Persistent Pelvic Adhesions and Fibrotic Remodeling Following Ectopic Lesion Ablation in a Mouse Model of Endometriosis

Kyeong A. So, Jung‐Yoon Yoo, Andrew M. Kelleher, Tae Hoon Kim, Jung‐Ho Shin, and Jae‐Wook Jeong conceived and designed the research. Sohyeon Yun, Kyeong A. So, Eunhee M. Jeong, Marissa G. Hoover, Abu Raihan, and Tae Hoon Kim performed the research and acquired the data. Sohyeon Yun, Kyeong A. So, Tae Hoon Kim, Jung‐Ho Shin, and Jae‐Wook Jeong analyzed and interpreted the data. Sohyeon Yun, Kyeong A. So, Jung‐Yoon Yoo, Tae Hoon Kim, Jung‐Ho Shin, and Jae‐Wook Jeong wrote the manuscript. Kyeong A. So, Lisa Brennaman, Jung‐Yoon Yoo, Tae Hoon Kim, Jung‐Ho Shin, and Jae‐Wook Jeong edited the manuscript. All authors read and approved the final paper.

This work was supported by HHS | NIH | Eunice Kennedy Shriver National Institute of Child Health and Human Development (NICHD), R01HD108895, R01HD101243, R01HD102170 (to J.W.J) and by a grant from Korea University (to K.A.S.). The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

To determine the efficiency of DT‐mediated inducible ablation of Pgr ‐expressing endometrial cells, Pgr cre/+ Rosa26 iDTR/mTmG mice received two intraperitoneal injections of DT (100 ng per mouse per injection) or PBS vehicle separated by an 8‐h interval (Figure  1A ). GFP‐positive (mG + ) uteri were examined by fluorescence imaging at 24 and 48 h after the final injection of DT or vehicle ( n  = 5 per treatment per time point). While vehicle‐treated mice retained GFP‐positive uteri, GFP fluorescence was barely detected in the DT‐treated group at either the 24‐ or 48‐h time point (Figure  1B ). After uterine dissection, fluorescence imaging confirmed a marked reduction in GFP‐positive uterine tissue in DT‐treated mice compared with vehicle controls, consistent with efficient ablation of DTR‐expressing cells (Figure  1C ). Quantitative analysis of fluorescence intensity revealed a significant decrease in GFP intensity in DT‐treated mice at 24 h ((0.44 ± 0.08) × 10 8 ) compared with vehicle controls ((1.31 ± 0.14) × 10 8 ; p  < 0.01) (Figure  1D ). A similar reduction was observed at 48 h, with lower GFP intensity in DT‐treated mice ((0.27 ± 0.04) × 10 8 ) relative to vehicle‐treated controls ((1.17 ± 0.10) × 10 8 ; p  < 0.001). DT‐mediated inducible ablation of Pgr ‐expressing endometrial cells in Pgr cre/+ Rosa26 iDTR/mTmG mice. (A) Experimental schematic illustrating the protocol used to determine the efficiency of DT‐mediated inducible ablation of Pgr ‐expressing endometrial cells. (B) Representative bright‐field and fluorescence (mT and mG) images from vehicle‐ and DT‐treated mice at 24 and 48 h post‐treatment. GFP‐positive uterine tissues (green) and tdTomato‐labeled tissues (red) are shown. DT treatment resulted in a marked reduction in GFP‐positive uterine tissues compared with vehicle controls. (C) Representative fluorescence images of GFP‐positive uterine tissues from vehicle‐ and DT‐treated mice at 24 and 48 h post‐treatment. (D) Quantification of maximum radiance (photons/s/cm 2 /sr) from GFP‐positive uterine tissues at 24 and 48 h post‐treatment. DT‐treated mice exhibited a significantly reduced GFP signal intensity compared with vehicle‐treated controls at both time points. (E) Representative hematoxylin and eosin (H&E)‐stained uterine sections from vehicle‐ and DT‐treated mice at 24 and 48 h post‐treatment. (F) Representative immunohistochemical images (low and high magnification) of progesterone receptor (PGR) expression in uterine tissues from vehicle‐ and DT‐treated mice at 24 and 48 h post‐treatment. (G) Quantification of PGR H‐scores in epithelial and stromal compartments at 24 and 48 h post‐treatment. Data are presented as mean ± SEM. n  = 5 per treatment per time point. ** p  < 0.01, *** p  < 0.001 (one‐way ANOVA followed by Tukey's multiple‐comparisons test). To assess potential DT‐associated toxicity, body weight and uterine weight were measured 24 and 48 h after the final DT or vehicle injection. No significant differences in body or uterine weight were observed between DT‐ and vehicle‐treated mice at either time point, and no overt signs of toxicity were observed during the experimental period (Figure  S3 ). Next, the histological effects of DT‐mediated ablation on the uterus were examined (Figure  1E ). In vehicle‐treated mice, uterine tissue architecture remained intact, with preserved myometrium as well as endometrial epithelial and stromal compartments at both time points. In contrast, DT‐treated mice exhibited a marked reduction in endometrial epithelial and stromal compartments at both 24 and 48 h compared with vehicle controls (Figure  1E ). Higher‐magnification images revealed thinning of the epithelial layer and reduced density of stromal cells following DT treatment. In contrast, the myometrial compartment remained largely intact in both groups. Next, the efficiency of inducible ablation of PGR‐positive endometrial cells was assessed by PGR immunohistochemistry (Figure  1F ). As expected, strong PGR expression was observed in both endometrial epithelial and stromal compartments of vehicle‐treated uteri, whereas PGR expression was nearly undetectable in the endometrium of DT‐treated mice. Quantitative H‐score analysis confirmed robust PGR expression in vehicle‐treated uteri within both the epithelial (145.50 ± 30.52 and 181.40 ± 9.14, respectively) and stromal (101.20 ± 25.12 and 95.48 ± 1.90, respectively) compartments at 24 and 48 h (Figure  1G ). In contrast, DT treatment resulted in a marked loss of PGR expression at 24 h in the epithelium (3.97 ± 2.21, p  < 0.001) and stroma (3.38 ± 1.05, p  < 0.01), which persisted at 48 h in both epithelial (2.78 ± 1.59, p  < 0.001) and stromal (9.07 ± 1.90, p  < 0.01) compartments (Figure  1G ). These results demonstrate efficient and rapid ablation of PGR‐expressing endometrial cells using the iDTR system. To assess the effects of ectopic lesion ablation on behavioral outcomes, pelvic adhesions, and fibrotic remodeling, established ectopic lesions were selectively ablated and subsequent changes in these outcomes were evaluated (Figure  2A ). Syngeneic endometriosis was induced in recipient Rosa26 iDTR/mTmG mice by intraperitoneal administration of minced uterine fragments from donor Pgr cre/+ Rosa26 iDTR/mTmG mice. After ectopic lesions were allowed to develop for 1 month, mice received two intraperitoneal injections of DT (100 ng per mouse per injection) or PBS vehicle separated by an 8‐h interval ( n  = 9 per treatment). Sham control mice received the same DT or PBS vehicle treatments ( n  = 6 per treatment), resulting in four experimental groups: vehicle‐treated sham, DT‐treated sham, vehicle‐treated endometriosis, and DT‐treated endometriosis. To confirm the effectiveness of ectopic lesion ablation, GFP‐positive donor‐derived lesions were evaluated by fluorescence imaging following behavioral assessment. Vehicle‐treated mice retained established GFP‐positive ectopic lesions, whereas no detectable GFP‐positive ectopic lesions