Abstract Background MII oocytes undergo time-dependent aging after ovulation, which is closely associated with impaired fertilization potential, poor embryo quality, and an increased risk of miscarriage. The apoptosis of cumulus cells and their secretion of TNF-α are identified as the primary contributors to postovulatory oocyte aging. Results In this study, we demonstrated that astaxanthin supplementation in culture medium effectively prevents postovulatory oocyte aging and extends oocyte lifespan in vitro. Importantly, this protective effect does not operate through the inhibition of cumulus cell apoptosis or TNF-α release. Notably, astaxanthin selectively binds to TNFR2 in oocytes, thereby preventing TNFR2 from interacting with TNF-α and inhibiting the activation of the TNF signaling pathway within oocytes. Furthermore, oocytes cultured with astaxanthin exhibit enhanced potential for early embryonic development and significantly increased IVF-ET litter size compared to controls. Conclusions Our findings, based on a mouse model, provide valuable insights into the potential clinical application of astaxanthin in mitigating post-ovulatory oocyte aging. Supplementary Information The online version contains supplementary material available at 10.1186/s12915-025-02292-x. Keywords: Astaxanthin, Aging, Oocytes, Cumulus cells, TNF-α, TNFR2 Background Oocytes are arrested at the metaphase of the second meiosis (MII) stage, representing a critical checkpoint in the cell cycle that demands stringent regulation. Such transient arrest is crucial for maintaining tight cell cycle regulation, ensuring optimal oocyte quality and proper post-fertilization development. In humans and most domestic animals, oocytes can retain their fertilization capability for no more than 24 h post-ovulation [[36]1]. MII oocytes following ovulation, if not fertilized in a timely manner, undergo a series of aging-related changes [[37]2, [38]3], including elevated early apoptosis and spindle-chromosome complexes (SCCs) morphological abnormalities [[39]4–[40]7]. Furthermore, postovulatory oocyte aging has detrimental effects on early embryonic development and offspring health [[41]8–[42]10]. Consequently, preventing postovulatory oocyte aging is of significant importance for enhancing the efficiency of assisted reproductive technology (ART) or somatic cell nuclear transfer (SCNT), which require extended manipulation of ovulated oocytes. Cumulus cells play pivotal roles during oocyte ovulation and fertilization [[43]11]. From the early stages of follicle formation, cumulus cells assume the critical function of sensing environmental conditions and communicating this information to the oocyte. Furthermore, cumulus cells significantly influence postovulatory oocyte aging [[44]12]. Oocytes experiencing postovulatory aging are promoted by cumulus cells’ apoptosis and the release of tumor necrosis factor α (TNF-α) [[45]13]. Although apoptosis is a tightly regulated fundamental biological phenomenon, cumulus cell apoptosis can negatively impact oocyte and embryo quality to some extent [[46]14]. TNF-α is a secretory factor that functions in its biologically active homotrimeric form [[47]15]. Primarily, TNF-α acts as a ligand to stimulate receptor oligomerization and activate downstream signaling pathways [[48]16, [49]17]. The receptors for TNF-α are tumor necrosis factor receptor 1 (TNFR1) and tumor necrosis factor receptor 2 (TNFR2) [[50]18], both of which are present in oocytes and implicated in cell apoptosis [[51]19]. Cell apoptosis can be categorized into three pathways based on their modality of initiation: mitochondrial, endoplasmic reticulum, and death receptor pathway-mediated apoptosis [[52]20, [53]21]. The TNF signaling pathway involves extracellular protein ligands (TNF-α) binding to and activating death receptors (TNFR1 or TNFR2) on the cell surface to induce apoptosis, thereby serving as a key member of the death receptor pathway in mediating cell apoptosis [[54]22, [55]23]. Astaxanthin (AX) is a potent ketone carotenoid with both hydrophilic and lipophilic molecular structures [[56]24, [57]25]. Extensive research has demonstrated its remarkable anti-apoptotic properties, including reducing apoptosis induced by cigarette smoke extract [[58]26], extending the lifespan of yeast Saccharomyces cerevisiae [[59]27], and exhibiting anti-apoptotic activity in a hyperoxia-induced retinopathy mouse model [[60]28]. Furthermore, transcriptomic analyses have revealed that AX enhances the anti-apoptotic capacity in crustaceans [[61]29]. These findings collectively highlight the strong anti-apoptotic potential of AX. Despite various approaches to prevent postovulatory oocyte aging, such as modifying culture conditions, mitochondrial replacement technology, and exogenous compound supplementation [[62]5, [63]30, [64]31], the anti-aging effects of AX remain unexplored. This study investigates the protective effects and underlying mechanisms of AX supplementation in preventing postovulatory oocyte aging. Our findings aim to provide novel insights and strategies to mitigate postovulatory oocyte aging, thereby enhancing the success rates of ART and SCNT. Results AX rescues developmental impairment in early embryos derived from postovulatory-aged oocytes Cumulus-oocyte complexs (COCs) were collected from mouse oviducts and randomly assigned to one of three parallel culture models—Control: COCs were fertilized or analyzed immediately without extended in vitro culture; Aging: COCs were cultured for 12 h in AX-free medium prior to fertilization or analysis; Aging + AX: COCs were cultured for 12 h in medium supplemented with AX (2.0 μg/mL) (Fig. [65]1a). To establish optimal in vitro culture durations, COCs were cultured for 0 (Control), 6, 12, 18, or 24 h. Control oocytes exhibited robust fertilization and blastocyst formation, whereas prolonged culture progressively impaired developmental competence. While oocytes