Abstract Background Combined cyclophosphamide (CTX) and busulfan (BUS) chemotherapy, while effective against malignancies, poses significant risks to male reproductive health. This study systematically investigates the proteomic basis underlying CTX + BUS-induced paternal reproductive toxicity and its impact on early embryonic development in mice. Methods We utilized a mouse model of acute CTX + BUS exposure. Systemic germ cell apoptosis, sperm parameters (concentration, motility, morphology), and fertilization rates were assessed. Testicular and sperm proteomic profiling was performed. Embryonic developmental outcomes were evaluated following fertilization with sperm from treated males. Results Acute CTX + BUS exposure induced systemic germ cell apoptosis and profound sperm head teratozoospermia, despite preserved sperm concentration and motility. Proteomic profiling revealed testicular dysregulation of ribosome biogenesis, DNA replication, and cell cycle control, alongside sperm-specific depletion of ribosomal proteins. Sperm from CTX + BUS-treated mice exhibited a reduced fertilization rate and induced severe embryonic developmental defects. Molecularly, ribosomal insufficiency in sperm is linked to lower ribosomal protein levels in early embryos, potentially compromising the paternal role in translational activation. Conclusions Our findings establish ribosome biogenesis defects as a novel pathway mediating paternal-derived embryotoxicity following CTX + BUS chemotherapy. They challenge the reliance on conventional semen parameters (motility, count) for fertility assessment, highlighting that significant molecular lesions in sperm can persist despite normal traditional metrics. These results advocate for incorporating protein biomarker analysis into clinical fertility risk stratification for cancer survivors. This work provides critical insights into the early embryonic consequences of chemotherapy and underscores the imperative to safeguard paternal genomic and epigenetic integrity in oncological care. Supplementary Information The online version contains supplementary material available at 10.1186/s12967-025-07215-6. Keywords: Chemotherapy, Ribosome, Perm proteomics, Teratozoospermia, Reproductive toxicity Introduction For decades, spermatozoa have been primarily regarded as passive vectors delivering paternal DNA to the oocyte during fertilization. However, accumulating evidence has revolutionized this paradigm, revealing that sperm contribute far more than genetic material to early embryogenesis. Beyond the haploid genome, sperm deliver a complex repertoire of components, including fertilization-activating factors, centrosomes, messenger RNAs (mRNAs), microRNAs (miRNAs), and epigenetic regulators, which collectively orchestrate critical post-fertilization events [[38]1–[39]5]. Notably, sperm-derived centrosomes govern mitotic spindle formation, while paternal RNAs and proteins are implicated in regulating chromatin remodeling, metabolic reprogramming, and the maternal-to-zygotic transition [[40]1–[41]3, [42]5]. Emerging studies further highlight that sperm-derived small non-coding RNAs can modulate embryonic gene expression and early embryo development [[43]2–[44]4]. Importantly, both genetic lesions and epigenetic aberrations in sperm have been linked to fertilization failure, embryonic developmental arrest, and transgenerational health risks [[45]3, [46]6, [47]7]. Despite these advances, the potential molecular alterations by which paternal non-genetic components coordinate embryogenesis, particularly under conditions of environmental or pharmacological stress remain poorly defined. Chemotherapy remains a cornerstone of cancer treatment, and with improving oncological outcomes, increasing attention has been directed toward its off-target toxicities, particularly reproductive impairment [[48]8, [49]9]. Alkylating agents such as cyclophosphamide (CTX) and busulfan (BUS) are widely used in conditioning regimens for hematopoietic stem cell transplantation and autoimmune diseases, exerting their antitumor effects through DNA crosslinking and oxidative damage to rapidly dividing cells [[50]10–[51]12]. While their therapeutic efficacy is well-established, both agents are highly gonadotoxic, inducing apoptosis in spermatogonia and spermatocytes, thereby disrupting spermatogenesis and leading to transient or permanent infertility [[52]11–[53]18]. Previous studies have characterized the individual reproductive toxicity profiles of CTX and BUS, However, in clinical practice, these agents are frequently administered in combination to maximize therapeutic synergism, yet the compounded impact of CTX + BUS co-treatment on spermatogenesis, sperm functional competence, and paternal contributions to embryonic development remains poorly understood. Chemotherapy exposure is well-documented to induce sperm DNA damage and epigenetic instability [[54]12, [55]19–[56]21]. However, its impacts on the functional sperm proteome and protein assembly machinery remain underexplored. While conventional biomarkers such as sperm concentration and motility are widely used to assess male fertility, emerging evidence highlights subtle molecular alterations in sperm non-genetic components [[57]19, [58]22–[59]25], including RNAs, chromatin architecture, and epigenetic modifiers, may impair embryonic development, even in the absence of overt semen parameter abnormalities. Survivors exposed to alkylating agents like CTX and BUS often retain residual sperm production; however, the functional integrity of these gametes and their capacity to support healthy offspring remain uncertain. Specifically, whether chemotherapy-induced dysregulation of sperm protein expression, particularly in processes such as ribosome biogenesis, chromatin remodeling, or centrosomal function, compromises paternal contributions to fertilization and early embryogenesis remains poorly characterized. In this study, we employed a multidimensional approach to investigate the impact of acute CTX + BUS exposure on testicular function, sperm integrity, and embryonic developmental competence in mice. Integrating