MRGPRX2 Mediates Mast Cell-Induced Endometriosis Pain Through the Sensitization of Sensory Neurons via Histamine/HRH1/TRPV1 Signaling Pathway

X.M., J.W., S.D., X.G., X.X., and X.Z. conceived the study. X.M., S.D., and X.G. were involved in developing the methodology. X.M., J.W., S.D., X.G., X.X., P.X., and X.Z. participated in the investigation. X.M., X.G., and X.X. performed the visualization. X.Z., S.D., and P.X. provided supervision and acquisition of funding. X.M. and J.W. wrote the original manuscript. X.M., J.W., S.D., X.G., X.X., P.X., and X.Z. reviewed and edited the manuscript. All authors reviewed and approved the manuscript.

This study was conducted according to the ethical policies and procedures approved by the Human Ethics Committee of Women's Hospital, School of Medicine, Zhejiang University (No. 20230091). All the animal experiments and the protocols were approved by the Ethics Committee of Zhejiang University (No. ZJU20230165).

Toluidine blue staining demonstrated that the density of mast cells was significantly greater in ectopic endometrial tissues than in eutopic endometrial tissues and controls (Figure  1A,B ). Also, the number of degranulated mast cells in the stroma of lesion samples from endometriosis patients was significantly greater than in samples of eutopic endometrium from patients with and without endometriosis (Figure  1A,C ), indicating that mast cells played a vital role in endometriosis. Mast cells mediated hyperalgesia in an endometriosis model, and the density of MRGPRX2‐positive mast cells was increased in endometriotic lesions. (A) Toluidine blue‐stained section of control, eutopic and ectopic endometria (Arrow: degranulated mast cells). (B) Number of mast cells per field of microscopic vision in the control ( n  = 10), eutopic ( n  = 10), and ectopic groups ( n  = 19). (C) Number of degranulated mast cells per field of microscopic vision in the control ( n  = 10), eutopic ( n  = 10), and ectopic groups ( n  = 19). (D) The mechanical threshold and (E) thermal latency were measured in three groups, including the WT sham, WT EMS, and Sash EMS groups ( n  = 10 in each group). (F) Images of double immunofluorescence staining for tryptase (green) and MRGPRX2 (red) in control, eutopic, and ectopic endometria. The white arrow indicates positive cells; 200×, scale bar = 50 μm. (G) Number of MRGPRX2‐positive mast cells per field of microscopic vision in the control ( n  = 8), eutopic ( n  = 10), and ectopic groups ( n  = 19). (H) Number of MRGPRX2‐positive mast cells per field of microscopic vision in the ectopic lesions of the pain ( n  = 10) and no pain ( n  = 9) groups. Control, endometrial tissue from patients without endometriosis; ectopic, endometriotic lesions from patients; eutopic, endometrial tissue from patients with endometriosis; no pain, endometriotic lesions from patients without endometriosis pain; pain, endometriotic lesions from patients with endometriosis pain.; Sash EMS, mast cell‐deficient mice with endometriosis surgery; WT EMS, wild‐type mice with endometriosis surgery; WT Sham, wild‐type mice with the sham operation. *Compared with the WT Sham group; # compared with the WT EMS group. ** p  < 0.01, *** p  < 0.001, **** p  < 0.0001,  ### p < 0.001, #### p < 0.0001, and ns, not significant. Our previous studies revealed that mast cells mediate endometriosis‐related pain in vitro and in a traditional mouse endometriosis model [ 7 , 31 ]. However, mast cell‐deficient mice should be a better model for investigating the role of mast cells in endometriosis because this mutant is profoundly deficient in mast cells in all organs and tissues [ 32 ]. Therefore, mast cell‐deficient mice (sash) were used, and endometriosis model surgery and behavior tests were performed to further confirm whether mast cells play a key role in endometriosis pain. WT mice served as controls. The results revealed pain‐related behavioral changes in both the sash and WT EMS groups at 1–4 weeks after surgery compared to the WT Sham group. The decreases in the mechanical pain threshold (MPT) and heat source latency (HSL) in the sash EMS group were smaller than those in the WT EMS group; therefore, the sash mice were less sensitive to mechanical and thermal stimuli (Figure  1D,E ). The Reflexive withdrawal tests in response to mechanical and thermal stimulation of the abdominal area were conducted in the WT sham, WT EMS, and Sash EMS groups. The results were consistent with those of the mechanical and thermal paw withdrawal tests (Figure  S1A,B ). The above data further confirmed that the mast cell deficiency might alleviate endometriosis‐induced pain in a mast cell‐deficient mouse model, indicating that mast cells mediate hyperalgesia in endometriosis. MRGPRX2 is an important mast cell receptor that plays a key role in neurogenic inflammation and pain [ 14 , 18 ]. However, whether MRGPRX2 mediates mast cell‐induced endometriosis pain is unclear. Double immunofluorescence staining was performed to investigate the density of MRGPRX2‐positive mast cells in endometriosis lesions. The results revealed that the density of MRGPRX2‐positive mast cells was significantly greater in ectopic endometrial tissues than in eutopic endometrial tissues and controls (Figure  1F,G ). Additionally, more MRGPRX2‐positive mast cells per field were observed in ectopic lesions from patients with pain than in those from patients without pain (Figure  1F,H ), indicating that the mast cell receptor MRGPRX2 might be involved in endometriosis pain. We established an MRGPRX2 knockout (MRGPRX2‐KO) HMC1.1 cell line via the CRISPR/Cas9 gene editing system to investigate the function of MRGPRX in mast cells (Figure  2A ). The efficiency of the knockout was verified by PCR using agarose gel electrophoresis, RT–qPCR, and western blotting (Figure  2B–D ), and the results revealed that the expression level of MRGPRX2 was significantly reduced in MRGPRX2‐KO HMC1.1 cells. In addition, the sequencing data for the MRGPRX2‐KO HMC1.1 cells were presented in Supporting Information  2 . If MRGPRX2 mediates endometriosis pain, HBD‐2, the MRGPRX2 ligand, might be involved in this process. Immunohistochemistry was used to detect its location and expression. As shown in Figure  2E,F , the expression of HBD‐2 was elevated in the ectopic group ( n  = 13) compared with the control ( n  = 21) and eutopic ( n  = 13) groups ( p  < 0.0001). Moreover, we found that the protein was mostly present in