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. 2026 Jun 15;59(10):2164–2179. doi: 10.1111/iej.70199

A Novel piRNA‐Mediated Epigenetic Axis: piR‐36241 Exacerbates Pulpitis by Silencing the Protective Receptor ADGRG2 in Human Dental Pulp Stem Cells

Wenmian Huang 1, Yue Chen 2, Ye Jiang 1, Mianfeng Yao 1, Lihong Wu 1, Xuechao Yang 1,✉
PMCID: PMC13569280  PMID: 42290388

ABSTRACT

Background

Pulpitis, a prevalent inflammatory dental disease, primarily results from bacterial infection, yet its epigenetic regulatory mechanisms remain poorly understood. PIWI‐interacting RNAs (piRNAs), a class of small non‐coding RNAs known to maintain genomic stability, have not been investigated in the context of pulpitis.

Objective

This study aimed to profile piRNA expression patterns in pulpitis and to elucidate the functional role and underlying mechanism of a specific piRNA, piR‐36241 and its target gene ADGRG2 in disease progression.

Methods

piRNA and mRNA sequencing were performed on pulp tissues from patients with irreversible pulpitis and healthy controls. Potential piRNA targets were predicted bioinformatically. The interaction between piR‐36241 and the 3′UTR of ADGRG2, an orphan adhesion G protein‐coupled receptor (GPCR), was validated using dual‐luciferase reporter assay, quantitative Real‐Time PCR (qRT‐PCR), fluorescence in situ hybridization (FISH), Western blot (WB), hematoxylin‐eosin staining (HE), immunohistochemistry (IHC) and enzyme‐linked immunosorbent assay (ELISA). Functional analyses were conducted in LPS‐stimulated human dental pulp cells (HDPCs) through gain‐of‐function and rescue experiments.

Results

We identified a distinct piRNA expression profile in pulpitis, with 21 piRNAs differentially expressed. piR‐36241 was notably upregulated. It directly targeted the 3′UTR of ADGRG2 and suppressed its expression. ADGRG2, hypothesised to exert protective and homeostatic functions, was downregulated in pulpitis tissues and LPS‐induced HDPCs. Overexpression of piR‐36241 inhibited ADGRG2 expression and promoted the secretion of pro‐inflammatory cytokines (IL‐6, IL‐8). Conversely, knockdown of ADGRG2 similarly enhanced inflammatory responses. Rescue experiments demonstrated that piR‐36241 primarily regulates inflammation through ADGRG2 silencing.

Conclusion

Our study reveals a novel pathogenic axis in pulpitis whereby upregulation of piR‐36241 exacerbates inflammatory progression by repressing the protective receptor ADGRG2. These findings provide the first evidence of piRNA‐mediated epigenetic regulation in pulpitis and highlight the piR‐36241/ADGRG2 pathway as a potential therapeutic target for preserving pulp vitality.

Keywords: AGRGR2, inflammation, piRNA, pulpitis

1. Introduction

Pulpitis, a common and painful inflammatory disease of the dental pulp, arises primarily from bacterial infection and significantly compromises patients' quality of life (Zanini et al. 2017; Bender 2000; Edwards et al. 2023). This condition is characterised by the local accumulation of inflammatory mediators, such as cytokines and chemokines, which drive a complex interplay between destructive and reparative processes (Pohl et al. 2024; Loo et al. 2025). The outcome—preservation of pulp vitality or progression to irreversible damage—hinges on the delicate balance within this inflammatory microenvironment (Arruda‐Vasconcelos et al. 2023; Selvakumar et al. 2021; Wu et al. 2019).

The dental pulp harbours a heterogeneous cellular population, including dental pulp stem cells, odontoblasts, fibroblasts, endothelial cells and neuronal cells, all embedded within a richly vascularised and innervated matrix (Nief 1980). Upon microbial invasion, resident and infiltrating immune cells (e.g., macrophages, lymphocytes) respond vigorously. Their interaction with pathogens and the subsequent signalling cascade critically shape the inflammatory landscape, affecting both immune responses and the survival of native pulp cells (Renard et al. 2016; Wang et al. 2025).

Current mainstay treatment for irreversible pulpitis, root canal therapy, often results in loss of tooth vitality, increasing long‐term risks of fracture (Salehrabi and Rotstein 2010; Herbst et al. 2022). While pulp regeneration holds promise, its clinical translation faces challenges related to scaffold design, cell survival and precise control of differentiation within the inflammatory milieu (Liu, Huang, et al. 2025; Fei et al. 2025). A deeper molecular understanding of how pulpitis perturbs various cell lineages is therefore essential for developing targeted therapies to preserve pulp function.

This context, non‐coding RNAs (ncRNAs) have emerged as key regulators of inflammatory and reparative processes. Altered expression of microRNAs (miRNAs), long non‐coding RNAs (lncRNAs) and circular RNAs (circRNAs) in pulpitis has been documented, revealing complex post‐transcriptional networks that influence disease progression (Tan et al. 2025; Guo, Ma, et al. 2025; Lu et al. 2025). However, the role of a crucial class of small ncRNAs, PIWI‐interacting RNAs (piRNAs), remains entirely unexplored in pulpal pathology.

piRNAs (24–31 nucleotides) guide PIWI proteins to regulate gene expression at transcriptional and post‐transcriptional levels, thereby governing genome stability and diverse cellular pathways (Mei et al. 2015; Iwasaki et al. 2015). Their functional importance extends to various diseases, including cancer (Ye et al. 2025), inflammatory disorders (Plestilova et al. 2016) and tissue repair (Chen et al. 2020). Notably, innate immune stimuli can induce piRNA/PIWI expression, suggesting a potential role in inflammation (Plestilova et al. 2016). Although piRNAs are present in oral tissue‐derived exosomes (Wang et al. 2020), their expression profile and function in dental pulp inflammation are unknown.

