Estrogen-Progesterone Sequential Exposure Induces the Fibrosis of Adenomyosis in Mice

Adenomyosis (ADS) is a common gynecological disorder characterized by dysmenorrhea, menorrhagia, and infertility, which markedly impair women’s quality of life. Repeated cyclic bleeding of ectopic endometrial lesions may sustain local inflammation and tissue remodeling, leading to a “menstruation—inflammation—fibrosis” cycle. This study aimed to establish a mouse model that mimics this process and to evaluate the effect of estrogen-progesterone (EP) sequential exposure on uterine fibrosis in adenomyosis. Methods: A tamoxifen-induced mouse model of adenomyosis was established, followed by EP sequential exposure to induce a menstruation-like process. Histological staining and immunohistochemistry were performed to evaluate lesion formation, inflammatory responses, and fibrosis-related changes in uterine tissues. Results: EP sequential exposure successfully induced menstruation-like changes in the uterus of mice with adenomyosis. Compared with controls, mice exposed to sequential EP showed more prominent endometrial breakdown, inflammatory responses, and fibrotic remodeling. Collagen deposition and the expression of fibrosis-associated markers, including α-SMA, were increased in uterine lesions, indicating aggravated fibrosis. Conclusion: EP sequential exposure promotes fibrosis in adenomyosis lesions in mice and may serve as a useful model for investigating the “menstruation—inflammation—fibrosis” cycle in adenomyosis. Similar content being viewed by others

Adenomyosis, a prevalent benign gynecological ailment, is typified by the pathological incursion of endometrial glands and stroma into the myometrium [1]. This disorder impacts a substantial portion of females, especially those within their reproductive phase, and is often linked to debilitating manifestations such as menorrhagia, dysmenorrhea, chronic pelvic pain, and infertility [2, 3]. Notwithstanding its clinical prevalence, the pathogenesis of ADS remains inadequately comprehended, which has, in turn, impeded the advancement of efficacious and targeted therapeutic approaches. The pathophysiological mechanisms underlying ADS are considered to be multifactorial, including theories ranging from tissue injury and repair (TIAR) caused by uterine hyperperistalsis to metaplasia of embryonic stem cells [4, 5]. A critical clinical discovery is the substantial correlation between the condition and events that disrupt the endometrial—myometrial interface, such as pregnancy termination, cesarean sections, and other uterine surgical operations. This suggests that repetitive TIAR processes play a central role in its onset and progression [6, 7]. A fundamental element of this injury model is the role of menstruation. In females suffering from ADS, it is hypothesized that retrograde menstruation deposits ectopic endometrial tissue within the myometrium, where it becomes established and proliferates[8]. More importantly, these ectopic lesions undergo cyclic bleeding in response to hormonal fluctuations, maintaining a local inflammatory reaction and subsequent tissue remodeling [9, 10]. A crucial consequence of this cyclical hemorrhage within the ectopic lesions is the accumulation of iron, a vital constituent of hemoglobin derived from extravasated red blood cells [11]. Although iron is indispensable for cellular function, an excess of it can be extremely cytotoxic. In the context of recurrent menstrual bleeding, iron can catalyze the generation of reactive oxygen species (ROS) through the Fenton reaction, resulting in oxidative stress, chronic inflammation, and the activation of fibroblasts [12, 13]. This cascade is a well—recognized driver of fibrogenesis in other organs, such as the liver [14, 15] and kidneys [16, 17]. In the uterus, fibrosis, characterized by the excessive deposition of collagen and other extracellular matrix (ECM) components, is a characteristic feature of advanced ADS, contributing to uterine stiffness, dysfunction, and pain [18, 19]. Nevertheless, the direct association between menstruation—induced iron overload and the exacerbation of fibrosis in ADS has not been systematically explored. To address this knowledge gap, suitable animal models are essential. The administration of tamoxifen (TAM) to neonatal mice has been recognized as a reliable model that reproduces the key characteristics of human ADS, such as the invasion of endometrial tissue into the myometrium [20]. Moreover, recent progress has facilitated the establishment of murine models that mimic the human menstrual cycle through the sequential administration of EP, followed by hormone withdrawal to trigger decidualization and disintegration, simulating a menstruation—like event [21]. Consequently, this research was formulated to clarify the function of iron metabolism in the advancement of ADS, with a particular emphasis on fibrosis. It was hypothesized that simulated menstruation would intensify the fibrotic pathology in ADS, and that this effect is mediated by iron overload. To verify this hypothesis, a tamoxifen—induced ADS model was initially established in mice. Subsequently, an artificial menstrual cycle protocol was implemented on both normal and ADS—affected mice to contrast the physiological repair response with the pathophysiological progression. Finally, the therapeutic potential of DFO, a potent iron chelating agent, in alleviating menstruation—exacerbated fibrosis was explored. The findings of this study offer novel perspectives on the "menstruation—inflammation—fibrosis" cycle in ADS and underscore iron chelation as a promising therapeutic approach.

