Fetal bovine serum (FBS) was purchased from Hyclone (Logan, United, United States). The Cells-to-cDNA™ II Kit was purchased from Ambion (Austin, TX, United States). All other experimental chemicals and reagents were obtained from Sigma–Aldrich (St. Louis, MO, United States).

Ovaries were collected from a local slaughterhouse in Guangzhou. The mesovarium was removed and the ovaries were rinsed with saline and placed in a physiological saline solution bath at 38 ° C. Porcine cumulus–oocyte complexes (COCs) were aspirated from 3–8 mm ovarian follicles using a 10-ml syringe with an 18-gauge disposable needle. Then, 20–30 COCs were cultured in 150 μl of IVM medium containing TCM-199, 10% (v/v) FBS, 10% (v/v) porcine follicular fluid (PFF), 10 IU/ml follicle-stimulating hormone (FSH), 10 IU/ml luteinizing hormone (LH), and 50 IU/ml penicillin and streptomycin (Cai et al., 2021); covered with mineral oil; and cultured for 42–44 h at 38.5° C in a 5% CO₂ incubator with 95% air. COC expansion was characterized by three grades. Grade 1: no significant morphological change was observed compared with freshly recovered COCs; Grade 2: COCs expanded slightly, but cell–to–cell contact was still relatively tight, with clustered cells still observed; Grade 3: COCs were extensively expanded, and cumulus cells were homogeneously spread (Kobayashi, Yamashita, and Hoshi 1994).

After the COCs were cultured for 42–44 h in vitro with a mature medium, the CCs were removed from the COCs with 0.1% hyaluronidase, which was diluted in TL-PVA-HEPES to harvest naked oocytes. Oocytes that had extruded the first polar body (PB) were cultured in a mature medium supplemented with (QUE-Aged) or without (Aged) QUE for another 24 h at 38.5°C in a 5% CO₂ incubator with 95% air. QUE (Q4951; Sigma, United States) was dissolved in 5 μl dimethyl sulfoxide (DMSO) and diluted to final concentrations of 5, 10, and 20 µM in a culture medium, according to a previously reported study (Cao et al., 2020). DMSO concentration was maintained below 0.05%. Oocytes with fragmented morphology, disjointed ooplasm, and parthenogenetic activation were judged as aged oocytes, based on previous studies (Kikuchi et al., 1995; Gable and Woods 2001). The aging rate was calculated as the percentage of both fragmented and parthenogenetic activated oocytes.

The 2′, 7′-dichlorodihydrofluorescein diacetate (DCFH-DA) fluorescent probe (Beyotime Biotechnology) was diluted with phosphate-buffered saline (PBS) at 1:1,000 to achieve a final concentration of 10 µM. DCFH-DA itself does not emit fluorescence; however, when DCFH is oxidized by intracellular ROS, it becomes fluorescent DCF. Oocytes were incubated with 10 µM DCFH-DA fluorescent probe solution for 40 min at 38 ° C in dark. The oocytes were then rinsed three times with PBS-PVA. Images were captured using a microscope (Eclipse Ti2-E, Nikon, Japan) with the same scanning settings for all groups. The oocyte fluorescence signal was analyzed using ImageJ software (National Institutes of Health, Bethesda, MD, United States).

The ATP assay kit (Beyotime Biotechnology, Shanghai, China) was used to measure ATP levels in porcine oocytes. Briefly, thirty oocytes were lysed with 20 μl of lysis buffer, followed by centrifugation at 400 × g for 5 min at 25 C to collect the supernatant. The ATP level was determined by mixing 20 μl of the supernatant with 100 μl of the luciferase working solution. The fluorescent signal was generated by catalysis of luciferin by firefly luciferase powered by ATP. The intensity of the fluorescent signal was linearly related to ATP concentration, which was measured using a microplate luminometer (BioTek, United States).

