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Research Article |
Department of Biomedical Sciences and Technologies,3
Center for Assisted Reproduction,4
Department of Basic and Applied Biology,5
Department of Experimental Medicine,6 University of L'Aquila, 67100 L'Aquila, Italy
ABSTRACT
To elucidate molecular mechanisms underlying oocyte senescence, we investigated whether oocytes from female mice of advanced reproductive age exhibit a precocious postovulatory aging that, in turn, may be responsible for the precocious activation of an apoptotic program. During a 9-h in vitro culture, the frequency of oocytes showing MII aberrations, spontaneous activation, and cellular fragmentation increased in old oocytes (P < 0.05), whereas it did not change in the young group. In old oocytes, the activities of MPF (a complex of the cyclin-dependent kinase cdc2 and cyclin B1) and MAPK (mitogen-activated protein kinase) decreased precociously, showing a first drop as early as 3 h after the beginning of in vitro culture (P < 0.05). Immunoblotting and immunocytochemical analysis revealed that, in oocytes of the old group, reduction of BCL2 expression at protein level occurred earlier than in the young group (P < 0.05) and was not associated to the loss of BCL2 transcripts detected by RT-PCR. These changes are followed by an abrupt increase of the rate of TUNEL-positive oocytes after 24 h of culture to a value of 67% ± 6%. Exposure of young oocytes to 20 µM roscovitine or 20 µM U0126, specific inhibitors of MPF and MAPK, resulted in the decreased percentage of oocytes showing positive immunostaining for BCL2 and in an increased rate of DNA fragmentation. Present results suggest that the developmental competence of oocytes ovulated by aging mice may be negatively influenced by a downregulation of MPF and MAPK activities that in turn induces the activation of a proapoptotic signaling pathway.
aging, apoptosis, fertilization, kinases, oocyte development
INTRODUCTION
Aging-related decline in female fertility is a common feature in females from most mammalian species, including humans. In women, reproductive senescence becomes clinically significant by the late 30s, ending in complete loss of fertility by the late 40s, even though there are still primary oocytes within the ovary [1]. Although age affects nearly every aspect of female reproduction, the oocyte is believed to be a major locus of reproductive aging. Besides reduction in terms of number of follicles, age is also responsible for a reduced oocyte quality, which refers to the oocyte ability to be fertilized and undergo normal embryo development [2]. The most direct evidence for oocyte senescence in humans is the finding that oocytes donated from young to older women in assisted-reproduction programs abrogate the effects of age on fertility [3]. However, how age affects the inherent features of the oocyte that determine its developmental potential is still poorly understood. It has been well recognized that oocyte aneuploidy is significantly increased with maternal age, mostly due to errors in chromosome segregation at the first meiotic division [4]. Also, anomalies in the meiotic spindle have been described and most likely contribute to the chromosome misalignment evidenced at both metaphase I and metaphase II [5]. Although these errors are passed on to monosomic and trisomic embryos and compromise embryo viability, an increase of euploid embryo abortuses has been observed during female aging. Moreover, in mice, the relationship between oocyte aneuploidy rate and aging-associated decrease in fertility remains still controversial [6–9], supporting the idea that, beyond nuclear defects, abnormalities in the ooplasm may also impact on embryo development during reproductive aging.
