Ulinastatin protects against cisplatin-induced ovarian damage via Nrf2/Keap1 pathway

Abstract

The administration of chemotherapeutic agents such as cisplatin frequently results in ovarian damage and premature ovarian failure (POF), severely compromising fertility and quality of life in young women. This study aimed to investigate the protective effects of ulinastatin, a compound with anti-inflammatory and antioxidant properties, against cisplatin-induced ovarian injury and to elucidate the underlying mechanisms through the Nrf2/Keap1 pathway. In vivo and in vitro models were established using cisplatin-treated rats and ovarian granulosa cells. Experimental results demonstrated that ulinastatin significantly alleviated cisplatin-induced ovarian structural damage, reduced follicular atresia, and inhibited ovarian fibrosis and inflammation. Furthermore, ulinastatin restored serum hormone levels (FSH, E2, AMH), mitigated oxidative stress by reducing ROS and MDA levels while increasing SOD and GSH levels, and suppressed apoptosis by downregulating cleaved-caspase-3 and BAX while upregulating Bcl-2. Most importantly, ulinastatin activated the Nrf2/Keap1 pathway. Keap1 silencing enhanced the protective effects of ulinastatin, whereas Keap1 overexpression attenuated these effects, confirming that the Nrf2/Keap1 pathway is the key mediator of ulinastatin’s protective role in the ovary. These findings underscore the dual function of ulinastatin in repairing cisplatin-induced ovarian damage and provide a promising therapeutic strategy for preserving fertility in female cancer patients undergoing chemotherapy.

Supplementary Information

The online version contains supplementary material available at 10.1186/s13048-025-01760-w.

Introduction

In recent years, with the continuous advancements in chemotherapy and improvements in supportive care, the survival rate of cancer patients has significantly increased, with approximately 90% of young female patients diagnosed at an early stage achieving long-term survival [1]. However, the risk of long-term complications associated with chemotherapy has become increasingly prominent [2–5]. In young gynecologic oncology patients who have undergone fertility-preserving surgery, the toxic side effects of chemotherapy often lead to premature ovarian failure (POF), resulting in infertility and seriously affecting the quality of life of women of reproductive age. With the increasing trend of cancer occurrence in younger populations and the improvement in the survival rate of young female cancer patients, finding effective personalized fertility preservation measures that can protect or restore ovarian function damaged by chemotherapy without affecting cancer treatment is an urgent issue [6].

Current studies have shown that chemotherapeutic drugs can cause ovarian tissue and structural disorders, affect follicle initiation, growth, and maturation processes, and lead to ovarian interstitial fibrosis [7–9]. However, the specific mechanism remains unclear, posing challenges in the development and research of targeted therapeutic drugs. At present, clinical approaches for preventing or treating chemotherapy-induced POF mainly include gonadotropin-releasing hormone therapy, hormone replacement therapy, and fertility cryopreservation techniques. Each method has certain limitations and has not yet been widely applied in clinical practice.

Ulinastatin is a broad-spectrum serine protease inhibitor with diverse anti-inflammatory, antioxidant, and anti-apoptotic properties. In recent years, its protective effects have been extensively investigated in various disease models, particularly in oxidative stress-associated tissue injury, where it has demonstrated significant protective efficacy [10, 11]. Oxidative stress is a key mechanism by which chemotherapeutic agents such as cisplatin induce organ damage. Cisplatin disrupts intracellular redox balance by inducing excessive production of reactive oxygen species (ROS), which subsequently triggers apoptosis and tissue fibrosis [12, 13]. Through its antioxidant capacity, ulinastatin effectively scavenges ROS and mitigates tissue damage caused by oxidative stress.

The Nrf2/Keap1 pathway is a central regulatory mechanism of the intracellular antioxidant defense system. Under physiological conditions, Nrf2 binds to its inhibitor Keap1 and remains inactive. However, under oxidative stress, Nrf2 dissociates from Keap1 and translocates into the nucleus, where it activates the expression of downstream antioxidant genes such as HO-1, NQO1, and SOD, thereby maintaining intracellular redox balance [14, 15]. Studies have shown that ulinastatin significantly enhances cellular antioxidant capacity and inhibits oxidative stress-induced apoptosis and tissue fibrosis by activating the Nrf2/Keap1 pathway [16, 17]. Therefore, ulinastatin may play a crucial role in the prevention and treatment of chemotherapeutic drug-induced ovarian damage by modulating the Nrf2/Keap1 pathway.

Based on the aforementioned research, this study focused on ulinastatin as the research subject and established a stable cisplatin-induced rat ovarian injury model and a rat ovarian granulosa cell injury model in vivo and ex vivo. The objective was to explore the potential pathways through which cisplatin induces ovarian damage and to investigate the feasibility and underlying mechanisms of ulinastatin in prophylactic protection and therapeutic repair of cisplatin-induced ovarian and granulosa cell injury in rats. The ultimate goal is to identify a novel approach that integrates both preventive protection and therapeutic repair for chemotherapy-induced ovarian damage, providing a new therapeutic strategy and theoretical foundation for the prevention and treatment of chemotherapy-induced ovarian injury, thereby offering hope to young female cancer patients seeking to preserve their reproductive function.