endometriotic epithelial cells and present in some of the stromal cells of endometriotic lesions. The concentrations of HBD‐2 in the cell supernatants of control ( n  = 3), eutopic ( n  = 3), and ectopic endometrial cells ( n  = 4) were evaluated by ELISA to explore whether HBD‐2 was secreted from endometrial cells (Figure  2G ). Our data revealed that HBD‐2 levels were increased in the supernatants of ectopic endometrial cells compared with those of the control and eutopic groups. These results indicated that endometriotic cells could release HBD‐2 in the endometriosis environment. Endometriotic cell‐derived HBD‐2 mediated histamine secretion from mast cells via the activation of MRGPRX2. (A) Strategy for MRGPRX2 knockout in a mast cell line (HMC1.1 cells). The expression of MRGPRX2 in HMC1.1 cells after knockout was examined by (B) PCR using agarose gel electrophoresis, (C) RT–qPCR and (D) western blotting. (E) IHC images showing HBD‐2 expression and distribution in control ( n  = 2 L), eutopic ( n  = 13), and ectopic endometria ( n  = 13); 400×, scale bars = 100 μm. (F) IHC scores for each group. (G) Concentrations of HBD‐2 in the supernatants of control ( n  = 3), eutopic ( n  = 3), and ectopic endometrial cells ( n  = 4). (H) Concentrations of histamine in the supernatants of HMC1.1 and MRGPRX2‐KO HMC1.1 cells, including HMC1.1, HMC1.1 + HBD‐2, MRGPRX2‐KO HMC1.1, and MRGPRX2‐KO HMC1.1 + HBD‐2 cells, after HBD‐2 treatment. The target concentration of HBD‐2 was 40 μg/mL. Control, endometrial tissue from patients without endometriosis; ectopic, endometriotic lesions from patients; eutopic, endometrial tissue from patients with endometriosis; KO, MRGPRX2‐KO HMC1.1 cells; WT, wild‐type HMC1.1 cells. * p < 0.05, ** p  < 0.01, *** p < 0.001, **** p  < 0.0001, and ns, not significant. HMC1.1 cells and MRGPRX2‐KO HMC1.1 cells were then treated with or without 40 μg/mL HBD‐2 for 1 h, and the concentrations of histamine in the supernatants of the four groups were evaluated by ELISA. Upon stimulation with HBD‐2, a significant increase in histamine release was detected in the supernatant of HMC1.1 cells ( p   0.05; Figure  2H ), indicating that HBD‐2/MRGPRX2 mediated histamine release from mast cells. Taken together, our data suggested that endometriotic cell‐derived HBD‐2 mediated histamine secretion from mast cells via the activation of MRGPRX2. We investigated the effect of histamine on dorsal root ganglion (DRG) neurons to explore whether and how histamine participates in endometriosis pain. First, agarose gel electrophoresis was performed to detect the expression of histamine receptors, including HRH1, HRH2, HRH3, and HRH4, in the rat dorsal root ganglion (DRG) cell line F11, and the results revealed that the expression of HRH1 was significantly higher than the other three receptors in F11 cells (Figure  3A ). Next, histamine (10 −7  mM) was used to treat F11 cells, and the results revealed that histamine upregulated the expression of the HRH1 mRNA and protein in F11 cells ( p  < 0.01, Figure  3B ; p  < 0.05, Figure  3C ). Consistent with these findings, HRH1 expression in the DRG tissues of EMS mice ( n  = 7) was significantly higher than that in the Sham group ( n  = 8, p  < 0.0001; Figure  3D,E ). Double immunofluorescence staining further revealed the colocalization of HRH1 (shown in red) and the peptidergic neuronal marker CGRP (shown in green; Figure  3F ), confirming that HRH1 was expressed on the peripheral nerves of endometriotic lesions. Histamine sensitized TRPV1 through HRH1 activation in endometriosis. (A) The expression of histamine receptors, including HRH1, HRH2, HRH3, and HRH4, in F11 cells was determined by PCR using agarose gel electrophoresis. (B) Changes in HRH1 gene expression in F11 cells after treatment with 10 −7  mM histamine for 24 h, as measured by RT–qPCR. (C) Changes in HRH1 protein expression in F11 cells after treatment with 10 −7  mM histamine for 24 h, as detected by western blotting. (D) IHC images showing HRH1 expression in DRG tissue from sham ( n  = 8) and EMS mice ( n  = 7); 200×, scale bar = 100 μm. The boxed region in the merged images is enlarged in the images on the right (original magnification 400×, bar = 100 μm). (E) IHC scores of the above two groups. (F) Images of IF staining for HRH1 (red), CGRP (green), and cell nuclei (DAPI, blue) in paraffin sections of endometriotic lesions; 400×, scale bar = 100 μm. (G) Fluo‐4 fluorescence intensity of F11 cells from the control, histamine, and histamine+DLT groups. (H) Increase in the Ca 2+ peak (DF/F0) in F11 cells from the control, histamine, and histamine+DLT groups ( n  = 3 in each group). Control (B, C, G, and H), F11 cells pretreated with the vehicle control; EMS, mice with endometriosis surgery; His, F11 cells pretreated with histamine (10 −7  mM); histamine, F11 cells pretreated with histamine (10 −7  mM) overnight; histamine + DLT, F11 cells pretreated with histamine (10 −7  mM) and desloratadine (1 μM) overnight; Sham, mice with the sham operation. * p  < 0.05, ** p  < 0.01, *** p < 0.001, **** p  < 0.0001, and ns, not significant. Studies have shown that histamine can mediate the sensitization of TRPV1, thereby inducing visceral hypersensitivity and abdominal pain in patients with irritable bowel syndrome [ 21 ]. Hence, we postulated that histamine might mediate TRPV1 sensitization, particularly via interaction with HRH1, in endometriosis. To this end, we tested the effect of the HRH1 antagonist desloratadine (DLT) on TRPV1 sensitization by histamine. The intracellular calcium (Ca 2+ ) response of F11 cells to capsaicin was used to study TRPV1 sensitization. Compared with the control, an overnight incubation with histamine (10 −7  mM) increased the Ca 2+ response to capsaicin (1 μM) ( p  < 0.0001; Figure  3G,H ), confirming that histamine could sensitize TRPV1. Additionally, the ability of histamine to potentiate the Ca 2+ response was reversed by DLT, indicating that TRPV1 sensitization occurred via HRH1 activation ( p  < 0.001; Figure  3H ). Taken together, these data suggested that histamine might sensitize TRPV1 through HRH1 activation in endometriosis. We further evaluated whether HRH1‐mediated TRPV1 sensitization could be replicated in the endometriosis environment. The mice were divided into four groups: the Sham, EMS‐Control, EMS‐DLT low‐dose, and EMS‐DLT high‐dose groups (Figure  4A ). Endometriosis‐induced thermal and mechanical hypersensitivity was tested in the