Here, we performed the first comprehensive analysis of piRNA and mRNA expression in human pulpitis. We identified a distinct piRNA signature and focused on a significantly upregulated piRNA, piR‐36241. Combining bioinformatic prediction with experimental validation, we demonstrated that piR‐36241 directly targets and silences the orphan adhesion G protein‐coupled receptor ADGRG2. Functional studies revealed that the piR‐36241/ADGRG2 axis plays a pivotal role in modulating the inflammatory response in human dental pulp cells. Our findings uncover a novel epigenetic mechanism in pulpitis and highlight a potential therapeutic target for preserving pulp vitality.

2. Materials and Methods

The manuscript of this laboratory study has been written following the Preferred Reporting Items for Laboratory Studies in Endodontology (PRILE) 2021 guidelines (Nagendrababu et al. 2021). The flowchart is presented in Figure 1.

FIGURE 1.

FIGURE 1

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Flowchart for Preferred Reporting Items for Laboratory Studies in Endodontology (PRILE) 2021 guidelines.

2.1. Sample Collection

Dental pulp tissue of the patients with pulpitis was obtained during a regular pulpectomy procedure from patients visiting the Department of Endodontics, Affiliated Stomatology Hospital of Guangzhou Medical University, Guangzhou, China. Pulp tissue was collected from nine pulpitis teeth of the third molar from nine patients and 27 healthy premolars from nine healthy individuals undergoing orthodontic treatment (three teeth per person). The demographics and clinical information of the patients and healthy controls are listed in Table S1. This study was approved by the Institutional Review Board at the Guangzhou Medical University (approved number: KY. 2019065). All participants signed informed consent. The age of pulpitis patients and healthy controls recruited in this study was between 18–60 years. The patients with diabetes, hypertension, heart disease and other systemic inflammatory diseases were excluded.

Following informed consent, local anaesthesia was administered. In teeth with healthy pulps, the tooth was extracted and the root was rinsed with normal saline containing 4% penicillin–streptomycin, after which the periodontal membrane attached to the root surface was removed. The root tip was then cut using sterilised forceps and the pulp tissue was extracted and stored. For teeth affected by irreversible pulpitis, a rubber dam was applied. The carious portion was excavated to access the root canal system and pulp tissue was harvested with a sterile barbed broach.

2.2. Bioinformatic Analysis of piRNAs and mRNAs

2.2.1. RNA Extraction

Pulp tissue from six patients and eight healthy controls was used for bioinformatic analysis. Total RNA from dental pulp tissue was extracted using Trizol (Invitrogen, Carlsbad, CA, USA).

2.2.2. Library Construction and Sequencing

The library construction and sequencing were completed at the center of NovelBio laboratory (Shanghai, China) as described previously (Guo et al. 2021). Briefly, the extracted total RNA was connected to the 3′ and 5′ ends and RT Primer was used for reverse transcription and PCR amplification to create a stable and easy‐to‐use library for sequencing. Then, 150 bp libraries were recovered and the concentration was detected. Libraries larger than 1.0 ng/L were considered as qualified libraries and sRNA libraries were formed. After mixing and denaturation, the Illumina HiSeq X 10 sequencing platform was used for high‐throughput parallel sequencing.

Under the conditions of FDR ≤ 0.05 and |log2FC| ≥ 1, relevant target genes of differentially expressed piRNAs were predicted using miRanda and RNAhybrid. piRNA targets were identified by first determining potential target sites using sequence complementarity between each piRNA and the mRNA of interest, followed by an estimation of thermodynamic stability of RNA duplexes based on these alignments.

The differentially expressed piRNAs and their negatively correlated target genes were further obtained using miRanda and RNAhybrid. The piRNAs‐mRNAs network was mapped according to the targeting relationship (Wei et al. 2025). The network was constructed using Cytoscape.

2.3. Construction of Mouse Pulpitis Model

Many factors, such as age, sex, lifestyle, eating behaviour and oral health status, are associated with the variation in the degree of pulpitis and inflammation in patients. A mouse model with the same age, sex, eating behaviour and physical status helps to reduce the variation in individuals. Therefore, we established a mouse model of pulpitis. The animal experimental protocol was approved by the Institutional Animal Care and Use Committee (IACUC) of the First Affiliated Hospital of Guangzhou Medical University of China (approved number: 2017‐67) and all experimental procedures were conducted in accordance with the National Institutes of Health (NIH) Guide for the Care and Use of Laboratory Animals. Experiments were performed using 48 male C57BL/6 mice (6 weeks old, bodyweight 20–25 g) supplied by Guangdong Medical Laboratory Animal Center, China (experimental group n = 24, control group n = 24).

The mouse pulpitis model was developed as described previously (He et al. 2017). Briefly, the mice were anaesthetised by intraperitoneal injection with 1% pentobarbital sodium (0.13 mg/kg body weight) and fixed in the supine position. A total of 48 mice had their bilateral maxillary first molar pulps exposed with the aid of an ultrasonic tooth cleaning machine (miniMaster FT‐155700, Switzerland) using a polishing bar (TC‐11EF, MANI, Japan). The exposed pulp was then left open to the oral environment. Control group mice were sacrificed immediately after pulp exposure by cervical dislocation. According to the preliminary experiment of our research group, the peak of pulpitis was 8 h after pulp opening, both in terms of inflammation degree and pathological process. So, in the pulpitis group, after exposure for 8 h, 20 mice were selected to extract the maxillary first molars on both sides. The periodontal tissue of the root surface was removed. These samples were stored in liquid nitrogen. The maxillae from mice were fixed with 4% paraformaldehyde and the paraformaldehyde‐fixed maxillary jaws were decalcified in EDTA decalcification fluid for 2 weeks at 25°C. The decalcification solution was changed every 3 days until the tissue was soft enough to be cut with a surgical blade.