were observed in DT‐treated mice (Figure  S4 ). These findings demonstrate effective ablation of detectable donor‐derived ectopic lesions following DT treatment. Effects of ectopic lesion ablation on endometriosis‐associated behavioral alterations and pelvic adhesions. (A) Experimental schematic illustrating the study design used to evaluate the effects of ectopic lesion ablation on functional behavioral outcomes and persistent pelvic adhesions. (B) Quantification of nesting scores in vehicle‐treated sham, DT‐treated sham, vehicle‐treated endometriosis (Vehicle‐Eosis), and DT‐treated endometriosis (DT‐Eosis) groups. (C) Quantification of weighted marble‐burying scores in the four experimental groups. Each marble was scored as 1 if fully buried, 0.5 if partially buried, and 0 if not buried. (D) Representative bright‐field and fluorescence (mT and mG) images demonstrating GFP‐positive donor‐derived ectopic lesions in vehicle‐treated mice and the absence of detectable GFP‐positive ectopic lesions following DT treatment. (E) Semiquantitative pelvic adhesion scores assessed using a predefined 0–3 macroscopic scoring system. Individual data points represent biologically independent mice, and data are presented as mean ± SEM. Adhesion scoring was performed by a single observer blinded to the experimental groups; n  = 6 per sham group and n  = 9 per endometriosis group. * p  < 0.05, ** p  < 0.01, *** p  < 0.001 (Kruskal–Wallis test followed by Dunn's multiple‐comparisons test for nesting and adhesion scores; one‐way ANOVA followed by Tukey's multiple‐comparisons test for marble‐burying scores). Eosis, endometriosis. To evaluate behavioral alterations associated with endometriosis and ectopic lesion ablation, we employed two non‐evoked behavioral assays, nesting and marble‐burying. The nesting test results showed that vehicle‐treated mice with endometriosis displayed significantly lower nesting scores (3.44 ± 0.24) compared with vehicle‐treated sham controls (4.67 ± 0.33, p  < 0.05) and DT‐treated sham controls (4.83 ± 0.16, p  < 0.01), indicating impaired nesting performance following endometriosis development (Figure  2B ). Nesting scores did not differ significantly between vehicle‐ and DT‐treated sham mice ( p  = 0.9797), indicating that DT treatment alone did not affect nesting performance. In mice with endometriosis, nesting scores also did not differ significantly between DT‐treated (4.33 ± 0.29) and vehicle‐treated (3.44 ± 0.24) mice ( p  = 0.077). The marble‐burying test assesses digging and burying behavior, with each marble scored as 1 if fully buried, 0.5 if partially buried, and 0 if not buried [ 22 ]. Vehicle‐ and DT‐treated mice with endometriosis exhibited significantly lower marble‐burying scores (3.28 ± 0.81 and 4.44 ± 1.07, respectively) compared with both vehicle‐treated sham controls (9.00 ± 1.32, p  < 0.01 and p  < 0.01, respectively) and DT‐treated sham controls (11.25 ± 1.07, p  < 0.001 and p  < 0.01, respectively). Marble‐burying scores did not differ significantly between vehicle‐ and DT‐treated sham mice ( p  = 0.5426), indicating that DT treatment alone did not affect marble burying performance. Similarly, marble‐burying scores did not differ significantly between DT‐ and vehicle‐treated mice with endometriosis ( p  = 0.8274) (Figure  2C ). These results indicate that endometriosis development is associated with impaired nesting and marble‐burying performance. Following ectopic lesion ablation, nesting performance was not significantly different from either sham controls or vehicle‐treated mice with endometriosis, whereas marble‐burying remained significantly reduced relative to sham controls, indicating variable behavioral responses to lesion ablation. We next evaluated whether pre‐existing pelvic adhesions persisted following ectopic lesion ablation. Because Pgr ‐Cre activity is not restricted to the uterus [ 21 ], we further evaluated potential off‐target ablation in recipient reproductive and endocrine tissues. Fluorescence analysis of the abdomen, ovary, uterus, mammary gland, and pituitary showed no detectable differences between vehicle‐ and DT‐treated recipient mice (Figure  S5 ), consistent with the Cre‐negative genotype of the recipients and the restriction of DT‐sensitive recombined cells to donor‐derived tissues in this model. Having confirmed effective ablation of detectable donor‐derived ectopic lesions, we next determined whether pre‐existing pelvic adhesions persisted following lesion ablation (Figure  2D ). Vehicle‐treated mice with persistent ectopic lesions exhibited severe pelvic adhesions associated with ectopic lesion sites, whereas sham control mice showed no evidence of pelvic adhesion. Notably, adhesions were also observed in DT‐treated mice despite the absence of detectable GFP‐positive ectopic lesions. For quantitative analysis, pelvic adhesions were evaluated using a standardized macroscopic scoring system from 0 (no adhesion) to 3 (severe adhesion), based on thickness, extent, and resistance to separation (Figure  S1 ). Semi‐quantitative scoring revealed persistently high adhesion scores in both vehicle‐treated (2.33 ± 0.33) and DT‐treated (2.00 ± 0.44) mice, whereas sham control mice consistently exhibited a score of 0 (Figure  2E ). These results indicate that ectopic lesion ablation does not resolve established pelvic adhesions, and that adhesion‐associated tissue persists despite the absence of detectable GFP‐positive ectopic lesions. To assess the impact of ectopic lesion ablation on molecular features of the persistent adhesion‐associated tissues, we first analyzed PGR and pSTAT3 expression in eutopic uteri from sham mice, ectopic lesions from vehicle‐treated mice, and adhesion‐associated tissues from DT‐treated mice. Stromal PGR expression was robustly detected in sham uterine tissues (177.05 ± 18.31), whereas significantly reduced expression was observed in vehicle‐treated ectopic lesions (90.91 ± 9.90, p  < 0.01) and was further reduced in adhesion‐associated tissues from DT‐treated mice (8.88 ± 3.89, p  < 0.001) (Figure  3A,B ). Similarly, epithelial PGR expression was strongly detected in sham uteri (261.59 ± 11.40) but was significantly reduced in ectopic lesions from vehicle‐treated mice (160.41 ± 9.93, p  < 0.001). No identifiable endometrial epithelial compartment remained within adhesion‐associated tissues following DT‐mediated ectopic lesion ablation. Therefore, epithelial PGR expression was not assessable by H‐score quantification in this group. These findings demonstrate marked PGR dysregulation in ectopic lesions and loss of the identifiable PGR‐positive endometrial‐like compartment following DT‐mediated ablation. PGR and pSTAT3 expression in ectopic lesions and adhesion‐associated tissues following ectopic lesion ablation. (A) Representative immunohistochemical images of progesterone receptor (PGR) expression in eutopic uteri from sham mice, ectopic lesions from vehicle‐treated mice, and adhesion‐associated tissues from DT‐treated mice. (B) Quantification of PGR H‐scores