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State Key Laboratory of Reproductive Biology,3 Institute of Zoology, Chinese Academy of Sciences, Beijing 100080, China
Guangzhou Second People's Hospital,4 Guangzhou 510150, China
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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aging, gamete biology, gametogenesis, MAD2, meiosis, meiosis II, oocyte, oocyte development, pig
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Aneuploidy is one of the most common characteristics associated with abnormal fertilization in mammals, and it results from abnormal chromosome separation during meiosis I and/or meiosis II [4, 5]. Kinetochores play a key role in chromosome separation during mitosis and meiosis and proteins associated with kinetochores are functionally responsible for the spindle formation, chromosome movement, and attachment to the spindles, controlling and monitoring the onset of anaphase during meiosis [10–12]. Any adverse effects on the structure and function of kinetochores affect equal distribution of chromosomes in daughter cells, resulting in aneuploidy [13–15]. Proteins associated with kinetochores can be classified into three types based on their functions: the first type comprises constitutive proteins, such as centromere protein (CENP)A, CENPB, CENPC, and CENPD. These proteins form the basic structure of kinetochores and establish the connection between microtubules and chromosomes; the second type comprises checkpoint proteins, such as mitotic arrest-deficient protein (MAD) 1, MAD2, and MAD3, budding uninhibited by benzimidazole (BUB) 1, BUB2, and BUB3 [14]. These proteins are transiently associated with kinetochores and monitor the defects of spindle integrity and chromosome alignment at the metaphase plate and produce signals to delay anaphase onset until these defects are corrected [11, 12, 16]; and the third type comprises motor proteins, such as CENPE, cytoplasmic dynein, and mitotic centromere-associated kinesin. These proteins play a role in the microtubule assembly and disassembly on kinetochores [15].
Recent studies indicate that mitotic spindle checkpoint protein MAD2 is also a meiotic spindle checkpoint protein during meiosis [11, 12, 17]. Different intracellular levels and localization of MAD2 were observed in mitosis as compared with meiosis [18]. Absence of MAD2 increased the frequency of meiosis I missegregation during meiosis [17], while microinjected MAD2 delayed the sister chromatid segregation at anaphase II in Xenopus oocytes [19]. The mRNA level of MAD2 is much lower in aged human oocytes than in young human oocytes [13], implying an age-related decrease in the activity of MAD2 in oocytes. Low activity of MAD2 in aged oocytes failed to inhibit the ability of the anaphase-promoting complex to ubiquinate its target proteins, inducing premature degradation of maturation/M-phase promoting factor (MPF) and the cohesin between sister chromatids, thus causing precocious separation of sister chromatids [19]. Reduced activity of MPF and increased premature centromere separation (PCS) in aged oocytes were also observed in some mammals [1, 20–23]. PCS predisposes sister chromatids to randomly segregate during anaphase II and increases the probability of aneuploidy in zygotes [4, 9, 20, 21, 24].
Mitogen-activated protein kinase (MAPK) is another key protein that plays an important role in the cell-cycle progression and metaphase II (M-II) arrest in mammalian oocytes [1]. High levels of MAPK activity in M-II oocytes are necessary for M-II arrest in the oocytes and their activities gradually reduced as oocytes are aging [2, 22, 25]. MAPK is also important for the spindle checkpoint in Xenopus egg extract [26]. Reduced levels of MPF and MAPK also lead to ubiquitination and degradation of the BCL2 protein, one of the critical antiapoptosis proteins in oocytes [1, 3]. BCL2 is present in mitochondria, nuclear membrane, and endoplasmic reticulum [27, 28]. Reduced BCL2 activity decreases the potential of mitochondria and downregulates the Ca2+ pump and its mRNA on endoplasmic reticulum [1, 27, 28], thereby reducing the ATP production, altering Ca2+ homeostasis [28], and increasing the level of cytochrome C and active oxygen in oocytes [1]. Although the aged oocytes are easily activated by chemical agents or sperm protein factor [29], this treatment is usually followed by abnormal [Ca2+]i oscillation and low pronuclear formation but high fragmentation in oocytes [8, 29–31].