Animals Ten late—gestational female Institute for Cancer Research (ICR) mice were provided by the Department of Laboratory Animal Science of Peking University Health Science Center (Beijing, China) and utilized for this study. All the mice delivered offspring within three days subsequent to their arrival at the laboratory animal facility. A total of 95 newborn mice were administered tamoxifen to induce ADS. Gender was determined four weeks after birth, and only female mice, amounting to 59 in total, were employed in the experiment. All mice were housed under controlled environmental conditions with a 12/12—hour light/dark cycle and had ad libitum access to food and water. All animal experiments were conducted in accordance with the Guide for the Care and Use of Laboratory Animals and were approved by the Institutional Animal Care and Use Committee (IACUC) of Peking University Health Science Center (Approval No. BCJI0251; approved on September 8, 2023). Experimental Design Among the 59 mice, 12 were euthanized at 6 weeks of age to confirm the success of the modeling. Specifically, 6 mice had tamoxifen-induced ADS, and 6 were treated with peanut oil vehicle only. The remaining 47 female mice were randomly allocated into 4 groups (11—12 mice per group) based on intervention combinations, as present in Table 1. Treatments were administered in stages during the neonatal, pubertal, and adult periods to establish an ADS model that simulates the menstruation—inflammation—fibrosis cycle and to explore the role of iron metabolism in fibrosis. Induction of ADS A well—established murine model of ADS was employed, with the exception of TAM − EP − Group. Oral administration was carried out via capillary tubing from postnatal day 2 to day 5 (with the day of birth defined as day 1) for four consecutive days. The administered dosage was 1 mg/kg of tamoxifen, which was suspended in a mixture of peanut oil, lecithin, and condensed milk at a volume—to—volume ratio of 2:0.2:3. The dosage volume was 5 μl/g of body weight [20, 22]. Randomly selected female neonatal control mice were fed in a similar manner with an equivalent amount of the solvent, excluding tamoxifen. When these female mice attained 4 weeks of age, they were weaned and separated from the dams. Induction of the Artificial Menstrual Cycle Except for TAM − EP −/TAM + EP − Group, all other experimental groups in the present study were subjected to the same murine menstruation model to ensure consistency in the experimental setup. The murine menstruation model utilized in this study was previously described by Cousins et al. [23]. As shown in the flow chart below, one or two weeks subsequent to ovariectomy, mice were administered a daily subcutaneous injection of 100 ng of 17β—estradiol (E2; Sigma—Aldrich) in arachis oil for a duration of 3 days. Following a three—day interval, a combination of E2 and progesterone (P; Sigma—Aldrich) was injected subcutaneously of each mouse for 3 days. The dosages of E2 and P were 10 ng and 1 mg, respectively. On the day of the final combined injection, 100 μl of arachis oil was randomly injected into one uterine horn of each mouse using an indwelling needle device (22G) to induce decidualization. One day after the uterine horn injection, mice received a daily subcutaneous injection of 500 ng of P in arachis oil for 4 days. The experimental design is shown in Fig. 1. Mouse Experiment Protocol In this section, the aforementioned mouse model simulating menstruation was employed to explore the function of iron ions in uterine fibrosis induced by menstruation. Twenty—three female ICR mice were selected for the experiment and randomly assigned to two groups: the TAM + EP + group (control group) and the TAM + EP + DFO group. Seventy-two hours before and at the time of progesterone withdrawal, the mice in the two groups were intraperitoneally (i.p.) injected with physiological saline and DFO (TargetMol, China), respectively. The deferoxamine mesylate was prepared by dissolving 0.06 g of the drug in 1.5 ml of physiological saline, with a daily dosage of 100 mg/kg. The administration dosages were determined based on previously published murine studies [24]. Twenty—four to forty—eight hours subsequent to progesterone withdrawal, all mice were euthanized and their uteri were harvested. The wet mass of the uteri was measured via an electronic balance, and the maximum longitudinal and transverse diameters were ascertained using a vernier caliper. Immunohistochemical analysis was conducted on mouse uterine tissue samples, and Masson's trichrome