Cell cycle progression of parthenogenetically activated mouse oocytes to interphase is dependent on the level of internal calcium

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1 Journal of Cell Science 103, (1992) Printed in Great Britain The Company of Biologists Limited Cell cycle progression of parthenogenetically activated mouse oocytes to interphase is dependent on the level of internal calcium C.VINCENT 1, T.R. CHEEK 2 and M.H. JOHNSON 1, * 1 Department of Anatomy, University of Cambridge, Downing Street,Cambridge CB2 3DY, England 2 Department of Zoology, University of Cambridge, Downing Street, Cambridge CB2 3EJ, England * Author for correspondence Summary Nuclear maturation of the mouse oocyte becomes arrested in metaphase of the second meiotic division (MII). Fertilization or parthenogenetic activation induces meiotic completion, chromosomal decondensation and formation of a pronucleus. This completion of meiosis is probably triggered by a transient increase in cytosolic calcium ions. When activated just after ovulation by a low concentration of the calcium ionophore A23187, the majority of the mouse oocytes go through a metaphase to anaphase transition and extrude their second polar body but they do not proceed into interphase; instead their chromatids remain condensed and a microtubular metaphase spindle reforms (metaphase III). However, a high percentage of these oocytes will undergo a true parthenogenetic activation assessed by the formation of a pronucleus, when exposed to a higher concentration of the calcium ionophore. The capacity of the mouse oocyte to pass into metaphase III is lost with increasing time post-ovulation. Direct measurement of intracellular calcium with Fura-2 reveals higher levels of cytosolic calcium in aged oocytes and/or using higher concentrations of calcium ionophore for activation. It is concluded that the internal free calcium level determines the transition to interphase. Key words: oocyte, parthenogenetic activation, cell cycle, calcium. Introduction Immature mouse oocytes are arrested at prophase of the first meiotic division until just prior to ovulation. The resumption of meiosis is characterized by germinal vesicle breakdown (GVBD), the condensation of chromosomes into distinct bivalents and subsequently the separation of homologous chromosomes and the emission of the first polar body (Donahue, 1968). The oocytes then enter a second meiosis but arrest their progression at metaphase II (referred to as MII). During normal maturation, chromosome decondensation does not occur between metaphase I (MI) and MII. Activation of an oocyte arrested in MII triggers the completion of meiosis, i.e. the extrusion of the second polar body, the decondensation of the chromosomes in the cytoplasm and the formation of a pronucleus. This progression of the cell cycle from MII to interphase can be induced by the spermatozoon at fertilization or by parthenogenetic activation. Fertilization is accompanied by a series of transient rises in the intracellular calcium concentration (Cuthbertson and Cobbold, 1985; Swann, 1990; Cheek et al. 1992), and parthenogenetic activation can result from the release of intracellular stores of calcium induced by calcium ionophore (Steinhardt et al. 1974; Kline and Kline, 1992). After exposure to parthenogenetic activating agents, the proportion of activated oocytes, assessed by the formation of a pronucleus, increases with the post-ovulatory age of the oocytes (Kaufman, 1983). Indeed, the response of oocytes activated immediately after ovulation can be defective: young mouse or rat oocytes extrude the second polar body but do not progress into interphase, arresting again in a new metaphase, called metaphase III (MIII; Keefer and Schuetz, 1982; Kubiak, 1989; Zernicka-Goetz, 1991). We have now used the calcium ionophore A23187 as a parthenogenetic activator and show that MIII formation depends not only on the age of the oocyte but also on the concentration of calcium ionophore used for activation: the majority of young oocytes which fail to undergo a full activation can nevertheless be forced to go into interphase by increasing the concentration of ionophore. By monitoring the intracellular calcium signal by the video-imaging of Fura-2 (Moreton, 1991), it is possible to obtain precise measurements of the calcium transient inside the oocytes for the two different ionophore concentrations used. Our results suggest that the cell cycle progression of oocytes activated early after ovulation depends on the level of internal calcium released during activation and that the capacity of the oocyte to release cytosolic calcium in response to ionophore increases with age.