in stromal and epithelial compartments. Epithelial PGR expression was not assessable by H‐score quantification in adhesion‐associated tissues from DT‐treated mice because no identifiable endometrial epithelial compartment remained following ectopic lesion ablation. (C) Representative immunohistochemical images of pSTAT3 expression in eutopic uteri from sham mice, ectopic lesions from vehicle‐treated mice, and adhesion‐associated tissues from DT‐treated mice. (D) Quantification of epithelial pSTAT3 H‐scores. Epithelial pSTAT3 expression was not assessable by H‐score quantification in adhesion‐associated tissues from DT‐treated mice because no identifiable endometrial epithelial compartment remained following ectopic lesion ablation. Data are presented as mean ± SEM. n  = 6 per group. ** p  < 0.01, *** p  < 0.001 (unpaired two‐tailed Student's t ‐test for epithelial PGR and pSTAT3 comparisons; one‐way ANOVA followed by Tukey's multiple‐comparisons test for stromal PGR comparisons). Because STAT3 phosphorylation is associated with inflammatory signaling in endometriosis, we next examined pSTAT3 expression across experimental groups. pSTAT3 levels were significantly increased in epithelial cells of ectopic lesions from vehicle‐treated mice (148.80 ± 15.02, p  < 0.001) compared with sham controls (12.36 ± 5.64) (Figure  3C,D ). Following DT‐mediated ectopic lesion ablation, adhesion‐associated tissues contained no identifiable endometrial epithelial compartment. Therefore, epithelial pSTAT3 expression was not assessable by H‐score quantification in this group. These results indicate that adhesion‐associated tissues persist following effective ablation of the donor‐derived ectopic epithelial compartment. To assess fibrotic remodeling in the eutopic uterus, ectopic lesions, and adhesion‐associated tissues, we conducted histopathologic analyses and examined fibrosis‐related markers in eutopic uteri from vehicle‐ and DT‐treated sham mice, eutopic uteri and ectopic lesions from vehicle‐treated mice with endometriosis, and eutopic uteri and adhesion‐associated tissues from DT‐treated mice without endometriosis. Vehicle‐ and DT‐treated sham control mice exhibited intact uterine architecture with well‐organized endometrial and myometrial layers and no evidence of pelvic adhesions. In contrast, vehicle‐treated mice displayed severe pelvic adhesions characterized by dense attachment of ectopic lesions to adjacent organs, including the uterus and intestine, accompanied by marked distortion of tissue architecture (Figure  4A ). Notably, in DT‐treated mice, uterine tissues remained firmly adherent to adjacent organs, despite the absence of detectable GFP‐positive ectopic lesions. Molecular characterization of adhesion‐associated tissues following ectopic lesion ablation. (A) Representative hematoxylin and eosin (H&E)‐stained sections showing the eutopic uterus and adjacent tissue from vehicle‐ and DT‐treated sham mice, the eutopic uterus and ectopic lesions from vehicle‐treated mice with endometriosis, and the eutopic uterus and adhesion‐associated tissues from DT‐treated mice without endometriosis. Relevant histological regions are indicated in the images. (B) Representative Masson's trichrome‐stained sections from the indicated tissue groups, highlighting collagen deposition and fibrotic remodeling. (C) Quantification of fibrosis based on Masson's trichrome staining. (D) Representative immunohistochemical images of collagen Type I alpha 1 (COL1A1) expression across the indicated experimental groups. (E) Quantification of COL1A1 H‐scores. (F) Representative immunohistochemical images of α‐SMA expression across the indicated experimental groups. (G) Quantification of α‐SMA H‐scores. Identifiable myometrial‐like structures were excluded from quantitative Masson's trichrome, COL1A1, and α‐SMA analyses of ectopic lesion‐associated fibrotic regions. Data are presented as mean ± SEM. n  = 6 per group. * p  < 0.05, ** p  < 0.01, *** p  < 0.001 (one‐way ANOVA followed by Tukey's multiple‐comparisons test). Next, MTS was performed to assess fibrotic remodeling in eutopic uteri, ectopic lesions from vehicle‐treated mice, and adhesion‐associated tissues from DT‐treated mice (Figure  4B,C ). Because ectopic lesions were generated from whole uterine fragments, occasional myometrial‐like structures were present. These identifiable regions were excluded from quantitative analysis. The fibrosis index was low in vehicle‐ and DT‐treated sham uteri (1.33 ± 1.30 and 2.07 ± 2.04, respectively). The fibrosis index in the eutopic endometrium of vehicle‐treated mice (18.08 ± 6.78) did not differ significantly from that of sham controls ( p  = 0.6718). In contrast, fibrosis was significantly increased in ectopic lesions from vehicle‐treated mice with endometriosis (162.07 ± 8.07, p  < 0.001) compared with sham controls. Following ectopic lesion ablation, the fibrosis index in the eutopic endometrium of DT‐treated mice (0.18 ± 0.06) did not differ significantly from that of DT‐treated sham controls ( p  = 0.5947). Although the fibrosis index in the eutopic endometrium was numerically lower in DT‐treated mice than in vehicle‐treated mice with endometriosis, this difference was not statistically significant. Adhesion‐associated tissues in DT‐treated mice showed a significantly lower fibrosis index (95.39 ± 16.16, p  < 0.001) compared with ectopic lesions from vehicle‐treated mice. However, substantial fibrotic areas persisted within adhesion‐associated tissues despite ectopic lesion ablation. To further characterize extracellular matrix deposition and α‐SMA‐positive fibrotic remodeling, we examined collagen Type I alpha 1 (COL1A1) and α‐SMA expression by immunohistochemistry. COL1A1 expression was low in the eutopic endometrium of vehicle‐ and DT‐treated sham mice (0.50 ± 0.38 and 0.34 ± 0.23, respectively) and was significantly increased in both the eutopic endometrium (56.07 ± 9.75, p  < 0.001) and ectopic lesions (92.38 ± 11.30, p  < 0.001) from vehicle‐treated mice compared with vehicle‐ and DT‐treated sham mice (Figure  4D,E ). COL1A1 levels in the eutopic endometrium of DT‐treated mice (23.50 ± 4.92) were significantly lower than those in vehicle‐treated mice (56.07 ± 9.75; p  < 0.05). However, COL1A1 expression in the DT‐treated eutopic endometrium remained numerically higher than that in DT‐treated sham controls (23.50 ± 4.92 vs. 0.34 ± 0.23), although this difference was not statistically significant. Similarly, COL1A1 levels in adhesion‐associated tissues from DT‐treated mice (46.93 ± 7.02) were significantly lower than those in ectopic lesions from vehicle‐treated mice (92.38 ± 11.30; p  < 0.001). However, substantial COL1A1 expression persisted within adhesion‐associated regions following ectopic lesion ablation. Together, these findings indicate that ectopic lesion ablation was associated with significantly lower eutopic COL1A1 expression, whereas the numerical decrease in the eutopic fibrosis index was not statistically significant. Because recipient mice were