staining and Sirius red staining were employed to quantitatively evaluate the extent of tissue fibrosis. Hematoxylin–Eosin Staining Tissue specimens were fixed in 4% (weight/volume) paraformaldehyde solution and embedded in paraffin. Serial sections with a thickness of 4 μm were prepared from each paraffin block. Subsequently, the slides underwent routine deparaffinization in xylene and rehydration via a graded alcohol series. The sections were stained with hematoxylin to visualize the cell nuclei and then counterstained with eosin to color the cytoplasm. After dehydration and clearing processes, the slides were mounted using a neutral resin. The first slide from each sample was examined under a light microscope to verify the pathological diagnosis of ADS. All histological evaluations were conducted by an investigator who was unaware of the experimental groups. Immunohistochemistry (IHC) Tissue specimens were fixed in 4% (w/v) paraformaldehyde and then embedded in paraffin. Serial 4-μm sections were prepared from each paraffin block. The initial slide was subjected to hematoxylin—eosin staining to verify the pathological diagnosis. Subsequently, the remaining slides were stained for E—cadherin (specific to epithelial cells), Collagen I, and α—SMA. Additionally, Masson's trichrome staining and Sirius red staining were performed. Routine deparaffinization and rehydration processes were carried out. For antigen retrieval, the slides were heated at 98 °C in an EDTA buffer (pH 9.0) for a cumulative duration of 30 min for staining of E—cadherin, Collagen I, and α—SMA, followed by natural cooling to room temperature. The primary antibodies against E—cadherin, Collagen I, and α—SMA were diluted to ratios of 1:8000, 1:1000, and 1:1000 respectively, and the sections were incubated with the primary antibodies overnight at 4 °C. After the slides were rinsed, the Rabbit Polymer Detection System (Zhongshan Goldenbridge Biotechnology Co., Ltd., China) was incubated at room temperature for 30 min. The bound antibody complexes were stained for 1—3 min or until suitable for microscopic examination using diaminobenzidine, followed by counterstaining with hematoxylin for 8 min. The slides underwent differentiation in hydrochloric acid alcohol and subsequent bluing with tap water. After dehydration and clearing, the slides were mounted. The names of the primary antibodies, along with their vendor names and the concentrations employed in this study, are presented in Table 2. Images were acquired using a microscope equipped with a digital camera (Olympus, Japan). Regarding other immunostained markers, quantification was conducted by randomly selecting 3—5 images per mouse at a magnification of 400 ×. Subsequently, the mean optical density value was obtained via Image J (USA). In the case of negative controls, mouse ectopic endometrium tissue samples were incubated with phosphate—buffered saline (PBS) rather than primary antibodies. To mitigate potential bias, the individual responsible for evaluating the slides was kept unaware of the group to which the slides pertained. Masson’s Trichrome Staining In the process of Masson's trichrome staining, tissue sections were subjected to deparaffinization in xylene and rehydration through a graded alcohol series. Subsequently, staining was carried out using the Masson Trichrome Staining solution (Nanjing Jiancheng Bioengineering Institute, China) in strict accordance with the manufacturer's instructions. The proportion of the area of the collagen fiber layer, stained blue, to the entire field of the ectopic implants was calculated via Image J. Sirius Red Staining For the evaluation of collagen deposition, Sirius red staining was carried out. After deparaffinization and rehydration as previously described, the tissue sections were stained with a Picro—Sirius Red solution (Solarbio, China) in strict compliance with the manufacturer's guidelines. Subsequently, the stained sections were observed under both standard light microscopy and polarized light microscopy. Under polarized light, type I collagen fibers present as orange—red or bright yellow, whereas thinner type III collagen fibers appear green. The ratio of the collagen fiber area (stained red) to the entire field of the ectopic implants was quantified using Image J software. Statistical Analysis All data were expressed as mean ± standard deviation (x̄ ± SD). One—way analysis of variance (One—way ANOVA) and LSD—t multiple comparison were performed using GraphPad Prism 9.0 software. A value of P < 0.05 was considered statistically significant.