Sham, EMS‐Control, EMS‐DLT low‐dose, and EMS‐DLT high‐dose groups to investigate the efficacy of desloratadine in alleviating hyperalgesia in endometriosis. Compared with those in the Sham group, pain‐related behavioral changes were detected in the EMS‐Control, EMS‐DLT low‐dose, and EMS‐DLT high‐dose groups 1–5 weeks after construction of an endometriosis model. When desloratadine treatment started in the 5th week after surgery, mechanical and thermal hyperalgesia were significantly alleviated at week 7 in the EMS‐DLT high‐dose group compared with the EMS‐Control group (Figure  4B,C ). The HRH1 antagonist desloratadine relieved hyperalgesia in the endometriosis model mice. (A) Schematic of the animal intervention experiment. Sham mice received sham surgery. EMS‐Control and EMS‐DLT mice underwent endometriosis surgery. Drug interventions were started at Week 5 and continued for the next 2 weeks by oral gavage. Behavioral tests were performed every week. The mice were sacrificed at Week 7, and DRG cells were isolated. Sensitivities to mechanical (B) and thermal (C) stimuli in the Sham, EMS‐Control, EMS‐DLT low‐dose, and EMS‐DLT high‐dose groups ( n  = 6 in each group). (D) Gross morphology of endometriotic lesions from EMS‐Control, EMS‐DLT low‐dose, and EMS‐DLT high‐dose mice. (E and F) The total weight and volume of endometriotic lesions in EMS‐Control, EMS‐DLT low‐dose, and EMS‐DLT high‐dose mice were assessed on Day 43 ( n  = 6 in each group). (G) Fluo‐4 fluorescence intensity of isolated DRG cells from the Sham, EMC‐Control, and EMS‐DLT high‐dose groups. (H) Increase in the Ca 2+ peak (DF/F0) in DRG cells isolated from the Sham, EMS‐Control, and EMS‐DLT high‐dose groups ( n  = 3 in each group). EMS‐Control, mice with endometriosis surgery and treated with the vehicle control; EMS‐DLT, mice with endometriosis surgery and treated with desloratadine; EMS‐DLT high‐dose, mice with endometriosis surgery and treated with desloratadine (20 mg/kg); EMS‐DLT low‐dose, mice with endometriosis surgery and treated with desloratadine (10 mg/kg); Sham, mice with the sham operation. *Compared with the Sham group; # compared with the EMS‐Control group. * p  < 0.05, ** p  < 0.01, *** p  < 0.001, **** p  < 0.0001, # p < 0.05 and ## p < 0.01. The reflexive withdrawal tests in response to mechanical and thermal stimulation of the abdominal area were also carried out in the Sham, EMS‐Control, EMS‐DLT low‐dose, and EMS‐DLT high‐dose groups. The results were in line with those of the mechanical and thermal paw withdrawal tests (Figure  S2A,B ). The effect of DLT on endometriotic lesions was also investigated. The results (Figure  4D–F ) revealed that the total volume and weight of the ectopic lesions in the high‐dose EMS‐DLT and low‐dose EMS‐DLT groups were both significantly lower than those in the EMS‐control group. Since the EMS‐DLT high‐dose group relieved hyperalgesia compared with the control group, the EMS‐DLT high‐dose group was selected for the subsequent trials. DRG cells were isolated from the Sham, EMS‐Control, and EMS‐DLT high‐dose groups. The intracellular calcium (Ca 2+ ) response of isolated DRG cells to capsaicin (1 μM) was then evaluated to investigate the role of desloratadine in TRPV1 sensitization. Capsaicin‐induced Ca 2+ influx was significantly potentiated in the EMS group compared with the Sham group ( p  < 0.05; Figure  4G,H ), indicating that TRPV1 was sensitized in endometriosis. However, the Ca 2+ response in the EMS‐DLT high‐dose group was weaker than that in the EMS group ( p  < 0.05; Figure  4H ). The data suggested that the HRH1 antagonist desloratadine could inhibit TRPV1 sensitization in endometriosis. Taken together, these data demonstrated that desloratadine weakened TRPV1 sensitization and alleviated endometriosis pain in a mouse endometriosis model, indicating that the HRH1 antagonist desloratadine could relieve hyperalgesia in endometriosis. An Mrgprb2 (the orthologue of human MRGPRX2) knockout mouse model was generated to further investigate the effect of MRGPRX2 on endometriosis pain in vivo. The strategies for generating the mice and genotyping are shown in Figure  5A . Specifically, the genotypes of the F1 generation were determined in 10‐day‐old mice. The primer pair F1/R1 was designed to specifically bind to the two flanking regions of the sequence targeted for deletion (PCR1) to indicate the knockout genotype, and another primer pair, F2/R2, was designed to bind to the two sides of the sequence and indicate the WT genotype (PCR2). The KO band is 5022 bp in PCR1 and 0 bp in PCR2, whereas the WT band is 707 bp in PCR1 and 483 bp in PCR2. The PCR results revealed the WT, heterozygous, and homozygous KO genotypes (Figure  5B ) via agarose gel electrophoresis. MRGPRX2 served as a therapeutic target for endometriosis‐induced hyperalgesia. (A) Mrgprb2 gene knockout strategy and Genotyping strategy in mice. Strategy of Genotyping: Wild type: ① PCR reaction obtains a single WT band; ② PCR reaction obtains a single WT band. Heterozygote: ① PCR reaction obtains a WT band and a KO band; ② PCR reaction obtains a WT band. Homozygote: ① PCR reaction obtains a single KO band; ② PCR reaction without product. (B) Genotypes, including the WT, heterozygous, and homozygous genotypes, were identified using agarose gel electrophoresis. (C) PMCs from WT and (D) Mrgrpb2 −/− mice were double‐stained with FcεRIα and c‐Kit antibodies and analyzed by FACS. (E) Mrgprb2 protein levels in PMCs from WT and KO mice. (F) Gross morphology of endometriotic lesions from WT and Mrgrpb2 −/− mice. (G and H) The total weight and volume of endometriotic lesions from WT and Mrgrpb2 −/− mice were assessed on Day 28 ( n  = 6 in each group). (I) Images of IHC staining for HRH1 in DRG tissues from WT EMS ( n  = 7) and Mrgrpb2 −/− EMS ( n  = 8), 200×, bar = 100 μm. The boxed regions in the merged images are enlarged in the image on the right (original magnification 400×, bar = 100 μm). (J) IHC scores for each group. (K) Histamine levels in peritoneal fluid from WT Sham ( n  = 5), WT EMS ( n  = 5), and Mrgrpb2 −/− EMS ( n  = 5) mice. (L) Toluidine blue‐stained section of uterus from WT Sham, lesions from WT EMS, and Mrgrpb2 −/− EMS. (Arrow: degranulated mast cells). (M) Number of mast cells per field of microscopic vision in the WT Sham ( n  = 6), WT EMS ( n  = 6), and Mrgrpb2 −/− EMS ( n  = 6) mice. (N) Number of degranulated mast cells per field of microscopic vision in the WT Sham ( n  = 