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Biology of Reproduction 59, 456-462 (1998)
©Copyright 1998 Society for the Study of Reproduction, Inc.

Oocyte Competence for In Vitro Maturation Is Associated with Histone H1 Kinase Activity and Is Influenced by Estrous Cycle Stage in the Mare1

Ghylène Goudeta, Jacqueline Bézarda, François Belina, Guy Duchampa, Eric Palmera, , and Nadine Gérard2,a

a Institut National de al Recerche Agronomique-Haras Nationaux, Equipe de Reproduction Equine, P.R.M.D., F-37380 Nouzilly, France


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The in vitro maturation rate of equine oocytes remains low, regardless of culture conditions. Our objective was to determine the reasons for failure of equine oocytes to resume meiosis during in vitro maturation and to ascertain the influence of the estrous cycle stage on meiotic competence. In 10 cyclic mares, 7 ultrasound-guided follicular punctures were performed alternately during the follicular phase (group DF; n = 3 punctures), at the end of the follicular phase (group EF; n = 2), and during the luteal phase (group DL; n = 2). We evaluated the competence of the oocytes for in vitro maturation and measured their maturation-promoting factor activity by histone H1 kinase assay.

Puncturing once at the end of the follicular phase and once during the luteal phase, or three times during the follicular phase, yielded about 11 cumulus-oocyte complexes per 22 days. The maturation rate was different between the groups, 51% in group EF, 34% in group DL (p < 0.05), and 15% in group DF (p < 0.01), and it increased with an increase in follicular diameter (p < 0.05). After in vitro culture, the H1 kinase activity was lower in oocytes that remained in germinal vesicle or dense chromatin stages than in oocytes that reached metaphase I or metaphase II (p < 0.05). The H1 kinase activity was not different between oocytes in germinal vesicle stage after in vitro maturation and immature oocytes that were not cultured in vitro, and was higher in preovulatory oocytes that reached metaphase II in vivo than in the oocytes that reached metaphase II after in vitro maturation (p < 0.001). This is the first report on kinase activity in the equine oocyte.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The conditions for in vivo oocyte maturation in the mare are somewhat different from those in other domestic mammals; the ovulatory LH surge occurs as a progressive rise that takes several days, with a maximum concentration occurring 1 day after ovulation [1, 2]. The conditions for in vitro oocyte maturation seem to be unusual too. In the classic media, adapted from those for bovine oocytes, the in vitro maturation rate remains low: 40–70% of the oocytes are in metaphase II at the end of the culture period ([3]; for review, see [4, 5]), whereas in the goat [6], the sow [7], and the cow [8], more than 90% of the oocytes are in metaphase II. The reasons for the failure of equine oocytes to resume meiosis during in vitro culture are not known.

Meiotic maturation is characterized by nuclear envelope breakdown, chromatin condensation, spindle assembly, and progression to the metaphase II stage. The occurrence of these events is accompanied, and probably regulated, by changes in the phosphorylation patterns of various cellular proteins [9–11]. An important component of this activity is the maturation-promoting factor (MPF) [12], which was found to be a universal cell cycle regulator of both mitosis and meiosis [13]. During meiotic maturation of oocytes from various mammalian species, MPF activity, as measured by phosphorylation of histone H1 [13], is very low during the germinal vesicle stage, and peaks at metaphase I and II stages (mice [14, 15], rabbits [16], pigs [17], goats [18], and cattle [19]). The phosphorylation-dephosphorylation activity during the meiotic cell cycle has never been studied in the equine oocyte, and no information about MPF activity is available in the mare. We hypothesized that an alteration in MPF activity may be responsible for the failure of equine oocytes to resume meiosis during in vitro culture.

The study of oocyte competence for in vitro maturation has been enhanced by the use of in vivo ultrasound-guided follicular puncture. Repeated punctures allow collection of a well-characterized population of oocytes and comparison of different populations within the same animal. Using this technique, we showed recently that the mare's reproductive status has an influence on oocyte competence for in vitro maturation: the percentage of competent oocytes tends to be higher at the end of the follicular phase than in the luteal phase [4]. In the present study, we also investigated the competence for in vitro maturation of oocytes collected during the follicular phase, before the emergence of the dominant follicle. We established an assay system for MPF activity in equine oocytes and investigated changes in activity in relation to meiotic competence.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Experimental Design

Ten cyclic mares (Selle Français) in good body condition, from 8 to 19 yr of age, housed indoors and fed with concentrates, were used in an experiment conducted from April to June. They received an initial synchronization treatment [20] with an intravaginal sponge containing 0.5 g altrenogest (Regumate; Roussel UCLAF, Romainville, France) plus 50 mg of estradiol benzoate (ß-estradiol 3-benzoate; Sigma, La Verpillère, France) during 1 wk. On the day of sponge removal, mares received a prostaglandin F2{alpha} analogue injection (cloprostenol [Estrumate], 250 µg/mare i.m.; Pitman-Moore, Meaux, France) to induce luteolysis, and all follicles larger than 5 mm were punctured to make the ovaries free of atretic follicles. Healthy follicles developed afterward. Ovarian activity was assessed by routine rectal ultrasound scanning [21] using an Aloka (Tokyo, Japan) 210 with a 5-MHz linear probe. Blood samples were withdrawn every 2 days, and plasma progesterone concentrations were measured using the RIA method described by Palmer and Jousset [22]. Luteinization of follicles was diagnosed by measurement of plasma progesterone levels higher than 2 ng/ml [23].