Materials and methods

Reagents

Ulinastatin (cat. HY-134616) was purchased from MCE (USA), and cisplatin (cat. SJ-MX0054) was purchased from Sparkjade (China). Antibodies against GAPDH (2118), TOM20 (42406), LC3I/II (4108), and SQSTM1/P62 (5114) were obtained from Cell Signaling Technology (USA). Anti-Syntaxin 17 (17815-1-AP), anti-Snap29 (12704-1-AP), anti-VAMP8 (15546-1-AP), and anti-SLC7A11/xCT (26864-1-AP) were purchased from Proteintech (USA). Anti-GPX4 (SAB5700944) was obtained from Sigma (USA), and anti-Lamp2b (ab118959) was obtained from Abcam (USA). Control siRNA and Keap1 siRNAs were purchased from RiboBio (China). The Keap1 cDNA was obtained from the Public Protein/Plasmid Library (China).

Animals

Three- and seven-week-old healthy female unmated Sprague-Dawley (SD) rats were purchased from SPF (Beijing) Biotechnology Co., Ltd. Feeding conditions: room temperature of 22–24℃, relative humidity of 45–60%, natural light, and air circulation. The rats had free access to water and food in the rearing cage, and the bedding was changed every three days. After one week of acclimatization, eight-week-old healthy female rats with a normal estrous cycle were included in the experiment. Twenty-one-day-old SD female rats, weighing approximately 40–50 g, were also purchased from SPF (Beijing) Biotechnology Co., Ltd. The rats were housed at a room temperature of 22–24℃, relative humidity of 50%, and natural light, with free access to food and water in the cage. Surgeries were performed under intraperitoneal injection of 1% pentobarbital sodium.

Ovarian granulosa cell extraction

Three-week-old female Sprague-Dawley (SD) rats were injected intraperitoneally with 80 IU of pregnant horse serum and euthanized after 48 h. The ovaries were removed and rinsed three times in pre-cooled phosphate-buffered saline (PBS), then placed in Petri dishes containing pre-cooled medium (Dulbecco’s Modified Eagle Medium/Nutrient Mixture F-12 (DMEM/F12) + 10% fetal bovine serum (FBS)). The follicles were punctured with a sterile 1 mL syringe needle under a stereo somatotropic microscope, and the follicular cavities were rinsed with the medium. The liquid was filtered through a sterile 200-mesh copper mesh, and the filtered ovarian tissue was centrifuged at 1000 rpm for 5 min. The supernatant was discarded, and the pellet was washed once with PBS. Next, 2 mL of 0.2% hyaluronidase was added, and digestion was performed at 37℃ for 30 min. The sample was then centrifuged at 1000 rpm for 5 min, and the supernatant was discarded. Complete medium (DMEM/F12 + 15% FBS + 1% Minimum Essential Medium (MEM) non-essential amino acids 100× + 1% Insulin-Transferrin-Selenium (ITS) 100× + 1% penicillin 100×) was added, and the cells were gently resuspended. The resuspended cells were counted using a cytometer plate, the concentration was adjusted, and the cells were uniformly inoculated into 6-well plates.

CCK-8 assay

Cells were passaged into 96-well plates at a density of 5 × 10³ cells/well. After 24 h, ovarian granulosa cells were treated with different concentrations of cisplatin and ulinastatin. After 48 h, the medium was discarded, and 100 µL of Cell Counting Kit-8 (CCK-8) reagent was added. Following a 2-hour incubation, absorbance was measured at 450 nm. Cell viability was calculated, and the optimal drug concentration was determined.

ELISA assay

Ovarian tissue was collected and cut into small pieces using sterile shears. One milliliter of lysis buffer was added to the tissue, which was then homogenized by shaking. The homogenate was centrifuged at 15,000 rpm at 4 °C, and the supernatant was collected for analysis. The levels of follicle-stimulating hormone (FSH), estradiol (E2), anti-Müllerian hormone (AMH), superoxide dismutase (SOD), glutathione (GSH), and malondialdehyde (MDA) were measured according to the instructions of the respective enzyme-linked immunosorbent assay (ELISA) kits. Optical density (OD) values were measured at 450 nm, and the different levels were calculated, with each measurement repeated three times. The concentration of each indicator was determined based on the standard protein concentration curve provided in the ELISA kit.

qPCR assay

Total RNA was extracted using the Trizol method, and RNA purity and concentration were assessed by spectrophotometry. The RNA was reverse-transcribed into cDNA using the SPARKscript II All-in-One RT SuperMix for quantitative polymerase chain reaction (qPCR) (with genomic DNA (gDNA) Eraser) Reverse Transcription Kit. Relative fluorescence was then detected using 2× SYBR Green qPCR Mix (with ROX) in a PCR reaction kit. The experiment was repeated three times. The primer sequences for human and rat Keap1, Nrf2, TGFβ-1, FN, C-caspase3, BAX, Bcl2, and β-actin are listed in Table 1.

Table 1.

Primers were used for our experiment

genetics	Positive (5′-3′)	Reverse (5′-3′)
keap1	CGGGGACGCAGTGATGTATG	TGTGTAGCTGAAGGTTCGGTTA
TGFβ-1	CCACCTGCAAGACCATCGAC	CTGGCGAGCCTTAGTTTGGAC
FN	GCTCAGCAAATCGTGCAGC	CTAGGTAGGTCCGTTCCCACT
C-caspase3	TGAAGGGGTCATTTATGGGACA	CCAGTCAGACTCCGGCAGTA
BAX	AGACAGGGGCCTTTTTGCTAC	AATTCGCCGGAGACACTCG
Bcl2	GAGAGCGTCAACAGGGAGATG	CCAGCCTCCGTTATCCTGGA
β-actin	GGCTGTATTCCCCTCCATCG	CCAGTTGGTAACAATGCCATGT

