A total of 50 8-week-old female 18–22 g BALB/c mice were purchased from Shanghai JST Laboratory Animal Co., Ltd., of which 20 were recipient mice, 20 were donor mice and 10 were control mice. All mice were subjected to controlled conditions under a 12 h light/dark cycle with a temperature of 22±2°C and relative humidity of 50±10%, as well as free access to food and water. All experiments were performed under the guidelines of the National Research Council Guide for the Care and Use of Laboratory Animals and were approved by the Institutional Laboratory Animal Review Board of Shanghai First Maternity and Infant Hospital (Shanghai, China; approval no. TJBG28625103).

A mouse model of endometriosis was established by intraperitoneal injection of endometrial debris (23). Briefly, after 1 week of adaptation, 20 donor mice were injected intramuscularly with 100 mg/kg estradiol 2–3 times per week (Hangzhou Animal Pharmaceutical (Hangzhou) Co., Ltd.). Euthanasia was performed after 1 week by cervical dislocation. The uteri were removed and collected, and the uterine tissue was cut up and injected intraperitoneally into the abdominal cavity of the recipient mice. The uterine horns of one donor mouse were injected into one recipient mouse. A total of 30 mice (20 recipient mice and 10 control mice) were randomly divided into three groups: i) Control group (CTL group), in which 10 mice received an intraperitoneal injection of saline in equal amounts to the mice in the established mouse endometriosis groups; ii) endometriosis with rapamycin treatment group (EM-R group), in which 10 mice received an intraperitoneal injection of endometrial debris and 250 µg rapamycin/mouse once per week at one 1 after endometriosis induction; and iii) endometriosis group (EM group), in which 10 mice received an intraperitoneal injection of endometrial debris only. Rapamycin was purchased from Sigma-Aldrich (Merck KGaA; cat. no. 553210). The present study referred to the study by Ren et al (24) for the dose of rapamycin used in mice. Body weight was measured and the hotplate test was performed in all mice at 1, 2, 3 and 4 weeks after endometriosis induction, and mice were executed 4 weeks after endometriosis induction by cervical dislocation.

The hotplate test was performed using a commercial hotplate analgesia meter (BME-480; Institute of Biomedical Engineering, Chinese Academy of Medical Sciences) to assess the degree of discomfort caused by endometriosis, as previously described (25). In summary, mice were given 10 min to acclimate before the test. The mouse had to hop on the hotplate or shake or lick its rear paws during 1 min of being placed within the cylinder to assess the withdrawal latencies to thermal stimulation. Over the course of 1 h, the latency was measured twice and then averaged by the two test results.

After the mice were executed, they were placed in a supine position with their limbs unfolded and fixed on a dissecting board. The abdomen was disinfected with 75% alcohol, the lower abdominal skin was lifted with ophthalmic forceps and a 2-cm incision was made along the abdominal midline with scissors. A total of 3–5 ml of sterile saline was injected into the abdominal cavity of the mice and the abdominal cavity of the mice was opened under aseptic conditions. The abdominal fluid was rinsed three times and the rinsed fluid was collected into centrifuge tubes for the detection of oxidative stress-related substances. Superoxide dismutase (SOD), malondialdehyde (MDA), glutathione peroxidase (GSH-PX) and 8-hydroxydeoxyguanosine (8-OHdG) were detected as the oxidative and antioxidant markers in the peritoneal fluid of mice. The SOD assay kit (cat. no. A001-3-2; WST-1 method), MDA assay kit (cat. no. A003-1-2; TBA method), GSH-PX assay kit (cat. no. A005-1-2; colorimetric method) and mouse 8-OHdG ELISA kit (cat. no. H165-1-2) were purchased from Nanjing Jiancheng Bioengineering Institute. The peritoneal fluid of PPARα and IGFBP2 in mice were detected using an ELISA kit (cat. nos. MM-0249M1 and MM-0029H2, respectively; Meimian Industry; Jiangsu ELISA Industrial Co., Ltd.). All operations are strictly in accordance with the kit instructions.

