Inhibition of protein kinases by 6-dimethylaminopurine accelerates the transition to interphase in activated mouse oocytes

Activation of the mouse oocyte is a complex process during which the oocyte passes from meiotic control of the cell cycle to mitotic control. The oocyte, previously arrested at the second meiotic metaphase (M II), extrudes the second polar body and enters a very peculiar period, characterized by the delay of some morphological events when compared to the embryonic mitotic transition from M-phase to interphase. In this period, the oocyte chromatin remains condensed and the interphase network of microtubules is not formed until at least one hour after the second polar body extrusion (M. Weber, J. Z. Kubiak and B. Maro, unpublished), despite maturation-promoting factor (MPF) inactivation (Weber et al., 1991). During the 3-to-4 h following activation, the sperm nucleus is remodelled completely, including the replacement of protamines by histones (Kopečný and Pavlok, 1975a; Kopečný and Pavlok, 1975b; Nonchev and Tsanev, 1990; Rodman et al., 1981). The condensed oocyte chromatin remains in the “telophase” state while the sperm chromatin decondenses, and then undergoes a transient recondensation (Adenot et al., 1991). This recondensed state of the male chromatin corresponds to the beginning of the formation of the male pronucleus, which occurs later than the formation of the female pronucleus. Both sets of chromatin then decondense almost synchronously. The mechanisms involved in the control of this long “telophase” state are not known.

When compared to interphase, M-phase is characterized by a high level of protein phosphorylation, due to the activity of many kinases activated by MPF. In the mouse oocyte, at the time of extrusion of the second polar body, the activity of MPF drops rapidly (Choi et al., 1991; Weber et al., 1991). The pattern of protein synthesis also changes after activation, mainly because of post-translational modifications (phosphorylation) of at least three groups of proteins: 30, 35 and 45 kDa (Howlett, 1986; Howlett and Bolton, 1985). All of these changes take place more slowly during entry into the first interphase, in contrast to the subsequent mitotic divisions, where they are rapid (Howlett, 1986). The 35 kDa protein complex remains phosphorylated for up to 4 h after sperm penetration. Moreover, the newly synthesized 35 kDa proteins become actively phosphorylated during this period, in contrast to the 30 kDa and 45 kDa polypeptides. Thus, the release from M II arrest is not characterized by a rapid and massive dephosphorylation but rather, by a slow and progressive inactivation of some protein kinases and activation of protein phosphatases.

We used the non-specific inhibitor of protein kinases, 6-dimethylaminopurine (6-DMAP; Néant and Guerrier, 1988) during the fertilization or parthenogenetic activation and subsequent culture of mouse oocytes to interfere with the normal course of protein phosphorylation during entry into the first interphase. We show that 6-DMAP is not able to induce MPF inactivation in M II-arrested oocytes, nor does it modify the kinetics of MPF inactivation after oocyte activation. However, following oocyte activation, 6-DMAP inhibits the phosphorylation of the 35 kDa complex, thus accelerating the disappearance of the phosphorylated form of these proteins. This is accompanied by the acceleration of morphological events, like oocyte chromosome decondensation, decondensation and recondensation of the sperm chromatin, pronucleus formation and formation of the interphase microtubule network.

Ovulated oocytes were collected from 9-11-week-old female mice from the OF1 or Swiss strains. Ovulation was induced by intraperitoneal injections of 5 i.u. of pregnant mare’s serum gonadotrophin (PMSG) and human chorionic gonadotrophin (hCG), 48 h apart. Oocytes were collected 16-17 h post-hCG in medium 2 (M2; Quinn et al., 1982). The follicle cells were removed with hyaluronidase (300 i.u./ml in M2). For in vitro fertilization, the zona pellucida was removed with α-chymotrypsin (30 μg/ml in M2) or with acid Tyrode’s solution (Nicolson et al., 1975).