Cre‐negative, the reduction in eutopic COL1A1 expression is consistent with the possibility that eutopic fibrotic remodeling is influenced by the presence of ectopic lesions rather than by direct DT‐mediated ablation of recipient uterine cells. α‐SMA is expressed by activated myofibroblasts involved in extracellular matrix deposition and tissue contractility [ 26 ]. Therefore, identifiable myometrial‐like regions were excluded when assessing α‐SMA expression in ectopic lesion‐associated fibrotic regions. Following exclusion of these regions, α‐SMA expression was significantly higher in ectopic lesions from vehicle‐treated mice (67.05 ± 4.93, p  < 0.001) and remained elevated in adhesion‐associated tissues in DT‐treated mice (30.78 ± 3.27, p  < 0.001) compared with vehicle‐ and DT‐treated sham controls (3.22 ± 1.64 and 3.28 ± 1.69, respectively) (Figure  4F,G ). Importantly, α‐SMA expression in adhesion‐associated tissues from DT‐treated mice was significantly lower than that in ectopic lesions from vehicle‐treated mice ( p  < 0.001). Collectively, these findings demonstrate that collagen‐rich, α‐SMA‐positive fibrotic remodeling persists within adhesion‐associated tissues despite effective ablation of detectable donor‐derived PGR‐expressing ectopic cells.

In this study, we show that DT‐mediated inducible ablation of donor‐derived Pgr ‐expressing endometrial epithelial and stromal cells in a syngeneic mouse model of endometriosis effectively eliminated detectable GFP‐positive ectopic lesions. Using this temporally controlled ablation approach, we found that nesting and marble‐burying performance did not differ significantly between DT‐ and vehicle‐treated mice with endometriosis. Although DT‐mediated ablation attenuated fibrosis in adhesion‐associated tissues relative to established ectopic lesions, pelvic adhesions persisted despite effective ectopic lesion ablation. These findings demonstrate that adhesion‐associated fibrotic remodeling persists after elimination of detectable donor‐derived ectopic lesions, whereas behavioral outcomes showed variable responses without a significant, uniform reversal following lesion ablation. Endometriosis is associated with extensive tissue remodeling, including inflammation, fibrosis, and adhesion formation, which are important features of disease progression and may contribute to chronic symptoms and functional impairment [ 27 ]. Fibrosis and adhesions are predominant features of endometriosis [ 28 ]. Although ectopic endometrium can initiate inflammation, the downstream fibrotic and adhesive remodeling is thought to characterize the chronic phenotype [ 29 ]. Transcriptomic profiling in a mouse endometriosis model has further supported this concept, showing consistent upregulation of pathways related to estradiol signaling, inflammation, angiogenesis, and fibrosis in ectopic lesions compared with eutopic endometrium [ 30 ]. Clinically, endometriosis‐associated pain is rarely fully eliminated after hormonal suppression or surgical excision of visible lesions [ 16 , 31 , 32 ]. Consistent with these observations, the behavioral effects of ectopic lesion ablation were variable across the two assays: nesting performance was not significantly different from sham controls, whereas marble‐burying remained significantly impaired relative to sham controls. In parallel, pelvic adhesions and collagen‐rich fibrotic remodeling persisted at prior lesion sites. These findings extend prior work implicating fibrosis and adhesions in chronic endometriosis‐associated tissue remodeling [ 27 ]. Dense and vascular adhesions have been reported to correlate more strongly with pain, consistent with the concept that involvement of pain‐sensitive structures and traction on adjacent organs can increase symptom severity [ 33 ]. The persistence of adhesions after selective ectopic lesion ablation in our model further supports the view that adhesions can be maintained independently of ongoing epithelial lesion activity, reinforcing the multifactorial nature of pain and tissue remodeling in endometriosis. The real‐world impact of endometriosis‐associated pain on social functioning is substantial. Clinically, patients with endometriosis demonstrate increased rates of activity limitation and absenteeism from school or work [ 34 , 35 ]. Consistent with these functional impairments, mice with endometriosis in our study exhibited reduced nesting and marble‐burying performance, which reflect motivation, well‐being, and affective or functional state. Because these assays are not specific measures of nociception, they should be interpreted as complementary readouts of endometriosis‐associated functional and affective impairment rather than direct measures of pain. A randomized study in human patients demonstrated that laparoscopic excision of endometriosis was more effective than placebo surgery in reducing pain and improving quality of life over 12 months [ 32 ]. Notably, approximately 20% of women did not experience pain improvement despite excision [ 32 ]. Surgical management can remove visible ectopic lesions, restore normal pelvic anatomy, and improve endometriosis‐associated pain [ 36 ]. Despite this, a substantial proportion of women continue to experience persistent pain after surgery; a systematic review reported that 11.8% of women who had laparoscopic excision surgery experienced no improvement in pain [ 37 ]. Adhesions have also been associated with increased pain levels [ 28 ]. In deep infiltrating endometriosis, which is often associated with more severe pain than superficial or ovarian endometriosis, adhesiolysis is frequently required [ 38 , 39 ]. Effective adhesion prevention during deep infiltrating endometriosis surgery can improve postoperative pain outcomes, including cycle‐independent pelvic pain and dysmenorrhea [ 40 ]. An important consideration is the biological nature of the adhesion‐associated tissues that persist following ectopic lesion ablation. Histological and immunohistochemical analyses showed that these tissues were collagen‐rich and contained α‐SMA‐positive fibrotic regions, but lacked an identifiable endometrial epithelial compartment and showed markedly reduced PGR expression. Thus, we do not interpret these persistent tissues as residual active ectopic lesions. Rather, they likely represent fibrotic and adhesive tissue remodeling that developed within and around established ectopic lesions and persisted after ablation of detectable donor‐derived PGR‐expressing ectopic cells. However, the present study does not establish the cellular origin of these persistent fibrotic cells. Potential contributors include recipient‐derived fibroblasts recruited to the lesion microenvironment, donor‐derived PGR‐negative stromal or perivascular cells that were not targeted by DT, and recruited immune cells involved in extracellular matrix remodeling. Lineage‐tracing studies combined with cell‐type‐specific markers will be required to distinguish among these possibilities. Similarly, the molecular pathways responsible for maintaining