In human oocytes, cytoplasmic aging has been associated with decline of mitochondrial functions, reduced content of transcripts from genes participating in spindle assembly checkpoint and increased frequency of apoptosis [10–12]. Similar age-related changes have been described in the mouse model, where a low fertilization competence has been related to oocyte abnormalities, including chromosome scattering, chromosome decondensation, alteration of gene expression pattern, and high rates of DNA fragmentation [13–15]. It is important to mention that some oocyte defects associated with maternal aging seem to resemble changes occurring during postovulatory aging in oocytes from young females [16, 17]. It is well known that the ovulated mature oocyte has a short life-span associated with a time-dependent decrease of its normal developmental competence after ovulation, so that delayed fertilization frequently fails or even causes long-term effects on conceptuses [16, 18]. Beyond cellular modifications, changes occurring after ovulation include the spontaneous decrease of the activities of critical enzymes involved in the maintenance of the meiotic arrest of oocytes at metaphase II (MII) [19–21]. MII-arrested oocytes exhibit a characteristic high activity of maturation promoting factor (MPF), a complex of activated cdc2 kinase (CDK1) and cyclin B that maintains MII state [22]. In addition to MPF, MII-arrested oocytes are also characterized by high activity of mitogen-activated protein kinase (MAPK)3/MAPK1 (also known as ERK1/ERK2). MAPK is a component of the cytostatic activity (CSF) that maintains MII arrest [23]. In order for fertilization to succeed, the oocytes must keep high MPF and MAPK activities and inactivate them, upon sperm penetration, in a coordinated, step-wise fashion. Therefore, downregulation of the molecular mechanisms that control MII can compromise normal entry into embryonic cell cycle following fertilization. A further biochemical change associated with postovulatory aging is the decreased expression of BCL2, an antiapoptotic regulator of the BCL2 family [21, 24]. BCL2 protein localizes in the outer mitochondrial membrane, nuclear membrane, and endoplasmic reticulum and regulates mitochondrial function, Ca2+ homeostatis, and cell survival [25, 26]. A reduced BCL2 activity may be responsible for the altered balance between survival and death-promoting factors, leading to programmed cell death in oocytes skipping or delaying sperm encounter, as occurs in somatic cells [27].
To elucidate molecular mechanisms underlying oocyte senescence, we investigated whether oocytes from females in advanced reproductive age exhibit a precocious postovulatory aging that, in turn, may be responsible for the precocious activation of an apoptotic program. In this study, we used the mouse model to compare the time-course of biochemical and cellular changes occurring during postovulatory in vitro culture in oocytes obtained from young and old animals. Then, to elucidate the possible role of MPF and MAPK in regulating oocyte survival, we studied whether treatment of oocytes from young mice with MPF and MAPK inhibitors can induce a decrease of BCL2 protein and an increased rate of DNA fragmentation.
MATERIALS AND METHODS
Oocyte Collection and Treatment
CD-1 mice were obtained from Charles River Italia s.r.l. (Calco, Italy). Animal care and experiments were carried out in accordance with the Guide for Care and Use of Laboratory Animals (1996). At the age of 4–8 wk (young mice) and 48–52 wk (old mice), females were superovulated by intraperitoneal injection of 10 IU of eCG (Folligon; Intervet-International, Boxmeer, Holland) and 10 IU of hCG (Profasi HP 2000; Serono, Roma, Italy) 48 h apart. MII-arrested oocytes were released at 15 h post-hCG from the oviducts into the medium M2 (Sigma, St. Louis, MO). Cumulus cells were dispersed by a brief exposure to 0.3 mg/ml hyaluronidase (Sigma) and oocytes were pooled and randomized before distribution into the experimental groups. Oocytes were processed immediately or cultured at 37°C, 5% CO2 in M16 medium (Sigma) for various times. For brevity, oocytes from animals of the two groups of age are called young oocytes and old oocytes. The mean ± SEM of oocytes ovulated per female were the following: 20.86 ± 1.76 young oocytes and 8.84 ± 1.18 old oocytes (P < 0.001, Student t-test).