Ovarian tissue samples were fixed in 4% paraformaldehyde for 24 h at 4°C and embedded in paraffin. Sections (4 µm) were used for subsequent staining. The first section was stained with hematoxylin and eosin to confirm the pathological diagnosis, and subsequent sections were stained with senescence-related markers such as p16, p21 and Lamin B1. Tissue sections were deparaffinized in xylene and rehydrated through a graded ethanol series. Antigen retrieval was performed in EDTA buffer (pH 9.0; Shanghai Sun Biotechnology Co., Ltd.) or citric acid solution (pH 6.0; Shanghai Sun Biotechnology Co., Ltd.) using a microwave heating method, then allowed to cool to room temperature. Following antigen retrieval and cooling, sections were washed three times with PBS. To block non-specific binding, sections were incubated in a solution of PBS containing 5% normal goat serum (cat. no. G1208; Wuhan Servicebio Technology Co., Ltd.) for 1 h at room temperature. The following antibodies were used in this experiment: p16 (1:50; cat. no. ab51243), p21 (1:1,000; cat. no. ab188224) and Lamin B1 (1:1,000; cat. no. ab16048). All antibodies were purchased from Abcam. All sections were incubated with the aforementioned primary antibodies overnight at 4°C. The sections were then incubated with horseradish peroxidase-labeled secondary antibody detection reagent (1:300; cat. no. A0303; Beyotime Biotechnology) for 1 h at room temperature. The bound antibody complexes were stained with diaminobenzidine for 3–5 min at room temperature, followed by hematoxylin for nuclear staining at room temperature. The development time was monitored under a microscope (typically between 3 to 5 min) and stopped immediately when optimal staining intensity was achieved with minimal background. Images were captured under a microscope (BX51; Olympus Corporation) equipped with a digital camera (DP70; Olympus Corporation). According to the manufacturer's instructions, distinct tissue slides were utilized for several antibodies as positive control. For negative controls, tissue samples were treated with mouse or rabbit serum (Wuhan Servicebio Technology Co., Ltd.) in lieu of primary antibodies.

Mouse ovarian tissues were cut into small pieces (soybean size), placed in a pre-cooled homogenizer, 1 ml RNA extract (Wuhan Servicebio Technology Co., Ltd.) was added per 100 mg of tissue, and the tissues were homogenized until no visible pieces were present. A total of 200 µl chloroform was added per 1 ml RNA extract, and the mixture was vortexed and mixed or shaken upside down vigorously for 15 sed, let stand at room temperature for 3–5 min, then centrifuged at 12,000 × g for 15 min at 4°C. The upper colorless aqueous phase was carefully transferred to a new centrifuge tube, aspirating 500–550 µl per ml of extract. A total of 500 µl isopropanol was added to the collected aqueous extract tube, which was centrifuged at 4°C for 10 min at 12,000 × g. The white precipitate visible at the bottom of the tube was the total RNA, and so supernatant was discarded. A total of 1 ml of 75% ethanol was then added, which was then mixed upside down (until the white precipitate floats) and centrifuged at 12,000 × g for 5 min at 4°C. The supernatant was discarded, then centrifugation was performed briefly at high speed (3–5 sec at 5,000 × g at 4°C) and the liquid was carefully aspirated with a micropipette. The centrifuge tube with RNA precipitate was left open for 3–5 min to allow the RNA to dry slightly, and then 20–50 µl DEPC-treated water (Wuhan Servicebio Technology Co., Ltd.) was added to fully dissolve the RNA. The NanoDrop^™ Microvolume UV–Vis spectrophotometer (Thermo Fisher Scientific, Inc.) was used to assess the quality control and quantification of total RNA. RNA with good integrity and purity (OD260/OD280 values of 1.8–2.0) was used for further analysis. Mouse tissue RNA was reverse transcribed and quantitative PCR (qPCR) was performed using forward and reverse primers. The cDNA synthesis was performed using a reverse transcription kit (Tiangen Biotech Co., Ltd.) at 42°C for 30 min. To evaluate the abundance of mRNA, real-time PCR was conducted using SYBR Premix Ex Taq (Takara Bio, Inc.). The thermocycling protocol consisted of an initial denaturation step at 95°C for 30 sec, followed by 40 cycles of 95°C for 5 sec and 60°C for 30 sec. GAPDH was used as a housekeeping gene. The sequences of senescence-related markers (p16, p21 and γH2AX), gonadotropin receptor [follicle-stimulating hormone receptor (FSHR) and luteinizing hormone receptor (LHR)] and PPARα and IGFBP2 primers are listed in Table I. All primers were designed by Sangon Biotech (Shanghai) Co., Ltd. The relative gene expression levels were calculated as ΔCq values (Cq ‘target gene’ minus Cq ‘housekeeping gene’) and 2^−ΔΔCq ratios indicated fold changes (26).