histopathology, proteomic profiling, ultrastructural analysis, and functional fertilization assays, we demonstrate that acute CTX + BUS treatment induces germ cell apoptosis, and profound ribosomal dysfunction in spermatozoa. Strikingly, while sperm concentration and motility remain largely unaffected, chemotherapy triggers severe sperm head teratozoospermia and ribosomal protein depletion, leading to decreased fertilization capacity and abnormal early embryonic development. Our findings identify ribosome biogenesis defects as novel molecular alterations underlying paternal-mediated embryotoxicity, highlighting the imperative to evaluate non-genetic sperm quality parameters in cancer survivors. This work advances our understanding of chemotherapy-induced reproductive risks and provides a foundation for developing strategies to mitigate paternal contributions to adverse developmental outcomes. Materials and methods Animals, treatments and samples collection Six-week-old male ICR mice were purchased from Vital River Laboratory Animal Technology Co., Ltd. (Vital River, China) and acclimatized for 7 days under standard housing conditions (temperature: 23 ± 2 °C, humidity: 55 ± 5%, 12-hour light/dark cycle) prior to experimental procedures. ICR male mice were randomly divided into two groups: the chemotherapy group receiving intraperitoneal injections of cyclophosphamide (120 mg/kg) and busulfan (30 mg/kg) dissolved in dimethylsulfoxide (DMSO) for 7 consecutive days, and a control group administered an equivalent volume of DMSO vehicle alone. All the animal procedures were approved by the Animal Care and Use Committee (ACUC) of Jiangxi Maternal and Child Health Hospital. Sperm motility analysis After 7 consecutive days of treatment, mice in both the control and chemotherapy groups were euthanized by cervical dislocation. Epididymides were then dissected, isolated, and subjected to spermatozoa collection via the swim-out method. This involved incubating epididymal tissues in pre-warmed IVF medium (Vitrolife, Sweden) for 10 min. Following tissue removal, sperm suspensions were centrifuged twice (300 × g, 5 min) with fresh IVF medium and resuspended. Sperm motility parameters were assessed using a computer-assisted sperm analysis (CASA) system (Hamilton-Thorne Biosciences, USA) as previously described [[60]26]. For each analysis, 5 µL of sperm suspension was loaded into a pre-warmed 20-µm depth GoldCyto counting chamber (Hamilton-Thorne, USA). A minimum of 200 spermatozoa per sample were analyzed to determine percentages of total motile and progressively motile spermatozoa. All experiments were performed in triplicate, with pooled data used for subsequent statistical analysis. Sperm collection After euthanasia by cervical dislocation, the epididymides were dissected from both the control and chemotherapy-treated mice. Spermatozoa were collected using the swim-out method by incubating the epididymal tissues in pre-warmed IVF medium (Vitrolife, Sweden) for 30 min at 37 °C under 5% CO₂. Following swim-up, the epididymal tissues were removed, and the sperm suspension was filtered through a 100-µm cell strainer to remove debris. The filtrate was then centrifuged at 10,000 × g for 15 min at 4 °C, and the supernatant was discarded. The sperm pellet was resuspended in 1x PBS (Gibco, USA) and centrifuged again under the same conditions. After the final centrifugation, the supernatant was removed, and the pellet was flash-frozen in liquid nitrogen and stored at − 80 °C for subsequent analysis. In vitro fertilization (IVF) and embryo culture Following cervical dislocation euthanasia, bilateral cauda epididymides were harvested from both control and chemotherapy-treated mice. Tissues were immediately immersed in pre-warmed IVF medium (Vitrolife, Sweden) and microscopically incised to facilitate sperm release. After a 30-minute incubation in a Cook mini-incubator (Cook Medical, USA) to promote sperm swim-out, sperm concentration was quantified using a Makler chamber. For IVF, spermatozoa were diluted to 1–2 × 10⁶ cells/mL in fertilization droplets and capacitated for 40 min. Concurrently, 8-week-old ICR females were euthanized 14–16 h post-hCG administration. Oviducts were rapidly excised and transferred to culture dishes, where cumulus-oocyte complexes (COCs) were isolated from ampullae under stereomicroscopic visualization using fine forceps. Matured COCs were co-incubated with capacitated sperm for 6 h, followed by sequential washing steps using a mouth pipette to eliminate excess sperm and cumulus cells. Total oocytes retrieved were categorized as immature (germinal vesicle-intact) or mature (extruded first polar body) based on morphological assessment. Embryonic development was monitored daily using an inverted microscope (Nikon, Japan) over 4 days of culture. Developmental milestones, including 4–8 cell embryo formation and blastocyst yield per group, were systematically recorded. Histology and transmission electron microscopy For paraffin embedding, testes and epididymides were fixed in Hartman’s fixative (Sigma, H0290) for 48 h. The tissues were then dehydrated through a gradient ethanol series (70% for 30 min, 80% for 30 min, 90% for 30 min, 95% for 30 min, and 100% for 30 min), cleared in xylene for 40 min, and infiltrated with liquid paraffin at 65 °C for 3 h. Subsequently, the tissues were embedded in paraffin within plastic molds. Sections of 5 μm thickness were cut, dewaxed at 65 °C, and dried for 4 h before storage at room temperature for further use. For histological analysis, the sections were rehydrated through a descending alcohol series, stained with hematoxylin and eosin (H&E), dehydrated in a gradient ethanol series, cleared in xylene, and finally mounted with resinous medium. For transmission electron microscopy analysis, tissues were fixed overnight at 4 °C in 5% glutaraldehyde buffered with 0.2 M sodium cacodylate. The samples were then sent to Wuhan Servicebio Technology Co., Ltd. for embedding and ultrathin sectioning. The sections were stained with uranyl acetate and lead citrate and examined under a transmission electron microscope (HITACHI, HT7800). TUNEL assay Paraffin-embedded tissue sections were dewaxed in xylene and rehydrated through a gradient ethanol series, followed by a 5-min rinse in phosphate-buffered saline (PBS; Gibco, 10010023). Next, sections were treated with 20 µg/mL Proteinase K (Sigma-Aldrich, P2308) in PBS at 37 °C for 20 min and washed twice with PBS. Endogenous peroxidase activity was quenched by incubating tissues in 3% H₂O₂ at room temperature for 20 min, followed by three PBS washes. The TUNEL assay was performed using the In Situ Cell Death Detection Kit (Roche, 11684817910) according to the manufacturer’s protocol. Briefly, 50 µL of Equilibration Buffer was applied to each section and incubated at room temperature for 10 min. Subsequently, 57 µL of TdT Reaction Mix was added to the tissues, and slides were incubated in a humidified chamber at 37 °C for 1 h. After three PBS washes, sections were treated with 100 µL of Streptavidin-HRP Conjugate (diluted 1:200 in PBS) at 37 °C for 30 min. Brown chromogenic signals were observed using the DAB Substrate Kit (Beyotime Biotechnology, P0203), and the reaction was immediately terminated by immersing the tissue slides in distilled water. Nuclei were counterstained with hematoxylin, and then the sections were dehydrated through an ascending ethanol series, cleared in xylene, and mounted with neutral balsam. Stained sections were examined under a light microscope (Nikon, Eclipse E100) equipped with a digital camera (Nikon, DS-Fi3). Apoptotic cells were identified by distinct brown-yellow nuclear staining, while viable cells exhibited blue hematoxylin-counterstained nuclei. Proteomic analysis Testicular tissues and sperm samples from both chemotherapy-treated and control group mice were subjected to quantitative proteomic analysis at Westlake Omics (Hangzhou, China) Inc. Specifically, after 7 consecutive days of treatment, testicular tissues were harvested from mice in both groups, and spermatozoa were collected from the epididymides of these mice for proteomic analysis. Briefly, proteins were extracted from the samples and enzymatically digested into peptides using trypsin. The resulting peptides were desalted, vacuum-dried, and quantified via A280 absorbance measurement. For spectral library construction, a Data-Dependent Acquisition (DDA) approach was performed. The peptides were fractionated by liquid chromatography (LC) and subsequently vacuum-dried. The prepared samples were then analyzed by liquid chromatography-tandem mass spectrometry (LC-MS/MS) on a high-resolution instrument. Proteomic data processing employed database searches against reference protein sequences for identification and quantification. The proteomics results for sperm and testicular tissue can be found in Supplementary material [61]S1 and [62]S2. Gene set enrichment analysis (GSEA) GSEA was performed on sperm proteomics data derived from chemotherapy-treated and control mice. The fold-change values of each protein between the two groups were used as input for GSEA. Pathways were ranked based on their normalized enrichment score (NES), and statistical significance was assessed through permutation testing. Only pathways meeting the threshold of P < 0.05 and an absolute NES >1 were considered significantly enriched. The analysis was conducted using the GSEA software (3.0.0) with the Molecular Signatures Database (MSigDB) as the reference gene set collection [[63]27, [64]28]. Protein-protein interaction (PPI) network analysis The significantly altered proteins identified in the testis or sperm proteomics data (chemotherapy-treated vs. control groups) were submitted to the STRING database (version 11.5; [65]https://string-db.org/) with Mus musculus as the reference species. A confidence score cutoff of ≥ 0.9 (high confidence) was applied to minimize false-positive interactions. The resulting interaction data, including protein nodes and edges, were exported as a TSV file and imported into Cytoscape (version 3.9.1) for further network visualization and analysis. Key topological parameters, such as degree centrality and betweenness, were calculated to identify hub proteins within the network. Real-time quantitative polymerase chain reaction (RT-qPCR) analysis Testicular tissues and sperm samples were collected from male mice assigned to two experimental groups: the control group and the CTX + BUS group, after 7 days of treatment. Total RNA was separately extracted from testis tissues and sperm using TRIzol Reagent (Invitrogen, USA) according to the manufacturer’s instructions. RNA concentration and purity were determined using a NanoDrop 2000 spectrophotometer (Thermo Fisher Scientific, USA). Complementary DNA (cDNA) was synthesized from 500 ng of total RNA per sample using a reverse transcription kit (R233-01, Vazyme, China). Quantitative real-time PCR was performed on a StepOnePlus Real-Time PCR System (Thermo Fisher Scientific, USA) with SYBR Green Kit (Q226-01, Vazyme, China). Primer sequences for apoptosis-related genes are listed in Supplemental Table [66]I. All primers were obtained from the PrimerBank database ([67]https://pga.mgh.harvard.edu/primerbank/) and had been previously validated for specificity and amplification efficiency. The relative mRNA expression levels of target genes were calculated using the 2^(–ΔΔCt) method, with alpha-tubulin (Tuba1a) serving as the internal reference gene for normalization. Western blot Western blot analysis was performed as previously described [[68]29]. Briefly, proteins were separated by SDS-polyacrylamide gel electrophoresis (SDS-PAGE) and transferred onto PVDF membranes (Millipore, USA). After blocking with 5% non-fat milk in PBS containing 0.1% Tween-20 (PBST) for 1 h at room temperature, membranes were incubated overnight at 4 °C with primary antibodies diluted in blocking buffer. The following primary antibodies were used: mouse monoclonal anti-α-TUBULIN (1:5000, Clone 5-B-1-2, Proteintech); Rabbit monoclonal anti-GAPDH (1:5000, 2118T, CST); Rabbit polyclonal anti-RPS9 (1:1000, 18215-1-AP, Proteintech); Rabbit monoclonal anti-RPS6 (1:1000, TN25622S, Abmart); Rabbit polyclonal anti-RPS10 (1:1000, 14894-1-AP, Proteintech); Rabbit polyclonal anti-RPS23 (1:1000, 29834-1-AP, Proteintech); Rabbit polyclonal anti-RPS25 (1:1000, 23599-1-AP, Proteintech); Rabbit polyclonal anti-RPS20 (1:1000, 15692-1-AP, Proteintech); Rabbit polyclonal anti-RPS19 (1:1000, 15085-1-AP, Proteintech); Rabbit polyclonal anti-RPS15 (1:1000, 14957-1-AP, Proteintech); Rabbit monoclonal anti-Cleaved PARP (1:1000, 9541T, CST); Rabbit monoclonal anti-Cleaved Caspase-3 (1:1000, 9664T, CST). Membranes were subsequently probed with horseradish peroxidase (HRP)-conjugated secondary antibodies for 1 h at room temperature. Protein signals were visualized using an enhanced chemiluminescence substrate (Vazyme, China) and captured with an automated chemiluminescence imaging system (Tanon 5200, China). Statistical analysis Data are presented as mean ± standard deviation (SD). To assess differences between two independent experimental groups, a two-tailed unpaired Student’s t-test was employed. Statistical significance was defined as P < 0.05 for all analyses. All statistical calculations and graphical representations were performed using GraphPad Prism 9 software (Version 9.5.1). Results Combined cyclophosphamide and busulfan treatment induces germ cell apoptosis To investigate the effects of cyclophosphamide (CTX) combined with busulfan (BUS) on testicular spermatogenesis, mice were intraperitoneally administered either saline (Control) or CTX + BUS for seven consecutive days, followed by analyses of testes and sperm parameters. Body weight was monitored daily throughout the treatment period. Starting from day 2, the CTX + BUS group exhibited a significant reduction in body weight compared to the control group (Fig. [69]1A). By day 7, the body weight of CTX + BUS-treated mice decreased by over 20% relative to their initial weight (Fig. [70]1A), indicating systemic toxicity induced by the drug treatment. Furthermore, our results revealed that both testicular size and weight in the CTX + BUS group were significantly reduced compared to control group (Fig. [71]1B and C). Interestingly, the testis-to-body weight ratio in the CTX + BUS group exhibited a modest but statistically significant increase (Fig. [72]1D). The asynchronous changes in testicular weight and body weight dynamics further underscore the necessity of integrating multidimensional evidence, including testicular weight, histopathological examination, and functional assays, for a comprehensive assessment of reproductive system injury. Histological examination of seminiferous tubules showed numerous deeply stained and highly condensed nuclei were observed in the CTX + BUS testes (Fig. [73]1E), indicative of impaired spermatogenesis. Similarly, the epididymal cauda in CTX + BUS-treated mice exhibited lots of densely stained nuclear debris (Fig. [74]1E), implying potential apoptotic events. To confirm apoptosis, TUNEL assays were performed on testicular and epididymal tissues (Fig. [75]1F). Quantitative analysis revealed a marked increase in TUNEL-positive germ cells within seminiferous tubules of the CTX + BUS group compared to controls (Fig. [76]1G and H). Furthermore, the epididymal cauda in CTX + BUS-treated mice displayed extensive apoptotic germ cell residues, consistent with the histological observations (Fig. [77]1I). To further explore the molecular mechanisms underlying CTX + BUS-induced apoptosis, we assessed the expression of apoptosis-related genes. RT-qPCR analysis of testicular tissues demonstrated that the mRNA levels of Bax, Parp, Caspase 3, Caspase 6, and Caspase 9 were significantly upregulated in the CTX + BUS group compared to the control (Fig. [78]1J). Similarly, elevated expression of these apoptosis-related genes was also observed in sperm samples from CTX + BUS-treated mice (Fig. [79]1K). Consistent with these findings, Western blot analysis revealed a significant increase in the protein levels of cleaved caspase-3 and cleaved PARP in both testicular (Fig. [80]1L) and sperm (Fig. [81]1M) samples from the CTX + BUS group, further confirming the activation of apoptotic pathways. These findings collectively demonstrate that CTX + BUS treatment induces systemic toxicity, testicular atrophy, and extensive germ cell apoptosis, severely compromising spermatogenesis. Fig. 1. [82]Fig. 1 [83]Open in a new tab Combined cyclophosphamide and busulfan treatment reduces testis size and enhances germ cell apoptosis. (A) Mice were intraperitoneally injected with saline (control) or a combination of cyclophosphamide (CTX) and busulfan (BUS) for 7 consecutive days, and body weight was recorded daily. n = 15 for control group, n = 20 for CTX + BUS group. For each time point, data are represented as mean ± SD. ^**P < 0.01, ^***P < 0.001. (B) Representative images of testes and epididymides from control and CTX + BUS-treated mice. (C) Quantitative analysis of testis weight in control and CTX + BUS groups. Error bars indicate SD, ^***P < 0.001. (D) Quantitative analysis of testis-to-body weight ratio. Data are represented as mean ± SD. ^*P < 0.05. (E) Hematoxylin and eosin (H&E) staining of testis and epididymis sections from control and CTX + BUS-treated mice. Condensed apoptotic nuclei were indicated by red arrows. The right panels showed magnified views of the areas marked by red dashed boxes. Scale bar: 50 μm. (F) Representative images of TUNEL staining in sections of testes and epididymides. Apoptotic cells are marked by black arrows. Scale bar: 50 μm. (G) Percentage of TUNEL-positive seminiferous tubules in testis from control and CTX + BUS-treated mice. n = 4 for each group. Data are represented as mean ± SD. ^**P < 0.01. (H) Quantification of TUNEL-positive cells per tubule in testis. Data are represented as mean ± SD. ^***P < 0.001. (I) Quantification of TUNEL-positive cells per tubule in the epididymis. Data are represented as mean ± SD. ^***P < 0.001. Analysis of apoptosis-related gene expression by RT-qPCR in the testes (J) and sperm (K) of mice following 7-day combined treatment with CTX and BUS. Data are represented as mean ± SD. ^*P < 0.05, ^**P < 0.01. Analysis of apoptosis-related protein expression by