2 390 C. Vincent and others Materials and methods Collection and treatment of oocytes Oocytes were collected from superovulated females 12.5 h after hcg injection and denuded of cumulus cells as described previously (Vincent et al. 1990). All the incubations were performed in medium H6 containing bovine serum albumin (BSA, 4 mg ml -1 ; Nasr-Esfahani et al. 1990) in cavity blocks pre-warmed to above 37 C. Removal of the zona pellucida was accomplished by a 10 min exposure to 0.001% chymotrypsin (Sigma Type II) (Boldt and Wolf, 1986), followed by two extensive washes. After a 5 to 15 min incubation in H6 + BSA free of calcium, oocytes were exposed to the calcium ionophore A23187 diluted in calcium-free medium at different concentrations according to schedules described in Results. As DMSO (dimethyl sulphoxide) was used as a solvent in the ionophore stock solution, an equivalent dilution of the highest concentration (0.1 or 0.25% DMSO) was used in control groups. After each treatment, oocytes were washed 5 times over a 30 min period. The first three washes were performed in calcium-free medium. Staining of microtubules and chromosomes Oocytes were carried through processing in specially designed chambers as described by Maro et al. (1984). Cells were fixed at 37 C for 30 min in 3.7% formaldehyde in PBS in the presence of 0.5% Triton X-100 (Sigma) for extraction and were washed in phosphate-buffered saline (PBS). Tubulin was visualized with a rat monoclonal anti-tubulin antibody (Kilmartin et al. 1982) followed by rhodamine-labelled anti-rat IgG. Chromosomes were stained by incubation in Hoechst dye (10 µg ml -1 in PBS) for 30 min. Air-dried chromosome spreads were prepared by the procedure of Tarkowski (1966) and stained with Giemsa for 20 min. Cortical granule staining The procedure used was derived from that used to stain hamster oocyte cortical granules (Cherr et al. 1988). Oocytes were fixed in 3% paraformaldehyde in PBS for 30 min and then washed extensively in a blocking solution of 1 mg ml -1 BSA, 100 mm glycine and 0.2% sodium azide in PBS. To visualize exclusively the content of the cortical granules after extrusion, oocytes were not permeabilized. Oocytes were incubated in 10 µg ml -1 Lens culnaris agglutinin conjugated to fluorescein isothiocyanate (FITC-LCA; United States Biochemical Corporation) in blocking solution for 15 min and then washed extensively in the blocking buffer. Loading oocytes with Fura-2 Oocytes were collected from superovulated females and denuded of cumulus cells and zona pellucida as described earlier. Oocytes were washed and transferred to H6+ polyvinylpyrrolidone (PVP; 6 mg ml -1 ) on a coverslip which had been precoated with concanavalin A (ConA; 0.2 mg/ml in PBS) and which formed the base of a metallic perfusion chamber (Moreton, 1991). Oocytes were then loaded with Fura-2 (2 µm; Molecular Probes) for 20 to 30 min and washed extensively with H6 + PVP. The chamber was then placed in a well on the stage of a Nikon Diaphot TMD inverted epifluorescence microscope for imaging. Incubations were all carried out via a system of continuous perfusion through the perfusion chamber maintained at 37 C. After a 5 min incubation in H6 + PVP free of added calcium, oocytes were exposed to the calcium ionophore A23187 diluted in the same buffer. The concentration of the calcium ionophore used and the time of exposure are described in Results. After ionophore treatment, oocytes were washed continuously in calcium-free medium. Fura-2 imaging Intracellular free calcium activity [Ca 2+ ]i was imaged through a Nikon CF-Fluor 20 objective and intensified CCD camera (Extended ISIS, Photonic Science, Robertsbridge, UK), by calculating the ratio of Fura-2 fluorescence at 510 nm, excited by UV light alternately at 340 and 380 nm from twin xenon arc lamps and grating monochromators. Excitation wavelengths were alternated by a rotating chopper mirror attached to a stepper-motor, which was driven in synchrony with the video signal from the camera, to switch wavelengths at the end of each video frame. The resulting video signals were combined by an Imagine digital image processor (Synoptics Ltd., Cambridge, UK) using a lookup table to implement the formula of Grynkiewicz et al. (1985). The calculation was done in real time, to give a live image of [Ca 2+ ]i, which was updated every 80 ms, and smoothed by recursive filtering with a 200 ms time-constant to reduce the noise (for further details, see Moreton, 1991; O Sullivan et al. 1989). The live image was recorded continuously on video tape, and subsequently played back and re-digitized into a frame-store, using software written in the semper language (Synoptics Ltd.) to sample selected oocytes and