cultured for 6 h showed no significant difference from controls, blastocyst formation rates plummeted in the 18- and 24-h groups. Consequently, the 12-h culture duration was selected for the Aging group (Additional File [66]1: Fig. S1a). Next, we determined the optimal AX concentration by culturing COCs for 12 h with 1.0, 1.5, 2.0, or 2.5 μg/mL AX. AX supplementation significantly improved embryonic development, with maximal efficacy at 2.0 μg/mL (designated Aging + AX) (Additional File [67]1: Fig. S1b). Morphological analysis revealed widespread embryo fragmentation and developmental arrest in the Aging group, both of which were markedly reduced by AX treatment (Fig. [68]1b). Quantitatively, fertilization rates in the Aging group were significantly lower than controls (P < 0.05), whereas AX restored early embryonic development rates to near-control levels (Fig. [69]1c). These results demonstrate that AX rescues the developmental impairment caused by postovulatory aging. To evaluate AX’s impact on offspring viability, we performed in vitro fertilization and embryo transfer (IVF-ET). Conception rates did not differ significantly among Control, Aging, and Aging + AX groups (Fig. [70]1d; P > 0.05). However, litter size was severely reduced in the Aging group but restored by AX treatment (Fig. [71]1e, f; P < 0.05). No differences were observed in offspring sex ratios, birth weight, weekly body weight (1–8 weeks), survival rates (0–8 weeks), or hematological parameters across groups (Additional File [72]1: Fig. S1c–n; P > 0.05). Collectively, these findings indicate that AX supplementation during extended in vitro culture mitigates postovulatory aging-induced defects in early embryo development. Fig. 1. [73]Fig. 1 [74]Open in a new tab AX reverses the impaired developmental potential of early embryos originating from postovulatory aging oocytes. a Experimental design. Ovarian cumulus-oocyte complexes (COCs) were collected from oviducts for the Control (no in vitro culture), Aging (cultured in vitro for 12 h), and Aging + AX groups (cultured in vitro for 12 h with AX supplementation). b Representative images of early embryonic development at different stages. Red, yellow, green, and blue arrows indicate stunted or fragmented embryos at the 2-cell, 4-cell, morula, and blastocyst stages, respectively. Scale bar = 100 μm. c Percentages of 2-cell, 4-cell, morula, and blastocyst oocytes; the number of oocytes is shown in parentheses. d Pregnancy rate; the number of pseudopregnant females is shown in parentheses. e Representative images of offspring generated from IVF-ET. f Litter size; the number of pregnant mice are shown in parentheses. Data are expressed as means ± SD. Significance markers: *, p < 0.05; **, p < 0.01; ***, p < 0.001; ****, p < 0.0001; ns, p > 0.05 AX mitigates apoptosis and spindle abnormalities in postovulatory-aged oocytes To elucidate how AX supplementation counteracts oocyte aging during extended in vitro culture, we first analyzed cumulus cell apoptosis and TNF-α dynamics. Flow cytometry revealed minimal apoptosis in Control cumulus cells, whereas both Aging and Aging + AX groups exhibited widespread apoptosis (Additional File [75]1: Fig. S2a, b). Immunofluorescence localized TNF-α predominantly to the cytoplasm of Control cumulus cells, yet signal intensity was markedly reduced in Aging and Aging + AX groups (Additional File [76]1: Fig. S2c). Quantification confirmed no significant differences in TNF-α fluorescence, protein, or mRNA levels between Aging and Aging + AX groups (Additional File [77]1: Fig. S2d–f; P > 0.05). ELISA further showed elevated TNF-α in culture media from Aging and Aging + AX groups compared to Controls, though no intergroup difference was observed (Additional File [78]1: Fig. S2g; P > 0.05). These results suggest that cumulus cells release TNF-α into the medium during prolonged culture, and AX neither suppresses cumulus cell apoptosis nor scavenges secreted TNF-α. To confirm TNF-α’s role in oocyte aging, exogenous TNF-α (10 or 100 ng/mL) was added to culture media. Both concentrations significantly impaired embryonic development, but AX co-treatment restored developmental rates (Additional File [79]1: Fig. S2h, i; P < 0.05). Oocyte perivitelline space width, a marker of cytoplasmic integrity, increased by 50% in the Aging group compared to Controls (P < 0.05), while AX restored it to baseline levels (Fig. [80]2a, b). Degeneration rates, characterized by cytoplasmic darkening and shrinkage, rose from 4.15 ± 0.53% (Control) to 19.24 ± 1.62% in the Aging group (P < 0.05), but AX reduced this to 14.82 ± 1.61% (Fig. [81]2c, d). Annexin-V staining revealed early apoptosis in 6.38 ± 3.02% of Control oocytes versus 46.11 ± 4.23% in the Aging group (P < 0.01), with AX lowering apoptosis to 27.00 ± 4.10% (Fig. [82]2e, f). Tubulin staining demonstrated abnormal spindle formations in 48.72 ± 5.33% of Aging group oocytes, which AX reduced to 26.98 ± 2.24% (Fig. [83]2g, h; P < 0.05). Spindle length-to-oocyte diameter ratios, a metric of structural integrity, declined in the Aging group but rebounded with AX treatment (Fig. [84]2i). Collectively, these findings demonstrate that AX ameliorates postovulatory aging by preserving oocyte morphology, reducing apoptosis, and stabilizing spindle architecture during extended in vitro culture. Fig. 2. [85]Fig. 2 [86]Open in a new tab AX reduces the occurrence of apoptosis and abnormal spindle-chromosome complexes configurations in postovulatory aging oocytes. a Representative images of perivitelline space. Square black frame indicates the region shown in detail. Scale bar = 20 μm. b Quantification of perivitelline space distance in the Control, Aging, and Aging + AX groups. c Representative images of degenerated oocytes. The red arrow indicates oocyte degeneration, and square black frame indicates the region shown in detail. Scale bar = 100 μm. d Ratio of degenerated oocytes. e Representative images of