Western blot in the testes (L) and sperm (M) of mice subjected to 7 consecutive days of combined CTX and BUS treatment Testicular proteomic profiling identifies CTX + BUS-induced dysregulation of DNA replication, ribosomal subunit assembly, and cell cycle control To investigate the impact of CTX + BUS treatment on testicular proteomic profiles, data-independent acquisition (DIA)-based proteomic analysis was performed with three biological replicates per group. Principal component analysis (PCA) demonstrated distinct clustering between the CTX + BUS-treated and control groups, indicating robust inter-group separation (Fig. [84]2A). A total of 10,043 proteins were identified by mass spectrometry. Differentially expressed proteins (DEPs) were screened using thresholds of |log2(foldchange)| >1 and p < 0.05, yielding 257 upregulated and 77 downregulated proteins (Fig. [85]2B). Heatmap revealed marked divergence in expression patterns between groups (Fig. [86]2C). Functional enrichment analysis was conducted to elucidate biological implications. Gene Ontology (GO) analysis revealed that downregulated proteins in the CTX + BUS group were predominantly associated with ribonucleoprotein complex biogenesis, ribosome biogenesis, and DNA replication (Fig. [87]2D). Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway analysis further corroborated these findings, identifying significant enrichment in DNA replication, ribosome biogenesis in eukaryotes, and cell cycle (Fig. [88]2E). Network-based pathway mapping using KEGG definitions confirmed primary involvement of three functional modules: DNA replication machinery, ribosomal function components, and cell cycle progression regulators (Fig. [89]2F). To investigate protein interaction dynamics, a protein-protein interaction (PPI) network was constructed using the STRING database and subsequently analyzed using the MCODE algorithm. The resulting network revealed functional modules centered on key hub proteins associated with ribosomal small subunit biogenesis (Fig. [90]3G). Collectively, these proteomic alterations suggest that CTX + BUS treatment perturbs testicular functions related to genomic stability maintenance, protein synthesis machinery, and cell cycle control. Fig. 2. [91]Fig. 2 [92]Open in a new tab Proteomic analysis of testes from mice treated with saline or CTX + BUS. (A) Principal component analysis (PCA) of protein expression patterns across treatment groups. CB: CTX + BUS. (B) Volcano plot displaying differentially expressed proteins (DEPs) between control and CTX + BUS groups, identified by Astral DIA. Proteins with a fold change > 2 and P < 0.05 were considered statistically significant. The top 20 most significantly altered DEPs are labeled. (C) Heatmap illustrating log2-transformed expression levels of DEPs in all samples from control versus CTX + BUS groups. (D) Bubble plot showing Gene Ontology (GO) enrichment analysis of downregulated proteins in the CTX + BUS group. (E) Bubble plot of KEGG pathway enrichment analysis for downregulated proteins in the CTX + BUS group. (F) Visualization of network-based pathway enrichment, where node color indicates statistical significance and node size corresponds to the gene count within each pathway. (G) Protein-protein interaction (PPI) network constructed via the STRING database and visualized using Cytoscape software, with key DEPs highlighted Fig. 3. [93]Fig. 3 [94]Open in a new tab CTX + BUS treatment induces sperm head abnormalities. (A) Sperm count from the cauda epididymis of control and CTX + BUS-treated mice. n = 6 per group. Data are presented as mean ± SD; ns, not significant. (B) Sperm parameters analyzed using a computer-assisted sperm analysis (CASA) system (n = 4 per group). Data are presented as mean ± SD; ns, not significant; *P < 0.05 (two-tailed unpaired t-test). (C) Transmission electron microscopy (TEM) showing the typical “9 + 2” microtubule structure in sperm flagella from control and CB groups. Scale bar: 1 μm. (D) TEM images of sperm head morphology in control and CTX + BUS (CB) groups. Scale bar: 1 μm. (E) Quantification of sperm with head abnormalities. Data are presented as mean ± SD; **P < 0.01 Acute CTX + BUS chemotherapy induces sperm head abnormalities with minimal impact on motility and count To evaluate the impact of CTX + BUS combinatorial chemotherapy on sperm quantity and functional integrity, we first assessed spermatozoa concentration in the cauda epididymis using a hemocytometer. No significant difference was observed in sperm density between the control group (27.77 ± 2.9 × 10⁶/mL) and the CTX + BUS group (23.94 ± 3.5 × 10⁶/mL; P > 0.05) (Fig. [95]3A), suggesting unaltered spermatogenic output after 7-day treatment. Sperm motility parameters were subsequently analyzed via computer-assisted sperm analysis (CASA). The results further demonstrated comparable total motility between groups (Control: 60.13% ± 4.9% vs. CTX + BUS: 53.78% ± 3.2%, P > 0.05) (Fig. [96]3B). Subpopulation analysis revealed a modest but statistically significant reduction in rapid progressive motility in CTX + BUS-treated males (Fig. [97]3B), while slow progressive and static sperm fractions remained unchanged (Fig. [98]3B). Ultrastructural examination via transmission electron microscopy (TEM) confirmed preserved integrity of the axonemal “9 + 2” microtubule architecture in CTX + BUS group sperm flagella (Fig. [99]3C), aligning with the observed minimal motility alterations. Strikingly, TEM revealed significant morphological abnormalities in sperm heads of the CTX + BUS group (Fig. [100]3D), with a 5.4-fold increase in malformations observed in this cohort compared to controls (Fig. [101]3E). Collectively, our findings demonstrate that acute CTX + BUS exposure preferentially induces sperm head teratozoospermia rather than overt motility deficits, highlighting a potential risk for fertilization failure or developmental anomalies despite preserved sperm quantity and gross motility metrics. Chemotherapy-induced sperm damage disrupts fertilization and early embryonic development To evaluate the impact of sperm head damage induced by CTX + BUS