Resveratrol attenuates doxorubicin-induced meiotic failure through inhibiting oxidative stress and apoptosis in mouse oocytes

RES alleviates impaired oocyte maturation in DXR-treated oocytes

Oocyte meiotic maturation is a unique asymmetric division, producing a large oocyte and a small first polar body. To assess the toxic effects of DXR on oocyte meiotic maturation, we cultured oocytes with increasing concentrations (100 nM, 200 nM, 400 nM) of DXR and calculated the rate of first polar body extrusion, which is the maker of oocyte meiotic maturation. After 12 h culture, as shown in Figure 1A, most oocytes reached the meiosis II (MII) stage (83.21 ± 1.61 %, n = 124) in control group, however, the maturation rate in DXR treatment group significantly decreased, and some oocytes exhibited bigger first polar body, which indicated abnormal cell division. Besides, we observed that when treated oocytes with a higher concentration of DXR (1 μM), the auto-fluorescent of DXR is detectable, accumulating at the chromosome of oocyte, which indicated that DXR directly bound to oocyte chromosome to exert its toxic effects (Supplementary Figure 1). The maturation rate in 400 nM group significantly reduced to 27.63 ± 2.91% (n = 122) (P < 0.01), and it was 57.84 ± 4.27% (n = 132) (P < 0.05) in 100 nM group and 35.93 ± 1.31% (n = 133) (P < 0.001) in 200 nM group (Figure 1B). So, 200 nM was used as the final DXR treatment group in the subsequent experiments.

Figure 1. Effect of different concentrations of DXR and RES on oocyte meiotic maturation. (A) Microscopy images of oocytes morphologies in control, DXR treatment and RES-supplemented group, oocytes exhibited bigger first polar body (black arrowhead) after DXR treatment. Bar = 200 μm. (B) The oocyte maturation rate was recorded in control and DXR treatment groups. (C) Effect of different concentrations of RES on oocyte maturation in DXR-treated oocytes. The number of oocytes used was shown above the according column. (D) The rate of abnormal meiosis was recorded in control, DXR and RES-supplemented oocytes. All experiments were repeated at least 3 times with more than 30 oocytes examined for each experimental condition. Results were represented as means ± SEMs. * means P < 0.05, **means P < 0.01, *** means P < 0.001.

To examine whether RES could mitigate the impaired oocyte maturation caused by DXR treatment, we cultured oocytes with increasing concentrations of RES (0.5 μM, 1 μM, 5 μM) while treated with 200 nM DXR. As shown in Figure 1, 1 μM RES significantly increased the rate of oocyte maturation (35.93 ± 1.31% vs. 48.20 ± 1.78%, P < 0.01) and reduced the proportion of abnormal meiotic oocytes comparable to that in control group (8.36 ± 1.34%, n = 97 vs. 26.29 ± 1.17%, n = 103, vs. 9.46 ± 1.83%, n = 95, P < 0.05). The results herein suggested that RES partially alleviated the impaired oocyte maturation caused by DXR.

RES restores the disrupted spindle structure and chromosome alignment in DXR-treated oocytes

To investigate the reduced meiotic maturation rate in DXR treatment group, we examined spindle morphology, actin localization and chromosome alignment in both groups. Oocytes were immunolabeled with anti-α-tubulin antibody to label the spindle and counterstained with Hoechst to observe the chromosome. As shown in Figure 2A, 2B, in control group, spindle had the typical barrel shape and chromosome well aligned on the equatorial plate, however, there was no significant difference in actin expression and localization in both groups (Supplementary Figure 2). On the contrary, spindle exhibited irregular configuration, even some aster spindle scattered in DXR treated oocytes, which might be caused by microtubule disassemble. The percentage of oocytes with aberrant spindle morphology (60.67 ± 6.06%, n = 101, vs. 25.67 ± 2.91%, n = 102; P < 0.01) and misaligned chromosomes (71.67 ± 4.05%, vs. 23.00 ± 3.51%, P < 0.01) were significantly increased in DXR treated group. After supplement with RES, the abnormal rates were significantly decreased (37.67 ± 3.76%, n = 100, P < 0.05, spindle; 45.67 ± 2.6%, n = 101, P < 0.05, chromosome), (Figure 2C, 2D). In summary, these results suggested that RES reduced the rate of abnormal meiosis by restoring DXR-induced spindle and chromosome disturbance.