Taken together, it appears that a complex network of checkpoint components participates in the generation of the spindle checkpoint signal at the kinetochores. Although morphological changes and developmental competence of embryos after fertilization of aged oocytes have been reported in some mammalian oocytes [31–33], biochemical events, especially those related to kinetochores within oocytes during aging are still unknown. In a previous study, we found that the kinetochores could be detected in pig oocytes during meiosis I and the changes in kinetochore location were associated with chromosome movement and spindle formation [34]. In the present study, experiments were conducted to compare the expression levels of some proteins associated with kinetochores and spindles, including tubulin (a microtubule constitutive protein), CENPB (a kinetochore constitutive protein), MAD2 (a spindle checkpoint protein), BCL2 (an antiapoptosis protein), and the activities of MAPK (plays as cytostatic factor) in young (newly matured) and aging pig oocytes. Some oocytes at different time intervals of culture were activated by calcium ionophore to allow examination of early events of anaphase II, embryo fragmentation, and apoptosis.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The collection and culture of oocytes were based on the procedures reported previously [35]. Briefly, ovaries were collected from prepubertal gilts at a local slaughterhouse and transported to the laboratory in 0.9% NaCl containing 75 µg penicillin G/ml and 50 µg streptomycin sulfate/ml maintained at 37°C. Oocytes were aspirated from medium-sized follicles (3
6 mm in diameter) with a 20-gauge needle fixed to a 20-ml disposable syringe. The oocytes with uniform ooplasm and also surrounded by a compact cumulus mass were selected and washed four times with maturation medium: tissue culture medium (TCM)-199 supplemented with 0.57 mM cysteine (Sigma Chemical Co., St Louis, MO), 10 ng epidermal growth factor/ml (Sigma), 10 IU eCG /ml (Sigma), 10 IU hCG/ml (Sigma), and 0.1% polyvinyl alcohol. A group of 50 oocytes was cultured in a 500-µl maturation medium at 39°C in an atmosphere of 5% CO2 in air and saturated humidity.
At 40, 48, 60, and 72 h after culture, cumulus cells were removed by 0.02% (w/v) hyaluronidase (Sigma) in the maturation medium. After 3–4 rinses, cumulus-free oocytes were used for the experiments.
Assessment of Nuclear Maturation
At 40–72 h of culture, the denuded oocytes were mounted and fixed for 48–72 h in 25% (v/v) acetic acid in ethanol at room temperature, stained with 1% (w/v) orcein in 45% (v/v) acetic acid and examined under a phase-contrast microscope at a magnification of 400x. The nuclear stages were classified as germinal vesicle (GV), prometaphase I (ProM-I), metaphase I (M-I), anaphase I (A-I), telophase I (T-I), and M-II. Oocytes at the stage of M-II were regarded as matured [36].
Mitochondrial Examination in Oocytes During Aging
Mitochondria distribution in the oocytes was examined by staining the oocytes with mitochondria probe MitoTracker. Briefly, oocytes were removed from enclosed cumulus cells after different times of culture and the denuded oocytes were cultured in TCM-199 containing 12.5 µM MitoTrackerRed (Molecular Probes, Eugene, OR) at 39°C for 30 min. After staining, the oocytes were washed in PBS twice and then mounted on slides and examined under a fluorescence microscope.
Calcium Ionophore Activation of Oocytes
The method used for oocyte activation by calcium ionophore A23187 (Sigma) was essentially the same as that reported by Wang et al. [37]. Cumulus-free oocytes were treated with 50 µM A23187 for 5 min in a fertilization medium [36]. After treatment, oocytes were washed six times in the fertilization medium and each group of 30 oocytes was cultured for 2 h for examination of anaphase II onset by staining spindle and chromosomes. To assess oocyte fragmentation, the artificially activated oocytes were washed thoroughly in North Carolina State University (NCSU)-23 medium containing 4 mg/ml BSA (A8022; Sigma) and then cultured in the same medium for 48 h at 39°C, 5% CO2 in air. Fragmentation was examined at 24, 36, and 48 h after culture.
