MSC-derived small extracellular vesicles promote dental pulp repair by alleviating inflammation in pulpitis via AKT and p38 MAPK pathways

Preparation and characterization of MSC-sEVs

MSC-sEVs were purchased from Paracrine Therapeutics Pte Ltd (Singapore) and derived from an immortalized E1-MYC 16.3 human embryonic stem cell (hESC)-derived MSC line.¹⁹ Briefly, cells were cultured in Dulbecco’s Modified Eagle Medium (DMEM) (Thermofisher Scientific, Waltham, MA, USA) supplemented with 10% fetal bovine serum (FBS; Thermofisher Scientific). For sEV preparation, the conditioned medium was prepared by growing 80% confluent cells in a chemically defined medium composed of DMEM supplemented with 1% non-essential amino acids, 1% glutamine, 1% insulin-transferrin-selenium-X, 1 mM sodium pyruvate, 0.05 mM β-mercaptoethanol, 5 ng/mL fibroblast growth factor (FGF)-2 (Thermofisher Scientific) and 5 ng/mL platelet-derived growth factor (PDGF)-AB (Cytolab Ltd, Rehovot, Israel) for 3 days.²⁰ The conditioned medium was then size-fractionated by tangential flow filtration, and concentrated 50 × using a 100 kDa molecular weight cutoff membrane (Sartorius, Göttingen, Germany). The sEV preparation was characterized according to the Minimal Information for Studies of Extracellular Vesicles (MISEV) guidelines^21,22 and more specifically with the identity and potency metrics proposed for MSC-sEV preparations.^23,24 EV morphology (Figure S1A, see online supplementary material for a color version of this figure) was examined by transmission electron microscopy (TEM; Talos L120C, Thermofisher Scientific), revealing the characteristic nanosized, membrane-bound vesicular structure. Protein concentration was determined using the Coomassie Plus (Bradford) assay (Thermofisher Scientific), particle size distribution (Figure S1B, see online supplementary material for a color version of this figure) and concentration were measured by ZetaView (Particle Metrix GmbH, Meerbusch, Germany), and CD73/ecto-5′-nucleotidase (NT5E) activity was assessed using the PiColorLock Gold Phosphate Detection System (Innova Biosciences, Cambridge, UK). Western blotting was performed to examine EV markers including HSP70, CD63, TSG101 CD9, and CD81 (Figure S1C, see online supplementary material for a color version of this figure). For this study, MSC-sEV preparation (AC108) was used, with a protein concentration of 1.49 mg/mL, particle concentration of 1.47 × 10¹¹ particles/mL, modal particle size of 135.2 nm (Figure S1B, see online supplementary material for a color version of this figure), and CD73/NT5E activity of 20.47 ± 0.7 mU/µg protein. MSC-sEVs were 0.22-µm filtered and stored at −20 °C until use.

Animal experiments

All procedures were approved by the Institutional Animal Care and Use Committee of the National University of Singapore (protocol number: R19-0502) and conducted in accordance with the institutional guidelines. A total number of 32 Sprague Dawley rats (8-week-old, male) with a mean weight of 265.6 ± 32.0 g (range 196-306 g) were used. Experimental pulpitis was induced in the bilateral maxillary first molars using LPS from Escherichia coli O111: B4 (Sigma, St Louis, MO, USA) as previously described.²⁵ LPS, a major constituent of the outer membrane of Gram-negative bacteria, plays a critical role in pulpal infection and inflammation and is widely used to establish experimental models of pulpitis.^26,27 Under general anesthesia, standardized Class I occlusal defects (1.5 mm diameter × 1.5 mm depth) were prepared on the bilateral maxillary first molars using a 1/4 RA carbide bur (Wave Dental, Ningbo, China) mounted on a dental handpiece (NSK, Tokyo, Japan). Subsequently, 2 μL of LPS solution (1 μg/mL) was applied to each defect, which was then temporarily sealed with a cotton pellet. After 24 h, cotton pellets were removed and the molars were randomly assigned to 4 groups (n = 8 defects/group): collagen sponge (CS), collagen sponge with 2 µl phosphate-buffered saline (CS/PBS), collagen sponge with 2 µL of MSC-sEVs (20 µg; CS/sEVs) and mineral trioxide aggregate (MTA). All defects were sealed with glass ionomer cement (GIC; Fuji GC, Tokyo, Japan). Post-operatively, animals received subcutaneous buprenorphine (0.2 mL/100 g body weight) twice daily for 3 days for analgesia. Rats were housed in pairs under standard laboratory conditions with unrestricted movement and provided food and water ad libitum. At 1 and 4 weeks post-treatment, animals were euthanized by carbon dioxide (CO₂) inhalation, and the maxillae were harvested for histological analysis (Figure 1).