Establishment of a Mouse ADS Model Induced by Tamoxifen To explore the pathological process of ADS, a mouse model was initially established through the oral administration of tamoxifen. In comparison with the normal control uteri, which presented as a slender, thread—like structure, exhibiting a pinkish hue and a supple texture, without any manifestations of hyperemia, edema, or petechiae, the uteri of tamoxifen—treated mice showed no evident thickening, with distinct vascular markings (Fig. 2A, Group TAM − EP − and Group TAM + EP −). Moreover, the weight of the tamoxifen—treated uterus was significantly lower than that of the normal control uteri, while no significant difference in body weight was observed between the two groups (290.58 ± 54.98 mg vs. 250.68 ± 30.05 mg, P = 0.0382; 42.30 ± 5.40 g vs. 41.29 ± 4.37 g, P = 0.6200; Fig. 2B/C, Group TAM − EP − and Group TAM + EP −). Histological examination (H&E staining) verified the invasion of endometrial glands and stroma into the myometrial layer, leading to a disrupted uterine architecture Fig. 2D. This successfully established model offered a reliable platform for subsequent investigations on disease progression and intervention. To further assess the fibrotic status at this early stage, Masson’s trichrome and Sirius Red staining were performed. The results revealed that even without subsequent hormonal stimulation, the TAM + EP − group already exhibited a significant increase in collagen deposition compared to the TAM − EP − group, as evidenced by a higher percentage of Masson—positive area (1.06 ± 0.18% vs. 2.01 ± 0.16%, P < 0.0001) and an elevated collagen volume fraction (CVF) (8.32 ± 0.74% vs. 13.37 ± 0.90%, P < 0.0001) (Figs. 2 and 4 A/B, Group TAM − EP − and Group TAM + EP −). Interestingly, at this initial modeling stage, immunohistochemical analysis showed no statistically significant differences between the TAM − EP − and TAM + EP − groups regarding the expression of the epithelial marker E—cadherin (119.69 ± 3.72 vs. 121.33 ± 7.52, P = 0.8212) or the pro—fibrotic markers Collagen I (120.37 ± 1.66 vs. 118.81 ± 2.82, P = 0.5174) and α—SMA (118.64 ± 3.41 vs. 114.72 ± 3.44, P = 0.0609) Figs. 3 and 4C - E. These findings suggest that while tamoxifen induction initiates structural damage and early collagen accumulation, the dramatic molecular dysregulation of epithelial integrity and myofibroblast activation is likely further triggered by the subsequent cycle of simulated menstruation. Simulated Menstruation Aggravates Uterine Fibrosis in Mice with ADS Upon the establishment of the model, we subsequently explored the influence of simulated menstruation on the progression of ADS in mice. Subsequent to sequential EP administration, the uterus of ADS exhibited thickening and congestion, presenting a vivid red hue. The surface blood vessels were thickened, and visible blood clots were observed in the uterine cavity, suggesting the successful induction of periodic withdrawal bleeding (Fig. 2A, TAM + EP + Group). Although there was no significant disparity in body weight between the two groups (41.29 ± 4.37 g vs. 41.62 ± 4.23 g, P = 0.8548; Fig. 2B, Group TAM + EP − and Group TAM + EP +), the uterine weight of the TAM + EP + group was significantly elevated compared to that of the TAM + EP − group (250.68 ± 30.05 mg vs. 279.48 ± 28.11 mg, P = 0.0240; Fig. 2C, Group TAM + EP − and Group TAM + EP +), indicating notable local pathological changes. Histological evaluation indicated a severe fibrotic reaction. Masson's trichrome staining exhibited extensive and dense blue collagen deposition within the myometrium of the TAM + EP + group, which was significantly more severe compared to that in the TAM + EP − controls (6.00 ± 0.19% vs. 2.01 ± 0.16%; P < 0.0001) (Figs. 3 and 4A Group TAM + EP − and Group TAM + EP +). This observation was verified by Sirius red staining, which also manifested a substantial elevation in collagen content (red areas) in the TAM + EP + group (29.07 ± 2.56 vs. 13.37 ± 0.90, P < 0.0001) (Figs. 3 and 4B, Group TAM + EP − and Group TAM + EP +). These outcomes confirm that simulated menstruation significantly aggravates extracellular matrix deposition and fibrosis in the presence of pre—existing ADS. Immunohistochemical analysis unveiled a distinct pathological molecular signature. In contrast to the coordinated repair response observed in normal mice, the expression of the epithelial adhesion protein E—cadherin was notably downregulated in the TAM + EP + group when compared with the TAM + EP − group (P = 0.0077; Figs. 3 and 4C, Group TAM + EP − and Group TAM + EP +). Simultaneously, the expression of the pro—fibrotic markers Collagen I (P = 0.0047) and α—SMA (P = 0.0029) was markedly and significantly upregulated in the TAM + EP + group (Figs. 3 and 4D/E, Group TAM + EP − and Group TAM + EP +). Collectively, these data indicate that simulated menstruation in ADS mice results in pathological exacerbation, which is manifested