6), WT EMS ( n  = 6), and Mrgrpb2 −/− EMS ( n  = 6) mice. (O) Mechanical threshold and (P) thermal latency in the WT sham, WT EMS, and Mrgrpb2 −/− EMS groups ( n  = 6 in each group). KO, Mrgrpb2 −/− mice; Mrgrpb2 −/− , Mrgrpb2 knockout mice; Mrgrpb2 −/− EMS, Mrgrpb2 −/− mice with endometriosis surgery; PMCs, mouse peritoneal mast cells; WT, wild‐type mice; WT EMS, wild‐type mice with endometriosis surgery; WT Sham, wild‐type mice with a sham operation. *Compared with the WT Sham group; # compared with the WT EMS group. * p  < 0.05, ** p  < 0.01, *** p  < 0.001, **** p  < 0.0001, # p < 0.05, ## p < 0.01 and #### p < 0.001. We isolated peritoneal mast cells (PMCs) from KO and WT mice to detect gene expression and confirm the efficiency of gene knockout in mast cells from the mouse model. The mast/stem cell growth factor receptor (also known as CD117 or c‐Kit) and FcεRIα were used for identity verification. The flow cytometry results revealed that 97.2% of WT PMCs and 97.9% of Mrgprb2 −/− PMCs were positive for both markers (Figure  5C,D ), confirming the purity of the isolated cells and the lack of difference in the generation of mast cells between the two models. The western blot results confirmed that the Mrgprb2 protein level in the PMCs of the KO mice was significantly lower than that in the PMCs of the WT mice ( p  < 0.01; Figure  5E ), further confirming the successful generation of the Mrgprb2 knockout mouse model. An endometriosis mouse model was established in WT and Mrgprb2 −/− mice, which were then sacrificed at 4 weeks after surgery. In the Mrgprb2 −/− group, both donor and recipient animals were knockouts. The total volume and weight of the ectopic lesions in the Mrgprb2 −/− group were both significantly lower than those in the WT group ( p  < 0.001 and p  < 0.05, respectively; Figure  5F–H ). We evaluated whether Mrgprb2 deficiency affected the expression of HRH1 by performing IHC on DRG tissues from the WT EMS and Mrgprb2 −/− EMS groups. The results revealed that HRH1 levels were decreased in DRG tissues from the Mrgprb2 −/− EMS group compared with the WT EMS group ( p  < 0.0001; Figure  5I,J ), indicating that the endometriosis‐induced upregulation of HRH1 levels in the DRG was reversed by Mrgprb2 deficiency. Furthermore, the concentration of histamine in the peritoneal fluid was detected using ELISA, and the level was significantly higher in the WT EMS group than in the WT Sham group ( p  < 0.0001; Figure  5K ). However, the histamine level was lower in the Mrgprb2 −/− EMS group than in the WT EMS group ( p  < 0.05; Figure  5K ). Toluidine blue staining demonstrated that the density of mast cells was significantly greater in lesions from WT EMS mice than in uterus from WT Sham mice and in lesions from Mrgrpb2−/− EMS mice (Figure  5L,M ). Additionally, the number of degranulated mast cells in WT EMS mice was significantly greater than in WT Sham mice and Mrgrpb2−/− EMS mice (Figure  5L,N ). Behavioral tests were performed in the WT sham, WT EMS, and Mrgprb2 −/− EMS groups to confirm whether Mrgprb2 played a vital role in endometriosis pain. Compared with those in the WT sham group, the thresholds of both mechanical and thermal hyperalgesia were significantly lower in the WT EMS group, whereas the thresholds in the Mrgprb2 −/− EMS group were elevated (Figure  5O,P ), indicating that Mrgprb2 deficiency could alleviate sensitivity to mechanical and thermal stimulation. The reflexive withdrawal tests in response to mechanical and thermal stimulation of the abdominal region were conducted in the WT sham, WT EMS, and Mrgrpb2−/− EMS groups. The findings were consistent with those observed in the mechanical and thermal paw withdrawal tests (Figure  S3A,B ). Collectively, these data suggested that Mrgprb2 could serve as a therapeutic target for endometriosis‐induced hyperalgesia. The effects of the Mrgprb2 antagonist QWF [ 33 ] on endometriosis‐induced thermal and mechanical hypersensitivity were tested to investigate whether a pharmacological antagonist of Mrgprb2 affects endometriosis pain. In these experiments, wild‐type mice were divided into four groups: the Sham, EMS‐Control, EMS‐QWF low‐dose, and EMS‐QWF high‐dose groups. The thermal and mechanical sensitivities (Figure  6A,B ) of the mice were tested at baseline (Week 0) and once a week following endometriosis surgery. EMS‐Control mice served as controls. The results revealed that pain‐related behavioral changes occurred in the EMS‐Control, EMS‐QWF low‐dose, and EMS‐QWF high‐dose groups 1–5 weeks after surgery compared with the Sham group. The animals received an intraperitoneal injection of QWF or vehicle beginning in the 7th week after surgery. Compared with those in the EMS‐Control group, mechanical and thermal hyperalgesia in the EMS‐QWF high‐dose group were significantly alleviated at Week 7. A pharmacological antagonist of Mrgprb2 alleviated hyperalgesia in the endometriosis model mice. Sensitivities to mechanical (A) and thermal (B) stimuli in the Sham, EMS‐Control, EMS‐QWF low‐dose, and EMS‐QWF high‐dose groups ( n  = 6 in each group). (C) Gross morphology of endometriotic lesions from mice in the EMS‐Control, EMS‐QWF low‐dose, and EMS‐QWF high‐dose groups. (D to E) The total weight and volume of endometriotic lesions in EMS‐Control, EMS‐QWF low‐dose, and EMS‐QWF high‐dose mice were assessed on Day 43 ( n  = 6 in each group). EMS‐Control, mice that underwent endometriosis surgery and received the vehicle control; EMS‐QWF high dose, mice that underwent endometriosis surgery and received QWF (20 mg/kg); EMS‐QWF low dose, mice that underwent endometriosis surgery and received QWF (10 mg/kg); QWF, antagonist of Mrgprb2. *Compared with the WT Sham group; # compared with the WT EMS group. * p  < 0.05, ** p  < 0.01, *** p  < 0.001, **** p  < 0.0001, # p < 0.05, ## p < 0.01, and #### p < 0.0001. The reflexive withdrawal tests in response to mechanical and thermal stimulation of the abdominal region were performed in the Sham, EMS‐Control, EMS‐QWF low‐dose, and EMS‐QWF high‐dose groups. The results were consistent with those observed in the mechanical and thermal paw withdrawal tests (Figure  S4A,B ). In addition, the total volume and weight of the ectopic lesions in the EMS‐QWF high dose and EMS‐QWF low dose groups were significantly lower than those in the EMS‐Control group (Figure  6C–E ). Taken together, these findings indicated that Mrgprb2 antagonism alleviated the hyperalgesia in endometriosis.