Oocytes with zonae pellucidae were activated by a 6 min exposure to 8% ethanol solution in M2 + BSA medium (Cuthberson, 1983). Spermatozoa collected from the caudae epididymides of an adult F ₁ (C57BL × CBA) male were capacitated for 1 h in 1 ml of Whittingham’s M16 medium modified by Fraser and Drury (1975), containing 32 mg/ml crystalline BSA (Sigma) and equilibrated for 24 h at 37.5°C in an atmosphere of 5% CO₂ in air under liquid paraffin. Zona-free oocytes were inseminated for 10-15 min in fertilization medium containing approximately 1 × 10⁶ sperm/ml, carefully washed and cultured in M2 + BSA.

Oocytes were labelled for 30 min with 5 ng/ml Hoechst 33342 in M2 + BSA, or M2 + BSA containing 6-DMAP, before activation or fertilization. For observation, they were individually distributed in small droplets of culture medium under liquid paraffin in specially prepared Petri dishes (for technical details see Debey et al., 1989). Eggs were observed at different times after activation or fertilization. Hoechst was removed carefully by washing after every observation and added again 0.5 h before the following observation. In the case of fertilization, the rapid motion of spermatozoa attached to the oocyte surface rendered observation of the chromatin in living oocytes impossible. Zygotes were therefore collected at selected times and fixed in EM fixative containing Hoechst. Observations were made with a Nikon (Diaphot) inverted microscope equipped with a 100 W mercury lamp (Osram HBO) and a thermostated box, connected to an intensified camera (Lhesa) and an image processing system (Quantel Sapphire; Debey et al., 1989). Chromatin surface in the focal plane was measured using an image analysis program (Quantel). The chromatin outline was defined either by thresholding the fluorescence intensity, or by manual drawing.

Oocytes and zygotes were fixed in a solution containing 2.5% glutaraldehyde, 0.7% paraformaldehyde in 0.075 M phosphate buffer, pH 7.2-7.4, containing 0.2-0.5% potassium ferricyanide (Yotsuyanagi and Szöllösi, 1981). They were washed in the same buffer and post-osmicated in 2% OsO₄in distilled water, washed three times in distilled water and stained overnight in toto in 0.5% aqueous uranyl acetate solution. They were then dehydrated in a series of increasing concentrations of ethanol solutions and embedded in Epon. Thin sections were stained with uranyl acetate and lead citrate.

Zona-free oocytes were placed in specially designed glass or stainless steel chambers, as described by Maro et al. (1984), except that the chambers were coated with 0.1 mg/ml concanavalin A (Sigma). The samples were centrifuged at 450 g for 8-10 min at 37°C, and left for 10 min at 37°C after centrifugation. They were then fixed as described in de Pennart et al. (1988) with 0.1% glutaraldehyde (Sigma) in PBS supplemented with 1% Triton X-100 (Boehringer Mannheim GmbH). After a 5 min wash in PBS they were extracted with 2% Triton X-100 for 30 min, incubated in 10 mg/ml NaBH₄ in PBS (three incubations of 10 min each), washed twice in PBS and processed for immunofluorescence as described in Maro et al. (1984).

We used the rat monoclonal antibody YL1/2 specific for tyrosinated α-tubulin (Kilmartin et al., 1982) and a fluorescein-labelled, anti-rat antibody (Miles) second layer. The chromatin was visualized using propidium iodide (Molecular Probes; 5 μg/ml in PBS).

The coverslips were removed from the chambers and samples were mounted in Citifluor (City University, London, UK) and viewed under a Leitz Diaplan microscope. Photographs were taken on Kodak T-max 400 film using a Leitz Orthomat photographic system. Confocal laser scanning microscopy was performed using a BioRad MRC-600, mounted on an Optiphot II Nikon microscope equipped with a 60× objective (plan apo; NA 1.4). For fluorescein, an argon ion laser adjusted to 488 nm wave length was used, close to the maximum of absorption of fluorescein, and for propidium iodide a helium-neon ion laser adjusted to 543 nm was used. The emitted light was separated by a dichroic mirror (540DF30), and a DR565 long pass filter was placed in front of the photomultiplier collecting the fluorescein emission and a EF600LP long pass filter in front of the photomultiplier collecting the propidium iodide emission. The adjustment of the confocal system allowed a field depth of about 0.6 μm. Double fluorescence images were acquired in two passes, fluorescein first and propidium iodide second, to avoid any bleeding from one chan-nel into the other. When necessary, the emitted signal was digitized by “photon counting” in order to increase the signal-to-noise ratio and each section was scanned 30-to-50 times. Photographs were taken on Kodak T-max 100 film using a Nikon F-301 camera mounted on a high resolution monitor.