MPF and MAPK Activity Assay
For kinase activity assays, nonfragmented oocytes (five for each experimental group, two replicates) were transferred to centrifuge tubes in 2 µl of collection buffer (PBS containing 1 mg/ml polyvinil alcohol, 5 mM EDTA, 10 mM Na3VO4, 10 mM NaF), immediately submerged in liquid N2, and stored at –80°C until the kinase assay was performed. The frozen oocytes were thawed in 4 µl of reaction mixture containing ß-glycerophosphate 54 mM, paranitrophenylphosphate 14.5 mM, morpholenepropanesulfonic acid 24 mM, MgCl2 14.5 mM, EGTA 14.5 mM, EDTA 0.12 mM, DTT 1 mM, leupeptin 1 µg/ml, aprotinin 1 µg/ml, ML-9 10 µM, genestein 75 µM, chimostatin 1 µg/ml, trypsin-chimotrypsin inhibitor 1 µg/ml, PKI 2,4 µM, 50 µCi/ml 
-[32P] ATP (Amersham Pharmacia Biotech, Uppsala, Sweden). To the reaction mixture were added histone H1 type III-S and myelin basic protein (MBP), specific substrates, respectively, of MPF and MAPK. The reagents for the collection buffer and the reaction mixture were obtained from Sigma. After 30 min at 37°C, assays were stopped by adding sample buffer 1:1 [28] and proteins in oocytes were denaturated by heating the reaction mixture at 95°C for 5 min. Samples were electrophoresed on 12% SDS polyacrylamide gels that were subjected to autoradiography. Autoradiographs were scanned and densitometric values for each treatment group were measured by the Adobe Photoshop software (Mountain View, CA). Values in each autoradiograph were standardized by setting oocytes from young mice at the beginning of in vitro culture (time 0) as 100% activity.
Analysis of Chromosomal Distribution, DNA Organization, and Cellular Morphology
The morphological appearance of oocytes was examined under a stereoscope microscope. Degenerated oocytes were discarded and the presence of oocytes with cellular fragmentation were evaluated. Chromosomal distribution and DNA organization were evaluated by DNA staining. Briefly, oocytes immediately recovered or cultured for different times in M16 medium at 37°C, 5% CO2 were fixed in 3.7% paraformaldehyde in PBS and then incubated with 3 µg/ml Hoechst 33342 (Sigma). Oocytes were mounted onto a slide under a coverslip in a mounting medium (glycerol 50%) and observed under an epifluorescence microscope with a 100x objective (Leitz Dialux with filter BP 340–380). A filter with an excitation wavelength of 330–380 nm and barrier filter of 420 nm was used to detect chromatin of oocytes stained by Hoechst 33342. Anomalies in the distribution of chromosomes in the MII spindle and DNA organization were classified as follows: 1) chromosome scattering if one or more chromosomes were separated from the metaphase plate and 2) chromosome decondensation if chromosomes showed varying degrees of DNA condensation. Oocytes showing anaphase-II or telophase-II configurations were considered activated.
Reverse Transcription and Polymerase Chain Reaction
Oocytes were washed in PBS containing 3 mg/ml polyvinylpyrrolidone (PBS-PVP) and transferred into a 0.6-ml tube in a volume less then 2 µl. Oocytes were immediately subjected to thermolysis and reverse transcription (RT) by means of cell-to-cDNA kit (Ambion Inc., Austin, TX), according to the manufacturer's instructions. Polymerase chain reaction (PCR) analyses were carried out in 50 µl containing the cDNA equivalent of 15 oocytes, PCR buffer 1x (Perkin Elmer Life And Analytical Sciences, Boston, MA), 1.5 mM MgCl2, 0.2 mM of each dNTP, 0.2 µM of each primer, and 1.25 U of Taq polymerase (all products from Perkin Elmer). Actb (beta-actin) was selected as a housekeeping gene. Primer sequences for beta-actin were derived from the published complementary DNA sequences provided by GenBank (Accession GI 38648901): 5'GTACCCCATTGAACACGGCA and 3'GCACAGTGTGGGTGACCCC. The beta-actin cDNAs were amplified for 35 cycles with the following profile: 94°C for 10 min for 1 cycle; 94°C for 50 sec, 60°C for 50 sec, 72°C for 1 min for 35 cycles; 72°C for 10 min. Only cDNA samples positive for beta-actin were processed for Bcl2 cDNA amplification. Primers sequences for BCL2 were derived from the published complementary DNA sequences provided by GenBank [29] (Accession M16506, L31532): 5'TACCGTCGTGACTTCGCAGAG and 3'GGCAGGCTGAGCAGGGTCTT. The Bcl2 cDNAs were amplified for 40 cycles with the following profile: 96°C for 1 min for 1 cycle; 96°C for 1 min, 56°C for 1 min, and 72°C for 15–25 sec for 2 cycles; 94°C for 1 min, 59°C for 1 min, 72°C for 15–25 sec for 18 cycles; then 94°C for 1 min, 59°C for 30 sec, 72°C for 15–25 sec plus 1 sec per cycle for 20 cycles. After amplification, the RT-PCR products were separated by agarose (1.5%) gel electrophoresis, stained by ethidium bromide, and visualized under UV.