treatment on fertilization and early embryogenesis, we conducted in vitro fertilization (IVF) experiments using mature MII oocytes collected from superovulated 6-week-old ICR female mice. Compared to the control group, spermatozoa from CTX + BUS-treated mice exhibited a significant 30% reduction in fertilization rate (Fig. [102]4A). Subsequent 4-day in vitro culture of zygotes revealed dramatic developmental impairments in the CTX + BUS group. The blastocyst formation rate showed a marked decline (Fig. [103]4B), while cleavage-stage embryos demonstrated substantially higher fragmentation rates (>50% fragmented blastomeres) compared to controls (Fig. [104]4C and E), indicating severely compromised embryo quality. Notably, approximately 23% of embryos in the CTX + BUS group remained arrested at the 2-cell stage following 4 days of culture, a phenomenon observed in fewer than 10% of control embryos (Fig. [105]4D and E), suggesting potential defects in zygotic genome activation (ZGA) triggered by chemotherapy exposure. To further investigate the underlying mechanisms of ZGA impairment, we collected 2-cell embryos derived from IVF using sperm from control and CTX + BUS-treated mice and analyzed the expression of key ZGA-related factors by RT-qPCR. The results showed significant downregulation of several critical ZGA regulators [[106]30, [107]31], including Btg4, Obox3, and Obox4, in the CTX + BUS group (Fig. [108]4F). In contrast, the expression levels of Obox1 and Obox5 remained comparable between the two groups (Fig. [109]4F). These findings further support the notion that CTX + BUS combination chemotherapy interferes with the initiation of ZGA, likely through the dysregulation of specific paternal factors essential for early embryonic transcription. Collectively, these findings demonstrate that CTX + BUS-induced sperm head abnormalities not only impair fertilizing capacity but also critically disrupt early embryonic development, likely through both physical damage to sperm components and dysregulation of paternal molecules that are essential for zygotic genome activation. Fig. 4. [110]Fig. 4 [111]Open in a new tab CTX + BUS treatment significantly reduces fertilization and blastocyst formation rates. (A) Fertilization rates were quantified using sperm from control and cyclophosphamide (CTX) + busulfan (BUS)-treated mice for in vitro fertilization (IVF) experiments with oocytes harvested from 6-week-old female ICR mice. Fertilization outcomes were assessed by microscopic evaluation of polar body extrusion and pronuclear formation (control: n = 8 mice; CTX + BUS: n = 7 mice). Data are presented as mean ± SD; ***P < 0.001. (B) Embryos were cultured for 4 days after IVF and blastocyst formation rates were calculated. Data are presented as mean ± SD; ***P < 0.001. (C) Comparison of embryos with > 50% fragmentation at cleavage stage between control and CTX + BUS groups. Data are presented as mean ± SD; ***P < 0.001. (D) Proportion of 2-cell embryos on day 4 post-IVF in control versus CTX + BUS groups. Data are presented as mean ± SD; ***P < 0.001. (E) Representative images of embryos from control and CTX + BUS groups after 4 days of in vitro culture following IVF. (F) RT-qPCR analysis of the expression of key regulatory factors for zygotic genome activation (ZGA) in two-cell embryos derived from fertilization of sperm from the control group and the CTX + BUS group. Data are presented as mean ± SD; *P < 0.05, **P < 0.01, ns: not significant Proteomic profiling reveals ribosome biogenesis defects as key drivers of CTX + BUS-induced reproductive toxicity To uncover the molecular alterations underlying CTX + BUS-induced sperm defects in fertilization and early embryonic development, we performed DIA-based proteomic profiling of spermatozoa from control and CTX + BUS-treated mice. Principal component analysis revealed distinct separation between the CTX + BUS and control groups, indicating substantial alterations in the sperm proteome following chemotherapy exposure (Fig. [112]5A). The DIA-based proteomics identified 8860 proteins, with 544 significantly downregulated and 29 upregulated in the CTX + BUS group (fold change > 2, P < 0.05). Volcano plot analysis highlighted the top 20 differentially, most of which were ribosomal proteins (Fig. [113]5B). A heatmap further confirmed the consistent downregulation pattern of these DEPs across control and CTX + BUS-treated samples (Fig. [114]5C). GO enrichment analysis of downregulated proteins demonstrated significant enrichment in biological processes related to cytoplasmic translation, ribosomal small subunit biogenesis, and ribosome assembly (Fig. [115]5D). Cellular component analysis revealed that these proteins were predominantly localized to the cytosolic ribosome, cytosolic ribosomal subunit, and ribosomal subunit. Molecular function annotation further demonstrated their involvement in Structural Constituent of Ribosome, rRNA binding, and translational regulator activity. KEGG pathway analysis corroborated these findings, identifying ribosome as the most significantly enriched pathway (Fig. [116]5G and H). Visualization of KEGG maps revealed that the majority of downregulated proteins clustered within both the large and small ribosomal subunits (Fig. [117]5I). Subcellular localization analysis further indicated that cytosolic proteins constituted the largest proportion of DEPs (Fig. [118]5J), aligning with their roles in cytoplasmic translation and ribosome biogenesis. Fig. 5. [119]Fig. 5 [120]Open in a new tab Proteomic analysis reveals impaired ribosomal function in sperm from CTX + BUS-treated mice. (A) PCA of protein expression profiles. Colored dots represent biological replicates from different treatment groups. (B) Volcano plot showing differentially expressed proteins between control and CTX + BUS groups. Significantly upregulated (orange), downregulated (blue), and unchanged (gray) proteins are indicated. The top 20 most significantly altered proteins are labeled. (C) Heatmap displaying expression patterns of DEPs across samples. (D-F) Gene Ontology enrichment analysis of DEPs: (D) biological