Histone H1 kinase activity was determined as described by Félix et al. (1989) in HK buffer (80 mM β-glycerophosphate, 20 mM EGTA, pH 7.3, 15 mM MgCl₂, 1 mM DTT, 1 mM PMSF, 10 μg/ml leupeptin, 10 μg/ml pepstatin, 10 μg/ml aprotinin), using exogenous histone H1 (H III-S from calf thymus, Sigma) as substrate. Samples, each containing 40 oocytes in 5 μl of water, were lysed by freezing and thawing three times, diluted twice in double concentrated HK buffer (2 × HK) and incubated for 15 min at 20°C in the presence of 3.3 mg/ml histone H1, 1 mM ATP and 0.25 mCi/ml [³²P]ATP. In order to study the effect of 6-DMAP on histone H1 kinase activity in vitro, the drug was added to the reaction tube at a final concentration of either 2.5 or 1.2 mM. The reaction was stopped by the addition of a similar volume of double concentrated sample buffer (Laemmli, 1970) and incubation for 2 min at 90°C. The samples were then electrophoresed on a 10% SDS-polyacrylamide gel (Laemmli, 1970). To test the specificity of the reaction, the p34^cdc2 kinase (histone H1 kinase) was removed by centrifugation from the control sample using p13^suc1-coated Sepharose beads. To calculate the relative percentages of activated oocytes, control groups were cultured for 5-6 h and scored for the presence of pronuclei. Alternatively, some samples were stained with Hoechst 33342 as described previously (Kubiak et al., 1991) and examined under the fluorescence microscope shortly after polar body extrusion.

Oocytes were sampled in groups containing 40 oocytes and incubated in the labelling medium (M2 + BSA containing 500 μCi/ml ³⁵S-methionine or phosphate-free M2 + BSA containing 500 μCi/ml ³²P-orthophosphate) for 1 h starting at 0, 1, 2 and 3 h after activation. Oocytes were then washed in three large drops of M2 + BSA and collected in 5 μl double strength sample buffer for analysis by 10% SDS-PAGE (Laemmli, 1970). For pulse-chase experiments, approximately 500-800 M II oocytes were labelled for 1 h as described above, washed in medium containing nonradioactive methionine (or phosphate), and then cultured in M2 + BSA. They were activated as described above and sampled in groups of 40 oocytes at the time indicated.

A6: oocytes were activated in the presence of 2.5 mM 6-DMAP and then cultured in the presence of the drug; 6A6: oocytes were cultured first in 2.5 mM 6-DMAP for 1 h and then activated and cultured in the presence of the drug; F6: oocytes were fertilized and cultured in the presence of 2.5 mM 6-DMAP. In addition, 0.6 mM and 1.2 mM 6-DMAP were used in certain experiments.

Oocytes were activated (A) or fertilized (F) and then cultured in M2 + BSA.

In every experimental and control group, Hoechst-stained oocytes were examined at 0.5, 1.0, 1.5, 3.0 and 4.5 h of culture. From each group observed, at least 3 specimens were studied by electron microscopy. In both experimental and control groups, only monospermic zygotes were taken for EM study.

The chromatin surface was measured in mono- and dispermic embryos because the development of both spermatozoa was comparable with the monospermic condition, as observed previously by Witkowska (1981).

The observations from living, parthenogenetically activated or fertilized oocytes stained with Hoechst showed that development was not synchronous and that the timing of development was not identical for both groups (Tables 1 and 2). Spindle rotation and polar body formation were observed earlier in fertilized than in artificially activated eggs. At the electron microscope level, during spindle rotation, we observed that one set of telophase female chromosomes remained in contact with a thick cortical actin layer (Fig. 6A), which was present also over the metaphase II spindle.