Western Blot Analysis
Three hundred oocytes for each experimental group were lysed in sample buffer. As a positive control, microsomes from mouse ovaries were used as previously described [24]. Briefly, tissue samples were homogenized in 0.3 M sucrose, 1 mM EDTA, 1 mM 2-mercaptoethanol, 0.1 mM PMSF, 2 µg/ml pepstatin, 2 µg/ml leupeptin, 50 mM Tris-HCl (pH 8.0; all chemicals from Sigma) and centrifuged at 2500 x g for 15 min. The supernatant was centrifuged at 100000 x g for 30 min. The microsomal precipitates were resuspended in 0.3 M sucrose, 1 mM EDTA, 1 mM 2-mercaptoethanol, 50 mM Tris-HCl (pH 8.0) and electrophoresed with oocytes samples onto 12% SDS-polyacrylamide gels. The resolved proteins were blotted onto nitrocellulose membranes, which were repeatedly washed in 10 mM Tris-HCl (pH 8), 150 mM NaCl, 0.05% Tween-20 (TBS-T). The membranes were then blocked in TBS-T supplemented with 6% (w/v) nonfat dry milk and then probed for BCL2 by incubation overnight at 4°C with an anti-BCL2 rabbit polyclonal antibody (1:400; Calbiochem, La Jolla, CA). After several washes in TBS-T, the membranes were incubated in a 1:3000 dilution of goat anti-rabbit secondary horseradish peroxidase-coupled antibody (Santa Cruz Biotechnology Inc., Santa Cruz, CA) in TBS-T for 1 h. The same blot was first probed with antibody to BCL2, then stripped and reprobed with anti-ACTB (beta-actin) rabbit polyclonal antibody (1:500; Sigma). The specific antibody binding was detected by using the enhanced chemiluminescence system according to the manufacturer's instructions (Amersham Biosciences, Little Chalfont, England). Autoradiographs were scanned and semiquantification of immunoreactive BCL2 was done using Adobe Photoshop 4.0. The intensity of the BCL2 bands in each autoradiograph was expressed relative to the intensity of the band in young oocytes processed at the beginning of in vitro culture (time 0).
Confocal Microscopy
Oocytes from each experimental group were exposed for about 1 min to Tyrode solution [30] to remove zona pellucida. Zona-free oocytes were fixed as above described, permeabilized for 5 min in 0.1% TritonX-100 in PBS-PVP, and washed in blocking buffer (PBS-PVP, 0.1% BSA, 0.01% Tween-20). Permeabilized oocytes were incubated with the BCL2 polyclonal antibody (1:100) for 1 h at room temperature, followed by incubation with 1:800 dilution of a secondary anti-rabbit IgG conjugated with Cy-3 (Molecular Probes, Eugene, OR). In each experiment, negative control samples in which the primary antibody was omitted were also evaluated. After washing in blocking solution, oocytes were treated with an antibleaching solution (Slow-Fade; Molecular Probes), mounted on a glass slide, and sealed with clear nail polish. Oocytes were observed using an Olympus FV500 confocal microscope equipped with a laser Green Helium-Neon (543 nm) and a PLAN-APO 60/1.4 oil objective (Olympus Optical Co. Gmbh, Hamburg, Germany).