early apoptotic cells. Early apoptotic oocytes exhibit distinct green membrane signals. Scale bar = 20 μm. f Percentages of oocytes undergoing early apoptosis. g Representative images of SCCs. The Control group shows normal SCCs (left column). The Aging groups display abnormal SCCs exhibited as spindle assemble disorder, chromosome condensation failure, and spindle chromosome malformation (three middle columns). The Aging + AX groups display normal and abnormal SCCs (two right columns). The circular white frame marks the oocyte edge, and the white square indicates the detailed region. Green, tubulin; blue, DNA. Scale bar = 20 μm. h Percentages of abnormal SCCs in the Control, Aging, and Aging + AX groups. i Ratio of spindle length (SL) to oocyte diameter (OD). Data are expressed as means ± SD. Significance markers: *, p < 0.05; **, p < 0.01; ***, p < 0.001; ****, p < 0.0001; ns, p > 0.05. The number of oocytes is indicated in parentheses Transcriptome analysis implicates TNF signaling in postovulatory oocyte aging To delineate AX’s anti-apoptotic mechanism, we performed Microcell transcriptome sequencing on Control, Aging, and Aging + AX groups. An UpSet plot highlighted distinct and overlapping gene expression patterns across groups, with unique and shared gene clusters color-coded for clarity (Fig. [87]3a). Heatmap analysis identified 73 differentially regulated signaling pathways, 14 of which, including MAPK, PI3K-Akt, and TNF signaling, were directly linked to apoptosis (Fig. [88]3b). Strikingly, the TNF signaling pathway emerged as the sole pathway dysregulated in both Aging vs. Control and Aging + AX vs. Aging comparisons, while remaining unaffected between Aging + AX and Control groups (Fig. [89]3c). Gene set enrichment analysis (GSEA; significance threshold: |NES|> 1, P < 0.05, q < 0.25) revealed marked suppression of TNF signaling in Aging vs. Control oocytes (NES = − 1.525, P = 0.025, q = 0.140; Fig. [90]3d). No pathway perturbation was observed between Aging + AX and Control groups (NES = − 1.018, P = 0.424, q = 0.064; Fig. [91]3e), whereas AX robustly reversed TNF signaling in Aging + AX vs. Aging comparisons (NES = 1.771, P = 0.002, q = 0.019; Fig. [92]3f). Heatmap profiling of TNF signaling components identified downregulation of Akt2, Akt1, Dab2ip, Cebpb, Atf6b, and Jag1 in Aging vs. Control oocytes. AX supplementation selectively rescued Dab2ip and Cebpb expression to control levels (Fig. [93]3g), a finding validated by qPCR (Additional File [94]1: Fig. S3a, b). Protein interaction network analysis further positioned DAB2IP and CEBPB as central nodes coordinating TNF-mediated apoptotic responses (Additional File [95]1: Fig. S3c). Fig. 3. [96]Fig. 3 [97]Open in a new tab Transcriptome analysis revealed abnormalities in the TNF signaling pathway in postovulatory aging oocytes. a Upset plot showing unique or common genes; dots of different colors represent unique genes, while lines of different colors represent shared genes. b Heatmap illustrating the signaling pathways enriched with genes in oocyte. c Spin plot showing the 14 signaling pathways directly related to oocyte apoptosis, with the TNF signaling pathway highlighted by a red frame. d GSEA diagram showing TNF signaling pathway activity in the Control vs. Aging groups. e GSEA diagram showing TNF signaling pathway activity in the Control vs. Aging + AX groups. f GSEA diagram showing TNF signaling pathway activity in the Aging vs. Aging + AX groups. g List of differentially expressed genes in TNF signaling pathway Astaxanthin suppresses TNF signaling activation in postovulatory-aged oocytes To validate AX’s impact on the TNF pathway, we analyzed key protein expression across Control, Aging, and Aging + AX groups. TNFR1 levels remained unchanged among all groups (Additional File [98]1: Fig. S4a, b; P > 0.05). In contrast, TNFR2 expression increased significantly in the Aging group (P < 0.05) but reverted to baseline with AX treatment (Fig. [99]4a, b). Downstream of TNFR2, TRAF2 expression rose markedly in the Aging group (P < 0.05) and declined to control levels upon AX supplementation (Fig. [100]4c, d). TRAF2 regulates DAB2IP, a critical apoptosis modulator. DAB2IP expression decreased in the Aging group (P < 0.05) but recovered with AX treatment (Fig. [101]4e, f). As TNFR2 primarily activates P38/JNK signaling, we assessed phospho-P38 (P-P38) and phospho-JNK (P-JNK). P-P38 levels surged in the Aging group (P < 0.05) and normalized with AX (Fig. [102]4g, h), whereas P-JNK remained unaffected (Additional File [103]1: Fig. S4c, d; P > 0.05). Elevated P-P38 drives caspase-dependent apoptosis, consistent with increased Caspase3 in Aging oocytes (P < 0.05) and its reduction by AX (Fig. [104]4i, j). BAX expression, a proapoptotic marker, rose significantly in the Aging group (P < 0.05) but decreased with AX treatment (Fig. [105]4k, l). Conversely, antiapoptotic BCL-2 levels showed no intergroup differences (Additional File [106]1: Fig. S4e, f; P > 0.05). These data demonstrate that AX counteracts TNF pathway activation in aged oocytes, restoring apoptotic protein homeostasis. Fig. 4. [107]Fig. 4 [108]Open in a new tab AX inhibits the activation of TNF signaling pathway in postovulatory aging oocytes. a TNFR2 protein expression in oocytes. b Quantification of TNFR2/TUBB relative intensity. c Expression of TRAF2 protein in oocytes. d Quantification of TRAF2/TUBB relative intensity. e Expression of DAB2IP protein in oocytes. f Quantification of DAB2IP/TUBB relative intensity. g Expression of P-P38 protein in oocytes. h Quantification of P-P38/TUBB relative intensity. i Expression of CASPASE3 protein in oocytes. j Quantification of CASPASE3/TUBB relative intensity. k Expression of BAX protein in oocytes. l Quantification of BAX/TUBB relative intensity. For protein detection, 200 oocytes were used per lane, with TUBB serving as the internal control. Data are expressed as means ± SD. Significance markers: *, p < 0.05; **, p < 0.01; ns, p > 0.05. Number of biological repeats is shown in parentheses AX preferentially binds to TNFR2 in oocytes and prevents TNF-α from binding with TNFR2 The detailed mechanism by which AX supplementation inhibits the activation of the TNF signaling pathway in postovulatory aging oocytes was explored. Immunofluorescence results showed that TNF-α was highly expressed in aging oocytes, while TNFR2 expression was also elevated. However, in AX supplemented oocytes, TNF-α membrane expression was significantly reduced and TNFR2 cytoplasmic expression decreased (Additional File [109]1: Fig. S5a–d). The ligand TNF-α exerts a biological effect in the form of a homologous trimer, and the receptor TNFR2 undergoes trimerization when stimulated by the ligand TNF-α. Immunofluorescence revealed that TNF-α combines with TNFR2 in the absence of AX and that TNF-α did not combine with TNFR2 in the presence of AX (Fig. [110]5a). Co-IP experiments confirmed the interaction between TNF-α and TNFR2 in oocytes (Fig. [111]5b). Also, the crystal structure shows that the trimer composed of monomer TNF-α (green) can combine with the trimer composed of monomer TNFR2 (red) (Fig. [112]5c). We hypothesized that AX supplementation preferentially binds to TNFR2 in oocytes and prevents TNF-α from binding with TNFR2. PyMOL software was used to separate the TNFR2 protein monomer structure (Fig. [113]5d). The AX molecular structure was downloaded from the ZINC database (Fig. [114]5e). Molecular docking was performed between TNFR2 and AX via Discovery Studio software. The domain location of the TNFR2 amino acid sequence includes signal peptide (1–22), extracellular peptide (23–257), helical peptide (258–287), and cytoplasmic peptide (288–461) (Fig. [115]5f). Molecular docking studies revealed that AX binds to the extracellular domain of TNFR2, forming van der Waals forces and hydrophobic interactions at TYR45, TYR46, ASP47, GLN48, THR49, PHE66, TRP89, PRO91, and GLU92 (Fig. [116]5g). Conservation analysis across species showed identical interaction sites at TYR45, TYR46, ASP47, and PHE66 in humans, mice, and rats (Fig. [117]5h). Experimental results demonstrated that AX treatment significantly reduced TNF-α trimer and monomer expression in aging oocytes (Fig. [118]5i–k). Drug affinity responsive target stability (DARTS) and cellular thermal shift assay (CETSA) experiments confirmed AX’s binding to TNFR2 (Fig. [119]5l). In the absence of AX, the TNFR2 protein level showed a gradual decline with increasing enzymolysis time at a pronase concentration of 1:500, whereas its stability was maintained in the presence of AX (Fig. [120]5m). Without AX, TNFR2 protein became undetectable after 15 min of enzymolysis, regardless of high (1:50) or low (1:5000) enzyme concentrations. However, when AX was present, TNFR2 expression exhibited a notable impact influenced by enzyme concentration (Fig. [121]5n). Furthermore, AX demonstrated effective protection against TNFR2 degradation under elevated temperatures (Fig. [122]5o). At 55 °C, AX at 2.0 μg/mL showed superior protective effects compared to 0.2 μg/mL (Fig. [123]5p). These results suggest that AX preferentially bind to TNFR2, preventing TNF-α from binding and inhibiting downstream TNF signaling pathways activation in aging oocytes. Fig. 5. [124]Fig. 5 [125]Open in a new tab AX preferentially binds to TNFR2 in oocytes and prevents TNF-α from binding with TNFR2. a Representative images showing TNF-α and TNFR2 colocalization in oocytes without or with AX treatment. Green, TNF-α; red, TNFR2; blue, DNA. Scale bar = 20 μm. b Co-IP assay demonstrating TNF-α binding to TNFR2 in oocytes. c Crystal structure of TNF-α combined with TNFR2. d Structure of the TNFR2 monomer. e Molecular structure of AX. f Structure model of AX binding to TNFR2. g Detailed binding location and force analysis of AX-TNFR2 interaction. Green, van der Waals; red, hydrophobic. h Amino acid sequences of TNFR2 in humans, mice, and rats. Key interaction sites are labeled in green (binding site 1) and red (binding site 2). i Expression levels of TNF-α monomer and trimer proteins in oocytes. j Quantification of TNF-α monomer/TUBB relative intensity. k Quantification of TNF-α trimer/TUBB relative intensity. l Schematic diagram illustrating the experimental workflow for validating AX-TNFR2 interaction using DARTS and CETSA. m DARTS (enzymatic hydrolysis time experiment): TNFR2 protein expression after treatment with pronase at 1:500 concentration for 5, 15, or 45 min. n DARTS (pronase concentration experiment): TNFR2 protein expression after pronase treatment at 1:50, 1:500, or 1:5000 concentrations for 15 min. o CETSA (temperature experiment): TNFR2 protein expression after treatment at 35 °C, 45 °C, 55 °C, or 65 °C. p CETSA (AX concentration experiment): TNFR2 protein expression after AX supplementation at 0.2 or 2.0 μg/mL at 55 °C. Data are expressed as means ± SD. Significance markers: *, p < 0.05; **, p < 0.01; ns, p > 0.05. The number of biological repeats is shown in parentheses Discussion This study demonstrates that AX supplementation enhances early embryonic development competence and litter size following IVF-ET in postovulatory aging oocytes, while concurrently reducing oocyte apoptosis. Mechanistically, AX exerts its protective effects by specifically targeting TNFR2 to inhibit the TNF signaling pathway within oocytes, providing a robust foundation for strategies to mitigate postovulatory aging in clinical and experimental contexts. ART remains the primary intervention for infertility caused by disease, environmental stressors, or psychological factors [[126]32, [127]33]. However, the prolonged in vitro manipulation poses a risk of accelerating postovulatory aging [[128]34]. This process is closely linked to cellular and molecular alterations that significantly compromise oocyte quality, embryonic developmental potential, and offspring health [[129]4, [130]9, [131]10]. Our