A total of 30 mice were sacrificed 4 weeks after endometriosis induction. After fixation with 4% paraformaldehyde in phosphate buffer at 4°C for 24 h, the tissues were embedded in paraffin and sectioned at 4 µm thickness. Ovarian tissue sections were stained with HE at room temperature and the number of follicles at different stages was counted for each mouse. The method of follicle counting involved 5-µm continuous sections, with 1/5 sections stained and counted (27). A total of five types of follicles were counted: i) Primordial follicles; ii) primary follicles; iii) secondary follicles; iv) antral follicles; and v) corpus luteum. Primordial follicles were defined as oocytes surrounded by a layer of squamous (flattened) granulosa cells; primary follicles were defined as one follicle containing several or all oocytes surrounded by a single layer of cuboidal granulosa cells; secondary follicles were defined as a follicle surrounded by >1 layer of cuboidal granulosa cells with no visible lumen; a follicle was identified as a antral follicle if it had a well-defined lumen and a layer of cumulus granulosa cells; and the corpus luteum was a post-ovulatory structure, which was filled with lutein cells that form only after ovulation (28). Images were captured using a microscope (BX51; Olympus Corporation) fitted with a digital camera (DP70; Olympus Corporation).

Mouse primary granulosa cells were extracted and cultured with reference to the study by Tian et al (29). Mouse ovaries were washed with sterile PBS and transferred to DMEM/F12 (cat. no. 11330-057; Invitrogen^™; Thermo Fisher Scientific, Inc.). Follicles were punctured with a 31-gauge needle under a dissecting microscope, and granulosa cells were released from the follicles, centrifuged at 1,000 × g for 5 min at room temperature, resuspended in fresh DMEM/F12 with 10% FBS, and incubated at 37°C and 5% CO₂. Cells were passaged every other day.

The granulosa KGN cell line was purchased from Shanghai Kanglang Biotechnology Co., Ltd. and cultured in DMEM/F12 supplemented with 10% FBS at 37°C and 5% CO₂. Cells were passaged every 5–6 days, and the number of passages was <15 for the experiments.

Small interfering (si)RNA of the human PPARα gene (siRNA-PPARα) and non-specific control siRNA (siRNAc) were purchased from Shanghai GenePharma Co., Ltd. and were transfected into cells at room temperature using Lipofectamine^™ 3000 (Invitrogen; Thermo Fisher Scientific, Inc.) for 48 h at a final concentration of 30 nM, according to the manufacturer's instructions. Subsequent experiments were conducted 48 h after cell transfection. The following siRNA sequences were used: siRNA-PPARα, 5′-GCUUUACGGAAUACCAGUAUU-3′; and siRNAc, (forward) 5′-UUCUCCGAACGUGUCACGUdTdT-3′ and (reverse) 5′-ACGUGACACGUUCGGAGAAdTdT-3′.

The PPARα overexpression plasmid [pCDNA3.1(+) plasmid vector Asian Vector Biotechnology Co., Ltd.] was transfected into granulocyte cells for 48 h using Lipofectamine 3000 and an empty plasmid (Asian Vector Biotechnology Co., Ltd.) was used as a negative control. Cells were transfected with 1.0 µg of either the PPARα overexpression plasmid or the empty control plasmid using 2.0 µl of Lipofectamine 3000 reagent per well in a 12-well plate, according to the manufacturer's instructions. Subsequent experiments were conducted 48 h after cell transfection.

To demonstrated the effects of PPARα knockdown or overexpression on cellular senescence, cells were treated with H₂O₂ following knockdown or overexpression. To demonstrated the anti-senescence effects of rapamycin, cells were treated with (50 nM) rapamycin (Sangon Biotech Co., Ltd.).

All statistical analyses were performed using GraphPad Prism 8 (Dotmatics). For comparisons among ≥3 groups, one-way analysis of variance (ANOVA) was employed, followed by Tukey's post hoc test for multiple comparisons when ANOVA indicated statistical significance (P<0.05). Unless otherwise stated, continuous variables are presented as mean ± standard deviation for normally distributed data. All tests were two-tailed, and P<0.05 was considered to indicate a statistically significant difference.

The findings demonstrated that there was no significant difference in body weight among the 3 groups of mice before the induction of endometriosis and their death (Fig. 1A). However, the results of the hotplate test were significantly different between the EM group and EM-R group, and the hotplate time from week 2 post-endometriosis induction was significantly lower in the mice of the EM group than in the EM-R group. There is no difference in hotplate test between CTL group and EM-R group (Fig. 1B). No lesions of endometriosis were found in the mice of the CTL group; however, obvious ectopic lesions were observed in the peritoneal cavity of mice in the EM-R and EM groups. Moreover, the weight of endometriotic lesions in mice in the EM group were significantly higher than that in the EM-R group (193.4±53.22 and 55.4±26.69 mg, respectively; P<0.01; Fig. 1C). This indicates that rapamycin may inhibit endometriosis.