Histology, immunohistochemistry, and histomorphometry

Tissue samples were fixed in 10% (v/v) neutral buffered formalin (Sigma) overnight and decalcified in 30% (v/v) buffered formic acid for 4 weeks. After decalcification, samples were dehydrated, embedded in paraffin, and serially sectioned at 5-μm thickness. Sections were stained with hematoxylin and eosin (HE) to evaluate general tissue morphology and with Masson’s Trichrome (MT) to assess reparative dentin formation. Stained sections were imaged using an inverted microscope (IX70, Olympus, Tokyo, Japan). Newly formed reparative dentin (osteodentin) was identified under 100 × magnification, and the area was measured using ImageJ software (National Institutes of Health, Bethesda, MD, USA). The percentage of reparative dentin was calculated relative to the total pulp defect or lesion area. Inflammation in the pulp tissue was evaluated on the HE-stained sections by 3 independent, blinded examiners using a modified histological grading system (Table 1).^28,29 The proportion of specimens within each grade was calculated relative to the total number of samples at each time point.

Immunohistochemistry was performed using a biotin-streptavidin Lab Vision UltraVision detection system (Thermofisher Scientific), as previously described.³⁰ Briefly, paraffin sections were deparaffinized, rehydrated, and subjected to antigen retrieval in sodium citrate buffer (pH 6.0) at 60 °C for 1 h. Endogenous peroxidase activity was quenched with peroxidase block for 15 min, followed by a 5-min incubation in Ultra V blocking solution. Sections were then incubated at room temperature for 1 h with the following primary antibodies: anti-DSPP (1:250, Santa Cruz Biotech, Dallas, TX, USA), anti-DMP1 (1:500, Biorbyt, Cambridge, UK), anti-IL-6 (1:250, Abcam, Cambridge, MA, USA), anti-MMP9 (1:2000, Abcam), and anti-NGF (1:500, R&D Systems, Minneapolis, MN, USA). After rinsing with PBS, sections were incubated with biotinylated secondary antibodies for 30 min. Antibody-antigen reaction was visualized using 3,3'-diaminobenzidine (DAB) chromogen for 1 min, followed by nuclear counterstaining with Mayer’s hematoxylin (Sigma). Stained sections were imaged using an inverted microscope (Olympus IX70). Quantification was performed at 100 × magnification by measuring the percentage of positively stained areas for DSPP or DMP1 across the pulp lesion, and for IL-6, MMP9 or NGF beneath the pulp lesion, normalized to the total image area using ImageJ software.

Isolation and culture of dental pulp cells

Dental pulp cells (DPCs) were isolated from the extracted rat incisors following an established protocol with modification.³¹ Briefly, the gingival soft tissues were removed, and incisors were extracted using sterile forceps. The incisors were washed with PBS supplemented with 2% penicillin/streptomycin (PS; Thermofisher Scientific). After removing the soft tissues from the root surface, the incisors were split open and the dental pulp tissues were carefully extracted. The isolated pulp tissues were minced into small pieces and enzymatically digested in a solution containing 2% dispase II and 0.2% collagenase I (Worthington, Lakewood, NJ, USA) at 37 °C for 1 h. The resulting cell suspension was filtered through a 40-μm cell strainer (Corning®, Corning, NY, USA) to obtain single cell suspension, which was then seeded at a density of 2 × 10⁴ cells/cm². Cells were cultured in complete growth medium comprising of DMEM-F12 supplemented with 10% fetal bovine serum (FBS) and 1% PS at 37 °C in a humidified atmosphere with 5% CO₂. The culture medium was changed every other day. Upon reaching confluence, DPCs were detached using TrypLe (Thermofisher Scientific) and passaged to passage (P) 3 for subsequent experiments.