this residual fibrotic compartment remain to be determined. Analysis of profibrotic signaling pathways, including TGF‐β/SMAD and CTGF signaling, will be important in future studies to define the mechanisms underlying persistent adhesion‐associated fibrosis after ectopic lesion ablation. The DT‐mediated ablation model also differs fundamentally from surgical excision of ectopic lesions in patients. DT treatment selectively eliminates donor‐derived PGR‐expressing ectopic cells without physically removing the surrounding tissue, allowing us to examine whether pre‐existing adhesion‐associated fibrotic remodeling persists after elimination of the ectopic cellular component. In contrast, surgical excision physically removes visible lesions together with variable amounts of surrounding fibrotic and adhesive tissue. Moreover, surgical excision itself causes tissue injury and initiates a wound‐healing response that may promote additional inflammation, extracellular matrix deposition, fibrosis, and adhesion formation. Thus, our model should not be considered equivalent to surgical lesion excision but instead provides a complementary experimental approach for distinguishing the consequences of ectopic cell ablation from those associated with physical tissue removal and surgical injury. Our findings provide a framework for interpreting these clinical observations. The variable behavioral responses observed following lesion ablation are consistent with the broader clinical observation that lesion‐directed therapies do not uniformly resolve endometriosis‐associated symptoms. This highlights the complex relationship between lesion burden, tissue remodeling, and functional outcomes. The persistence of adhesions and collagen‐rich fibrosis despite effective ablation of detectable donor‐derived ectopic cells suggests that additional pharmacologic or surgical interventions may need to address established fibrotic and adhesive remodeling to achieve more durable symptom control. Collectively, these results support further investigation of therapeutic strategies that target both ectopic lesions and the surrounding fibrotic and adhesive microenvironment. This study establishes an experimental platform to examine how defined lesion‐associated cell populations contribute to tissue remodeling and functional behavioral outcomes in endometriosis. Although ectopic lesions trigger inflammatory and fibrotic responses in surrounding tissue, the cellular mechanisms linking lesion activity, adhesion formation, fibrosis, and persistent symptoms remain incompletely understood. By enabling temporally controlled ablation of donor‐derived Pgr ‐lineage ectopic cells, this model can be used to examine pathways that connect progesterone resistance with inflammation, extracellular matrix remodeling, and functional behavioral alterations. Future studies can leverage this platform to determine whether targeting fibrosis‐ and adhesion‐associated pathways, alone or as adjuncts to lesion‐directed therapies, improves functional outcomes more effectively than lesion removal alone. Strengths include the use of a genetically controlled Pgr cre/+ Rosa26 iDTR/mTmG donor system combined with Cre‐negative recipients, enabling temporally controlled ablation of donor‐derived Pgr ‐lineage ectopic cells, together with complementary behavioral, histological, and quantitative imaging analyses. An important limitation is that nesting and marble‐burying are not specific measures of nociceptive pain. Nesting reflects general well‐being, motivation, and functional state, whereas marble‐burying can reflect digging, repetitive, and anxiety‐related behaviors. Thus, the behavioral alterations observed in our model should be interpreted as affective and functional impairments associated with endometriosis rather than as direct measures of pain. Future studies incorporating validated evoked pain assays, including abdominal mechanical sensitivity using von Frey testing, together with complementary non‐evoked assays, will be important to directly determine the effects of ectopic lesion ablation on nociceptive pain. Although Cre‐negative recipients restrict DT sensitivity to donor‐derived Pgr‐lineage cells, we cannot completely exclude the possibility that PGR‐expressing donor‐derived immune cells transplanted with the uterine fragments were also ablated by DT and contributed to the observed phenotypes. Therefore, the effects of DT‐mediated ablation cannot be attributed exclusively to donor‐derived endometrial epithelial and stromal cells. In addition, although the syngeneic mouse model recapitulates key features of endometriosis, species‐specific differences in immune and hormonal regulation may limit direct extrapolation to patients. Because the model uses transplanted uterine fragments containing both endometrial and myometrial components, myometrial‐like structures can occasionally be present within established ectopic lesions. Although identifiable myometrial‐like regions were excluded from quantitative histological and immunohistochemical analyses, a contribution of donor‐derived myometrial components to fibrotic remodeling cannot be completely excluded. Importantly, mice do not spontaneously menstruate, and the transplantation‐based model bypasses retrograde menstruation and the natural establishment of ectopic lesions, which should be considered when translating these findings to human endometriosis. Estrous cycles were not synchronized and cycle stage was not determined at the time of tissue collection. In the adult mouse uterus, PGR expression is estrous‐cycle dependent, with distinct patterns of regulation in the epithelial and stromal compartments. Thus, cycle‐dependent hormonal variation may have contributed to inter‐animal variability in PGR, pSTAT3, COL1A1, and other molecular readouts. However, such physiological variation is unlikely to account for the marked loss of PGR‐positive epithelial and stromal cells following DT‐mediated ablation. Similarly, hormonal status may influence STAT3 signaling and COL1A1 expression, although robust pSTAT3 activation was observed in established ectopic lesions relative to the low levels observed in control uterine tissues. Nevertheless, the lack of estrous‐cycle staging should be considered when interpreting inter‐animal variability in these molecular endpoints. The relatively modest cohort sizes used for histological and behavioral analyses may also limit statistical power, particularly for detecting partial differences in behavioral outcomes. In summary, our findings demonstrate that pelvic adhesions and adhesion‐associated fibrotic remodeling persist despite effective ablation of detectable donor‐derived ectopic lesions, whereas behavioral outcomes show variable responses without a significant, uniform reversal. These findings suggest that targeting ectopic lesions alone may be insufficient to resolve established fibrotic and adhesive remodeling and support further investigation of mechanisms that sustain these pathological changes following lesion ablation.