Apoptosis Assays
The apoptosis signals in the parthenogenetic embryos were examined with a TUNEL procedure after Days 2 and 5 of culture of the oocytes being activated with A23187. In brief, embryos were washed three times in PBS/PVP (PBS with 0.1% polyvinylpyrrolidone) and then fixed in 4% paraformaldehyde/PBS/PVP for 24 h at room temperature. After three washes, each for 5 min, the embryos were treated with 0.1% Triton X-100 in 0.1% citrate solution for 1 h at room temperature. After being washed in PBS/PVP, the embryos were incubated in TUNEL reaction medium for 1 h at 39°C, then washed and stained with 10 µg propidium iodide (PI)/ml for 20 min at room temperature for examination of nuclei. The embryos were mounted on slides with antifade solution (0.5% n-propyl gallate in 20 mM Tris, pH 8.0, with 90% glycerol) and examined with a fluorescence microscope. As a positive control for TUNEL, embryos resulting from oocytes cultured for 40 h were incubated in RQ1 Rnase-free Dnase (5 µl/50 µl of PBS) for 25 min at 39°C before TUNEL assay.
Immunofluorescence Staining of Kinetochores, Chromosomes, and Microtubules
The methods for staining of kinetochores, chromosomes, and microtubules were the same as our previous study [18]. Briefly, for labeling of kinetochores and chromosomes, cumulus-free oocytes at various culture times or after activation treatment were fixed with 4% formaldehyde in a PHEM buffer (60 mM PIPES, 25 mM HEPES, 10 mM EGTA, 4 mM MgSO4, pH 7.0) for 20 min at room temperature. After being washed three times in PBS, each for 5 min, the oocytes were treated for 10 min in 1% Triton X-100 in PHEM. To suppress nonspecific binding of IgG, the oocytes were incubated with 1% BSA in PHEM for 1 h at room temperature. Then the oocytes were incubated in human nuclear ANA-centromere autoantibody (Cortex Biochem, Sanleandro, CA) diluted 1:500 in PHEM containing 1% BSA overnight at 4°C. After four washes in PBS with 0.05% Tween-20 (PBST), the oocytes were incubated in fluorescein isothiocyanate (FITC)-conjugated goat antihuman IgG (Jackson ImmunoResearch; West Grove, PA) diluted 1:200 in PHEM with 1% BSA for 45 min at room temperature. The oocytes were then washed and stained for 2 min with 10 µg PI/ml in PBST for chromosome examination before being mounted on slides.
For labeling of microtubules and chromosomes, the oocytes were treated with 0.5% Triton-X-100 in PHEM for 5 min, then fixed in 4% formaldehyde in PHEM for 20 min at room temperature. After being washed in PBS, the oocytes were blocked with 1% BSA in PHEM for 1 h at room temperature and incubated in monoclonal mouse anti-
-tubulin (Sigma) diluted 1:10 000 in PHEM with 1% BSA overnight at 4°C. The oocytes were washed three times in PBST and then stained with FITC-conjugated goat anti-mouse IgG (Jackson ImmunoResearch) diluted 1:200 in PHEM with 1% BSA for 45 min at room temperature. After four washes in PBST, the oocytes were finally stained with PI for chromosome examination employing the methods described above.
Confocal Microscopy of Stained Oocytes
After staining, oocytes were washed briefly in PBST and then mounted on slides with antifade solution (0.5% n-propyl gallate in 20 mM Tris, pH 8.0, with 90% glycerol) and examined with a Leica confocal laser scanning microscope (TCS-4D) on the same day.
