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Keywords:

  • oocyte;
  • meiosis;
  • caspases;
  • apoptosis;
  • Z-VAD-fmk;
  • doxorubicin

Abstract

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. RESULTS
  5. DISCUSSION
  6. EXPERIMENTAL PROCEDURES
  7. Acknowledgements
  8. REFERENCES

Several studies have shown that apoptotic pathways control fragmentation of unfertilized ovulated oocyte, induced by doxorubicin. But very few have investigated the basis of this process, from prophase I to later stages. Our results revealed the presence of caspase-2L, caspase-9, and caspase-3 in their zymogen and cleaved forms in the oocyte before meiosis resumption. Caspase-2L and caspase-9 were detected in the nucleus of GV-oocytes in a distribution related to chromatin configuration. The inhibition of caspase activity by Z-VAD-fmk accelerated the transition from metaphase I to metaphase II, and caspase-9 and caspase-3 were detected along the meiotic spindle. Surprisingly, Western blot analysis revealed that the three cleaved caspases were present in similar amounts in healthy and fragmented oocytes and caspase inhibition did not prevent doxorubicin-induced apoptosis. Our results suggest that, if cleaved, caspases may be dispensable for final oocyte death and they could be involved in regulating the maturation process. Developmental Dynamics 237:3892–3903, 2008. © 2008 Wiley-Liss, Inc.


INTRODUCTION

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. RESULTS
  5. DISCUSSION
  6. EXPERIMENTAL PROCEDURES
  7. Acknowledgements
  8. REFERENCES

Programmed cell death is an essential regulatory process in the mammalian ovary. Whereas few follicles pursue their growth and reach the ovulatory stage, most degenerate by atresia during the animal's lifetime, together with the oocytes they enclose. The fate of an individual follicle, growth/ovulation or atresia, depends on a delicate balance between factors promoting follicular cell proliferation, growth, and differentiation, or programmed cell death (Craig et al.,2007). It is now widely accepted that the somatic follicular cells surrounding the oocyte die by apoptosis (Hughes and Gorospe,1991; Tilly et al.,1991; Johnson and Bridgham,2002; Matsuda-Minehaya et al.,2006). Moreover, if not fertilized at the optimal time for fertilization, eggs may be aged, leading to abnormal development or apoptosis, as shown in mice (Fujino et al.,1996). Most of the studies on oocyte death, realized on unfertilized ovulated oocytes, bring evidence that the final oocyte fragmentation is an apoptotic process. Transcripts of several members of the Bcl2 family as well as those of caspases are present in ovulated oocytes (Exley et al.,1999) and both caspase-2 (Bergeron et al.,1998) and Bax (Perez et al.,1997) are necessary to apoptosis induced by a chemotherapeutic agent, doxorubicin (DXR), whereas oocytes overexpressing Bcl-2 were resistant to this treatment (Morita et al.,1999). It has also been suggested that caspases are involved in oocyte apoptosis since the general caspase inhibitor Z-VAD-fmk has been demonstrated to be sufficient to inhibit DXR-induced fragmentation in ovulated oocytes (Perez et al.,1997). Apoptosis has also been studied in mouse fetus ovaries, in which active caspase-2 is involved (Lobascio et al.,2007; Hanoux et al.,2007), and in adult rodent ovaries in which active caspase-3 has been found in the oocytes during atresia of the smallest follicles (Ortiz et al.,2006; Fenwick and Hurst,2002). However, the early processes leading oocytes to death during antral follicle atresia are still unknown.

In the ovary, oocytes remain arrested at prophase I and resume meiosis in healthy preovulatory follicles under the influence of the gonadotrophin surge. However, in atretic follicles, the release of meiotic arrest is also one of the first visible signs preceding oocyte degeneration (Lefèvre et al.,1989), and the mechanisms regulating meiosis resumption in these two opposite contexts have to be more precisely studied and compared. As the mammalian oocyte has the particularity to spontaneously resume meiosis in vitro, the few studies on the early regulation of apoptotic oocyte death carried out to date have focused on the in vitro matured oocyte (IVM) model. In cattle, poor quality oocytes have been shown to have higher levels of the pro-apoptotic factor Bax, and lower levels of the anti-apoptotic factor Bcl-2. If the Fas/FasL system has been shown not to be involved in cattle oocyte death (Sakamaki et al.,1997; José de Los Santos et al.,2000; Rubio Pomar et al.,2004), an increase in caspase activity, cell fragmentation, and TUNEL labeling have been observed after a heat shock during IVM (Roth and Hansen,2004a,b), and sphingosine 1-phosphate, which inhibits ceramide-induced apoptosis, protects oocytes against the effects of heat shock, as do caspase inhibitors (Roth and Hansen,2004b). Moreover, these in vitro matured oocytes are more difficult to fertilize and often degenerate during embryo development (Yang and Rajamahendran,2002). The caspase activity has been observed to decline dramatically during maturation in the bovine (Yuan et al.,2005) or to be very low in immature and mature bovine oocytes (Wasielak and Bogacki,2007). In addition, in vitro oocyte apoptosis has been demonstrated to be caspase-3-dependent in the rat (Chaube et al.,2005) as well as in the unfertilized starfish oocytes (Sasaki and Chiba, 2005; Sakaue et al.,2006). All these results suggest the involvement of caspases via the apoptotic intrinsic pathway. However, caspase involvement needs to be confirmed.