Endometriosis is a chronic gynecological disorder defined by the presence of endometrial‐like tissue outside the uterus and is frequently associated with pelvic pain, infertility, reduced quality of life, and substantial socioeconomic burden [ 1 , 2 , 3 ]. In addition to severe dysmenorrhea, women with endometriosis are at increased risk for other pain conditions, including back pain, headaches or migraines, and joint pain [ 4 ]. Endometriosis‐associated pain is closely linked with anxiety and depression and significantly impairs work productivity and social functioning [ 5 , 6 ]. Pain generation in endometriosis is multifactorial. Ectopic lesions may exert mass effects, adhere to adjacent pelvic organs such as the bowel, and generate a pro‐inflammatory microenvironment through the release of cytokines, prostaglandins, and neurotrophic factors that sensitize nociceptive pathways [ 7 , 8 ]. In parallel, progressive fibrosis and adhesion formation distort pelvic anatomy, generate mechanical traction, and may involve pain‐sensitive structures, further contributing to pain [ 9 ]. However, the relationship between adhesions and pain severity remains controversial. While some studies report a positive correlation between adhesion extent and pain severity [ 10 , 11 ], others find no significant association between pelvic adhesions and pain [ 12 , 13 ]. This inconsistency highlights an incomplete understanding of how lesion‐associated tissue remodeling contributes to pain generation and persistence. Current first‐line management of endometriosis is hormonal suppression [ 14 ]. Therapies such as combined oral contraceptives, progestins, and gonadotropin‐releasing hormone (GnRH) agonists or antagonists reduce ovarian hormone production, suppress lesion activity, and alleviate pain. However, these therapies do not eradicate ectopic lesions, allowing disease activity to persist or recur after treatment discontinuation. Notably, 11%–19% of individuals with endometriosis experience no pain improvement with medical therapy, while 25%–34% experience recurrent pain following treatment cessation [ 15 , 16 ]. These clinical observations underscore the clinical challenge of achieving sustained pain relief and suggest that lesion activity alone may not fully account for persistent symptoms. There is therefore a critical need to identify cellular and molecular mechanisms that drive lesion persistence, tissue remodeling, and pain. Progesterone signaling, mediated by the progesterone receptor (PGR), is essential for endometrial homeostasis through regulation of epithelial differentiation, stromal‐epithelial communication, and cyclical tissue remodeling [ 17 ]. Loss or dysregulation of PGR‐expressing cells is a hallmark of endometriosis and contributes to progesterone resistance in both eutopic endometrium and ectopic lesions, thereby facilitating disease progression [ 18 ]. Despite the recognized role of progesterone resistance in endometriosis pathogenesis, the contribution of donor‐derived Pgr ‐lineage ectopic cells to the maintenance of established lesions and associated tissue remodeling remains poorly understood. Animal models that permit selective and temporally controlled ablation of defined cell populations provide a powerful approach to address this gap. The inducible diphtheria toxin receptor (iDTR) system enables targeted ablation of genetically defined cell populations following diphtheria toxin (DT) administration [ 19 ]. In the present study, we combined the iDTR system with a syngeneic mouse model of endometriosis to selectively ablate donor‐derived Pgr ‐lineage ectopic cells after lesion establishment [ 19 , 20 , 21 ]. This approach allowed us to directly determine whether elimination of established ectopic lesions is sufficient to resolve persistent pelvic adhesions, fibrotic remodeling, and endometriosis‐associated behavioral alterations. We therefore evaluated ectopic lesion ablation, persistent pelvic adhesions, fibrotic remodeling, and functional behavioral outcomes to determine whether elimination of detectable donor‐derived ectopic lesions is sufficient to resolve established disease‐associated tissue and behavioral alterations.

Jae‐Wook Jeong serves as an editorial board member for the FASEB Journal and was recused from any editorial processes during the peer review of this manuscript, including the final decision on potential publication.

All animal experiments were approved by the Animal Care and Use Committee (ACUC) at the University of Missouri under protocol number 65323. Mice were maintained in the designated animal care facility in accordance with ACUC guidelines. Pgr cre/+ mice were crossed with Rosa26 iDTR/iDTR and Rosa26 mTmG/mTmG mice to generate Pgr cre/+ Rosa26 iDTR/mTmG and Rosa26 iDTR/mTmG mice. Eight‐week‐old female Pgr cre/+ Rosa26 iDTR/mTmG and Rosa26 iDTR/mTmG mice were used for all experiments. For endometriosis experiments, Pgr cre/+ Rosa26 iDTR/mTmG mice were used as uterine tissue donors, whereas Cre‐negative Rosa26 iDTR/mTmG mice were used as recipients. To selectively ablate Pgr ‐expressing cells, mice received two intraperitoneal injections of DT (100 ng per mouse per injection; Sigma‐Aldrich, St. Louis, MO, USA) separated by an 8‐h interval. Control mice received vehicle injections (phosphate‐buffered saline; PBS) following the same schedule. Estrous cycles were not synchronized, and estrous‐cycle stage was not determined at the time of euthanasia. At designated experimental time points following DT or vehicle treatment, uteri were carefully dissected, trimmed of surrounding fat and connective tissue, and processed for downstream analyses. For histological and immunohistochemical analyses, uterine tissues were fixed in 4% paraformaldehyde overnight at 4°C, dehydrated, and embedded in paraffin. In vivo fluorescence imaging was performed to assess changes in reporter signal following DT or vehicle treatment. Mice were anesthetized and imaged at 24 and 48 h after treatment. Fluorescence signals were acquired under identical imaging settings for all experimental groups. GFP fluorescence was detected using the AMI HTX imaging system and analyzed with Aura Imaging Software (Spectral Instruments Imaging, Tucson, AZ, USA). Regions of interest (ROIs) were manually drawn around the uterus after tissue collection. The maximum radiance value (photons/s/cm 2 /sr) within each ROI was extracted and used for quantification. Radiance values were calculated after background subtraction using a consistent threshold across all samples. Syngeneic endometriosis was induced by introducing uterine tissue from donor Pgr cre/+ Rosa26 iDTR/mTmG mice into Cre‐negative recipient Rosa26 iDTR/mTmG mice. Both donor and recipient mice were 8‐week‐old adult females. Donor Pgr cre/+ Rosa26 iDTR/mTmG mice received 17β‐estradiol (E2; 0.1 μg per mouse in sesame oil) via subcutaneous