findings reveal that AX significantly improves the developmental capacity of postovulatory-aged oocytes. These results align with Liu et al.’s work, which reported that 12-h postovulatory aging reduced 2-cell and blastocyst formation rates to 13.6% and 1.6%, respectively, while 1 μM auranetin treatment restored these rates to 47.6% and 26% [[132]35]. This underscores the potential of exogenous compounds to rescue fertilization outcomes in aged oocytes. In IVF-ET experiments, blastocysts derived from aged oocytes produced markedly smaller litters in surrogate mothers, whereas AX supplementation restored litter size. We hypothesize that while aged oocytes can form blastocysts in vitro, their intrinsic quality limitations hinder post-implantation development. AX likely enhances both blastocyst quality and quantity, thereby improving reproductive outcomes. Notably, offspring from AX-treated oocytes exhibited no hematological abnormalities, though comprehensive pharmacokinetic and toxicological studies are required to confirm AX’s safety profile. Cumulus cells exhibit poor in vitro survival and undergo early apoptosis [[133]36], which accelerates oocyte aging via TNF-α secretion [[134]13]. While we observed similar trends, AX did not significantly reduce cumulus cell apoptosis or TNF-α release. This suggests that cumulus cell apoptosis precedes oocyte apoptosis in vitro, with apoptotic cumulus cells releasing TNF-α into the culture medium. AX neither mitigates cumulus cell apoptosis nor acts as an efficient TNF-α scavenger, resulting in elevated TNF-α levels that exacerbate oocyte aging. Postovulatory aging frequently induces oocyte degeneration [[135]37], a phenotype ameliorated by AX in our study. We attribute this improvement to AX-mediated suppression of TNF-α’s degenerative effects, delaying oocyte degeneration. Prior studies corroborate that exogenous compounds attenuate postovulatory aging by reducing early apoptosis [[136]38, [137]39], consistent with our findings. TNF-α also drives structural alterations in SCCs [[138]37], which AX supplementation significantly mitigated, aligning with Li et al.’s observations [[139]40]. Collectively, these data suggest that AX extends oocyte viability by curbing apoptosis during postovulatory aging. MicroRNA sequencing, a sensitive transcriptomic tool for detecting low-abundance genes [[140]41], highlighted the TNF signaling pathway as central to AX’s anti-aging effects. While somatic cells typically activate TNFR1 in response to TNF-α [[141]42], oocytes predominantly express TNFR2, which exhibits heightened sensitivity to TNF-α in ovarian tissue [[142]19]. This receptor selectivity may underlie the unique responses observed in oocytes. Our findings demonstrate that TNF-α binding TNFR2 activates the P38 pathway via TRAF2 and DAB2IP [[143]43]. Furthermore, DAB2IP also plays a pivotal role in P38 activation [[144]44, [145]45]. These observations align with our results, underscoring that the TNF-α-TNFR2-TRAF2-DAB2IP-P38 axis represents a key and altered component of the TNF signaling pathway. Results of expression levels of CASPASE3 and BAX, proteins associated with apoptosis, mirror those of resveratrol in its ability to prevent postovulatory oocyte aging [[146]46]. Exogenous compounds have been shown to mitigate apoptotic protein expression. Notably, AX supplementation has been reported to enhance oocyte quality by alleviating oxidative stress and reducing CASPASE3 activity in aging oocytes [[147]47]. While our experimental outcomes share similarities with previous studies, our work provides novel insights into the detailed molecular mechanisms by which AX supplementation in culture medium exerts its anti-apoptotic effects to prevent postovulatory oocyte aging. The interaction between AX and TNFR2 was determined using molecular docking technology (as no accurate mouse TNFR2 protein structures are currently available, the human TNFR2 protein structure was used for analysis). While the amino acid sites of TNFR2 differ between humans and mice, nearly half of the interaction sites between AX and TNFR2 are conserved, suggesting these sites are highly conserved and may play a more significant role. DARTS and CETSA, established methodologies for validating small molecule-protein interactions [[148]48]. were employed to confirm these findings. For instance, Wu et al. demonstrated that protocatechuic acid protects cardiac cells from ischemic injury through PKM2 modulation [[149]49], while bavachinin targets PCNA to protect liver cells in nonalcoholic fatty liver disease [[150]48]. Our DARTS and CETSA experiments confirmed that AX supplementation prevents TNFR2 degradation caused by elevated temperature or prolonged enzymatic hydrolysis. This suggests that TNFR2 is the most likely target of AX. In the culture medium, AX preferentially binds TNFR2, thereby preventing TNF-α- TNFR2 interaction and inhibiting TNF signaling pathway activation in oocytes. This interaction may explain AX’s role in preventing postovulatory oocyte aging. However, further in vivo experiments are needed to validate this mechanism. Notably, mice, as a species capable of rapid meiotic resumption and polyovulation, exhibit significant physiological differences from humans, who experience prolonged meiotic arrest and monoovulation. These species-specific differences suggest that this approach may not be directly applicable to the human assisted reproductive system. Nevertheless, it provides valuable insights and a foundation for future research on anti-aging strategies. Conclusions In summary, our findings in a mouse model suggest that AX supplementation has the potential to prolong the in vitro culture time of oocytes. Moreover, AX inhibits the activation of the TNF signaling pathway by specifically targeting TNFR2, thereby reducing oocyte apoptosis. This work lays a foundation for gaining a deeper understanding of the detailed molecular mechanism by which