2.4. Histological and Immunohistochemical (IHC) Analysis

For H&E staining, human and mouse specimens were dehydrated, cleared, embedded in paraffin and sectioned into 4 μm slices. The sections were then stained with haematoxylin and eosin. Each slide was scanned using an automated digital slide scanning system (Precise 500B, China). The parameters of inflammatory cells and blood vessels numbers in human and mouse pulpitis tissue were performed independently by two blinded observers.

For IHC, tissue blocks were embedded in paraffin, sectioned and deparaffinised with xylene (Chongqing Chuangdong Chemical CO. LTD, China). Tissue sections were dehydrated using a graded ethanol series. Antigen retrieval was performed via high‐pressure heat treatment using a citrate buffer (pH 6.0; Beyotime, China). Blocking was carried out with 3% H2O2 (20 min, room temperature; Boster, China) and 5% bovine serum albumin (30 min, room temperature; Boster, China). The sections were subsequently incubated with primary antibodies overnight at 4°C, followed by incubation with corresponding horseradish peroxidase‐conjugated secondary antibodies (1 h, room temperature; Proteintech, China). The sections were then developed with diaminobenzidine and counterstained with haematoxylin (Beyotime, China). The antibodies used and their respective dilutions were as follows: IL‐6 (1:200, Proteintech, China), IL‐8 (1:200, Proteintech, China) and ADGRG2 (1:200, Affinity Biosciences, China).

IHC images were captured using the microscope image system (Servicebio, China). The pulp samples from three humans and mice (n = 3) of each group were employed for the semi‐quantitative analysis. Five random regions of interest per section were selected and imaged at 40× magnification. Semi‐quantitative analysis was conducted using ImageJ software (NIH, USA). All measurements were performed in a double‐blind manner by two independent investigators.

2.5. Isolation and Primary Culture of HDPCs

Dental pulp tissues were minced into small pieces and washed three times with PBS (Gibco, USA) containing 4% Penicillin–Streptomycin (P/S) (Gibco, USA). The tissue fragments were then digested using 3 mg/mL Collagenase Type I (Gibco, USA) and 4 mg/mL Dispase (Yuanye, China) at 37°C for 30 min. The digest was centrifuged at 1000 rpm for 5 min at 20°C. The resulting pellet was resuspended in α‐MEM (Gibco, USA) supplemented with 15% Fetal Bovine Serum (FBS) (Gibco, USA) and 1% P/S and incubated at 37°C in a 5% CO2 atmosphere. Cells from passages 2 to 3 were used for subsequent experiments.

2.6. Establishment of an In Vitro Pulpitis Model

HDPCs (passages 3–5) were seeded into six‐well plates at a density of 1 × 105 cells per well and cultured until 80% confluence. After 36 h, the cells were incubated in serum‐free medium containing 1 μg/mL lipopolysaccharide (LPS; from Escherichia coli O111:B4, L2630, Thermo Fisher, USA) at 37°C in a humidified atmosphere of 5% CO2 for 24 or 48 h.

2.7. RNA Extraction and Quantitative Real‐Time PCR (qRT‐PCR)

Total RNA was extracted with TRIzol reagent. Then, cDNAs for piRNAs and mRNAs were reversed with PrimeScript RT reagent Kit (TaKaRa, Japan). piRNAs' reverse transcription primers and primers for qPCR were purchased from RiboBio (Guangzhou, China). The expression levels of piRNAs and target genes were determined by TB Green Premix Ex Taq II (TaKaRa, Japan). Amplification was performed as follows: 95°Cfor 10 s, followed by 40 amplification cycles of 95°C for 5 s, 60°C for 20 s and 95°C for 60 s, 60°C for 30 s, 95°C for 30 s. All reactions were performed in triplicate. U6 was used as an internal reference for piRNAs, while GAPDH was used as the internal reference for target genes. The data were analysed by the 2−ΔΔCT method. Sequences of primers used for RT‐qPCR are shown in Table 1.

TABLE 1.

Primer sequence for quantitative reverse transcription polymerase chain reaction.