to record and plot mean [Ca 2+ ]i readings at regulat time intervals. In all cases data were sampled at 4 s intervals. Results Parthenogenetic activation of mouse oocytes after treatment with different concentrations of calcium ionophore Freshly ovulated zona-free oocytes (recovered 12.5 h after the hcg injection) were pipetted several times in order to remove the first polar body and then exposed to calcium ionophore at two concentrations (2 µm for 2 min and 5 µm for 5 min) in calcium-free medium. After washing, each oocyte was cultured in an individual microdrop and examined periodically for evidence of the second polar body extrusion, which generally took place within an hour. Apart from experiment (4), the percentage of oocytes showing polar body extrusion after exposure to 5 µm ionophore appeared high (from 90 to 98%), but only slightly superior to that observed after 2 µm treatment (from 70 to 87%) (Table 1). The state of the chromatin organization was analysed at least 4 hours after the ionophore treatment. Those oocytes which had not shown any second polar body extrusion (whether control or ionophore-treated) were in MII and their condensed chromosomes were organized in a metaphase plate (Fig. 1A, A, A ). The oocytes, from which polar body extrusion had been recorded, showed different behaviour not only according to the experiment but also according to the concentration of ionophore used (Table 1). Regardless of the concentration of ionophore, the incidence of pronucleus formation varied from one experiment to another. However, apart from experiment (4), the percentage of pronucleus formation in each experiment was always significantly higher (P < 0.005; χ 2 test) after exposure to 5 µm ionophore (from 27 to 76%) than after exposure to 2 µm ionophore (0 to 19%) (Table 1). Thus after exposure to a low concentration of ionophore (2 µm), most of the oocytes that had extruded their second polar body did not form pronuclei. As these oocytes did not undergo full acti-

3 Cell cycle of activated oocytes 391 Table 1. Response of mouse oocytes to different concentrations of calcium ionophore No. of oocytes with polar body No. of oocytes With condensed chromatin Showing polar With pronucleus (% of these that show a Incubation conditions Observed body II (%) Observed (%) metaphase spindle) (1) Control (0.25% DMSO) 16 0 (0) 2 µm ionophore (86) 41 6 (15) 35 (80) 5 µm ionophore (93) (68)* 7 (43) (2) Control (0.25% DMSO) 2 µm ionophore (80) 32 6 (19) 26 (58) 5 µm ionophore (92) (53)* 21 (67) (3) Control (0.25% DMSO) 71 0 (0) 2 µm ionophore (71) 49 0 (0) 49 (100) 5 µm ionophore (90) (27)* 42 (98) (4) Control (0.25% DMSO) 32 3 (9) 2 µm ionophore (54) (45) 12 (100) 5 µm ionophore (87) (48) 22 (95) (5) Control (0.25% DMSO) 48 0 (0) 2 µm ionophore (87) 53 8 (15) 5 µm ionophore (98) (76)* *Significant difference between 2 and 5 µm ionophore; P<0.005 (χ 2 test). Oocytes were exposed to the 5 µm ionophore for 30 min. vation, their position in the cell cycle was examined by both microtubule and chromatin staining at different times after ionophore activation. Metaphase III formation in oocytes that did not undergo full activation Before ionophore exposure (Fig. 1 A, A, A ), the chromosomes consisting of two chromatids (Fig. 1A ) were organized on a metaphase plate (MII, Fig. 1A ), and the microtubules were organized in a short anastral barrelshaped spindle (Fig. 1A) (Maro et al. 1985). After ionophore treatment, the oocytes underwent cell cycle progression through anaphase, separation of the sister chromatids and emission of the second polar body (Fig. 1B, B, B ). Thereafter, in the majority of the oocytes (Table 1) a new spindle appeared (MIII spindle; Fig. 1C) around the unichromatid chromosomes (Fig. 1C ). In the majority of the cases, the chromosomes were scattered along the spindle (Fig. 1C ) which appeared longer and thinner (Fig. 1C) than that observed in the MII arrested oocytes (Fig. 1A), and sometimes the MIII spindle was abnormally shaped (pointed-shape spindle, detached microtubules, multipolar spindle). Observations 24 hours later (data not shown) showed only a disorganization of the spindle microtubules but no further progression through the cell cycle, emphasizing that the arrest of oocytes at this MIII stage is durable, as it is in MII before activation. Metaphase III spindles were observed in oocytes that had not undergone pronuclear formation after activation in either 2 or 5 µm ionophore (Table 1). Metaphase III formation does not occur in aged oocytes Both the proportion of activated oocytes (assessed by the extrusion of the second polar body) and the proportion of oocytes showing a pronucleus increased with the post-ovulatory age of the oocytes: 