by notable uterine enlargement, severe collagen deposition, and a dysregulated molecular profile. This profile is characterized by the loss of epithelial integrity (downregulation of E—cadherin) accompanied by a marked pro—fibrotic response (upregulation of Collagen I and α—SMA). Iron Chelation by DFO Attenuates Menstruation—Exacerbated Fibrosis in ADS To explore the functional significance of iron metabolism in the exacerbation of fibrosis, we administered the iron—chelating agent DFO to ADS mice subjected to simulated menstruation (TAM + EP + DFO group). Macroscopically, the distinct uterine enlargement noted in the TAM + EP + group was markedly alleviated through DFO treatment (Fig. 2A, Group TAM + EP + DFO). Although there was no significant disparity in body weight between the two groups (41.65 ± 4.44 g vs. 38.23 ± 4.61 g, P = 0.0801; Fig. 2B, Group TAM + EP + and Group TAM + EP + DFO), the uterine weight in the DFO—treated group was significantly lower compared to that in the untreated TAM + EP + group (277.95 ± 29.60 mg vs. 225.65 ± 29.37 mg, P = 0.0003; Fig. 2C, Group TAM + EP + and Group TAM + EP + DFO), suggesting a local therapeutic efficacy. Histopathological evaluation verified the anti—fibrotic efficacy of DFO. Masson's trichrome staining indicated a significant decline in the scope and intensity of blue—stained collagen fibers within the myometrium of the TAM + EP + DFO group when compared to the TAM + EP + group (Fig. 3A, Group TAM + EP + and Group TAM + EP + DFO). This observable enhancement was further supported by Sirius red staining, which also exhibited a decrease in collagen content (red areas) following DFO treatment (Fig. 3B, Group TAM + EP + and Group TAM + EP + DFO). Quantitative assessments confirmed these findings, revealing a statistically significant reduction in both the fibrotic area determined by Masson's trichrome staining P < 0.0001; Fig. 4A and the collagen volume fraction measured by Sirius red staining P < 0.0001; Fig. 4B in the DFO—treated group. At the molecular level, the TAM + EP + DFO group demonstrated a significantly enhanced immunoreactivity for E—cadherin. A more intense and continuous staining pattern was observed in the endometrial glands (Fig. 3C, Group TAM + EP + DFO). The expression level of E—cadherin in the TAM + EP + DFO group was significantly higher than that in the untreated group (P = 0.0008; Fig. 4C, Group TAM + EP + and Group TAM + EP + DFO). Moreover, treatment with DFO effectively reversed the up—regulation of key fibrotic markers. Immunohistochemical staining revealed that the expression of Collagen I in the TAM + EP + DFO group was significantly lower compared to the TAM + EP + group. Quantitative analysis of the average optical density (AOD) confirmed a substantial reduction in Collagen I expression (P < 0.0001; Figs. 3 and Fig. 4 D, Group TAM + EP + and Group TAM + EP + DFO). Similarly, the expression of α—SMA was significantly decreased by DFO intervention, along with a significant decline in its AOD value (P = 0.0090; Figs. 3 and 4E, Group TAM + EP + and Group TAM + EP + DFO). In conclusion, treatment with DFO effectively alleviated the menstruation—exacerbated fibrotic pathological changes in ADS mice. This was manifested by a decrease in uterine weight, a reduction in histological collagen deposition, and a down—regulation of the expression of pro—fibrotic markers, namely Collagen I and α—SMA. These findings imply that iron chelation represents a potential therapeutic approach for attenuating fibrosis in ADS.

This research offers robust evidence for a crucial pathophysiological axis in ADS, namely the "menstruation—iron—fibrosis" cycle. Through the establishment of a novel murine model that combines neonatal tamoxifen—induced ADS with an artificial menstrual cycle, we have effectively replicated the essential characteristic of recurrent bleeding and iron deposition in ectopic lesions. Our primary discovery is that simulated menstruation notably aggravates uterine fibrosis in ADS, and this aggravation is predominantly propelled by iron overload, a process that can be effectively targeted and reversed via iron chelation. A crucial discovery of this study is the disparate responses to simulated menstruation between normal uteri and those affected by ADS. Human menstruation is a complex and highly regulated physiological phenomenon wherein the endometrium, under the influence of cyclic fluctuations in estrogen and progesterone, undergoes a series of coordinated processes: proliferation, differentiation, disintegration, and subsequent restoration [25]. In contrast to humans, laboratory mice do not undergo spontaneous menstruation. To surmount this significant translational obstacle, our experimental design was guided by established methodologies in the relevant field [21, 26, 27]. This entailed a bilateral ovariectomy to eliminate interference from endogenous ovarian hormones, followed by the sequential administration of exogenous