Western blot

Frozen ovarian tissue was lysed in radioimmunoprecipitation assay (RIPA) digestion buffer. Protein concentrations in the supernatants were determined using the bicinchoninic acid (BCA) protein analysis kit after centrifugation. Proteins in equal quantities were electrophoretically separated in a sodium dodecyl sulfate-polyacrylamide gel (SDS-PAGE) and subsequently transferred to a nitrocellulose (NC) membrane. The membranes were blocked with Rapid Closure Solution for 45 min, followed by incubation with primary antibodies against Keap1, Nrf2, TGFβ-1, FN, C-caspase3, BAX, Bcl2 and β-actin overnight at 4 °C. The membranes were washed with tris-buffered saline with Tween-20 (TBST) and then incubated with the primary antibodies for 1 h at 4 °C. After washing with TBST, the membranes were incubated with horseradish peroxidase (HRP)-labeled anti-rabbit antibody for 2 h at 25 °C. Finally, the membranes were analyzed using a developer.

ROS detection

Cells were passaged into 12-well plates at a density of 2 × 10⁵ cells/well and cultured in an incubator for 24 h. Then, 1 mL of serum-free DMEM was added, followed by PBS containing different concentrations of cisplatin and ulinastatin. After 48 h, 1 mL of DCFH-DA staining solution (10 µM) was added, and the cells were incubated for 30 min in 5% CO₂ at 37℃ in the dark. The cells were then washed three times with PBS. The coverslips containing the cells were inverted onto slides, and the cells were finally observed under a fluorescence microscope.

Immunofluorescence assay

Cells were seeded into 12-well plates with sterile coverslips and incubated at 37 °C in a constant-temperature incubator containing 5% CO₂ for 24 h. When the cells reached a density of 60-70%, 1 mL of complete medium containing different drugs was added, and incubation was continued for 48 h. After 48 h, the medium was removed, and 500 µL of 4% paraformaldehyde was added to each well for 30 min to fix the cells. Following fixation, the wells were washed three times with PBS, the paraformaldehyde was discarded, and 1 mL of immunostaining blocking solution was added to each well for 1 h. After aspirating the blocking solution and washing three times with PBS, 1 mL of diluted primary antibody was added to each well and incubated overnight on a shaker at 4 °C, with the shaker speed set to 20 rpm. Following incubation, the wells were washed three times with PBS. Subsequently, the cells were incubated for 1 h at room temperature in the dark with an Alexa Fluor 488-conjugated secondary antibody. After incubation, the cells were washed three times with PBS in the dark. Coverslips were carefully removed using forceps, and 10 µL of DAPI staining solution was added. Images were captured in a darkroom using a fluorescence microscope.

Animal experiment

A rat model of cisplatin-induced ovarian injury was established by intraperitoneal injection of cisplatin at a concentration of 1.5 mg/kg for six consecutive days, while the control group received an equal volume of saline. For pharmacological treatment, 15 mg/kg of ulinastatin (low-dose treatment group) and 30 mg/kg of ulinastatin (high-dose treatment group) were administered via intraperitoneal injection on days 7 and 14. A total of 48 female SD rats were randomly divided into four groups of 12 rats each: control, model, low-dose treatment, and high-dose treatment groups. On days 14 and 35 of the experiment, serum, uterine tissue, and ovarian tissue were collected and fixed for hematoxylin and eosin (H&E) staining, Masson’s staining, and RNA and protein level analysis (Figure S1A).

After H&E staining, morphological changes in ovarian and uterine tissues at various time points were examined under a light microscope. Ovarian follicles were classified and counted according to the method described by Myers et al. [18]. The criteria for identifying follicles at different developmental stages were as follows:

(1) Primordial follicles: Located in the superficial layer of the ovarian cortex, consisting of a primary oocyte surrounded by a single layer of flattened pre-granulosa cells. (2) Primary follicles: Comprising a primary oocyte surrounded by a single layer of cuboidal granulosa cells. When both flattened and cuboidal granulosa cells are present, and the latter predominates, the follicle is classified as a primary follicle. (3) Secondary follicles: Consist of an oocyte surrounded by two or more layers of cuboidal granulosa cells, without a follicular cavity. (4) Mature follicles: The largest among all stages, characterized by a substantial increase in follicular fluid that displaces the oocyte and surrounding granulosa cells to one side.

Fresh ovarian tissues were prepared as frozen sections, then thawed and incubated with DHE dye at 37 ℃ in the dark for 30 min. The slides were subsequently placed in PBS (pH 7.4) and washed three times in the dark on a decolorization shaker, each wash lasting 5 min. After gently drying the sections, DAPI staining solution was added, followed by incubation at room temperature in the dark for 10 min. The slides were then placed in PBS and washed three additional times on a decolorization shaker in the dark, each for 5 min. After gently drying the sections again, they were mounted with an anti-fluorescence quenching mounting medium. Finally, the slides were observed and photographed using a fluorescence microscope.

Statistical analysis

Statistical significance between groups was determined using GraphPad Prism, version 8.0 (GraphPad Software, San Diego, CA). Data were presented as mean ± SEM for all experiments and analyzed using Student’s t-test or one-way analysis of variance (ANOVA) followed by Dunnett’s post hoc test. A P < 0.05 was considered statistically significant.