In vitro DPC spheroid model of pulpitis

An in vitro 3-D spheroid culture model of DPCs was established to evaluate the therapeutic effects of MSC-sEVs under inflammatory conditions. This scaffold-free culture strategy enhances cell–cell interactions and multilineage regenerative potential, thereby preserving the intrinsic properties of DPCs and more closely mimicking the native pulp tissue microenvironment.^32,33 Pulpal inflammation was induced by LPS stimulation. Briefly, rat DPCs were seeded into 96-well U-bottom plates at a density of 2.5 × 10⁵ cells/well and centrifuged at 250 × g for 5 min to facilitate spheroid formation. Spheroids were maintained in low-serum culture medium comprising of DMEM-F12 supplemented with 0.5% FBS and 1% PS at 37 °C in a humidified atmosphere containing 5% CO₂. Upon spheroid formation, DPCs were stimulated with LPS from E. coli O111: B4 (10 μg/mL; Sigma) for 24 h to induce an inflammatory response.^34,35 Subsequently, spheroids were treated with MSC-sEVs at concentrations of 1, 5 and 10 μg/mL, or vehicle (PBS) for an additional 48 h. All in vitro experiments were performed in quadruplicates (n = 4) in at least 2 independent experiments.

Treatment of sEVs and inhibitor studies

DPC spheroids were stimulated with LPS (10 μg/mL) for 24 h, followed by treatment with MSC-sEVs (10 μg/mL) or PBS as a vehicle control, and incubated for an additional 48 h. To investigate the role of adenosine receptors in the activation of protein kinase B (AKT), extracellular signal-regulated kinase (ERK), and p38 mitogen-activated protein kinase (MAPK) signaling pathways, LPS-stimulated spheroids were pretreated for 1 h with theophylline (1 mM; a nonselective adenosine receptor antagonist; Sigma),³⁶ wortmannin (1 μm; AKT inhibitor), U0126 (10 μm; ERK inhibitor), or SB203580 (10 μm; p38 MAPK inhibitor) (Cell Signaling Technology, Danvers, MA, USA) prior to sEV treatment. For CD73 inhibition experiments, cells were co-treated with sEVs and 10 nM PSB12379 (CD73 inhibitor; Tocris Biosciences, CO, USA) for 1 h. All in vitro experiments were performed in quadruplicates (n = 4) in at least 2 independent experiments.

Nitric oxide and DNA quantification

Nitric oxide (NO) levels in culture supernatants from DPC spheroids were quantified using the Griess Reaction Assay Kit (Thermofisher Scientific) according to the manufacturer’s instructions. Absorbance was measured at 548 nm using a microplate reader (Infinite® 200 PRO, Tecan, Männedorf, Switzerland). NO concentrations were normalized to the DNA content of the corresponding DPC spheroids and expressed as NO/DNA.

Enzyme-linked immunosorbent assays

The concentrations of IL-6, MMP9 and NGF in culture supernatants from DPC spheroids were measured using commercially available rat IL-6 and MMP9 enzyme-linked immunosorbent assay (ELISA) kits (MyBiosource, San Diego, CA, USA) and a rat NGF ELISA kit (Sigma), following the manufacturers’ protocols. Absorbance was read at 450 nm using a microplate reader (Infinite® 200 PRO, Tecan™). Total protein concentration was determined using the Coomassie (Bradford) Protein Assay Kit (Thermofisher Scientific), with absorbance measured at 595 nm. IL-6, MMP9, and NGF concentrations were normalized to total protein content and presented as IL-6/Protein, MMP9/Protein and NGF/Protein.

Western blot analysis

Total protein was extracted and separated by SDS-PAGE using 4%-12% polyacrylamide gels (Thermofisher Scientific), followed by electro-blotting onto a nitrocellulose membrane (GE Healthcare, Chicago, IL, USA). Membranes were blocked and incubated with primary antibodies followed by horseradish peroxidase (HRP)-conjugated secondary antibodies. The primary antibodies used were anti-HSP70, anti-CD9, anti-CD63, anti-TSG101, and anti-CD81 (all 1:1000, Thermofisher Scientific), anti-CD81 (1:1000, Proteintech), anti-AKT (pan), anti-phospho-AKT (Ser473), anti-ERK1/2, anti-phospho-ERK1/2 (Thr202/Tyr204), anti-p38 MAPK, and anti-phospho-p38 MAPK (Thr180/Tyr182) (all 1:1000, Cell Signaling Technology), as well as mouse anti-GAPDH (1:10000, Abcam). Following incubation with appropriate HRP-coupled secondary antibodies (GE Healthcare), immunoreactive bands were visualized using SuperSignal West Pico Chemiluminescent Substrate (Thermofisher Scientific) and imaged with the ChemiDoc MP System (Bio-Rad Laboratories, Hercules, CA, USA).