EP to mimic the human menstrual cycle. To simulate the receptive state and decidual transformation of the endometrium prior to menstruation, an intrauterine injection of oil was administered during the peak phase of estrogen—progesterone, thereby inducing decidualization. Subsequently, the withdrawal of progesterone triggered menstruation—like events [26, 27]. This multi—step protocol effectively replicated the key hormonal and morphological transitions of menstruation in mice, offering a valuable means for investigating endometrial reconstruction processes, particularly the dynamics of disintegration and regeneration. Within the framework of this model, extant literature delineates a distinct mechanistic cascade in normal mice subsequent to progesterone withdrawal. The primary trigger instigates the generation of ROS, which subsequently activates the NF—κB signaling pathway. This activation facilitates the expression of cyclooxygenase—2 (COX—2), thereby propelling prostaglandin synthesis, leukocyte recruitment, and ultimately culminating in endometrial disintegration [28]. Crucially, the ensuing repair phase entails a coordinated Mesenchymal—Epithelial transition (MET), which is indispensable for prompt re—epithelialization and the restoration of tissue integrity [23]. It is within this well—established repair window that our experimental observations are situated. Conversely, within adenomyotic uteri, the identical stimulus incited pathological fibrosis. This was substantiated by the down—regulation of E—cadherin, which signified the disruption of epithelial integrity, along with a notable up—regulation of Collagen I and α—SMA. The manifestation of fibrosis in the ADS mouse model aligns with previous findings [29, 30]. Moreover, the menstruation model induced through sequential administration of EP demonstrates more pronounced fibrotic alterations. The underlying mechanisms propelling this enhanced fibrotic response are presumably multifactorial, potentially encompassing hormonal imbalance [31], the EMT—FMT—SMM (EMT: Epithelial—Mesenchymal Transition; FMT: Fibromuscular Transformation; SMM: Smooth Muscle Metaplasia) vicious cycle, the amplified inflammation—fibrosis axis, as well as aberrant angiogenesis and local hypoxia [32]. Beyond the cellular and molecular alterations, the most immediate consequence of inducing menstruation in adenomyotic mice is the cyclic bleeding within ectopic lesions, leading to the degradation of extravasated red blood cells and subsequent iron accumulation. While the pathogenic role of iron in ADS has not been extensively documented, compelling evidence from endometriosis research confirms significant iron deposition surrounding ectopic endometrium, with iron levels directly correlating with the severity of fibrosis [33]. Crucially, iron overload is a well—established driver of fibrogenesis in endometriosis. Excess iron catalyzes the generation of ROS via the Fenton reaction, promoting oxidative stress and chronic inflammation, which in turn activates the pivotal TGF—β1 signaling pathway, a master regulator of collagen production and fibroblast—to—myofibroblast differentiation [11, 34]. The most direct evidence supporting a causal role for iron in our model comes from our therapeutic intervention. Administration of the iron chelator DFO significantly attenuated the menstruation-exacerbated fibrotic pathology. This striking anti—fibrotic effect, demonstrated by reduced collagen deposition and downregulation of pro—fibrotic markers, strongly implies that iron-driven pathways are instrumental in the progression of ADS—related fibrosis. Despite these promising findings, this study is constrained by limitations that chart a clear path for future research. The current observations are based on a single artificial menstrual cycle; the cumulative impact of long—term, recurrent bleeding on fibrosis progression and potential plateau phases warrants exploration within a long—term multi—cycle cohort. Secondly, direct quantification of tissue iron concentrations and ROS levels was not conducted, which would enhance mechanistic interpretations. Future studies should utilize Prussian blue staining and malondialdehyde assays to verify local iron overload and oxidative stress. Additionally, the cell—specific contributions of macrophages and fibroblasts, especially their expression of iron—handling proteins such as ferritin and ferroportin, need to be clarified through single—cell sequencing or lineage tracing. Moreover, although the doses of DFO were effective, they were determined based on acute toxicity pre—tests and lacked a comprehensive dose—response curve. Future investigations should incorporate gradient dose optimization and compare systemic and intrauterine localized drug delivery in terms of efficacy and safety.