When we studied the changes in the organization of the microtubule network by immunofluorescence with an antitubulin antibody, we observed that an interphase network of microtubules started to form in control oocytes approximately 1 h after the beginning of the second polar body extrusion (i.e. 1.5-1.7 h after ethanol treatment; Weber, Kubiak and Maro, in preparation). Before this time, microtubules are not usually detected in the cytoplasm and the only microtubules present were found in the region of the midbody joining both sets of chromosomes (Fig. 2A-D).

The dense chromatin mass observed after telophase differed from telophase chromosomes in that it was round or oval, smaller than in telophase and heterogeneous in fluorescence (compare Fig. 1B and D). This dense chromatin mass was characterized by heterogeneous chromatin density and the presence of cytoplasmic vesicles attached to the chromatin surface, indicating nuclear envelope formation (Fig. 1C). This stage was observed 1.5 h after fertilization but generally later in artificially activated oocytes (Tables 1 and 2). At the same time, in fertilized eggs, the sperm was decondensed in 50% of eggs and recondensed in the remaining 50% (Fig. 4A, B). 3h after fertilization, the chromatin from the sperm was recondensed in 64% of eggs and a male pronucleus with nucleoli was formed in the others. The measurement of chromatin surface demonstrated a phase of recondensation of the sperm chromatin during development (Fig. 3).

In the presence of 1.2 mM or 2.5 mM 6-DMAP, we observed large cytoplasmic microtubule asters as soon as anaphase II was completed (about 20 min after ethanol treatment). In addition, numerous long microtubules radiated from the vicinity of chromosomes. These microtubules rapidly formed the interphase network (Fig. 2E-H). At the lowest drug concentration tested (0.6 mM), the interphase microtubules appeared slightly later than at higher concentrations, but still sooner than in the controls.

Oocytes activated (parthenogenetically or by fertilization in vitro) in the presence of 6-DMAP and then cultured further in the presence of the drug, developed faster than the con-trols (Tables 1, 2; Fig. 3). One hour after activation, the female chromatin was already at the dense mass stage, with formation of a nuclear envelope. 1.5 h after activation, the first female pronuclei containing a few nucleoli, were already observed. When oocytes pretreated with 6-DMAP for 1h were activated in the presence of the drug, the events following activation were accelerated further (Table 1). 30 min after activation, one or two very close chromatin dense masses were observed (Fig. 4E) which usually formed a single big pronucleus later (Fig. 4F).

Decondensation and recondensation of the sperm nuclei in Hoechst-stained zygotes resembled controls, except that all phases were accelerated in the presence of 6-DMAP. Further as sperm nuclei remodelling was asynchronously accelerated, a decrease in the chromatin surface was not obvious in 6-DMAP-treated zygotes when the surface was measured as a function of time (Fig. 3). In some decondensed sperm heads, the Hoechst image showed more intensely fluorescing spots, which were bigger than those observed in the controls (Fig. 4D). The fertilization cones were not formed, and the male pronucleus had usually formed after 1.5 h of culture, much earlier than in the controls, and their growth was also faster (Table 2, Fig. 3).

The dense chromatin mass, corresponding to the forming female pronucleus, was often very irregular in shape and surrounded by cytoplasmic vesicles. When the oocyte was treated with 6-DMAP before activation, this mass usually occupied the entire space left by the spindle between two microtubule organizing centers (MTOCs). It looked rather like a group of aggregated chromosomes, enclosing many vesicles and annulate lamellae (Fig. 5A). The ultrastructure of the decondensed pronuclei found at later times was normal.