Results

Protective effect of ulinastatin against ovarian damage

After inducing ovarian injury with cisplatin, the ovaries were observed for 35 days. On both days 14 and 35, the control group ovaries exhibited an ovoid shape with vesicular elevations of varying sizes on the surface. In contrast, the ovaries in the model group were significantly smaller and had fewer vesicular elevations compared to the control group. There was no significant difference in ovarian size between the ulinastatin low-dose treatment group and the model group. However, the ovaries in the ulinastatin high-dose treatment group were slightly larger than those in the model group, although still smaller than those in the control group (Fig. 1A). Histological analysis using H&E and Masson’s staining revealed that on day 14 after cisplatin administration, the control group displayed a normal ovarian structure with a well-defined corpus luteum, a higher number of follicles, a thicker granulosa cell layer, orderly cell arrangement, and no signs of degeneration, inflammatory cell infiltration, or fibrotic tissue proliferation in the ovarian interstitium (Fig. 1C and E). In contrast, the model group exhibited severe ovarian damage, characterized by a significant reduction in the number of normal follicles and an increase in atretic follicles, as indicated by yellow arrowheads. Additionally, a large number of inflammatory cells and proliferating fibrotic tissues were observed in the ovarian interstitium, as shown by red arrows. The ulinastatin low-dose treatment group showed a slight improvement compared to the model group, with an increased number of normal follicles, a reduction in atretic follicles, and a decrease in inflammatory cells and fibrotic tissue within the interstitium. The ulinastatin high-dose treatment group exhibited a further reduction in ovarian damage, with a notable decrease in inflammatory cells and fibrotic tissue. On day 35 after cisplatin administration, the control group continued to maintain a normal ovarian structure (Fig. 1D-E). However, the model group showed increased ovarian damage, ovarian atrophy, and a large number of atretic follicles, as indicated by yellow arrows. These changes were accompanied by extensive inflammatory cell infiltration and fibrotic tissue hyperplasia, as shown by red arrowheads. In the ulinastatin low-dose treatment group, ovarian damage was significantly alleviated, with a largely normal follicular structure and only a small number of inflammatory cells and fibrotic tissues in the interstitium. The high-dose ulinastatin treatment group exhibited the least damage, with minimal inflammation and fibrosis. In addition, for the comparison of the number of follicles at all levels, it was found that the number of primordial follicles, primary follicles, secondary follicles and mature follicles in the model group was significantly lower than that in the control group (P < 0.05); while the number of follicles at all levels in the ulinastatin-treated group was lower than that in the control group, but significantly higher than that in the model group (P < 0.05) (Fig. 1F). These findings demonstrate that ulinastatin treatment significantly improved cisplatin-induced ovarian structural damage.

Fig. 1.

Ulinastatin ameliorates ovarian tissue damage. (A) Comparison of the gross appearance of rat ovaries on days 14 and 35 using cisplatin. (B) Comparison of the gross appearance of rat uterus on days 14 and 35 using cisplatin. (C-D) H&E staining results of rat ovaries treated with cisplatin on days 14 (C) and 35 (D). The yellow arrow indicates a decrease in the number of normal follicles and an increase in the number of atretic follicles; the red arrow indicates the presence of a large number of inflammatory cells and fibrous tissue proliferation in the stroma. (E) Masson staining results of rat ovaries on days 14 and 35 following cisplatin treatment. Increased blue staining indicates elevated collagen fiber content in the tissue and a higher degree of fibrosis. (F) Quantitative analysis of primordial, primary, secondary, and maturing follicles per slide in each group. (G-H) H&E staining results of rat uterus treated with cisplatin on days 14 (G) and 35 (H). The black arrow indicates uterine cavity dilatation and epithelial cell apoptosis; the yellow arrow indicates inflammatory cell infiltration in the lamina propria; the green arrow indicates visible neovascularization. Scale bar = 200 and 50 μm. Compared with the model group, * * * P < 0.001, * * P < 0.01, * P < 0.05

Additionally, we examined the appearance and histomorphology of the rat uterus. In the model group, although the uterus retained a Y-shape, the uterine horns were significantly thinner compared to the control group (Fig. 1G-H). In contrast, ulinastatin treatment resulted in a significant thickening of the uterine horns. H&E staining of uterine tissues showed that on day 14, the control group exhibited a normal uterine structure, with an undilated uterine cavity, closely and orderly arranged epithelial cells, regular cytoplasmic morphology without apparent degeneration, and a greater number of densely distributed glands in the lamina propria without inflammatory cell infiltration. In the model group, the uterine cavity was severely dilated, with vacuolar degeneration and cytoplasmic loosening of epithelial cells in some areas, as indicated by red arrows. A small number of apoptotic cells were observed, characterized by fixed and deeply stained nuclei (black arrows); the lamina propria appeared slightly loose, with mild infiltration of inflammatory cells (yellow arrows). In the low-dose ulinastatin group, the degree of tissue damage was ameliorated compared to the model group. Although the uterine cavity remained dilated, the number of degenerated epithelial cells was reduced (black arrows), and mild inflammatory cell infiltration was still observed in the lamina propria (yellow arrows). The high-dose ulinastatin group exhibited a further improvement, with reduced uterine cavity dilation and fewer degenerated epithelial cells and inflammatory cells in the lamina propria. By day 35, the control group maintained a normal uterine structure without evident pathological changes. In contrast, tissue damage in the model group was exacerbated, characterized by uterine cavity dilation, extensive epithelial cell apoptosis, deeply stained nuclei, loss of cellular contours (black arrows), loosening of the lamina propria structure, inflammatory cell infiltration (yellow arrows), and marked neovascularization (green arrows). In the low-dose ulinastatin group, the extent of injury was significantly reduced, with only a few degenerated epithelial cells, dense lamina propria structure, and occasional inflammatory cell infiltration. In the high-dose ulinastatin group, the uterine structure appeared nearly normal, with epithelial cells arranged in a closely packed and orderly manner, no evident degeneration, and only sporadic inflammatory cell infiltration in the lamina propria.