AX delays postovulatory oocyte aging. Methods Experimental design All experimental procedures adhered to the National Research Council Guide for the Care and Use of Laboratory Animals and were approved by the Institutional Animal Care and Use Committee at Inner Mongolia University (Approval number: SYXK2020-0006). ICR mice were purchased from SBF Biotechnology Co., Ltd., and housed in a specific pathogen-free (SPF) animal facility under controlled conditions (20–22 °C, 40–70% humidity, 12-h light/dark cycle) at the university’s animal research center. COCs at MII stage were collected from 8-week-old female mice. The collected COCs were randomly assigned to three groups: (1) Control group (no in vitro culture), (2) Aging group (cultured in vitro for 12 h), and (3) Aging + AX group (cultured in vitro for 12 h with 2.0 μg/mL AX supplementation). Where necessary to assess the effect of TNF-α on embryo development, a specific concentration of TNF-α (HY-P7090, MCE, NJ, USA) was added to the culture medium. Cumulus cell isolation and oocyte collection The female mice underwent intraperitoneal injection of 5 IU of equine chorionic gonadotropin (eCG; San Sheng, Ningbo, Zhejiang, China), followed by the administration of 5 IU of human chorionic gonadotropin (hCG; San Sheng) 48 h later. The COCs were collected 14 h after hCG injection by releasing them from oviducts and placed into M2 medium containing 0.3 mg/mL hyaluronidase. The cumulus cells and oocytes were then collected for subsequent experimental studies. IVF, embryo culture and embryo transfer Adult male ICR mice (12–14 weeks of age) were used for sperm collection. Sperm suspensions were capacitated in 200 µL of T6 medium supplemented with 10 mg/mL bovine serum albumin (BSA) for 2 h. MII oocytes were fertilized by incubating with the capacitated spermatozoa at a concentration of 1 × 10^6/mL in 200 µL of T6 medium containing 20 mg/mL BSA for 6 h. For embryo culture, zygotes were collected and cultured in Chatot-Ziomek-Bavister (CZB) medium supplemented with 3 mg/mL BSA without glucose under a humidified atmosphere of 5% CO[2] at 37 °C for the first 48 h. Once embryos reached the 4-cell stage, they were transferred to CZB medium supplemented with 5.5 mM glucose. Embryonic development was monitored at 24, 48, 72, and 96 h post-fertilization to determine the percentages of embryos at the 2-cell, 4-cell, morula, and blastocyst stages, respectively. For embryo transfer, male mice with good mating records were selected for vasectomy. The male mice were anesthetized via intraperitoneal injection of isoflurane and underwent vasectomy according to the method of Bermejo-Alvarez [[151]50]. After a two-week recovery period, pseudopregnant recipient female mice were obtained by natural mating with vasectomized males. The vaginal mating plugs of female mice were checked daily. Females with mating plugs were selected and recorded as pseudopregnant recipients at 0.5 days post coitum (dpc). Blastocyst transfer was performed on pseudopregnant recipients at 2.5 dpc using the NSET device as described [[152]51, [153]52]. Each NSET device was loaded with 7 blastocysts, and the tip of the device was inserted into the speculum. By fully depressing the plunger, the blastocysts were released into one uterine horn. The procedure was repeated to transfer an additional 7 blastocysts into the opposite uterine horn. In total, 14 blastocysts were transplanted into each recipient mouse. Offspring assessments All offspring in this study were derived from IVF-ET. Pregnancy rates were determined at delivery, and parameters including litter size, birth weight, sex ratio, survival rate of mice from 0 to 8 weeks of age, and growth status from 1 to 8 weeks were evaluated. For blood analysis, samples were collected via ocular puncture and transferred to EDTA-containing anticoagulant tubes. The tubes were gently inverted to ensure proper mixing. Following the manufacturer’s instructions for the ABX PENTRA80 series hematology analyzer (Horiba Medical, Kyoto, Japan), 50 μL of blood was aspirated and analyzed. The instrument was programmed with appropriate detection parameters to assess hematological parameters such as large immature cell percentage, RBC count, hematocrystallin, basophil percentage, phagocytic neutrophil percentage, lymphocyte ratio, hematocrit, and platelet count. Immunofluorescence and cell staining assay Immunofluorescence detection was conducted as previously described [[154]53]. The following primary antibodies were used: mouse anti-tubulin (1:1000; ab6046; Abcam, Cambridge, UK), mouse anti-TNF-α (1:500; ab1793; Abcam), and rabbit anti-TNFR2 (1:500; 10366-1-AP; Proteintech). For secondary antibodies, DyLight 488-conjugated goat anti-mouse (1:1000; Jackson ImmunoResearch Laboratories, West Grove, PA, USA) or TRITC-conjugated donkey anti-rabbit (1:1000; Jackson ImmunoResearch Laboratories) antibodies were used. The cell staining assay was conducted as previously described [[155]53]. For apoptosis experiments, cells were stained with Annexin-V (G003-1; Jian Cheng Bioengineering Institute) and DNA was stained with Hoechst 33342 (ab228551, Abcam). After washing, the samples were mounted onto glass slides and examined using a confocal laser-scanning microscope (Nikon A1R). RNA-seq and bioinformatics analysis Forty oocytes from each of the Control, Aging, and Aging + AX groups were collected as biological replicates for RNA transcript sequencing using the Smart-Seq 2 method. To ensure optimal library quality, cDNA library quantification was performed using qPCR, with a minimum effective concentration of 2 nM required. qPCR was employed to precisely quantify the library by amplifying fragments with complete adapter connections at both ends, ensuring accurate measurement of target DNA. The library quantification process included preparing a standard curve, preliminary quantification using a Qubit fluorometer, and fragment size analysis with an Agilent 