16 h post-hcg, treatment with 2 µm ionophore induced 94% (49/52) polar body extrusion and 98% (38/39) pronucleus formation. It is important to note that in the case of such aged oocytes, decreasing the concentration of ionophore used (1 µm and 0.5 µm) lessened the percentage of oocytes showing the extrusion of the second polar body (respectively, 32% (11/34) and 3% (1/39)) but failed to induce the formation of MIII; all the oocytes which were activated formed a pronucleus. Cortical granule extrusion Cortical granule exocytosis is an event that takes place within the few minutes following fertilization (Cran, 1988; Fukuda and Chang, 1978) or parthenogenetic activation (Gulyas and Yuan, 1985). However, recent studies reported that the exocytosis of cortical granules might occur in oocytes without parthenogenetic activation, either during in vitro maturation (Ducibella et al. 1990) or during exposure to 1.5 M DMSO (Vincent et al. 1991). We have investigated the occurrence of cortical granule extrusion when full parthenogenetic activation was not achieved. The study was performed on freshly ovulated oocytes after 2 µm ionophore treatment when the incidence of MIII formation was high (Table 1) and after 0.5 µm ionophore treatment when activation assessed by polar body extrusion did not occur (data not shown). The oocytes were not permeabilized before the lectin staining in order to assess directly the amount of cortical granule content extruded onto the oocyte surface (Table 2). After exposure to 2 µm ionophore (Fig. 2C,C ), an extensive extrusion of granules (++) had occurred in the majority of the oocytes (Table 2), while no evidence of extrusion (Fig. 2A,A ; ( ) Table 2) or a low-density cortical granule extrusion (Fig. 2B,B ; (+) Table 2) was found in the majority of the control oocytes or in oocytes treated with 0.5 µm ionophore. Calcium profile in oocytes activated at different ages and with different concentrations of ionophore After collection, the oocytes were either loaded with Fura- 2 in the chamber for analysis (freshly ovulated oocytes) or left in culture in H6 + BSA under oil before loading (aged

4 392 C. Vincent and others Metaphase II Extrusion of the second polar body Metaphase III Fig. 1. Analysis of the MII to MIII transition of mouse oocytes after exposure to 2 µm ionophore. Each oocyte was fixed and double stained with both anti-tubulin antibody (A,B,C) and DNA-specific dye Hoechst (A,B,C ) or air-dried prepared and stained for chromatin with Giemsa (A,B,C ). (A,A,A ) Control oocytes before treatment. The oocyte presents a barrel-shaped spindle of microtubules (A) around the chromosomes organized on a metaphase plate (A ). Each chromosome consists of two chromatids (A ). (B,B,B ) Oocytes observed 1.5 h after ionophore treament. The oocyte has gone through an anaphase, separation of the sister chromatids and extrusion of the second polar body. (C,C,C ) Oocytes observed 4 h after ionophore treatment. The oocytes are arrested in a new metaphase (MIII). The spindle is rather long and thin (C). The chromosomes are dispersed along the spindle (C ) and composed of single chromatids (C ). oocytes). After 1 min incubation in H6 + PVP containing 1.2 mm calcium, oocytes were washed extensively with calcium-free medium for 5 min before the calcium ionophore A23187 was added. The same concentrations of calcium ionophore (2 µm and 5 µm) and the same times of exposure (respectively, 2 min and 5 min) as the ones used for the activation study (Table 1) were used for the calcium analysis (Table 3; Fig. 3). After the ionophore treatment, oocytes were washed continuously in calcium-free medium. The results presented in this section came from a single experiment. However, similar results have been obtained in other replicates with similar numbers of oocytes in each group. Exposure to control levels (0.25%) of DMSO did not induce a calcium response. As a result of the ionophore exposure in calcium-free medium, only the internal stores of calcium are mobilized for release (Stauderman and Pruss, 1989). Consequently, one calcium transient is seen (Fig. 3) and the length of time during which the oocytes are incubated with the ionophore Table 2. Comparison of the cortical granule extrusion occurring in mouse oocytes after treatment with different concentrations of calcium ionophore Number of oocytes according to the density of cortical granule extrusion Total + ++ Control (0.1% DMSO) µm ionophore µm ionophore Oocytes were classified according to the density of the cortical granule contents observed on their surface., Almost no cortical granules (Fig. 2A,A ); +, very low density of cortical granules (Fig. 2B,B ); ++, high cortical granule density (Fig. 2C,C ).