Effect of ulinastatin on ovarian function

To evaluate the effect of ulinastatin on ovarian function in cisplatin-induced ovarian injury, serum levels of FSH, E2, and AMH were measured in each group of rats at different time points (Fig. 2A-C). The results showed that in the in vivo experiments assessing the repair of cisplatin-induced ovarian injury by ulinastatin, there were no significant differences in FSH, E2, and AMH levels among the groups before cisplatin injection on day 1 (P > 0.05). By day 7, before treatment, E2 and AMH levels were significantly lower, while FSH levels were significantly higher in both the treatment and model groups compared to the control group. Moreover, E2 and AMH levels were significantly lower, and FSH levels were significantly higher in the model group compared to the control group. However, no significant difference was observed between the treatment and model groups (P > 0.05). The therapeutic effect of ulinastatin became evident on days 14, 28, and 35 after cisplatin administration. E2 and AMH levels were significantly higher, and FSH levels were significantly lower in the treatment group compared to the model group (P < 0.05 or P < 0.01) (Fig. 2). These results indicate that ulinastatin improves ovarian function in cisplatin-induced ovarian injury.

Fig. 2.

Ulinastatin improves ovarian function. (A) Serum E2 levels in rats in each group after treatment with ulinastatin. (B) Serum FSH levels in rats treated with ulinastatin. (C) Serum AMH levels in rats treated with ulinastatin. (D) TUNEL staining of ovarian tissues of rats treated with ulinastatin. Scale bar = 20 μm. * * * P < 0.001, * * P < 0.01, * * P < 0.05 compared with the model group

To investigate whether cisplatin accelerates follicular atresia and impairs ovarian function by inducing ovarian cell apoptosis, the apoptosis of ovarian tissue cells in each group was assessed on days 14 and 35 after cisplatin administration using the TUNEL assay (Fig. 2D). Fluorescence microscopy revealed minimal apoptosis in the ovarian tissue cells of the control group. In contrast, the model group exhibited a significant increase in apoptotic ovarian tissue cells (green fluorescence represents apoptotic cells). However, apoptosis was significantly reduced in the ulinastatin treatment group (Fig. 2). These results suggest that ulinastatin treatment effectively inhibits cisplatin-induced apoptosis in ovarian tissues.

Effect of ulinastatin on the antioxidant action of the ovary

Ulinastatin is a broad-spectrum serine protease inhibitor with a strong ability to regulate oxidative stress. Cisplatin promotes oxidative stress, ultimately leading to apoptosis. To investigate this phenomenon, we analyzed the inhibitory effect of ulinastatin on cisplatin-induced oxidative stress in the ovary. First, the levels of MDA, SOD, and GSH in ovarian tissues were measured (Fig. 3A-C). The results showed that in the model group, SOD and GSH levels were significantly decreased, while MDA levels were significantly increased on day 14 after cisplatin administration. These changes were significantly alleviated by ulinastatin treatment, and this trend persisted until day 35 of cisplatin administration. Subsequently, fluorescence detection of ROS in ovarian tissues was performed (Fig. 3D). The results indicated that the ROS positivity rate in the model group increased significantly on day 14 after cisplatin administration, while the addition of ulinastatin markedly suppressed ROS production. On day 35 of cisplatin administration, the ROS positivity rate in the model group increased even further, whereas ulinastatin treatment effectively maintained ROS levels within a lower range. These findings suggest that cisplatin induces oxidative stress in the ovary and that ulinastatin significantly inhibits this oxidative stress response.

Fig. 3.

Ulinastatin has antioxidant properties. (A) SOD levels in the ovarian tissues of rats in each group after treatment with ulinastatin. (B) GSH levels in ovarian tissues of rats treated with ulinastatin. (C) MDA levels in ovarian tissues of rats treated with ulinastatin. (D) ROS staining of ovarian tissues of rats treated with ulinastatin. Scale bar = 20 μm. * * * P < 0.001, * * P < 0.01, * P < 0.05 compared with the model group

Under physiological conditions, Nrf2 is coupled to its inhibitor protein Keap1 and bound to the actin cytoskeleton in an inactive state. However, under oxidative stress, Nrf2 is activated, translocated into the nucleus after uncoupling from Keap1, and binds with other proteins to form a heterodimer. This heterodimer then binds to the antioxidant response element (ARE), activating the downstream expression of antioxidant genes such as HO-1, NQO1, and SOD, thereby maintaining redox homeostasis [14, 15]. Therefore, Nrf2 activation is an effective strategy to inhibit oxidative stress and prevent fibrosis. It has been found that cisplatin inhibits Nrf2 activation. Additionally, our previous results demonstrated that cisplatin increased ovarian fibrosis, while the addition of ulinastatin reduced fibrosis levels. Therefore, we investigated whether ulinastatin alleviates cisplatin-induced ovarian damage and oxidative stress by activating Nrf2. First, we examined the expression of antioxidant and apoptosis-related proteins at the mRNA level. The results showed that in the model group, Keap1, TGFβ-1, FN, cleaved-caspase3, and BAX expression levels were significantly increased from day 14 of cisplatin administration, whereas Nrf2 and Bcl2 expression levels were significantly decreased (Fig. 4A). The addition of ulinastatin reversed these changes, producing the opposite effect. Similar results were observed on day 35 of cisplatin administration (Fig. 4B). Subsequently, immunohistochemistry and Western blot (WB) analysis were performed on ovarian tissues, yielding results consistent with the mRNA levels (Fig. 4C-E). These findings suggest that ulinastatin significantly inhibits oxidative stress and reduces cisplatin-induced ovarian injury, a process that may be mediated through the Nrf2/Keap1 pathway.