2100 Bioanalyzer. The final library concentration was calculated using the formula: original library concentration (nM) = (standard length (bp)/library average length (bp)) × diluted library concentration (pM) × dilution factor/1000. Libraries were pooled according to sequencing requirements and sequenced on the Illumina NovaSeq 6000 platform. Differentially expressed genes were identified based on P values < 0.05, FDR < 0.05, and log2(FPKM) > 1.5. Pathway enrichment analysis was conducted using GSEA and KEGG, with all bioinformatics analyses performed on the BMK Cloud Platform. The gene expression data for oocytes are summarized in Additional file [156]2: Table S1. Protein‒protein interaction network (PPI) construction The PPI network was constructed using the Search Tool for the Retrieval of Interacting Genes (STRING) database (version 11.5, [157]https://cn.string-db.org/) and visualized with Cytoscape software (version 3.7.2). Real-time quantitative PCR To validate the transcriptomic gene expression data, total RNA was extracted from oocytes (Control, Aging, and Aging + AX groups) using the TaKaRa MiniBEST Universal RNA Extraction Kit (KIT0204, Thermo Fisher Scientific, Waltham, MA, USA). RNA concentrations and quality ratios were assessed using a Nanodrop 1000 spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA). cDNA was synthesized from 500 ng of total RNA using the PrimeScript RT Reagent Kit (RR047A, TaKaRa, Dalian, Liaoning, China) in a 20-μl reaction volume following the manufacturer’s protocol. The reverse transcription process included a single cycle with incubation periods of 65 °C for 5 min, 25 °C for 10 min, and 50 °C for 50 min, followed by final incubation at 37 °C for 20 min. qPCR reactions were performed on a LightCycler 480 system (Roche Applied Science, Indianapolis, IN, USA) using SYBR Green I Master Mix (Roche Applied Science). Primers were designed using Primer 3.0 software, with GAPDH as the reference gene. The primer sequences for Dab2ip and Cebpb are provided in Additional file [158]2: Table S2. Primer efficiencies were evaluated by amplifying serially diluted cDNA samples and calculated using the equation: E = (10^(− 1/slope) − 1) × 100. Efficiencies ranged from 90 to 110%. Each reaction mixture (15 μl) contained 7.5 μl of SYBR Green I Master Mix, 10 μM of each primer, and 20 ng of cDNA. All reactions were performed in triplicate, with non-template controls and a calibrator included in each run. The qPCR protocol consisted of an initial denaturation at 95 °C for 10 min, followed by 45 cycles of 95 °C for 10 s, 60 °C for 10 s, and 72 °C for 10 s. Melting curve analysis was conducted to confirm the specificity of the qPCR products. Gene expression levels were normalized to GAPDH expression using the 2 − ΔΔCt method, following the MIQE guidelines [[159]54]. The MIQE Checklist used in this study is provided as Additional file [160]2: Table S3. Immunoprecipitation (IP) and immunoblotting IP was performed using the Co-Immunoprecipitation Kit (88828, Thermo Fisher Scientific) according to the manufacturer’s protocol. A total of 1000 MII-stage oocytes were lysed in IP buffer (20 mM Tris-HCl, 10 mM EDTA, 1 mM EGTA, 150 mM NaCl, 0.05% Triton X-100, 0.05% NP-40, 1 mM PMSF) supplemented with protease inhibitors (Sigma-Aldrich P8340, 1:100) and phosphatase inhibitors (Sigma-Aldrich P5726, 1:500). Protein A/G agarose beads were washed and incubated with anti-TNF-α or anti-TNFR2 antibodies at 4 °C for 4 h. The antibody-bead complexes were collected and incubated with the lysate at 4 °C overnight. After washing three times, the beads were boiled in SDS-PAGE sample buffer for 5 min. SDS-PAGE was performed, and immunoblotting was conducted as follows: MII-stage oocytes were lysed in 2 × loading buffer containing protease inhibitors, separated on 12–15% bis-tris gels, and transferred onto nitrocellulose membranes. The membranes were blocked in non-fat milk at 37 °C for 1 h and incubated overnight at 4 °C with primary antibodies against TNF-α (1:300; ab1793, Abcam), TNFR1 (1:500; 21574-1-AP, Proteintech), TNFR2 (1:500; 19272-1-AP, Proteintech), TRAF2 (1:500; 26846-1-AP, Proteintech), DAB2IP (1:500; 23582-1-AP, Proteintech), P-P38 (1:500; WLP1576, Wanleibio), P-JNK (1:500; WL01813, Wanleibio), CASPASE3 (1:500; WL04004, Wanleibio), BAX (1:500; WL01637, Wanleibio), BCL-2 (1:500; WL01556, Wanleibio), or Tubulin (1:1000; ab6046, Abcam). Membranes were washed in TBST and incubated with horseradish peroxidase-conjugated secondary antibodies for 1.5 h at room temperature. Proteins were visualized using the ECL Plus system (GE, Piscataway, NJ, USA) and imaged with a Tanon 3900 system (Tanon, Beijing, China). Flow cytometry detection Cumulus cells were collected and resuspended to a concentration of 1 × 10⁶ cells/mL. The cell suspension was centrifuged at 1000 rpm for 5 min, and the supernatant was discarded. This washing procedure was repeated twice with pre-cooled PBS buffer. The washed cell pellet was resuspended in 100 μL of Binding Buffer, followed by gentle mixing. According to the Annexin V-FITC/PI apoptosis detection kit instructions (C1062M, Byotime, Shanghai, China), 5 μL of Annexin V-FITC and 5 μL of PI were added to the cell suspension, which was then incubated in the dark at room temperature for 15 min. After incubation, 400 μL of Binding Buffer was added, and the sample was gently mixed. Flow cytometry analysis was performed using a Cytoflex LX system (Beckman Coulter, CA, USA), with appropriate laser settings and detector parameters configured. Annexin V-FITC was excited at 488 nm and detected in the FITC channel (FL1), while PI was excited at 488 nm and detected in the PI channel (FL3). Compensation was set using single-stained cells to eliminate fluorescence overlap. A flow rate of 10–30 μL/min was used to acquire fluorescence signals from at least 10,000 cells. Data analysis was performed using CytExpert software, with cell gating applied