In the present study, our findings delineated a new signaling pathway in which the mast cell receptor MRGPRX2 is critically involved in endometriosis pain by mediating histamine production from mast cells, and the potential mechanism of histamine in endometriosis‐induced hyperesthesia was explored. Future clinical studies are needed to elucidate the potential of MRGPRX2 as a strong biomarker or therapeutic target for patients suffering from endometriosis pain. Here, we showed that the ablation of mast cells alleviated endometriosis‐induced hyperalgesia in mast cell‐deficient mice, further suggesting that mast cells contribute to endometriosis pain, as reported in our previous studies [ 7 , 13 ]. Recently, the mast cell receptor MRGPRX2 has received increasing attention because several antagonists of MRGPRX2 have been developed for the potential treatment of a variety of mast cell‐mediated diseases, such as EP262, which was developed by Escient Pharmaceuticals. MRGPRX2 and its murine orthologue Mrgprb2 have been detected in the skin, colon, and lungs [ 15 , 19 , 34 ], which contain mast cells. Notably, MRGPRX2 is upregulated in inflammatory disorders. Chen et al. reported that MRGPRX2 accumulates in the colonic mast cells of ulcerative colitis patients [ 15 ]. Jia revealed that the atopic dermatitis severity and type 2 cytokine release are associated with MRGPRX2 expression levels [ 35 ]. Here, we further explored the relationship between MRGPRX2 and endometriosis pain based on our previous research [ 36 ]. By performing immunofluorescence staining, we observed that MRGPRX2 expression was significantly elevated in endometriosis tissues, revealing a potential role for MRGPRX2 in endometriosis. Recent evidence has indicated that MRGPRX2 functions as a mediator of the interaction between mast cells and nociceptors, thus regulating pain and itch [ 37 , 38 , 39 ]. Green et al. reported that MRGPRX2, a crucial driver of neuropeptide SP‐induced immune cell recruitment and the release of multiple proinflammatory cytokines from mast cells, led to incision‐induced inflammatory pain [ 18 ]. In addition, Sbei et al. showed that MRGPRX2 was potently activated by PACAP and resulted in migraine‐like pain [ 16 ]. Here, we found that the density of MRGPRX2‐positive mast cells in endometriosis lesions from EMS patients with pain was greater than that in lesions from EMS patients without pain, indicating that MRGPRX2 might be involved in endometriosis pain. Increasing evidence suggests that cationic antimicrobial peptides, including β‐defensins and LL‐37, can activate mast cells through MRGPRX2 and could be endogenous ligands for MRGPRX2 [ 40 , 41 ]. HBD‐2, which is a β‐defensin, can activate mast cells and elicit histamine release by binding to MRGPRX2 [ 42 , 43 ]. HBD‐2 was found to be involved in inflammatory disorders and pain. Wijesundara et al. reported that HBD‐2 is an inflammatory mediator produced by tonsil cells and mediates an inflammatory response in pharyngitis [ 44 ]. Additionally, Shulman et al. showed that increased HBD‐2 concentrations in children with IBS appear to be related to symptoms of abdominal pain [ 45 ]. Here, we reported that HBD‐2 accumulated in endometriotic lesions, which is consistent with the findings of previous studies [ 46 ]. Our study revealed that this secretory protein was present mostly in endometriotic epithelial cells of endometriotic lesions and could be secreted from endometriotic cells, indicating that HBD‐2 was produced mainly by endometriotic epithelial cells in the endometriosis environment. Furthermore, we generated an MRGPRX2 KO HMC1.1 cell line and discovered that MRGPRX2 could induce histamine production from human mast cells after HBD‐2 binding. However, James et al. reported that histamine production from mast cells was mediated mainly by FcεRI and that Mrgprb2 rarely evoked this response [ 45 ]. Recently, evidence has suggested that activated MRGPRX2 alone can elicit histamine release [ 19 ]. Our data provide evidence that histamine levels in peritoneal fluid are elevated in an endometriosis model and that Mrgprb2 deficiency significantly reduces the concentration of histamine in endometriosis model mice, indicating that the release of histamine from mast cells is mediated by MRGPRX2 activation in endometriosis. TRPV1, expressed on a subclass of nociceptors, has been widely studied for its involvement in pain sensation. The process of sensitization (increased activity) of nociceptors is called hyperalgesia [ 47 ]. Sensitization of the TRPV1 channel contributes to the development of visceral pain and arthritis pain [ 48 , 49 ]. TRPV1 sensitization‐induced endometriosis pain has previously been suggested [ 50 ], but the possible mechanism of TRPV1 sensitization in endometriosis is still unclear. Previous studies have shown that histamine can sensitize TRPV1 channels, thus leading to increased visceral pain perception [ 21 , 51 ]. Our data revealed that incubation with histamine significantly increased the sensitivity of DRG neurons to capsaicin, a specific agonist of the transient receptor potential channel TRPV1, confirming that the TRPV1 channel can be sensitized by histamine. We previously reported that ketotifen, which is not only a mast cell stabilizer but also an HRH1 antagonist, reduces hyperalgesia in a rodent model of surgically induced endometriosis [ 31 ]. Here, we showed that the histamine receptor HRH1 was expressed in the sensory neurons around endometriotic lesions. Thus, the speculation that HRH1 is involved in endometriosis pain is reasonable. Aguilera‐Lizarraga et al. reported that the HRH1 antagonist pyrilamine could prevent TRPV1 sensitization in DRG neurons treated with the colonic supernatant of mice with visceral hypersensitivity (VHS) and reduce the pain response to colorectal distention in VHS mice [ 48 ], suggesting an effect of this HRH1 antagonist on TRPV1 desensitization. Desloratadine (DLT) is a second‐generation HRH1 antagonist that is a highly efficient HRH1 inhibitor that blocks chronic urticaria, breast cancer, and bladder cancer [ 52 , 53 , 54 ]. Because DLT does not penetrate the blood–brain barrier, it does not impair cognitive or psychomotor performance and is also nonsedating [ 55 ]. Here, we showed that DLT prevented the increased TRPV1‐mediated response in DRG neurons after treatment with histamine. We used DRG neurons isolated from mice for further study to investigate this phenomenon in a mouse endometriosis model. Our findings confirmed that DLT could desensitize the TRPV1 channel in DRG neurons from EMS mice, demonstrating that HRH1 is involved in the sensitization of the TRPV1 channel in endometriosis. Furthermore, DLT alleviated hyperalgesia in endometriosis model mice, indicating that DLT may be a novel modality for the treatment of endometriosis‐related pain. In this study, we generated Mrgprb2 −/− mice via CRISPR/Cas9 technology to further investigate the effect of Mrgprb2 on endometriosis pain in vivo. Our data showed that genetic ablation or pharmacological antagonism of Mrgprb2 is effective in treating the development of endometriosis pain in mice, accompanied by a significant reduction in lesion size. However, the loss of Mrgprb2 significantly attenuated but did not completely abolish hyperalgesia compared with that in the WT endometriosis model mice. Similar to the findings of studies with Mrgprb2 −/− mice, the ablation of mast cells significantly attenuated but did not abolish endometriosis‐induced mechanical and thermal hypersensitivity in mice. These findings potentially indicate that other effectors contribute to neuroinflammation in endometriosis, including macrophages, T lymphocytes, B lymphocytes, neutrophils, and NK cells [ 56 , 57 , 58 , 59 ]. One of the limitations of our study is that the functions of human MRGPRX2 in endometriosis‐induced pain behaviors were based on mouse orthologues and animal experiments, which might lead to some biases because MRGPRX2 displays only 53% sequence identity with Mrgprb2 [ 60 ]. A humanized MRGPRX2 knock‐in model, therefore, needs to be generated to investigate the role of MRGPRX2 in endometriosis pain in future research. Another limitation is that the source of the MRGPRX2 and Mrgprb2 band was still present in the knockout cells and animals, which might be attributed to the antibody recognition site. Specifically, the target epitope of the MRGPRX2 antibody (Thermo Fisher) we utilized corresponds to the amino acid sequence 286–330. However, the knocked gene regions might not fully cover the epitope regions recognized by the antibody. Unfortunately, within the domain of commercial antibodies, it remains highly challenging to identify one that fully satisfies the specified requirements. This may explain why, even if truncated proteins are produced, Western blotting (WB) still detects signals, thereby failing to accurately reflect the complete loss of the protein. To address this issue, it will be necessary to develop a customized antibody for the future. In addition, the molecular mechanism underlying the functional interaction between HRH1 and TRPV1 remains elusive. Gao et al. reported that HRH1 could directly bind to the carboxy‐terminal region of TRPV1 at residues 715–725 and 736–749 to mediate histaminergic itch [ 61 ]. This result is not consistent with the study by Shim et al., who reported that HRH1 activation leads to the accumulation of phospholipase A2 and 12‐lipoxygenase, thus activating TRPV1 [ 62 ]. Therefore, additional mechanisms need to be explored to better understand how the HRH1‐TRPV1 interaction improves the TRPV1 response to histamine in endometriosis pain. Although some MRGPRX2 antagonists have been designed and tested in animal experiments and clinical trials [ 63 , 64 ], they have not been widely used in the clinic. Additionally, the efficacy and safety of MRGPRX2 antagonists for the treatment of endometriosis pain need to be investigated in clinical trials. Furthermore, more MRGPRX2 antagonists will be developed for the modulation of MRGPRX2 in endometriosis pain.