Fig. 4.

Ulinastatin regulates antioxidant and apoptosis-related proteins. (A) qPCR results of Keap1, Nrf2, TGFβ-1, FN, Cleaved-caspase3, BAX, and Bcl2 in cells of each group after treatment with ulinastatin on day 14 using cisplatin. (B) qPCR results of Keap1, Nrf2, TGFβ-1, FN, Cleaved-caspase3, BAX, and Bcl2 in cells of each group after treatment with ulinastatin on day 35 using cisplatin. (C) WB results of Keap1, Nrf2, TGFβ-1, FN, Cleaved-caspase3, BAX, and Bcl2 in cells of each group after treatment with ulinastatin on days 14 and 35 using cisplatin. (D) Immunohistochemical results of Keap1, Nrf2, TGFβ-1, and FN in cells of each group after treatment with ulinastatin on day 14 of cisplatin use. (E) Immunohistochemical results of Keap1, Nrf2, TGFβ-1, and FN in cells of each group after treatment with ulinastatin on day 35 of cisplatin use. Scale bar = 20 μm. Compared with the model group, * * * P < 0.001, * * P < 0.01, * P < 0.05

Effect of ulinastatin on ovarian damage and antioxidant effects

To verify the pathway through which ulinastatin repairs cisplatin-induced ovarian damage, ovarian granulosa cells were first extracted. Subsequently, the ovarian function was analyzed after the addition of cisplatin and ulinastatin to the granulosa cells. The results showed that after cisplatin treatment, E2 and AMH levels were significantly decreased, while FSH levels were significantly increased (Fig. 5A-C). However, with the addition of ulinastatin, E2 and AMH levels increased, and FSH levels decreased. Next, the antioxidant effects of ulinastatin were analyzed. The results demonstrated that cisplatin treatment significantly decreased SOD and GSH levels while increasing MDA levels in the cells (Fig. 5D-F). In contrast, the addition of ulinastatin increased SOD and GSH levels while reducing MDA levels. Furthermore, ROS levels were examined, and staining results showed that cisplatin treatment significantly increased ROS release, whereas the addition of ulinastatin significantly inhibited ROS production (Fig. 5G). The expression of antioxidant and apoptosis-related proteins was assessed at the mRNA and protein levels (Fig. 5H-I). The results indicated that in cisplatin-treated cells, the expression of Keap1, TGFβ-1, FN, cleaved-caspase3, and BAX was significantly upregulated, while the expression of Nrf2 and Bcl2 was significantly downregulated. However, the addition of ulinastatin reversed these changes, yielding results similar to those of the control group. These findings suggest that ulinastatin effectively inhibits oxidative stress and alleviates cisplatin-induced ovarian damage.

Fig. 5.

Ulinastatin ameliorates ovarian granulosa cell damage. (A) Levels of E2 secreted in ovarian granulosa cells after treatment with ulinastatin. (B) Levels of FSH secreted in ovarian granulosa cells after treatment with ulinastatin. (C) Levels of AMH secreted in ovarian granulosa cells after treatment with ulinastatin. (D) SOD levels in ovarian granulosa cells after treatment with ulinastatin. (E) GSH levels in ovarian granulosa cells after treatment with ulinastatin. (F) MDA levels in ovarian granulosa cells after treatment with ulinastatin. (G) ROS staining in ovarian granulosa cells after treatment with ulinastatin. Scale bar = 20 μm. (H) qPCR results of Keap1, Nrf2, TGFβ-1, FN, Cleaved-caspase3, BAX, and Bcl2 in ovarian granulosa cells treated with ulinastatin. (I) WB results of Keap1, Nrf2, TGFβ-1, FN, Cleaved-caspase3, BAX, and Bcl2 in ovarian granulosa cells after treatment with ulinastatin. Compared with the model group, * * * P < 0.001, * * P < 0.01

Ovarian protection by ulinastatin through the Nrf2/Keap1 pathway

After confirming that the effects of ulinastatin in ovarian granulosa cells were consistent with those observed in the ovary, we further analyzed the role of the Nrf2/Keap1 pathway in ulinastatin-mediated repair of cisplatin-induced ovarian damage. First, the role of ulinastatin was examined by silencing the Keap1. The results showed that Keap1 silencing further increased E2 and AMH levels and decreased FSH levels, suggesting that Keap1 plays an inhibitory role in ulinastatin-mediated ovarian repair (Fig. 6A-C). Since Nrf2 is negatively regulated by Keap1, this also indicates that Nrf2 promotes the action of ulinastatin. Next, changes in antioxidant effects were assessed. Keap1 silencing further increased SOD and GSH levels and decreased MDA levels in the ulinastatin-treated group (Fig. 6D-F). Additionally, ROS staining results showed that Keap1 silencing further reduced ROS release in the ulinastatin-treated group (Fig. 6G). At the mRNA and protein levels, Keap1 silencing further reduced the levels of apoptotic proteins, decreased the levels of TGFβ-1 and FN, and increased the levels of Nrf2 in the ulinastatin-treated group (Fig. 6H-I). These results suggest that Keap1 silencing enhances the repair capacity and antioxidant ability of ulinastatin.

