Cadmium Exposure of Female Mice Impairs the Meiotic Maturation of Oocytes and Subsequent Embryonic Development

Cadmium Exposure of Female Mice Impairs the Meiotic Maturation of Oocytes and Subsequent... Abstract Cadmium is one major pollutant that is highly toxic to animals and humans. The mechanism of cadmium toxicity on the female reproductive system, particularly oocyte maturation and fertility, remains to be clarified. In this study, we used a mouse model to investigate the effects of cadmium in the drinking water on the meiotic maturation of oocytes and subsequent embryonic development, and the underlying mechanisms associated with the impairment of oocyte maturation such as mitochondrial distribution and histone modifications. Our results show that cadmium exposure decreased the number of ovulated oocytes and impaired oocyte meiotic maturation rate both in vivo and in vitro. The embryonic development after fertilization was also impaired even when the potential hazards of cadmium on the spermatozoa or the genital tract have been excluded by fertilization and embryonic development in culture. Cadmium exposure disrupted meiotic spindle morphology and actin filament, which are responsible for successful chromosome segregation and the polar body extrusion during oocyte maturation and fertilization. ATP contents, which are required for proper meiotic spindle assembly in the oocyte, were decreased, consistent with altered mitochondrial distribution after cadmium exposure. Finally, cadmium exposure affected the levels of H3K9me2 and H4K12ac in the oocyte, which are closely associated with the acquisition of oocyte developmental competence and subsequent embryonic development. In conclusion, cadmium exposure in female mice impaired meiotic maturation of oocytes and subsequent embryonic development by affecting the cytoskeletal organization, mitochondrial function, and histone modifications. cadmium, mouse oocyte, maturation, embryos Cadmium is widely used in the steel industry and manufacture of plastics or batteries, and is easily released into the environment via wastewater, contaminated fertilizers and atmospheric air. As a common pollutant, cadmium is readily accumulated in crops, aquatic organisms and other food-chains (Fan et al., 2017). Human exposure occurs mainly from consumption of contaminated food, active or passive inhalation of tobacco smoke and air inhalation in a metal industry area. Drinking water can be contaminated with cadmium as an impurity in the zinc of galvanized pipes and solder in fittings, water heaters, water coolers or taps. Cadmium is classified as a human carcinogen and is known to have numerous adverse effects on health in both experimental animals and humans. Cadmium-induced toxicity is very well known across the world. Long-term exposure to cadmium through air, water, soil, and food leads to systemic organ toxicity, in the hepatic, renal, skeletal, reproductive, cardiovascular, nervous, and respiratory systems (Rafati Rahimzadeh et al., 2017). The molecular mechanisms of cadmium toxicity are multiple and complex. We have previously reported that disruption of gap junction intercellular communication and activation of mitogen-activated protein kinase pathways are possible mechanisms of cadmium-induced hepatotoxicity (Zou et al., 2015). Furthermore, cadmium disrupts the F-actin cytoskeleton in rat renal mesangial cells, which is mediated by gelsolin translocation, programmed cell death, and F-actin depolymerization through formation of a CAP1-cofilin-F-actin complex (Apostolova et al., 2006; Liu and Templeton, 2013, 2017b). It has been demonstrated that mitochondria copy numbers and mitochondrial function are involved in acute or chronic renal cadmium toxicity (Nair et al., 2015). In addition, cadmium-induced cytotoxicity in primary rat proximal tubular cells can be attributed to the inhibition of the cytosolic Ca2+ dependent autophagosome-lysosome fusion (Liu et al., 2017a). We also found that cadmium-induced apoptosis and autophagy are involved in the cadmium neurotoxicity (Rani et al., 2014; Wang et al., 2015a; Yuan et al., 2016). The extensive deleterious effects of cadmium on the reproductive system and developing embryos have been described, especially in the male (de Angelis et al., 2017; Thompson and Bannigan, 2008; Zhao et al., 2017). Cadmium can disrupt the blood-testis barrier in the testis even at low concentrations by damaging the testis vasculature and exerting the cytotoxicity on Sertoli and Leydig cells, resulting in a decrease in sperm quality, fertilization capacity and subsequent early embryonic development (de Angelis et al., 2017; Zhao et al., 2017). It has been demonstrated that cadmium levels in the ovary increase with age, associated with a failure in ovulation, retardation of trophoblastic outgrowth and embryonic development, placental necrosis, implantation delay, and early pregnancy loss (Thompson and Bannigan, 2008). Furthermore, cadmium exposure inhibits follicular development and increases the number of atretic follicles (Wang et al., 2015b; Zhang et al., 2017). However, it is not well understood how cadmium affects oocyte quality, particularly on meiotic maturation, which is critical for fertilization and embryo development. In most previous studies, the toxicity of cadmium was tested by directly adding it to isolated organs or cells in culture. However, any organ like the ovary has a microenvironment in vivo, through which its viability and functions are regulated. Accordingly, in this study, we administered cadmium in the drinking water to female mice for 35 days and tested its toxic effects on the oocytes both in vivo and in vitro. The environmental concentration of cadmium can reach 75 mg/l, depending on the source of cadmium and the degree of industrialization and pollution (Fan et al., 2017; Li et al., 2015). The dietary intake of cadmium via the food chain in mining areas of China is estimated to be 0.179 mg/day/person on average, which far exceeds the provisional tolerable monthly intake for cadmium of 25 μg/kg body weight specified by the World Health Organization (Du et al., 2013). Moreover, the biological half-life of cadmium is 10–35 years, resulting in long-term accumulation of cadmium in organs. It has been confirmed that long-term exposure to cadmium contributes to the development of lung cancer. The mean blood level of cadmium was 24 μg/l for the local resident in the cadmium high exposure area, which had significantly positive correlations with the risk of all cancer and mortality from all cancers (Wang et al., 2011). In the current study, we chose the cadmium dose 32 mg/l according to our preliminary experiment and the previous report (Yamanobe et al., 2015). The dosage used is in the range found in the environment. The average volume of drinking water was estimated to be 4 ml/day/mouse, so the intake of cadmium was 0.128 mg/day/mouse, which is comparable to the human intake in the mining area of China. Throughout the study, the average body weight of a mouse was estimated to be 35 g, which was used to calculate the cadmium dosage of 3.657 mg/kg/day. This cadmium dose administered was 6% of the LD50 for mice, which was 60 mg/kg/day (Yamanobe et al., 2015; Zhai et al., 2013). In this study, we used the mouse model described earlier to investigate the effects of cadmium on ovulation, meiotic progression in the oocytes during in vivo and in vitro maturation (IVM), and subsequent early embryonic development after in vitro fertilization (IVF). We examined spindle conformation and actin filament localization in the matured oocyte. We also investigated possible mechanisms of cadmium toxicity on mitochondrial distribution, ATP levels, and histone modifications in the oocyte. MATERIALS AND METHODS Materials and chemicals Chemicals, M16 media, bovine serum albumin (BSA), and mouse antiα-tubulin-FITC were purchased from Sigma-Aldrich (St Louis, Missouri). Rabbit monoclonal antiH3K9me2 antibody and polyclonal antiH4K12ac antibody were purchased from AbCam (Cambridge, UK). Donkey antirabbit IgG-Alexa Fluor-488, Rhodamine-phalloidin, Mitotracker Green-FM, Antifade Mountant with DAPI, and MEM-alpha medium were purchased from Thermo Fisher (Waltham, Massachusetts). Cadmium chloride 2.5-hydrate (purity 99.99%) was purchased from Suyi Chemical Company (Shanghai, China). Equine gonadotropin (eCG) and human chorionic gonadotropin (hCG) were obtained from Ningbo Second Hormone Factory (Ningbo, China). KSOMaa medium was purchased from Millipore (Burlington, Massachusetts). Enhanced ATP Assay Kit was purchased from Beyotime Biotechnology Company (Nantong, China). Ethics statement This study was conducted in accordance with the recommendations of the Guide for the Care and Use of Laboratory Animals of the National Research Council. All procedures executed in this study were reviewed and approved by the Animal Care and Use Committee of Yangzhou University (approval ID: SYXK [Su] 2007-0005). Healthy ICR mice (Experimental Animal Center of Yangzhou University), 6–12 weeks-of-age, were used for all experiments. The mice were housed in an air-conditioned animal house (26°C ± 2°C) and exposed to 10–12 h of light per day. Animals were maintained on a standard diet and had free access to water. Animal treatment The mice were randomly divided into 2 groups in each experiment. Every mouse was only used for 1 experiment. The cadmium treatment was carried out according to our preliminary experiment and the previous report. The cadmium dose administered in this study, 32 mg/l, is 6% of the LD50 in mice (Yamanobe et al., 2015). There was no lethality in all cadmium-treated mice throughout the study. For the experimental group, female mice were given ad libitum Milli-Q water containing 32 mg/l cadmium for 35 days. For the control group, Mill-Q water was given ad libitum. Collection of in vivo matured oocytes by superovulation Female mice after cadmium treatment were injected intraperitoneally each with 10 IU eCG, followed by injection with 10 IU hCG 45-47 h later, and sacrificed 14–15 h later. The oocytes enclosed in cumulus cells, named cumulus-oocyte complexes (COCs), were flushed out from the oviduct. After being treated with 300 μg/ml hyaluronidase for 5 min at 37°C and being denuded of cumulus cells by repeated pipetting through a narrow glass needle in the M2 medium, the number of oocyte per mouse was counted. The stage of each oocyte was determined under stereomicroscope, and the number of oocyte in each stage from different mice was pooled in 1 experiment. Collection of germinal vesicle stage oocytes and IVM Female mice after cadmium treatment were injected intraperitoneally each with 10 IU eCG and were sacrificed 48 h later. COCs were isolated by puncturing large preantral and antral follicles with a pair of 26-gauge needles in the M2 medium. For the denuded-oocytes (DOs), COCs were