There was no significant difference between the CTL and EM-R groups for the levels of oxidative stress-related molecules, SOD (8.60±1.71 and 7.93±1.32 U/ml, respectively; P=0.49), GSH-PX (18.99±2.32 and 17.30±2.25 µmol/l, respectively; P=0.17), MDA (1.11±0.13 and 1.39±0.32 nmol/ml, respectively; P=0.06) and 8-OHdG (4.34±2.31 and 5.91±2.68 ng/l, respectively; P=0.45). The antioxidant molecules, SOD and GSH-PX, in mouse peritoneal fluid were significantly lower in the EM group compared with in the EM-R group (SOD, 1.65±0.69 vs. 7.93±1.32 U/ml; GSH-PX, 4.62±1.48 vs. 17.30±2.25 µmol/l; both P<0.01). By contrast, the markers of increased oxidative capacity, MDA and 8-OHdG, were significantly higher in the EM group than in EM-R group (MDA, 4.79±0.32 vs. 1.39±0.32 nmol/ml; 8-OHdG, 24.31±3.48 vs. 5.91±2.68 ng/l; both P<0.001) (Fig. 2). This suggests the presence of oxidative stress, decreased antioxidant capacity and increased oxidative function in the peritoneal cavity of endometriosis mice without rapamycin treatment. Moreover, levels of activation of intraperitoneal oxidative stress were reduced in rapamycin-treated endometriosis mice.

Increased expression of senescence-associated markers, p16, p21 and γH2AX, and decreased expression of Lamin B1 suggest that tissue or cellular senescence is occurring (30). The immunohistochemical experiments in the present study demonstrated that senescence-related markers expression in ovarian granulosa cells and oocyte did not differ significantly between the CTL and EM-R group. However, the expression of senescence-related markers, p16 and p21, in ovarian granulosa cells and oocytes was significantly higher in the EM group than in the EM-R group (p16, 0.42±0.14 vs. 0.12±0.03; p21, 0.37±0.04 vs. 0.11±0.02; both P<0.01). Another senescence-related marker, Lamin B1, was significantly lower in ovarian granulosa cells and oocytes of the EM group compared with the EM-R group (0.13±0.05 vs. 0.37±0.07; P<0.01) (Fig. 3A). This suggests that ovarian tissues in endometriosis mice without rapamycin treatment undergo senescence, and rapamycin could decrease expression of senescence markers in ovaries. Similarly, ovarian tissue PCR experiments revealed significantly higher gene expression of p16, p21 and γH2AX in mice in the EM group than in mice in the EM-R group (p16, 3.57±0.36 vs. 0.94±0.04; p21, 3.80±0.39 vs. 0.93±0.06; γH2AX, 5.63±1.37 vs. 1.61±0.61; all P<0.01). Moreover, gonadotropin receptor (FSHR and LHR) gene expression was significantly higher in the EM-R group than the EM group (FSHR, 0.95±0.04 vs. 0.23±0.11; LHR, 0.98±0.04 vs. 0.25±0.07; both P<0.01) (Fig. 3B).

The number of follicles at different stages was not significantly different between the CTL and EM-R groups. However, primordial follicles of the EM group were significantly higher than in the EM-R group (595.6±13.66 vs. 392.1±13.89; P<0.01), whilst primary follicles, secondary follicles, antral follicles and corpus luteum were significantly lower in the EM group than in the EM-R group (primary follicles, 172.2±24.15 vs. 278.6±23.08; secondary follicles, 94.9±10.62 vs. 145.3±9.81; antral follicles, 48±8.56 vs. 106.2±7.88; corpus luteum, 37.8±5.73 vs. 103.6±8.5; All P<0.01). This suggests that endometriosis mice follicles undergo senescence in response to oxidative stress, which leads to impaired development and significant increase in the number of mature follicles in endometriosis mice treated with rapamycin (Fig. 4).

The concentration of PPARα and IGFBP2 in the peritoneal fluid of mice in the CTL group were not significantly different from those of the EM-R group; however, the concentration of PPARα and IGFBP2 in the EM group were significantly lower than those in the EM-R group (PPARα, 46.2±19.33 vs. 191.6±56.37 pg/ml; IGFBP2, 7.90±4.04 vs. 35.4±5.89 ng/ml; both P<0.01) (Fig. 5A). Furthermore, the results revealed a positive correlation between SOD and PPARα, and between SOD and IGFBP2, compared with a negative correlation between MDA and PPARα, and between MDA and IGFBP2 in both the EM-R and EM groups (Fig. 5B). In addition, the gene expression of PPARα and IGFBP2 in the ovaries of mice in the EM-R group was not significantly different from that of mice in the CTL group; however, the expression of PPARα and IGFBP2 in the ovaries of mice in the EM group was significantly lower than that of mice in the EM-R group (PPARα, 0.90±0.07 vs. 0.45±0.03; IGFBP2, 0.91±0.09 vs. 0.46±0.06; both P<0.01). This indicated that the expression of PPARα and IGFBP2 in the ovaries of endometriosis mice was significantly decreased, and treatment with rapamycin activated PPARα and IGFBP2 expression (Fig. 5C).