Our findings ultimately support a model (Figure  7 ) in which endometriotic cell‐derived HBD‐2 mediates histamine secretion from mast cells by binding to MRGPRX2. Secreted histamine promotes TRPV1 sensitization via HRH1 activation in pain sensory neurons, thereby aggravating hyperalgesia in endometriosis. Mrgprb2 deficiency effectively alleviates hyperalgesia and reduces the size of endometriotic lesions. In conclusion, MRGPRX2 mediates mast cell‐induced hyperalgesia and can serve as a novel therapeutic target for endometriosis pain. Schematic summary of the mechanisms of MRGPRX2‐mediated endometriosis pain. Endometriotic cell‐derived HBD‐2 mediated the secretion of histamine from mast cells by binding to MRGPRX2. Histamine sensitized TRPV1 via the activation of HRH1 in pain sensory neurons, thus aggravating hyperalgesia in an endometriosis model.

Endometriosis (EMS) is a common chronic inflammatory gynecological disorder characterized by pain and infertility. The pain symptoms associated with endometriosis include dysmenorrhea, dyspareunia, dysuria, and chronic pelvic pain [ 1 , 2 ], affecting up to 73% of endometriosis patients [ 3 ]. However, currently, no definitive cure is available for endometriosis pain because the mechanism of endometriosis pain is poorly understood. Increasing evidence suggests that endometriosis pain is a type of inflammatory and neuropathic pain [ 4 ]. Large numbers of macrophages, mastocytes, and neutrophils are recruited to endometriotic lesions [ 5 ]. Inflammatory factors secreted by immune cells activate the abnormally distributed nerve fibers in lesions [ 6 , 7 ], triggering nerve sensitivity and leading to hyperalgesia. Mast cells (MCs) are multifunctional immune cells that are distributed throughout the body [ 8 ]. Mast cells play key roles in inflammatory diseases, tumors, and injury repair [ 9 , 10 , 11 ]. Our previous studies revealed that mast cells degranulate and secrete inflammatory mediators or cytokines around lesions, contributing to the development of endometriosis [ 12 , 13 ]. Moreover, mast cells located in close proximity to nerve fibers promote nerve growth in endometriotic lesions [ 7 ]. However, studies on how MCs mediate endometriosis pain are still limited. Mas‐related G protein‐coupled receptor‐X2 (MRGPRX2) is a receptor expressed on mast cells [ 14 ]. MRGPRX2 is involved in various conditions, including inflammation, itch, and pain [ 15 , 16 , 17 ]. In mice, the MRGPRX2 orthologue is called Mrgprb2, and Mrgprb2 has been shown to contribute to neuroimmune interactions in a mouse injury model [ 18 ]. MRGPRX2 activation can trigger mast cells to release various mediators, including histamine [ 19 ]. Histamine can act on cells that express one or more of the four known histamine G protein‐coupled receptors (HRH1‐4) [ 20 ]. Histamine and HRH1 have been found to play crucial roles in nerve sensitization and thus lead to visceral hypersensitivity in patients with irritable bowel syndrome [ 21 ]. Previous studies have shown that antihistamine drugs could potentially be used for endometriosis management [ 22 , 23 ]. However, whether and how pain is mediated by the MRGPRX2‐induced release of histamine in mast cells of endometriotic lesions remains unclear. Here, we demonstrated that MRGPRX2 could mediate mast cells to induce pain sensitization in endometriosis. Following elevated expression levels of MRGPRX2 and its ligand, beta‐defensin‐2 (HBD‐2), in endometriotic lesions, we found that HBD‐2 mediated histamine secretion from mast cells via MRGPRX2. We further characterized the roles of histamine and HRH1 in pain sensitization by conducting in vitro and in vivo experiments. A mouse endometriosis model was established via Mrgprb2 knockout, and subsequent behavioral tests revealed that MRGPRX2 could serve as a potential target to alleviate endometriosis pain.

The authors declare no conflicts of interest.

This study was conducted according to the ethical policies and procedures approved by the Human Ethics Committee of Women's Hospital, School of Medicine, Zhejiang University (No. 20230091). All patients signed an informed consent form to participate in this study, and they were able to opt out of the study for the whole duration. In accordance with the surgical findings, the patients were divided into two groups, namely, the endometriosis ( n  = 19) and the control ( n  = 21) groups. The severity of pain was assessed using a visual analogue scale (VAS). Self‐reported endometriosis‐associated pain was measured by the VAS on a scale ranging from 0 to 10. A score of 1–3 was considered mild, a score of 4–6 was considered moderate, and a score > 6 was considered severe pain. None of the patients received sex hormone therapy 6 months before surgery. Patients who suffered from other gynecological or autoimmune diseases were also excluded. The patients' characteristics are listed in Table  S1 . Mast cell‐deficient W‐sash c‐kit mutant Kit W‐sh/W‐sh (Kit) mice were purchased from The Jackson Laboratory. Homozygous (Mrgprb2 −/− ) mice were constructed (GemPharmatech Co. Ltd., Nanjing, Jiangsu, China) to investigate the effects of Mrgprb2 deficiency. Six‐ to eight‐week‐old female C57BL/6J wild‐type control mice were purchased from Shanghai SLAC Laboratory Animal Co. Ltd. (Shanghai, China). All the mice were housed under identical conditions, with a maximum of 5 animals per cage, on a 12‐hour light–dark cycle. Food and water were provided ad libitum. All the animal experiments were performed in accordance with the Guide for the Care and Use of Laboratory Animals published by the United States National Institutes of Health (NIH Publication, revised 2011), and the protocols were approved by the Ethics Committee of Zhejiang University (No. ZJU20230165). The human mast cell line HMC1.1 was obtained from Merck Millipore (Billerica, MA, USA) and grown in Iscove's medium (Gibco, Grand Island, NY, USA) supplemented with 10% FBS (Gibco), 1× penicillin/streptomycin (Gibco) and 1.2 mM thioglycerol (Sigma–Aldrich, St. Louis, MO, USA). The DRG cell line (F11) was obtained from the European Collection of Authenticated Cell Cultures (Porton Down, Salisbury, UK). F11 cells are a somatic cell hybrid of a rat embryonic DRG and the mouse neuroblastoma cell line N18TG2. The line retains both rat and mouse chromosomes and synthesizes both rat and mouse isoenzymes. They were maintained in DMEM/F12 (Gibco) supplemented with 10% FBS. Dorsal root ganglion neurons from adult (postnatal weeks 12–14) mice were isolated and cultured as described elsewhere [ 24 ]. The extraction and culture of peritoneal mast cells from the mice were performed as described previously [ 25 ]. Primary endometrial cells were isolated and cultured as described in our previous study [ 26 ]. In brief, endometrial tissues and cyst wall specimens were collected from patients with or without endometriosis during laparoscopy and hysteroscopy. Following triple PBS rinsing, the biological samples were sectioned into 1‐mm fragments and enzymatically processed using 0.1% collagenase type I (Thermo Fisher Scientific) for 70–80 min at 37°C. Cells were subsequently isolated through 5‐min centrifugation cycles and reconstituted in DMEM/F‐12 basal medium (Gibco) without additives. Sequential