DEVELOPMENTAL

BIOLOGY

143,162-172

(1991)

Meiotic Competence Acquisition Is Associated with the Appearance of M-Phase Characteristics in Growing Mouse Oocytes DINELI Tufts University

WICKRAMASINGHE,

KARL

M. EBERT,

AND DAVID

F. ALBERTINI’

of Anatomy and Cellu,lar Biology, Schools of Medicine Health Science Schools, Department and Veterinary Medicine, 136 Harrison Avenue, Boston, Massachusetts 02111

Accepted October 5, 1YW To determine whether the acquisition of meiotic competence during the growth phase of oogenesis is associated with the appearance of M-phase characteristics, oocytes obtained from 13. to 30-day-old mice were evaluated by fluorescence microscopy with respect to chromatin and microtubule organization, in vitro maturation ability, and the distribution of M-phase phosphoproteins. Meiotically incompetent oocytes were distinguished from their competent counterparts in displaying elaborate interphase-like arrays of cytoplasmic microtubules and dispersed germinal vesicle chromatin. Meiotically competent oocytes were larger in size, exhibited condensation of chromatin around the nucleolus, and displayed a progressive diminution of cytoplasmic microtubules in conjunction with the appearance of multiple microtubule organizing centers. After 24 hr in culture, medium- to large-sized oocytes exhibiting perinucleolar chromatin condensation resume meiosis whereas smaller meiotically incompetent oocytes retain GVs with diffuse chromatin. Moreover, indirect immunofluorescence studies using the M-phase phosphoprotein specific monoclonal antibody MPM-2 indicate that the appearance of reactive cytoplasmic foci is directly correlated with nuclear changes characteristic of meiotically competent oocytes. Thus, the earliest transition to a meiotically competent state during oocyte growth in the immature mouse ovary is characterized by stage-specific and coordinated modifications of nuclear and cytoplasmic 0 1991 Academic Press, Inc components. INTRODUCTION

Meiotic competence is defined as the ability of mammalian oocytes to spontaneously resume and progress through meiosis, a process initiated during germinal vesicle breakdown (GVBD)’ and completed after extrusion of the first polar body and arrest at metaphase of meiosis-2. The expression of meiotic competence in viva is normally held in abeyance by a combination of intrafollicular factors (Schultz, 1988). In response to periovulatory hormonal signals, meiotic maturation ensues in the hours normally preceding ovulation (Thibault et al., 1987). The ability of isolated oocytes to resume meiosis spontaneously in culture has been exploited experimentally to define when during the course of oogenesis the oocyte acquires meiotic competence. In the mouse, oocytes from animals 15 days and older in age exhibit an increased frequency of meiotic resumption in culture (Bachvarova et al, 1980; Eppig, 1977; Szybek, 1972) that correlates with the extent of oocyte growth (Sorensen and Wassarman, 1976). While small oocytes liberated from juvenile mouse ovaries cannot resume meiosis,

1 To whom correspondence should be addressed. ’ Abbreviations used: GVBD, germinal vesicle breakdown; MPF, maturation promoting factor; PBS, phosphate-buffered saline; GV, germinal vesicle; BMOC-2, Brinster’s modified oocyte culture medium. 0012-1606/91 $3.00 Copyright All rights

0 1991 by Academic Press, Inc. of reproduction in any form reserved.

Sorensen and Wassarman (1976) noted further that many growing oocytes that do undergo GVBD are unable to form polar bodies and these authors made the suggestion that competence acquisition is expressed in two phases relative to the extent of oocyte growth and differentiation. This observation suggests that developing oocytes must first acquire the capacity to reinitiate meiosis and, at a later time, acquire the capacity to complete meiosis-l and progress to metaphase of meiosis-2. While there is evidence to suggest that the biochemical or structural properties of developing oocytes are altered at either of these transitional states (Bornslaeger et al., 1988; Schultz and Wassarman, 1977; Wassarman and Josefowicz, 1978), very little is known with regard to modifications in the expression of cell cycle control elements during the protracted period of meiotic prophase. The importance of maturation promoting factor (MPF) at the onset of M-phase in both meiotic and mitotic cells has been well established in recent years particularly with respect to critical protein phosphorylation events that drive cells to the metaphase state (Kishimoto et al,, 1982; Murray and Kirschner, 1990). Among the events involving protein phosphorylation, chromatin condensation and microtubule assembly at sites of centrosome activation are known to be temporally coordinated during entry into mitotic M-phase (Vandre and Borisy, 1989); however, these events have 162