Figure 2. Effect of RES treatment on spindle morphology and chromosome alignment in DXR-treated oocytes. Oocytes were immunolabeled with anti-α-tubulin antibody to visualize spindle and counterstained with Hoechst to observe the chromosomes. (A) Representative images of spindle morphologies and chromosome alignment in control, DXR-treated and RES-supplemented oocytes. Green, α-tubulin; blue, DNA. Bar = 20 μm. (B) Representative images of chromosomes alignment in control, DXR-treated and RES-supplemented oocytes. Blue, DNA. Bar = 5 μm. (C) Rate of aberrant spindle was recorded control, DXR-treated and RES-supplemented oocytes. (D) Rate of misaligned chromosomes was recorded in control, DXR-treated and RES-supplemented oocytes. All experiments were repeated at least 3 times with more than 30 oocytes examined in each experimental condition. Results were represented as means ± SEMs. *means P < 0.05, **means P < 0.01.

RES mitigates oxidative stress in DXR-treated oocytes

Previous studies in different cell lines have demonstrated that ROS is one of the major toxic effects of DXR. To testify whether this was the case in oocytes as well, we detected the ROS level by DCFH-DA fluorescent probe. As shown in Figure 3A, the green signals were considerably stronger in DXR exposure oocytes. Statistical data showed that the fluorescence intensity of ROS was higher in DXR treated group compared with that in control group, but reduced in RES administration group (25 ± 3.61, n = 105 vs. 72 ± 4.16, n = 93 vs 40 ± 3.61, n = 95, P< 0.01) (Figure 3B). We next assessed mRNA expression of antioxidant genes Cat, Sod1, Sod2 and Gpx3. As shown in Figure 3C, the mRNA levels of Cat, Sod1, and Gpx3 were significantly increased while the mRNA levels of Sod2 had no significant difference between the control and DXR treatment groups. Instead, RES reduced ROS levels and elevated the related antioxidant genes expression (Figure 3A–3C). Thus, these results suggested RES prevented oocytes against DXR-induced oxidative stress.

Figure 3. Effect of RES treatment on ROS levels in DXR-treated oocytes. (A) Representative images showed ROS levels in control, DXR-treated and RES-supplemented oocytes, Bar = 100 μm. (B) The fluorescence intensity of ROS was measured in control, DXR-treated and RES-supplemented oocytes. *means P < 0.05, **means P < 0.01. (C) Relative expressions of antioxidant genes Cat, Sod1, Sod2 and Gpx3 mRNA levels in oocytes. β-actin was used as a housekeeping gene. *means P < 0.05, **means P < 0.01. All experiments were performed in triplicates and the data were represented as the means ± SEM.

RES rescues DXR-induced apoptosis by DNA-damage-P63-Caspase3 pathway

Accumulating evidence has demonstrated that excessive ROS leads to DNA damage. Since DXR treatment caused severe oxidative stress in mouse oocytes, we wonder whether DXR treatment would cause DNA damage to oocytes. H2A histone family member X (H2AX) is one of several genes coding for histone H2A. γ-H2AX is widely used as a marker for DNA DSBs damage. We first examined γ-H2AX expression by immunofluorescence staining. As shown in Figure 4A, 4B, DXR treatment caused severe DNA damage to oocytes, whereas RES reduced DNA damage signal to an indistinguishable level compared with that in control group according to quantitative analysis of fluorescence intensity (70.67 ± 2.96, n = 95 vs. 13.00 ± 2.08, n = 102 vs 20.00 ± 2.52, n = 93, P <0.05). To further explore the possible pathway responsible for DXR-induced DNA damage, we then examined the protein expression of P63. Previous studies in ovarian follicles demonstrated that P63 was involved in cisplatin-induced DNA damage in mouse oocytes [19]. As shown in Figure 4C, P63 expression was markedly increased in DXR treated oocytes. Since DNA damage often triggered apoptotic pathways, we next examined active Caspase3 in both groups. As shown in Figure 4C, compared with control group, active Caspase3 expression was significantly increased in DXR-treated group, which indicated DXR treatment induced cell death in oocytes. As expected, RES rescued DXR-induced apoptosis by DNA-damage-P63-Caspase3 pathway (Figure 4A–4C).

Figure 4. RES rescued DXR-induced apoptosis through DNA-damage-P63-Caspase3 pathway in mouse oocytes. (A) Representative immunofluorescence images showing the expression of γ-H2AX in mouse oocytes. Green, γ-H2AX, Blue, DNA, Bar = 20 μm. (B) The relative immunofluorescence intensity of γ-H2AX was measured in control, DXR-treated and RES-supplemented oocytes. Experiments were repeated at least 3 times with more than 30 oocytes examined for each group. Data were presented as means ± S.E.M of three independent experiments. **means P < 0.01, *** means P < 0.001. (C) Protein levels of γ-H2AX, P63 and Active-Caspase3 were examined by Western blotting in control, DXR-treated and RES-supplemented oocytes. GAPDH was used as a loading control. The clean backgrounds for the active-Caspase-3, γ-H2AX and GAPDH is due to the exposure.