Human gene Primer sequence (5′‐3′) Mice gene Primer sequence (5′‐3′)
DCX F‐CAGGGAGTGCGTTACATTT Spock1 F‐AACAAGCCCTTCGACCA
R‐CAGACCAGTTGGGATTGAC R‐ATTGCCCTTCTTCTGCCT
FAM19A3 F‐GCAGCAGTGGACACAAAG Fam19a3 F‐CTTTTGGACAGCACTGGAA
R‐TGGCTGATGACTGGATACC R‐TGGAGGGGTGATTCTGAC
GFRA1 F‐CGATCCCAGACAAACCC Gfra1 F‐CAGCTACGGGATGCTCTT
R‐CAAGTTCAGCTCCAAATGC R‐GGCCTCTCTCGTTCTTCAT
IGF2BP1 F‐GGAGATGGTGCAGGTGTT Ajap1 F‐GCTGCGCTCATCACAACT
R‐AGGTGTTTCGGGTGGTG R‐TCTGGCAACTCTCCTCCTG
LYPD6 F‐GTGCCTGGAGGAGGATTT Lypd6 F‐ATCCTTGCACCTTGGCT
R‐GGTGCCAAGTCAAGGAGA R‐GGAAGAGAACTGGCACTCA
MAPT F‐ATTCGGTTCCCTGTCTCCT Mapt F‐CTGGCTTCCTTCCCTCTAA
R‐AGGGCTCTCTCCATGTCAA R‐CCCTCATCCCCACTTACTT
NOS1 F‐ATCGTTTCCTACCGCACTCG Nos1 F‐GGAGGATGCTGGTGTGTT
R‐GCAGGGGTTCATTCCTTTGT R‐AGGCGGTTGGTCACTTC
SMPD3 F‐GGGTACATCGCCTGCAC Smpd3 F‐CGCTGTGGTTTGCCTGT
R‐CGCTGGACGAGGAGGTA R‐ATCCCTGTGTCGGCCTT
SNAP91 F‐GCCAGCAAAGGTGGATT Snap91 F‐CACCGATTTCAGACCCA
R‐TCACTGGAGATGGGGAAG R‐GCAAAGGCATCTCCAAAG
TFPI2 F‐CAGGAAATAACGCGGAGAT Adgrg2 F‐CAATGGCACACCAGATGA
R‐TCCCAGGTGTAGAAATTGTTG R‐GAACCAAGACCACGATGAA
ADGRG2 F‐TGCATCAGGCGTCAAAC Gapdh F‐AAGAAGGTGGTGAAGCAGG
R‐GGGGAACAGTGCTTTCTTT R‐GAAGGTGGAAGAGTGGGAGT
AJAP1 F‐CATCACCGTCTCCCTCATC
R‐TGCTGGTTGGTCTTCCG
CDH4 F‐GACCCCGTAACCAACGAG
R‐CGTGGACTGGAAGGACATC
SPOCK1 F‐AAGAGGATGAGGTCGGGTA
R‐CCTTGGTGCCTTGGAGT
GAPDH F‐GGACCTGACCTGCCGTCTAG
R‐GTAGCCCAGGATGCCCTTGA
Name of primer Primer sequence (5′‐3′)
piR‐36241 GGGGGATTAGCTCAAATGGTAGAGCGCTCG
piR‐36340 GGTCGCTGGTTCGTTTCCGGCTCGAAGGACC
piR‐36252 GGGGGTGTAGCTCAGTGGTAGAGAGCGTGCT
piR‐31686 ATCCCCTGGTCGTCTAGTGGTTAGGATTCGGC
U6 F‐GGAACGATACAGAGAAGATTAGC
R‐TGGAACGCTTCACGAATTTGCG
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Abbreviations: F, forward; R, reverse.

2.8. Fluorescence In Situ Hybridisation (FISH) Assay

The FISH experiment was carried out to detect the subcellular localisation of piR‐36241. A FISH kit (Servicebio, China) was employed, following the manufacturer's instructions. The hybridisation process involved the utilisation of Cy3‐labelled piR‐36241 probes from Servicebio, China. DAPI staining solution, which contained a fluorescence quencher (Servicebio, China), was applied to the slides and coverslips were placed over them for sealing. Finally, the results were analysed through an upright fluorescence microscope (Nikon, Japan).

2.9. Western Blotting Assay

HDPCs were washed three times with ice‐cold PBS. The cells were then lysed using RIPA lysis buffer (Beyotime, China) supplemented with PMSF (Sigma, USA). Protein concentration was determined with a BCA protein assay kit (Beyotime, China). The proteins were separated by 10% sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS‐PAGE) (Epizyme, China) and transferred onto a PVDF membrane. Subsequently, the membrane was blocked with 5% skimmed milk for 1 h at room temperature, followed by an overnight incubation at 4°C with the following primary antibodies: anti‐GAPDH (1:10000, Proteintech, China) and anti‐ADGRG2 (1:500, Abnova, China). The next day, the membrane was incubated with appropriate secondary antibodies for 1 h at room temperature. Finally, the protein bands were visualised using BeyoECL Plus substrate (Beyotime, China).

2.10. Enzyme‐Linked Immunosorbent Assay

Supernatants from HDPCs under different treatment conditions were collected. The secretion of IL‐6 and IL‐8 inflammatory cytokines was detected by the commercial ELISA kit (FindTest, China). Briefly, following incubation of the supernatant with the specific substrate for 4 h at 37°C, the change in OD at 450 nm was detected with a microplate analyser.

2.11. Transient Cell Transfection

Once the population of HDPCs reaches 70%–80% confluence, we transfected piR‐36241 mimic and its negative control (mimic NC), piR‐36241 inhibitor and its negative control (inhibitor NC), along with siADGRG2 and its negative control (siNC) into HDPCs. This was done using Lipofectamine 2000 Transfection Reagent (Thermo Fisher, USA) at a final concentration of 100 nmol, following the manufacturer's instructions.

2.12. Dual‐Luciferase Reporter Assay

The miRanda database was used to predict the putative target sites of pir‐36241 and ADGRG2. The mutant sequence of ADGRG2 was designed according to the predicted results. The four groups (ADGRG2 3'UTR‐WT+mimic‐NC, ADGRG2 3'UTR‐WT+piR‐36241 mimic, ADGRG2 3'UTR‐MUT+mimic‐NC, ADGRG2 3'UTR‐MUT+piR‐36241 mimic) were transfected into HEK‐293 T cells, respectively. The fluorescence intensity of each group was measured using Dual Luciferase Reporter Gene Assay Kit (Beyotime, China). GraphPad Prism software was used to analyse the data.