As most oocytes enclosed in atretic follicles in vivo resume meiosis, and as most IVM oocytes have lower rates of fertilization and embryonic development than oocytes matured in vivo (Yang and Rajamahendran,2002), we hypothesized that, during IVM, some oocytes could enter meiosis via pathways similar to those inducing meiosis resumption in oocytes enclosed in atretic follicles. This hypothesis is supported by the following similarity between these two situations. In vitro, oocytes are not under the control of follicular granulosa and cumulus cells. In vivo, in atretic follicles, oocytes are no more connected to apoptotic cumulus cells. We tried to determine in vitro whether effector caspases of the intrinsic pathway, caspase-2L and caspase-9, and the initiator caspase-3 were involved in controlling meiosis resumption and oocyte maturation. We have chosen the caspase-2L isoform, as it is the alternative splicing product of caspase-2 transcript, which has a proapoptotic function (Wang et al.,1994; Kumar et al.,1997), and which is the only isoform found in the ovary (Bergeron et al.,1998). We determined the meiosis stage at which molecular markers for apoptosis begin to appear, and the involvement of these markers in mouse oocyte evolution in vitro. For this purpose, prophase I oocytes were maintained in culture, and followed during the early stage of meiosis (GVBD), the transition from metaphase I (MI) to metaphase II (MII), and final fragmentation.

RESULTS

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. RESULTS
  5. DISCUSSION
  6. EXPERIMENTAL PROCEDURES
  7. Acknowledgements
  8. REFERENCES

Caspase-2L, Caspase-9, and Caspase-3 Are Present in GV-Oocytes

The mRNAs for caspase-2L, caspase-9, and caspase-3 were clearly detected in GV-oocytes (Fig. 1a). Western blots were carried out on healthy oocytes before meiosis resumption or after fragmentation induced by 55 hr of culture in the presence of doxorubicin (DXR), known to induce apoptosis of ovulated oocytes (Fig. 1b). Batches of 200 oocytes (≈ 4.6 μg proteins) were sufficient for the detection of caspase-2L, whereas 400 oocytes were required to detect caspase-9 and caspase-3. Large amounts of procaspase-2L, procaspase-9, and procaspase-3 were present in healthy and fragmented oocytes. Faint bands were detected for the cleaved forms of these three caspases and only the intermediate cleaved forms of caspase-2L (30 and 18 kDa) were detected. However, the labeling intensity of the cleaved bands was similar in fragmented and healthy oocytes.

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Figure 1. Presence of caspase-2L, caspase-9, and caspase-3 transcripts and proteins in oocytes. a: Caspase-2L, caspase-9 and caspase-3 transcripts were detected in GV-oocytes (right column). Pools of 5 oocytes were analyzed by RT-PCR for caspase gene expression. Testicular cell RNA was used as a positive control (left column). b: Western blots analysis on batches of 200-400 oocytes at the GV-stage or fragmented after 55 h of culture in 200 nM DXR. Molecular weigh marker is indicated at the left. Procaspase-2L was detected as its 48 kDa, 30 and 18 kDa forms; mouse procaspase-9 was detected as the 51 kDa form and the cleaved 37 kDa form; procaspase-3 was clearly present as a 32-kDa molecule, whereas its active form gave only a weak signal at 19 kDa.

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Effect of the Induction of Apoptosis by DXR or Inhibition of Caspases by Z-VAD-fmk

Meiosis resumption kinetics.

We analyzed the effects of a pancaspase inhibitor, Z-VAD-fmk, and of an apoptotic inducer, doxorubicin (DXR), on the kinetics of meiosis resumption in 4 experiments. The inhibitor of caspases as well as the apoptotic inducer had no effect on GVBD kinetics (Fig. 2a), as the frequency of GVBD oocytes after 2.5 hr in culture was similar in treated and control groups (89.8 ± 6.2% vs. 87.7 ± 5.8 or 90.8 ± 0.8%, respectively).

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Figure 2. Effect of Z-VAD-fmk and/or DXR treatments on in vitro oocyte meiosis and fragmentation. GV-oocytes were incubated in culture medium supplemented with either Z-VAD-fmk, DXR, or both and compared to control. a: GVBD kinetics during the first 3 hr of culture. b: Kinetics of MI to MII transition between 7 and 11 hr of culture. Oocytes were or were not subjected to pretreatment with Z-VAD-fmk and cultured in medium supplemented with Z-VAD-fmk, DXR or both. c: Kinetics of oocyte fragmentation between 24 and 72 hr of culture. d: Rate of fragmentation of ovulated oocytes after 24 hr of culture. Mean values are given ± S.E.M referring to the number of experiments; *P < 0.05; **P 0.01.