WICKRAMASINGHE,EBERT,ANDALBERTINI

not been evaluated systematically with respect to the expression of M-phase characteristics or meiotic competence in developing mammalian oocytes. It is of interest to note that progressive changes in chromatin condensation occur in developing mammalian oocytes during meiotic prophase (Chouinard, 1975; Lefevre et ul., 1989; Parfenov et al., 1989). For example, in the mouse, changes in chromatin organization appear to be coordinated with cytoplasmic microtubule alterations (Mattson and Albertini, 1990), and in the pig, progressive changes in chromatin organization have been correlated with the expression of meiotic competence (McGaughey et al., 1979). Since the juvenile mouse ovary has been a useful model for studies on the expression of meiotic competence and the developmental capability of cultured growing oocytes (Eppig, 1977; Eppig and Schroeder, 1989), we have used this system to characterize two aspects of cell cycle control during the developmental expression of meiotic competence. First, the extent of M-phase progression during spontaneous meiotic maturation is shown to occur in two sequential steps; that is, initially oocytes undergo GVBD and progress up to, but not beyond, metaphase of meiosis-l and subsequently oocytes acquire the ability to progress to metaphase of meiosis-z. Second, a series of in Virgo alterations, in chromatin condensation, microtubule reorganization, and changes in the distribution of cytoplasmic phosphoproteins, which resemble changes observed during mitotic prophase, occur concomitant with the expression of meiotic competence. These data suggest that prophase alterations are required for the expression of meiotic competence in growing mouse oocytes. MATERIALS Collectior~

AND METHODS

and Culture of Mome Oocytes

Oocytes were obtained by enzymatic dissociation of ovaries from 13- to 30-day-old CD-1 mice (Charles River Breeding Laboratories, Wilmington, MA). Ovaries were excised and washed in sterile phosphate-buffered saline (PBS), cut into four to eight pieces, and treated for 10 min at 25°C with a follicle-digesting enzyme solution consisting of 0.01% DNase I, 0.05% collagenase, and 0.05% trypsin (Sigma) in Hanks’ balanced salts solution (Ebert et al., 1984). Follicle puncture with sterile needles was used in conjunction with enzyme digestion to isolate oocytes from 25 to 30-day-old mice since large oocytes from these animals were not readily released upon enzyme treatment alone. Oocytes containing intact germinal vesicles (GV) were identified using a Zeiss stereo microscope, washed three times, and subsequently cultured in medium (BMOC-2) supplemented with 10% heat-inactivated and dialyzed fetal calf serum, 1% Lglutamine, 1% essential amino acids, and 1% nonessen-

Acquisitiovr

163

qf Meiotic Cmpetence

tial amino acids (GIBCO, Grand Island, NY) as described previously (Brinster, 1972; Ebert et ab, 1984). Oocytes were cultured in loo-p1 drops (20 per drop) of medium under heavy mineral oil (Fisher, Lot 875080) for 24 hr in 5% 0,, 5% CO,, and 90% N, atmosphere in a Billups-Rothenberg incubator chamber maintained at 37°C. Oocyte Size ClassiJication CorLfiguratiofl

and Evaluation

of Chromatin

The diameters of freshly isolated oocytes were determined using a calibrated ocular micrometer mounted in a Zeiss stereo dissecting microscope. Measurements were made of the diameter between paired points at the vitellus using a final stage magnification of 50x. For some experiments, individual oocytes from different age groups were size classed according to these measurements, subdivided into three groups referred to as small (80 pm), and fixed either immediately or 24 hr after culture. To evaluate GV chromatin organization, freshly isolated or cultured oocytes were fixed and subjected to the triple stain protocol as described below. Previous studies have identified four discrete patterns of GV chromatin organization, as evaluated by fluorescence microscopy in Hoechst 33258-stained samples of gonadotropinprimed mice, which represent sequential stages in mouse oogenesis (Mattson and Albertini, 1990). These are referred to as stage I-IV GVs and are distinguished from each other based upon the degree of chromatin association with the nucleolus. Using this staging scheme, an analysis of GV stages was performed on both size- and age-separated oocytes for at least two experiments (Days 13, 18, 25, 30) using 6-10 ovaries per time point. Samples were analyzed at the time of isolation and 24 hr after culture. All oocytes were stained for chromatin, f-actin, and tubulin as described below and characterized with respect to the degree of meiotic progression based on chromosome configuration, spindle organization and location, and the presence or absence of polar bodies. Moreover, the distinction between meiosis-l and meiosis-2 stage oocytes was facilitated using fluorescence microscopy since the size of metaphase chromosomes and the presence of polar body chromatin on the oocyte surface was readily discernible. Fizution and Labeling Microscopy

