Precise capture of circulating endometrial cells in endometriosis

Endometriosis (EM) refers to the appearance of endometrial tissue (glands and stroma) outside the endometrium. [ 1 ] EM is a benign disease; however, it has malignant biological behaviors, such as implant erosive growth, distant metastasis, and easy recurrence, and can invade any part of the body. EM is a hormone-dependent disease and affects 10–15% of women of reproductive age, although the prevalence may actually be higher due to diagnostic difficulties and delays in diagnosis. It is estimated that approximately 190 million women worldwide suffer from EM. [ 2 ] Clinical manifestations include pelvic pain, dysmenorrhea, infertility, pelvic nodules, and masses. [ 3 ] The prevalence rate of EM among symptomatic women is 35–100%. [ 4 ] In addition, there is the possibility of malignant transformation, and the malignant transformation rate is approximately 1–4%. [ 5 , 6 ] The gold standard for EM diagnosis is laparoscopy, and the degree of adhesion and the color, sizes, and shapes of EM lesions and nodules can be seen during surgery. [ 7 ] At present, the possibility of EM is judged mainly by clinical diagnostic criteria, but at least 18% of patients with clinically diagnosed EM have no histological evidence to support EM. [ 8 ] Imaging tests such as transvaginal ultrasonography have limited utility and poor specificity in the diagnosis of EM. [ 9 ] The most commonly used serum marker of EM is carbohydrate antigen 125 (CA125), but its specificity is also poor. [ 10 ] Due to the heterogeneity of EM lesions, the efficacy of treatment in patients also varies, and the recurrence rate is high. [ 11 ] Among EM patients with the same clinical stage, the clinical effects of the same dose of gonadotropin-releasing hormone therapy are significantly different, which also increases the difficulty of disease treatment, and so stratified management and individualized treatment of EM patients are also particularly important. [ 12 , 13 ] In 2017, our research group detected circulating endometrial cells (CECs) in the peripheral blood of EM patients with the microfluidic chip method, and the detection rate was 89.5% (17/19), [ 14 ] suggesting that CECs may be useful as a new type of biomarker for the diagnosis of EM. However, the principle of the above method is based on cell size, and the diameter of the filtered cells is greater than 8 μm, which may cause our target cells to be missed. Moreover, there are many circulating vascular endothelial cells (CVECs) in human peripheral blood. These cells, like CECs, are often estrogen receptor/progesterone receptor (ER/PR) positive and are mixed in the final cell population. Therefore, in this study, we used the more advantageous method of subtraction enrichment and immunostaining fluorescence in situ hybridization (SE-iFISH) to capture all cell populations and at the same time increase CD31 staining to distinguish CVECs to clearly analyze the number, size, and ploidy of CECs and the relationship between CECs and the clinical course characteristics of EM patients.

This study was supported by grants from the National Key Research and Development Program of China (No. 2022YFC2704000) and the National Natural Science Foundation of China (No. 81971360).