injection once daily for three consecutive days before tissue collection, according to our previously validated protocol [ 20 ]. Following E2 priming, one uterine horn was harvested from each donor mouse, minced into small fragments, suspended in 0.5 mL of PBS, and introduced into the peritoneal cavity of one recipient Rosa26 iDTR/mTmG mouse by intraperitoneal injection to establish endometriosis. Thus, tissue derived from one uterine horn was used for each recipient mouse. Sham control mice received an equivalent volume of PBS without uterine tissue by intraperitoneal injection. Recipient mice were allowed to develop ectopic lesions for 1 month following endometriosis induction. Within the sham and endometriosis cohorts, mice were randomly assigned to DT‐ or vehicle‐treatment groups. At 1 month post‐induction, mice received two intraperitoneal injections of DT (100 ng per mouse per injection) or PBS vehicle separated by an 8‐h interval ( n  = 9 per treatment). Sham control mice received the same DT or PBS vehicle treatments ( n  = 6 per treatment) and served as controls. Mice were maintained for an additional month after treatment, after which behavioral assessments were performed, followed by euthanasia and assessment of ectopic lesions under a fluorescence stereomicroscope based on mTmG reporter expression. To assess functional and affective behavioral alterations associated with endometriosis, nesting and marble‐burying tests were performed 1 month after DT or vehicle treatment. These non‐evoked behavioral assays were used as complementary measures of functional and affective impairment and were not interpreted as direct measures of nociception. Nesting and marble‐burying tests were performed to assess goal‐directed behavior and repetitive/digging behavior, respectively [ 22 , 23 , 24 ]. For the nesting test, the mice were individually placed in clean home cages with a single nestlet of cotton tissue (5 × 5 cm, mean weight 2.5 g) 1 h before the dark phase. Nest‐building ability was assessed the next morning according to a 5‐point rating scale from 1 to 5, as previously described by Deacon [ 23 ]. Nest‐building scores were assigned by an observer blinded to the experimental groups. For the marble‐burying test, a Plexiglas cage (25 cm width × 40 cm length × 20 cm height) containing 5 cm‐deep‐fresh, unscented corn cob bedding was used. Mice were individually placed in the Plexiglas cage for 10 min before the test. Mice were then removed from the testing cage and 20 glass marbles (15 mm diameter) were carefully placed equidistant from each other in a 4 × 5 arrangement on the surface of the bedding. Each mouse was placed in a corner of the cage and allowed to bury marbles for 30 min. The marbles were imaged and scored by an observer blinded to the experimental groups. Each marble was assigned a score of 1 when fully buried, 0.5 when partially buried, and 0 when not buried, and individual scores were summed to generate the total marble‐burying score for each mouse [ 24 ]. Pelvic adhesions were evaluated macroscopically at necropsy by a single observer blinded to the experimental groups. Adhesion severity was assessed using a standardized 0–3 scoring system based on adhesion thickness, extent, and resistance to separation: 0, no adhesion; 1, mild adhesion; 2, moderate adhesion; and 3, severe adhesion. The detailed scoring criteria are illustrated in Figure  S1 . In DT‐treated mice, adhesion assessment was performed after confirming the absence of detectable GFP‐positive ectopic lesions to determine whether pre‐existing pelvic adhesions persisted following ectopic lesion ablation. Hematoxylin and eosin (H&E) staining was performed to evaluate tissue morphology. Paraffin‐embedded tissue sections (5‐μm‐thickness) were deparaffinized in xylene and rehydrated through a graded ethanol series followed by rinsing in distilled water. Sections were stained with hematoxylin to visualize nuclei, rinsed in running tap water, and differentiated as needed. After bluing, sections were counterstained with eosin to visualize cytoplasmic and extracellular structures. Following staining, sections were dehydrated through graded ethanol, cleared in xylene, and coverslipped using a permanent mounting medium. Paraffin‐embedded tissue sections were deparaffinized and rehydrated through sequential immersion in xylene and graded alcohol solutions (xylene, 5 min each for three rounds; 100% ethanol, 3 min each for three changes; 95% ethanol, 2 min each for two changes; 70% ethanol, 2 min each for two changes). To block nonspecific binding, sections were incubated with 10% normal goat serum (NGS; Vector Laboratories, Burlingame, CA, USA; #S‐1000‐20) in PBS (pH 7.5) for 2 h at room temperature in a humidified chamber. Primary antibody incubation was performed overnight at 4°C using antibodies against PGR (1:500 dilution, Cell Signaling Technology Inc., Danvers, MA, USA; #8757), COL1A1 (1:500 dilution, Cell Signaling Technology Inc.; #72026), α‐SMA (1:1000 dilution, Abcam, Cambridge, UK; #ab5694), and pSTAT3 (Tyr705) (1:1000 dilution, Cell Signaling Technology Inc.; #9131) diluted in 10% NGS in PBS (pH 7.5). The following day, sections were washed in PBS and incubated with a biotinylated anti‐rabbit IgG secondary antibody (Vector Laboratories, Burlingame, CA, USA; #BA‐9200) for 1 h at room temperature, followed by incubation with an HRP‐streptavidin conjugate (Invitrogen, Waltham, MA, USA; #43–4323; 1:1000) for 45 min. Signal development was performed using the Vectastain Elite DAB kit (Vector Laboratories, Burlingame, CA, USA; #SK‐4100). Sections were counterstained with hematoxylin (Volu‐sol, Sterling, VA, USA; #VMH‐032) followed by 1% lithium carbonate (Poly Scientific, Bay Shore, NY, USA; #S127‐1GL). Slides were dehydrated and coverslipped using an automated Gemini staining system and ClearVue coverslipper (Epredia Kalamazoo, MI, USA). Masson's trichrome staining (MTS) was performed to evaluate tissue fibrosis using a modified one‐step trichrome staining protocol, as previously described [ 25 ]. Paraffin‐embedded tissue sections were deparaffinized in xylene (three changes, 5 min each) and rehydrated through graded ethanol solutions (100%, 95%, and 70%), followed by rinsing in distilled water. Sections were fixed in Bouin's solution (Sigma‐Aldrich, St. Louis, MO, USA) overnight at room temperature to enhance staining intensity and then rinsed thoroughly in distilled water until the tissue appeared colorless. Nuclei were stained using modified Mayer's hematoxylin (Sigma‐Aldrich) for 1 min. Sections were subsequently stained using a one‐step trichrome staining solution (Sigma‐Aldrich) for 1 min, with optimization performed as needed depending on tissue age. After staining, sections were briefly rinsed in distilled water, dehydrated through graded ethanol, cleared in xylene, and coverslipped using Permount