treated with 300 μg/ml hyaluronidase for 5 min at 37°C and oocytes were denuded of cumulus cells by repeated pipetting through a narrow glass needle in the M2 medium. Ten COCs or DOs in a group were cultured in a prewarmed 20 μl droplet of the M16 medium under mineral oil at 37°C with 5% CO2 in a humidified atmosphere. After 18 h, COCs were denuded of cumulus cells, the stage of each oocyte was determined under stereomicroscope. The oocytes in each stage from different mice were pooled in each experiment. IVF and embryonic development Spermatozoa were collected from the caudal epididymis of males that were not treated with cadmium at 10–12 weeks of age and capacitated in the MEM-alpha medium supplemented with 0.9% BSA under mineral oil at 37°C for 60 min. The ovulated oocytes (in vivo matured) were transferred into an IVF medium (MEM-alpha supplemented with 0.4% BSA) containing spermatozoa under oil and further incubated for 5 h at 37°C with 5% CO2 in a humidified atmosphere. After washing in the modified KSOMaa medium, the fertilized eggs were cultured in the fresh modified KSOMaa medium for up to 5 days. For the in vivo fertilized oocytes, female mice that were injected with 10 IU eCG and 45–47 h later with 10 IU hCG were left with male mice overnight. The female mice found with vaginal plugs next morning were considered to have successfully copulated. After 0.5 day, the in vivo fertilized eggs were flushed out from the oviduct, and cultured as above for up to 5 days. The stage of each embryo was determined under stereomicroscope every day, and the embryos in each stage from different experiments were pooled. Confocal microscopy analysis of meiotic spindles and actin filaments Immunofluorescence staining was done as previously described in Zhu et al. (2017). Oocytes were fixed with 4% paraformaldehyde in phosphate-buffered saline (PBS) for 30 min at room temperature. Then, oocytes were blocked in PBS containing BSA (3%) and Triton-X-100 (0.1%) (named blocking solution) for 1 h at room temperature and incubated with mouse antiα-tubulin-FITC (1:500) and Rhodamine-phalloidin (1:200) overnight at 4°C. After 3 washes in PBS, oocytes were transferred into Antifade Mountant with DAPI on glass slides. Fluorescence signals were examined under the confocal laser-scanning microscope (Leica TCS SP8). A rugby ball shape of α-tubulin was defined as the normal meiotic spindle. A region of interest (ROI) of actin was defined as a dotted line to cross the center of oocyte and midzone of spindle. Higher fluorescence intensity of ROI in the oolemma than that in the ooplasm was defined as a normal actin distribution. The proportion of abnormal oocytes was counted in each experiment. Measurement of ATP contents ATP contents in oocytes were determined by using Enhanced ATP Assay Kit according to the manufacturer’s protocol. 40 oocytes in a group were added to ice-cold Cell ATP-Releasing Reagent, following the addition of ATP Assay Reagent. The bioluminescence of each sample was measured using a high-sensitivity luminometer (BioTek). Examination of mitochondrial distribution Oocytes were fixed in 4% paraformaldehyde in PBS for 25 min. After washing, oocytes were incubated in 100 nM Mitotracker Green-FM in the M2 medium for 35 min in dark. After washing, oocytes were transferred into Antifade Mountant with DAPI on glass slides. Fluorescence signals were examined under the confocal laser-scanning microscope (Leica TCS SP8). Mitochondrial distribution patterns were classified according to the practices used in previous reports (Moawad et al., 2014; Wang et al., 2009). The proportion of abnormal oocytes was calculated in each experiment. Immunofluorescence staining of H3K9me2 and H4K12ac Oocytes were fixed as described earlier, and incubated with rabbit monoclonal antiH3K9me2 antibody (1:200) or rabbit polyclonal antiH4K12ac antibody (1:200) overnight at 4°C. After washing, oocytes were incubated with donkey-antirabbit IgG- Alexa Fluor-488 (1:500) at room temperature for 1 h. After washing, oocytes were transferred into Antifade Mountant with DAPI on glass slides and were examined under the confocal laser-scanning microscope. Oocytes in the experimental and control groups were mounted on the same glass slide, and the constant scanning setting was used for sample scanning. Leica microscope software (Leica LAS AF) was used to assess oocyte fluorescence intensity levels. A ROI was defined as encompassing all the chromosomes in each oocyte, and the average fluorescence intensity for this ROI was determined. The relative fluorescence intensity of H3K9me2 and H4K12ac in each oocyte was normalized against the fluorescence intensity of DAPI. Statistical analysis At least 20 oocytes were examined in 1 experiment. All experiments were repeated at least 3 times. Data are presented as mean ± SEM. Statistical differences among pooled results were analyzed by 2-way ANOVA using SPSS software. Statistical differences between groups from separate experiments were analyzed by t test using SPSS software. A p < .05 was considered statistically significant. RESULTS Cadmium Exposure Reduces the Number of Ovulated Oocytes We first tested the effects of cadmium in drinking water on ovulation. As shown in Figure 1, the number of ovulated oocytes from the cadmium-treated group (Cd) was significantly less than that from the control group (Con) (42.77 ± 2.44 vs 52.43 ± 3.06/female; p < .05). These results show that cadmium exposure reduces the number of ovulated oocytes after superovulation. Figure 1. View largeDownload slide Cadmium exposure (Cd) reduces the number of ovulated oocytes. Data are presented as means ± SEM. The total number of females examined is given in parentheses at the bottom of each column. *Statistical differences between the oocytes from 2 groups by t test at p < .05. Figure 1. View largeDownload slide Cadmium exposure (Cd) reduces the number of ovulated oocytes. Data are presented as means ± SEM. The total number of females examined is given in parentheses at the bottom of each column. *Statistical differences between the oocytes from 2 groups by t test at p < .05. Cadmium Exposure Impairs the Oocyte Maturation In Vivo We next examined the morphology of ovulated oocytes under the stereo microscope. As shown in Figure 2A, most oocytes were at the MI-stage without the first polar body in the Cd group (50.58% ± 2.60%); compared with the control group (24.29% ± 3.55%) with a significant difference (p < .01). Conversely, the percentage of MII-oocytes that had extruded the first polar body in the Cd group was significantly smaller than that in the control group (29.65% ± 3.17% vs 72.60% ± 3.63%; p < .01). These results indicate that cadmium exposure impaired the oocyte maturation in vivo. Figure 2. View largeDownload slide Cadmium exposure impairs the oocyte maturation both in vivo and in vitro. A, In vivo maturation rate of oocytes were significantly reduced after female mice were exposed to cadmium. In vitro maturation rate of oocytes with (B) or without (C) cumulus cells was significantly reduced after female mice were exposed to cadmium prior to COCs collection. For the DOs, COCs were denuded of cumulus cells before in vitro culture. In vivo maturation and in vitro maturation were repeated 10 and 5 times, respectively. *,**Statistical differences of each stage between the oocytes from 2 groups by t test at p < .05 and .01, respectively. Figure 2. View largeDownload slide Cadmium exposure impairs the oocyte maturation both in vivo and in vitro. A, In vivo maturation rate of oocytes were significantly reduced after female mice were exposed to cadmium. In vitro maturation rate of oocytes with (B) or without (C) cumulus cells was significantly reduced after female mice were exposed to cadmium prior to COCs collection. For the DOs, COCs were denuded of cumulus cells before in vitro culture. In vivo maturation and in vitro maturation were repeated 10 and 5 times, respectively. *,**Statistical differences of each stage between the oocytes from 2 groups by t test at p < .05 and .01, respectively. Cadmium Exposure Impairs the Oocyte Maturation In Vitro After Separating From the Follicular Microenvironment Oocyte competence for meiotic progression depends on the ovarian follicle microenvironment during the oocyte growth phase, whereas the continual presence of neighboring cumulus cells facilitate oocyte maturation and competence for embryonic development (Eppig, 2001; Villemure et al., 2007; Xu et al., 2014). In the above study, both the oocyte and its surrounding somatic cells were exposed to cadmium throughout the growth and maturation of oocytes, so we asked whether the systematic presence of cadmium directly affected the oocyte meiotic maturation or the ovarian follicle microenvironment which in turn impaired oocyte maturation. We isolated the oocytes from follicular environment and matured them in vitro. Furthermore, a group of oocytes were left surrounded by cumulus cells (COC group), while another group of oocytes was denuded of cumulus cells (DO group) before maturation in vitro. As shown in Figures 2B and 2C, the percentages of MII-oocytes were significantly smaller (p < .05) in both COC and DO groups from the Cd-treated females (85.64% ± 3.00% and 46.11% ± 5.84%, respectively), compared with the control females (94.70% ± 1.25% and 70.62% ± 4.97%, respectively). Conversely, percentages of MI-oocytes were significantly larger in the Cd group compared with the control group. However, the percentages of MII-oocytes in the Cd IVM group were significantly higher than those in Cd in vivo maturation group (Figure 2, COC 85.64% ± 3.00% and DO 46.11% ± 5.84% vs 29.65% ± 3.17%). These results indicate that meiotic maturation of oocyte was partially improved when the oocyte was separated from the ovarian follicle microenvironment. We conclude that cadmium exposure affected the ovarian follicle microenvironment which then continually impaired the maturation of oocyte in vivo. Cadmium Exposure Impairs Embryonic Development We next compared the development of eggs from both cadmium-treated female mice and the control female mice. The results are summarized in Figure 3B. In the Cd group, the rate of the first cleavage (2-cell) was comparable with that of the control group. However, the rate of 4-cell stage embryo development after 2 days in culture (57.9%, n = 242) was significantly lower and that of morula after 3 days culture (46.7%) was also lower, although without statistical difference compared with the control group (82.2% and 68.0%, respectively, n = 309). After 4 and 5 days in culture, 42.2% and 48.8% of fertilized eggs reached the blastocyst-stage in the Cd group, while 66.0% and 75.4% did so in the control group (p < .05 and .01, respectively). When the proportions of embryos at various stages were statistically analyzed, significant differences between the 2 groups