Our study was approved by the Human Research Ethics Committee of Henan Provincial People’s Hospital (Zheng-zhou, China). Sixteen healthy females aged 32.46±5.96 years without prior medical interventions and 43 endometriosis patients aged 34.87±6.73 years were enrolled and the informed written consent was obtained. Exclusion criteria included females undergoing hormone treatment, those with abnormal uterine bleeding, submucous myoma, endometrial polyps, benign and malignant tumors, autoimmune diseases, diabetes, hypertension, and other serious systemic diseases. The ectopic and eutopic endometrium tissues were preserved in a mixture composed of 50% Dulbecco’s modified Eagle’s medium and 50% Ham’s F-12 medium (DMEM/F-12; HyClone) or fixed in 4% paraformaldehyde. A portion of the tissues was transported to the lab in DMEM/F-12, washed in phosphate buffer saline (PBS; HyClone), and used for primary cell isolation. The remaining tissues were fixed in paraformaldehyde for 24 hours and then embedded in paraffin for immunohistochemistry analysis.

The simplified immunohistochemistry protocol used is as follows: Paraffin sections (3 μm thick) of the ectopic and eutopic endometrium were dewaxed in xylene and ethanol, incubated in 3% hydrogen peroxide and goat serum albumin to remove endogenous peroxidase. The sections were incubated overnight with the following antibodies: 10 μg/ml rabbit anti-human vimentin, 10 μg/ml rabbit anti-human CK7, and 25 μg/ml mouse anti-human NOD1. PBS was used as a negative control. Subsequently, the sections were incubated within a moist chamber at the temperature of 4℃, followed by washing for 3 times. Next, the expression levels of proteins were determined utilizing AB complex (streptavidin/peroxidase) method (SP-9001; Zhongshanjinqiao). Color development was achieved using 3,3’-diaminobenzidine, followed by counterstaining with hematoxylin.

RNA was extracted from both tissues and cells using TRIzol reagents (Invitrogen), followed by reverse transcription into cDNA using a PrimeScript RT reagent Kit (TaKaRa) in an Applied Biosystems pattern recognition receptor (PCR) machine (Applied Biosystems). The cDNA of each sample was diluted and amplified for real time quantitative PCR (RT-qPCR) in a final volume of 20 μl, containing 2 μl of the cDNA template, SYBR Premix Ex Taq II, and Rox reference dye (TaKaRa). The forward and reverse primers were purchased from Sangon Biotech. The primer pairs used for gene amplification of glyceral-dehyde-3phosphate dehydrogenase (GAPDH), NOD1, transforming growth factor-β1 (TGF-β1), vascular endothelial growth factor A (VEGFA), matrix metalloproteinase 2 (MMP2), MMP3, and IL-8 are as follow:

GAPDH: forward 5’-GCACCGTCAAGGCTGAGAAC-3’, reverse 5’-TGGTGAAGACGCCAGTGGA-3;

NOD1: forward 5’-TACTGAAAAGCAATCGGGAACT-3’, reverse 5’-GTAGAGGAAGAACTCGGACACC-3’;

TGF-β1: forward 5’-GGCCAGATCCTGTCCAAGC-3’,reverse 5’-GTGGGTTTCCACCATTAGCAC-3’;

VEGFA: forward 5’-CGGATCAAACCTCACCAAG-3’, reverse 5’-ACGCTCCAGGACTTATACC-3’,

MMP2: forward 5’-GCAATACCTGAACACCTTCTATG-3’, reverse 5’-ATTCTGGTCAAGATCACCTGTC-3’,

MMP3: forward 5’-GGTCTCTTTCACTCAGCCAACAC-3’, reverse 5’-CAGGCGGAACCGAGTCAGG-3’;

IL-8: forward 5’-ACTGAGAGTGATTGAGAGTGGAC-3’, reverse 5’-AACCCTCTGCACCCAGTTTTC-3’.

The reactions were performed at 95℃ for 30 seconds, followed by 40 cycles at 95℃ for 5 seconds and 60℃ for 30 seconds. Each sample was analyzed in triplicate using an ABI Prism 7500 Sequence Detector (Applied Biosystems). The amplified sequences were quantified against a standard curve. The expressions of the genes were normalized to that of GAPDH.

Stromal cells and MSCs were isolated from the eutopic endometrium and ectopic endometriosis tissues using collagenase II digestion (Sigma-Aldrich). The primary cells were seeded into DMEM/F-12 (Gibco) complete medium supplemented with 10% inactivated fetal bovine serum (FBS; Gibco), 100 U/ml penicillin, and 100 mg/ml streptomycin. The cells were incubated at 37℃ with 5% CO₂. Non-adherent hematopoietic cells were removed by washing with PBS after 24 hours to isolate pure stromal cells. When the cells reached approximately 90% confluence, they were passaged using 0.25% Trypsin-ethylene diamine tetraacetic acid (EDTA; Gibco) at a 1：2 ratio. Differentiation studies were conducted on cells from passages 3 and 4.