2.13. Statistical Analysis

All statistics were analysed using GraphPad prism 7. Mean ± standard deviation (SD) was used to show the data spreading. The two‐tailed student's t‐test was used to test for statistical significance between the two cohorts; a p‐value of < 0.05 was considered to indicate a statistically significant difference.

3. Result

3.1. Histological Changes in Human and Mouse Pulpitis Samples

H&E staining results showed a well‐arranged odontoblast layer and spindle‐shaped pulp cells in healthy dental pulp. In contrast, inflamed pulp tissues exhibited a disorganised odontoblast layer, vasodilation and extravasated red blood cells (Figure 2a). Compared with the control group, human pulpitis samples showed a significant increase in the number of dilated blood vessels. Moreover, the number of inflammatory cells in the pulpitis group was 4.43 times higher than that in normal pulp tissues (Figure 2b). These pathological alterations were similarly pronounced in mouse pulpitis tissue sections (Figure 2a,c). Immunohistochemical staining further revealed significantly elevated expression levels of IL‐6 (Figure 2d–f) and IL‐8 (Figure 2g–i) in inflamed human and mouse pulp tissues.

FIGURE 2.

FIGURE 2

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Histopathological features and inflammatory marker expression in pulpitis. (a–c) Representative H&E staining of human and mouse pulp tissues. Inflamed pulp exhibits a disorganised odontoblast layer, vasodilation and increased inflammatory infiltrate compared to the well‐arranged structure of healthy pulp. Immunohistochemical analysis reveals significantly elevated expression levels of the pro‐inflammatory cytokines IL‐6 (d–f) and IL‐8 (g–i) in human (scale bar, 100 μm) and mouse pulpitis tissues (scale bar, 200 μm). *p < 0.05i, **p < 0.01, ***p < 0.001. Each test was repeated at least three times.

3.2. Differentially Expressed piRNAs in Dental Pulp Tissues of Pulpitis Patients

In total, 210 known piRNAs were identified from the pulp tissues. Among these, 111 known piRNAs were detected in the pulpitis group and 99 in the healthy control group. As shown in Figure 3a, the differentially expressed piRNAs exhibited distinct clustering between the groups. We further analysed the differences in piRNA expression between the pulpitis and healthy control groups. Volcano plot analysis revealed 21 differentially expressed piRNAs in the pulpitis group compared to the healthy controls (Figure 3b). Among these, 15 piRNAs were significantly upregulated and six were significantly downregulated (|log2FC| ≥ 1, FDR ≤ 0.05) (Figure 3a,b).

FIGURE 3.

FIGURE 3

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Comparison and high‐throughput sequencing analysis of human healthy and pulpitis pulp tissue. (a) Shows the heatmap indicating differentially regulated piRNAs expression in pulpitis tissue (|log2FC| ≥ 1, FDR ≤ 0.05). (b) Volcano plot identifying 21 differentially expressed piRNAs (15 up, 6 down; |log2FC| ≥ 1, FDR ≤ 0.05). 4 piRNAs with |log2FC| ≥ 2 are labelled. (c) RT‐qPCR validation confirms significant upregulation of piR‐36241 in human (1) and mouse (2) pulpitis tissues. (d) Interaction network depicting piR‐36241 and its predicted target mRNAs. (e) A heatmap of differentially expressed target mRNAs in pulpitis tissue (log2FC ≤ −1, FDR ≤ 0.05). (f) RNA‐seq analysis shows expression levels of 14 candidate target mRNAs of piR‐36241 in pulpitis. (g) RT‐qPCR validation of target genes that are consistently downregulated in human pulpitis. (h) RT‐qPCR validation of seven key target genes (ADGRG2, APAJ1, GFRA1, MAPT, NOS1, SMPD3, SPOCK1) that are consistently downregulated in mouse pulpitis. *p < 0.05, **p < 0.01, ***p < 0.001 and NS (not significant). Each test was repeated at least three times.

Target gene prediction performed using miRanda and RNAhybrid indicated that the 15 upregulated piRNAs targeted 81 genes. Similarly, the six downregulated piRNAs targeted 332 genes. Interaction network demonstrated that all target genes of the differentially upregulated piRNAs were downregulated in pulpitis tissues compared to healthy pulp (Figure S1). Conversely, all target genes of the differentially downregulated piRNAs were upregulated in pulpitis tissues.

Among the differentially expressed piRNAs, piR‐36241, piR‐36340 and piR‐36252 were significantly upregulated, while piR‐31686 was significantly downregulated (|log2FC| ≥ 2, FDR ≤ 0.05) (Figure 3b). We further validated the expression of these piRNAs in human and mouse pulp tissues by RT‐qPCR. The expression level of piR‐36241 was significantly upregulated in human inflammatory pulp tissue samples and in a mouse model of pulpitis compared to their respective healthy controls (Figure 3c, panels 1 and 2). In contrast, the expression levels of piR‐36340, piR‐36252 and piR‐31686 showed no significant differences between inflammatory and healthy human pulp tissues (Figure S2).

PiRNAs are known to silence gene expression in a sequence‐dependent manner in conjunction with PIWI proteins during development (Iwasaki et al. 2015). Consequently, piRNA expression levels are typically negatively correlated with their target mRNAs. An interaction network for piR‐36241 (Figure 3d) and its predicted targets and heatmap indicating target mRNA was constructed (log2FC ≤ −1, FDR ≤ 0.05) (Figure 3e). Analysis of RNA‐seq data revealed that 14 target mRNAs of piR‐36241 were expressed at lower levels in pulpitis (Figure 3f). These mRNA targets of piR‐36241 were further validated by RT‐qPCR in human and mouse samples. Compared to control samples, seven target genes were significantly reduced in both human and mouse pulpitis samples (Figure 3g,h). These seven genes are: ADGRG2, APAJ1, GFRA1, MAPT, NOS1, SMPD3 and SPOCK1.