Immunoblotting for Quantities of -Tubulin, MAPK, BCL2, MAD2, and CENPB in the Oocytes
The immunoblotting procedures were carried out according to the methods reported previously [34]. A total of 200 oocytes from each culture point were collected in SDS sample buffer and heated to 100°C for 4 min. After being cooled on ice and centrifuged at 12 000 x g for 4 min, samples were frozen at –20°C until use. For examination of -tubulin and CENPB, the proteins were separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) with a 4% stacking gel and a 10% separating gel for 2.5 h at 120 V and then electrophoretically transferred onto nitrocellulose membrane for 2.5 h at 200 mA at 4°C. After washing in TBS (20 mM Tris, 137 mM NaCl, pH 7.4) for 30 min, the membrane was blocked for 2.5 h in TBST buffer (TBS with 0.1% Tween-20) containing 5% low-fat milk at room temperature. The membrane was then incubated in mouse anti--tubulin (1:4000 in TBST) or human nuclear ANA-centromere autoantibody (1:4000 in TBST) overnight at 4°C. After being washed three times, each for 10 min in TBST, the membrane was treated in peroxidase (HRP)-conjugated goat anti-mouse IgG (1:3000 in TBST) or HRP-conjugated goat anti-human IgG (1:20 000 in TBST) for 1 h at 37°C.
For MAPK, MAD2, and BCL2 analyses, proteins were separated with a 4–12.5% gel. After being blocked in 5% low-fat milk for 1 h at room temperature, the membrane was incubated with mouse antiactive MAPK antibody (1:500 in TBST), goat anti-MAD2 antibody (1:200 in TBST), or rabbit anti-BCL2 antibody (1:500 in TBST) overnight at 4°C. To check MAPK, the washed membrane was incubated with HRP-conjugated goat anti-mouse IgG (1:5000 in TBST) for 1 h at room temperature. For MAD2, the membrane was treated with HRP-conjugated rabbit anti-goat IgG (1:20 000 in TBST) for 1 h at 37°C. For analysis of BCL2, the membrane was incubated in HRP-conjugated goat anti-rabbit IgG (1:5000 in TBST) for 1 h at room temperature. Finally, the membranes were washed three times in TBST and then the specific proteins were visualized using chemiluminescence detection system. Immunoblot density was determined by the system of Personal Densitometer SI and FragmeNT Analysis software produced by Molecular Dynamics Inc. (Sunnyvale, CA).
Statistical Analyses
For immunofluorescence staining of oocytes, each experiment was repeated four times and at least 15 oocytes were examined each time. The experiments for immunoblotting of various proteins were repeated three times. The experiments for examination of oocyte nuclear stages and oocyte activation with ionophore were repeated four times. All percentage data were subjected to arc-sine transformation before statistical analysis. Data were analyzed by ANOVA.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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As shown in Table 1, when oocytes were cultured for 40 h, most (79.7%) progressed to the M-II stage. These M-II oocytes were designated as young oocytes to be distinguished from aging oocytes. As the culture time was prolonged to 48, 60, and 72 h, the numbers of oocytes at M-II stage were not significantly different from those at 40 h. However, the numbers of degenerated oocytes were increased (from 1.9% to 6.5%) as the culture time was prolonged, although no statistical differences were observed.
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Characteristics of Spindle, Kinetochores, and Chromosomes in Pig Oocytes During Aging
As shown in Table 2 and Figure 1, when oocytes were examined at 40 h of culture, 90.5% of oocytes at M-II stage had a barrel-shaped spindle and the spindles were organized vertically to the cortex of oocytes. The chromosomes were tightly arranged in one line on the spindle equatorial plane, while the kinetochores were aligned in two rows on the outside of the chromosome line (Fig. 1, A and B). As oocytes aged, the proportion of oocytes with normal spindle morphology and/or chromosome alignment decreased from 90.5% at 40 h to 57.1% at 72 h, and accordingly, the oocytes with abnormal spindle morphology and/or chromosome alignment increased from 9.5% at 40 h to 42.9% at 72 h (Table 2). The abnormal spindle morphology and/or chromosome alignment included reduced distance between spindle poles, chromosomes that were not aligned at the spindle equator, some chromosomes stretched out of the chromosome rows, kinetochores not kept in two orderly lines between chromosomes and spindle, and the axis of the kinetochore/chromosome arrangement vertical to the cortex of the oocytes (Fig. 2, G–H).