In summary, this research posits iron overload as a pivotal mechanistic connection between menstruation and fibrosis in ADS. Through the introduction of a physiologically relevant model and the demonstration of the effectiveness of iron—targeted interventions, we not only enhance the comprehension of ADS pathogenesis but also uncover a promising novel therapeutic approach intended to disrupt the pernicious "menstruation—inflammation—fibrosis" cycle. Data Availability The datasets generated and/or analyzed during the current study are available from the first author on reasonable request. Code Availability Not applicable. Abbreviations - α-SMA: - Alpha-smooth muscle actin - ADS: - Adenomyosis - CVF: - Collagen volume fraction - COX-2: - Cyclooxygenase-2 - DFO: - Deferoxamine mesylate - E2: - 17β-Estradiol - ECM: - Extracellular matrix - EMT: - Epithelial–mesenchymal transition - EP: - Estrogen and progesterone - FMT: - Fibromuscular transformation - IACUC: - Institutional Animal Care and Use Committee - ICR: - Institute for Cancer Research - IHC: - Immunohistochemistry - i.p.: - Intraperitoneally - MET: - Mesenchymal–epithelial transition - P4: - Progesterone - PBS: - Phosphate-buffered saline - ROS: - Reactive oxygen species - SMM: - Smooth muscle metaplasia - TAM: - Tamoxifen - TIAR: - Tissue injury and repair

Bird CC, McElin TW, Manalo-Estrella P. The elusive adenomyosis of the uterus-revisited. Am J Obstet Gynecol. 1972;112:583–93. https://doi.org/10.1016/0002-9378(72)90781-8. Chu LH, Liao CC, Liew PL, et al. Epigenomic analysis reveals the KCNK9 potassium channel as a potential therapeutic target for adenomyosis. Int J Mol Sci. 2022. https://doi.org/10.3390/ijms23115973. Gordts S, Grimbizis G, Campo R. Symptoms and classification of uterine adenomyosis, including the place of hysteroscopy in diagnosis. Fertil Steril. 2018;109(3):380-388.e1. https://doi.org/10.1016/j.fertnstert.2018.01.006. Guo SW. Cracking the enigma of adenomyosis: an update on its pathogenesis and pathophysiology. Reproduction. 2022;164(5):R101–21. https://doi.org/10.1530/REP-22-0224. García-Solares J, Donnez J, Donnez O, et al. Pathogenesis of uterine adenomyosis: invagination or metaplasia? Fertil Steril. 2018;109(3):371–9. https://doi.org/10.1016/j.fertnstert.2017.12.030. Leyendecker G, Wildt L. A new concept of endometriosis and adenomyosis: tissue injury and repair (TIAR). Horm Mol Biol Clin Investig. 2011;5(2):125–42. https://doi.org/10.1515/HMBCI.2011.002. Leyendecker G, Wildt L, Mall G. The pathophysiology of endometriosis and adenomyosis: tissue injury and repair. Arch Gynecol Obstet. 2009;280(4):529–38. https://doi.org/10.1007/s00404-009-1191-0. Chen YJ, Li HY, Huang CH, et al. Oestrogen-induced epithelial-mesenchymal transition of endometrial epithelial cells contributes to the development of adenomyosis. J Pathol. 2010;222(3):261–70. https://doi.org/10.1002/path.2761. Kobayashi H, Kishi Y, Matsubara S. Mechanisms underlying adenomyosis-related fibrogenesis. Gynecol Obstet Invest. 2019;85(1):1–12. https://doi.org/10.1159/000502822. Bourdon M, Santulli P, Jeljeli M, et al. Immunological changes associated with adenomyosis: a systematic review. Hum Reprod Update. 2021;27(1):108–29. https://doi.org/10.1093/humupd/dmaa038. Wyatt J, Fernando SM, Powell SG, et al. The role of iron in the pathogenesis of endometriosis: a systematic review. Hum Reprod Open. 2023;2023(3):hoad033. https://doi.org/10.1093/hropen/hoad033. Defrère S, Lousse JC, González-Ramos R, et al. Potential involvement of iron in the pathogenesis of peritoneal endometriosis. Mol Hum Reprod. 2008;14(7):377–85. https://doi.org/10.1093/molehr/gan033. Maus M, López-Polo V, Mateo L, et al. Iron accumulation drives fibrosis, senescence and the senescence-associated secretory phenotype. Nat Metab. 2023;5(12):2111–30. https://doi.org/10.1038/s42255-023-00928-2. Chen J, Li X, Ge C, et al. The multifaceted role of ferroptosis in liver disease. Cell Death Differ. 2022;29(3):467–80. https://doi.org/10.1038/s41418-022-00941-0. Allameh A, Niayesh-Mehr R, Aliarab A, et al. Oxidative stress in liver pathophysiology and disease. Antioxidants. 2023;12(9):1653. https://doi.org/10.3390/antiox12091653. Patera F, Gatticchi L, Cellini B, et al. Kidney fibrosis and oxidative stress: from molecular pathways to new pharmacological opportunities. Biomolecules. 2024;14(1):137. https://doi.org/10.3390/biom14010137. Lv W, Booz GW, Fan F, et al. Oxidative stress and renal fibrosis: recent insights for the development of novel therapeutic strategies. Front Physiol. 2018;9:105. https://doi.org/10.3389/fphys.2018.00105. Vannuzzi V, Bernacchioni C, Muccilli A, et al. Sphingosine 1-phosphate pathway is dysregulated in adenomyosis. Reprod Biomed Online. 2022;45(1):15–8. https://doi.org/10.1016/j.rbmo.2022.03.026. Huang Q, Liu X, Critchley H, et al. How does the extent of fibrosis in adenomyosis lesions contribute to heavy menstrual bleeding? Reprod Med Biol. 2022;21(1):e12442. https://doi.org/10.1002/rmb2.12442. Green AR, Styles JA, Parrott EL, et al. Neonatal tamoxifen treatment of mice leads to adenomyosis but not uterine cancer. Exp Toxicol Pathol. 