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Kinetics of the transition from metaphase I to metaphase II.

By contrast, in the 3 performed experiments, Z-VAD-fmk alone significantly accelerated the MI to MII transition (Fig. 2b): after 8 hr in culture, 42.3 ± 3.9% of oocytes were at the MII stage, vs. 15.0 ± 5.8% for control oocytes (P < 0.05); after 9 hr, the corresponding percentages were 72.6 ± 7.9% vs. 55.0 ± 5.0% (P < 0.05), but the percentages became similar for the treated and control groups after 10.5 hr in culture (82.7 ± 9.9% vs.75.0 ± 2.9%, respectively).

The apoptotic inducer DXR strongly delayed the passage in MII stage (Fig. 2b). After 9 hr in culture, only 20.0 ± 7.6% of the treated oocytes had extruded the first polar body, vs. 55.0 ± 5.0% of control oocytes (P < 0.05). Inhibition remained significant after 10.5 hr, as only 43.3 ± 14.5% of the treated oocytes were at the MII stage, vs. 75.0 ± 2.9% in the control group (P < 0.05).

Nevertheless, 10 μM Z-VAD-fmk did not prevent the slow-down induced by 200 nM DXR, as oocytes cultured in a medium supplemented with both DXR and Z-VAD-fmk had kinetics similar to those of oocytes cultured with DXR alone.

Fragmentation kinetics.

In IVM oocytes (Fig. 2c), Z-VAD-fmk alone had no significant effect on oocyte fragmentation, as the frequencies of fragmented oocytes were similar in treated and control groups for each observation time (in particular, at 72 hr, 35.0 ± 5.4% vs. 27.5 ± 5.2%). The percentage of fragmented oocytes was evaluated taking into account only the total number of viable oocytes in the 4 performed experiments.

The percentage of fragmented oocytes after DXR treatment was significantly higher than that in the control group, regardless of time in culture (in particular, at 72 hr, 56.5 ± 3.1% vs. 27.5 ± 5.2%, P < 0.05). Z-VAD-fmk did not prevent DXR-induced fragmentation (at 72 hr: 46.2 ± 3.7% vs. 56.5 ± 3.1%).

As previously described for ovulated oocytes (Perez et al.,1999), DXR induced a very large increase in oocyte fragmentation rate after 24 hr of culture (54.5 ± 5.4% vs. 8.2 ± 3.5%, P < 0.05). By contrast, Z-VAD-fmk did not significantly prevent the effects of DXR, regardless of dose (50 or 100 μM) or the presence or absence of a 30-min incubation before DXR treatment (Fig. 2d) and these results were obtained from 7 experiments.

Immunolocalization of the Three Caspases During the Different Stages of In Vitro Oocyte Meiosis and Fragmentation

Caspase localization was analyzed at different stages of oocyte meiosis and fragmentation as shown in Figure 3. We have also taken into account the chromatin configuration as already described in GV oocytes (Bouniol-Baly et al.,1999; Zuccotti et al.,2002) where the chromatin may be dispersed in the nucleus (NSN) or condensed around the nucleolus (SN). The SN oocytes are considered as more ready to resume meiosis and more able to be fertilized and to have a good embryo development (for review see Lefèvre,2008).

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Figure 3. Different stages of oocyte meiosis and fragmentation.

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Caspase-2L (Fig. 4).

At the time of their recovery from ovarian follicles, numerous small bright dots were visible in the GV of oocytes exhibiting a SN chromatin configuration (n = 26), and 51.9% had also several large dots more often around the nucleolus. By contrast, when the oocytes exhibited a NSN configuration (n = 4), the caspase-2L staining was homogenous in both the nucleus and the cytoplasm. Whatever the chromatin configuration, caspase-2L was always uniformly dispersed throughout the cytoplasm as small dots.

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Figure 4. Intracellular localization of caspase-2L. In GV oocytes, the staining was homogeneous in the cytoplasm and nucleus when chromatin was diffuse, whereas strong dots were observed in the nucleus (arrow) when the chromatin was condensed around the nucleolus. In MI and MII oocytes, the staining was homogeneously dispersed in the cytoplasm. In fragmented oocytes, caspase-2L was detected with numerous small dots of staining dispersed in the cytoplasm of the fragments (arrow). The red labeling (TRITC) corresponds to caspases (right column); the green labeling (SYTOX Green) corresponds to DNA (left column: merged files).

