Proteomic analysis of cisplatin-induced spermatogenesis defects in mice

Abstract

Background

Cisplatin is a crucial chemotherapeutic agent used for treating various cancers; however, its excessive use can cause irreversible damage to the reproductive system, and the protein expression profile of cisplatin-induced testicular injury remains unclear.

Methods

Male C57BL/6 mice were treated with cisplatin at various doses, and testes were collected for histological, immunofluorescence, and proteomic analyses. Germ cell loss and apoptosis were assessed using H&E staining, TUNEL assays, and immunofluorescence for LIN28A, SYCP3, MVH, and CDK1. Label-free quantitative proteomics identified differentially expressed proteins, which were analyzed for functional enrichment and protein–protein interactions.

Results

We observed that cisplatin treatment led to smaller testes, reduced sperm count, and a significant decrease in the number of spermatocytes and spermatids in mice. Label-free quantitative proteomic analysis revealed that cisplatin significantly reduced the expression of cyclin-dependent kinase 1 (CDK1), a key spermatogenesis regulator, in the testes. Reduction in CDK1 expression is correlated with spermatogenic arrest, particularly in spermatocytes.

Conclusion

These findings highlight the critical role of CDK1 in cisplatin-induced spermatogenic dysfunction and provide new insights into fertility preservation strategies for patients with cancer undergoing chemotherapy.

Introduction

Maintenance of male fertility relies on spermatogenic homeostasis in the testes [1]. Various medical interventions, such as radiotherapy and chemotherapy, can severely disrupt male reproductive homeostasis, leading to irreversible reproductive damage [2,3]. Germ cells in the testes are particularly sensitive to chemotherapeutic agents and pose a risk of lifelong infertility in patients [4]. Although sperm cryopreservation and assisted reproductive technologies offer options for adult males to preserve fertility, these methods are not applicable to prepubertal males that do not produce mature sperm. Due to the unclear gene expression profiles that regulate testicular repair after spermatogenic damage, it remains challenging to predict, prevent, and treat infertility in these patients.

In cancer treatment, single or combined chemotherapeutic agents often cause prolonged azoospermia. Some drugs, such as anthracyclines (e.g. doxorubicin), microtubule inhibitors (e.g. vincristine), and metabolic synthesis inhibitors (e.g. methotrexate), primarily affect differentiated spermatogonia without significantly reducing the number of spermatogonial stem cells (SSCs), leading to temporary reductions in sperm count [4–7]. In contrast, alkylating agents (e.g. busulfan and cyclophosphamide) cause severe spermatogenic damage at high doses, often resulting in permanent azoospermia [8,9]. Cisplatin is a chemotherapeutic agent commonly used to treat various solid tumors including lung, breast, testicular, and ovarian cancers. Its primary mechanism of action involves the formation of irreversible DNA crosslinks in proliferating cells, leading to DNA damage and apoptosis [10,11]. This mechanism makes cisplatin particularly detrimental to testicular spermatogenesis, often resulting in azoospermia and male infertility [12,13]. Numerous studies have shown that cisplatin reduces germ cells in seminiferous tubules, impairs Leydig cell function, and causes abnormal sperm morphology [14,15]. Although the effects of cisplatin on male reproduction are well-known, its biological mechanisms are not well understood. Elucidating the molecular mechanisms underlying cisplatin-induced spermatogenic damage may provide potential strategies for preserving clinical fertility.

Cyclin-dependent kinase 1 (CDK1) is a highly conserved serine/threonine kinase that plays an essential role in regulating cell-cycle progression, particularly the G2/M transition. During spermatogenesis, CDK1 activity ensures proper entry into meiosis and accurate chromosomal segregation [16]. Disruption of CDK1 function causes meiotic arrest, germ cell apoptosis, and infertility in several mammalian models [17–19]. In addition, CDK1 coordinates with cyclins and other checkpoint regulators to maintain genomic stability during the complex chromosomal dynamics of meiotic prophase [20]. Given its pivotal role in meiotic progression and germ cell survival, we hypothesized that cisplatin disrupts meiotic progression in the testes by downregulating key cell-cycle regulators, such as CDK1, ultimately leading to spermatogenic arrest and infertility.

In this study, we investigated the inducement of spermatogenic dysfunction and germ cell loss in mice due to cisplatin, as well as changes in CDK1 and other proteins. Label-free quantitative proteomic analysis revealed a significant reduction in CDK1 and other proteins. Investigating the reproductive side effects of cisplatin and the disruption of spermatogenesis could offer important insights into the rational clinical use of cisplatin.