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Quantification of -Tubulin, Active MAPK, BCL2, MAD2, and CENPB in Oocytes During Aging
As shown in Figure 2, there was no significant difference in CENPB in oocytes cultured for 40–72 h. The quantity of -tubulin (Fig. 3) in the oocytes expressed a gradual reduction tendency as the maturation time was prolonged. However, the levels of active MAPK (Fig. 4), BCL2 (Fig. 5), and MAD2 (Fig. 6) in the oocytes cultured for 40 h were significantly (P < 0.01–0.001) higher than that in the oocytes cultured for 60 and 72 h. A gradual reduction tendency in the levels of MAPK, MAD2, and BCL2 was observed as the oocytes underwent aging.
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Mitochondria Distribution in Pig Oocytes During In Vitro Aging
As shown in Figure 7A, mitochondria were evenly distributed in 95.8–98.4% of oocytes cultured for 40 and 48 h. However, when the culture time was extended to 60 and 72 h, it was found that the mitochondria aggregated to the center of the oocytes (Fig. 7B). The proportion of oocytes with aggregated mitochondria was increased significantly from 1.6% and 4.2% at 40 and 48 h to 65.2% and 69.3% at 60 and 72 h (Fig. 7C), respectively.
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Characteristics of Anaphase II in Pig Oocytes Activated by A23187 During Aging
As shown in Table 3, when oocytes were activated by calcium ionophore and examined 2 h after activation, 16.7– 24.1% of oocytes were found at the anaphase II stage. It was found that 13 out of 14 young oocytes (92.9%) display normal anaphase II, in which chromosomes were separated in the spindle in the direction to two spindle poles (Fig. 8A). Two sets of chromosomes were observed in late anaphase II stage (Fig. 8, A and B) and kinetochore staining was not distinct (Fig. 8B). However, when aged oocytes were activated, the proportion of oocytes with normal anaphase II decreased significantly (P < 0.001) and dropped to 23.8–30.8% after 60–72 h of culture (Table 3). Abnormal anaphase II includes triple or multiple nuclear separation (Fig. 8, C, E, and G). The proportion of oocytes with triple and multiple nuclear separation increased from 7.1–14.3% at 40–48 h of culture to 69.2–76.2% at 60–72 h of culture (Table 3). Some chromosomes remained near the spindle equator during the anaphase II stage, while the majority of sister chromatids moved to the spindle poles (Fig. 8D). Some chromosomes were clearly ungrouped from the others or the separation was delayed. Therefore, single kinetochore staining was observed in these oocytes (Fig. 8, D and F).
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Fragmentation of Activated Oocytes by Calcium Ionophore
As shown in Figure 9, when the oocytes were examined 24 h after activation, activated oocytes cultured for 40–48 h had 2 cells (Fig. 9A) and fragmentation was observed in 2.8–4.3% of the embryos. However, 50–55.3% of the activated oocytes cultured for 60–72 h had fragmentation with two blastomeres and more than 25–30% of fragmentation (Fig. 9B). As the oocytes were examined at 48 h after activation, fragmentation increased in all groups, but with significantly higher (P < 0.001) fragmentation rates in the oocytes cultured for 60–72 h (67.9–76.6%) than in the oocytes cultured for 40–48 h (12.5–13%) (Fig. 9C).