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After 8 hr in culture, all the oocytes remaining at the GV stage (n = 6) had a NSN chromatin configuration and a caspase-2L staining similar in the nucleus and in the cytoplasm. When the oocytes have resumed meiosis (GVBD-oocytes), the staining was diffuse in the whole oocyte (n = 10). In oocytes at the MI and MII stage, small dots of caspase-2L staining scattered the cytoplasm (n = 51) whatever the absence or presence of DXR in the culture medium. In fragmented oocytes, abundant dots of caspase-2L staining were observed in the perivitelline space and in the cytoplasm of all cell fragments sometimes with one fragment more strongly stained (n = 46).

Caspase-9 (Fig. 5)

At the time of their recovery from ovarian follicles, caspase-9 staining in the GV appeared as numerous small dots, which were more tightly packed in NSN (n = 12) than in SN oocytes (n = 46). It was also noticeable that in 41.3% of the SN-GV oocytes, caspase-9 was present along the nuclear envelope. However, it was diffuse in the cytoplasm of all these oocytes.

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Figure 5. Intracellular localization of caspase-9. In GV oocytes, the staining was faint and homogeneous in the cytoplasm whereas in the nucleus, numerous small dots of stain were more packed when chromatin was diffuse (NSN) than when it was condensed around the nucleolus (SN). In MI and MII oocytes, the staining was strong on the meiotic spindle (arrow) and also at the periphery of MII oocytes. In fragmented oocytes, the staining was observed to be stronger in the fragments containing some DNA associated with what remained of the spindle (arrow). The red labeling (TRITC) corresponds to caspases (right column); the green labeling (SYTOX Green) corresponds to DNA (left column: merged files).

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When the oocytes have just resumed meiosis (GVBD-oocytes), the staining was diffuse in the whole oocytes, but the staining was always stronger in the region of the condensed chromatin, whatever the oocytes were cultured in the absence (n = 16) or the presence (n = 18) of DXR.

Similarly, in oocytes at the MI and MII stage, whatever the absence or presence of DXR, the caspase-9 staining was diffuse throughout the cytoplasm of all the oocytes and more strongly visible along the meiotic spindle in 53.4% of them (n = 43). In fragmented oocytes (n = 26), the caspase-9 staining was also more often diffuse in the cytoplasm of all the fragments (61.5%), and sometimes clearly detected on the dispersed elements of the spindle.

Caspase-3 (Fig. 6).

At the time of their recovery from ovarian follicles, no particular staining was observable in the GV either with a SN chromatin configuration (n = 19) or a NSN one (n = 6), whatever the antibody used; by contrast, a strong labeling along the nuclear envelope was visible in a third of them whatever the chromatin configuration. A faint staining was observed in the cytoplasm with the antibody recognizing both procaspase-3 and its cleaved forms in all the tested oocytes, whereas, besides a faint staining of the cytoplasm, a strong labeling of the subcortical region was observed in all the oocytes treated with the CM1 antibody recognizing only cleaved caspase-3 (n = 25). After 2–5 hr of culture, when the oocytes have resumed meiosis, caspase-3 was always observed diffuse in the cytoplasm and more strongly present around the condensed chromatin whatever the presence (n = 43) or absence (n = 36) of the apoptotic inducer, DXR. In oocytes at the MI and MII stage, the caspase-3 staining was dense mostly along the meiotic spindle. The caspase-3 (n = 18) and its cleaved form (n = 24) were diffuse throughout the cytoplasm of all the studied fragmented oocytes, the cleaved caspase-3 being more strongly present at the periphery of most fragments (78.3%).

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Figure 6. Intracellular localization of caspase-3 and cleaved caspase-3. In GV-oocytes, no significant staining was observed whatever the chromatin configuration and whatever the antibody used. The active caspase-3 labeling was strong in the subcortical region (arrow). In MI and MII oocytes, the staining was strong on the meiotic spindle (arrow) whatever the antibody used. In fragmented oocytes, staining for caspase-3 and active caspase-3 was observed in the fragments containing some DNA associated with what remained of the spindle (arrow). Active caspase-3 was also present at the periphery of some fragments (arrowhead). The red labeling (TRITC) corresponds to caspases (left column); the green labeling (SYTOX Green) corresponds to DNA (right column: merged files).

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Moreover, it was also observed that after DXR treatment, the chromosomes appeared abnormal in numerous oocytes (60.7%, n = 28) (Fig. 7).

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Figure 7. Doxorubicin induced a disorganization of the chromosomes during metaphase formation. The green labeling (SYTOX Green) corresponds to DNA.

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DISCUSSION

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. RESULTS
  5. DISCUSSION
  6. EXPERIMENTAL PROCEDURES
  7. Acknowledgements
  8. REFERENCES

We investigated the stage of meiosis at which molecular markers for apoptosis could be detected, and the eventual involvement of these markers in the progression of mouse oocytes in vitro. We focused on two steps during oocyte maturation: the resumption of meiosis (i.e., GVBD) and the transition from MI to MII, and finally on oocyte fragmentation.