Detection of Apoptosis by TUNEL Assay
DNA fragmentation was detected by TUNEL method by means of in situ cell death kit (Roche Diagnostic Gmbh, Mannheim, Germany). The oocytes were fixed and permeabilized as above described. Positive control was represented by young oocytes treated with 50 U/ml DNase. The experimental groups and positive control group were incubated at 37°C for 1 h in 25 µl of TUNEL reaction mixture containing dUTP-fluorescein isothiocyanate (FITC), terminal deoxynucleotydil transferase (TdT) enzyme, and reaction buffer. Negative control was represented by young oocytes incubated in TUNEL reaction mixture without dUTP-transferase enzyme. After washing three times in PBS-PVP, the oocytes were incubated in Hoescht 33342 at room temperature and mounted on slides. The samples were observed using an epifluorescence microscope at 400x magnification. Two standard filter sets were used, a filter with an excitation wave length of 450–490 nm and a barrier filter of 520 nm were used to detect FITC alone. A filter with an excitation wavelength of 330–380 nm and a barrier filter of 420 nm were used to detect the nuclear status of oocytes stained with Hoechst 33342. Oocytes were considered TUNEL-positive when Hoechst33342 staining was associated with FITC staining.
Roscovitine and U0126 Treatments
Roscovitine and U0126 (Calbiochem) were prepared as 25 mM stock solutions in DMSO and stored at –20°C until use. Oocytes from young mice were transferred in M16 medium containing 20 µM roscovitine or 20 µM U0126. For control treatments, oocytes were cultured in M16 containing 0.05% DMSO.
Statistical Analysis
Values were compared using one-way ANOVA and Student t-test. Multiple comparison of values was carried out using Student-Newman-Keul test. All analyses were performed using the SigmaStat software (Jandel Scientific Corporation, San Rafael, CA). Statistically significance was reached at P < 0.05.
RESULTS
MII Aberrations, Spontaneous Activation, and Cellular Fragmentation in Old Oocytes During Postovulatory In Vitro Culture
In the first set of experiments, we compared, in young and old oocytes, the ability to maintain a normal MII configuration during postovulatory in vitro culture. To this end, we monitored the presence of oocytes with abnormal MII or spontaneously activated after different times of in vitro culture. As shown in Figure 1A, when cells were scored for the presence of abnormal MII at the beginning of in vitro culture, old oocytes exhibited a frequency of this pattern higher than that monitored in young oocytes (P = 0.022). Subsequent in vitro culture resulted in an abrupt increase of this alteration in old oocytes (time 0 vs. time 3 h, P = 0.014), whereas it had only a slight, but not significant, effect on young oocytes (P = 0.3). Moreover, in the old group, the percentage of oocyte activation increased from a value of 7.38 ± 4 to 22.75 ± 2.12 after 9 h of culture (P = 0.04), a value about twofold higher than that observed in the young group at the corresponding time point (Fig. 1B). By contrast, the frequency of oocyte activation in the young group did not change significantly after 9 h of in vitro culture (Fig. 1B). When cellular fragmentation was assessed through morphological analysis under a phase-contrast microscopy, old oocytes revealed greater values than those observed in the young group at all time points investigated. Also, unlike the young group, where the frequency of cellular fragmentation increased slowly during in vitro culture, in old oocytes, a 9-h in vitro culture resulted in a significant increase of this pattern from 21% to 35% (P = 0.017; Fig. 1C).
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Changes of MPF and MAPK Activities in Old Oocytes During Postovulatory In Vitro Culture
We compared in old and young oocytes the time course of the spontaneous decrease of the enzymatic activities of MPF and MAPK that occurs after ovulation. The activities of MPF and MAPK, evaluated as oocyte ability to phosphorylate histone H1 and MBP substrates, respectively, were monitored following different times of in vitro culture. As shown in Figure 2, old oocytes at the beginning of in vitro culture exhibit only a slight decrease in the activities of both kinases when compared with the young group. However, as in vitro culture progressed, MPF and MAPK activities in old oocytes were significantly lower than those detected in the young group at any time point investigated (P < 0.05).
