PV was purchased from the Korea Plant Extract Bank at the Korea Research Institute of Bioscience and Biotechnology (Chungju-si, Chungchungbuk-do, South Korea). The extract was dissolved in phosphate buffered saline (PBS) for the in vitro enzyme activity assay along with dimethyl sulfoxide (Sigma-Aldrich Chemical Co., Louis, Mo, United States), before being diluted using culture media for cell line experiments, or with corn oil immediately before use in animal studies.

Five-week-old female C57BL/6 mice were purchased from Orient Bio (Seongnam, South Korea) and were maintained in polycarbonate cages during the experimental period at an animal facility at the Institute for Laboratory Animals of Pusan National University. A 12 h day-night cycle, room temperature of 22 ± 2°C, and humidity of 50–60% were automatically maintained in the animal facility. Isoflurane and CO₂ inhalation was used for anesthesia and sacrifice, respectively. All experimental procedures were approved by the Institutional Animal Care and Use Committee of Pusan National University (Pusan, Republic of Korea; PNU-2020-2679).

Endometriosis was induced based on a previous protocol (Choi et al., 2018) with slight modification. Briefly, six-week-old C57BL/6 female mice were ovarectomized and recuperated before estrogen injection. After 14 days, mice were subcutaneously injected with 100 mg/kg β-estradiol (Santa Cruz Biotechnology; Dallas, TX, United States) suspended in corn oil every week. The uteri of donor mice were isolated and chopped using Gentle Max (Miltenyi Biotec, Bergisch Gladbach, Germany) and intraperitoneally inoculated into recipient mice at a 1:1 ratio. One day after uterine injection, the mice were orally administered PV 5 days per a week, and 100 mg/kg β-estradiol was subcutaneously injected once a week for 3 weeks. The dosages of PV given in mice were calculated based on 50% growth inhibition concentration at 48 h after PV treatment in 12Z cells and these two dosages (0.820 and 4.100 mg/kg) were smaller than the dosages calculated from formula provided by FDA (Nair and Jacob, 2016). Three weeks after uterine injection, the mice were euthanized, and endometriotic lesions were excised from the surrounding tissue to evaluate their number and weight. The number of mice per group was five.

Immortalized human endometrial stromal cells (T-HESCs) and immortalized human endometriotic epithelial (12Z) cells were purchased from the American Type Culture Collection (ATCC; #CRL-4003, Rockville, MD, United States) and Applied Biological Materials (#T0764, Richmond, Canada), respectively. T-HESCs were maintained in phenol red-free DMEM/F12 (Welgene, Daegu, South Korea) supplemented with 10% inactivated charcoal (Sigma-Aldrich)-stripped fetal bovine serum (FBS, Invitrogen, Waltham, MA, United States), 1% ITS premix (BD Biosciences, San Jose, CA), and 1% penicillin/streptomycin (Gibco, Rockville, MD, United States). 12Z cells were maintained in DMEM/F12 (Welgene, Daegu, Korea) supplemented with 10% non-inactivated charcoal-stripped FBS and 1% penicillin/streptomycin (Thermo Fisher Scientific, Waltham, MA, United States). Both cell types were cultured at 37 °C in a humidified atmosphere containing 5% CO₂.

Cell proliferation and apoptosis were analyzed using 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) (Sigma) and propidium iodide (PI)/Annexin V (BD Biosciences, Franklin Lakes, NJ, United States) methods 24 h after PV treatment. Both 12Z cells and T-HESCs were plated onto 96 well at 1 × 10⁴ cells/well and treated with the indicated concentrations of PV. Cell viability was analyzed by measuring absorbance at 450 nm using a Spectramax M2 spectrofluorometer (Molecular Devices, San Jose, CA, United States). For PI/Annexin V detection, 3 × 10⁵ cells were plated to 6 wells and cell apoptosis was measured at excitation (Ex) 494/emission (Em) 525 nm for Annexin V and Ex 535/Em 617 nm for PI using Attune X (Thermo Fisher Scientific), and analysis was conducted using the FlowJo program (BD Bioscience).