Caspase-2L, Caspase-9, and Caspase-3 Do Not Seem To Be Involved in Oocyte Fragmentation

Our first surprise was to discover that, at least in our hands, the three studied caspases did not appear to be involved in the control of oocyte death. Nevertheless, caspase-2L, caspase-9, and caspase-3 transcripts, which have never been studied at this stage before, were detected in GV-oocytes. Except caspase-9 mRNA, they have already been detected in ovulated oocytes (Exley et al.,1999). Moreover, Western blot analysis showed the three procaspases-2L, -9, and -3 to be present in large amounts, whereas their cleaved forms were only slightly detected. Even if the apoptotic inducer, DXR, increased significantly the rate of fragmented oocytes, it did not amplify the detection of the studied caspases, as well as it did not modify their intracellular localization. Caspase-2L was homogeneously dispersed in the cytoplasm of most of the cellular fragments, whereas caspase-9 and caspase-3 were co-localized with the remaining fragments of the spindle.

Unlike most somatic cell types, MII arrested female germ cells lack an efficient DNA integrity checkpoint control, and damaged DNA or absence of DNA leads to cyto-fragmentation after oocyte activation (Liu et al.,2002). As competent oocytes isolated from preovulatory follicles naturally degenerate after a long period in culture, we used DXR to accelerate this process. However, caspase inhibition by Z-VAD-fmk did not prevent the spontaneous or DXR-induced fragmentation of IVM oocytes. Our results conflict with published data for freshly ovulated oocytes, in which Z-VAD-fmk significantly decreased DXR-induced oocyte fragmentation (Perez et al.,1997,1999). We checked whether this was due to differences between IVM and ovulated oocytes, by carrying out the same experiment on unfertilized ovulated oocytes. But, the results obtained were the same as for IVM oocytes. Even if it was more than improbable, we could not, however, totally exclude that this discrepancy may be due to the strain of mouse used as the susceptibility of ovulated oocytes and the pathways controlling apoptosis have been demonstrated to be variable (Perez et al.,2007). However, even when we checked once on the same strain as Perez et al. (1997), Z-VAD-fmk did not prevent DXR-induced fragmentation (B6C3F1/CrlBR mice, data not shown).

These results suggest that, even if cleaved forms of caspase-2L, caspase-9, and caspase-3 are present, caspases may not be necessary for natural and DXR-induced death. We cannot exclude the hypothesis that procaspases are involved without being cleaved. The absence of a requirement for caspase-3 for ovulated oocyte death has already been demonstrated in Casp-3−/− mice (Matikainen et al.,2001). This highlights the complexity of oocyte death control, and other studies on ovulated mouse oocytes have also reported conflicting observations. Casp-2 null mice ovulated oocytes were reported to be resistant to DXR-induced cell death, suggesting a role for caspase-2 (Bergeron et al.,1998). However, Casp-2 and Casp-3 double-mutant mice produce ovulated oocytes highly sensitive to DXR-induced apoptosis, with compensatory up-regulation of caspase-12 (Takai et al.,2007). Thus, although oocyte fragmentation reflects the programmed cell death process, the control of this pathway remains unclear.

Caspase Inhibition and/or Apoptosis Induction Have Different Effects on the Meiosis Process That Do Not Seem to Be Related

Our second noticeable observation was that whereas the process of oocyte fragmentation was not disturbed by caspase inhibition, the kinetics of meiosis was, but not at the time of reinitiation. Indeed, when caspases were inhibited by Z-VAD-fmk, no significant modification in the percentage of oocytes having resumed meiosis in the first 3 hr of culture has been observed in comparison to the control. These data mean that caspases do not seem to be necessary for meiosis resumption or that they do not need to be cleaved to act, as already demonstrated (Nicholson et al.,1995; Read et al.,2002; Baliga et al.,2004). This also reveals that DXR-induced apoptosis did not affect the first step of meiosis since no modification of GVBD occurrence was observed. By contrast, it strongly slowed down the second step, the MII formation, as well as it modified chromosome appearance in more than half of the oocytes. In these damaged oocytes, the meiosis mechanism could thus be blocked by the known DXR-induced DNA damages (Lee et al.,2005). Even if the pancaspase inhibitor was unable to prevent DXR-effects, it significantly accelerated the oocyte transition from MI to MII, as well as it was able to prevent apoptosis of neonatal male germ cells in our lab (unpublished data). As a consequence, this suggests that either caspases do not need to be cleaved to act, or they are not necessary in the case of damage induced by DXR.