2005;56(4–5):255–63. https://doi.org/10.1016/j.etp.2004.10.001. Finn CA, Pope M. Vascular and cellular changes in the decidualized endometrium of the ovariectomized mouse following cessation of hormone treatment: a possible model for menstruation. J Endocrinol. 1984;100(3):295–300. https://doi.org/10.1677/joe.0.1000295. Parrott E, Butterworth M, Green A, et al. Adenomyosis - a result of disordered stromal differentiation. Am J Pathol. 2001;159(2):623–30. https://doi.org/10.1016/S0002-9440(10)61733-6. Cousins FL, Murray A, Esnal A, et al. Evidence from a mouse model that epithelial cell migration and mesenchymal-epithelial transition contribute to rapid restoration of uterine tissue integrity during menstruation. PLoS ONE. 2014;9(1):e86378. https://doi.org/10.1371/journal.pone.0086378. Dongiovanni P, Valenti L, Ludovica Fracanzani A, et al. Iron depletion by deferoxamine up-regulates glucose uptake and insulin signaling in hepatoma cells and in rat liver. Am J Pathol. 2008;172(3):738–47. https://doi.org/10.2353/ajpath.2008.070097. Garry R, Hart R, Karthigasu KA, et al. A re-appraisal of the morphological changes within the endometrium during menstruation: a hysteroscopic, histological and scanning electron microscopic study. Hum Reprod. 2009;24(6):1393–401. https://doi.org/10.1093/humrep/dep036. Brasted M, White CA, Kennedy TG, et al. Mimicking the events of menstruation in the murine uterus. Biol Reprod. 2003;69(4):1273–80. https://doi.org/10.1095/biolreprod.103.016550. Milligan SR, Cohen PE. Silastic implants for delivering physiological concentrations of progesterone to mice. Reprod Fertil Dev. 1994;6(2):235–9. https://doi.org/10.1071/rd9940235. Wu B, Chen X, He B, et al. ROS are critical for endometrial breakdown via NF-κB-COX-2 signaling in a female mouse menstrual-like model. Endocrinology. 2014;155(9):3638–48. https://doi.org/10.1210/en.2014-1029. Shen M, Liu X, Zhang H, et al. Transforming growth factor β1 signaling coincides with epithelial-mesenchymal transition and fibroblast-to-myofibroblast transdifferentiation in the development of adenomyosis in mice. Hum Reprod. 2016;31(2):355–69. https://doi.org/10.1093/humrep/dev314. Ye Z, Zhao A, Li X, et al. Nociceptive nerve-derived CGRP exacerbates uterine fibrogenesis in adenomyosis by promoting CD140b+ CD146+ fibroblast differentiation. Adv Sci (Weinh). 2025;12(45):e07128. https://doi.org/10.1002/advs.202507128. Zhang Y, Yu P, Sun F, et al. Expression of oxytocin receptors in the uterine junctional zone in women with adenomyosis. Acta Obstet Gynecol Scand. 2015;94(4):412–8. https://doi.org/10.1111/aogs.12595. Hsu CY, Hsieh TH, Tsai CF, et al. MiRNA-199a-5p regulates VEGFA in endometrial mesenchymal stem cells and contributes to the pathogenesis of endometriosis. J Pathol. 2014;232(3):330–43. https://doi.org/10.1002/path.4295. Li G, Lin Y, Zhang Y, et al. Endometrial stromal cell ferroptosis promotes angiogenesis in endometriosis. Cell Death Discov. 2022;8(1):29. https://doi.org/10.1038/s41420-022-00821-z. Li Y, He Y, Cheng W, et al. Double-edged roles of ferroptosis in endometriosis and endometriosis-related infertility. Cell Death Discov. 2023;9(1):306. https://doi.org/10.1038/s41420-023-01606-8.

The authors thank all members of the Department of Obstetrics and Gynecology, Peking University Third Hospital, for their support in this study. Author information Authors and Affiliations Contributions Molin Wang, Haojie He, and Xinran Gao contributed equally to this work. Xinran Gao and Haojie He were responsible for conceptualization, methodology, investigation, and writing of the original draft. Molin Wang performed formal analysis, data curation, visualization, and writing, review and editing. Qinkun Sun and Lu Liu contributed to data curation. Chunliang Shang and Hongyan Guo supervised the study, and Hongyan Guo provided resources. All authors read and approved the final manuscript. Corresponding authors Ethics declarations Ethical Approval All animal experiments were conducted in accordance with the Guide for the Care and Use of Laboratory Animals and were approved by the Institutional Animal Care and Use Committee (IACUC) of Peking University Health Science Center (Approval No. BCJI0251; approved on September 8, 2023). Consent to Participate Not applicable. Consent for Publication Not applicable. Competing Interests The authors declare that they have no competing interests. Additional information Publisher's Note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. Rights and permissions Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article’s Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by/4.0/. About this article Cite this article Wang, M., He, H., Gao, X. et al. Estrogen-Progesterone Sequential Exposure Induces the Fibrosis of Adenomyosis in Mice. Reprod. Sci. (2026). https://doi.org/10.1007/s43032-026-02225-1 Received: Accepted: Published: Version of record: DOI: https://doi.org/10.1007/s43032-026-02225-1