were found (2-way ANOVA, p < .001). These results indicate that cadmium treatment of females made their eggs incompetent for embryonic development even when the eggs were separated from the oviduct and the uterus which had been exposed to cadmium. Figure 3. View largeDownload slide Cadmium exposure impairs the embryonic development of naturally fertilized eggs (in vivo) and in vitro fertilized eggs (IVF). A, Examples of embryos at 1–5 days in culture. Bar: 50 μm. “Others” includes dead or fragmented embryos (arrow). Each column indicates the percentages of embryos at different stages at 1–5 days in culture from in vivo (B) and in vitro fertilized eggs (C). The total number of embryos examined is given in parentheses at the top of each graph. The data were pooled from 4 (in vivo) and 6 (IVF) experiments. Significant differences between the 2 groups by 2-way ANOVA. *,**Statistical differences of each stage between the 2 groups at each day by t test at p < .05 and .01, respectively. Figure 3. View largeDownload slide Cadmium exposure impairs the embryonic development of naturally fertilized eggs (in vivo) and in vitro fertilized eggs (IVF). A, Examples of embryos at 1–5 days in culture. Bar: 50 μm. “Others” includes dead or fragmented embryos (arrow). Each column indicates the percentages of embryos at different stages at 1–5 days in culture from in vivo (B) and in vitro fertilized eggs (C). The total number of embryos examined is given in parentheses at the top of each graph. The data were pooled from 4 (in vivo) and 6 (IVF) experiments. Significant differences between the 2 groups by 2-way ANOVA. *,**Statistical differences of each stage between the 2 groups at each day by t test at p < .05 and .01, respectively. Considering the potential effects of cadmium on spermatozoa motility and fertilization through contact with the oviduct and the uterus (Zhao et al., 2017), we used IVF. As shown in Figure 3C, the percentage of blastocyst development was significantly smaller in the Cd group than that of the control group (33.68% vs 48.41%, p < .05) after 5 days in culture. In addition, more embryos were found dead or fragmented (others in Figure 3) in the Cd group. When the proportions of embryos at various stages after IVF were statistically analyzed, significant differences between the 2 groups were also found (2-way ANOVA, p < .001). These results demonstrate that the embryonic development was impaired when oocytes were collected from cadmium-treated female mice. Cadmium Exposure Disrupts Meiotic Spindle Morphology and Actin Accumulation and Distribution Next, we examined the cytoskeletal integrity of ovulated oocytes, which indicate quality of the oocyte. As shown in Figure 4A, the meiotic spindle labeled with α-tubulin antibody in the control group typically had a rugby ball shape, but some had a bulky shape with at least 1 pole wider than the midzone. The proportion of oocytes with aberrant spindles such as these in the Cd group was significantly higher than in the control group (Figure 4B; 50.00% ± 10.94% vs 12.87% ± 3.44%, p < .01). Figure 4. View largeDownload slide Cadmium exposure disrupts meiotic spindle morphology and actin accumulation and distribution. A, The MII-oocyte was stained with antiα-tubulin-FITC (green), rhodamine-phalloidin (actin, red) and DAPI (blue). In the control group, the typical meiotic spindle (α-tubulin) was seen in a rugby ball shape, and the actin filaments (actin) were localized along the oolemma and concentrated close to the MII-spindle as well as around the first polar body. Fluorescence intensity of actin along in the dotted line shows that actin accumulation in the oolemma was significantly higher than that in the ooplasm. However, in the Cd group, the spindle was bulky (at least 1 pole was wider than the midzone), and the actin accumulation in the oolemma was absent even when the fluorescence signals actin in the ooplasm was overexposed. Bar: 20 μm. (B, C) The rate of aberrant spindle and lack of actin localization in the Cd group was significantly higher than in the control group. The total number of oocytes examined, from 4 experiments, is given in parentheses at the right of the graph. **Statistical differences between the 2 groups by t test at p < .01. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.) Figure 4. View largeDownload slide Cadmium exposure disrupts meiotic spindle morphology and actin accumulation and distribution. A, The MII-oocyte was stained with antiα-tubulin-FITC (green), rhodamine-phalloidin (actin, red) and DAPI (blue). In the control group, the typical meiotic spindle (α-tubulin) was seen in a rugby ball shape, and the actin filaments (actin) were localized along the oolemma and concentrated close to the MII-spindle as well as around the first polar body. Fluorescence intensity of actin along in the dotted line shows that actin accumulation in the oolemma was significantly higher than that in the ooplasm. However, in the Cd group, the spindle was bulky (at least 1 pole was wider than the midzone), and the actin accumulation in the oolemma was absent even when the fluorescence signals actin in the ooplasm was overexposed. Bar: 20 μm. (B, C) The rate of aberrant spindle and lack of actin localization in the Cd group was significantly higher than in the control group. The total number of oocytes examined, from 4 experiments, is given in parentheses at the right of the graph. **Statistical differences between the 2 groups by t test at p < .01. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.) We also examined the actin filaments, which play a role in the polar body extrusion. In the control group, actin filaments were localized along the oolemma and concentrated over the MII-spindle in addition to the first polar body; however, the actin filaments were no longer seen along the oolemma in the Cd group even when the fluorescence signals were enhanced (Figure 4A). The proportion of oocytes that lacked actin localization on the oolemma in the Cd group was significantly higher than that in the control group (Figure 4C; 37.04% ± 5.63% vs 6.93 ± 1.08%, p < .01). We did not find any difference in chromosome condensation or alignment by labeling with DAPI within the oocytes in the 2 groups. These results indicate that cadmium exposure disrupted the meiotic spindle assembly and actin localization, resulting in the loss of the cortical polarity. Cadmium Exposure Reduces ATP Contents by Disrupting the Mitochondrial Distribution in the Oocyte ATP is mainly synthesized by mitochondria in the oocyte and plays a critical role in proper spindle assembly and fertilization (Van Blerkom et al., 1995; Xu et al., 2014; Zhang et al., 2006). We measured ATP contents in the MII-oocytes from the cadmium-treated female mice. The results showed that the ATP contents in the Cd group were significantly lower than those in the control group (2.03 ± 0.11 vs. 3.00 ± 0.10 μmol/l, p < .01) (Figure 5). It is also known that mitochondrial distribution in the MII-oocytes is associated with the competence for embryonic development (Bavister and Squirrell, 2000; Wilding et al., 2001). Accordingly, we examined the mitochondrial distribution with Mitotracker Green-FM staining, and classified the results into 3 different patterns: accumulation around the MII-spindle (Figure 6Ai), homogeneous distribution over the ooplasm (Figure 6Aii), and peripheral accumulation with overall weak fluorescence signals (Figure 6Aiii). As shown in Figure 6B, the majority of oocytes in the control group showed accumulation around the MII-spindle (pattern i) (76.92 ± 3.91%); however, the proportion of oocytes in this pattern was significantly lower in the Cd group (56.44% ± 4.96%, p < .01). Instead, a considerable number of oocytes in the Cd group showed peripherally accumulated mitochondria with overall weak fluorescence signals (pattern iii), and these were found at a significantly larger percentage than in the control group (33.66% ± 4.73% vs 8.55% ± 2.60%, p < .01). These results indicate that cadmium exposure disrupts the mitochondrial distribution, which may be responsible for lower ATP contents in the MII-oocytes of the Cd group. Figure 5. View largeDownload slide Cadmium exposure reduces ATP contents in MII-oocytes. This experiment was repeated 9 times. **Statistical differences between the 2 groups by t test at p < .01. Figure 5. View largeDownload slide Cadmium exposure reduces ATP contents in MII-oocytes. This experiment was repeated 9 times. **Statistical differences between the 2 groups by t test at p < .01. Figure 6. View largeDownload slide Cadmium exposure disrupts the mitochondrial distribution in MII-oocytes. A, The MII-oocytes were labeled with Mitotracker Green-FM to visualize mitochondrial localization. Mitochondrial distribution was classified using 3 different patterns: accumulation around the MII-spindle (i), homogeneous distribution over the ooplasm (ii), and peripheral concentration with overall weak signals (iii). Bar: 20 μm. (B) Quantification of oocytes in each mitochondrial distribution pattern. The total number of oocytes examined is given in parentheses at the right of the graph, from 3 experiments. ** Statistical differences between the 2 groups by t test at p < .01. Figure 6. View largeDownload slide Cadmium exposure disrupts the mitochondrial distribution in MII-oocytes. A, The MII-oocytes were labeled with Mitotracker Green-FM to visualize mitochondrial localization. Mitochondrial distribution was classified using 3 different patterns: accumulation around the MII-spindle (i), homogeneous distribution over the ooplasm (ii), and peripheral concentration with overall weak signals (iii). Bar: 20 μm. (B) Quantification of oocytes in each mitochondrial distribution pattern. The total number of oocytes examined is given in parentheses at the right of the graph, from 3 experiments. ** Statistical differences between the 2 groups by t test at p < .01. Cadmium Exposure Increases H3K9me2 and H4K12ac Levels in MII-Oocytes Histone modifications, particularly dimethylation of H3K9 (H3K9me2) and deacetylation of H4K12 (H4K12ac), during meiotic maturation of oocytes are essential for the developmental competence of oocytes (Racedo et al., 2009; van den Berg et al., 2011). As shown in Figure 7, the relative fluorescence intensity of H3K9me2 in the Cd group was significantly increased, compared with that of the control group (0.72 ± 0.02 vs 0.54 ± 0.03, p < .01). The H4K12ac intensity in the Cd group was also significantly increased, compared with that of the control group (Figure 8, 0.65 ± 0.03 vs 0.54 ± 0.02, p < .01). These results indicate that cadmium exposure increases both H3K9me2 and H4K12ac levels in MII-oocytes. Figure 7. View largeDownload slide Cadmium exposure increases H3K9me2 levels in MII-oocytes. A, MII-oocytes were stained with antiH3K9me2 antibody (green) and DAPI (blue). A constant setting of confocal laser-scanning microscope was used for all samples. The average fluorescence intensity for a ROI (white box) surrounding MII-chromatids was measured. Bar: 10μm. B, The relative fluorescence intensity of H3K9me2 in the MII-oocyte was normalized against the fluorescence intensity of DAPI. The total number of oocytes examined is given in parentheses at the bottom of each column. **Statistical differences between the 2 groups by t test at p < .01. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.) Figure 7. View largeDownload slide Cadmium exposure increases H3K9me2 levels in MII-oocytes. A, MII-oocytes were stained with antiH3K9me2 antibody (green) and DAPI (blue). A constant setting of confocal laser-scanning microscope was used for all samples. The average fluorescence intensity for a ROI (white box) surrounding MII-chromatids was measured. Bar: 10μm. B, The relative fluorescence intensity of H3K9me2 in the MII-oocyte was normalized against the fluorescence intensity of DAPI. The total number of oocytes examined is given in parentheses at the bottom of each column. **Statistical differences between the 2 groups by t test at p < .01. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.) Figure 8. View largeDownload slide Cadmium exposure increases H4K12ac levels in MII-oocytes. A, MII-oocytes were stained with antiH4K12ac antibody (green) and DAPI (blue). A constant setting of confocal laser-scanning microscope was used for all samples. The average fluorescence intensity for a ROI (white box) surrounding MII-chromatids was measured. Bar: 10 μm. B, The relative fluorescence intensity of H4K12ac in the MII-oocyte was normalized against the fluorescence intensity of DAPI. The total number of oocytes examined is given in parentheses at the bottom of each column. **Statistical differences between the 2 groups by t test at p < .01. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.) Figure 8. View largeDownload slide Cadmium exposure increases H4K12ac levels in MII-oocytes. A, MII-oocytes were stained with antiH4K12ac antibody (green) and DAPI (blue). A constant setting of confocal laser-scanning microscope was used for all samples. The average fluorescence intensity for a ROI (white box) surrounding MII-chromatids was measured. Bar: 10 μm. B, The relative fluorescence intensity of H4K12ac in the MII-oocyte was normalized against the fluorescence intensity of DAPI. The total number of oocytes examined is given in parentheses at the bottom of each column. **Statistical differences between the 2 groups by t test at p < .01. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.) DISCUSSION In this study, we investigated the effects of cadmium treatment of female mice on meiotic maturation of their oocytes and subsequent embryonic development, and the possible mechanisms responsible for the adverse effects. Our results show that cadmium exposure impairs the meiotic maturation of oocytes by disturbing the ovarian follicle microenvironment as well as the ooplasmic components which are essential for meiotic spindle assembly, actin filament localization, ATP metabolism, mitochondrial distribution, and histone modifications. Acute exposure of cadmium is known to affect the hypothalamic-pituitary-ovarian axis and decrease FSH, LH and basal progesterone levels, which are responsible for ovulation (Paksy et al., 1989; Piasek and Laskey, 1994; Thompson and Bannigan, 2008). Our current study demonstrated that cadmium exposure reduced the number of ovulated oocytes after superovulation by approximately 20%. We did not determine if the long-term treatment of cadmium at such a low dosage affected the hormone levels. Nonetheless, stronger effects of cadmium exposure were found in meiotic and developmental competence of oocytes. Our results show that cadmium exposure impaired the oocyte meiotic maturation both in vivo and in vitro. It is known that the oocyte becomes competent for meiotic progression by the end of growth phase (GV-stage) in normal females. Our results showed that the rate of maturation was decreased in the oocytes from cadmium-treated females after either in vivo or IVM. Therefore, cadmium exposure exerted adverse effects on meiotic competence of oocytes at the GV-stage or earlier. However, the impairment was more severe after maturation in vivo than in vitro, indicating that continuous exposure to the cadmium-treated ovarian microenvironment during oocyte maturation has additive adverse effects. Alternatively, the cadmium accumulated in the oocyte during the growth phase may have diffused out during oocyte maturation in vitro. Since the cumulus cells in COCs were also previously exposed to cadmium, we removed them from the oocyte prior to maturation in vitro. The removal of CCs not only decreased the maturation rate in the control group, but also exacerbated the impairment of maturation in the Cd group. These results suggest that the presence of neighboring CCs, despite their exposure to cadmium, is beneficial for oocyte maturation. Consequently, cadmium treatment of female mice impaired the embryonic development, independent of the potential hazardous effects of cadmium in the female genital tract on the spermatozoa. To determine the underlying mechanisms for the impairment of oocyte maturation, we investigated the accumulation and localization of spindle microtubules and actin filaments, which are 2 key components of the cytoskeleton. Spindle microtubules align chromosomes, while actin filaments regulate MII-spindle movements, establish the cortical polarity, and prevent interaction between sperm DNA and meiotic spindle (Laband et al., 2017; Panzica et al., 2017; Yi and Li, 2012). Our results showed that the previous cadmium exposure disrupts meiotic spindle assembly and actin filament accumulation, leading to a loss of the cortical polarity which is critical for the polar body extrusion. However, cadmium exposure did not disrupt the chromosome condensation or alignment at the MII-stage. It has been reported that sufficient ATP levels in oocytes are required for the MII-spindle assembly and embryonic development following IVF (Xu et al., 2014; Zhang et al., 2006). We found that cadmium exposure dramatically reduced the ATP content in the oocyte. The low ATP levels may indicate a functional decline in mitochondria, likely reflected in their abnormal distribution. Our results showed that cadmium exposure disrupted the mitochondrial distribution in the MII-oocytes, which may be partly responsible for the aberrant spindles. Weak fluorescence staining of mitochondria indicates that the number of mitochondria may have been decreased in the oocytes from cadmium-treated females. And it has been reported that cadmium decreased mitochondrial mass and mitochondrial DNA content in human hepatocellular carcinoma cells (Guo et al., 2014). Together, we propose that cadmium exposure disrupted the mitochondrial distribution and reduced ATP contents in oocytes, which may be partly responsible for the impairment of oocyte maturation and subsequent embryonic development. H3K9 methylation has broad roles in transcriptional repression, gene silencing, maintenance of heterochromatin, and serves as an epigenetic marker for the stable heredity of heterochromatin (Krishnan et al., 2011; Martin and Zhang, 2005; Stewart et al., 2006). The increased H3K9me2 in the oocyte would promote the formation of heterochromatin in the female pronucleus after fertilization, resulting in late DNA replication (Liu et al., 2004). Acetylation of histone 4 lysine 12 (H4K12ac) has an important role in the regulation of gene expression and the establishment of an open chromatin configuration, and its persistence is associated with epigenetic-mediated infertility (Paradowska et al., 2012; Vieweg et al., 2015). H4K12 was deacetylated during the first and second meiosis, but it was temporarily acetylated around the time of the first polar body extrusion (Akiyama et al., 2004). Defective deacetylation of H4K12 in the oocytes is correlated with misaligned chromosomes, which results in a high incidence of aneuploidy and embryonic death (Akiyama et al., 2006; van den Berg et al., 2011). Maintenance of H3K9me2 and low levels of H4K12ac during meiotic maturation is closely associated with the developmental competence of oocytes (Racedo et al., 2009; van den Berg et al., 2011). It has been reported that cadmium increases global H3K9me2 in human bronchial epithelial cells (Xiao et al., 2015). Our results showed that cadmium exposure increased the levels of H3K9me2 and H4K12ac, which may result in late DNA replication and aneuploidy after fertilization. This may be partly responsible for the failure of embryonic development when the oocytes were from cadmium-treated female mice. In conclusion, our results indicated that cadmium exposure of female mice impairs the fertility of their oocyte by reducing meiotic maturation capability and subsequent embryonic development. The underlying mechanisms of cadmium toxicity can be attributed to the ovarian follicle microenvironment as well as the cytoplasmic components of oocytes that affect the cytoskeletal organization, mitochondrial function, and histone modifications. FUNDING This work was supported by the National Natural Science Foundation of China (Grant No. 31672620), the Natural Science Foundation of Jiangsu Province (13KJB230003), the Innovation Foundation of Yangzhou University (yzucx2016-6c), Yangzhou University research foundation to Jia-Qiao Zhu, and a project funded by the Priority Academic Program Development of Jiangsu Higher Education Institutions (PAPD). ACKNOWLEDGMENTS We thank Dr Teruko Taketo (McGill University, Montreal, Quebec, Canada) for valuable comments on the article. We also thank Patricia Willers (University of California, Davis, California) for review. REFERENCES Akiyama T. , Kim J. M. , Nagata M. , Aoki F. ( 2004 ). Regulation of histone acetylation during meiotic maturation in mouse oocytes . Mol. Reprod. Dev. 69 , 222 – 227 . Google Scholar CrossRef Search ADS PubMed Akiyama T. , Nagata M. , Aoki F. ( 2006 ). Inadequate histone deacetylation during oocyte meiosis causes aneuploidy and embryo death in mice . Proc. 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A lack of coordination between sister-chromatids segregation and cytokinesis in the oocytes of B6.YTIR (XY) sex-reversed female mice . Sci. Rep. 7 , 960. Google Scholar CrossRef Search ADS PubMed Zou H. , Liu X. , Han T. , Hu D. , Wang Y. , Yuan Y. , Gu J. , Bian J. , Zhu J. , Liu Z. P. ( 2015 ). Salidroside protects against cadmium-induced hepatotoxicity in rats via GJIC and MAPK pathways . PloS One 10 , e0129788. Google Scholar CrossRef Search ADS PubMed © The Author(s) 2018. Published by Oxford University Press on behalf of the Society of Toxicology. All rights reserved. For permissions, please e-mail: journals.permissions@oup.com This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/about_us/legal/notices) http://www.deepdyve.com/assets/images/DeepDyve-Logo-lg.png Toxicological Sciences Oxford University Press