Hypothetical Roles of the Caspases According to Their Intracellular Localizations During Meiosis

At prophase I stage, whereas procaspase-2L and/or cleaved caspase-2L as well as procaspase-9 and/or cleaved caspase-9 were observed in the nucleus of the oocytes, neither procaspase-3 nor cleaved caspase-3 were seen in this cell compartment. Moreover, whereas caspase-2L was strongly present as dot-like structures when the chromatin was condensed around the nucleolus, the staining was homogeneous both in the cytoplasm and the nucleus when the chromatin was dispersed in the nucleus. The prodomain-dependent nuclear localization of caspase-2 has already been observed in somatic cells as dot-like structures (Colussi et al.,1998). It has been suggested that these structures could be a visual indicator of caspase-2 activity and consequent cell death (Baliga et al.,2003). As chromatin condensation around the nucleolus indicates that the oocyte is ready to reinitiate meiosis (Bouniol-Baly et al.,1999; Zuccotti et al., 1999), caspase-2L could be involved in controlling GVBD, the first step of meiosis. If Z-VAD-fmk had no effect on the GVBD process, this could be due to the fact that this inhibitor does not enter easily through the nucleus envelope. Caspase-9 also was present in the GV, but as small dots, which were more tightly packed when the chromatin was dispersed (NSN) than when chromatin was condensed around the nucleolus (SN). Procaspase-9 has also been observed in the nucleus of mammary epithelial cells (Ritter et al.,2000). Since it appears less dense in the nucleus of oocytes ready to resume meiosis, it can be hypothesized that it must leave the nucleus to participate in meiosis induction either in a normal pathway or an apoptotic one. It remains difficult, however, to distinguish between what could be a “normal” or an “apoptotic” GVBD control. Nevertheless, important cellular processes such as cell cycle control are controlled by caspases (Algeciras-Schminich et al.,2002; Gulyaeva,2003; Schwerk and Schulze-Osthoff,2003; Lamkanfi et al.,2007). Moreover, caspase-2, caspase-9, and caspase-3 play an important role in chromatin condensation and nuclear disintegration, two important steps of erythroblast differentiation (Zermati et al.,2001). Caspase-3 is also known to cleave Nup153, a protein of the nuclear pore complex, and thus to be involved in nuclear envelope breakdown (Buendia et al.,1999), and caspase-3 was also observed along the nuclear envelope of the oocytes. Moreover, the nuclear reorganization that precedes the demolition phase correlates with caspase-3 sensitivity of lamina proteins (Raz et al.,2006).

In MII oocytes, procaspase-2L was evenly distributed in the cytoplasm, whereas both procaspase-9 or its cleaved form and cleaved caspase-3 were strongly present along the meiotic spindle. Such caspase localization has never been shown by immunolabeling before, although caspase-3-like activity along the spindle of mouse oocytes has been previously reported (Perez et al.,1999; Papandile et al.,2004). Since the pancaspase inhibitor accelerated this process, this suggests a “brake” role of caspase-9 and caspase-3, slowing MII development and polar body extrusion. Consistent with this result, DXR had an opposite effect; it strongly slowed this step of meiosis. The negative regulator role of caspases on the cell cycle has already been described in somatic cells. In particular, the caspase-3 gene disruption in mice does not prevent apoptosis but induces B cell proliferation (Woo et al.,2003). On the other hand, in the oocyte, the spindle checkpoint is known to control the transition from metaphase I to anaphase I (Wassmann et al.,2003; Brunet et al., 2004). So we can hypothesize that caspases, in particular caspase-9 and then caspase-3, may have a negative regulator role at this step of meiosis.

In summary, we showed for the first time that procaspases and their cleaved forms are present in the oocyte before meiosis resumption and that they could be involved in regulation of the meiotic rhythm. Correlation between caspase-2L and caspase-9 localization in the nucleus and chromatin configuration in prophase I oocytes suggests that these proteases have a role in the initiation of maturation. The presence of caspase-9 and caspase-3 on the meiotic spindle of metaphase II oocytes suggests that they may be involved in the regulation of this step of meiosis, by either participating in normal meiotic progression or guiding early oocytes to death. It will be necessary to compare the role of caspases in vivo in healthy and atretic follicles to determine whether caspases are involved in the orientation of oocytes toward ovulation or apoptosis.

EXPERIMENTAL PROCEDURES

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. RESULTS
  5. DISCUSSION
  6. EXPERIMENTAL PROCEDURES
  7. Acknowledgements
  8. REFERENCES

Oocyte Recovery and Culture

Housing and experiments were in compliance with the relevant French laws (European Act 2001-246, June 6). For experiments performed on prophase I oocytes, 6-week-old female CD1 mice (Charles River, L'Arbresle, France) were injected with 5 IU pregnant mare serum gonadotropin (PMSG, Chronogest; Intervet International, Boxmeer, Holland) and sacrificed 40 hr later. Their ovaries were harvested; fully-grown oocytes were released from the ovarian follicles, and denuded from their surrounding cells by repeated aspiration through a glass micropipette. Denuded instead of cumulus-enclosed oocytes were chosen for the following reasons: (1) they resume meiosis spontaneously as well as cumulus-enclosed oocytes; (2) the kinetics of meiosis and the general aspect of the oocyte were easier to follow under a stereomicroscope during the culture period. All oocytes were at prophase I of meiosis characterized by the presence of a large nucleus, the germinal vesicle (GV).