The MMP in porcine oocytes was measured using an MMP assay kit (Beyotime Biotechnology, Shanghai, China), according to the manufacturer’s instructions. Briefly, 50 μl JC-1 (200 ×) was mixed with 8 ml DEPC water through a strong vortex, and then 2 ml JC-1 staining buffer (5×) was added to form the JC-1 staining working solution. A total of eight oocytes from each group were added into the working solution stained at 38 ° C for 30 min. Carbonyl cyanide m-chlorophenylhydrazone (CCCP) was used as the positive control. CCCP is a reversible mitochondrial uncoupler. Treatment of porcine oocytes with 10 μM CCCP for 20 min resulted in the loss of mitochondrial membrane potential, and JC-1 stained green. Images were captured using a microscope (Eclipse Ti2-E, Nikon, Japan) with the same scan settings across multiple groups. Oocyte fluorescence signal was analyzed using ImageJ software.

The occurrence of early apoptosis in porcine oocytes was tested using an Annexin V-FITC/PI apoptosis measurement kit (Vazyme, China) according to the manufacturer’s instructions. Briefly, 5 μl of annexin V-FITC and 400 μl of 1× binding buffer were gently mixed to form the working solution. A total of eight oocytes were placed in the annexin V-FITC working solution and incubated at 38° C for 30 min. Annexin V is a Ca²⁺-dependent phospholipid-binding protein that has a high affinity for phosphatidylserine, whereas, in normal cells, phosphatidylserine is only distributed on the inner side of the lipid bilayer of the cell membrane, and in the early stage of apoptosis, phosphatidylserine in the cell membrane is turned from the inside to the outside of the lipid membrane. Therefore, annexin V is recognized as a sensitive indicator of early apoptosis in cells. Images were captured using a microscope (Eclipse Ti2-E, Nikon, Japan) with the same scan settings across multiple groups. The oocyte fluorescence signal was analyzed using ImageJ software.

The oocytes were collected into PCR tubes containing 8 μL cell–to–cDNA™ II lysis buffer. Next, oocyte RNA was extracted and reverse-transcribed to cDNA using the SuperScript ™ II Reverse Transcriptase Kit (Invitrogen, Carlsbad, CA, United States) following the manufacturer’s instructions. Briefly, the following is the 20-µl mixture of reverse transcription system containing 1 µL of DNase I, 1.3 µl of DNase I buffer, 1 µl of EDTA, 1 µl of dNTP, 2 µl of random primer, 4 µl of 5x First-Strand Buffer, 0.5 µl of RNase inhibitor, 2 µl of DTT, and 0.3 µl of FS RT (reverse transcriptase). Next, quantitative PCR was carried out to detect cDNA, and all primers used for mRNAs (Supplementary Table S1) were designed using Oligo 7.0 and compounded by Sangon Biotech (Sangon Biotech, Shanghai, China). The levels of gene transcripts were quantified using 2× Taq Pro Universal SYBR qPCR Master Mix (Vazyme, Nanjing, China), and fluorescence signals were acquired using a fluorescence proportion PCR instrument (Applied Biosystems, QuantStudio 7 Flex, Singapore). The sample was analyzed three times, and the relative gene expression was computed using the 2^−ΔΔCt method with glyceraldehyde-3-phosphate dehydrogenase (GAPDH) as the housekeeping gene.

Oocytes were fixed with 4% PFA at 4 ° C overnight and then rinsed thrice with a blocking solution (PBS containing 3% bovine serum albumin [BSA] and 7.5% glycine). The oocytes were permeabilized with 1% Triton X-100 in PBS for 30 min at 25 ° C and blocked with 1% BSA at 25 ° C. The oocytes were then washed thrice with 1% Triton X-100 and 3% BSA in PBS. Next, oocytes were incubated with an anti-α-tubulin FITC primary antibody at an attenuation of 1:200 and an anti-LC3B antibody at an attenuation of 1:200 overnight at 4 ° C (Supplementary Table S2). After that, the oocytes were washed with PBS-PVA three times and incubated with a secondary antibody (Supplementary Table S2) for 1.5 h at 25 ° C. For spindle structure analysis, oocytes were counter-stained with 10 μg/ml 2-(4-amidinophenyl)-6-indolecarbamidine dihydrochloride for 15 min and then washed three times with PBS-PVA. Finally, fluorescent images were captured using a microscopy system (LAS—Leica TCS-SP5, Germany) using the same scanning settings across different samples, and the fluorescent signal of the oocytes was analyzed using ImageJ software.