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Apoptosis in Parthenogenetic Embryos
As shown in Figure 10, apoptosis signals were observed in 16 out of 18 positive control embryos (Fig. 10A, from three replications). When the embryos were examined on Day 2, no positive signal was observed in all embryos (15 embryos at each group) irrespective of the embryos having resulted from oocytes with different culture times. When the embryos were cultured for 5 days, there was no positive signal in the embryos resulting from the oocytes cultured for 40 h (0/18 embryos, n = 3) and 48 h (0/16 embryos, n = 3). However, positive signals were observed in the embryos resulting from the oocytes cultured for 60 h (Fig. 10C; 3/16 embryos, n = 3) and 72 h (Fig. 10D; 4/17 embryos, n = 3).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The kinetochore plays a key role in spindle formation, chromosome alignment, and correct onset of anaphase in mitosis and meiosis [10]. When the kinetochore's function is blocked in the oocytes, the chromosome movement and separation become random, thereby producing aneuploidy [38]. In the present study, although no difference in the quantification of CENPB and tubulin was observed between young and aged oocytes, the dynamics of kinetochores in aged oocytes were different from that in young oocytes, suggesting that the ability of kinetochores to attach microtubules and recruit other transient proteins was affected during aging [39]. Because CENPB and tubulin are constitutive proteins in the kinetochores and microtubules, their amount may not be affected by aging or early aging as examined in the present study. However, their functions appear affected, such as abnormal spindle formation in the aged oocytes. These results may also indicate that functional changes occur before amount changes in these proteins. The distance between sister kinetochores became larger in aged oocytes; this was in accordance with the fact that the level of premature centromere segregation gradually increased during oocyte aging in some mammals [4, 9, 20, 21, 24] and may be related to the precocious degradation of cohesion between sister chromatids in aged oocytes [2, 4, 24]. A recent report showed that premature separation of sister kinetochores during meiosis I resulted in lagging chromosomes during meiosis II in maize [40] and produced aneuploidy [41]. In aged porcine oocytes, some chromosomes were also observed left near the spindle equator during the anaphase II stage, and the proportions of oocytes with triple or multiple separation gradually increased with oocyte aging from 7.1% in the oocytes at 48 h to 76.2% in the oocytes at 72 h.
As one of the transient checkpoint proteins in the kinetochores, MAD2 is able to check spindle deformation, chromosome disorder, and chromosome attachment tension and delay anaphase onset [11, 12, 24, 42]. It has been found that absence of MAD2 reduced the accuracy of chromosome segregation in meiosis [17] and the mRNA level of human MAD2 was lower in aged oocytes than in young ones [13], suggesting that MAD2 may also participate in spindle checkpoint during meiosis as that in mitosis. We also found that expression level of MAD2 increased from GV to M-II stage and reached the peak at the M-II stage in pig oocytes during meiotic maturation (unpublished data). However, in the present study, we found that MAD2 expression reduced in the aged oocytes and reduced MAD2 expression may not have been able to check abnormal chromosomes attachment and distribution in the oocytes. Taken together, these results indicate an age-dependent loss in the function of spindle checkpoint system.
The reasons for the impaired surveillance system during oocyte aging are still not clear. Some crucial molecules of protection are affected in aged oocytes [3]. Our results showed that the amount of MAPK decreased as oocytes aged. MAPK is a family of Ser/Thr protein kinases that are widely distributed in eukaryotic cells. MAPK plays pivotal roles in regulating the meiotic cell cycle progression of oocytes [43]. After GVBD, MAPK is involved in the regulation of microtubule organization and meiotic spindle assembly and ensures asymmetric division of meiosis I. The activation of this kinase is essential for the maintenance of metaphase II arrest, while its inactivation is a prerequisite for pronuclear formation after fertilization or parthenogenetic activation [26, 43, 44]. In the present study, as shown in previous studies [43], two isoforms of MAPK, extracellular-regulated kinase (ERK) 1 and ERK2, are expressed in both young and aging oocytes. However, their activities were significantly reduced in the aging oocytes, which was consistent with other mammalian oocytes [1, 4, 25]. Because MAPK is responsible for progression from the GV to the M-II stage and also is important for oocytes to remain arrested at the M-II stage [1, 22], high levels of MAPK are necessary for the chromosomes to orderly align at the spindle equator [1]. A low level of MAPK seems to be associated with a high level of fragmentation formation and apoptosis [present study, 22]. Our results suggest that a low level of MAPK may cause instability of chromosomes in the spindles and may alternate the precious relationship between microtubules and chromosomes.