5 + Cell cycle of activated oocytes Fig. 2. Fluorescence micrographs of cortical granule contents on the surface of mouse oocytes. Each oocyte was fixed and double stained with Lens culnaris agglutinin (A, A, B, B, C, C ) and DNA-specific dye Hoechst (A,B,C ). (A,B,C) oocytes viewed en face; (A,B,C ) cross-section of the same oocytes. (A,A,A ) Almost no cortical granules are visible at the surface of the MII mouse oocyte ( ). (B,B,B ) Few cortical granules are detected at the surface of the MII mouse oocyte (+). (C, C,C ) High cortical granule concentration is visible at the surface of the mouse oocyte after activation with 2 µm ionophore (++). does not seem to influence the response; free cytosolic calcium declines even though the ionophore is still present (Fig. 3B and D). In order to analyse the response of the oocytes incubated in different conditions, two parameters were chosen to characterize the shape of the calcium spike: in addition to the amplitude (peak calcium reached; [Ca 2+ ] i ), the response duration was quantified by measuring the width of the transient at 75% of the peak [Ca 2+ ] i reached (full-width, 3/4 maximum, FWTM) (Thager and Miller, 1990). These parameters indicate differences in the total amount of [Ca 2+ ] i release during the incubation with the calcium ionophore. Table 3. Characteristics of the calcium transient in freshly ovulated ( h post-hcg) and aged ( h post-hcg) mouse oocytes activated by low (2 M) or high (5 M) concentration of ionophore Measurements from individual oocyte Ionophore concentration (1) (2) (3) (4) (5) Mean Freshly ovulated oocytes 2 µm Ca2+ (nm) (a,b) FWTM (s) (c) 5 µm Ca2+ (nm) (a) FWTM (s) (d) Aged oocytes 2 µm Ca2+ (nm) (b) FWTM (s) (c) 5 µm Ca2+ (nm) FWTM (s) (d) Means with similar letters are significant different (Student s t-test): (a) P=0.001; (b) P=0.04; (c) P=0.009; (d) P=0.05.

6 394 C. Vincent and others A B Time (s) Time (s) C D Time (s) Time (s) Fig. 3. [Ca 2+ ] responses induced in freshly ovulated ( h post-hcg) and aged ( h post-hcg) mouse oocytes by low (2 µm) or high (5 µm) concentrations of ionophore. The graph plotted for each cell is representative of a typical response in each group (Table 3). (A) Freshly ovulated oocyte activated by 2 µm ionophore. (B) Freshly ovulated oocyte activated by 5 µm ionophore. (C) Aged oocyte activated by 2 µm ionophore. (D) Aged oocyte activated by 5 µm ionophore. The arrowheads indicate the transition from medium containing calcium (1.2 mm) to calcium-free medium. The bar represents the length of time during which the calcium ionophore was applied (2 min for 2 µm ionophore, 5 min for 5 µm ionophore). The amplitude of the spike was significantly higher in the presence of 5 µm ionophore (Fig. 3B and D; Table 3) than in the presence of 2 µm (Fig. 3A and C; Table 3), but only for fresh, not aged, oocytes. The same concentration of ionophore, whether 2 µm or 5 µm, stimulates a significantly wider calcium transient in oocytes analysed after in vitro ageing (Fig. 3C and D; Table 3) than in oocytes analysed shortly after ovulation (Fig. 3A and B; Table 3). Discussion The formation of the MIII stage has been reported in freshly ovulated mouse oocytes activated by ethanol (Kubiak, 1989). In that report, it was inferred that such oocytes were not able to undergo the transition to interphase when activated by ethanol. Our results show that fresh ovulated oocytes may also fail to complete a normal cell cycle when activated by a short exposure to a low concentration of ionophore. However, the majority of such freshly ovulated oocytes could be forced into interphase by elevating the concentration of ionophore used. Measurement of the calcium in the cytosol supports the idea that use of higher ionophore concentrations causes release of more calcium inside the cell. This result suggests that the MIII formation seen in the majority of freshly ovulated oocytes which fail to undergo a full activation can be related to a failure to release enough cytosolic calcium. However, the limited calcium transient did successfully induce the release of cortical granules as is observed after fertilization or complete parthenogenetic activation. The conclusion that entry to interphase requires a larger calcium transient is reinforced by the results on older oocytes. It has been known for a long time that the proportion of activated oocytes increases with post-ovulatory age (Kaufman, 1983; Whittingham and Siracusa, 1978). We confirm, as previously reported by Kubiak (1989), that the appearance of MIII is a phenomenon observed only in freshly ovulated oocytes and that the