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In young oocytes, relative MPF activity decreased significantly after 9 h of culture (6 h vs. 9 h, P = 0.020), whereas in old oocytes, a loss of this activity could be detected as early as 3 h after the beginning of in vitro culture (0 h vs. 3 h, P = 0.047; Fig. 2A). After 24 h, MPF activity reached in old oocytes a value about half of that monitored in the young group.
As shown in Figure 2B, a first drop in relative MAPK activity occurred after 6 h of culture in young oocytes (3 h vs. 6 h, P = 0.048) and after 3 h of culture in old oocytes (0 h vs. 3 h, P = 0.025), where a second drop could be observed at 9 h (6 h vs. 9 h, P = 0.025). MAPK activity decreased further in this group, ending at 32% residual value after 24 h of culture, when young oocytes exhibited a value about twofold higher.
Effect of Postovulatory In Vitro Culture on BCL2 Expression and DNA Fragmentation in Old Oocytes
It is known that expression of the survival factor BCL2 at RNA and protein level is spontaneously impaired during postovulatory in vitro culture in the presence of constant levels of Bax transcripts [24]. Then, it is possible that the susceptibility to spontaneous apoptosis of old oocytes [15] correlates with a reduced expression of BCL2. To accomplish this, we monitored the presence of Bcl2 mRNA in oocytes from young and old mice, soon after their collection and after 9 h of in vitro culture. RT-PCR experiments showed that old oocytes, similarly to young oocytes, still maintained detectable levels of Bcl2 transcripts at least until 9 h from the beginning of in vitro culture (Fig. 3).
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Then, protein levels of BCL2 were monitored by means of immunoblotting and immunocytochemistry. Results from Western blot analysis revealed that the levels of BCL2 protein, at the beginning of culture period, were not significantly different comparing old and young oocytes of the groups of age. However, after a prolonged in vitro culture, the level of BCL2 protein in the young group decreased significantly, to about half of the initial level, whereas in the old group, it became undetectable (Fig. 4). This trend was more clearly evident when single oocytes were analyzed by immunofluorescence with the same primary antibody used in immunoblot analysis. Confocal analysis revealed that oocytes exhibited a diffuse punctuate staining. However, by using the same confocal settings, a different intensity of fluorescence could be observed. As shown in Figure 5A, three different types of immunostaining were established: type I, high staining; type II, moderate staining; type III, negative staining (equivalent to negative control). Data in Figure 5B show that the frequency of young and old oocytes showing type I and type II staining (BCL2-positive staining) decreased more rapidly in the old group, exhibiting after 9 h of culture about 50% ± 11% of BCL2-positive oocytes (young vs. old, P = 0.027). Conversely, at this time, the young group still maintained a value not significantly different from that monitored at the beginning of the culture period (P = 0.359). After 24 h, the percentage of BCL-positive oocytes decreased from 86 ± 6 to 17 ± 9 (P = 0.004) in the old group and from 100% to 61% ±7% (P = 0.012) in the young group.
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To relate these changes with the activation of an apoptotic pathway, we monitored DNA fragmentation during in vitro culture in the two groups of age. We found that all fragmented oocytes displayed FITC-fluorescent chromatin in at least one oocyte fragment (Fig. 6A). Unlike the young group, where the rate of TUNEL-positive oocytes increased with a slow time course from 5% ± 4% to 18% ± 4%, in the old group, most of oocytes entered the final stage of apoptosis by 24 h, when the percentage of TUNEL-positive oocytes increased from to 14% ± 5% at time 0 to 67% ± 6% (Fig. 6B).