Materials and methods

Animals

The C57BL/6J mice used in this study were purchased from Charles River. Male C57/BL6 mice aged two months were intraperitoneally injected with cisplatin at doses of 15, 20, 30, or 40 mg/kg, or normal saline (0.9% NaCl solution), based on body weight. Testes were collected 30d after treatment, cleaned, and weighed. No deaths were observed in any of the experimental groups, including the high-dose (40 mg/kg) cisplatin group. A slight decrease in body weight was observed in the 40 mg/kg cisplatin group, but did not exceed 10% of the initial body weight, and the mice showed normal activity levels. No abnormal behaviors were observed in any of the groups. Epididymides were minced in phosphate-buffered saline (PBS) and incubated at 37 °C for 20 min to release the sperm, which were counted under a microscope. All animal experiments were approved by the Committee on Animal Care of the Institute of Basic Medical Sciences, Chinese Academy of Medical Sciences, and Peking Union Medical College. All mice were housed and bred under specific pathogen-free conditions (temperature: 22–26 °C, humidity: 40–55%, 12-h light/dark cycle) in the animal facility at the Institute of Basic Medical Sciences. Mice were randomly assigned to either the control or cisplatin-treated groups. To minimize bias, all outcome assessments were performed by investigators who were blinded to the treatment groups. All animal procedures were approved and conducted in accordance with the guidelines, standards, and regulations of the Chinese Academy of Medical Sciences Animal Care and Use Committee of the Chinese Academy of Medical Sciences (Approval Number: ACUC-XMSB-2024-065). The animal experiments were conducted from 2024 to 2025. Animal studies were performed in compliance with the ARRIVE guidelines. Animals were euthanized by cervical dislocation, following the AVMA Guidelines for the Euthanasia of Animals.

Immunofluorescence histology

Mouse testes were fixed in 4% paraformaldehyde for 24 h at 4 °C, followed by paraffin embedding and sectioning at 5 µm. Antigen retrieval was performed using citrate buffer (pH 6.0) for 20 min at 95 °C. Sections were blocked with 5% BSA and 0.1% Triton X-100 for 1 h at room temperature to reduce nonspecific binding. Primary antibodies used were: anti-LIN28A (R&D Systems, AF3757) and anti-ZBTB16 (Santa Cruz, sc-28319) at a 1:100 dilution, anti-SYCP3 (ab97672; Abcam) and anti-CDK1 (19532-1-AP; Proteintech) at a 1:200 dilution, and anti-MVH (ab27591; Abcam) and anti-PNA (Invitrogen, L32459) at a 1:150 dilution. Primary antibodies were incubated overnight at 4 °C. For secondary antibody detection, Alexa Fluor^® 488 (Thermo Fisher, A-21206), Alexa Fluor^® 594 (Thermo Fisher, A-11058), and Alexa Fluor^® 647 (Thermo Fisher, A-31573) were used at a 1:500 dilution. Membranes were then incubated with secondary antibodies for 1 h at room temperature. Following incubation, the sections were washed thrice in PBS for 10 min each and counterstained with DAPI (Sigma, D3571) for 5 min. Fluorescent images were acquired using a Zeiss 780 confocal microscope.

Western blotting

Testicular proteins were extracted using RIPA lysis buffer supplemented with 1 mM PMSF and protease inhibitors. After sodium dodecyl sulfate–polyacrylamide gel electrophoresis, the proteins were transferred to polyvinylidene fluoride membranes and blocked with 5% non-fat milk for 1 h. Membranes were incubated with primary antibodies (CDK1: Proteintech,19532-1-AP; TUBULIN: Abcam, ab7291) overnight at 4 °C, followed by peroxidase-conjugated secondary antibodies at 37 °C for 1 h. Protein bands were visualized using ECL.

Protein sequencing

Extracted proteins were dissolved in 100 mM ammonium bicarbonate containing 1% sodium deoxycholate, reduced with 5 mM tris (2-carboxyethyl) phosphine at 55 °C for 10 min, and alkylated with 10 mM iodoacetamide in the dark at room temperature for 15 min. Proteins were digested with sequencing-grade trypsin at 37 °C. After SDC removal with 2% trifluoroacetic acid, the tryptic peptides were desalted using a C18 column and dried using a vacuum concentrator. Peptides were separated using an EASY-nLC1200 nanoUPLC system and analyzed using a Q-Exactive mass spectrometer. Raw data were processed using MaxQuant software (version 1.6.1.0). Differentially regulated proteins were identified based on a fold change ≤ 1.5 and p-value ≤ 0.05, and were visualized in volcano plots and cluster heatmaps. Gene ontology (GO) and Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway analyses were performed to predict the biological processes, cellular components, and molecular functions of the differentially expressed proteins in the cisplatin-treated mouse testes. Protein–protein interactions (PPIs) were predicted using the STRING database, and the resulting interaction network was visualized using Cytoscape (version 3.10), which helped to identify key hub proteins involved in cisplatin-induced testicular damage.