Two hundred oocytes were lysed with 120 μl radioimmunoprecipitation assay lysis buffer (Beyotime Biotechnology, Shanghai, China) in an EP tube. The protein concentration was determined using a protein assay kit (Beyotime Biotechnology, Shanghai, China). Next, 30 μl of 5× loading solution per lysed sample was added. A total of 30 μg of protein was transferred to a nitrocellulose membrane using a semi-drying transfer system (Bio-Rad, Hercules, CA, United States). After blocking for 1.5 h with 3% BSA in PBS, the membrane was incubated with a primary antibody (Supplementary Table S2) overnight at 4 °C. Following three 10-min washes in Tris-buffered saline containing 0.1% (v/v) Tween-20, the membrane was incubated with a secondary antibody in Supplementary Table S2 for 1 h at 25 °C. The membrane was washed with Tris-buffered saline containing 0.1% (v/v) Tween-20 and then covered with a 1:1 mixture of enhancer solutions A and B (Millipore, Darmstadt, Germany) for 3 min. Protein bands were visualized using an imaging system (Bio-Rad, Hercules, CA, United States), and the intensity of the bands was analyzed using ImageJ software. Protein levels were standardized by comparison with GAPDH.

All experiments were performed in triplicate. Statistical analysis was performed using SPSS software version 17.0, and quantitative data were presented as the mean ± standard error of the mean. Statistical differences between two groups were analyzed using the t-test, and Duncan’s test was used for analysis of variance in more than two groups. p-values < 0.05 were considered to be statistically significant, and p-values < 0.01 were considered to be extremely statistically significant.

The process of porcine COC in vitro maturation and aging is shown in Figure 1A. The COCs of the germinal vesicle (GV) stage were cultured in a mature medium supplemented with 0, 5, 10, and 20 µM for 42–44 h, and the COC expansion and first polar body extrusion were evaluated. We found that 5 µM QUE did not affect COC expansion, but 10 and 20 µM QUE significantly promoted COC expansion (Figures 1B–D). Furthermore, we found that 5 µM QUE significantly improved the PB1 extrusion rate of porcine oocytes, and 10 µM QUE improved the PB1 extrusion rate to a greater extent; however, 20 µM QUE had a lower effect on PB1 extrusion than 5 µM QUE ( Figures 1E,F ).

An excellent fresh oocyte presents a spherical structure, with a homogeneous and translucent cytoplasm surrounded by a uniform pellucid zone, whereas aged in vitro cultured oocytes tend to have a loosened and fragmented cytoplasm ( Figure 2A). We found that 10 µM QUE significantly reduced oocyte cytoplasmic fragmentation (Figure 2A), thereby reducing the aging rate of in vitro cultured porcine oocytes (Figures 2F,G). Further analysis of the maturation-related promotion factors revealed that the expression levels of bone morphogenetic protein 15 (BMP15), growth differentiation factor 9 (GDF9), Moloney sarcoma oncogene (MOS), and cyclin-dependent kinase 2 (CDK2) were significantly downregulated in aged oocytes compared to those in fresh oocytes (Figures 2B–E). Supplementation with 10 µM QUE restored the expression of these maturation-related promoting factors (Figures 2B–E).

Oxidative stress derived from the accumulation of ROS during oocyte in vitro aging has been considered a key factor affecting oocyte quality. To examine whether QUE can reduce oxidative stress in in vitro aged oocytes, a DCFH-DA fluorescent probe was used to detect intracellular ROS levels in oocytes. The results showed that the fluorescence intensity of DCFH-DA was significantly higher in aged oocytes than in fresh oocytes (Figures 3A,B). QUE supplementation significantly reduced the ROS levels in aged oocytes, as reflected by the 43% decreased fluorescence intensity of DCFH-DA (Figures 3A,B). These results indicate that QUE could alleviate oxidative stress. Molecular analysis of the antioxidant enzymes showed that the expression levels of both CAT and SOD2 decreased significantly in aged oocytes, and QUE supplementation increased the expression of both enzymes (Figures 3C,D). These findings indicate that QUE supplementation could restore the antioxidant capacity of aged oocytes.