The reduction of NADH (β-nicotinamide adenine dinucleotide disodium salt hydrate; reduced form, Tokyo Chemical Industry, Tokyo, Japan) was used to validate the LDHA activity of PV. One unit of LDH activity is defined as the amount of enzyme that catalyzes the conversion of lactate into pyruvate to generate 1.0 μM of NADH per minute at 37°C. Briefly, various concentrations of PV were incubated in buffer containing 2 mM pyruvate, 20 μM NADH, 20 mM HEPES-K⁺ (pH 7.2), and 10 ng of purified recombinant human LDHA protein for 20 min. The fluorescence of NADH was measured at Ex 340/Em 460 nm wavelength using a spectrofluorometer (Spectramax M2; Molecular Devices, Sunnyvale, CA, United States).

For protein analysis, total proteins were extracted using RIPA buffer and 1% NP-40 lysis buffer containing a protease inhibitor cocktail (Roche, Basel, Switzerland). The extracted proteins were quantified using the Bradford assay and subjected to immunoblot analysis. Briefly, the proteins were separated using SDS-PAGE and transferred to a PVDF membrane (Amersham Bioscience, Uppsala, Sweden). The first antibody, anti-human PARP (#9542s; Cell Signalling), Caspase-3 (#9665s, Cell Signalling), Caspase-9 (#9508s; Cell Signalling), Bax (NB100-56095; Novusbio), Bcl-2 (NB100-56098; Novusbio), PDHA (sc-377092; Santa Cruz), phospho-PDHA (ab177461; Abcam), LDHA (ab84716; Abcam), PDK1 (ADI-KAP-PK112; Enzo), PDK3 (ab182574, Abcam), and GAPDH (sc-32233; Santa Cruz). HRP-conjugated anti-rabbit IgG or anti-rat IgG (all from Invitrogen) were used as secondary antibodies. The specific bands were developed using a western blot detection kit (Bio-Rad, Hercules, CA, United States) using the ImageQuant LAS 4000 chemiluminescence imaging system (GE Healthcare, Munich, Germany).

The oxygen consumption rate and lactate production in 12Z cells were measured after 4 h of PV treatment to clarify the influence of PV on metabolic activity. First, the oxygen consumption rate was analyzed using an oxygen consumption rate assay kit (Abcam, Cambridge, MA, United States). 12Z cells were seeded in a black 96-well microplate at a density of 3 × 10⁴ cells. The next day, various concentrations of PV (0, 62.5, 125, and 250 μg/ml) were added, and the experiment was conducted according to the manufacturer’s instructions. Second, lactate production was measured using a lactate fluorometric assay kit (Biovision, CA, United States). After 4 h of PV treatment, the media was replaced with phenol red- and serum-free DMEM and incubated for 1 h at 37°C. Then, 1 μl of the medium from each well was assessed by fluorescence at Em 535/Ex 590 nm using the Spectramax M2 spectrofluorometer. Dichloroacetate (DCA) was used as a positive control for lactate production.

The MMP and mitoROS were analyzed using the cationic and lipophilic fluorescent dye, Tetramethylrhodamine (TMRM), and red mitochondrial superoxide indicator, MitoSOX™ (Invitrogen, Eugene, OR, United States), respectively. The 12Z cells were stained with TMRM 18 h after PV treatment using flow cytometry. After pre-treatment with PV for 4 h, 12Z cells were washed twice with pre-warmed PBS for TMRM staining or Hank’s balanced salt solution for MitoSOX staining. To measure MMP, TMRM was added at a final concentration of 20 nM and incubated for 30 min at 37°C. To assess mitoROS, 5 μM MitoSOX was added and incubated for 10 min at 37°C. After staining, 12Z cells were washed three times, and then the intensity of fluorescence was analyzed using Attune X (Thermo Fisher Scientific) and the geometric mean fluorescence intensity was calculated using the FlowJo program (BD Bioscience).