Statistical analysis

Sample sizes are described in the figure legends. All statistical analyses were performed using GraphPad Prism 9.0 software (GraphPad Software, San Diego, CA). Prior to statistical testing, data distributions were evaluated for normality using the Shapiro–Wilk test, and homogeneity of variance was assessed using Levene’s test. For comparisons between the two groups, unpaired two-tailed Student’s t-tests were used when the data met normality and equal variance assumptions. In cases where assumptions were not met, non-parametric alternatives (Mann–Whitney U test) were used. Data are presented as mean values ± standard error of the mean.

Results

Cisplatin-induced spermatogenic disruption in mouse testes

To simulate the reproductive damage observed in patients with cancer after chemotherapy, we induced spermatogenic dysfunction in mice using different cisplatin doses and studied protein expression associated with infertility (Figure 1(A)). Following treatment with 15, 20, 30, and 40 mg/kg cisplatin, hematoxylin and eosin staining was performed (Figure 1(B)). The results showed that germ cell loss in the testes increased at higher cisplatin doses, with many seminiferous tubules exhibiting vacuolated structures and cell layer detachment. We measured the cross-sectional area of the seminiferous tubules and found that it decreased with cisplatin-induced damage. In particular, the cross-sectional area of the seminiferous tubules in the 40 mg/kg group was only 30% of that in the control group (Figure 1(C)). The ratio of testis weight to body weight also decreased after cisplatin treatment, indicating impaired testicular development (Figure 1(D)). Additionally, sperm counts decreased progressively with cisplatin treatment. Sperm count in the 40 mg/kg group was 50% of that in the control group (Figure 1(E)). Additionally, to evaluate the impact of cisplatin on spermatid development, PNA immunofluorescence staining was performed to visualize acrosomal structures in mouse testes. Abundant PNA⁺ cells were observed in the seminiferous tubules of the control group, indicating normal spermatid formation. However, the number of PNA⁺ spermatids markedly decreased with increasing doses of cisplatin. In the 15 mg/kg group, partial loss of PNA⁺ signals was detected, whereas in the 20 mg/kg group, only a few scattered PNA⁺ cells remained. Nearly complete depletion of PNA⁺ spermatids was observed in the 40 mg/kg group, suggesting severe impairment of spermatid differentiation. Quantitative analysis confirmed a significant, dose-dependent reduction in the proportion of seminiferous tubules containing PNA⁺ cells (Supplementary Figure 1(A and B)). These results demonstrated that cisplatin induced spermatogenic dysfunction and developmental arrest in mouse testes.

Figure 1.

Spermatogenic failure in cisplatin-treated mice. (A) Schematic illustration of the workflow for mice following cisplatin or sterile normal saline (0.9% NaCl solution) induction. (B) Hematoxylin and eosin (H&E) staining of paraffin-embedded sections from the testis and epididymis of mice treated with different doses of cisplatin; * indicates degenerated tubules. Scale bar = 100 µm. (C) Average cross-sectional area of seminiferous tubules in mice treated with different doses of cisplatin (n > 50 tubules from each of three independent males). Student t-test; *P < 0.05, ***P < 0.001. (D) Assessment of testis to body weight ratio (mg/g) from mice treated with different doses of cisplatin; n = 3, Student t-test; *P < 0.05, **P < 0.005, ***P < 0.001. (E) Sperm count from cauda epididymides of mice treated with different doses of cisplatin; n = 3, Student t-test; ns.: no significance; **P < 0.005, ***P < 0.001.