of Oocytes for Fluorescence

Oocytes were fixed and extracted for 20 min at 37°C in a microtubule stabilizing buffer (0.1 M Pipes, pH 6.9, 5 mM MgCI,, 2.5 mM EGTA) containing 2.0% formaldehyde, 0.1% Triton X-100, 1 yM taxol, 10 units/ml aprotinin, and 50% deuterium oxide (Herman et aZ., 1983). Fixed oocytes were stored at 4°C for l-5 days in PBS

164

DEVELOPMENTAL BIOLOC:Y

containing 2% bovine serum albumin, 2% normal goat serum, 0.2% powdered milk, 0.1 M glycine, and 0.01% Triton X-100. Oocytes were subsequently incubated with either a 1:lOO dilution in PBS of a rat IgG monoclonal antibody (YOL 34) to cr-tubulin (Kilmartin et al., 1982) or a 150 dilution in PBS of a mouse IgG monoclonal antibody to P-tubulin (Accurate Chemical Co., Lot N122088) for 60 min at 37”C, washed three times in PBS, and further incubated in a 1:50 dilution of fluoresceinated goat anti-mouse IgG (Cooper Biomedical, Inc., Lot 22378) for 60 min at 37°C. The fluoresceinated secondary antibody cross-reacts with both mouse and rat immunoglobulins. Following three washes in PBS, the oocytes were then labeled with rhodamine-phalloidin (1.0 unit/ml, Molecular Probes, Inc., Eugene, OR) for 30 min at 37”C, washed in PBS, labeled with 1 pg/ml of Hoechst 33258 (Polysciences, Inc.) for 5 min at 25°C and finally washed and mounted in a 50% glycerol-PBS solution containing 25 mg/ml sodium azide as an antifading reagent (Bock et ab, 1985). The mouse monoclonal IgG antibody MPM-2 (Davis et al, 1983) was used to stain oocytes at a 1:50 dilution of ascites fluid in PBS, for 60 min at 37”C, washed, stained with Hoechst 33258, and mounted as described above. Oocytes were labeled with secondary antibody alone as a control for specific tubulin and MPM-2 staining. No staining was observed under these conditions. An additional control for MPM2 localization was performed by alkaline phosphatase treatment of oocytes subsequently treated with MPM-2 and secondary antibody; no staining was observed under these conditions. Using this fixation method, MPM-2 staining of metaphase figures in primary cultures of mouse granulosa cells showed staining at the spindle poles as reported by Vandre and Borisy (1989). Labeled oocytes were analyzed by epifluorescence and phase microscopy on a Zeiss Photomicroscope II equipped with fluorescein (Zeiss 487709) rhodamine (Zeiss 487714) and Hoechst (Zeiss 487702) selective filter sets and a 50 W mercury arc lamp. Photographs were taken using 40x Neofluar (0.9 na.) or 63x Neofluar (1.3 n.a.) objective lenses and were recorded on Tri-X-pan film processed with acufine developer for 5.15 min at 25°C. Exposure times ranged from 1 to 8 sec. RESULTS

To document oocyte growth during oogenesis in the CD-l strain of mice mean diameters of oocytes obtained from mice of different ages were measured. Moreover, to determine accurately the extent of meiotic progression in freshly isolated and cultured oocytes triple fluorescence staining was used to monitor nuclear and cytoplasmic organization. Ovaries from 13- to 30-day-old CD-l mice yield oocytes exhibiting mean diameters which range from 60 pm (13 days old) to 85 pm (30 days