filtration was performed through progressively smaller nylon filters (100 μm followed by 40 μm) to differentially isolate epithelial and stromal cell populations. Both cell types were maintained in growth medium consisting of DMEM/F‐12 enriched with 12% FBS. To ensure cellular purity, the initial culture medium was refreshed after 12 h to eliminate non‐adherent cellular components. All these cells were maintained at 37°C with 5% CO 2 , and the medium was changed every 2–3 days. Histamine was purchased from MedChem Express (MCE, Monmouth Junction, NJ, USA) and was used to treat cells at a concentration of 10 −7  mM. The guide RNA sequences for CRISPR/Cas9 were designed using the online design tool CRISPOR ( http://crispor.tefor.net/ ). The common CDS region of all transcripts of MRGPRX2 (Gene ID: 117194) was selected, and exon 2, where the common CDS region is located, was identified for target site design. The oligonucleotides used are listed in Table  S2 . HMC‐1.1 cell transfection was performed as described in our previous study [ 7 ], and the cells were analyzed by sequencing. The sequences of primers used for identification are listed in Table  S3 . Direct PCR Universal Master Mix (Thermo Fisher Scientific, Waltham, MA, USA) was used to lyse the toes of 7–10‐day‐old mice, followed by PCR. The PCR products were resolved by electrophoresis on 2% agarose gels and then visualized using a UV lamp in Fusion FX7 (Vilber Lournar, Paris, France). FCM analysis was applied to validate peritoneal mast cells. The isolated cells were incubated with APC‐conjugated mouse CD117/KIT and FITC‐conjugated mouse FcεRIα. The antibodies used for flow cytometry are listed in Table  S6 . Next, the cells were washed with staining buffer twice and resuspended for FCM analysis with a BD FACSVerse flow cytometer (BD Biosciences, San Jose, CA, USA). Endometriosis was induced as previously described [ 13 ]. Briefly, bilateral ovariectomy was performed seven days before the operation, followed by subcutaneous injection of 0.5 mg of oestradiol benzoate dissolved in 60 mL of corn oil every 5 days. On Day 0, the uterine tissue of each donor was dissected and cut into fragments less than 1 mm 3 in size. These fragments were then suspended in 0.9% sterile saline and injected intraperitoneally into two other recipient mice. The Sham operation group received an equivalent volume of 0.9% sterile saline. WT mice were separated into four groups to investigate the role of HRH1 in endometriosis‐induced hyperalgesia: the Sham group, drug administration groups (low‐dose and high‐dose groups), and control group. The mice in the Sham and control groups were injected with the vehicle control containing 10% DMSO, 40% PEG400, 5% Tween‐80 and 45% saline, whereas those in the drug administration groups were administered the proven HRH1 antagonist desloratadine (DLT, MCE) (i.g.) at a dose of 10 or 20 mg/kg once a day [ 27 , 28 ]. WT mice were separated into four groups to investigate the role of Mrgprb2 in endometriosis‐induced hyperalgesia: the Sham group, low‐dose and high‐dose drug groups, and the control group. The mice in the Sham and control groups were injected intraperitoneally with the vehicle control containing 10% DMSO, 40% PEG400, 5% Tween‐80 and 45% saline, while those in the drug administration group were injected with the Mrgprb2 antagonist QWF (Tocris Bioscience) (i.p.) at a dose of 10 mg/kg or 20 mg/kg once a day. The mice were anesthetized via the inhalation of CO2 and euthanized via cervical dislocation on Days 28 or 49 after the induction of endometriosis. The peritoneal fluid and DRG (T11‐L2 segment) were preserved for further experiments. Behavioral tests included mechanical and thermal paw withdrawal tests and reflexive withdrawal tests to mechanical and thermal stimulation of the abdominal area. Behavioral tests were performed to assess hyperalgesia in the mice before and every week after surgery, as described previously [ 29 , 30 ]. The tests were performed by personnel who were blinded to the genotypes or drugs injected. Each test was repeated three times. For Mechanical and thermal paw withdrawal tests, each mouse was placed in a chamber with a wire mesh floor and allowed to acclimate for at least 30 min before testing began. Mechanical allodynia was evaluated using the von Frey test, which was performed using an electronic device (Model 2390, IITC/Life Science Instruments, Lowell, CA, USA) as previously described. The right hind paw was poked three times with a flexible probe, and the paw withdrawal threshold was determined by averaging these three measurements. The apparatus automatically recorded the numerical value of the force needed to cause paw withdrawal, which was defined as the mechanical pain threshold (MPT). An apparatus (Model 33B, IITC/Life Science Instruments, Lowell, CA, USA) was used to measure the paw withdrawal latency in response to noxious thermal stimuli. The mice were placed in a Plexiglas chamber on a glass plate containing a light box. Radiant thermal stimulation was applied by passing a light beam through a hole in the light box aimed at the heel of the right hind paw through the glass sheet. The beam was turned off when the mouse lifted its foot. This test was repeated three times at 5‐minute intervals at room temperature (20°C). The average value from the three measurements was obtained. To assess nociceptive responses in the abdominal region, mice were individually placed in a chamber with a wire mesh floor for a minimum 30‐minute acclimatization period prior to experimental procedures. Quantitative evaluation of mechanical hypersensitivity was conducted using the von Frey test, which was performed using an electronic device (Model 2390, IITC/Life Science Instruments, Lowell, CA, USA). Mechanical hypersensitivity in the abdominal region was assessed by quantifying the frequency of withdrawal responses (including abdominal withdrawal from von Frey filament stimulation, subsequent licking of the abdominal area, or whole‐body withdrawal) evoked by normally innocuous or subthreshold mechanical stimuli. The abdominal region was stimulated three times with a flexible probe, and the reflexive withdrawal threshold was calculated as the mean of these three measurements. The apparatus automatically documented the precise force value required to elicit a reflexive withdrawal response. Thermal hyperalgesia was assessed by measuring the latency of withdrawal responses to noxious thermal stimuli using a thermal and pain sensitivity tester (Ugo Basile, Varese, Italy). Mice were placed in Plexiglas chambers on an elevated glass platform, through which a high‐intensity light source was directed. A radiant heat stimulus was applied by focusing a beam of light through a 1 cm × 1 cm aperture in the light box onto the abdominal region. The timer and light beam were automatically terminated upon the animal's withdrawal response, enabling the measurement of the time interval between the onset of the stimulus and the withdrawal event. The abdominal skin of the animals remained in close contact with the glass surface (separation distance ≤ 1 mm). These studies also demonstrated that the rate of temperature increase within this proximity did not vary significantly. Each trial