Fig. 6.

Activation of Nrf2 promotes the protective effect of ulinastatin on ovarian granulosa cells. (A) Levels of E2 secreted in cells in each group after silencing Keap1. (B) Levels of FSH secreted in cells after silencing Keap1. (C) Levels of AMH secreted in cells after silencing Keap1. (D) SOD levels in cells of each group after silencing Keap1. (E) GSH levels in cells after silencing Keap1. (F) MDA levels in cells after silencing Keap1. (G) ROS staining results in Keap1-silenced cells. Scale bar = 20 μm. (H) qPCR results of Keap1, Nrf2, TGFβ-1, FN, Cleaved-caspase3, BAX, and Bcl2 in cells of each group after silencing Keap1. (I) WB results of Keap1, Nrf2, TGFβ-1, FN, Cleaved-caspase3, BAX and Bcl2 in cells of each group after silencing Keap1. Compared with the model group, * * * P < 0.001, * * P < 0.01. Between groups, # P < 0.05

Next, the effects of ulinastatin were analyzed by overexpressing Keap1. The results showed that Keap1 overexpression significantly inhibited the ovarian repair effects of ulinastatin (Fig. 7A-C). Additionally, Keap1 overexpression markedly suppressed the antioxidant effects of ulinastatin, as indicated by the levels of SOD, GSH, and MDA (Fig. 7D-F), as well as ROS detection (Fig. 7G). At the mRNA and protein levels, Keap1 overexpression significantly increased the expression of apoptotic proteins, elevated the levels of TGFβ-1 and FN, and decreased Nrf2 expression (Fig. 7H-I). These findings suggest that Keap1 overexpression inhibits the repair and antioxidant capacities of ulinastatin. The combined results from Keap1 silencing and overexpression further indicate that the inhibition of oxidative stress and reduction of cisplatin-induced injury by ulinastatin are closely related to the Nrf2/Keap1 pathway.

Fig. 7.

Inhibition of Nrf2 reduces the protective effect of ulinastatin on ovarian granulosa cells. (A) Levels of E2 secreted in cells of each group after overexpression of Keap1. (B) Levels of FSH secreted in cells after overexpression of Keap1. (C) Levels of AMH secreted in cells after overexpression of Keap1. (D) SOD levels in cells after overexpression of Keap1. (E) GSH levels in cells after overexpression of Keap1. (F) MDA levels in cells after overexpression of Keap1. (G) ROS staining results in cells after overexpression of Keap1. Scale bar = 20 μm. (H) qPCR results of Keap1, Nrf2, TGFβ-1, FN, Cleaved-caspase3, BAX, and Bcl2 in cells after overexpression of Keap1. (I) WB results of Keap1, Nrf2, TGFβ-1, FN, Cleaved-caspase3, BAX, and Bcl2 in cells of each group after overexpression of Keap1. Compared with the model group, * * * P < 0.001, * * P < 0.01. Between groups, # P < 0.05

Discussion

The gonadal toxicity of antitumor therapy in young women is a problem that cannot be ignored. Research on reducing the gonadal toxicity of chemotherapeutic drugs has been a major focus of clinical concern. Current studies have identified three main pathways through which chemotherapeutic drugs induce ovarian damage: they can directly damage primordial follicles or indirectly reduce their number by damaging growing follicles [7]; they can directly cause oocyte damage or indirectly damage oocytes by affecting granulosa cells [19, 20]; and they can damage the ovarian mesenchyme and vasculature, leading to the loss of follicles at all developmental stages [9, 21]. Among these mechanisms, DNA damage and apoptosis of oocytes and granulosa cells, fibrosis of the ovarian interstitium, and increased activation of primordial follicles may be important mechanisms by which chemotherapeutic agents cause a decrease in ovarian reserve [7, 22–24].

Cisplatin, as a commonly used chemotherapeutic agent in the treatment of gynecological malignancies, has significant gonadotoxic effects [25]. Due to the ovarian toxicity of cisplatin, young female patients often experience amenorrhea, POF, and infertility after treatment, which seriously affects their quality of life [26]. Some small-molecule antioxidants have been found to have a protective effect against cisplatin-induced ovarian damage. Ulinastatin, a protease inhibitor with anti-inflammatory, antioxidant, and anti-apoptotic properties, has been widely used for the repair and protection of various organ injuries. For example, it alleviates lung and kidney injuries by inhibiting inflammatory responses and oxidative stress [27, 28] and protects liver and cardiac function by inhibiting apoptosis and reducing the release of inflammatory mediators [29, 30]. Therefore, theoretically, ulinastatin may prevent chemotherapy-induced ovarian damage, but relevant studies remain limited.

In the present study, we observed that cisplatin significantly reduced the number of normal follicles in the ovary, increased the number of atretic follicles, and triggered interstitial inflammation and fibrosis. These results are consistent with previous studies, which have shown that cisplatin can lead to the destruction of ovarian structures by inducing oxidative stress and apoptosis [31, 32]. In contrast, treatment with ulinastatin significantly improved ovarian structure, reduced the number of atretic follicles, and suppressed inflammation and fibrosis. This finding aligns with the protective effects of ulinastatin observed in other tissues, suggesting its broad anti-inflammatory and antifibrotic properties.