BCL2 is crucial in preventing apoptosis in cells. Overexpression of BCL2 prevents cells from undergoing apoptosis in response to a variety of stimuli [27]. One possible role of BCL2 in preventing apoptosis is to block cytochrome C release from mitochondria. Reduced BCL2 is also associated with dysfunctional mitochondria [1, 27–29]. In the present study, we found that oocyte aging caused aggregation of mitochondria in the oocytes and reduced BCL2 expression. These results suggest that mitochondrial aggregation and a reduced level in BCL2 are two traits related to oocyte aging [30].
MAPK is also important for the spindle checkpoint in Xenopus egg extracts [26]. Reduced MAPK accelerated the degradation of the BCL2 in mouse and human oocytes [1, 3]. All these results indicate that reduced MAPK and BCL2 in aged porcine oocytes directly or indirectly change the spindle checkpoint signal and alter spindle structure and the connection between chromosomes and microtubules. In the present study, we found that not only the activities of MAPK but also the expression levels of MAD2 and BCL2 were reduced during oocyte aging, which results in abnormal meiosis II initiation. The present study indicates that the optimal fertilization time of pig oocytes is about 40 h and no more than 48 h. After 48 h, the activities of MAPK and the expression of BCL2 and MAD2 were reduced and significantly higher rates of oocytes with abnormal meiosis II were observed. A higher number of aged oocytes were activated as compared with young oocytes [6, 21, 33], indicating that the activities of some components associated with M-II arrest, such as MAPK, were reduced. However, it is possible that, in the aged oocytes, other surveillance systems responsible for accurate progression of the cell cycle are affected [9, 24]; thus, normal embryo production may be decreased when aged oocytes are used.
In the present study, when calcium ionophore, which can mimic sperm to induce porcine oocyte activation [45, 46], was used to examine meiosis II and subsequent development of activated oocytes, we found that abnormal anaphase II was observed in most oocytes cultured for 60–72 h and the activated oocytes also showed significantly higher degrees of fragmentation. It was suggested that fragmentation is an early indicator of apoptosis [47]. However, apoptosis was not observed in pig embryos produced by in vitro fertilization and nuclear transfer until Day 5 of culture [47]. The same results were observed in the present study and we found that the apoptosis signal was not observed in the embryos from aging oocytes before Day 5. However, after Day 5 of culture, significantly more embryos showed apoptosis signal in the embryos resulting from the aged oocytes (60–72 h) as compared with young oocytes (40– 48 h). These results, together with the protein expression levels and MAPK activities examined in the aging oocytes, indicate that oocyte aging increases embryo fragmentation and apoptosis.
In conclusion, our present study indicates that postovulatory aging reduces the expression of BCL2. Due to the low level of the antiapoptotic protein BCL2, apoptosis is not prevented in the oocytes, which in turn causes (directly or indirectly) reduced activities of MAPK and expression of MAD2. Reduced MAPK activity may induce an abnormal relationship between microtubules and chromosomes and reduced MAD2 level may not be able to detect these abnormalities in spindles and chromosomes; thus, abnormal meiosis II is initiated when oocytes are activated. To produce normal embryos, optimal timing for insemination of oocytes is important. From the present study, it is suggested that 40 h of culture and no more than 48 h is the optimal insemination period for pig oocytes matured in vitro. Aggregated mitochondria distribution in the oocytes and fragmentation degree in the early embryos may be the morphological markers for oocyte aging.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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2 Correspondence. FAX: 86 10 6265 0042; wangweihua11{at}yahoo.com
Received: 14 April 2004.
First decision: 10 May 2004.
Accepted: 16 September 2004.
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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