Approval was obtained from the Ethics Committee of Peking University People’s Hospital (No. 2019PHB108-01), written informed consent was obtained from patients before blood sampling, and clinical studies were carried out in accordance with the principles of the Declaration of Helsinki . Human peripheral venous blood used to detect CECs and CVECs was taken from EM patients ( n = 34) hospitalized in the Department of Gynecology of Peking University People's Hospital for surgical treatment. The inclusion criteria were as follows: (1) diagnosis of EM by laparoscopy; (2) women who were not pregnant, breastfeeding, or menopausal at the time of blood collection; (3) no history of other malignant or benign tumors; (4) no hematological diseases or white blood cell abnormalities; and (5) no use of steroidal hormone drugs (including oral contraceptives) during the 3 months before blood collection or subcutaneous implantation of contraceptives (e.g., intrauterine release devices for progesterone). There were 12 participants in the control group, 9 of whom underwent surgical treatment during the same period and were pathologically diagnosed with hydrosalpinx or other benign masses. The other 3 participants were healthy volunteers and had no symptoms related to EM and no abnormalities on gynecological color Doppler ultrasound. Peripheral blood was collected from one patient with EM and two healthy volunteers for testing during the menstrual, proliferative, and secretory phases of the menstrual cycle. The stage of the menstrual cycle at the time of preoperative blood collection was determined by an experienced pathologist through histological methods, and EM staging was performed according to the revised American Fertility Society (r-AFS) classification (1996). The general clinical data of participants are shown in Supplementary Table 1, http://links.lww.com/CM9/B785 ; the complete clinical data, such as symptoms, surgical records, and pathology, are shown in Supplementary Table 2, http://links.lww.com/CM9/B785 . All peripheral blood samples were collected from preoperative blood in the morning of the operation day. A blood collection tube containing acid citrate dextrose (ACD) anticoagulant (Becton Dickinson, Franklin Lakes, NJ, USA) was used to draw 6 mL of peripheral venous blood from each participant, and all blood samples were drawn after other clinical blood samples were collected to prevent endothelial cells from contaminating the blood samples. After completion, the contents in the tube were mixed by inverting the tube eight times. The tube was then stored at room temperature in the dark and processed within 24 h. SE of CEC and CVEC was performed according to the manufacturer's instructions (Cytelligen, San Diego, CA, USA). [ 15 , 16 ] After the blood collection tube was leveled and mixed upside down, it was centrifuged at room temperature for 15 min (400 × g ). The supernatant was discarded up to 5 mm above the brown–red precipitate, the cleaning solution 1 × CRC (Cytelligen, San Diego, CA, USA) was added to the yellow line of the blood collection tube label, the tube cap was covered, and the context was mixed by inverting the tube 10 times. A total of 3 mL of the sample density separation solution (Cytelligen, San Diego, CA, USA) was added to 50-mL centrifuge tube A, and an electric pipette was used to slowly add the blood cells in the blood collection tube to the top of the separation solution along the liquid surface of centrifuge tube A. After trimming, the cells were centrifuged at room temperature for 6 min (350 × g ). After centrifugation in tube A, all the liquid in layers 1 and 2 was transferred to 50-mL centrifuge tube B, and the liquid was mixed by shaking. The magnetic bead buffer (Cytelligen, San Diego, CA, USA) was mixed thoroughly, followed by slow addition to centrifuge tube B at a ratio of 300 μL per tube. The centrifuge tube was shaken while adding the buffer to mix the magnetic bead buffer. Then, the centrifuge tube was fixed on a shaker at an angle of 35°–40° and shaken at room temperature for 20 min at 125 r/min (the highest scale of liquid level shaking was between 30 mL and 35 mL). Next, tube B was placed on the magnetic stand, a 5-mL pipette tip was used to reach the bottom of the tube, and the tube was gently pipetted twice at 1 min. Then, the same pipette tip was used to carefully transfer the liquid to 50-mL centrifuge tube C along the center of tube B after 2 min. Next, washing solution 1 × CRC was added to 45 mL in 50-mL centrifuge tube C, the tube was balanced and inverted to mix its contents, and the contents were centrifuged at room temperature for 5 min (500 × g ). Then, the supernatant was discarded after preserving 500 μL of supernatant, the cells were gently shaken in a vortex mixer, washing solution 1 × CRC was added up to 45 mL, and the tube was balanced and inverted to mix its contents. Finally, the tube was centrifuged at room temperature for 5 min (400 × g ), and the supernatant was discarded after preserving 100 μL of supernatant. A total of 2 μL of Antigen Retrieval Buffer (Cytelligen, San Diego, CA, USA) was added to the liquid surface of centrifuge tube C. A shaking mixer was used to gently shake and mix the pelleted cells, and the suspension was allowed to stand at room temperature for 10 min. At this time, the staining solution was prepared to aspirate the staining solution at 2–8°C, and the volume used was calculated according to each slide. The staining solution comprised 200 μL of blood cell analysis diluent (Cytelligen, San Diego, CA, USA), with 4 μL of Cy5-labeled blood cell analysis staining solution for CD45 (Cytelligen, San Diego, CA, USA), 1 μL of Cy7-labeled blood cell analysis staining solution for CD31 (Cytelligen, San Diego, CA, USA), 1 μL of Alexa Fluor 488-labeled blood cell analysis staining solution for ER (Abcam, Cambridge, UK), and 1 μL of Alexa Fluor 594-labeled blood cell analysis staining solution for PR (Abcam, Cambridge, UK). For the testing of antibodies in cell lines, see Supplementary Table 3, http://links.lww.com/CM9/B785 and Supplementary Figure 1, http://links.lww.com/CM9/B785 . All 200 μL of the abovementioned mixture was added to tube C under dark conditions, gently shaken and mixed, and incubated in the dark at room temperature for 20 min (gently mixed to precipitate the cells after 10 min). After washing, the supernatant was discarded after preserving 100 μL of supernatant, 100 μL of tissue fixative Cytelligen was added, and each tube of specimen liquid was applied to the Cytelligen slide specimen frame (one person per slide). Then, the specimen was placed in a 30–33°C drying oven for subsequent testing. FISH was performed according to the kit instructions. A total of 20 μL of tissue fixative FR1 (Cytelligen, San Diego, CA, USA) and 180 μL of sample diluent FR2 (Cytelligen, San Diego, CA, USA) were mixed thoroughly by shaking, and the mixture was then added to the specimen, which was allowed to stand for 10 min in the dark at room temperature. Then, the mixture was aspirated. Next, 200 μL of buffer 1 × FR3 (Cytelligen, San Diego, CA, USA) was added dropwise to wash the specimen twice. Then, 200 μL of absolute ethanol was added dropwise, immediately aspirated and discarded, and this step was repeated twice. Next, the slide was inserted into absolute ethanol and allowed to stand for 2 min. After it was completely dry, 10 μL of chromosome 8 centromere probe (Cytelligen, San Diego, CA, USA) was immediately added, and the slide was mounted and placed into the hybridization machine. Hybridization conditions were as follows: denaturation at 76°C for 10 min and hybridization at 37°C for 3 h. After hybridization, the slides were placed in 1 × FR3 buffer preheated to 37°C and shaken gently until the cover glass fell off. Then, 200 μL of antibody was removed and added dropwise. The solution was aspirated twice and blown dry. Afterward, 10 μL of 4',6-diamidino-2-phenylindole (DAPI) staining solution was added for blood cell analysis, the cover glass was placed on the specimen while avoiding air bubbles, and the slide was mounted. Finally, image scanning and analysis were performed. The slides were automatically loaded onto a Zeiss fluorescence microscope (AXIO Imager Z2, Zeiss, Oberkochen, Germany) and then subjected to automated X–Y scanning with cross Z-sectioning of all cells performed at 1-μm steps of depth. XYZ 3D scanning was performed in each of the six fluorescence color channels. Positivity for the target cell was defined as follows: DAPI+/CD45–/CD31–/ER+ or PR+ chromosome 8 diploidy or aneuploidy for CECs and DAPI+/CD45–/CD31+/ER+ or –/PR+ or-chromosome 8 diploidy or aneuploidy for CVECs. After high-throughput scanning, cell images were collected and processed, and the phenotype, cell size, and ploidy of chromosome 8 of each target cell were statistically analyzed. Comparisons between two groups were performed by Student's t -test for continuous variables with a normal distribution and by the non-parametric Mann–Whitney test for continuous variables with a skewed distribution. Graphs were generated with GraphPad Prism 8.0 software (GraphPad Software Inc., San Diego, CA, USA). All data analyses were performed using SPSS v25 software (SPSS Inc., Chicago, IL, USA). P values <0.05 were statistically significant.