VOLUME 143,1991 90

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FIG. 1. Relationship between animal age and oocyte size, ability to resume meiosis, and extent of meiotic progression. (A) Mean diameters for all oocytes measured at the time of isolation; error bars correspond to standard errors of the mean for data pooled from three separate experiments at each time point (total number of oocytes evaluated = 1507). (B) Oocytes were cultured for 24 hr, fixed, and evaluated for GVBD as described in text; % GVBD was calculated for all oocytes at each time point from pooled data. (C) Extent of meiotic progression was evaluated by triple stain analysis of oocytes fixed after a 24-hr culture period; bars illustrate percentage of oocytes at metaphase of meiosis-l (hatched) or meiosis-2 (open) and percentages are derived only from oocytes that had undergone GVBD.

old; Fig. 1A). Oocytes obtained from 13- to 30-day-old animals were cultured for 24 hr to determine at what developmental age they were able to resume meiosis and subsequently progress to metaphase-2. Meiotic competence was evaluated in oocytes fixed at the end of the 24-hr culture period by analyzing chromatin, meiotic spindle, and cortical actin organization after triple fluo-

WICKRAMASINGHE,EBERT,ANDALBERTINI

rochrome labeling. As shown in Fig. lB, the ability to resume meiosis, as evidenced by GVBD, was first observed in a small fraction (~10%) of oocytes obtained from 15-day-old animals; with increasing animal age, a progressively greater percentage of oocytes undergo GVBD, reaching a maximum of approximately 70% by Day 30. Triple fluorochrome labeling analysis indicates that the extent of meiotic progression in vitro, for oocytes undergoing GBVD, is related to both oocyte size and animal age (Figs. lA, 1C). This is evidenced by the observation that the majority of oocytes obtained from 15- to 25-day-old animals arrest at metaphase of meiosis-l, while a progressive increase in the percentage of oocytes reaching metaphase of meiosis-2 occurs from 15 to 30 days of development. Representative triple fluorochrome labeling results of a 24-hr eulture of oocytes obtained from a 25-day-old animal are shown in Fig. 2 and illustrate the cellular heterogeneity of meiotic progression within these cultures. Of particular note, the oocytes that undergo GVBD and arrest at meiosis-l exhibit aligned chromosomes on barrel-shaped metaphase spindles (Figs. ZA, 2B) that subtend a well-differentiated f-actin cortical domain (Fig. ZC). These results indicate that a large proportion of growing mouse oocytes are unable to proceed past the metaphase-anaphase juncture of meiosis-l. In an attempt to distinguish competent from incompetent oocytes, we next evaluated freshly isolated oocytes with respect to nuclear and microtubule organization. Since the trend in oocyte growth and animal age is correlated with an increase in meiotic competence, we examined oocytes obtained from 13-, 18, 25-, and 30day-old animals to determine whether the acquisition of meiotic competence was associated with discernible changes in nuclear or cytoskeletal organization. GV stage oocytes fixed immediately after isolation were found to exhibit marked alterations in chromatin and microtubule organization at each of the ages examined (Fig. 3). Four discrete patterns of GV chromatin organization were observed based on the degree of heterochromatin association with the nucleolus and these were evaluated using the stage I-IV GV classification scheme described in a previous report (Mattson and Albertini, 1990). Oocytes from 13-day-old animals exhibit heterochromatic foci throughout the nucleoplasm (stage I) and characteristically possess an elaborate cytoplasmic microtubule complex (Figs. 3A, 3B). In l&day-old animals, the majority of oocytes exhibit two to five heterochromatic foci at the nucleolar periphery (stage II) and a somewhat diminished microtubule network (Figs. 3C, 3D), although a distinct perinuclear band of microtubules is present. Two other types of GV stage oocytes are noted in 18 to 30-day-old animals. The first is characterized by partial envelopment of the nucleolus with heterochromatin (stage III) in which fewer cytoplasmic mi-