Granulosa cells, which closely surround the oocyte, are essential for supporting oocyte development (31). To assess cellular senescence in these cells, primary granulosa cells were isolated from three experimental groups of mice. PCR analysis revealed that granulosa cells from the EM group exhibited significantly elevated levels of senescence-associated markers, including p16, p21 and γH2AX, compared with in the EM-R group (p16, 2.95±0.41 vs. 1.11±0.11; p21, 5.13±1.02 vs. 1.09±0.25; γH2AX, 3.74±0.87 vs. 1.01±0.09; all P<0.01; Fig. 6A). Notably, the expression of these markers in the EM-R group was comparable with that of the CTL group, with no statistically significant differences observed between them (Fig. 6A). By contrast, the expression levels of PPARα and IGFBP2 were significantly reduced in the EM group compared with the EM-R group (PPARα, 0.35±0.55 vs. 0.97±0.04; IGFBP2, 0.34±0.03 vs. 0.93±0.04; both P<0.01; Fig. 6B).

To evaluate the functional relationship between PPARα and IGFBP2, knockdown and overexpression of PPARα experiments were performed using KGN cells. First, PCR experiments were performed to assess the expression of PPARα after transfecting cells with siRNA and overexpression plasmids. The results demonstrated that PPARα expression was significantly lower after siRNA transfection (0.37±0.05 vs. 1.04±0.06; P=0.0002) and significantly higher after overexpression plasmid transfection (4.50±0.73 vs. 1.06±0.11; P=0.0013) than in the respective control groups (Fig. 7A). Notably, silencing PPARα significantly reduced IGFBP2 expression, whereas PPARα overexpression significantly increased IGFBP2 levels (Fig. 7B and C). This indicated that IGFBP2 is a downstream target of PPARα.

To assess the role of PPARα in cellular senescence, granulosa cells were treated with H₂O₂ following PPARα knockdown or overexpression. The siRNA-PPARα + H₂O₂ group exhibited the highest expression of senescence markers (p16, p21 and γH2AX), significantly exceeding levels in the siRNA-control + H₂O2 group (p16, 5.96±0.90 vs. 3.03±0.23; p21, 5.10±0.34 vs. 2.89±0.30; γH2AX, 4.79±0.27 vs. 2.66±0.48; all P<0.01; Fig. 7B). Conversely, the pcDNA-PPARα + H₂O₂ group exhibited lower expression of the senescence markers (p16, p21 and γH2AX) than the pcDNA + H₂O₂ group, (p16, 2.56±0.41 vs. 5.13±0.17; p21, 3.13±0.13 vs. 6.15±0.17; γH2AX, 2.40±0.25 vs. 4.63±0.35; all P<0.01; Fig. 7C). PPARα overexpression attenuated H2O2-induced senescence, further supporting the protective role of PPARα against cellular aging. Moreover, cells with knockdown or overexpression of PPARα were treated with H₂O₂ for 24 h followed by the addition of 30 µM rapamycin (32) for 48 h, revealed that rapamycin significantly increased PPARα and IGFBP2 expression and significantly decreased the expression of senescence-related markers, p16, p21 and γH2AX. The siRNA-PPARα + H₂O₂ group exhibited higher expression of the senescence markers (p16, p21 and γH2AX) than the siRNA-PPARα + H₂O₂ + rapamycin group (p16, 0.97±0.06 vs. 0.35±0.05; p21, 0.95±0.09 vs. 0.21±0.10; γH2AX, 0.94±0.08 vs. 0.31±0.09; all P<0.01; Fig. 7D). The pcDNA-PPARα + H₂O₂ group exhibited higher expression of the senescence markers (p16, p21 and γH2AX) than the pcDNA-PPARα + H₂O₂ + rapamycin group (p16, 0.99±0.08 vs. 0.29±0.04; p21, 0.97±0.06 vs. 0.26±0.07; γH2AX, 0.98±0.06 vs. 0.22±0.02; all P<0.01; Fig. 7E) These findings suggest that rapamycin counteracted senescence by upregulating the PPARα-IGFBP2 axis.