Cisplatin exerted the greatest toxicity on spermatocytes in testes

To identify the specific germ cell types affected by cisplatin-induced spermatogenic disruption, we performed immunofluorescence staining at various germ cell stages. We used LIN28A to label undifferentiated spermatogonia, SYCP3 to label spermatocytes, and MVH to label germ cells. Compared with the control group, the number of MVH⁺ germ cells decreased progressively with cisplatin treatment, with the most significant loss observed at higher doses. Almost all germ cells were lost in the 40 mg/kg cisplatin group (Figure 2(A,B)). Further analysis showed that SYCP3⁺ spermatocyte loss was the most severe in the spermatogenic tubules. The number of SYCP3⁺ spermatocytes in the 15 mg/kg cisplatin group was 55% of that in the control group, and that in the 20 mg/kg cisplatin group was 20% of that in the control group. The SYCP3⁺ spermatocyte count was reduced by > 90% in the 40 mg/kg cisplatin group compared to that in the control group (Figure 2(C,D)). Following treatment with different doses of cisplatin, the total number of LIN28A⁺ undifferentiated spermatogonia was also reduced. Compared with the 15 mg/kg and 20 mg/kg groups, a further decrease in LIN28A⁺ cell number was observed in the high-dose (40 mg/kg) group (Figure 2(E,F)). Abundant ZBTB16⁺cells were detected along the basal membrane of seminiferous tubules in the control group, indicating normal maintenance of the spermatogonial stem cell pool. However, the number of ZBTB16⁺ cells progressively decreased with increasing cisplatin doses. In the 15 mg/kg group, the density of ZBTB16⁺ spermatogonia was moderately reduced, whereas in the 20 mg/kg and 40 mg/kg groups, ZBTB16⁺ cells were rarely observed within the seminiferous tubules. Quantitative analysis confirmed a significant, dose-dependent reduction in the number of ZBTB16⁺ cells per tubule (Supplementary Figure 2(A,B)). TUNEL staining was used to observe the number of testicular cell deaths after cisplatin treatment, and cisplatin was found to induce germ cell death in a dose-dependent manner (Figure 2(E,G)). These results indicated that germ cells in the seminiferous tubules were highly sensitive to cisplatin, with spermatocytes being the most vulnerable.

Figure 2.

Spermatocytes are the most sensitive to cisplatin toxicity. (A) Immunostaining of MVH (green) in mice treated with different cisplatin doses. Nuclei were stained with DAPI (blue). Scale bar = 50 µm. (B) Statistical results for the number of MVH⁺ cells per tubule in mice treated with different cisplatin doses (n > 50 tubules from each of three independent males); Student t-test; **P < 0.005, ***P < 0.001. (C) Immunostaining of SYCP3 (green) in mice treated with different cisplatin doses. Nuclei were stained with DAPI (blue). Scale bar = 50 µm. (D) Statistical results for the number of SYCP3⁺ cells per tubule in mice treated with different cisplatin doses (n > 50 tubules from each of three independent males); Student t-test; ***P < 0.001. (E) TUNEL (red) and LIN28A (yellow) staining to detect testicular cell apoptosis and undifferentiated spermatogonium in mice treated with different cisplatin doses. Nuclei were stained with DAPI (blue). Scale bar = 50 µm. (F) Statistical results for the number of LIN28A⁺ cells per tubule in mice treated with different cisplatin doses (n > 50 tubules from each of three independent males); Student t-test; ***P < 0.001. (G) TUNEL assay quantification in LIN28A⁺ cells, revealing a significant increase in apoptosis in the treated groups relative to the control. Data are mean values ± standard error of the mean. *P < 0.05, ***P < 0.001 indicates a statistically significant difference.

Proteomic analysis of cisplatin-induced spermatogenic damage in mouse testes

To identify the proteins associated with cisplatin-induced spermatogenic damage, we performed label-free quantitative proteomic analysis of testes from mice treated with 20 mg/kg cisplatin. Nuclear proteins constituted 49.43% of all the detected proteins (Figure 3(A)). The volcano plot and clustering heatmap show that, compared to the control group, 412 proteins were significantly downregulated in the cisplatin-treated group (fold change ≤ −1.2; p ≤ 0.05), while only 242 proteins were upregulated (fold change ≥ 1.2; p ≤ 0.05) (Supplementary Table 1). This suggests that cisplatin primarily reduces the cellular protein levels following testicular injury (Figure 3(B,C)).

Figure 3.

Proteomic analysis of cisplatin-treated mouse testes. (A) Distribution of proteins in label-free quantitative proteomic analysis. (B) Volcano plot displaying differentially expressed proteins between control group and cisplatin-treated mice. Fold change ≥1.2 or Fold change ≤ −1.2 and P value <0.05 were used as cut-offs. Red represents upregulated proteins, blue represents downregulated proteins, gray represents non-critical proteins. (C) Heatmap of protein expression comparing control (Con) and cisplatin-treated groups (Cis). Left panel displays all significant differential proteins levels between groups, with upregulated proteins shown in red and downregulated proteins in blue. Right panel displays the top 10 upregulated (red) and top 10 downregulated (blue) proteins from these differentially expressed proteins.