Cadmium Exposure of Female Mice Impairs the Meiotic Maturation of Oocytes and Subsequent Embryonic Development

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Oxford University Press
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© The Author(s) 2018. Published by Oxford University Press on behalf of the Society of Toxicology. All rights reserved. For permissions, please e-mail: journals.permissions@oup.com
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10.1093/toxsci/kfy089
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Abstract

Abstract Cadmium is one major pollutant that is highly toxic to animals and humans. The mechanism of cadmium toxicity on the female reproductive system, particularly oocyte maturation and fertility, remains to be clarified. In this study, we used a mouse model to investigate the effects of cadmium in the drinking water on the meiotic maturation of oocytes and subsequent embryonic development, and the underlying mechanisms associated with the impairment of oocyte maturation such as mitochondrial distribution and histone modifications. Our results show that cadmium exposure decreased the number of ovulated oocytes and impaired oocyte meiotic maturation rate both in vivo and in vitro. The embryonic development after fertilization was also impaired even when the potential hazards of cadmium on the spermatozoa or the genital tract have been excluded by fertilization and embryonic development in culture. Cadmium exposure disrupted meiotic spindle morphology and actin filament, which are responsible for successful chromosome segregation and the polar body extrusion during oocyte maturation and fertilization. ATP contents, which are required for proper meiotic spindle assembly in the oocyte, were decreased, consistent with altered mitochondrial distribution after cadmium exposure. Finally, cadmium exposure affected the levels of H3K9me2 and H4K12ac in the oocyte, which are closely associated with the acquisition of oocyte developmental competence and subsequent embryonic development. In conclusion, cadmium exposure in female mice impaired meiotic maturation of oocytes and subsequent embryonic development by affecting the cytoskeletal organization, mitochondrial function, and histone modifications. cadmium, mouse oocyte, maturation, embryos Cadmium is widely used in the steel industry and manufacture of plastics or batteries, and is easily released into the environment via wastewater, contaminated fertilizers and atmospheric air. As a common pollutant, cadmium is readily accumulated in crops, aquatic organisms and other food-chains (Fan et al., 2017). Human exposure occurs mainly from consumption of contaminated food, active or passive inhalation of tobacco smoke and air inhalation in a metal industry area. Drinking water can be contaminated with cadmium as an impurity in the zinc of galvanized pipes and solder in fittings, water heaters, water coolers or taps. Cadmium is classified as a human carcinogen and is known to have numerous adverse effects on health in both experimental animals and humans. Cadmium-induced toxicity is very well known across the world. Long-term exposure to cadmium through air, water, soil, and food leads to systemic organ toxicity, in the hepatic, renal, skeletal, reproductive, cardiovascular, nervous, and respiratory systems (Rafati Rahimzadeh et al., 2017). The molecular mechanisms of cadmium toxicity are multiple and complex. We have previously reported that disruption of gap junction intercellular communication and activation of mitogen-activated protein kinase pathways are possible mechanisms of cadmium-induced hepatotoxicity (Zou et al., 2015). Furthermore, cadmium disrupts the F-actin cytoskeleton in rat renal mesangial cells, which is mediated by gelsolin translocation, programmed cell death, and F-actin depolymerization through formation of a CAP1-cofilin-F-actin complex (Apostolova et al., 2006; Liu and Templeton, 2013, 2017b). It has been demonstrated that mitochondria copy numbers and mitochondrial function are involved in acute or chronic renal cadmium toxicity (Nair et al., 2015). In addition, cadmium-induced cytotoxicity in primary rat proximal tubular cells can be attributed to the inhibition of the cytosolic Ca2+ dependent autophagosome-lysosome fusion (Liu et al., 2017a). We also found that cadmium-induced apoptosis and autophagy are involved in the cadmium neurotoxicity (Rani et al., 2014; Wang et al., 2015a; Yuan et al., 2016). The extensive deleterious effects of cadmium on the reproductive system and developing embryos have been described, especially in the male (de Angelis et al., 2017; Thompson and Bannigan, 2008; Zhao et al., 2017). Cadmium can disrupt the blood-testis barrier in the testis even at low concentrations by damaging the testis vasculature and exerting the cytotoxicity on Sertoli and Leydig cells, resulting in a decrease in sperm quality, fertilization capacity and subsequent early embryonic development (de Angelis et al., 2017; Zhao et al., 2017). It has been demonstrated that cadmium levels in the ovary increase with age, associated with a failure in ovulation, retardation of trophoblastic outgrowth and embryonic development, placental necrosis, implantation delay, and early pregnancy loss (Thompson and Bannigan, 2008). Furthermore, cadmium exposure inhibits follicular development and increases the number of atretic follicles (Wang et al., 2015b; Zhang et al., 2017). However, it is not well understood how cadmium affects oocyte quality, particularly on meiotic maturation, which is critical for fertilization and embryo development. In most previous studies, the toxicity of cadmium was tested by directly adding it to isolated organs or cells in culture. However, any organ like the ovary has a microenvironment in vivo, through which its viability and functions are regulated. Accordingly, in this study, we administered cadmium in the drinking water to female mice for 35 days and tested its toxic effects on the oocytes both in vivo and in vitro. The environmental concentration of cadmium can reach 75 mg/l, depending on the source of cadmium and the degree of industrialization and pollution (Fan et al., 2017; Li et al., 2015). The dietary intake of cadmium via the food chain in mining areas of China is estimated to be 0.179 mg/day/person on average, which far exceeds the provisional tolerable monthly intake for cadmium of 25 μg/kg body weight specified by the World Health Organization (Du et al., 2013). Moreover, the biological half-life of cadmium is 10–35 years, resulting in long-term accumulation of cadmium in organs. It has been confirmed that long-term exposure to cadmium contributes to the development of lung cancer. The mean blood level of cadmium was 24 μg/l for the local resident in the cadmium high exposure area, which had significantly positive correlations with the risk of all cancer and mortality from all cancers (Wang et al., 2011). In the current study, we chose the cadmium dose 32 mg/l according to our preliminary experiment and the previous report (Yamanobe et al., 2015). The dosage used is in the range found in the environment. The average volume of drinking water was estimated to be 4 ml/day/mouse, so the intake of cadmium was 0.128 mg/day/mouse, which is comparable to the human intake in the mining area of China. Throughout the study, the average body weight of a mouse was estimated to be 35 g, which was used to calculate the cadmium dosage of 3.657 mg/kg/day. This cadmium dose administered was 6% of the LD50 for mice, which was 60 mg/kg/day (Yamanobe et al., 2015; Zhai et al., 2013). In this study, we used the mouse model described earlier to investigate the effects of cadmium on ovulation, meiotic progression in the oocytes during in vivo and in vitro maturation (IVM), and subsequent early embryonic development after in vitro fertilization (IVF). We examined spindle conformation and actin filament localization in the matured oocyte. We also investigated possible mechanisms of cadmium toxicity on mitochondrial distribution, ATP levels, and histone modifications in the oocyte. MATERIALS AND METHODS Materials and chemicals Chemicals, M16 media, bovine serum albumin (BSA), and mouse antiα-tubulin-FITC were purchased from Sigma-Aldrich (St Louis, Missouri). Rabbit monoclonal antiH3K9me2 antibody and polyclonal antiH4K12ac antibody were purchased from AbCam (Cambridge, UK). Donkey antirabbit IgG-Alexa Fluor-488, Rhodamine-phalloidin, Mitotracker Green-FM, Antifade Mountant with DAPI, and MEM-alpha medium were purchased from Thermo Fisher (Waltham, Massachusetts). Cadmium chloride 2.5-hydrate (purity 99.99%) was purchased from Suyi Chemical Company (Shanghai, China). Equine gonadotropin (eCG) and human chorionic gonadotropin (hCG) were obtained from Ningbo Second Hormone Factory (Ningbo, China). KSOMaa medium was purchased from Millipore (Burlington, Massachusetts). Enhanced ATP Assay Kit was purchased from Beyotime Biotechnology Company (Nantong, China). Ethics statement This study was conducted in accordance with the recommendations of the Guide for the Care and Use of Laboratory Animals of the National Research Council. All procedures executed in this study were reviewed and approved by the Animal Care and Use Committee of Yangzhou University (approval ID: SYXK [Su] 2007-0005). Healthy ICR mice (Experimental Animal Center of Yangzhou University), 6–12 weeks-of-age, were used for all experiments. The mice were housed in an air-conditioned animal house (26°C ± 2°C) and exposed to 10–12 h of light per day. Animals were maintained on a standard diet and had free access to water. Animal treatment The mice were randomly divided into 2 groups in each experiment. Every mouse was only used for 1 experiment. The cadmium treatment was carried out according to our preliminary experiment and the previous report. The cadmium dose administered in this study, 32 mg/l, is 6% of the LD50 in mice (Yamanobe et al., 2015). There was no lethality in all cadmium-treated mice throughout the study. For the experimental group, female mice were given ad libitum Milli-Q water containing 32 mg/l cadmium for 35 days. For the control group, Mill-Q water was given ad libitum. Collection of in vivo matured oocytes by superovulation Female mice after cadmium treatment were injected intraperitoneally each with 10 IU eCG, followed by injection with 10 IU hCG 45-47 h later, and sacrificed 14–15 h later. The oocytes enclosed in cumulus cells, named cumulus-oocyte complexes (COCs), were flushed out from the oviduct. After being treated with 300 μg/ml hyaluronidase for 5 min at 37°C and being denuded of cumulus cells by repeated pipetting through a narrow glass needle in the M2 