3.3. Screening and Verification of ADGRG2

Subsequently, pro‐inflammatory treatment of HDPCs confirmed that piR‐36241 is localised in the cytoplasm and simultaneously upregulated the expression of piR‐36241, IL‐6 and IL‐8 (Figure 4a,b). Concurrently, only three target mRNAs showed significantly reduced expression: ADGRG2, GFRA1 and SPOCK1 (Figure 4c). Western blot results demonstrated that only ADGRG2 protein expression was significantly downregulated (Figure 4d–f). Consistent with these findings, IHC revealed a significant reduction of ADGRG2 expression in human and mouse pulpitis tissues (Figure 4g).

FIGURE 4.

FIGURE 4

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Screening and verification of ADGRG2. (a) FISH was performed to detect the localisation of piR‐36241 in HDPCs (scale bar, 20 μm). (b) Expression levels of piR‐36241, IL6 and IL8 in LPS‐treated HDPCs is determined by RT‐qPCR (1) and ELISA (2). (c) RT‐qPCR analysis identifies GFRA1, SPOCK1 and ADGRG2 as significantly downregulated mRNAs. (d‐f) Western blot analysis of GFRA1, SPOCK1 and ADGRG2 protein expression in LPS‐treated HDPCs. The results confirm the specific downregulation of ADGRG2. (g) Immunohistochemistry validates decreased ADGRG2 expression in human and mouse pulpitis tissues (scale bar, 100 μm). *p < 0.05, **p < 0.01, ***p < 0.001 and NS (not significant). LPS‐HDPCs, Lipopolysaccharide‐induced HDPCs. Each test was repeated at least three times.

3.4. PiR‐36241 Negatively Regulates ADGRG2 to Promote the Development of Pulpitis

To further validate interaction of piR‐36241 and ADGRG2, a dual‐luciferase reporter assay was performed. The results showed that piR‐36241 binding significantly reduced luciferase activity when paired with the wild‐type ADGRG2 3′UTR. In contrast, no such reduction was observed when paired with the mutant ADGRG2 3′UTR (Figure 5a).

FIGURE 5.

FIGURE 5

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Validation of the relationship between piR‐36241 and ADGRG2 in pulpitis. (a) Dual‐luciferase reporter assay in 293T cells confirms that piR‐36241 directly binds to ADGRG2. (b, c) In LPS‐HDPCs, RT‐qPCR shows piR‐36241 mimics downregulate ADGRG2 and upregulate IL‐6 and IL‐8, while its inhibitor has opposite effects. (d) Western blot analysis of ADGRG2 expression in LPS‐HDPCs transfected with piR‐36241 mimics or inhibitor. (e, f) IL‐6 and IL‐8 expression determined by ELISA in LPS‐HDPCs transfected with piR‐36241 mimics or inhibitor. *p < 0.05, **p < 0.01, ***p < 0.001. LPS‐HDPCs, Lipopolysaccharide‐induced HDPCs. Each test was repeated at least three times.

Subsequently, we transfected cells with piR‐36241 mimics or inhibitor to further investigate the regulatory role of ADGRG2 in pulpitis progression.

In LPS‐treated HDPCs transfected with piR‐36241 mimics, the expression of ADGRG2 was downregulated, while the expression of IL‐6 and IL‐8 was upregulated, indicating that increased levels of piR‐36241 promote inflammatory development. Conversely, transfection with the piR‐36241 inhibitor in LPS‐treated HDPCs yielded opposite results (Figure 5b–f).

3.5. Functional Validation of a Mechanistic Link Between piR‐36241 and ADGRG2

We then knocked down ADGRG2 using siRNA interference. The knockdown efficiency was confirmed by RT‐qPCR (Figure 6a) and Western blotting (Figure 6b). Knockdown of ADGRG2 led to increased expression of IL‐6 and IL‐8 in LPS‐treated HDPCs (Figure 6c,d).

FIGURE 6.

FIGURE 6

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Validation of the relationship between piR‐36241 and ADGRG2. Knockdown efficiency of ADGRG2 siRNA, confirmed at mRNA (a) and protein (b) levels. Knockdown of ADGRG2 alone in LPS‐HDPCs increases the expression of IL‐6 and IL‐8 by RT‐qPCR (c) and ELISA (d). Rescue experiments: Inhibition of piR‐36241 suppresses IL‐6 and IL‐8 expression in the level of mRNA (e) and protein (f), but this effect is abolished when ADGRG2 is co‐knockdown. *p < 0.05, **p < 0.01, ***p < 0.001. LPS‐HDPCs, Lipopolysaccharide‐induced HDPCs. Each test was repeated at least three times.

Furthermore, inhibition of piR‐36241 alone in LPS‐treated HDPCs significantly reduced the expression of IL‐6 and IL‐8. However, when ADGRG2 was simultaneously knocked down, this suppressive effect on IL‐6 and IL‐8 expression was abrogated and their levels increased instead (Figure 6e,f). This experiment showed that silencing of ADGRG2 reverses the functional effects of piR‐36241 knockdown on pulp inflammation.