Stromal cells were seeded into 6-well plates until they reached 90% confluency. Subsequently, the cells were digested using 0.25% trypsin, centrifuged at 1,000 rpm for 5 minutes, and the supernatant was discarded. The cells were then washed twice with PBS and suspended in 1 ml of sheath solution. Afterwards, the cell concentration was adjusted to 1×10⁶/ml. Next, 100 μl of the cell suspension was transferred to each tube and incubated with antibodies against CD31, CD44, CD73, CD140b, CD146, SUSD2, and CD271 for 30 minutes. Following the incubation, 500 μl of sheath solution was added. The antibody-labeled cells were subsequently analyzed and sorted using a FACSCalibur Flow Cytometer (BD Biosciences) within 1 hour. The obtained data was analyzed using FlowJo 10 software (FlowJo, LLC).

To evaluate the differentiation capacities of osteogenic, adipogenic, and chondrogenic phenotypes, three lineage differentiation detection kits (Cyagen) were utilized following the manufacturer’s instructions. The MSCs were seeded into a 24-well plate (Corning) with 500 µl of complete culture medium at a density of 5×10⁴ cells per well. Once the cells reached 80% confluency, the complete medium was replaced with the differentiation medium. The medium was changed every 3 days over a period of 21 days. Staining with Oil Red O, Alizarin Red, and Alcian Blue dye solutions was performed to evaluate the adipocyte, osteoblast, and chondrocyte genetic capacity of the MSCs.

Cells were seeded into 6-well plates at a density of 1×10³ cells per well. eut-MSCs and ect-MSCs were pretreated with various concentrations of NOD1 ligand γ-D-glutamyl-meso-diaminopimelic acid (Tri-DAP) (Invitrogen) or NOD1 inhibitor ML-130 for 10 days until obvious colony-forming unit (CFU) formation (more than fifty cells per CFU) was observed. Subsequently, the cells were fixed and stained using Giemsa’s stain (Solarbio).

To evaluate MSC proliferation, a cell counting assay kit (Cell Counting Kit-8, CCK-8; Tongren) and immunofluo-rescence for Ki67 were used. The CCK-8 kit utilizes a water-soluble tetrazolium salt that is reduced through cellular dehydrogenase activity to present a yellow formazan dye, in proportion to living cell number: the MSCs were seeded into 96-well plates at a density of 1×10³ cells per well. eut-MSCs were pretreated with various concentrations of Tri-DAP for 72 hours, while ect-MSCs were pretreated with various concentrations of ML-130. After-ward, CCK-8 assay (10 μl) was supplemented to each well, followed by the incubation of the mixture for 2.5 hours. Subsequently, the optical density of each well was determined at 450 nm using an enzyme-linked immuno-sorbent assay (ELISA) Reader (Molecular Devices) and the number of cells were calculated. Ki67 immunofluo-rescence staining: The primary cells cultured on chamber slides were fixed using cold 4% paraformaldehyde and then permeabilized with 0.1% Triton X-100 for 15 minutes. After washing using PBS, the cells were incubated overnight with mouse anti-human Ki67 (diluted 1：200; ZSGB-BIO) as a primary antibodies, and then the slides were incubated with Alexa Fluorconjugated (green fluorescence) as secondary antibodies (Abcam) for another 1 hours at room temperature and then counter stained with 4’,6-dia-midino-2-phenylindole (blue color, P36966; Invitrogen). The stained cells were viewed using laser scanning confocal microscope (LSM780; Zeiss). The proliferative cell index was calculated as the percentage of green nucleus cells in each total cell number (green and blue nucleus staining cells).

MSCs were seeded into 12-well transwell plates (Corning) at a density of 5×10⁴ cells in 200 μl per well and cultured with various concentrations of ML-130 (0.28 or 0.56 μM) for 24 hours. The cells on the lower membranes of the wells were then fixed and stained with Giemsa’s stain. The number of cells that had successfully migrated and invaded was calculated to analyze their migration and invasion capacities.

Apoptosis Detection Kit I (BD Biosciences) was used to evaluate the ratio of apoptotic cells. The cells were seeded into 12-well plates (Corning) at a density of 1×10⁶ cells per well and cultured with or without ML-130 for 24 and 48 hours. Cells were harvested using 0.25% trypsin without EDTA and collected with PBS into individual polystyrene round-bottom tubes 2045 (BD Falcon). Then, they were stained with Annexin-V and propidium iodide (PI) according to the manufacturer’s instructions simplified as follows: to each cell suspension, 10 μl Alexa Fluor 488-conjugated Annexin-V and 10 μl PI reagent were added. Cells were incubated in the dark for 15 minutes at room temperature. At the end of the incubation, the 400 μl binding buffer was added. Analysis was conducted using a fluorescence activated cell sorter (FACS) (FACSCalibur Flow Cytometer) as well as CellQuest software (BD Bioscie-nces). The percentage of early and late apoptotic cells was calculated using Annexin-V and PI double-positive cell staining, respectively.