Acquisition

of Mt~iotir Comp~tw~c~

165

crotubules are found that appear shorter in length and emanate from microtubule organizing centers (Figs. 3E, 3F). Finally, GVs are observed with complete nucleolar rims of heterochromatin (stage IV) in which antitubulin staining is confined to variable numbers of microtubule organizing centers located near the GV from which short microtubules radiate (Figs. 3G, 3H). These results show that as oocytes grow in size, discrete modifications in perinucleolar chromatin and cytoplasmic microtubules occur. To determine whether these nuclear/cytoskeletal alterations are developmentally regulated and associated with the expression of meiotic competence, the distribution of stage I to IV GVs was quantitated in oocytes obtained from size-classed 13-, 18, 25-, or 30day-old animals both before and after culture. As shown in Table 1, oocytes obtained from animals at later ages are generally larger in size, confirming results summarized in Fig. 1A. This analysis reveals that independent of animal age, small oocytes consistently exhibit stage I or II GVs, whereas few large oocytes were observed to contain stage I GVs (Table 1). Stage III and IV GVs were predominantly found in medium- and large-sized oocytes obtained from 18-, 25-, and 30-dayold animals. However, it should be noted that a variable percentage of stage II GVs was observed in mediumand large-sized oocytes. These data illustrate a gradual developmental transition in GV morphology that is generally correlated with oocyte growth. To address more specifically the relationship between GV stages and meiotic competence acquisition, size-classed oocytes were cultured for 24 hr and evaluated with respect to the degree of meiotic progression. Table 2 summarizes data derived from 24-hr cultures of oocytes according to oocyte size and animal age. Regardless of animal age, it is clear that all small oocytes are meiotically incompetent and remain at either stage I or II with respect to GV morphology. Moreover, these incompetent oocytes retain an extensive cytoplasmic microtubule complex not unlike that observed in freshly isolated stage I and II oocytes as shown in Fig. 3A-3D. It should also be noted that most medium and large oocytes that were unable to resume meiosis were arrested at GV stages I or II. Significantly, medium and large oocytes were rarely observed in either stage III (17%) 18 days, medium; 14%, 18 days, large; 3% 30 days, medium) or stage IV (0% all ages and sizes), suggesting that only oocytes that previously existed in either stage III or IV (Table 1) were able to resume meiosis. Finally, mediumand large-sized oocytes were found to undergo more complete meiotic progression with advancing age as evidenced by the observations that oocytes from 18-day-old animals exhibit few metaphase-2 figures (13% medium and 10% large) when compared to those obtained from 30-day-old animals which exhibit proportionately greater numbers of metaphase-2 figures (44%: medium

FIG. 2. Triple stain analysis of meiotic progression of a 24-hr culture of oocytes obtained from a 25-day-old animal. (A) Hoechst 33258 staining pattern showing intact GV (GV), metaphase of meiosis-l (Ml), and metaphase of meiosis-2 (M2) stage oocytes. (B) Antituhulin staining pattern illustrating the characteristic barrel shape and subcortical positioning of meiotic spindles; note normal polar body (pb) formation in M2 oocyte. (C) Rhodamine-phalloidin staining pattern illustrating the distribution of cortical f-actin; note the localization of intense staining to the cortical regions overlying meiotic spindles (arrowheads). The scale bar in C corresponds to 20 pm.

FIG. 3. Correlative Hoechst 33258 (A, C, E, G) and antitubulin (B, D, F, H) staining patterns in mouse oocytes isolated from mice aged 13-30 days. (A) Stage I GV containing diffuse chromatin and heterochromatin spots (arrowheads). (B) Same oocyte as in A depicting extensive array of cytoplasmic microtubules; note that nucleoli (n) always appear unstained in tubulin-labeled preparations. (C, D) Stage II GV with perinucleolar chromatin foci (arrowheads) and dense perinuclear array of cytoplasmic microtubules. (E, F) Stage III GV in which chromatin is more condensed (compare to C) and partially envelops the nucleolus; note reduction in cytoplasmic microtubules and presence of a microtubule organizing center (arrowhead, F). (G, H) Stage IV GV with perinucleolar chromatin; one microtubule organizing center (arrowhead, H) is located adjacent to the nucleolus while the other is out of the plane of focus. The scale bar in H corresponds to 20 wrn and all photographs were processed identically. 166

168

DEVELOPMENTAL BIOLOGY TABLE 1 DISTRIBUTION OF GV STAGES IN SIZE-SEPARATED OOCYTES FROM JUVENILE MICE % GV stage

Age (days)

Size (N)

I

II

III

IV

13

Small (146) Medium (73) Large (0) Small (24) Medium (64) Large (79) Small (41) Medium (127) Large (82) Small (27) Medium (105) Large (85)