Electron microscopy showed abnormalities during remodelling of the sperm heads in oocytes exposed to 6-DMAP. In some sperm heads, the chromatin recondensation had already begun in some regions before chromatin decondensation had been completed in other regions. In such sperm heads, classified using Hoechst staining as decondensed, primarily condensed chromatin remained associated with the implantation fossa of the flagellum (Fig. 5B, C). The entire area of the decondensed chromatin was surrounded by vesicles originating from the spermatozoon nuclear envelope (Fig. 5D), that in control oocytes had disappeared very rapidly. Inside the decondensed chromatin, islands of more condensed chromatin were present, surrounded by vesicles with a few nuclear pores present between them (Fig. 5E). The presence of pores in this area suggests strongly the formation of a new nuclear envelope. The part of the old nuclear envelope from the spermatozoon (the redundant segment containing nuclear pores) was found in the same specimen on other serial sections (Fig. 5C). Within the decondensed chromatin, tightly packed straight or undulating bundles of chromatin filaments were present for a long time (Fig. 5D), while they disappeared earlier in controls. These bundles of filaments were still pre-sent in some recondensed sperm chromatin in the presence of the drug (Fig. 5F). The incorporation cone was usually absent above the decondensed or recondensed sperm chromatin, but a thick actin layer was located along the inner surface of the plasma membrane. Finally, the ultrastructure of the male pronucleus formed in the presence of the drug did not differ from controls.

In 96.5% (136/141) of activated and 80% (68/85) of fertilized oocytes the second polar body was not extruded in the presence of 2.5 mM 6-DMAP and, in most cases, the spindle did not rotate. In these eggs, two dense chromatin masses and subsequently, two female pronuclei remained in the egg, usually very close to one another (Fig. 6B, C). In some eggs, a small surface protuberance formed, and then usually disappeared within one hour, although in some cases it remained until the pronuclear stage (Fig. 6C, D). The second polar body was never extruded in oocytes preincubated with 2.5 mM 6-DMAP. Although the polar body did not form, an actin layer was found in the area overlying the spindle and when the spindle did not rotate, or turned slightly, the chromatin was found at a distance from this cortical layer (Fig. 6E). At lower concentrations of 6-DMAP (1.2 mM and 0.6 mM), polar body formation was impaired in most cases.

Since karyokinesis and cytokinesis seemed to be impaired in the presence of 2.5 mM 6-DMAP, we looked at the spindle structure by immunofluorescence using an anti-tubulin antibody in M II-arrested oocytes (Fig. 7A, B). Even after a 3 h treatment with three concentrations of the drug (0.6 mM, 1.2 mM and 2.5 mM), the spindles looked almost normal: we only observed a slight, non significant, decrease in the spindle volume (Fig. 7C) and the presence of some astral microtubules at the spindle poles (Fig. 7B). However, numerous cytoplasmic asters were observed and their number increased when higher doses of the drug were used (Fig. 7B, D). All these effects were clearly dosedependent. These observations also suggested that the oocytes remained in M-phase despite the presence of 6-DMAP.

This was confirmed when we checked the histone H1 kinase activity in 6-DMAP-treated, M II-arrested oocytes. We found that although 6-DMAP is able to inhibit the mouse H1 kinase activity in our in vitro assay, it did not induce a drop in the kinase activity when oocytes were treated for 3 h with the drug (Fig. 8A). This is in agreement with observation that the metaphase II oocytes stay in M-phase, even during prolonged culture in the presence of 6-DMAP (Rime et al., 1989; Szöllösi et al., 1991).

To follow the effects of 6-DMAP on protein phosphorylation we used two different approaches. The first was to examine the pattern of ³⁵S-methionine-labelled proteins, since it has been shown that the major band shifts observed after fertilization or activation are due to dephosphorylation events (Howlett, 1986). The second was to look directly at the pattern of ³²P-phosphate incorporation in oocytes cultured in the presence or absence of 6-DMAP. We did not see any changes in the pattern of newly synthesized proteins after ³⁵S-labelling in M II-arrested oocytes treated for 3 h with 6-DMAP (Fig. 9). However, a decrease in ³²P-phosphate incorporation was observed after 3 h of treatment (Fig. 10). This shows that 6-DMAP is able to decrease the turnover of phosphate in M II-arrested oocytes. However, it is not able to inhibit completely phosphate incorporation as some proteins are still phosphorylated.