processes, (E) cellular components, and (F) molecular functions. Top 10 enriched pathways are shown as bubble plots ranked by P-value. (G) Bar plot of KEGG pathway enrichment analysis for DEPs. (H) Network visualization of enriched KEGG pathways. (I) KEGG Pathview mapping showing downregulated ribosomal proteins (blue) in the CTX + BUS group. (J) Subcellular localization analysis of DEPs Collectively, these results demonstrate that CTX + BUS treatment induces a pronounced decline in sperm ribosomal function, characterized by systemic downregulation of ribosome-associated proteins. This ribosomal insufficiency likely disrupts critical paternal contributions to zygotic translation and early embryogenesis, providing a mechanistic explanation for the observed fertilization defects and embryonic developmental arrest. GSEA unveils systemic multidimensional functional disruption in chemotherapy-exposed sperm To complement the differential expression analysis and gain a systems-level perspective on proteomic alterations, we performed Gene Set Enrichment Analysis (GSEA) on the entire sperm proteome dataset. The most significantly enriched gene sets were associated with Ribosome, and complement and coagulation cascades (Fig. [121]6A), consistent with the DEP-focused GO/KEGG findings. Notably, GSEA uncovered additional impaired pathways related to chromatin remodeling, transcription regulator complex, spliceosomal complex, spermatogenesis, Ras signaling pathway, and chemokine signaling pathway (Fig. [122]6A), suggesting comprehensive functional disruption in chemotherapy-exposed sperm. Fig. 6. [123]Fig. 6 [124]Open in a new tab Functional analysis of differentially expressed proteins in CTX + BUS-treated sperm. (A) Gene set enrichment analysis (GSEA) comparing control and CTX + BUS treatment groups, revealing significantly altered biological pathways. (B) Protein-protein interaction (PPI) network constructed using STRING database and visualized with Cytoscape. Node size and color intensity represent the degree of protein connectivity (number of interacting partners). (C) Heatmap showing expression patterns of key ribosomal proteins identified through PPI network analysis. Color gradient indicates relative protein expression levels across samples. (D) Western blot analysis of the protein expression levels of the screened hub proteins in sperm from both the control group and the chemotherapy-treated group. (E) Western blot analysis of the expression levels of the screened hub proteins in two-cell zygotes derived from fertilization of normal mature oocytes with sperm from either the control or chemotherapy-treated group PPI networks uncover chemotherapy-induced ribosomal dysfunction in sperm and its paternal contribution to early embryonic developmental defects To explore the functional and physical interactions among DEPs following chemotherapy exposure, we constructed a protein-protein interaction (PPI) network using the STRING database and visualized it via Cytoscape. The hub protein screening demonstrated that ribosomal proteins formed the most highly connected nodes within the PPI network (Fig. [125]6B), suggesting they may represent critical functional hubs following chemotherapy exposure. This observation aligned with the GO/KEGG enrichment findings, further supporting that CTX + BUS-induced spermatogenic toxicity primarily perturbs ribosomal and translational machinery. A heatmap visualization of protein expression within this PPI network demonstrated a consistent downregulation pattern in the CTX + BUS group compared to controls (Fig. [126]6C). To validate these findings at the protein level, western blot analysis confirmed significantly reduced expression of key hub proteins (e.g., RPS6, RPS9, RPS10, RPS15, RPS19) in CTX + BUS-treated sperm (Fig. [127]6D). Notably, Western blot analysis of day 2 post-fertilization embryos derived from CTX + BUS-exposed sperm demonstrated persistently downregulated expression of ribosomal proteins (e.g., RPS6, RPS9) that were reduced in paternal sperm (Fig. [128]6E), indicating paternal transmission of ribosomal impairment to preimplantation embryos. These results collectively implied sperm ribosomal impairment as a critical mediator of early embryonic developmental defects, highlighting a novel paternal contribution of ribosomal integrity to embryogenesis. Discussion The present study elucidates the multifaceted reproductive toxicity of combined CTX + BUS chemotherapy, revealing profound impacts on testicular function, sperm integrity, and paternal contributions to early embryogenesis. Our findings demonstrate that acute CTX + BUS exposure induces systemic germ cell apoptosis, disrupts testicular proteostasis, and triggers sperm head teratozoospermia, ultimately compromising fertilization competence and embryonic developmental potential. Notably, while conventional semen parameters such as sperm concentration and motility remained largely unaffected, proteomic and functional analyses uncovered critical defects in ribosomal biogenesis and chromatin remodeling pathways, implicating paternal non-genetic components as key mediators of chemotherapy-induced embryotoxicity. A central finding of this work is the pronounced downregulation of ribosomal proteins in CTX + BUS-exposed spermatozoa, accompanied by impaired ribosome assembly and cytoplasmic translation pathways. Beyond their canonical role in protein synthesis, ribosomal proteins have been shown to critically regulate spermatogenesis, oogenesis, and embryogenesis [[129]32–[130]38]. The observed depletion of ribosomal subunits in CTX + BUS-exposed sperm, coupled with reduced ribosomal protein abundance in 2-cell stage embryos derived from these sperm, mechanistically links paternal ribosomal insufficiency to zygotic genome activation defects. Such insufficiency may disrupt the synthesis of transcriptional activators required for embryonic genome activation. Previous reports indicate that CTX monotherapy (200 mg/kg for 5 days) primarily induces germ cell apoptosis within seminiferous tubules [[131]39], and prolonged exposure leads to significant reductions in sperm