47 81 0 58 5 1 17 12 5 48 9 6

53 19 0 33 41 27 83 41 32 52 50 25

0 0 0 8 44 59 0 37 44 0 29 55

0 0 0 0 10 13 0 10 19 0 12 14

18

25

30

Note. Oocytes were isolated from animals at the designated ages and size separated as described under Materials and Methods; after sizing, all samples were fixed immediately and processed for fluorescence microscopy. The percentage of oocytes in each GV stage is indicated and the number of oocytes examined in each size class for each age group is shown in parentheses (N). Data are pooled from at least two experiments for each age group.

and 52% large). These data corroborate results described earlier indicating that competence acquisition is developmentally regulated at two transition points that first support progression to metaphase of meiosis-l and subsequently progression to metaphase of meiosis-2 (Fig. 1C). Since this functional transition is correlated with chromatin and microtubule alterations that resemble changes in somatic cells upon entry into M-phase, it is possible that the growing oocyte may undergo a change in its cell cycle status during this phase of oogenesis. In view of this possibility, MPM-2, an antibody that recognizes M-phase specific phosphoproteins, was used to analyze the presence of M-phase epitopes in competent and incompetent oocytes by indirect immunofluorescence microscopy. Oocytes obtained from 18- to 30-day-old animals were fixed at the time of isolation, labeled with MPM-2 antibody and stained with Hoechst 33258 to precisely define the GV stage (Figs. 4A-4H). Stage I and II GV oocytes labeled with MPM-2 exhibit low levels of punctate staining in both the nucleus and cytoplasm (Figs. 4A4D). In stage III and IV oocytes, prominent multiple cytoplasmic foci react with the MPM-2 antibody while the punctate staining is retained in the nucleus and cytoplasm (Figs. 4E-4H). Consistently, stage IV oocytes display MPM-2-reactive foci associated with the nucleolus as well (Fig. 4H). Treatment of stage III and IV oocytes with alkaline phosphatase prior to antibody labeling was found to abolish completely cytoplasmic and nuclear staining with MPM-2. These data suggest that

VOLUME 143.1991

while the expression of some M-phase-specific phosphoprotein epitopes is present in stage I/II oocytes, the transition from stage II to stages III/IV is marked by the appearance of prominent cytoplasmic foci, which coincides with the acquisition of meiotic competence in growing mouse oocytes. DISCUSSION

The point at which mammalian oocytes acquire the capacity to reinitiate and complete meiosis during the course of oogenesis is of obvious importance to normal development particularly in relation to the temporal coordination of meiotic maturation, ovulation, and fertilization. Oocytes obtained from the ovaries of prepubertal mice provide a unique opportunity to evaluate the developmental expression of meiotic competence since the capacity of oocytes to resume meiosis under in vitro conditions has been shown to first occur at or about Day 15 of postnatal development (Szybek, 1972; Bachvarova et ab, 1980). Using this model system, two aspects of competence acquisition during oogenesis in the mouse have been studied. We first demonstrate that modifications in nuclear and cytoplasmic organization occur coincident with the acquisition of meiotic competence and are correlated with the ability to resume meiosis in vitro. These modifications may represent the structural manifestations of a GB/M-like cell cycle transition for

TABLE 2 GERMINAL VESICLE STAGES AND MATURATION PROGRESSION IN OOCYTES CULTURED FOR 24 hr % GV or maturation Age (days) M2

Size (N)

13

Small (257) Medium (108) Large (0) Small (13) Medium (30) Large (28) Small (27) Medium (158) Large (173) Small (15) Medium (77) Large (53)

18

25

30

87 77 0 85 23 11 63 15 7 67 6 0

stage

I

II

III

IV

Ml

13 23 0 15 33 29 37 34 8 33 20 25

0 0 0 0 17 14 0 0 0 0 3 0

0 0 0 0 0 0 0 0 0 0 0 0

0 0 0 0 13 36 0 27 24 0 21 23

0 0 0 0 13 10 0 24 61 0 44 52

Note. Oocytes were isolated from animals at the designated ages and size separated as described under Materials and Methods; all samples were then cultured for 24 hr, fixed, and processed for fluorescence microscopy. The data illustrate the percentage of oocytes in each category (GV stages I-IV or metaphases of meiosis-l [Ml] or meiosis-2 [M2]). The number of oocytes evaluated in each size class for each age group is indicated in parentheses (N). Data are pooled from at least two experiments for each age group.