6-DMAP did not inhibit either parthenogenetic activation or fertilization, since more than 90% of the oocytes were released from the M II arrest in both cases. We chose artificial activation, which is easy to produce, as a model in which to study the influence of 6-DMAP on histone H1 kinase activity. We found that the activity of histone H1 kinase was not changed in activated oocytes (control and drug-treated) during the first 13-15 min following ethanol treatment (the presumed time of the metaphase-anaphase transition in control activated oocytes). A drop in H1 kinase activity was observed just after this time point in all groups (controls and experimental, preincubated with or without 6-DMAP). These observations suggested that histone H1 kinase activity drops during anaphase II and that both events are not accelerated by 6-DMAP. In order to verify this hypothesis, we collected at the same time a group of 40 oocytes to measure the histone H1 kinase activity and a group of 20 oocytes that were fixed immediately and stained with Hoechst to assess cytologically the mitotic status of the oocytes during the first 20 min following ethanol treatment. The results shown in Fig. 8B confirmed that the drop of histone H1 kinase activity takes place at anaphase II, and that 6-DMAP does not interfere with the timing of both processes.

When 35S-methionine-labelled proteins were observed in control eggs, we confirmed the observations of Howlett (1986). The newly synthesized 35 kDa complex was still phosphorylated 3 h after activation (upper and middle bands still present), whilst the newly synthesized 30 kDa polypeptide was not phosphorylated 1 h after activation (lower band present; Fig. 9). The chase experiments showed that the 35 kDa complex phosphorylated in M-phase was not dephosphorylated, whereas the 30 kDa protein was slowly dephosphorylated during this 3 h period (Fig. 9). In the presence of 6-DMAP, the phosphorylation of the newly synthesized 35 kDa polypeptides was progressively inhibited and the dephosphorylation of the 30 kDa protein and 35 kDa complex was accelerated.

³²P-labelling allowed us to show that 6-DMAP inhibited the phosphorylation of the major phosphoprotein bands, the 35 kDa complex and a band at around 71 kDa, and accelerated their dephosphorylation in activated oocytes (Fig. 10).

After fertilization (or activation) of the mouse oocyte, a very distinctive period takes place during the transition between meiosis and the first mitotic interphase. After the completion of karyokinesis, the chromosomes remain condensed without formation of a nuclear envelope and the interphase network of microtubules is not formed. It is only at the dense chromatin mass stage (see Results) that the nuclear envelope forms and that the cytoplasmic network of microtubules appears. The major finding described in this paper is that this transition can be accelerated by the kinase inhibitor 6-DMAP.

We show that there is a differential effect of 6-DMAP between M II-arrested and activated oocytes. 6-DMAP clearly inhibits in vitro histone H1 kinase in M II oocyte extracts. However, extracts made from 6-DMAP-treated M II-arrested oocytes still retain a full level of histone H1 kinase activity (when 6-DMAP has not been added into the reaction tube). This suggests that 6-DMAP is not able to induce MPF destruction in vivo in M II-arrested oocytes. It is possible that 6-DMAP does not enter the metaphase oocytes, but we observed that the drug induced (1) the formation of microtubule asters; (2) a decrease in phosphate incorporation in the 35 kDa and 71 kDa phosphoproteins and (3) a further acceleration of the transition to interphase in oocytes that had been preincubated with the drug before activation. These observations strongly suggest that 6-DMAP enters M II-arrested oocytes; however, we cannot exclude that the penetration of the drug is lower in M II when compared to the transitional period. It might be expected that inhibition of kinase enzymes by 6-DMAP during M-phase, when their activity is normally high (Howlett and Bolton, 1985; Karsenti et al., 1987), would lead to the dephosphorylation of numerous proteins (since the activity of protein phosphatases should remain unchanged) and consequently, induce the transition to interphase. However, to our knowledge, there are no reports showing that 6-DMAP induces this transition, when added to metaphase cell-free extracts. In addition, there is no evidence to show that inhibition of the cdc2 kinase by 6-DMAP in M-phase extracts induces cyclin destruction and thus MPF inactivation. On the contrary, the cdc2 kinase itself has been shown to trigger cyclin degradation in interphase extracts of amphibian eggs (Félix et al., 1990). 6-DMAP only inhibited changes in microtubule dynamics, when added together with active cdc2 kinase to interphase extracts (Verde et al., 1990, 1991). We observed the formation of asters in M II-arrested oocytes, suggesting that 6-DMAP stabilizes microtubules in the oocyte, as expected from the in vitro experiments. We must also point out that mouse oocytes incubated with 6-DMAP just after germinal vesicle breakdown form interphase nuclei within 7 h, whereas the application of 6-DMAP just after the metaphase I/anaphase I transition leads to the formation of nuclei within 1.5 h of culture (Szöllösi et al., 1991; Szöllösi and Debey, unpublished data). Taken together, these data suggest that 6-DMAP can only induce the formation of interphase nuclei once the MPF fall has been triggered independently from 6-DMAP action.