7 Cell cycle of activated oocytes 395 proportion of oocytes showing a pronucleus increases with the post-ovulatory age of the oocytes. Older oocytes also give a wider calcium transient and show a higher calcium amplitude in response to a lower ionophore concentration, but the reason for these results is unknown. It is possible that the ionophore partitions more effectively into the membrane lipid bilayers as the oocytes get older, increasing the quantity of calcium released from the internal store to the cytosol. Implicit in this conclusion is a change in the lipid composition of the oocyte membranes occuring with aging (Pratt, 1978). Indeed, when we perfused the oocytes at the end of each run with a buffer containing calcium, it was possible to see a small transient of [Ca 2+ ] i in the group of old oocytes (but not in fresh oocytes) that had been activated with 5 µm ionophore (results not shown), indicating that ionophore is retained in old oocytes, but not fresh ones, after washing. Direct analysis of the first calcium transient induced in mouse oocytes by a fertilizing spermatozoon shows a duration of between 2 and 3 min and an average amplitude of 300 nm (Cheek et al. 1992). Thus this single transient provides a sufficient rise in intracellular calcium to activate entry into interphase, and the subsequent calcium transients observed after fertilization in mammals might have some other role (Ozil, 1990). We can conclude from our experiments that the cell cycle progression of mouse oocytes appears to be dependent on the profile of internal calcium release. In Xenopus, the calcium transient that follows oocyte activation is thought to be involved in the inactivation of MPF (maturation-promoting factor) activity due to the degradation of the cyclin B (Murray et al. 1989), and of CSF (cytostatic factor) activity, linked to the destruction of the c-mos proto-oncogene product (Watanabe et al. 1989). There is evidence from studies on Xenopus oocyte extracts in vitro that cyclin B destruction can occur at a lower calcium concentration than that which is required to destroy c-mos (Lorca et al. 1991). Moreover, whereas a Ca 2+ -calmodulin-dependent process has been implicated in cyclin B destruction, the proteolysis of c-mos is probably caused by the calcium-dependent cysteine protease calpain (Lorca et al. 1991; Watanabe et al. 1989). In addition, work on Xenopus (Lorca et al. 1991; Watanabe et al. 1991) and mouse (Weber et al. 1991) oocytes released from meiosis has shown that the CSF inactivation is not the primary cause of MPF inactivation. A mouse oocyte which passes into metaphase III must have exited from the metaphase II arrest, and to achieve this should have decreased its MPF activity. However, the fact that its chromatin remains condensed argues that sufficient CSF activity is still present to stabilize residual or newly formed MPF activity. Thus, the differential sensitivity of MPF and CSF to destruction by calcium observed in in vitro extracts of Xenopus (Lorca et al. 1991) may have an in vivo counterpart in the mouse metaphase III model. We thank M. George and B. Doe for technical assistance, J. Bashford and colleagues for photographic work, J. Kilmartin for the anti-tubulin antibody, and S.J. Pickering, M.J. Berridge, R.B. Moreton and J. McConnell for helpful discussion. This work was supported by a Medical Research Council programme grant to M.H. Johnson and P.R. Braude, and by a grant to M.H. Johnson from the Wellcome Trust. References Boldt, J. and Wolf, D. P. (1986). 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