mounting medium. Stained sections were used for downstream image acquisition and fibrosis quantification. MTS for all experimental groups was performed in the same staining batch, and images were acquired under identical imaging conditions. Digital image analysis was performed using Visiopharm software (Visiopharm Integrator System, Hoersholm, Denmark), which incorporates artificial intelligence‐based algorithms for quantitative tissue analysis. For immunohistochemical quantification, epithelial and stromal compartments were manually delineated using user‐drawn regions of interest (ROIs) based on tissue morphology. The deep‐learning‐based Nuclear Detection, AI App (Visiopharm, Hoersholm, Denmark; #10169) was optimized using manually annotated representative sections from this study to ensure accurate nuclear segmentation across tissues with different cellularity and architecture. Following optimization, the same nuclear detection and classification criteria were applied consistently across all experimental groups. Representative quality‐control images demonstrating nuclear segmentation in eutopic, ectopic, and adhesion‐associated tissues are shown in Figure  S2 . More than 500 epithelial cells and 1000 stromal cells were analyzed per sample within these defined ROIs. The same anatomical and morphological criteria for ROI selection were applied consistently across all experimental groups to minimize selection bias. Following nuclear detection, nuclei were classified into staining intensity categories: weak (+1), moderate (+2), and strong (+3). H‐scores were calculated using the formula: H‐score = (1 × % weak) + (2 × % moderate) + (3 × % strong), yielding a range of 0–300. Cytoplasmic staining intensity was quantified using a separate area‐based DAB H‐score approach. Whole‐slide images were acquired under standardized bright‐field conditions and analyzed using a custom‐built Visiopharm application. ROIs (> 20 000 μm 2 ) were manually selected to include relevant tissue compartments while excluding artifacts and background. Cytoplasmic DAB signal was classified into four intensity categories (negative, +1 low, +2 medium, and +3 high), and the percentage of stained area within each ROI was calculated. The area‐based H‐score was computed using the formula: H‐score = (1 × % low) + (2 × % medium) + (3 × % high). Fibrosis quantification was performed on Masson's trichrome‐stained sections using Visiopharm image analysis. All images were analyzed under identical illumination and color‐balance settings to minimize batch effects. Collagen‐rich regions were identified using threshold‐based segmentation of the blue channel (RGB range: Red 0–85, Green 0–150, Blue 110–255). ROIs (20000–30 000 μm 2 ) were manually drawn within relevant stromal and fibrotic tissue regions according to predefined anatomical criteria. Because transplanted uterine fragments contained both endometrial and myometrial components, identifiable myometrial‐like structures within ectopic lesions were excluded from ROI selection. Quantitative analyses were focused on fibrotic regions associated with ectopic lesions and adhesion‐associated tissues. For each ROI, mean blue intensity, positive fibrosis area, total ROI area, and integrated intensity (mean blue intensity × positive area) were quantified. A normalized fibrosis index was calculated as: Fibrosis index = Integrated Intensity/ROI Area. Statistical analyses were performed using GraphPad Prism version 10.2.0 (GraphPad Software, San Diego, CA, USA). Comparisons between two independent groups were performed using an unpaired two‐tailed Student's t ‐test. Ordinal nesting and pelvic adhesion scores were analyzed using the Kruskal–Wallis test followed by Dunn's multiple‐comparisons test. For continuous data containing more than two groups, one‐way ANOVA followed by Tukey's multiple‐comparisons test was used. Data are presented as mean ± SEM, and p  < 0.05 was considered statistically significant.

Figure S1: Representative macroscopic images for pelvic adhesion scores. Representative images illustrating the macroscopic adhesion scoring system. Images depict examples of no adhesion (score 0), mild adhesion (score 1), moderate adhesion (score 2), and severe adhesion (score 3). Red boxes indicate the adhesion‐associated regions used to illustrate each score. Figure S2: Quantification of immunohistochemistry using Visiopharm software. (A–C) Representative immunohistochemical images showing manually annotated regions of interest (ROIs) for the luminal epithelium (blue), glandular epithelium (red), and stroma (green) in the eutopic endometrium using Visiopharm software. (D–F) Representative images showing nuclear segmentation and staining‐intensity classification using the Visiopharm Nuclear Detection AI App for H‐score quantification. Nuclear outlines indicate staining intensity as follows: blue, negative (0); yellow, weak (+1); orange, moderate (+2); and red, strong (+3). Figure S3: Assessment of potential DT‐associated toxicity following acute DT treatment. Body weight (A) and uterine weight (B) were assessed at 24 and 48 h after the final vehicle or DT injection in the same experimental cohorts used for the DT‐mediated ablation analysis in Figure 1. Mice received two intraperitoneal injections of DT (100 ng per mouse per injection) or PBS vehicle separated by an 8‐h interval. n  = 5 mice per treatment group at each time point. Individual data points represent biologically independent mice. Data are presented as mean ± SEM. No significant differences were observed between DT‐ and vehicle‐treated mice at either time point. Figure S4: Bright‐field and fluorescence images of ectopic lesions following DT‐mediated ablation in the syngeneic mouse model of endometriosis. Representative bright‐field (BF), merged fluorescence (Texas Red and GFP), and GFP‐only images of the peritoneal cavity from vehicle‐treated (Vehicle‐Sham) and DT‐treated sham mice (DT‐Sham) and mice with endometriosis. GFP‐positive donor‐derived ectopic lesions were detectable in vehicle‐treated mice with endometriosis (Vehicle‐Eosis), whereas no detectable GFP‐positive ectopic lesions were observed in DT‐treated mice with endometriosis (DT‐Eosis). Figure S5: Confirmation of the absence of detectable off‐target ablation in recipient mice. Representative bright‐field and merged fluorescence (Texas Red and GFP) images of the abdomen, ovary, uterus, mammary gland, and pituitary gland from vehicle‐ (Vehicle‐Eosis) and DT‐treated mice with endometriosis (DT‐Eosis) with endometriosis. Pgrcre/+Rosa26 iDTR/mTmG mice were used as positive controls to demonstrate GFP reporter expression in Pgr‐lineage tissues. No detectable differences in GFP fluorescence were observed between vehicle‐ and DT‐treated recipient mice, supporting the absence of detectable DT‐mediated off‐target ablation in the examined recipient tissues.