medium, the number of oocyte per mouse was counted. The stage of each oocyte was determined under stereomicroscope, and the number of oocyte in each stage from different mice was pooled in 1 experiment. Collection of germinal vesicle stage oocytes and IVM Female mice after cadmium treatment were injected intraperitoneally each with 10 IU eCG and were sacrificed 48 h later. COCs were isolated by puncturing large preantral and antral follicles with a pair of 26-gauge needles in the M2 medium. For the denuded-oocytes (DOs), COCs were treated with 300 μg/ml hyaluronidase for 5 min at 37°C and oocytes were denuded of cumulus cells by repeated pipetting through a narrow glass needle in the M2 medium. Ten COCs or DOs in a group were cultured in a prewarmed 20 μl droplet of the M16 medium under mineral oil at 37°C with 5% CO2 in a humidified atmosphere. After 18 h, COCs were denuded of cumulus cells, the stage of each oocyte was determined under stereomicroscope. The oocytes in each stage from different mice were pooled in each experiment. IVF and embryonic development Spermatozoa were collected from the caudal epididymis of males that were not treated with cadmium at 10–12 weeks of age and capacitated in the MEM-alpha medium supplemented with 0.9% BSA under mineral oil at 37°C for 60 min. The ovulated oocytes (in vivo matured) were transferred into an IVF medium (MEM-alpha supplemented with 0.4% BSA) containing spermatozoa under oil and further incubated for 5 h at 37°C with 5% CO2 in a humidified atmosphere. After washing in the modified KSOMaa medium, the fertilized eggs were cultured in the fresh modified KSOMaa medium for up to 5 days. For the in vivo fertilized oocytes, female mice that were injected with 10 IU eCG and 45–47 h later with 10 IU hCG were left with male mice overnight. The female mice found with vaginal plugs next morning were considered to have successfully copulated. After 0.5 day, the in vivo fertilized eggs were flushed out from the oviduct, and cultured as above for up to 5 days. The stage of each embryo was determined under stereomicroscope every day, and the embryos in each stage from different experiments were pooled. Confocal microscopy analysis of meiotic spindles and actin filaments Immunofluorescence staining was done as previously described in Zhu et al. (2017). Oocytes were fixed with 4% paraformaldehyde in phosphate-buffered saline (PBS) for 30 min at room temperature. Then, oocytes were blocked in PBS containing BSA (3%) and Triton-X-100 (0.1%) (named blocking solution) for 1 h at room temperature and incubated with mouse antiα-tubulin-FITC (1:500) and Rhodamine-phalloidin (1:200) overnight at 4°C. After 3 washes in PBS, oocytes were transferred into Antifade Mountant with DAPI on glass slides. Fluorescence signals were examined under the confocal laser-scanning microscope (Leica TCS SP8). A rugby ball shape of α-tubulin was defined as the normal meiotic spindle. A region of interest (ROI) of actin was defined as a dotted line to cross the center of oocyte and midzone of spindle. Higher fluorescence intensity of ROI in the oolemma than that in the ooplasm was defined as a normal actin distribution. The proportion of abnormal oocytes was counted in each experiment. Measurement of ATP contents ATP contents in oocytes were determined by using Enhanced ATP Assay Kit according to the manufacturer’s protocol. 40 oocytes in a group were added to ice-cold Cell ATP-Releasing Reagent, following the addition of ATP Assay Reagent. The bioluminescence of each sample was measured using a high-sensitivity luminometer (BioTek). Examination of mitochondrial distribution Oocytes were fixed in 4% paraformaldehyde in PBS for 25 min. After washing, oocytes were incubated in 100 nM Mitotracker Green-FM in the M2 medium for 35 min in dark. After washing, oocytes were transferred into Antifade Mountant with DAPI on glass slides. Fluorescence signals were examined under the confocal laser-scanning microscope (Leica TCS SP8). Mitochondrial distribution patterns were classified according to the practices used in previous reports (Moawad et al., 2014; Wang et al., 2009). The proportion of abnormal oocytes was calculated in each experiment. Immunofluorescence staining of H3K9me2 and H4K12ac Oocytes were fixed as described earlier, and incubated with rabbit monoclonal antiH3K9me2 antibody (1:200) or rabbit polyclonal antiH4K12ac antibody (1:200) overnight at 4°C. After washing, oocytes were incubated with donkey-antirabbit IgG- Alexa Fluor-488 (1:500) at room temperature for 1 h. After washing, oocytes were transferred into Antifade Mountant with DAPI on glass slides and were examined under the confocal laser-scanning microscope. Oocytes in the experimental and control groups were mounted on the same glass slide, and the constant scanning setting was used for sample scanning. Leica microscope software (Leica LAS AF) was used to assess oocyte fluorescence intensity levels. A ROI was defined as encompassing all the chromosomes in each oocyte, and the average fluorescence intensity for this ROI was determined. The relative fluorescence intensity of H3K9me2 and H4K12ac in each oocyte was normalized against the fluorescence intensity of DAPI. Statistical analysis At least 20 oocytes were examined in 1 experiment. All experiments were repeated at least 3 times. Data are presented as mean ± SEM. Statistical differences among pooled results were analyzed by 2-way ANOVA using SPSS software. Statistical differences between groups from separate experiments were analyzed by t test using SPSS software. A p < .05 was considered statistically significant. RESULTS Cadmium Exposure Reduces the Number of Ovulated Oocytes We first tested the effects of cadmium in drinking water on ovulation. As shown in Figure 1, the number of ovulated oocytes from the cadmium-treated group (Cd) was significantly less than that from the control group (Con) (42.77 ± 2.44 vs 52.43 ± 3.06/female; p < .05). These results show that cadmium exposure reduces the number of ovulated oocytes after superovulation. Figure 1. View largeDownload slide Cadmium exposure (Cd) reduces the number of ovulated oocytes. Data are presented as means ± SEM. The total number of females examined is given in parentheses at the bottom of each column. *Statistical differences between the oocytes from 2 groups by t test at p < .05. Figure 1. View largeDownload slide Cadmium exposure (Cd) reduces the number of ovulated oocytes. Data are presented as means ± SEM. The total number of females examined is given in parentheses at the bottom of each column. *Statistical differences between the oocytes from 2 groups by t test at p < .05. Cadmium Exposure Impairs the Oocyte Maturation In Vivo We next examined the morphology of ovulated oocytes under the stereo microscope. As shown in Figure 2A, most oocytes were at the MI-stage without the first polar body in the Cd group (50.58% ± 2.60%); compared with the control group (24.29% ± 3.55%) with a significant difference (p < .01). Conversely, the percentage of MII-oocytes that had extruded the first polar body in the Cd group was significantly smaller than that in the control group (29.65% ± 3.17% vs 72.60% ± 3.63%; p < .01). These results indicate that cadmium exposure impaired the oocyte maturation in vivo. Figure 2. View largeDownload slide Cadmium exposure impairs the oocyte maturation both in vivo and in vitro. A, In vivo maturation rate of oocytes were significantly reduced after female mice were exposed to cadmium. In vitro maturation rate of oocytes with (B) or without (C) cumulus cells was significantly reduced after female mice were exposed to cadmium prior to COCs collection. For the DOs, COCs were denuded of cumulus cells before in vitro culture. In vivo maturation and in vitro maturation were repeated 10 and 5 times, respectively. *,**Statistical differences of each stage between the oocytes from 2 groups by t test at p < .05 and .01, respectively. Figure 2. View largeDownload slide Cadmium exposure impairs the oocyte maturation both in vivo and in vitro. A, In vivo maturation rate of oocytes were significantly reduced after female mice were exposed to cadmium. In vitro maturation rate of oocytes with (B) or without (C) cumulus cells was significantly reduced after female mice were exposed to cadmium prior to COCs collection. For the DOs, COCs were denuded of cumulus cells before in vitro culture. In vivo maturation and in vitro maturation were repeated 10 and 5 times, respectively. *,**Statistical differences of each stage between the oocytes from 2 groups by t test at p < .05 and .01, respectively. Cadmium Exposure Impairs the Oocyte Maturation In Vitro After Separating From the Follicular Microenvironment Oocyte competence for meiotic progression depends on the ovarian follicle microenvironment during the oocyte growth phase, whereas the continual presence of neighboring cumulus cells facilitate oocyte maturation and competence for embryonic development (Eppig, 2001; Villemure et al., 2007; Xu et al., 2014). In the above study, both the oocyte and its surrounding somatic cells were exposed to cadmium throughout the growth and maturation of oocytes, so we asked whether the systematic presence of cadmium directly affected the oocyte meiotic maturation or the ovarian follicle microenvironment which in turn impaired oocyte maturation. We isolated the oocytes from follicular environment and matured them in vitro. Furthermore, a group of oocytes were left surrounded by cumulus cells (COC group), while another group of oocytes was denuded of cumulus cells (DO group) before maturation in vitro. As shown in Figures 2B and 2C, the percentages of MII-oocytes were significantly smaller (p < .05) in both COC and DO groups from the Cd-treated females (85.64% ± 3.00% and 46.11% ± 5.84%, respectively), compared with the control females (94.70% ± 1.25% and 70.62% ± 4.97%, respectively). Conversely, percentages of MI-oocytes were significantly larger in the Cd group compared with the control group. However, the percentages of MII-oocytes in the Cd IVM group were significantly higher than those in Cd in vivo maturation group (Figure 2, COC 85.64% ± 3.00% and DO 46.11% ± 5.84% vs 29.65% ± 3.17%). These results indicate that meiotic maturation of oocyte was partially improved when the oocyte was separated from the ovarian follicle microenvironment. We conclude that cadmium exposure affected the ovarian follicle microenvironment which then continually impaired the maturation of oocyte in vivo. Cadmium Exposure Impairs Embryonic Development We next compared the development of eggs from both cadmium-treated female mice and the control female mice. The results are summarized in Figure 3B. In the Cd group, the rate of the first cleavage (2-cell) was comparable with that of the control group. However, the rate of 4-cell stage embryo development after 2 days in culture (57.9%, n = 242) was significantly lower and that of morula after 3 days culture (46.7%) was also lower, although without statistical difference compared with the control group (82.2% and 68.0%, respectively, n = 309). After 4 and 5 days in culture, 42.2% and 48.8% of fertilized eggs reached the blastocyst-stage in the Cd group, while 66.0% and 75.4% did so in the control group (p < .05 and .01, respectively). When the proportions of embryos at various stages were statistically analyzed, significant differences between the 2 groups were found (2-way ANOVA, p < .001). These results indicate that cadmium treatment of females made their eggs incompetent for embryonic development even when the eggs were separated from the oviduct and the uterus which had been exposed to cadmium. Figure 3. View largeDownload slide Cadmium exposure impairs the embryonic development of naturally fertilized eggs (in vivo) and in vitro fertilized eggs (IVF). A, Examples of embryos at 1–5 days in culture. Bar: 50 μm. “Others” includes dead or fragmented embryos (arrow). Each column indicates the percentages of embryos at different stages at 1–5 days in culture from in vivo (B) and in vitro fertilized eggs (C). The total number of embryos examined is given in parentheses at the top of each graph. The data were pooled from 4 (in vivo) and 6 (IVF) experiments. Significant differences between the 2 groups by 2-way ANOVA. *,**Statistical differences of each stage between the 2 groups at each day by t test at p < .05 and .01, respectively. Figure 3. View largeDownload slide Cadmium exposure impairs the embryonic development of naturally fertilized eggs (in vivo) and in vitro fertilized eggs (IVF). A, Examples of embryos at 1–5 days in culture. Bar: 50 μm. “Others” includes dead or fragmented embryos (arrow). Each column indicates the percentages of embryos at different stages at 1–5 days in culture from in vivo (B) and in vitro fertilized eggs (C). The total number of embryos examined is given in parentheses at the top of each graph. The data were pooled from 4 (in vivo) and 6 (IVF) experiments. Significant differences between the 2 groups by 2-way ANOVA. *,**Statistical differences of each stage between the 2 groups at each day by t test at p < .05 and .01, respectively. Considering the potential effects of cadmium on spermatozoa motility and fertilization through contact with the oviduct and the uterus (Zhao et al., 2017), we used IVF. As shown in Figure 3C, the percentage of blastocyst development was significantly smaller in the Cd group than that of the control group (33.68% vs 48.41%, p < .05) after 5 days in culture. In addition, more embryos were found dead or fragmented (others in Figure 3) in the Cd group. When the proportions of embryos at various stages after IVF were statistically analyzed, significant differences between the 2 groups were also found (2-way ANOVA, p < .001). These results demonstrate that the embryonic development was impaired when oocytes were collected from cadmium-treated female mice. Cadmium Exposure Disrupts Meiotic Spindle Morphology and Actin Accumulation and Distribution Next, we examined the cytoskeletal integrity of ovulated oocytes, which indicate quality of the oocyte. As shown in Figure 4A, the meiotic spindle labeled with α-tubulin antibody in the control group typically had a rugby ball shape, but some had a bulky shape with at least 1 pole wider than the midzone. The proportion of oocytes with aberrant spindles such as these in the Cd group was significantly higher than in the control group (Figure 4B; 50.00% ± 10.94% vs 12.87% ± 3.44%, p < .01). Figure 4. View largeDownload slide Cadmium exposure disrupts meiotic spindle morphology and actin accumulation and distribution. A, The MII-oocyte was stained with antiα-tubulin-FITC (green), rhodamine-phalloidin (actin, red) and DAPI (blue). In the control group, the typical meiotic spindle (α-tubulin) was seen in a rugby ball shape, and the actin filaments (actin) were localized along the oolemma and concentrated close to the MII-spindle as well as around the first polar body. Fluorescence intensity of actin along in the dotted line shows that actin accumulation in the oolemma was significantly higher than that in the ooplasm. However, in the Cd group, the spindle was bulky (at least 1 pole was wider than the midzone), and the actin accumulation in the oolemma was absent even when the fluorescence signals actin in the ooplasm was overexposed. Bar: 20 μm. (B, C) The rate of aberrant spindle and lack of actin localization in the Cd group was significantly higher than in the control group. The total number of oocytes examined, from 4 experiments, is given in parentheses at the right of the graph. **Statistical differences between the 2 groups by t test at p < .01. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.) Figure 4. View largeDownload slide Cadmium exposure disrupts meiotic spindle morphology and actin accumulation and distribution. A, The MII-oocyte was stained with antiα-tubulin-FITC (green), rhodamine-phalloidin (actin, red) and DAPI (blue). In the control group, the typical meiotic spindle (α-tubulin) was seen in a rugby ball shape, and the actin filaments (actin) were localized along the oolemma and concentrated close to the MII-spindle as well as around the first polar body. Fluorescence intensity of actin along in the dotted line shows that actin accumulation in the oolemma was significantly higher than that in the ooplasm. However, in the Cd group, the spindle was bulky (at least 1 pole was wider than the midzone), and the actin accumulation in the oolemma was absent even when the fluorescence signals actin in the ooplasm was overexposed. Bar: 20 μm. (B, C) The rate of aberrant spindle and lack of actin localization in the Cd group was significantly higher than in the control group. The total number of oocytes examined, from 4 experiments, is given in parentheses at the right of the graph. **Statistical differences between the 2 groups by t test at p < .01. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.) We also examined the actin filaments, which play a role in the polar body extrusion. In the control group, actin filaments were localized along the oolemma and concentrated over the MII-spindle in addition to the first polar body; however, the actin filaments were no longer seen along the oolemma in the Cd group even when the fluorescence signals were enhanced (Figure 4A). The proportion of oocytes that lacked actin localization on the oolemma in the Cd group was significantly higher than that in the control group (Figure 4C; 37.04% ± 5.63% vs 6.93 ± 1.08%, p < .01). We did not find any difference in chromosome condensation or alignment by labeling with DAPI within the oocytes in the 2 groups. These results indicate that cadmium exposure disrupted the meiotic spindle assembly and actin localization, resulting in the loss of the cortical polarity. Cadmium Exposure Reduces ATP Contents by Disrupting the Mitochondrial Distribution in the Oocyte ATP is mainly synthesized by mitochondria in the oocyte and plays a critical role in proper spindle assembly and fertilization (Van Blerkom et al., 1995; Xu et al., 2014; Zhang et al., 2006). We measured ATP contents in the MII-oocytes from the cadmium-treated female mice. The results showed that the ATP contents in the Cd group were significantly lower than those in the control group (2.03 ± 0.11 vs. 3.00 ± 0.10 μmol/l, p < .01) (Figure 5). It is also known that mitochondrial distribution in the MII-oocytes is associated with the competence for embryonic development (Bavister and Squirrell, 2000; Wilding et al., 2001). Accordingly, we examined the mitochondrial distribution with Mitotracker Green-FM staining, and classified the results into 3 different patterns: accumulation around the MII-spindle (Figure 6Ai), homogeneous distribution over the ooplasm (Figure 6Aii), and peripheral accumulation with overall weak fluorescence signals (Figure 6Aiii). As shown in Figure 6B, the majority of oocytes in the control group showed accumulation around the MII-spindle (pattern i) (76.92 ± 3.91%); however, the proportion of oocytes in this pattern was significantly lower in the Cd group (56.44% ± 4.96%, p < .01). Instead, a considerable number of oocytes in the Cd group showed peripherally accumulated mitochondria with overall weak fluorescence signals (pattern iii), and these were found at a significantly larger percentage than in the control group (33.66% ± 4.73% vs 8.55% ± 2.60%, p < .01). These results indicate that cadmium exposure disrupts the mitochondrial distribution, which may be responsible for lower ATP contents in the MII-oocytes of the Cd group. Figure 5. View largeDownload slide Cadmium exposure reduces ATP contents in MII-oocytes. This experiment was repeated 9 times. **Statistical differences between the 2 groups by t test at p < .01. Figure 5. View largeDownload slide Cadmium exposure reduces ATP contents in MII-oocytes. This experiment was repeated 9 times. **Statistical differences between the 2 groups by t test at p < .01. Figure 6. View largeDownload slide Cadmium exposure disrupts the mitochondrial distribution in MII-oocytes. A, The MII-oocytes were labeled with Mitotracker Green-FM to visualize mitochondrial localization. Mitochondrial distribution was classified using 3 different patterns: accumulation around the MII-spindle (i), homogeneous distribution over the ooplasm (ii), and peripheral concentration with overall weak signals (iii). Bar: 20 μm. (B) Quantification of oocytes in each mitochondrial distribution pattern. The total number of oocytes examined is given in parentheses at the right of the graph, from 3 experiments. ** Statistical differences between the 2 groups by t test at p < .01. Figure 6. View largeDownload slide Cadmium exposure disrupts the mitochondrial distribution in MII-oocytes. A, The MII-oocytes were labeled with Mitotracker Green-FM to visualize mitochondrial localization. Mitochondrial distribution was classified using 3 different patterns: accumulation around the MII-spindle (i), homogeneous distribution over the ooplasm (ii), and peripheral concentration with overall weak signals (iii). Bar: 20 μm. (B) Quantification of oocytes in each mitochondrial distribution pattern. The total number of oocytes examined is given in parentheses at the right of the graph, from 3 experiments. ** Statistical differences between the 2 groups by t test at p < .01. Cadmium Exposure Increases H3K9me2 and H4K12ac Levels in MII-Oocytes Histone modifications, particularly dimethylation of H3K9 (H3K9me2) and deacetylation of H4K12 (H4K12ac), during meiotic maturation of oocytes are essential for the developmental competence of oocytes (Racedo et al., 2009; van den Berg et al., 2011). As shown in Figure 7, the relative fluorescence intensity of H3K9me2 in the Cd group was significantly increased, compared with that of the control group (0.72 ± 0.02 vs 0.54 ± 0.03, p < .01). The H4K12ac intensity in the Cd group was also significantly increased, compared with that of the control group (Figure 8, 0.65 ± 0.03 vs 0.54 ± 0.02, p < .01). These results indicate that cadmium exposure increases both H3K9me2 and H4K12ac levels in MII-oocytes. Figure 7. View largeDownload slide Cadmium exposure increases H3K9me2 levels in MII-oocytes. A, MII-oocytes were stained with antiH3K9me2 antibody (green) and DAPI (blue). A constant setting of confocal laser-scanning microscope was used for all samples. The average fluorescence intensity for a ROI (white box) surrounding MII-chromatids was measured. Bar: 10μm. B, The relative fluorescence intensity of H3K9me2 in the MII-oocyte was normalized against the fluorescence intensity of DAPI. The total number of oocytes examined is given in parentheses at the bottom of each column. **Statistical differences between the 2 groups by t test at p < .01. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.) Figure 7. View largeDownload slide Cadmium exposure increases H3K9me2 levels in MII-oocytes. A, MII-oocytes were stained with antiH3K9me2 antibody (green) and DAPI (blue). A constant setting of confocal laser-scanning microscope was used for all samples. The average fluorescence intensity for a ROI (white box) surrounding MII-chromatids was measured. Bar: 10μm. B, The relative fluorescence intensity of H3K9me2 in the MII-oocyte was normalized against the fluorescence intensity of DAPI. The total number of oocytes examined is given in parentheses at the bottom of each column. **Statistical differences between the 2 groups by t test at p < .01. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.) Figure 8. View largeDownload slide Cadmium exposure increases H4K12ac levels in MII-oocytes. A, MII-oocytes were stained with antiH4K12ac antibody (green) and DAPI (blue). A constant setting of confocal laser-scanning microscope was used for all samples. The average fluorescence intensity for a ROI (white box) surrounding MII-chromatids was measured. Bar: 10 μm. B, The relative fluorescence intensity of H4K12ac in the MII-oocyte was normalized against the fluorescence intensity of DAPI. The total number of oocytes examined is given in parentheses at the bottom of each column. **Statistical differences between the 2 groups by t test at p < .01. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.) Figure 8. View largeDownload slide Cadmium exposure increases H4K12ac levels in MII-oocytes. A, MII-oocytes were stained with antiH4K12ac antibody (green) and DAPI (blue). A constant setting of confocal laser-scanning microscope was used for all samples. The average fluorescence intensity for a ROI (white box) surrounding MII-chromatids was measured. Bar: 10 μm. B, The relative fluorescence intensity of H4K12ac in the MII-oocyte was normalized against the fluorescence intensity of DAPI. The total number of oocytes examined is given in parentheses at the bottom of each column. **Statistical differences between the 2 groups by t test at p < .01. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.) DISCUSSION In this study, we investigated the effects of cadmium treatment of female mice on meiotic maturation of their oocytes and subsequent embryonic development, and the possible mechanisms responsible for the adverse effects. Our results show that cadmium exposure impairs the meiotic maturation of oocytes by disturbing the ovarian follicle microenvironment as well as the ooplasmic components which are essential for meiotic spindle assembly, actin filament localization, ATP metabolism, mitochondrial distribution, and histone modifications. Acute exposure of cadmium is known to affect the hypothalamic-pituitary-ovarian axis and decrease FSH, LH and basal progesterone levels, which are responsible for ovulation (Paksy et al., 1989; Piasek and Laskey, 1994; Thompson and Bannigan, 2008). Our current study demonstrated that cadmium exposure reduced the number of ovulated oocytes after superovulation by approximately 20%. We did not determine if the long-term treatment of cadmium at such a low dosage affected the hormone levels. Nonetheless, stronger effects of cadmium exposure were found in meiotic and developmental competence of oocytes. Our results show that cadmium exposure impaired the oocyte meiotic maturation both in vivo and in vitro. It is known that the oocyte becomes competent for meiotic progression by the end of growth phase (GV-stage) in normal females. Our results showed that the rate of maturation was decreased in the oocytes from cadmium-treated females after either in vivo or IVM. Therefore, cadmium exposure exerted adverse effects on meiotic competence of oocytes at the GV-stage or earlier. However, the impairment was more severe after maturation in vivo than in vitro, indicating that continuous exposure to the cadmium-treated ovarian microenvironment during oocyte maturation has additive adverse effects. Alternatively, the cadmium accumulated in the oocyte during the growth phase may have diffused out during oocyte maturation in vitro. Since the cumulus cells in COCs were also previously exposed to cadmium, we removed them from the oocyte prior to maturation in vitro. The removal of CCs not only decreased the maturation rate in the control group, but also exacerbated the impairment of maturation in the Cd group. These results suggest that the presence of neighboring CCs, despite their exposure to cadmium, is beneficial for oocyte maturation. Consequently, cadmium treatment of female mice impaired the embryonic development, independent of the potential hazardous effects of cadmium in the female genital tract on the spermatozoa. To determine the underlying mechanisms for the impairment of oocyte maturation, we investigated the accumulation and localization of spindle microtubules and actin filaments, which are 2 key components of the cytoskeleton. Spindle microtubules align chromosomes, while actin filaments regulate MII-spindle movements, establish the cortical polarity, and prevent interaction between sperm DNA and meiotic spindle (Laband et al., 2017; Panzica et al., 2017; Yi and Li, 2012). Our results showed that the previous cadmium exposure disrupts meiotic spindle assembly and actin filament accumulation, leading to a loss of the cortical polarity which is critical for the polar body extrusion. However, cadmium exposure did not disrupt the chromosome condensation or alignment at the MII-stage. It has been reported that sufficient ATP levels in oocytes are required for the MII-spindle assembly and embryonic development following IVF (Xu et al., 2014; Zhang et al., 2006). We found that cadmium exposure dramatically reduced the ATP content in the oocyte. The low ATP levels may indicate a functional decline in mitochondria, likely reflected in their abnormal distribution. Our results showed that cadmium exposure disrupted the mitochondrial distribution in the MII-oocytes, which may be partly responsible for the aberrant spindles. Weak fluorescence staining of mitochondria indicates that the number of mitochondria may have been decreased in the oocytes from cadmium-treated females. And it has been reported that cadmium decreased mitochondrial mass and mitochondrial DNA content in human hepatocellular carcinoma cells (Guo et al., 2014). Together, we propose that cadmium exposure disrupted the mitochondrial distribution and reduced ATP contents in oocytes, which may be partly responsible for the impairment of oocyte maturation and subsequent embryonic development. H3K9 methylation has broad roles in transcriptional repression, gene silencing, maintenance of heterochromatin, and serves as an epigenetic marker for the stable heredity of heterochromatin (Krishnan et al., 2011; Martin and Zhang, 2005; Stewart et al., 2006). The increased H3K9me2 in the oocyte would promote the formation of heterochromatin in the female pronucleus after fertilization, resulting in late DNA replication (Liu et al., 2004). Acetylation of histone 4 lysine 12 (H4K12ac) has an important role in the regulation of gene expression and the establishment of an open chromatin configuration, and its persistence is associated with epigenetic-mediated infertility (Paradowska et al., 2012; Vieweg et al., 2015). H4K12 was deacetylated during the first and second meiosis, but it was temporarily acetylated around the time of the first polar body extrusion (Akiyama et al., 2004). Defective deacetylation of H4K12 in the oocytes is correlated with misaligned chromosomes, which results in a high incidence of aneuploidy and embryonic death (Akiyama et al., 2006; van den Berg et al., 2011). Maintenance of H3K9me2 and low levels of H4K12ac during meiotic maturation is closely associated with the developmental competence of oocytes (Racedo et al., 2009; van den Berg et al., 2011). It has been reported that cadmium increases global H3K9me2 in human bronchial epithelial cells (Xiao et al., 2015). Our results showed that cadmium exposure increased the levels of H3K9me2 and H4K12ac, which may result in late DNA replication and aneuploidy after fertilization. This may be partly responsible for the failure of embryonic development when the oocytes were from cadmium-treated female mice. In conclusion, our results indicated that cadmium exposure of female mice impairs the fertility of their oocyte by reducing meiotic maturation capability and subsequent embryonic development. The underlying mechanisms of cadmium toxicity can be attributed to the ovarian follicle microenvironment as well as the cytoplasmic components of oocytes that affect the cytoskeletal organization, mitochondrial function, and histone modifications. FUNDING This work was supported by the National Natural Science Foundation of China (Grant No. 31672620), the Natural Science Foundation of Jiangsu Province (13KJB230003), the Innovation Foundation of Yangzhou University (yzucx2016-6c), Yangzhou University research foundation to Jia-Qiao Zhu, and a project funded by the Priority Academic Program Development of Jiangsu Higher Education Institutions (PAPD). ACKNOWLEDGMENTS We thank Dr Teruko Taketo (McGill University, Montreal, Quebec, Canada) for valuable comments on the article. We also thank Patricia Willers (University of California, Davis, California) for review. REFERENCES Akiyama T. , Kim J. M. , Nagata M. , Aoki F. ( 2004 ). Regulation of histone acetylation during meiotic maturation in mouse oocytes . Mol. Reprod. Dev. 69 , 222 – 227 . Google Scholar CrossRef Search ADS PubMed Akiyama T. , Nagata M. , Aoki F. ( 2006 ). Inadequate histone deacetylation during oocyte meiosis causes aneuploidy and embryo death in mice . Proc. 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Toxicological SciencesOxford University Press

Published: Apr 19, 2018

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