The peripheral blood of 34 EM patients and 12 non-EM patients was evaluated by SE-iFISH [Figure 1 ]. The results accurately captured DAPI+/CD45–/CD31–/ER+ or PR+ CECs and DAPI+/CD45–/CD31+/ER+ or –/PR+ or – CVECs [Figure 2 A]. Using leukocytes as a control, many small cells (<5 μm) and large cells (≥5 μm) could be seen among CECs under the microscope. The statistical results of the morphological analysis showed that in the peripheral blood of patients with active EM, small cells comprised 63.5% (40/63) of the CEC population, and cells with larger diameters than those of white blood cells comprised the other 36.5% (23/63) [Figure 2 B]. General process of detecting CECs by the SE-iFISH method. Six milliliters of venous blood is collected from EM patients, and a non-hypotonic method is used to remove RBCs from the blood of EM patients. A combination of various anti-leukocyte antibodies, including CD45, is coupled to specially coated magnetic beads to remove leukocytes to the greatest extent. Then, endometrium-specific antibodies against ER and PR are used for identification, and CECs and CVECs are distinguished according to the characteristics of CD31 immunofluorescence staining. Finally, the ploidy of chromosome 8 is identified by chromosome in situ hybridization. CECs: Circulating endometrial cells; CEP8: Centromere of chromosome 8;CVECs: Circulating vascular endothelial cells; EM: Endometriosis; ER: Estrogen receptor; PR: Progesterone receptor; RBCs: Red blood cells; SE-iFISH: Subtraction enrichment and immunostaining fluorescence in situ hybridization. Representative microscopic images and sizes of CECs and CVECs. (A) Representative microscopic fluorescence images of CECs and CVECs. Positivity for the target cell was defined as follows: DAPI+/CD45–/CD31–/ER+ or PR+ for CECs and DAPI+/CD45–/CD31+/ER+ or –/PR+ or – for CVECs. Scale bars, 5 μm. (B) Representative microscopic fluorescence images of small and large CECs. The proportions of small cells (diameter <5 μm) and large cells (diameter ≥5 μm) among CECs. Scale bars, 5 μm. CECs: Circulating endometrial cells; CVECs: Circulating vascular endothelial cells; ER: Estrogen receptor; PR: Progesterone receptor. The statistical results showed that the detection rate of ER/PR-positive cells (CEC + CVEC) in patients with EM was 85.3% (29/34), which was similar to that with the previous microfluidic chip method (89.5%, 17/19) [ 13 ] [Figure 3 A]. The number of cells was significantly higher in EM patients than in controls (8.24 vs. 2.00 cells/6 mL, P = 0.0477) [Figure 3 B]. After exclusion of the interference of CVECs, the detection rates of CECs were 58.8% (20/34) in EM patients and 16.7% (2/12) in the control group [Figure 3 A], and the number of cells was significantly higher than that in the control group (1.94 vs. 0.17 cells/6 mL, P = 0.0071) [Figure 3 B]. At the same time, we also measured the detection rate and number of CVECs and found that the detection rates of CVECs were 73.5% (25/34) in EM patients and 66.7% (8/12) in the control group [Figure 3 A]; but there was no significant difference in number of cells (6.29 vs. 1.83 cells/6 mL, P = 0.2655) [Figure 3 B]. Analysis of the correlation between the number of CECs and CVECs showed that the correlation coefficient was 0.6704 ( P = 0.0004); when the number of CECs increased, the number of CVECs also increased [Figure 3 C]. Comparison of the positive rates and numbers of CECs and CVECs in the EM and control groups. (A) The positive rates of CECs and CVECs and the sum of the two in the EM and control groups. (B) The numbers of CECs and CVECs in the EM and control groups and the sum of the two. * P <0.05 vs . non-EM, † P 0.05 vs . non-EM. (C) The correlation coefficient for CECs and CVECs is 0.6704 ( P <0.05). CECs: Circulating endometrial cells; CVECs: Circulating vascular endothelial cells; EM: Endometriosis. We analyzed the clinical characteristics of CEC-positive patients and