Oleanolic acid (OA) (99% purity) and rosmarinic acid (RA) (98% purity) were purchased from Sigma-Aldrich, and ursolic acid (UA) (92.84% purity) was purchased from the Resource Bank of Korean Medicine, National Institute for Korean Medicine Development. The compounds were dissolved in methanol at a concentration of 10 mM for use as controls, according to the methods described in previous studies (Qiang et al., 2011; Chen et al., 2012). The PV extract was dissolved to 3.12 mg/ml, sonicated using ultrasound for 20 min, and filtered through a 0.45 μm syringe filter (Millipore Sigma, Burlington, MA, United States). HPLC analysis was conducted on an Agilent 1200 series system (Agilent Technologies, Santa Clara, CA, United States) equipped with a Shiseido UG120 column (5 μm, 6.4 × 250 mm) maintained at 25°C. The mobile phase was methanol (90%): 0.5% ammonium acetate (10%) for OA, UA, and methanol (52%), and 0.01% phosphoric acid (48%) for RA. Samples (20 μl) were analyzed at a flow rate of 0.6 ml/min using an ultraviolet detector at 210 nm for OA and UA, and at 310 nm for RA.

Values are expressed as the mean ± SEM. A two-tailed Student’s t-test was used for comparisons of two different groups and one-way ANOVA followed by Tukey’s post-hoc test was used for comparisons of multiple groups. These were performed using GraphPad Prism Software (GraphPad, San Diego, CA, United States).

To evaluate if PV is beneficial for people with endometriosis, we first applied PV to a mouse endometriosis model. PV extract was administered orally at doses of 0.820 mg/kg and 4.100 mg/kg body weight or with 1 mg/ml of dienogest five times a week for 3 weeks after intraperitoneal inoculation of donor uterus tissue (Figure 1A). Dienogest, the first-line drug for endometriosis (Andres Mde et al., 2015; Liu et al., 2021), was used as a control to evaluate the efficiency of PV in endometriosis. The PV group showed a significant reduction in cyst-like ectopic lesion volume compared to the untreated mice group. However, a significant reduction in the volume and number of endometriotic lesions was observed in the dienogest-treated mouse group (Figures 1B,C). In addition, to clarify the safety of PV, kidney and liver tissues were examined using H&E staining. During histological examination, the doses of PV used in the in vivo experiment were deemed non-toxic (Supplementary Figure S1). These results indicate that the growth of ectopic endometriotic tissues can be ameliorated by oral administration of PV.

To determine the inhibitory mechanism of endometriotic tissue growth in mouse experiments (Figure 1C), we first assessed the apoptosis of 12Z cells and T-HESCs after treatment with increasing doses of PV (0, 50, 100, 250, 500, and 1000 μg/ml) for 24 h. The colorimetric cell viability assay showed that PV induced cell death in both 12Z and T-HESCs in a dose-dependent manner, but the 12Z cells (50% growth inhibition concentration (GI₅₀) 210.6 μg/ml) were more susceptible to PV-induced cell death than T-HESCs (GI₅₀ 453.9 μg/ml) (Figure 2A). Similar to the colorimetric cell viability assay, the percentage of apoptotic cells, including necroptic, early, and late apoptotic cells, started to increase at a concentration of 150 μg/ml, and these apoptotic cell populations were dramatically increased at a concentration of 250 μg/ml in 12Z cells (Figure 2B and Supplementary Figure S2).

Apoptosis occurred through a signaling cascade; therefore, we next analyzed the changes in the intracellular apoptosis signaling pathway in 12Z cells by western blotting. 12Z cells were exposed to various concentrations of PV, and the protein levels and cleaved forms of caspase-3 and -9 and PARP were examined. PV induced activation of procaspase-9 into cleaved caspase-9 and cleavage of procaspase-3 into its active form. PARP, downstream of the activation of caspase-3, is a well-known characteristic of apoptosis, and cleavage of the PARP protein was also detected in a dose-dependent manner (Figure 2C). These data suggest that PV effectively induces 12Z cell apoptosis by activating the caspase-3 and -9 and PARP pathways, indicating the activation of the mitochondrial pathway for apoptosis.

Mitochondria play an important role in the initiation and regulation of apoptosis. The mitochondrial-dependent apoptosis pathway involves changes in MMP, mitoROS generation, and ATP synthesis by oxidative phosphorylation, which is a phenomenon of mitochondria disruption (Nicholls, 2004; Martinez-Reyes and Chandel, 2020). The change in the mitochondrial membrane potential causes mitochondrial outer membrane permeabilization, resulting in caspase-dependent cell death following the release of cytochrome C into the cytosol (Li et al., 1997). Therefore, membrane depolarization and excessive production of mitoROS are good indicators of mitochondrial dysfunction (Abraham et al., 2008; Dykens et al., 2008).

