidation features in DPSCs. Mechanistically, the protective effects are functionally linked to a FOXM1-centered program, as FOXM1 knockdown attenuated the anti-senescence and redox-protective responses. By integrating scRNA-seq analysis, in vitro oxidative stress experiments, bulk RNA-seq, and human tissue validation, this work provides a mechanistically informed but cautious framework for understanding how selenium metabolism contributes to DPSC aging. These findings support further investigation of SEPP1/selenium-based redox-supportive strategies for preserving pulp vitality in aging or stress-compromised microenvironments.
The authors appreciate ProMab Biotechnologies for supplying antibodies and molecular detection platforms.
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