4. Discussion

Pulpitis represents a dynamic host inflammatory response to microbial invasion of the dental pulp, where the balance between tissue destruction and repair dictates clinical outcomes (Brizuela et al. 2025; Liu, Li, et al. 2025). ncRNAs have emerged as pivotal regulators of such inflammatory processes. While roles for miRNAs, lncRNAs and circRNAs in pulpitis are established (Xu et al. 2025; Hu et al. 2025; Lin et al. 2025; Jiang et al. 2023), the involvement of piRNAs remained unknown. piRNAs, typically 24–31 nucleotides in length, complex with PIWI proteins to silence transposable elements and regulate gene expression epigenetically and post‐transcriptionally (Portell‐Montserrat et al. 2025; De et al. 2025; Claro‐Linares and Rojas‐Rios 2025). The biogenesis of piRNA begins in the cytoplasm, where large precursor transcripts are cleaved by endonucleases to process piRNA precursors, generating the 5′ and 3′ ends of mature piRNAs (Huang et al. 2017; Lee et al. 2025). In the cytoplasm, piRNAs function primarily by amplifying transposon silencing via the ping‐pong cycle, post‐transcriptionally repressing target mRNAs through cleavage or translational inhibition and regulating cell maintenance and differentiation (Portell‐Montserrat et al. 2025). This study provides the first comprehensive analysis of piRNA expression in human pulpitis, revealing significant dysregulation of specific piRNAs, including the markedly upregulated piR‐36241. The biological functions of piRNAs were first characterised in germline cells (Gao et al. 2025; Gan et al. 2025), where they mediate transposon silencing to protect genomic integrity in mammalian testes and influencing the post‐meiotic transcriptome (Stallmeyer et al. 2024), disruption of piRNA biogenesis can therefore impair these essential processes. Beyond the germline, their functional relevance in somatic inflammation is increasingly recognised, as seen in airway diseases (Corsello et al. 2025) and rheumatoid arthritis (Ren et al. 2023). Recent work reveals a pro‐inflammatory role for the overexpressed piRNA, piENOX2, which operates in collagen‐induced arthritis models by degrading Alkbh5 mRNA to alter macrophage activation and inflammation (Feng et al. 2025). The pathogenesis of pulpitis is driven by excessive production of pro‐inflammatory cytokines (Sabeti et al. 2025, 2024; Bas et al. 2024). Bacteria release substantial amounts of toxic lipopolysaccharide (LPS) during replication and death. LPS activates the NF‐κB and MAPK signalling pathways and induces immune responses via reactive oxygen species (ROS), leading to the robust synthesis and release of pro‐inflammatory factors. It has been demonstrated that these pro‐inflammatory factors can be released by odontoblasts (Zhou et al. 2021), dental pulp fibroblasts (Coil et al. 2004), stem cells and tissue‐resident immune cells (Zhang et al. 2025; Xu et al. 2024). Additionally, immune cells, including neutrophils (Guo, Wu, et al. 2025), monocytes (Erdogan et al. 2023) and macrophages (Wang et al. 2026), participate in the pathophysiology of pulpitis, where they release inflammatory cytokines and amplify the inflammatory response. Consistent with this, studies have shown that the levels of IL‐6 and IL‐8 in the pulp of patients with irreversible pulpitis are significantly higher than those in patients with caries‐exposed or normal teeth (Elsalhy et al. 2013; Sabeti et al. 2024). Studies have shown that IL6 and IL8 are secreted by HDPCs (He et al. 2013; Widbiller et al. 2018) which have a high capacity for differentiation, the ability to regenerate a dentine/pulp‐like complex and great potential for various applications in stem cell research and regenerative medicine (Nakashima and Iohara 2017; Jeong et al. 2020). They are also characterised by easy accessibility and well‐defined properties. Therefore, we selected HDPCs as the research object in this study. Reducing the release of these inflammatory factors could not only help alleviate patient pain but also hold promise for restoring pulp cell vitality.

Our data position piR‐36241 as a key modulator of this inflammatory cascade. Its upregulation, likely a maladaptive response to infection, initiates a pathogenic sequence by silencing the expression of ADGRG2. This suppression of a putative protective receptor may disrupt crucial homeostatic checks, thereby facilitating the progression of inflammation.

G protein‐coupled receptors (GPCRs), which play diverse physiological roles and represent major targets for pharmaceutical research, possess a long extracellular N‐terminus involved in cell–cell and cell‐matrix interactions, which is coupled to the classic GPCR signalling domain (Khan et al. 2025; Gurevich and Gurevich 2025; Fan et al. 2024). As the second‐largest family within the GPCR superfamily, adhesion GPCRs (aGPCRs) hold promise as therapeutic targets for a wide spectrum of diseases spanning obesity (Suchy et al. 2020), psychiatric disorders (Zhu et al. 2025), cancer (Liu, Tian, et al. 2025; Mooney et al. 2024) and inflammation (Chu et al. 2022; Hsiao et al. 2018). ADGRG2 is an orphan aGPCR whose functions extend beyond its established role in male fertility (Tan et al. 2022; Zhang et al. 2020) to include somatic processes such as tissue development, cell adhesion and migration (Peeters et al. 2015; Richter et al. 2013; Grunddal et al. 2021). Notably, emerging evidence links ADGRG2 to inflammatory regulation through pathways like cAMP‐CREB and calcium signalling and it shows a strong correlation with neutrophil activity (Balenga et al. 2017; Wu et al. 2023). The function of ADGRG2 appears context‐dependent, exhibiting both pro‐ and anti‐inflammatory roles across different tissues (Karagiannis et al. 2013; Galligan et al. 2007). Our findings—consistent downregulation of ADGRG2 in pulpitis tissues and LPS‐HDPCs—strongly suggest it serves a protective, homeostatic function within the dental pulp. We hypothesise that ADGRG2 signalling helps fine‐tune the local immune response, possibly by promoting resolution pathways. Thus, its silencing by piR‐36241 could remove a critical brake on inflammation, leading to uncontrolled tissue damage. This loss compromises the structural defence of the pulp, its ability to control inflammation and its repair capacity, collectively contributing to disease aggravation. Therefore, restoring ADGRG2 function could represent a novel therapeutic strategy aimed at preserving pulp vitality, a hypothesis that warrants rigorous future investigation.