found that the number of CECs was related to the length of the disease course and the size of the lesions in EM patients; that is, 85.0% (17/20) of CEC-positive EM patients had pelvic pain or dysmenorrhea in the past 6 months, progressive aggravation of symptoms, or the presence of a significant increase in ovarian mass >2 cm [Supplementary Table 4, http://links.lww.com/CM9/B785 ]. To better explore the relationship between CEC and other rare cells in the blood and the onset and clinical symptoms of EM, we analyzed EM-related symptoms, onset priority data, gynecological color Doppler ultrasound data, gynecological examination data, and intraoperative lesions. The patients were divided into two groups, "active EM" and "dormant EM". The specific manifestations of the "active EM" were as follows. The gynecological examination and imaging examination led to a diagnosis of suspected EM. Among these patients, (1) the symptoms of pelvic pain or dysmenorrhea progressively worsened over the past 6 months, and (2) there were obvious ovarian masses on gynecological examination and imaging over the previous 6 months with size increases >2 cm. Patients with signs in line with any of the above were considered to have active EM. The specific manifestations of the "dormant EM" were as follows. The gynecological examination and imaging studies led to suspected EM. Among these patients, (1) there was no pelvic pain or dysmenorrhea over the previous 6 months or there was mild pain without progressive aggravation, and (2) there was no obvious increase in tumor size over the past 6 months (<2 cm). If both conditions were met, patients were considered to have dormant EM. The results showed that the positive rate of CECs in the peripheral blood of active EM patients was 94.4% (17/18), while the positive rates in dormant EM patients and non-EM controls were 18.8% (3/16) and 16.7% (2/12), respectively [Figure 4 A]. At the same time, the positive rate of CVECs in the peripheral blood of active EM patients was 83.3% (15/18), and the positive rates in dormant EM patients and non-EM controls were 62.5% (10/16) and 66.7% (8/12), respectively. The number of CECs and CVECs in active EM was higher than that in dormant EM [Figure 4 B]. The average number of CECs in active EM patients was 3.50 cells/6 mL, which was significantly higher than that in dormant EM patients (3.50 vs. 0.21 cells/6 mL, P <0.0001) and significantly higher than that in the control group (3.50 vs. 0.20 cells/6 mL, P 0.9999). The average number of CVECs in active EM patients was 9.22 cells/6 mL, which was significantly higher than that in dormant EM patients (9.22 vs. 3.00 cells/6 mL, P = 0.0407) and significantly higher than that in the control group (9.22 vs. 1.83 cells/6 mL, P = 0.0297), and there was no significant difference in the number of CVECs between the latter two groups ( P = 0.8290). The average number of the two in active EM patients was 12.72 cells/6 mL, significantly higher than that in the dormant EM patients (12.72 vs. 3.19 cells/6 mL, P = 0.0014) and significantly higher than that in the control group (12.72 vs. 2.00 cells/6 mL), P = 0.0007), and there was no significant difference between the latter two groups ( P = 0.9190). Comparison of the positive rates and numbers of CECs and CVECs in different groups. (A) The positive rate of CECs, that of CVECs, and the sum of the two in the active EM, dormant EM, and non-EM control groups; (B) CECs, CVECs, and the sum of the two in the active EM, dormant EM, and non-EM groups. † P <0.05 vs . dormant EM; * P <0.05 vs . non-EM, † P <0.01 vs . dormant EM, ‡ P <0.001 vs . non-EM, § P <0.0001 vs . non-EM. CECs: Circulating endometrial cells; CVECs: Circulating vascular endothelial cells; EM: Endometriosis. Because the operation time is the proliferative or secretory phase of menstruation and considering that the exclusion of surgery and postoperative drugs may affect the test