to identify target cells based on FSC vs. SSC scatter plots. Cells were categorized into four regions on the Annexin V-FITC/PI scatter plot: (1) Annexin V-FITC⁻/PI⁻ (normal cells), (2) Annexin V-FITC⁺/PI⁻ (early apoptotic cells), (3) Annexin V-FITC⁺/PI⁺ (late apoptotic/necrotic cells), and (4) Annexin V-FITC⁻/PI⁺ (necrotic cells). The percentage of cells in each category was calculated. All procedures followed MIFlowCyt guidelines and ISAC requirements [[161]55, [162]56]. The MIFlowCyt Checklist used in this study is provided as Additional file [163]2: Table S4. Enzyme-linked immunosorbent (ELISA) assay According to the instructions of the TNF-α ELISA kit (CSB-EQ023955MO, CUSABIO, Wuhan, Hubei, China), the standard product was diluted using the provided dilution buffer. A pre-coated TNF-α antibody microplate was set up with designated wells for standards, samples, and blank controls. The blank control wells were filled with 100 μL of sample dilution buffer, while the standard wells were sequentially loaded with different concentrations of the standard product (100 μL per well). Sample wells received 100 μL of processed samples. The microplate was gently mixed and sealed with a plate sealer film, then incubated at 37 °C for 90 min. After incubation, the sealing film was carefully removed, and the liquid in the wells was discarded by flicking. Each well was washed five times with 350 μL of wash buffer, soaking for 1 min each time, followed by blotting on absorbent paper. Next, 100 μL of detection antibody working solution was added to each well, and the plate was resealed and incubated at 37 °C for 60 min. The washing steps were repeated to ensure complete removal of unbound detection antibodies. Then, 100 μL of enzyme conjugate working solution was added to each well, and the plate was sealed and incubated at 37 °C for 30 min. After incubation, the washing steps were repeated to remove any unbound enzyme conjugates. Subsequently, 90 μL of substrate solution was added to each well, and the plate was gently mixed and incubated at room temperature in the dark for 15 min, during which the solution gradually changed color. Next, 50 μL of stop solution was added to each well, and the plate was gently mixed. Within 15 min after adding the stop solution, the microplate was placed in a microplate reader set to a detection wavelength of 450 nm to measure the optical density (OD) values of each well. A standard curve was constructed by plotting the standard concentrations against their corresponding OD values using professional graphing software. The concentration of TNF-α in the samples was calculated based on the standard curve equation using the OD values from the sample wells. Molecular docking The binding of AX to TNFR2 was analyzed using molecular docking and virtual screening technology. Briefly, the AX molecular structure (ZINC database: ZINC000100042059) was based on the human TNFR2 structure (PDB code: 3ALQ). Using Discovery Studio 2018 software (Accelrys, San Diego, CA, USA), the receptor-binding pocket was identified, the small-molecule structure was prepared, and accurate docking was performed. The results were visualized and optimized using Chimera software. Drug affinity-responsive target stability MII oocytes were lysed at low temperatures in a lysis buffer containing protease inhibitors for 30 min. The homogenate was assayed for protein concentration via a BCA assay. The protein was diluted to the corresponding concentration using a mixture of 10 × TNC (500 mM Tris HCl, 500 mM NaCl, and 100 mM CaCl2, pH 8.0) and aliquoted into six equal tubes. Each tube was treated with either AX or DMSO alone, mixed thoroughly, and incubated at room temperature for 2 h. Pronase solution (10 mg/mL) was diluted at ratios of 1:50, 1:500, or 1:5000. The AX homogenate, with or without AX, was incubated with each pronase dilution for 5, 15, or 45 min. The reactions were terminated by adding loading buffer and boiling sample, and the results were analyzed by western blotting. To ensure reliable and reproducible results, three biological replicates were performed in the DARTS assay. Cellular thermal shift assay MII oocytes were lysed at low temperatures in a lysis buffer containing protease inhibitors for 30 min. The homogenate was incubated with AX (dissolved in DMSO) or DMSO alone for 2 h. The samples were heated at different temperatures (35, 45, 55, and 65 °C) for 3 min using a PCR instrument, followed by cooling at room temperature for 3 min. The homogenate was transferred to new microtubes, and loading buffer was added. The reaction was immediately terminated by boiling. The results were analyzed by western blotting. To ensure reliable and reproducible results, three biological replicates were performed for the CETSA experiment. Statistical analysis All key experiments were conducted with ≥ 3 independent biological replicates, with exact n-values indicated in each figure panel. Data analysis was performed using GraphPad Prism 9.0 software (GraphPad Software Inc., La Jolla, CA, USA). Shapiro–Wilk normality tests and Brown-Forsythe variance homogeneity tests were performed for all one-way ANOVA analyses. Following the ANOVA, Tukey’s multiple comparisons test was applied. Results are presented as means ± standard deviations (SD). For the pregnancy rate and offspring sex ratio, chi-square tests were conducted in Microsoft Excel software (Microsoft Corporation, Redmond, WA, USA). Specific P values for other data are labeled in the figures. Supplementary Information [164]Additional file 1. Fig. S1-S5.^ (3.3MB, docx) [165]12915_2025_2292_MOESM2_ESM.zip^ (1.5MB, zip) Additional file 2. Table S1. Gene expression of oocytes. Table S2. Primers used in this study. Table S3. MIQE Checklist. Table S4. MIFlowCyt Checklist. [166]12915_2025_2292_MOESM3_ESM.zip^ (8.1MB, zip) Additional file 3. File S1. Original western blot. File S2. Antibody efficacy verification experiment. Acknowledgements