count, motility, normal morphology, and DNA integrity [[132]40, [133]41]. Similarly, Busulfan treatment induces orchitis, disrupts the blood-testis barrier, and triggers spermatocyte apoptosis within two weeks [[134]16, [135]42, [136]43]. Prolonged exposure leads to a near-complete depletion of germ cells, resulting in seminiferous tubules populated primarily with Sertoli cells (SCs) [[137]10, [138]16, [139]42]. Therefore, both the dosage of the drug and the duration of exposure can lead to significant differences in germ cell damage. Future studies employing a range of drug doses will be essential to clarify dose-dependent effects on sperm molecular composition and embryonic developmental potential. Notably, our combined CTX + BUS regimen, despite a shorter exposure period (7 days), elicited a unique phenotype: severe sperm head teratozoospermia and a profound depletion of ribosomal proteins from spermatozoa, without a significant reduction in sperm concentration or gross motility (Fig. [140]3). This contrasts with the overt oligozoospermia and motility deficits commonly reported for individual agents. At the molecular level, however, while individual treatments primarily impact DNA integrity and inflammatory pathways [[141]42], our proteomic profiling uncovered ribosomal biogenesis dysfunction as a novel and central pathway disrupted by the combination therapy. This suggests a synergistic effect wherein CTX and BUS converge on mechanisms critical for ribosomal assembly and post-fertilization translational activation, culminating in paternal-derived embryotoxicity despite the preservation of conventional semen parameters. Thus, our findings underscore that combination chemotherapy can manifest unique molecular lesions that are not predictable from the toxicity profiles of single agents, highlighting the critical need to assess non-genetic sperm quality in patients receiving multi-drug regimens. Notably, our data challenge the clinical reliance on conventional semen parameters for fertility assessment in chemotherapy survivors. Despite preserved sperm counts and motility, CTX + BUS-exposed sperm exhibited severe head malformations and epigenetic instability (Fig. [142]3), correlating with reduced fertilization rates and blastocyst formation (Fig. [143]4). While this study identifies a strong correlation between chemotherapy-induced sperm ribosomal protein deficiencies and early embryonic arrest, the exact causal mechanisms linking sperm head abnormalities to these adverse fertility outcomes require further elucidation. Specifically, the co-occurrence of severe sperm head malformations with reduced fertilization competence raises critical questions about whether these morphological defects directly drive fertilization failure or early developmental defects, independently of or in conjunction with molecular lesions like epigenetic instability or ribosomal protein deficiencies. Future studies will address this gap by employing advanced sperm selection techniques (e.g., microfluidic sorting based on morphological criteria) or intracytoplasmic sperm injection (ICSI) with morphologically classified sperm to disentangle the effects of sperm head morphology from underlying molecular contributions. These observations align with growing evidence that sperm DNA fragmentation and non-coding RNA dysregulation can impair embryogenesis independently of obvious motility defects [[144]44–[145]49]. The persistence of morphologically abnormal sperm with latent molecular lesions underscores the importance of integrating advanced diagnostics, such as proteomic profiling or chromatin integrity assays into fertility risk stratification for cancer survivors. Several limitations warrant consideration. First, our acute exposure model (7-day regimen) may not fully replicate the chronic or cumulative effects observed in human chemotherapy protocols. Notably, as a complete spermatogenic wave in mice takes approximately 35 days, our 7-day post-treatment assessment, which focuses on acute injury induced by combined CTX and BUS chemotherapy, does not evaluate the long-term effects of the treatment on the entire spermatogenic cycle, such as sustained impairments in spermatogonial stem cell function, sperm quality stability, or long-term fertility outcomes. This represents a key limitation of the current study, and investigating these long-term impacts will be a primary focus of our future research. Second, while ribosomal dysfunction emerged as a central theme, we acknowledge that our current work establishes a correlation rather than direct causality between sperm ribosomal protein deficiencies and impaired early embryonic development. However, the exact mechanistic pathways through which these specific ribosomal protein deficiencies disrupt early development, particularly zygotic genome activation (ZGA), remain to be fully delineated. In the future, rescue experiments could be performed by microinjecting mRNAs encoding the downregulated ribosomal proteins into embryos derived from CTX + BUS-exposed sperm to test whether this approach can alleviate developmental defects. Lastly, the translational relevance of these findings requires validation in human sperm samples, particularly given species-specific differences in spermatogenesis and embryonic genome activation timelines. In conclusion, this study establishes ribosome biogenesis defects as novel molecular alterations underlying chemotherapy-induced paternal reproductive toxicity. By linking sperm ribosomal insufficiency to embryonic developmental arrest, we provide a framework for understanding how environmental and iatrogenic insults disrupt intergenerational health. These findings advocate for expanded male fertility assessment protocols that evaluate molecular sperm quality, ultimately guiding interventions to mitigate paternal contributions to adverse developmental outcomes in cancer survivors. Supplementary Information Below is the link to the electronic supplementary material. [146]Supplementary Material 1^ (3.3MB, xlsx) [147]Supplementary Material 2^ (3.6MB, xlsx) [148]Supplementary Material 3^ (10.9KB, xlsx) Acknowledgements