After oocyte activation, the inhibitory effect of 6-DMAP on kinase activity leads to a dramatic acceleration of the transition between the second meiotic M-phase and the first mitotic interphase. This is correlated with modifications in the pattern of phosphorylation of a 35 kDa protein complex. This complex is phosphorylated during the metaphase II arrest and slowly dephosphorylated after activation, although phosphorylation of the newly synthesized 35 kDa proteins is still observed during the first few hours after oocyte activation (Howlett, 1986; Howlett and Bolton, 1985). Our results show that 6-DMAP inhibits this phosphorylation, probably by inhibiting directly the corresponding kinase(s).

6-DMAP inhibits spindle rotation and second polar body formation. These two events are controlled by interactions between the chromosomes and the oocyte cortex, resulting in the formation of a microfilament-rich domain overlying the spindle (Maro et al., 1986). During normal activation, a close association between one of the rotating spindle poles and the cortical actin layer can be observed. The lack of this contact in the presence of 6-DMAP suggests that protein phosphorylation is involved in the maintenance of the spindle pole-cortex association and successful spindle rotation. Extrusion of the first polar body was also often inhibited by this drug during mouse oocyte maturation (Szöllösi et al., 1991), as well as the formation of both polar bodies in maturing oocytes of Echinoderms (Néant et al., 1989). 6-DMAP does not visibly change actin organization. The thick microfilament layer overlying the M II spindle persists and a thick layer of microfilaments forms over the decondensing sperm in the presence of the drug, as during normal fertilization (Karasiewicz and Soltyńska, 1985; Maro et al., 1984). Cytochalasin D, a microfilament inhibitor, prevents the extrusion of the second polar body and the formation of the incorporation cone (Maro et al., 1984), as does 6-DMAP. This suggests that 6-DMAP inhibits contractile activity in the cortex after activation. This may be linked to the absence of phosphorylation at the activating phosphorylation site of the myosin II regulatory light chain at telophase, thus decreasing contractile activity in the furrow (Satterwhite et al., 1989; Satterwhite and Pollard, 1992). In addition, 6-DMAP changes the outline of the cell surface in mature and activated eggs, causing it to become folded very irregularly, like in maturing oocytes (Szöllösi et al., 1991), suggesting an uncontrolled cortical activity.

In metaphase II-arrested oocytes, 6-DMAP modifies slightly the organization of the microtubule network. Astral microtubules can be observed at the spindle poles and cytoplasmic asters are present, suggesting that the drug facilitates microtubule polymerization in the oocyte. This is consistent with the observations of Verde et al. (1990) in Xenopus egg extracts, where 6-DMAP was used to inhibit the p34cdc2 kinase and was able to block the shrinking of microtubules induced by the p34cdc2 kinase. In control, activated mouse eggs, the interphase network of microtubules does not form until 1 h after the beginning of the second polar body extrusion. During this period, the microtubules behave as if they were still in M-phase, despite the drop in H1 kinase activity (Weber, Kubiak and Maro, unpublished). 6-DMAP accelerates the formation of the interphase network, suggesting that it inhibits a kinase involved in the regulation of the microtubule network, such as the MAP kinase. It has been shown that this serine-threonine kinase is able to induce the interphase-metaphase transition of the microtubule network in interphase extracts from Xenopus eggs (Gotoh et al., 1991). The observed behaviour of microtubules in 6-DMAP-treated, M II-arrested oocytes and activated eggs is consistent with an inhibitory effect of the drug on MAP kinase.