The statistical tests used in this study were identified unambiguously. The t-test with one or two tails, linear correlation in the parametric (Pearson) or non-parametric (Spearman) method were employed. Statistical analysis was carried out utilizing SPSS Statistics 19.0 software (IBM Corp.), while Prism 5.0 (GraphPad) was adopted for graph plotting. Quantitative variables between groups were compared using Student’s t-test (for normal distribution) or Mann–Whitney U-test (for non-normal distribution), and one-way or two-way ANOVA was used for multiple com-parisons. Pearson χ² test or Fisher’s exact test was utilized to compare qualitative variables. Survival curves were generated using the Kaplan–Meier method and compared using the log-rank test. p<0.05 was considered statistically significant.

To identify the ectopic and eutopic endometrium tissues, HE staining was performed (Supplementary Fig. S1A, S1E), followed by CK7 and vimentin staining to distinguish between endometrium and endometriosis. The endometrium exhibited diffuse glands in the stromal cells (Supplementary Fig. S1A-S1D), while the endometriosis primarily consisted of stromal cells (Supplementary Fig. S1E-S1H).

RT-qPCR and immunohistochemistry were employed to compare the mRNA and protein expression levels of NOD1 between ectopic and eutopic endometrium, respectively. The immunohistochemistry results demonstrated strong positive expression of NOD1 in the endometriosis (Fig. 1A-1D). Moreover, NOD1 mRNA expression was higher in the extopic endometrium (ect-EM) tissue/endometriosis with statistical significance compared to the eutopic endometrium (eut-EM) tissue/endometrium (ect-EM tissue and eut-EM tissue were paired samples and derived from the same patient) (Fig. 1E). Notably, for patients with endometriosis, NOD1 expression in their eut-EM tissue did not differ significantly from that observed in the endometrium (EN) from healthy females (Fig. 1E, EN vs. eut-EM, p=0.738).

To explore whether and how NOD1 is functionally expressed in MSC-assisted endometrial ectopic implants, MSCs were isolated from the endometrial and endometriosis tissues. Adherent eutopic and ectopic ESCs were grown (Supplementary Fig. S2A, S2C). CD45(−) & CD31(−) & CD44(＋) stromal cells were selected as MSCs, achieving a purity of approximately 98% after 72 hours of culture (Supplementary Fig. S2B, S2D). The eut-MSCs and ect-MSCs exhibited spindle-shaped or short and conical morphology. Adipogenic, osteogenic, and chondrogenic differentiation successfully induced multilineage differentiation potential of CD44(＋) MSCs. Immunostaining revealed the presence of NOD1 protein in both CD44(＋) eut-MSCs and ect-MSCs (Fig. 2A, 2B). Additionally, an increasing trend was observed in NOD1 mRNA expression among the ect-MSCs, although the trend was not statistically significant (Fig. 2C).

Stimulation with the various concentration of NOD1 ligand, Tri-DAP, increased the number of CFUs and the proliferation of eut-MSCs (Fig. 3A, 3B), which was consistent with their increased viability and proliferation rates in Ki67 assays (Fig. 3C). The significantly higher proportion of proliferative cells were observed in the proliferation phase compared to the control cells (pretreated with PBS).

Since MSCs originating from endometrial tissue are implanted outside the uterine cavity, they need to tolerate the local microenvironment properly. To explore the differences in chemokine/cytokine profiles between eut-MSCs and ect-MSCs, we measured the transcription levels of IL-8 (Fig. 3D). The ect-MSC samples exhibited significantly higher IL-8 expression compared to the eut-MSC group (p<0.05). Moreover, the ect-MSC samples also performed significantly enhanced migratory capacity compared to the eut-MSC group (Supplementary Fig. S3) (p<0.0001).

After pretreatment with ML-130, the inhibitory effect on ect-MSCs was evaluated at 24, 48, and 72 hours. Nota-bly, after 48 hours, the proliferation of ect-MSCs was significantly inhibited (Fig. 4A, right panel). In addition, clonogenesis, invasion, and migration abilities decreased following ML-130 pretreatment, as shown in Fig. 4A left panel and 4B. The transcription and secretion levels of IL-8 were higher in the ect-MSCs group with statistical significance compared to the eut-MSCs group (Fig. 4C, 4D). However, ML-130 pretreatment partially inhibited the upregulated expression of IL-8 (Fig. 4C, 4D).

Primary ect-MSCs were stimulated with varying concentrations (0, 0.28, or 0.56 μM) of ML-130 for a duration of 48 hours (Fig. 5). Although there was a noticeable trend towards early apoptosis in the ect-MSCs, the observed difference between pretreatment cells and control cells were not statistically significant (Fig. 5B-5E) (p>0.05).