Ovarian damage induced by chemotherapeutic agents is typically characterized by decreased serum E2 and AMH levels and abnormally elevated FSH levels. Therefore, the effects of protective drugs on chemotherapy-induced ovarian injury are usually evaluated based on these serum hormone levels. In the present study, by detecting serum FSH, E2, and AMH levels, we found that ulinastatin significantly decreased FSH levels and significantly increased E2 and AMH levels, thereby ameliorating cisplatin-induced ovarian impairment. In addition, results from the TUNEL assay and protein expression analysis demonstrated that ulinastatin significantly reduced cisplatin-induced ovarian cell apoptosis, further supporting its role in protecting ovarian function. Furthermore, beyond inducing apoptosis, cisplatin also elevated MDA levels, reduced SOD and GSH levels, and significantly increased ROS levels in ovarian tissues by inducing oxidative stress. In contrast, ulinastatin treatment effectively reversed these changes, demonstrating its potent antioxidant capacity.

The Nrf2/Keap1 pathway is a crucial intracellular anti-oxidative stress signaling pathway that maintains intracellular redox homeostasis by regulating the expression of downstream antioxidant genes. In this study, we found that cisplatin-induced ovarian injury was associated with inhibition of the Nrf2 signaling pathway and elevated oxidative stress levels. In contrast, ulinastatin significantly activated the Nrf2 pathway and reduced the expression of Keap1, leading to enhanced antioxidant capacity and attenuation of oxidative stress-induced damage in ovarian tissues.

To further verify the mechanism of action of ulinastatin, this study isolated and cultured rat ovarian granulosa cells in vitro. Based on cellular characterization, a stable in vitro model of cisplatin-induced ovarian granulosa cell injury was established to explore the reparative effects of ulinastatin on cisplatin-induced ovarian injury. The results showed that ulinastatin significantly increased E2 and AMH levels, decreased FSH levels in ovarian granulosa cells, and inhibited cisplatin-induced oxidative stress and apoptosis.

Subsequently, we further explored the mechanism by which ulinastatin repairs cisplatin-induced ovarian damage through the Nrf2/Keap1 pathway by silencing and overexpressing Keap1. Keap1 is a negative regulator of Nrf2, inhibiting Nrf2 activity by binding to it and promoting its ubiquitination and degradation under normal conditions. However, under oxidative stress, Nrf2 dissociates from Keap1, translocates into the nucleus, and activates the expression of downstream antioxidant genes, thereby exerting antioxidant and anti-apoptotic effects [33].

We found that the protective effect of ulinastatin was further enhanced by silencing Keap1. After Keap1 silencing, Nrf2 activity was significantly increased, leading to the upregulation of downstream antioxidant genes (e.g., HO-1, NQO1, SOD), which resulted in a significant reduction in oxidative stress levels in ovarian tissues. This was evidenced by a significant increase in SOD and GSH levels and a significant decrease in MDA and ROS levels. In addition, Keap1 silencing significantly reduced apoptosis in ovarian tissues. These results are consistent with previous studies, suggesting that Nrf2 activation plays a key role in antioxidant and anti-apoptotic mechanisms. To further validate the role of the Nrf2/Keap1 pathway in ulinastatin-mediated protection, we found that Keap1 overexpression significantly inhibited its effects. Keap1 overexpression resulted in decreased Nrf2 activity, downregulated antioxidant gene expression, and a significant increase in oxidative stress levels. Additionally, Keap1 overexpression significantly increased apoptosis in ovarian tissues. These findings further confirm the critical role of the Nrf2/Keap1 pathway in the protective effects of ulinastatin.

In summary, this study confirmed the significant role of ulinastatin in repairing cisplatin-induced ovarian damage and elucidated its mechanism of regulating oxidative stress and apoptosis through the Nrf2/Keap1 pathway. These findings provide a theoretical basis for the clinical application of ulinastatin, particularly in protecting ovarian function in chemotherapy patients, highlighting its important potential value.

Electronic supplementary material

Below is the link to the electronic supplementary material.

Abbreviations

SD

Sprague-Dawley

AMH

Anti-Müllerian hormone

E2

Estradiol

FN

Fibronectin

FSH

Follicle-stimulating hormone

GSH

Glutathione

Keap1

Kelch-like ECH-associated protein 1

MDA

Malondialdehyde

Author contributions

Conceived and designed the experiments: Zeming Fu and Yiquan Li. Performed the experiments: Liping Zhao , Yishi Wu, Xiwen Zhang and Yaru Li. Analyzed the data: Yishi Wu and Xiwen Zhang. Contributed reagents/materials/analysis tools: Yaru Li and Yiquan Li. Wrote the paper: Liping Zhao and Yiquan Li. Revised the paper: Liping Zhao and Zeming Fu. All authors read and approved the final manuscript.

Funding

This work was supported by the Natural Science Foundation of Jilin Province (Grant No. YDZJ202201ZYTS293).

Data availability

No datasets were generated or analysed during the current study.

Declarations

Ethics approval and consent to participate

The animal study was reviewed and approved by the Animal Experiment Ethics Committee of Jilin University (Approval No. SY2025-07-008). All applicable international, national, and/or institutional guidelines for the care and use of animals were followed.

KT202103225 is the animal ethics approval number for our preliminary experiments conducted during the project（Grant No. YDZJ202201ZYTS293） application phase. SY2025-07-008 is the animal ethics approval number for our formal animal experiments.

Consent for publication

Not applicable.

Competing interests

The authors declare no competing interests.