WICKRAMASINGHE, EBERT, AND ALBERTINI

FIG. 4. Correlative Hoechst 33258 (A, C, E, G) and MPM-2 (B, D, F, H) staining patterns in stage I-IV mouse oocytes obtained from 13- to 25-day-old animals. (A, B) Punctate GV with cytoplasmic staining in a stage I GV oocyte which is similar to that observed in stage II GV oocytes (C, D). (E, F) Stage III GV in which MPM-2 antibody intensely stains four foci (arrowheads) dispersed through the cytoplasm. (G, H) Approximation of some MPM-2 reactive foci (arrowheads) to nucleoli. The scale bar in H corresponds to 20 pm.

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progressing through meiotic prophase. Moreover, these studies confirm and extend the findings of Sorensen and Wassarman (1976) in demonstrating that although mouse oocytes acquire the ability to reinitiate meiosis and undergo GVBD around Day 15 of postnatal development, full meiotic competence, that is, the ability to progress through meiosis-l and arrest at metaphase of meiosis-2, is not acquired until later in development. Specifically, a deficiency in meiotic progression occurs in partially competent mouse oocytes at the metaphaseanaphase transition of meiosis-l. Thus, cell cycle progression is developmentally regulated during oogenesis with respect to the acquisition, modulation, and expression of meiotic competence under in vi?)0 and in vitro conditions. Three lines of evidence bear on the question of whether the changes observed during progressive steps in meiotic prophase represent cell cycle control points that are coupled to the acquisition of meiotic competence. From a comparative perspective, condensation of GV chromatin, including perinucleolar chromatin, has been observed in oocytes from a number of mammalian species at progressive stages of oogenesis (Chouinard, 1975; Lefevre et ah, 1989; McGaughey et ah, 1979; Mattson and Albertini, 1990; Parfenov et uh, 1989). Moreover, in the pig (McGaughey et al., 1979) and primate (Lefevre et al., 1989) changes in chromatin condensation have been correlated with the expression of meiotic competence in cultured oocytes. While nuclear modifications emerge as a useful indicator of prophase progression, are such changes relevant to meiotic competence and the cell cycle status of growing oocytes? Growing mouse oocytes undergo concomitant alterations in cytoplasmic microtubule organization and GV chromatin when they acquire meiotic competence. It is noteworthy that the conversion of an elaborate interphase-like network of microtubules into a system of perinuclear microtubule organizing centers bears a striking similarity to changes in the cytoplasmic microtubule complex known to occur in somatic cells at the G2/M transition point of the cell cycle when microtubules are converted from long to short polymers (Vandre and Borisy, 1989). Verde et al. (1990) have recently shown that long, interphase-like microtubules form in vitro from isolated centrosomes in the absence of cdc2 kinase, a component of maturation promoting factor (MPF) and that short, M-phase-like microtubules assemble from centrosomes in the presence of cdc2 kinase. Since direct microinjection of cdc2 kinase into cultured interphase somatic cells also causes an M-phase like alteration in the microtubule network (Lamb et ul., 1990), it is tempting to speculate that the alteration in cytoplasmic microtubule organization observed presently during competence acquisition not only signifies a GUM-like cell cycle transition at this stage of oocyte