Quantitative real-time reverse transcription polymerase chain reaction

Total RNA was isolated using PureLink® RNA Mini kit (Thermofisher Scientific) according to the manufacturer’s instructions. Complementary DNA (cDNA) was synthesized using iScript Reverse Transcription Supermix (Bio-Rad). Quantitative real-time reverse transcription polymerase chain reaction (qRT-PCR) was performed using iTaq Universal SYBR® Green Supermix (Bio-Rad) and gene-specific primers (Table 2) on a CFX Connect Real-Time PCR System (Bio-Rad). The PCR cycling conditions comprised an initial denaturation at 95 °C for 30 s, followed by 40 cycles of amplification consisting of denaturation at 95 °C for 15 s and extension at 60 °C for 30 s. Relative mRNA expression levels were normalized to glyceraldehyde 3-phosphate dehydrogenase (GAPDH) and calculated using the comparative ΔCt method.³⁷ Data are presented as fold change relative to control.

Statistical analysis

All quantitative data are presented as mean ± standard deviation (SD). For normally distributed data, Student’s t-test or one-way analysis of variance followed by Scheffe’s post-hoc test was used. For non-normally distributed data, Mann-Whitney or Kruskal-Wallis followed by Dunn-Bonferroni post-hoc test was applied. Pearson’s correlation was performed to assess association between variables, with results reported as the Pearson correlation coefficient (r), sample size (n), and Pvalue. Statistical analysis was conducted using IBM SPSS Statistics version 25.0 (IBM, Chicago, IL, USA). Statistical significance was set as P < .05.

MSC-sEVs enhance reparative dentin formation in rat pulpitis

To determine whether MSC-sEVs enhance pulp repair in vivo, reparative dentin formation was evaluated in an experimentally induced rat pulpitis model. As early as 1 week post-treatment, all pulp lesions in the CS/sEVs group exhibited robust reparative dentin bridge formation, characterized by a dense collagenous matrix covering >50% of the exposed pulp area, closely resembling the response observed in the MTA-treated group (Figure 2A). In contrast, the CS group displayed only residual collagen sponge without discernible dentin-bridge formation, while CS/PBS group showed limited and discontinuous hard tissue deposition (Figure 2A). Quantitative histomorphometric analysis confirmed these observations. The CS/sEVs group demonstrated a significantly higher percentage areal deposition of reparative dentin (29.80% ± 0.43%) compared with the CS group (0.99% ± 0.03%; P < .001) and CS/PBS group (4.77% ± 1.05%; P < .01) and was comparable to the MTA group (34.34% ± 1.64%; P > .05) (Figure 2B).

By 4 weeks, reparative dentin bridges in the CS/sEVs group completely covered the pulp area and exhibited a collagen-rich architecture, comparable to the MTA group (Figure 2A). In contrast, minimal hard tissue deposition persisted in the CS group, while the CS/PBS group formed only a thin and incomplete layer of reparative dentin. Quantitative analysis showed that the CS/sEVs group had a significantly greater areal deposition of reparative dentin (38.13% ± 1.68%) than the CS (2.87% ± 0.82%; P < .01) and CS/PBS groups (10.37% ± 0.71%; P < .01), with no significant difference compared to MTA group (36.89% ± 1.40%; P > .05) (Figure 2B). Notably, reparative dentin formation in the CS/sEVs group increased significantly from 1 to 4 weeks (P < .01), indicating progressive and sustained pulp repair.

Consistent with enhanced dentinogenesis, immunohistochemical analyses revealed significantly elevated expression of odontogenic markers in the CS/sEVs group. At 1 week, DSPP expression was markedly higher in the CS/sEVs group (11.29% ± 1.36%) compared with the CS (2.41% ± 0.84%; P < .01) and CS/PBS groups (4.13% ± 1.02%; P < .05), and comparable to the MTA group (11.31% ± 1.32%; P > .05) (Figure 2C). In contrast, DMP1 expression did not differ significantly among groups except MTA at this early time point (Figure 2D).