6-DMAP accelerates the formation of both pronuclei. Nuclear envelope assembly around the female chromatin takes place very rapidly, sometimes even before the formation of clear telophase masses. However, after fertilization, 6-DMAP interferes with the remodelling of the sperm nucleus into a male pronucleus. The nuclear envelope breaks down in small vesicles, but these are not dispersed in the cytoplasm and stay around the decondensing chromatin. Mouse sperm nuclei contain protamines but lack somatic histones and transition proteins (Pogany et al., 1981). Chromatin decondensation proceeds first by reduction of the sperm nuclear disulfide bonds, and then by destabilization of the DNA/protamine complex, probably as a result of protamine phosphorylation (Wiesel and Schultz, 1981) that may be due to the protein kinase C (Nishiyama et al., 1988). Chromatin recondensation is accelerated in the presence of 6-DMAP. However, the bundles of chromatin filaments present at the beginning of the decondensation period disappear rapidly in controls but remain for a long time in the presence of the drug. They are even found in the recondensed sperm chromatin after formation of a new nuclear envelope. The last part of the sperm chromatin to decondense in presence of 6-DMAP is the posterior part, perhaps due to the link that exists between the chromatin and the nuclear annulus (Ward and Coffey, 1989). This is also the area of chromatin that condenses last during spermatogenesis (Czaker, 1987). It was still condensed when the rest of the chromatin was already decondensed and had started the transitional recondensation. These two anomalies suggest that 6-DMAP might perturb the exchange of the DNA-associated proteins. We will test this hypothesis by studying the presence of protamine and histone by immunofluorescence during the sperm nuclear remodelling in the presence of the drug.

The remodelling of the sperm nucleus was also accelerated in oocyte fragments penetrated in vitro 3 h after parthenogenetic activation (Borsuk, 1991; Borsuk and Tarkowski, 1989; Szöllösi, Borsuk and Szöllösi, unpublished). This coincides with the normal period of dephosphorylation of the 35 kDa protein complex (Howlett and Bolton, 1985; this paper). These data support a possible role of this protein complex in the transformation of the sperm head into a pronucleus after fertilization. If the dephosphorylation of these proteins is involved in the regulation of the dynamics of sperm nucleus remodelling in the oocyte, their slow dephosphorylation during normal fertilization might prevent a too rapid remodelling of the sperm nucleus, that may lead to abnormalities (as it is the case in the presence of 6-DMAP). It seems clear that the kinase(s) implicated in the phosphorylation of the 35 kDa complex is (are) inhibited by 6-DMAP, but only after MPF inactivation. Thus, it is tempting to speculate that two different subpopulations of kinases able to phosphorylate the 35 kDa complex are active during M-phase and after activation, some of those active during the M II arrest being insensitive to 6-DMAP.

In conclusion, our data show that 6-DMAP inhibits some unidentified kinases, and accelerates some post-fertilization events in mouse oocytes. Inhibition of these kinases does not have an impact on MPF activity during the M II arrest, or on the dynamics of MPF inactivation during oocyte activation. Some of these 6-DMAP-sensitive kinases are involved in the regulation of the polymerization of microtubules, nuclear envelope assembly and chromatin remodelling.

We are grateful to Dr N. Winston for critical reading of the manuscript, Ms D. Huneau for providing high quality thin sections for the electron microscope study and to R. Scandolo, R. Schwartzmann and J. Hamel for their expert photographic work. Part of this work was supported by grants from the Institut National pour la Santé et la Recherche Medicale, the Ligue Nationale contre le Cancer, the Association pour la Recherche contre le Cancer and the Fondation pour la Recherche Médicale (FRM) to Bernard Maro and the FRM to Pascale Debey. Jacek Kubiak is the recipient of a fellowship from the FRM and Henri de Pennart from the Ligue Nationale contre le Cancer.