Groups of 20 to 25 denuded GV-oocytes were cultured at 37°C in 50 μl M2 medium drops (Sigma, Saint-Quentin-Fallavier, France) under mineral oil. M2 medium was preferred to other media for the following reasons: these experiments were realized within the context of the “Nuclear Toxicology and Environment” project for which the putative apoptotic effects of uranium on the female gamete must be analyzed. This medium contains a low level of bicarbonate (2.5 mM) such that it does not induce uranium precipitation. It also contains HEPES, allowing the oocytes to be observed repeatedly on the heating plate of the stereomicroscope without the necessity of gassing throughout the process.

In this study, the oocytes were cultured in the presence or absence of DXR (200 nM; Sigma), known to induce apoptosis of ovulated oocytes. Caspase involvement during in vitro oocyte maturation was assessed by adding 10 μM N-benzyloxycarbonyl-Val-Ala-Asp-fluoromethylketone, a broad-spectrum caspase inhibitor (Z-VAD-fmk; MP Biomedicals), to culture media. The Z-VAD-fmk concentration (10 μM) used was based on published data (Perez et al.,1997). DXR stock solutions were made up in 0.9% NaCl, whereas Z-VAD-fmk stock solutions were made up in DMSO. For control oocytes, the culture medium was supplemented with DMSO at the same concentration as the Z-VAD-fmk final solution (0.05 %). Meiotic stages, fragmentation, and death were observed under a stereomicroscope during the first 3 hr for GVBD, then regularly between 8 and 12 hr for transition from MI to MII, and after 32, 48, 52, 56, and 72 hr for fragmentation. Oocytes were removed from the culture at specific times, and were stored or fixed as described below for further analysis.

For experiments performed on ovulated oocytes, 6-week-old female CD1 mice were injected with PMSG and treated 48 hr later with 5 IU human chorionic gonadotropin (hCG, Chorulon; Intervet International). Ovulated MII oocytes were recovered 15 hr later from the oviducts, freed of cumulus cells by incubation for 10 s in 0.3 mg/ml hyaluronidase (Sigma), and washed three times in culture medium. Groups of 10 MII-oocytes were cultured for 30 min in 50 μl M2 medium supplemented with 0, 50, or 100 μM Z-VAD-fmk, then for 23.5 hr after the addition of 200 nM DXR, as previously described (Perez et al.,1999). Another group of oocytes was cultured for 24 hr in a medium supplemented with both 200 nM DXR and 100 μM Z-VAD-fmk. Control oocytes were cultured for 24 hr in M2 medium, containing 0.1 % DMSO. At the end of the culture period, the number of fragmented oocytes was noted.

Single-Step RT-PCR

Reverse transcription polymerase chain reactions (RT-PCR) were carried out with the OneStep RT-PCR Kit (Qiagen, Hilden, Germany), according to the manufacturer's protocol from 10 pooled GV-oocytes or from 3.7 × 104 total testicular cells as a positive control. Specific primers were used for caspase-2L (F: ccaggccaaagggggcagtttcag, R: gtgtggttctttccatcttgctgg) (Coureuil et al.,2006), caspase-9 (F: catccttgtgtcctactccacc, R: cagctttttccggaggaagt), caspase-3 (F: gagcactggaatgtcatctcg, R: ttgcgtggaaagtggagtc), and β-actin (F: acccacactgtgcccatctac, R: cttcatggtgctaggagccag) as loading control.

Primers have been previously assayed for the absence of genomic DNA amplification (Coureuil et al.,2006) and all reactions have been performed in parallel with negative controls lacking cell lysates (data not shown).

Western Blot Analysis

Batches of GV-oocytes or fragmented oocytes cultured for 55 hr with 200 nM DXR were collected, rinsed twice in phosphate-buffered saline (PBS; Pierce, Rockford, IL), and resuspended in 5 μl of lysis buffer (25 mM Hepes, pH 7.3, 10 mM EDTA, 0.1% SDS, 125 mM NaCl, 10 mM NaF, 0.5% sodium deoxycholate, and 1% Triton) supplemented with 1% (v/v) protease inhibitor cocktail (Sigma) during 10–15 min.