Animals

CD-1 female mice (21-23 days old) were purchased from the Laboratory Animal Center of the Institute of Genetics and Developmental Biology (Beijing, China). All procedures were approved by the Animal Care and Use Committee of China Agricultural University and were performed following the Animal Research Institute Committee guidelines. Mice were raised in a temperature and light-controlled room, water and food were feed ad libitum.

Oocyte collection and culture

Immature female CD-1 mice were killed by cervical dislocation to collect ovaries. Germinal vesicle intact oocytes were picked with a pipetted tube, each group contains at least 30 oocytes, washed several times in M2 medium, then cultured under mineral oil in M16 medium h at 37°C in 5% CO₂ atmosphere.

Reagents and antibodies

Doxorubicin hydrochloride and resveratrol were purchased from J&K Chemical Ltd. (Shanghai, China). Throughout this paper, the abbreviation DXR and RES were used to refer to doxorubicin hydrochloride and resveratrol respectively. DXR and RES were dissolved in DMSO, then diluted to the corresponding working concentrations, with the final concentration of DMSO not more than 0.1% of the culture medium. Concentrations range of doxorubicin in the study were determined from the relevant plasma level (50-200 nM) in cancer patients [35]. Finally, 200 nM was chosen as the final dosage in the following experiments.

Mouse anti-α-tubulin-FITC antibody (#F2168) was purchased Sigma-Aldrich (MA, USA); rabbit polyclonal anti-γ-H2AX antibody (#NB100-2280) was purchased from Novus (CO, USA), rabbit polyclonal anti-P63 antibody (#BS1279) was purchased from Bioworld (MO, USA), Rabbit polyclonal anti-active-Caspase3 antibody was purchased from Beyotime, (Shanghai, China). Alexa Fluor 488 goat anti-rabbit antibody, Alexa Fluor 594 goat anti-mouse antibody were from Invitrogen (CA, USA).

Immunofluorescence

Oocytes were fixed 30 min in 4% paraformaldehyde, then permeabilized with 0.5% Triton X-100 for 20 min, and blocked 1 h in blocking buffer (PBS containing 1% BSA) at room temperature, oocytes were then probed with different primary antibodies (1:400 for anti-α-tubulin-FITC; 5 μg/ml Phalloidin-TRITC; 1:200 for γ-H2AX) at 4°C overnight, after extensive washing, oocytes were then stained with the corresponding secondary antibodies. Finally, oocytes were stained with Hoechst for 10 min to visualize the DNA, then transfer oocytes to a glass slide. The slides were examined and photographed using Nikon Eclipse 80i digital fluorescence microscopy.

The fluorescence intensity was assessed using Image J software (NIH). Samples from different groups were mounted on the same glass slide and scanned with the same parameters of fluorescence microscopy.

Analysis of intracellular ROS generation

Intracellular ROS levels were measured by detecting ROS as green fluorescent signals of DCFH diacetate (DCHFDA; Beyotime Biotechnology, Shanghai, China). After 9 h culture, oocytes from both groups were incubated at 37 °C for 30 min in D-PBS that contained 10 μM DCHFDA. Then, the oocytes were washed 3 times with D-PBS and placed on glass slides and covered with coverslips. After that, each sample was observed immediately under a Nikon fluorescent microscope with the same scan settings.

Western blotting analysis

At least 100 oocytes were collected in each group, dissolved in SDS sample buffer then boiled at 100°C for 10 min. Protein lysates were separated by SDS-PAGE and electrophoretically transferred to PVDF membranes (Millipore, MA, USA). The membranes were blocked in Tris-buffered saline containing 0.1% Tween 20 and 5% no fat dry milk for 1 hour, then probed with primary antibodies, overnight at 4°C. After several times washing, the membranes were incubated with a horseradish peroxidase-linked secondary antibody. Finally, bands on the membranes were detected using the Super-Signal West Pico chemiluminescent detection system (Prod 34080, Thermo, USA).

RNA extraction and quantitative real-time RT-PCR

RNA was extracted from at least 150 oocytes for different groups using Trizol Reagent according to the manufacturer’s protocol. Quantitative PCR was performed on a Light Cycler instrument (Roche, Mannheim, Germany). Gene expression changes were analyzed by the 2^-ΔΔCt method and normalized with β-actin. The primers used for testing genes are listed in Supplementary Table 1.

Statistical analysis

Each experiment repeated at least 3 times with independent samples. Data are presented as means ± SEMs and analyzed by one-way ANOVA. P value < 0.05 was considered significant.