At 4 weeks, DSPP expression in the CS/sEVs group (13.82% ± 0.92%) remained significantly higher than in the CS (3.43% ± 1.72%; P < .001) and CS/PBS groups (6.52% ± 2.17%; P < .01), although it was modestly lower than in the MTA group (18.6% ± 0.93%, P < .01) (Figure 2C). Importantly, DMP1 expression was significantly increased in the CS/sEVs group (12.25% ± 1.45%) relative to the CS (3.91% ± 1.03%; P < .001) and CS/PBS groups (4.19% ± 1.39% P < .01) and was comparable to MTA group (10.79% ± 0.80%, P > .05) (Figure 2D). DMP1 expression increased significantly from 1 to 4 weeks in the CS/sEVs group (P < .01), consistent with progressive odontoblastic maturation.

MSC-sEVs alleviate pulpal inflammation

Given the central role of inflammation in pulpitis, inflammatory responses were assessed histologically. At 1 week, ∼88% of samples in the CS/sEVs group exhibited only mild inflammatory cell infiltration, whereas ≥50% of samples in the CS and CS/PBS groups displayed severe inflammation (P < .001) (Figure 2E). Consistent with these findings, immunohistochemical analysis showed that expression of pro-inflammatory cytokine IL-6, a key mediator of inflammation in pulpitis,²⁶ was significantly lower in the CS/sEVs group (3.50% ± 0.62%) compared with the CS (5.83% ± 1.00%; P < .05) and CS/PBS groups (6.69% ± 0.43%; P < .01), and was comparable to that observed in the MTA group (3.40% ± 1.33%; P > .05) (Figure 3A and B). Similarly, expression of MMP9, which contributes to extracellular matrix (ECM) degradation during pulpal inflammation,^38,39 was markedly reduced in the CS/sEVs group (5.65% ± 0.91%) relative to the CS (11.94% ± 1.90%; P < .05) and CS/PBS groups (12.47% ± 0.99%; P < .01), and was comparable to the MTA group (4.68% ± 0.46%; P > .05) (Figure 3A and C).

At 4 weeks, ∼60% of samples in the CS/sEVs group exhibited minimal or normal inflammatory responses, whereas ≥50% of samples in the CS and CS/PBS groups showed mild to moderate inflammation (P < .01); these findings were comparable to the MTA group, in which 50% of samples demonstrated normal inflammatory responses (Figure 2F). Consistently, IL-6 expression in the CS/sEVs group remained significantly lower (3.23% ± 0.54%) than in the CS (8.57% ± 0.99%; P < .001) and CS/PBS groups (9.68% ± 1.37%; P < .01), and was comparable to that in the MTA group (3.66% ± 0.37%; P > .05) (Figure 3B). Likewise, MMP9 expression was significantly reduced in the CS/sEVs group (7.61% ± 1.10%) compared with the CS (15.00% ± 1.82%; P < .01) and CS/PBS groups (16.61% ± 0.91%; P < .001), and did not differ from the MTA group (7.42% ± 1.62%; P > .05) (Figure 3C). Pearson’s correlation analysis revealed that lower expression levels of IL-6 (r = 0.618, n = 32, P = .032) and MMP9 (r = 0.606, n = 32, P = .037) were positively correlated with improved scores for reparative dentin formation.

To assess potential pain modulation, NGF expression was examined (Figure 3A). At 1 week, NGF expression in the CS/sEVs group (3.14% ± 0.62%) was significantly higher than that observed in the CS (0.81% ± 0.38%; P < .05), CS/PBS (1.95% ± 0.10%; P < .01), and MTA groups (0.57% ± 0.24%; P < .05) (Figure 3D). In contrast, at 4 weeks, NGF expression in the CS/sEVs group was significantly reduced (0.72% ± 0.15%) compared with the CS (1.48% ± 0.24%; P < .05) and CS/PBS groups (4.06% ± 0.75%; P < .01), and was comparable to that in the MTA group (0.62% ± 0.23%; P = .992) (Figure 3D). Pearson’s correlation analysis further revealed that reduced NGF expression was positively correlated with improved reparative dentin formation scores (r = 0.630, n = 32, P = .002).