In future studies, we will focus on evaluating the feasibility of using ADGRG2 as a diagnostic biomarker, for instance, in gingival crevicular fluid or saliva. Meanwhile, local delivery of ADGRG2 via hydrogels or nanoparticles represents a promising therapeutic strategy to maintain pulp vitality. Nevertheless, several challenges remain, including delivery efficiency, off‐target effects and the need for further preclinical validation. Overall, we believe that this complementary perspective provides a tangible and forward‐looking view of the clinical relevance of our findings.

The piR‐36241/ADGRG2 axis was rigorously validated through multiple lines of evidence. Dual‐luciferase reporter assay confirmed direct binding of piR‐36241 to the ADGRG2 3′UTR. An inverse correlation between piR‐36241 upregulation and ADGRG2 downregulation was observed in human clinical samples, mouse pulpitis samples and cellular models. Functionally, piR‐36241 mimics suppressed ADGRG2 and enhanced pro‐inflammatory cytokine release in HDPCs, while ADGRG2 knockdown replicated this inflammatory phenotype. Critically, rescue experiments demonstrated that the pro‐inflammatory effect of piR‐36241 is primarily mediated through its targeting of ADGRG2.

In conclusion, our study unveils a novel epigenetic regulatory axis in pulpitis, where upregulation of piR‐36241 exacerbates disease progression by repressing the protective receptor ADGRG2. These findings not only expand the known ncRNA network in pulp biology but also identify the piR‐36241/ADGRG2 pathway as a promising therapeutic target. Strategies aimed at inhibiting piR‐36241 or restoring ADGRG2 function could potentially modulate the inflammatory milieu, offering a new avenue for preserving pulp vitality and moving beyond conventional root canal therapy.

5. Conclusions

In summary, this study reveals, for the first time, the significant involvement of piRNAs in the pathogenesis of pulpitis. We have identified a novel pathogenic axis wherein upregulation of piR‐36241 exacerbates pulpal inflammation by directly silencing the expression of the orphan receptor ADGRG2. Our findings, substantiated by clinical tissue analysis, in vitro functional experiments and mechanistic validation, establish the piR‐36241/ADGRG2 pathway as a critical regulator of inflammatory homeostasis in the dental pulp. The suppression of ADGRG2 disrupts a key protective mechanism, thereby unleashing an unrestrained inflammatory cascade. These insights not only expand the understanding of epigenetic regulation in pulpitis but also highlight this pathway as a potential therapeutic target for preserving pulp vitality.

Author Contributions

W.H., Y.C., L.W. and X.Y. contributed to the study design. W.H., Y.C., Y.J. and M.Y. performed the experiments and were responsible for clinical sample collection. W.H., Y.C. and Y.J. established the in vitro and in vivo pulpitis models. W.H. and Y.C. conducted the statistical analysis and prepared the diagrams and figures. W.H., Y.C. and M.Y. were responsible for H&E and immunohistochemical staining. W.H. performed the cell transfection experiments. W.H. and Y.C. wrote the initial draft of the manuscript. L.W. and X.Y. participated in manuscript revision. X.Y. supervised and consulted on the study, participated in drafting the manuscript and assisted in its revision. All authors have read and approved the final manuscript.

Funding

This work was supported by the Natural Science Foundation of Guangdong Province (No. 2023A1515012777), the Featured Clinical Technique of Guangzhou (No. 2023C‐TS58), the Special projects in key fields of Guangdong Colleges and Universities (No. 2021ZDZX2058), the Plan on enhancing scientific research in GMU (No. GMUCR2024‐02024) and the Science and Technology Planning Project of Guangdong Province of China (No. 2025A0505020087).

Ethics Statement

This study was approved and supervised by the Ethics Committee of the Affiliated Stomatology Hospital of Guangzhou Medical University, Guangzhou, China (KY. 2019065). Animal experiments were approved by the Institutional Animal Care and Use Committee (IACUC) of the First Affiliated Hospital of Guangzhou Medical University of China (approved number: 2017‐67).

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Figure S1: Interaction networks analysis in human pulp tissues.

Figure S2: Expression levels of piR‐36 340 (a), piR‐36 252 (b) and piR‐31 686 (c) in human pulp tissues.

Table S1: Patients' demographic and clinical information.

IEJ-59-2164-s001.docx (676.6KB, docx)

Data Availability Statement

The data that support the findings of this study are available on request from the corresponding author. The data are not publicly available due to privacy or ethical restrictions.

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Associated Data

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

Supplementary Materials

Figure S1: Interaction networks analysis in human pulp tissues.

Figure S2: Expression levels of piR‐36 340 (a), piR‐36 252 (b) and piR‐31 686 (c) in human pulp tissues.

Table S1: Patients' demographic and clinical information.

IEJ-59-2164-s001.docx (676.6KB, docx)

Data Availability Statement

The data that support the findings of this study are available on request from the corresponding author. The data are not publicly available due to privacy or ethical restrictions.

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