Consistent with Figure 2C, we attempted to clarify whether the mitochondrial apoptotic pathway is a major factor in PV-induced apoptosis. First, we monitored both MMP and mitoROS using TMRM and Mitosox staining-based assays, respectively. The results showed that MMP was markedly reduced but mitoROS dramatically increased after 18 h of treatment with PV extract (Figures 3A,B). Because a loss of MMP promotes the release of cytochrome c from the mitochondria to the cytosol, we next examined the protein levels of Bax and Bcl-2, which are pro- and anti-apoptotic Bcl-2 members, respectively (Salah-Eldin et al., 2003). As shown in Figure 3C, the expression of Bax protein was increased, whereas the expression of Bcl-2 protein decreased upon PV treatment in 12Z cells. The increased ratio of Bax/Bcl-2 implies that PV favors the occurrence of apoptosis in 12Z cells. These results indicate that PV induces apoptosis of 12Z cells through the Bcl-2/Bax balance and ROS-mediated mitochondrial damage.

In glycolytic cells, excessive pyruvate influx into the mitochondria leads to an accelerated tricarboxylic acid cycle, resulting in impaired mitochondria (Martinez-Reyes and Chandel, 2020). As shown in Figure 3, PV could lead to mitochondrial malfunction, so we next determined the influence of PV on glycolysis in 12Z cells. Therefore, we measured the activity and expression of LDHA, PDHA and PDK1/3. As shown in Figures 4A–C, PV significantly suppresses the in vitro enzymatic and intracellular LDHA activity, while the expression of LDHA did not change. In addition, PV markedly inhibited the expression of PDK1/3 and phosphorylation of PDHA (Figure 4D). These data imply that the mitochondrial apoptosis signal induced by PV is associated with enhanced PDH and suppressed LDHA activity. As LDHA and PDH are essential enzymes for the conversion of pyruvate into lactate or acetyl-CoA, respectively (Koukourakis et al., 2005; Zhang et al., 2015; Urbanska and Orzechowski, 2019), we next analyzed the lactate production and oxygen consumption rate after 4 h of PV treatment. The results showed that PV significantly decreased the amount of extracellular lactate but increased oxygen consumption in a dose-dependent manner (Figures 4E,F). Taken together, these results indicate that PV leads to 12Z cell apoptosis through the mitochondrial apoptosis signaling pathway, particularly glycolytic metabolism.

To validate the purity of the PV extract used in this study, high-performance liquid chromatography analysis was performed. As shown in Figure 5, the PV extract was well separated within 15 min. OA, UA, and RA are well known as the main active substances in PV extract (Cheung and Zhang, 2008; Wang et al., 2019). Therefore, we estimated three substances by the retention times of the peaks with those of three standard solutions for OA, UA, and RA under the same conditions. To evaluate the reproducibility of the extraction efficiency, the extraction was repeated three times using the same sample. The results showed that the contents of OA, UA, and RA in 1 g of PV were 14.290 ± 0.148 mg, 69.852 ± 2.457 mg, and 9.373 ± 0.304 mg, respectively (Figure 5C). Based on GI₅₀ of PV in 12Z cells, the amount of OA, UA, and RA were 6.72 μM (3.10 μg/ml), 33.14 μM (15.16 μg/ml), and 5.42 uM (1.95 μg/ml), respectively. This result indicated that UA was the most dominant compound in PV among the three substances.

We next examined the effect of each substance on cell viability and the LDHA activity of each substance to determine the main active compounds in PV-led 12Z cell death in terms of dysregulated glycolysis. In the colorimetric assay, RA did not induce cell apoptosis even at a dose of 250 μM and showed the smallest effect on LDHA activity (IC₅₀ 42.93 μM) among the three substances. In contrast to RA, UA and OA induced apoptosis of 12Z cells (GI₅₀ 87.58 and 218.6 μμM, respectively) and showed significant effects on LDHA activity (IC₅₀ 1.76 and 0.64 μμM, respectively) (Figure 6). Because the content of OA and UA in 24.84 μg/ml of PV (IC₅₀) were 0.79 and 3.91 μμM, respectively, it can be deduced that UA is the most effective substance in PV for inducing 12Z cell apoptosis via inhibition of LDHA activity among the three compounds.