GO enrichment analysis of the downregulated proteins indicated significant enrichment in pathways related to meiosis, transcriptional regulation, and reproductive development, suggesting severe disruption of meiotic processes and spermatogenic arrest (Figure 4(A)). KEGG pathway analysis revealed the potential involvement of the meiosis, cell cycle, and P53 signaling pathways in cisplatin-induced testicular damage. Notably, the P53 signaling pathway, including the downregulated proteins CDK1 and GTSE1, was identified as a potential mechanism of action of cisplatin (Figure 4(B)). Gene set enrichment analysis further highlighted that pathways related to the cell cycle, reproductive development, and male spermatogenesis were affected by cisplatin treatment (Figure 4(C)). Using the STRING database, we predicted PPIs among the differentially expressed proteins and constructed a network map of cisplatin-induced testicular damage (Figure 5(A)). The Cytoscape HUBGENE function identified CDK1 as the highest-scoring core interaction protein, suggesting that it played a critical role in cisplatin-induced spermatogenic dysfunction (Figure 5(B)).

Figure 4.

Differentially expressed proteins enrichment analysis in cisplatin-treated mouse testes. (A) Gene ontology (GO) analysis results of down-regulated differentially expressed proteins; selected GO categories. (B) KEGG pathway analysis of down-regulated differentially expressed proteins. (C) GSEA plots show marked enrichment of the cell cycle, reproduction, and spermatogenesis pathway.

Figure 5.

Protein interactions in cisplatin-treated mouse testes. (A) Protein-Protein Interaction (PPI) network for down-regulated differentially expressed proteins, with nodes representing proteins and edges representing predicted interactions. (B) PPI network showing the interactions of differentially expressed proteins and hub proteins, indicating important roles in the network.

Cisplatin downregulated CDK1 protein levels in testes

To validate CDK1 expression in cisplatin-induced spermatogenic dysfunction, we performed immunofluorescent staining of testicular sections from mice treated with different cisplatin doses. CDK1 was primarily expressed in spermatogonia and spermatocytes, and the number of CDK1⁺ cells decreased as cisplatin dose increased (Figure 6(A,B)). Western blot analysis confirmed that cisplatin treatment significantly reduced CDK1 protein levels in mouse testes (Figure 6(C,D)). Our findings suggest that cisplatin-induced spermatogenic dysfunction may have been associated with the downregulation of CDK1 and related cell cycle regulators.

Figure 6.

CDK1 protein was significantly reduced in cisplatin-treated mouse testis. (A) Representative immunofluorescence images showing the co-localization of SYCP3 (green) and CDK1 (red) in testicular sections from control and cisplatin-treated mice. Nuclei were stained with DAPI (blue). Scale bars = 50 μm. B. Statistical results for the number of CDK1⁺ cells per tubule in mice treated with different doses of cisplatin, n > 50 tubules from each of three independent males; Student t-test; ***P < 0.001. (C) Representative Western blot images showing CDK1 (33 kDa) and TUBULIN (55 kDa, loading control) expression levels in testes from control and cisplatin-treated mice (15, 20, and 40 mg/kg). (D) Quantification of relative CDK1 protein levels normalized to TUBULIN. Data are presented as mean ± SEM from three independent biological replicates (n = 3). Statistical significance was determined using Student’s t-test; *P < 0.05, ***P < 0.001.