MSC-sEVs suppress LPS-induced inflammation in DPC spheroids in vitro

To corroborate the in vivo findings, we examined the effects of MSC-sEVs on LPS-induced inflammatory responses in DPC spheroids (Figure 4A). LPS stimulation markedly increased NO production by DPCs; however, this response was suppressed by MSC-sEVs (Figure 4B). At 48 h, sEV treatment with 10 µg/mL reduced NO production by ∼50% (P < .01), restoring level comparable to that of untreated control (Figure 4B). Consistent with these findings, MSC-sEVs significantly reversed LPS-induced increases in IL-6 and MMP9 production (Figure 4C and D). IL-6 level was reduced by ∼50% following treatment with 10 μg/mL sEVs (P < .001), reaching baseline level observed in untreated control (Figure 4C). MMP9 production was significantly reduced at 1 µg/mL sEVs (P < .001), with no additional suppression observed at higher concentrations (Figure 4D). Similarly, NGF production was reduced by ∼40% following treatment with 1 µg/mL sEVs (P < .01), restoring level to that of untreated control (Figure 4E). Gene expression analysis further supported these findings, demonstrating significant downregulation of genes associated with oxidative stress (iNOS), inflammation (iNOS, IL-6, IL-8, and MCP1), ECM degradation (MMP3 and MMP9), pain (NGF, Substance P, TrkA, CGRP, and P75NTR), and apoptosis (CCP3) following sEV treatment (P < .05, Figure 4F). In contrast, genes related to cell proliferation (PCNA) and odontogenesis, including DSPP and DMP1, were significantly upregulated with sEV treatment, exceeding the baseline levels observed in untreated controls (P < .01, Figure 4F). Together, these results demonstrate that MSC-sEVs effectively suppressed LPS-induced production of NO, IL-6, MMP9, and NGF in DPC spheroids, consistent with their anti-inflammatory effects observed in vivo, while simultaneously promoting dental pulp repair and reparative dentin formation.

MSC-sEVs modulate AKT and p38 MAPK signaling to mediate anti-inflammatory effects

To elucidate the underlying mechanisms, we examined the effects of MSC-sEVs on key intracellular signaling pathways in LPS-stimulated DPC spheroids (Figure 5A). Treatment with MSC-sEVs significantly increased AKT phosphorylation while concomitantly reducing p38 MAPK phosphorylation (Figure 5B and F). Importantly, these effects were partially reversed by pharmacological inhibition of CD73 with PSB12379 (Figure 5B-E), suggesting a critical role for sEV-associated CD73 activity. Similarly, treatment with theophylline, a nonselective adenosine receptor antagonist, attenuated MSC-sEV-induced AKT phosphorylation and reversed the suppression of p38 MAPK phosphorylation (Figure 5F-I). Together, these findings indicate that the modulatory effects of MSC-sEVs on AKT and p38 MAPK signaling are mediated, at least in part, through CD73-dependent adenosine receptor activation. In contrast, ERK phosphorylation was reduced by CD73 inhibition with PSB12379 but not by theophylline inhibition of the adenosine receptors (Figure 5B and F). Functionally, theophylline significantly attenuated the sEV-mediated suppression of NO, IL-6, MMP9, and NGF production (P < .05-.01; Figure 5J-M), further supporting the role of adenosine receptor-dependent signaling in mediating the anti-inflammatory effects of MSC-sEVs.

Pharmacological inhibition studies were performed to delineate pathway specificity. Inhibition of AKT signaling with wortmannin (Figure 5N and O) partially reversed the sEV-mediated suppression of IL-6 and MMP9 production (P < .05; Figure 5U and V), while exerting only modest effects on NO and NGF levels (Figure 5T and W). In contrast, inhibition of ERK signaling by U0126 (Figure 5P and Q) did not significantly affect the sEV-mediated suppression of NO, IL-6, MMP9, or NGF production (Figure 5T-W). Notably, inhibition of p38 MAPK using SB203580 further reduced p38 MAPK phosphorylation (Figure 5R and S) and potentiated the suppressive effects of MSC-sEVs on NO, IL-6, MMP9, and NGF production (P < .05-.01; Figure 5T-W).

Collectively, these findings indicate that MSC-sEVs suppress inflammation, ECM degradation, and pain-associated signaling in DPC spheroids primarily through inhibition of p38 MAPK and, to a lesser extent, activation of AKT, while ERK signaling appears dispensable. This mechanistic framework aligns with the observed in vivo effects of MSC-sEVs in promoting resolution of pulpal inflammation and enhancing reparative dentin formation.