results, preoperative blood collection results were not included in the analysis of patients with EM. The analysis results showed that in the peripheral blood of patients with active EM, the average number of CECs in the proliferation phase was 4.8 cells/6 mL, and the number in the secretion phase was 1.4 cells/6 mL. The average was 1.9 cells/6 mL. The sum of the two types of cells was an average of 18.7 cells/6 mL in the proliferation phase and 3.3 cells/6 mL in the secretion phase; the differences were significant ( P = 0.0161) [Figure 5 A]. In the peripheral blood of dormant EM patients, the average number of CECs in the proliferation phase was 0.3 cells/6 mL, and the average number in the secretory phase was 0.2 cells/6 mL. Additionally, the average number of CVECs in the proliferation phase was 1.7 cells/6 mL, and the average number in the secretion phase was 4.0 cells/6 mL. The sum of the two types of cells was an average of 2.0 cells/6 mL in the proliferation phase and 4.1 cells/6 mL in the secretion phase, and the difference was not significant ( P = 0.5500) [Figure 5 B]. In the peripheral blood of the non-EM control group, the average number of CECs in the proliferation phase was 0.1 cells/6 mL, and the average number in the secretion phase was 0.3 cells/6 mL. Additionally, the average number of CVECs in the proliferation phase was 1.9 cells/6 mL, and the average number in the secretion phase was 1.8 cells/6 mL. The sum of the two types of cells averaged 2.0 cells/6 mL in the proliferative phase and 2.0 cells/6 mL in the secretory phase, and the difference was not significant ( P >0.9999) [Figure 5 C]. The relationships of CECs and CVECs with menstrual cycle and EM stage. (A–C) The relationship of the number of CECs, number of CVECs, or the sum of the two with menstrual cycle phase in the active EM group ( * P <0.05, ‡ P <0.001), as well as the dormant EM and non-EM control groups. (D, E) Number of CECs, number of CVECs, and the sum of the two in active EM and dormant EM and their relationships with the stages of EM. CECs: Circulating endometrial cells; CVECs: Circulating vascular endothelial cells; EM: Endometriosis; ns: Not significant. We analyzed the relationship between the number of CECs in patients with EM and the stage of EM, and the results showed that in the peripheral blood of patients with active EM, the average number of CECs in stage I–II EM was 1.9 cells/6 mL, and the average number in stage III–IV EM was 4.9 cells/6 mL; the average number of CVECs was 10.6 cells/6 mL in stage I–II EM and 9.1 cells/6 mL in stage III–IV EM. The average number of the sum of CECs and CVECs was 12.4 cells/6 mL in stage I–II EM and 14.0 cells/6 mL in stage III–IV EM, and the difference was not significant ( P = 0.7942) [Figure 5 D]. In the peripheral blood of patients with dormant EM, CECs were not detected in stage I–II EM, but an average of 0.3 cells/6 mL was detected in stage III–IV EM. CVECs averaged 2.8 cells/6 mL in stage I–II EM and 3.1 cells/6 mL in stage III–IV EM. The sum of the two averaged to 2.8 cells/6 mL in stage I–II EM and 3.4 cells/6 mL in stage III–IV EM, and the differences were not significant ( P = 0.6465) [Figure 5 E]. Therefore, the number of CECs and CVECs in the peripheral blood of EM patients, whether active or dormant, has nothing to do with the EM stage. We analyzed the ploidy status of chromosome 8 in all CECs. An example image of the cells under the microscope is shown in Figure 6 A. It was found that 44.4% of the CECs had the characteristics of aneuploidy on chromosome 8, and tetraploidy and polyploidy were the main types [Figure 6 B]. Chromosome 8 aneuploidy and subtype analysis of CECs. (A) Representative fluorescence images of aneuploid CECs. (B) Proportions of different chromosome 8 ploidy subtypes; most CECs were normal diploid, 4.1% were haploid, 4.8% were triploid, 3.1% were tetraploid, and 32.4% were polyploid. CECs: Circulating endometrial cells.