development but may be induced by the appearance of MPF components. Further support for this idea derives from our studies on the expression of M-phase specific phosphoproteins in growing mouse oocytes. Entry into meiotic or mitotic M-phase is known to involve the phosphorylation of many proteins by kinases, one intrinsic to MPF is the cdc2 kinase mentioned earlier, and others presumably activated downstream by MPF itself (Kishimoto et ab, 1982; Murray and Kirschner, 1990). Previous studies attempting to analyze protein phosphorylation events associated with entry into M-phase have utilized the antibody MPM-2. This monoclonal antibody raised against mitotic HeLa cells recognizes phosphorylated M-phase-specific proteins on Western blots (Davis and Rao, 1987) and labels centrosomes in mitotic cells (Centonze and Borisy, 1990). Interestingly, microinjection of MPM-2 into Xenopz~s oocytes prevents progesterone-induced GVBD (Kuang et al., 1989) suggesting that it recognizes MPF or MPF-regulating protein(s). In the present studies MPM-2 was found to label meiotically competent oocytes at multiple cytoplasmic foci that converged on the GV at more advanced stages of oocyte development (Fig. 4). In contrast, meiotically incompetent oocytes exhibit moderate levels of punctate cytoplasmic and nuclear staining, the latter of which is characteristic of the staining described in somatic interphase cells. The expression of MPM-2 epitopes at cytoplasmic foci in meiotically competent oocytes is correlated spatially and temporally with the microtubule alterations noted earlier and, based upon the reactivity of this antibody with M-phase centrosomes (Centonze and Borisy, 1990), lends further credence to the idea that a cell cycle transition to a M-phase state has occurred at this specific point in oocyte development. Studies are underway to define these MPM-2 reactive foci as active centrosomes. Conversely, the persistence of interphase characteristics in freshly isolated or cultured meiotitally incompetent oocytes may be due to a lack of expression of specific cell cycle proteins or requisite posttranslational modifications of these proteins such as phosphorylation/dephosphorylation. Support for this idea derives from the observations of Bornslaeger et al. (1988) in which meiotically incompetent growing oocytes were shown to display qualitative and quantitative differences in protein phosphorylation patterns compared to meiotically competent oocytes. Moreover, because microinjection of MPF or fusion of MPF containing cytoplasm causes GVBD in incompetent growing oocytes in both Xenojvus and mouse (Sadler and Maller, 1983; Balakier, 1978; Motlik and Fulka, 1986; Fulka et al., 1986) it is tempting to suggest that the presence of MPF and its associated role in M-phase specific phosphorylation is required for the expression of meiotic competence. These data, in conjunction with the

WICKRAMASINGHE, EBERT, AND ALBERTINI

present findings showing an association between microtubule reorganization and MPM-2 reactivity, are consistent with the idea that the ability to express active MPF in Gvo is required for the oocyte to make the transition from an incompetent to a competent state. An issue that remains unresolved is why so many oocytes capable of reinitiating meiosis in culture are unable to complete maturation. Sorensen and Wassarman (1976) previously proposed that developing oocytes acquire the capacity to mature in a two-step process enabling GVBD competence prior to the ability to progress to metaphase-2 based on the observation that oocytes from juvenile mice arrest at metaphase-1. The present studies confirm and extend this observation by showing that the block to meiotic progression occurs specifically at the metaphase-anaphase transition point of meiosis1 (Figs. 1 and 2) and establish further that the block is not due to a disturbance in either meiotic spindle assembly or the ability of the spindle to interact with the oocyte cortex. Spindle interactions with the oocyte cortex induce structural remodeling of the cortex that is thought to be required for the initiation of polar body formation at anaphase onset (Long0 and Chen, 1984). The observation that oocytes from juvenile mice arrest at the metaphase-anaphase transition of meiosis-l may be related to recent studies on the regulation of cell cycle progression in animal oocytes and embryos. There is mounting evidence to suggest that the timely destruction of the cyclin component of MPF is responsible for the metaphase-anaphase transition in both meiotic and mitotic cells (Draetta et al., 1989; Murray and Kirschner, 1989; Standart et al., 1987; Westendorf ef al., 1989). It seems plausible, then, that the failure of growing mouse oocytes to progress past metaphase of meiosis-l may be due to a quantitative or qualitative deficiency in protease activity that degrades cyclin or other metaphase arresting factors, such as c-mos (Sagata ef al., 1989), that appear to trigger anaphase onset. In conclusion, these studies document the developmentally coordinated expression of meiotic competence and nuclear/cytoplasmic modifications during progressive phases of oocyte growth in immature mice. Elucidation of the physiological mechanisms that control cell cycle behavior and meiotic competence acquisition in nivo and iu vitro will require objective staging methods, such as those described presently using fluorescent staining techniques in conjunction with a biochemical analysis of the relevant regulatory and target proteins of the MPF kinase. The protracted series of events currently recognized to occur during meiotic prophase in growing oocytes from juvenile mice further establishes the utility of this model system for future studies aimed at understanding the regulation of the cell cycle in the acquisition of meiotic competence.

Acyllisitiorl We are generous spectively, tute, NIH, HD20068 critically tions and

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grateful to Drs. John V. Kilmartin and Potu N. Rao for their provisions of YOL34 antitubulin and MPM-2 antibodies, reand to Dr. Matthew Suffness of the National Cancer Instifor supplying taxol. This work was supported by NIH Grant (D.A.). We thank Tom Ducibella and Susan Messinger for reading the manuscript and for providing helpful suggescomments.

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