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Effects of MPF and MAPK Inhibitors on MII Configuration, BCL2 Protein Level, and DNA Fragmentation in Young Oocytes
We tested the hypothesis that perturbation of MPF and MAPK activities in oocytes from young mice may cause destabilization of metaphase arrest and chromosomal configuration and a decrease of BCL2 protein. To this end, we have used U0126, a specific inhibitor of the MAP kinase cascade [31], and roscovitine, an inhibitor of CDK1 [32], to negatively affect MAPK and MPF activities in young oocytes, as previously described [33]. To this end, we exposed oocytes from young mice to low doses of the inhibitors (20 µM roscovitine or 20 µM U0126) for 9 h. After 3 and 9 h from the beginning of incubation, oocytes were scored for the presence of abnormal MII configuration. After 9 h of culture, about 40% of U0126-treated oocytes exhibited abnormal MII configuration related to the presence of chromosomes scattered and/or decondensed. Conversely, the percentage of roscovitine-treated oocytes with abnormal MII configuration was not significantly different from that of the control group (Fig. 7A). As expected, both the inhibitors induced a low but significant activation response (Fig. 7B).
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As shown in Figure 8A, after 9 h from the beginning of treatments, both the inhibitors were found to induce a significant reduction in the rate of oocytes positive for BCL2, reaching a value of 44 ± 5 and 33 ± 5 in the roscovitine and U0126 groups, respectively. These values declined further after 24 h of culture to 16% ± 9% and 13% ± 2% in the oocytes treated with U0126 or roscovitine, respectively. Both the inhibitors induced an increase in the rate of TUNEL-positive oocytes that, after 24 h of culture, reached a value of 60% ± 2% in the roscovitine group and a value of 53% ± 5% in the U0126 group (Fig. 8B).
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DISCUSSION
The main point about the observations reported in the present article is the following: postovulatory aging exerts a deleterious effect on oocytes ovulated from old mice. We found that, in old oocytes cultured in vitro after ovulation, both MPF and MAPK activities decreased more rapidly than in young oocytes. In parallel, we observed a more rapid increase of abnormal MII, spontaneous oocyte activation, and cellular fragmentation. This reduced ability to control MII state seems to have a strong influence on survival of old oocytes after ovulation. An increased susceptibility to fragmentation and cell death during reproductive aging has been previously described in both human and mouse oocytes [12, 15]. In our study, we found that accelerated BCL2 degradation and activation of apoptotic pathway are both associated with an accelerated decline of meiotic kinases. Moreover, by comparing results in Figures 1 and 6, it turns out that, in the old group, massive entry in the final stage of apoptosis occurred after nearly all the oocytes had lost their normal MII configuration. This temporal correlation cannot be observed in young oocytes, where postovulatory changes occurred more slowly and gradually. It is worth noting that this comparison between oocytes with different susceptibility to apoptosis is the first evidence that meiotic kinases play an important role in the regulation of oocyte survival after ovulation, as in the model proposed by Fissore et al. [17].
As reproductive aging is associated with mitochondrial dysfunctions [10], it is reasonable to conjecture that the activities of meiotic kinases or their upstream regulators decline as a result of a reduced energy supply and/or oxidative stress, as already reported [34, 35]. Nevertheless, precocious inactivation of MPF and MAPK may reflect an aging of the old oocyte before ovulation due to an accelerated meiosis I or to an asynchrony between meiosis and ovulation. Indeed, old mouse oocytes during in vitro maturation progress faster than young oocytes to first anaphase and to metaphase II [6]. This behavior could be related to defects in the meiotic machinery, as suggested by the finding that old oocytes exhibited a reduced amount of transcripts involved in cell cycle, as compared with young oocytes [14]. Moreover, a loss of coordination between the events leading to ovulation may be caused by age-associated raised basal LH level during the follicular phase that could trigger meiosis resumption before the LH surge [16].
In the present study, we show that age-related changes of BCL2 expression at protein level were not accompanied by the loss of Bcl2 transcripts. This finding further supported the view that BCL2 degradation could be due to loss of posttranslational signals that occurred in the old oocyte more rapidly than in the young oocyte. Several works in somatic cells provide evidence that survival-promoting signals targeting BCL2 may come from the cell-cycle machinery that control M phase. BCL2 contains several consensus protein kinase sites and its phosphorylation universally occurs at the G2-M phase of the cell cycle [36]. Phosphorylation of BCL2 at residues Thr-56, Thr-74, and Ser-87 prevents proteasome targeting and confers resistance against the induction of apoptosis [37]. MAPK is believed to play a major role in this process because dephosphorylation of the MAPK site Ser-87 within BCL2 triggers its ubiquitin-dependent degradation [38]. Moreover, the observation that mitotic BCL2 phosphorylation is inhibited by roscovitine suggests that CDK1 or CDK2 are involved in this process [39].