EM is a very complex condition, and its diagnosis and mechanistic research has faced many obstacles. [ 17 ] The gold standard for the diagnosis of EM is laparoscopy; thus far, there is still no one or groups of effective biomarkers, and the diagnosis is typically delayed by approximately 7.5 years. [ 18 , 19 ] At the same time, there is still no theory that can fully explain the pathogenesis of EM. [ 20 ] The discovery of CECs provides a new direction for clinical and basic research on EM. [ 14 ] However, the CEC detection method used previously was the microfluidic chip method we used in the early stage or the porous polycarbonate membrane filtration method with a pore size of 8 μm used by Bobek et al. [ 21 ] The ScreenCell device used in the Spanish study conducted in 2009 is based on the physical characteristics of the cell size, that is, a diameter of the filtered cell greater than 8 μm. [ 22 ] Thus, cells smaller than 8 μm may be missed, and this aspect of the method type may be more important in terms of the cell type being analyzed in EM, similar to the small-cell circulating tumor cells (CTCs) in research. [ 23 ] In addition, there is interference of CVECs mixed in the final cell population. [ 24 ] In this study, we analyzed the size of the target cells and confirmed that there are indeed a large number of cell populations smaller than 8 μm in the target cells. The SE-iFISH method achieves the enrichment of all target cells, is not based on the physical characteristics of cell size, is not limited by cell size, does not miss smaller cells, is efficient, time-saving, and labor-saving, and is more suitable for clinical application. Regarding the staining identification of CECs, we did not choose CD10 used in other studies mentioned above but chose ER and PR, which are highly expressed in 90% of endometrial epithelium and stroma, to judge whether they are target cells, because in EM, the cell type of interest present in the patient's peripheral blood may not be mesenchymal cells. With this new method, we more accurately captured CECs in the peripheral blood of EM patients and differentiated CVECs. With the improvement in the accuracy of CEC detection, we found that not all EM patients have CECs in their peripheral blood. CECs were present mainly in the peripheral blood of EM patients whose dysmenorrhea symptoms were aggravated or whose lesions increased rapidly over the previous 6 months, namely, active EM patients. Interestingly, the CECs and CVECs significantly increased in the proliferative phase only in the peripheral blood of active EM patients, and the test results in different menstrual cycles were in line with this trend; however, there was no increase in the proliferative phase of the cycles in the control group. We speculate that when endometrial fragments are shed during menstruation, there may be more endometrial cells entering the blood in patients with active EM, which then implant in the open sinusoids and form lesions in suitable places through blood circulation. Therefore, the detection of CECs may have clinical guiding significance for identifying which patient groups need timely attention and for early diagnosis and treatment to alleviate the pain caused by the disease. Due to the particularity of EM disease, the correlation between its severity and clinical symptoms is poor, and the appearance of EM lesions is also significantly different, including the degree of adhesion, color, size, and shape, which vary from person to person. As a complex disease with various lesions, various clinical manifestations, unclear mechanisms, and unstable treatment effects, it is very important to stratify patients in the study of EM. In addition, with regard to the detection of the ploidy number of chromosome 8, it was found that CECs were mainly normal diploid in EM patients, but there were also a large number of aneuploid cells, and the main type was tetraploid. The more characteristic triploid types in malignant tumors are less common in EM patients. [ 25 ] We speculate that this finding also reflects to some extent that EM is a benign disease but has the characteristics of malignant biological behavior. The detection of aneuploidy in CECs may be of great significance in the malignant transformation of EM into ovarian cancer. This study has certain limitations. The study participants mainly had ovarian EM, and the number of cases is still low. In the future, the sample size will be further expanded, and research of other types of EM will be performed. In addition, for patients with dormant EM with detected CECs, it is not clear whether new lesions will develop and progress to active EM. Early detection and dynamic tracking of CECs for dormant EM patients and even adolescent patients with dysmenorrhea will be very important. In adolescents, since early-onset dysmenorrhea is a risk factor for EM and adolescent EM is difficult to identify and differentiate through imaging or CA125, CECs may be used in the detection of adolescent dysmenorrhea and early prediction or detection of EM, thereby further reducing diagnostic delays. In conclusion, we captured CECs more precisely in this study, overcoming the limitations of previous methods. Moreover, CECs are related mainly to active EM and may be used as a biomarker in the early diagnosis and treatment, personalized management, and prediction of malignant transformation of EM in the future.

None.