ROS accumulation can induce mitochondrial dysfunction. Mitochondrial function can be indicated by the mitochondrial membrane potential (ΔΨm), which is critical for maintaining the physiological function of the respiratory chain to generate ATP. Therefore, we used the JC-1 probe to determine whether the mitochondrial function was impaired in aged oocytes. The significantly reduced red/green ratio compared with that in fresh oocytes (Figures 4A,B) implied impaired mitochondrial function in aged oocytes. QUE supplementation efficiently restored the impaired mitochondrial function in aged oocytes, as reflected by a 25% increased red/green ratio (Figures 4A,B), indicating that impaired mitochondrial function was restored. Further detection of significantly reduced ATP content in aged oocytes confirmed impaired mitochondrial function (Figure 4C), while supplementation with QUE increased 40% ATP production in aged oocytes (Figure 4C), indicating that impaired mitochondrial function was restored.

Oxidative stress can induce early apoptosis during post-ovulatory oocyte aging (Lord and Aitken 2013). Early apoptotic oocytes express phosphatidylserines on the outer leaflet of the plasma membrane, which can be stained with fluorophore-labeled annexin-V (Guemra et al.). We found that fresh oocytes could not be stained by annexin-V (Figure 5A), but aged oocytes were strongly stained by annexin-V (Figure 5A). QUE supplementation reduced 23% annexin-V staining in aged oocytes (Figures 5A,B). We further detected the mRNA expression levels of the pro-apoptotic gene, CASPASE3, and the anti-apoptotic gene, B-cell lymphoma-2 (BCL2), using RT-qPCR. The results demonstrated that CASPASE3 was significantly upregulated in aged oocytes and QUE supplementation reduced CASPASE3 expression in aged oocytes (Figure 5C), which was confirmed by Western blot analysis of the expression of cleaved caspase 3 protein and cleaved caspase 9 protein (Figures 5E,H,I). In contrast, the expression of BCL2 was significantly downregulated in aged oocytes, and QUE supplementation upregulated BCL2 expression in aged oocytes (Figure 5D), which was confirmed by Western blot analysis of BCL2 protein expression (Figures 5E,F). Additionally, Western blotting analysis of the pro-apoptotic factor, Bcl-2-associated X (BAX), showed that it was significantly upregulated in aged oocytes, and QUE supplementation downregulated BAX expression to a certain extent in aged oocytes (Figures 5E,G). These results suggest that QUE supplementation inhibits early apoptosis during oocyte in vitro aging.

A complex interplay between oxidative stress and autophagy has been found in the context of aged oocytes (Wang et al., 2021). Therefore, we investigated whether QUE affects autophagy in porcine oocytes in vitro. Through immunofluorescent staining of the autophagic marker LC3B, we found few positive staining dots in fresh oocytes but universally distributed positive staining dots in aged oocytes (Figure 6A), indicating that autophagy had occurred in aged oocytes. QUE supplementation reduced the quantity of LC3B positive dots (Figure 6A), implying that QUE can attenuate autophagic activity in aged porcine oocytes. This was confirmed by Western blotting analysis of the autophagic activity reporters p62 and LC3B (LC3B-I and LC3B-II), where QUE supplementation attenuated the upregulation of these proteins (Figures 6B–E).

Accumulated ROS levels cause cytoskeletal defects that may lead to spindle abnormalities in aged oocytes (Ren et al., 2012). Therefore, we further examined whether QUE could protect in vitro aged oocytes from spindle organization and chromosome alignment abnormalities. Spindle morphology and chromosome alignment were analyzed (Figure 7A). We found that the percentage of normal spindle/chromosome structures decreased significantly in aged oocytes, and QUE supplementation increased the 19% normal spindle in aged oocytes (Figure 7B). This result suggests that QUE supplementation can reduce spindle/chromosomal abnormalities in in vitro aged porcine oocytes.