In the present study, we found that drugs that inhibit MPF and MAP kinase activities seem to mimic an aging effect because young oocytes undergo morphological and biochemical changes similar to those observed in old oocytes. We used roscovitine or U0126 at low doses to avoid the abrupt decrease in the activity of these kinases described when oocytes are exposed to concentrations of these drugs associated with a high rate of oocyte activation [33]. As expected, both the inhibitors induced a low but significant activation response and, in oocytes exposed to U0126, we also observed an increased percentage of inactivated oocytes with chromosomes scattered or decondensed. Moreover, both the inhibitors induced a precocious decrease in BCL2 protein and a significant increase in the rate of DNA fragmentation. Because we did not determine the activities of meiotic kinases, we cannot correlate the extent of downregulation of kinase activity with the effects on the oocytes. Furthermore, we are not able to identify separate roles for MPF and MAPK because these kinases modulate their activities with each other [33]. It is likely that both MPF and MAPK participate in survival-promoting signaling in the MII oocyte. Indeed, in porcine oocytes, drugs preventing spontaneous MPF decrease can prevent cellular fragmentation [40] and, in amphibian oocytes, a MAPK pathway protects them from apoptosis [41]. This conclusion is in accordance with the view that the MII state confers protection from apoptosis, increasing survival chances after ovulation [42]. Moreover, it can be speculated that MAPK is required to exert a protective role also upon release of MII arrest by accompanying the fertilized oocytes until the entry in the first embryonic cell cycle.
Beyond downregulation of MPF and MAPK, many factors can influence the oocyte developmental competence and survival after ovulation [43, 44]. Therefore, here we suggest that the cross-talk between meiotic kinases and BCL2 may play an important role in the deadly decision in oocytes where a survival disequilibria already exists. This could be the case of oocytes from aging females, where mitochondrial dysfunction and oxidative stress have arisen during their prolonged stay in the ovary and/or from a compromised oxygen supply [45]. Oxidative stress damage has been detected in the resting oocyte and follicle pool of the aged ovary [46] and reduced antioxidant defenses have been detected in the mature oocyte [47], as well as in the follicular environment of the periovulatory follicle [48].
In conclusion, the present study provides evidence that oocytes ovulated by aging females exhibit a reduced temporal window for normal fertilization because of a failure in the mechanisms that control MII state in the mature oocytes that, in turn, might result in the decay of positive signals targeted on survival factors. Such a connection between machineries that control cell cycle and apoptosis may represent a surveillance system that guarantees the precocious elimination after ovulation of oocytes with a low ability to support normal fertilization and embryo development.
Even though results on animal models should be interpreted with caution, our findings are relevant to both the knowledge and the treatment of human subfertility associated with aging. Indeed, it is known that older couples are likely to be exposed to an increased risk of delayed fertilization because of a decreased coital frequency [49]. Moreover, our observations may be helpful for improving IVF treatment of old patients by encouraging IVF operators to avoid a prolonged interval between oocyte recovery and insemination.
ACKNOWLEDGMENTS
The authors thank the generous technical support of R. De Carolis and G. Ciccone (Animal Care Unit, Faculty of Medicine, University of L'Aquila).
FOOTNOTES
1 Supported by a PRIN grant from Ministero dell'Università e della Ricerca Scientifica e Tecnologica (prot. 2002068388). 
2 Correspondence. FAX: 39 0862 433433; ctatone{at}univaq.it
Received: 28 July 2005.
First decision: 14 August 2005.
Accepted: 24 October 2005.
REFERENCES
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