Discussion

Current research on testicular spermatogenic damage has primarily focused on busulfan-induced injury, identifying several key genes and intervention strategies [21–23]. However, the effect of cisplatin—a first-line chemotherapeutic agent used to treat multiple cancers—on testicular damage remains unclear. Previous studies have demonstrated that cisplatin-induced testicular toxicity can be exacerbated by co-treatment with other agents such as cetuximab, which intensifies the damage caused by cisplatin [24]. The present study further supports these findings, highlighting that cisplatin treatment at higher doses (15, 20, and 40 mg/kg) led to significant decreases in the sperm count and spermatocyte number in seminiferous tubules. This finding aligns with the known sensitivity of meiotically active cells to DNA-damaging agents, as spermatocytes undergo complex chromosomal dynamics during meiosis, rendering them particularly vulnerable to cisplatin-induced DNA crosslinks [25–27]. In contrast to studies focusing on the effects of busulfan, which primarily affects spermatogonia, our study highlights the pronounced spermatocyte sensitivity to cisplatin [21]. This observation is consistent with the work of Liu et al. who noted similar susceptibility in spermatocytes exposed to chemotherapy drugs, resulting in meiotic arrest and infertility [9]. Furthermore, the partial preservation of undifferentiated spermatogonia at lower cisplatin doses in our study suggested a degree of resistance to cisplatin-induced damage in SSCs [28]. Interestingly, although spermatocytes were the most severely affected, we observed a reduction in the number of undifferentiated spermatogonia at higher cisplatin doses. However, the significant reduction in LIN28A-positive cells at higher doses raises concerns regarding the potential for permanent infertility. This suggested that cisplatin may have a dual effect on spermatogenesis: acute toxicity to actively dividing spermatocytes, and a subtle long-term effect on the SSC pool. The latter is particularly concerning, as it can impair the regenerative capacity of the testis, leading to prolonged or even permanent infertility.

Furthermore, cisplatin disrupted spermatogenesis by downregulating CDK1 expression. CDK1 is the key spermatogenesis regulator in multiple species [16,17]. In the context of spermatogenesis, CDK1 activity is tightly regulated to ensure proper progression through meiosis, where errors can lead to chromosomal mis-segregation and subsequent germ cell apoptosis [18,19]. Previous studies have shown that disruption of CDK1 expression in mouse spermatocytes leads to meiotic arrest and infertility, which is consistent with our findings. In our study, cisplatin treatment caused spermatocyte death and reduced the CDK1 protein levels, indicating significant changes in CDK1 and related proteins during cisplatin-induced spermatogenic dysfunction. The observed reduction in CDK1 protein levels may explain the spermatocyte depletion observed in our study, as CDK1 is essential for the G2/M transition and proper progression through meiosis. CDK1 and related proteins may serve as biomarkers for cisplatin-induced spermatogenic damage. However, the molecular regulatory mechanisms and signaling pathways involving CDK1 remain to be elucidated and require further investigation. The enrichment of P53 signaling pathways in our proteomic analysis provides further insights into the action mechanisms of cisplatin. P53 activation in response to DNA damage can lead to cell cycle arrest or apoptosis, potentially explaining the observed germ cell loss [29]. The interplay between CDK1 downregulation and P53 activation warrants further investigation as it may represent a key regulatory node in cisplatin-induced spermatogenic damage.

This study had certain limitations. Proteomic analysis can only establish associations, but not causality, between cisplatin treatment and altered protein expression. Although CDK1 was identified as a central node in our analyses and its downregulation was confirmed experimentally, the functional validation of its role in spermatogenic disruption remains indirect. Further studies employing genetic or pharmacological approaches are required to establish the mechanistic CDK1 contribution in cisplatin-induced infertility. Moreover, this study primarily focused on the short-term effects of cisplatin on spermatogenesis and the molecular mechanisms underlying testicular injury, without a comprehensive evaluation of long-term male fertility following cisplatin treatment. Mating trials and hormone assays could provide a more thorough assessment of male reproductive capacity after chemotherapy, offering valuable insights into the potential for permanent infertility or recovery. These evaluations would provide essential information on the effectiveness of fertility preservation strategies for patients with cancer undergoing chemotherapy.

In conclusion, this study provides novel insights into the molecular mechanisms underlying cisplatin-induced spermatogenic damage. The identification of CDK1 as a key regulator of this process offers potential targets for fertility preservation strategies. Further research on the interplay between CDK1, P53, and other cell-cycle regulators in the context of cisplatin toxicity may lead to the development of protective interventions for cancer patients undergoing chemotherapy.

Funding Statement

This study was supported by grants from the National Natural Science Foundation of China [82260327], the Science and Technology Program of the Joint Fund of Scientific Research for the Public Hospitals of Inner Mongolia Academy of Medical Sciences [2024GLLH0297], and the Inner Mongolia Autonomous Region Science and Technology Plan-Innovation Platform Construction Program [2025KYPT0069]

Disclosure statement

No potential conflict of interest was reported by the author(s).

Data availability statement

The data supporting the findings of this study have been deposited in the ProteomeXchange Consortium via the PRIDE partner repository with the dataset identifier PXD061652 (http://www.proteomexchange.org). For any inquiries regarding the data, please contact Yingnan Zhang.