was separated by a 45‐minute interval, and three trials were averaged for analysis. A withdrawal response to radiant heat stimulation of the abdomen was defined as an abdominal retraction (either contraction of the abdominal musculature or elevation of the abdomen via postural adjustment), accompanied by head turning toward the stimulus and licking of the abdominal area. Total RNA was extracted from cells using a Total RNA Isolation Kit (ES Science, Shanghai, China), and 1000 ng of total RNA was used for subsequent reverse transcription. Reverse Transcription used the PrimeScript RT reagent kit (Takara, Tokyo, Japan) according to the manufacturer's instructions. The reaction conditions involved 37°C for 2 min, 85°C for 5 s, and 4°C maintenance. Specific primers were synthesized by Generay (Shanghai, China). The sequences of the primers used are listed in Table  S4 . Real‐time PCR was performed in an Applied Biosystems 7900HT system (Applied Biosystems, USA) with the SYBR Premix Ex Taq TM kit (Takara). The thermal cycling program for qRT–PCR typically included the following steps: 95°C for 30 s of initial denaturation, followed by 40 amplification cycles (95°C for 5 s, 60°C for 30 s), and 4°C maintenance. Next, the 2 −△△Ct method was used to determine the fold change. A total of 1000 ng RNA was used as the template for reverse transcription. Total RNA was reverse transcribed using the PrimeScript RT reagent kit (Takara), and the reaction conditions involved 37°C for 2 min, 85°C for 5 s, and 4°C maintenance. PCR amplification used Green Taq PCR Master Mix (Thermo Fisher Scientific) and thermal cycling conditions included 95°C for 2 min of initial denaturation, followed by 35 amplification cycles (95°C for 30 s, 55°C for 30 s, 72°C for 1 min), and 72°C for 10 min. The specific primers used for amplification were synthesized by Generay, and the sequences of the primers are listed in Table  S5 . Agarose gels were prepared with 0.5× TBE containing 2.0% agarose, and the mixture was melted by heating in a microwave. Next, the gel was exposed to UV light to capture an image of the DNA fluorescent bands with Fusion FX7 (Vilber Lournar). The samples were homogenized in RIPA buffer (Beyotime, Shanghai, China) supplemented with protease and phosphatase inhibitors (Beyotime). 20–30 μg protein was loaded per lane for Western blot analysis. Proteins were separated by SDS‐PAGE, transferred to PVDF membrane, and blocked with 5% non‐fat dry milk in TBST. The membranes were incubated with antibodies against MRGPRX2 (Invitrogen, Carlsbad, CA, USA; 1:1000), beta‐tubulin (Abcam, Cambridge, MA, USA; 1:500), HRH1 (Invitrogen; 1:1000) and GAPDH (Proteintech, Wuhan, China; 1:50000). Table  S6 lists the antibodies used for western blotting. Blocking buffers for western blot were used. Following incubation with an HRP‐conjugated antibody against rabbit IgG (Abcam; 1:5000) or mouse IgG (Abcam; 1:5000), the blots were not stripped and reprobed. The membranes were imaged on a Fusion FX7 (Vilber Lournar) using EZ‐ECL (Biological Industries, Israel). The detected protein levels were quantified using ImageJ and normalized to the expression of the loading controls, such as beta‐tubulin or GAPDH. HBD‐2 and histamine levels were quantified using a Human BD‐2/beta‐Defensin‐2 ELISA Kit (Arigo Biolaboratories, Hsinchu, Taiwan, China) and Histamine ELISA Kit (Abcam, Cambridge, MA, USA) in accordance with the manufacturers' recommendations. To identify mast cells in each specimen, paraffin sections were stained using a standardized method with toluidine blue (Konno et al., 2003). Given that the granules within mast cells contain heparin and sulfated glucosaminoglycans, they exhibit metachromatic staining with toluidine blue. Sections were stained for 30 min in a solution containing 0.05% toluidine blue dissolved in citric acid phosphate buffer (pH 2.51) and subsequently examined under light microscopy. On toluidine blue‐stained sections, degranulated and intact mast cells were observed under light microscopy. Mast cells with densely packed cytoplasmic granules were classified as intact, whereas those exhibiting both typical granule release and scattered granules were categorized as degranulated. In each sample, the number of cells per field (x20 objective, x10 ocular) was counted in five randomly selected fields. IHC staining and analysis of HBD‐2 and HRH1 were performed as previously described [ 13 ]. Briefly, the tissues were fixed with 4% paraformaldehyde, embedded in paraffin, and cut into 3–5 μm sections. The sections were then stained for 60 min at room temperature with a diluted primary antibody against HBD‐2 or HRH1. The antibodies used for IHC are listed in Table  S6 . The sections were then incubated with the corresponding secondary antibody for 60 min. The results were visualized and examined by two blinded observers. The scores for HBD‐2 or HRH1 expression were defined as the sum of the percentage score and intensity score. The average IHC score of five different fields was considered the protein expression level of the gene. Sections of endometriotic lesions and endometrial tissues were permeabilized with 0.1% Triton X‐100 for 10 min and blocked with 8% BSA at room temperature for 1 hour. F11 cells were washed with PBS, fixed with 4% formaldehyde, permeabilized with 0.1% Triton X‐100, and blocked with 5% BSA. The tissues or cells were then incubated with antibodies against tryptase (Abcam; 1:200), MRGPRX2 (Invitrogen; 1:200), CGRP (Abcam; 1:100), and HRH1 (Invitrogen; 1:200) overnight at 4°C. The antibodies used for IF are listed in Table  S6 . The next day, the sections and cells were incubated with the corresponding fluorescent dye‐conjugated secondary antibody (1:200 of Goat Anti‐Rabbit IgG H&L (Alexa Fluor 647; Abcam) or 1:200 of Goat Anti‐Mouse IgG H&L (Alexa Fluor 488; Abcam)) at room temperature for 1 h. Following staining with DAPI (Abcam), the tissues and cells were observed under a confocal microscope (Olympus, Tokyo, Japan). F11 cells or isolated DRG neurons were loaded with Fluo‐4 AM (Invitrogen, Carlsbad, CA, USA) for 20 min at 37°C, washed three times with HBSS (Gibco), and incubated for an additional 30 min. An Olympus FV1200 confocal microscope (Olympus) was used to record calcium influx images, and during the process, capsaicin (1 μM, MCE) was added. The fluorescence intensity in manually drawn regions of interest was quantified using FV10‐ASW Viewer software and plotted against time. The absolute calcium concentration was calculated from the fluorescence DF/F0, where F0 is the baseline fluorescence and DF is the peak amplitude poststimulus. Statistical analyses were performed using GraphPad Prism 9.0 for Windows (Version X; La Jolla, CA, USA, www.graphpad.com ). All experiments were performed at least three times, and all the results are presented as the means ± standard errors of the means (SEMs). Two‐tailed Student's t tests were performed for comparisons between two groups. For comparisons of multiple groups, one‐way ANOVA was used for normally distributed variables, and the Kruskal–Wallis test with Dunn's multiple comparison test was used for variables with skewed distributions. *P values < 0.05 were considered statistically significant.

Figure S1. Table S1. Data S1.