Whole oocyte lysate was diluted in 5 μl 2× Laemmli Buffer (Sigma), boiled for 2 min and loaded on a 12% polyacrylamide 0.1% SDS gel. Electrophoresis was carried out at a constant voltage (200 V) for about 30 min in 192 mM glycine, 25 mM Tris, and 0.1% SDS. Separated proteins were electroblotted onto nitrocellulose membrane (Millipore, Saint-Quentin-en-Yvelines, France) in liquid medium (192 mM glycine, 25 mM Tris and 20% [v/v] ethanol) for 1.5 hr at 100 V. Membranes were rinsed in PBS and nonspecific sites were blocked by incubation for 1 hr in SuperBlock buffer in PBS and 0.1% Tween 20 (Pierce). Membranes were incubated overnight at 4°C in blocking buffer, with anti-caspase-2L rabbit polyclonal antibody (sc-626; 1:300; Santa Cruz, CA), anti-caspase-9 mouse monoclonal antibody (9508; 1:500; Cell Signaling), anti-caspase-3 rabbit monoclonal antibody (9665; 1:1,000; Cell Signaling, Boston, MA), or anti-β-actin mouse monoclonal antibody for control (C-15; 1:5,000; Sigma). Membranes were washed three times, for 10 min each, in 0.1% Tween 20 in PBS (PBS-T), and proteins were tagged by incubation with ImmunoPure Peroxidase–conjugated goat anti-rabbit or anti-mouse IgG secondary antibody (for caspase-2L, 1:2,500; for caspase-9 and caspase-3, 1:1,000; for β-actin, 1:2,000; both from Pierce), diluted in blocking buffer for 1.5 hr at room temperature. Membranes were washed twice, for 5 min each, in PBS-T, once for 5 min in PBS, and specific labeling was detected with SuperSignal West Femto Maximum Sensitivity Substrate (Pierce), according to the manufacturer's instructions. MagicMark XP Western Protein Standard (Invitrogen) was used as molecular weight markers.

Immunocytochemistry on Whole Oocytes

Collected oocytes were fixed by incubation in 2% paraformaldehyde (Sigma), 0.5% saponin (Sigma) in PBS for 1 hr at 37°C after culture in the presence of DXR or not. Fixed oocytes were rinsed three times, for 10 min each, at room temperature, in 1% BSA (Sigma) and 0.5% saponin in PBS. They were incubated for 15 min at room temperature in 0.3% ammonium chloride, 0.5% saponin in PBS, and then for 1 hr at room temperature in the blocking buffer (1.5% BSA, 0.05% Tween 20 and 0.5% saponin in PBS). Finally, they were incubated overnight at 4°C with primary antibody diluted in blocking buffer. Caspase-2L was detected with the rabbit polyclonal antibody (sc-626; 1:40; Santa Cruz), caspase-9 with the rabbit polyclonal antibody (cs-9504; 1:40; Cell Signaling), caspase-3 with the rabbit monoclonal antibody (9665; 1:30; Cell Signaling), and cleaved caspase-3 with the rabbit polyclonal antibody (CM1; 1:500; BD Biosciences, San Jose, CA). Oocytes were rinsed for 10 min in blocking buffer at room temperature and incubated for 45 min at 37°C in tetramethylrhodamine isothiocyanate (TRITC)–conjugated anti-rabbit IgG secondary antibody (1:40; Sigma) diluted in blocking buffer. They were rinsed for 10 min in blocking buffer at room temperature and then maintained in PBS. DNA was labeled just before observation, by incubation for 20 min in 500 nM SYTOX Green (Invitrogen-Molecular Probes, Eugene, OR) in PBS at room temperature. Control oocytes were similarly treated, however, either without the primary antibody or with isotype IgG.

Labeled oocytes were observed with an inverted confocal microscope (LSM5 Pascal; Carl Zeiss SA, Le Pecq, France) in a single optical plane through the GV or the metaphase plate, using a 40× objective (Plan-APOCHROMAT 0.95 Korr). A multi-channel laser configuration was used and images were recorded using a four frame-average mode. SYTOX Green fluorescence was excited with a 458-nm argon laser and recorded through a 560-nm long-pass filter. TRITC fluorescence was excited with a 543-nm helium-neon laser and recorded through a 560-nm long-pass filter. The same intensity of excitation was used for control and caspase-labeled oocytes.

Statistical Analysis

In vitro experiments were carried out at least in triplicate. Oocytes recovered from four mice were randomly assigned to the control and treated groups. For the immunohistochemistry data, the percentages were compared using the Chi-square test and for meiosis kinetics the data presented as means ± Standard Error of the Mean (S.E.M) and were compared using the Student's t-test. They were considered to be significantly different if P < 0.05.

Acknowledgements

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. RESULTS
  5. DISCUSSION
  6. EXPERIMENTAL PROCEDURES
  7. Acknowledgements
  8. REFERENCES

This work was supported by a “Toxicologie Nucléaire et Environnement” contract and by the EDF, through the CEA, and by INSERM. Emilie Arnault, PhD student, was supported by the ToxNucE program of the CEA. We thank “Alex Edelman & Associates” for English editorial assistance.

REFERENCES

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  3. INTRODUCTION
  4. RESULTS
  5. DISCUSSION
  6. EXPERIMENTAL PROCEDURES
  7. Acknowledgements
  8. REFERENCES
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