Follicular Puncture and Oocyte Recovery

For each of the 10 mares, 7 ultrasound-guided punctures were performed alternately during the follicular phase (termed group DF; n = 3; see below), at the end of the follicular phase (termed group EF; n = 2; see below), and during the luteal phase (termed group DL; n = 2; see below). In order to avoid any seasonal effect, 5 mares were punctured according to the rhythm EF, DL, EF, DL, DF, DF, DF, and 5 according to DF, DF, DF, EF, DL, EF, DL.

When punctures were performed at the end of the follicular phase (EF), an injection (i.v.) of 25 mg of crude equine gonadotropin (CEG; [24]) was administered when the largest follicle reached 35 mm to induce ovulation; all follicles larger than 5 mm were punctured 34 h later, i.e., just before the expected ovulation. For punctures done during the luteal phase (DL), a CEG injection was administered when the largest follicle reached 16 mm to increase the in vitro maturation rate [4]; all follicles larger than 5 mm were punctured 34 h afterward. On the day of the puncture in the luteal phase, prostaglandin F2{alpha} was administered to suppress the corpus luteum. During the follicular phase (DF), the punctures were performed before the emergence of the dominant follicle. A CEG injection was administered when the largest follicle reached 16 mm, to increase the in vitro maturation rate; all follicles larger than 5 mm were punctured 34 h afterward.

During the follicular puncture attempts, mares were sedated with detomidine (0.6 mg/100 kg BW i.v. Domosedan; Smithkline & French, Courbevoie, France), and the rectum was relaxed with prifinium bromide (45 mg/100 kg BW i.v. Prifinial; Vétoquinol, Lure, France). The follicles were punctured using a transvaginal ultrasound-guided follicular aspiration technique [25] with a 7.5-MHz sectorial probe (Kretz; Soframed, Truchtersheim, France). To improve the oocyte recovery rate, two types of aspiration needles were used [25]. A single lumen needle (length, 600 mm; outer diameter, 1.8 mm; Thiebaud Frères, Jouvernex Margencel, France) was used for puncturing follicles > 25 mm, and a two-way needle (length, 700 mm; outer diameter, 2.3 mm; internal diameter, 1.35 mm; CASMED, Cheam Surrey, England) was used for follicles <= 25 mm. After follicular fluid aspiration, the follicle was flushed with PBS (Dulbecco "A"; Unipath, Dardilly, France) containing heparin (50 IU/ml; LEO S.A., St-Quentin Yvelines, France) at 37°C. After puncture, the mares received an antibiotic injection (Mixtencilline: 1.6 x 106 IU penicillin/100 kg BW and 1.3 g dihydrostreptomycin/100 kg BW i.m.; Rhône Mérieux, Lyon, France). All aspirated fluids were individually examined under a stereomicroscope for oocyte recovery.

Oocyte Culture and Nuclear Examination

At recovery, oocytes were individually classified according to cumulus aspect as previously described [4]: expanded cumulus, compact corona radiata, and compact cumulus and cells from the follicular wall.

Compact cumulus-oocyte complexes (COCs) (compact corona radiata plus compact cumulus and cells from the follicular wall) were cultured individually in humidified atmosphere (95% air:5% CO2) at 38.5°C for 30 h in 500 µl of maturation medium: Medium 199 with Earle's salt, 2.2 g/L NaHCO3, and L-glutamine (Gibco, Eragny, France) supplemented with 20% FCS (inactivated fetal calf serum; Gibco), antibiotics (100 IU/ml penicillin, 100 µg/ml streptomycin, and 0.25 µg/ml fungizone; Gibco), CEG (9.5 µg/ml equine FSH and 15 µg/ml equine LH; [24]), and estradiol-17ß (1 µg/ml; Sigma). After culture, the COCs were stripped with small glass pipettes in 500 µl of PBS solution supplemented with 87.5 IU/ml hyaluronidase (type III, 875 IU/mg; Sigma) at 37°C. Totally denuded oocytes were rinsed in PBS with 1% FCS at 37°C, stained with 1 µg/ml bis-benzamide (Hoechst 33342; Sigma) in PBS for 5 min at 37°C for DNA detection, and evaluated with a fluorescence microscope. They were classified according to chromatin configuration as "germinal vesicle," "dense chromatin," "metaphase I," "metaphase II," or "degenerated" as previously described [4]. They were then rinsed in maturation medium and kept at -80°C in 5 µl of maturation medium.

Expanded COCs at recovery were immediately stripped of their cumulus cells, stained, classified, and kept as described above.

In order to have immature controls, 12 compact COCs were collected on the day of sponge removal from follicles smaller than 20 mm. They were stripped of their cumulus cells just after collection, rinsed, and kept at -80°C without any culture time.

Histone H1 Kinase Assay

The assay of histone H1 kinase activity was adapted from the one previously described in the mouse [26]. All products were purchased from Sigma unless otherwise specified. Briefly, each oocyte in 5 µl maturation medium was added to 5 µl of buffer containing 160 mM ß-glycerophosphate, 40 mM EGTA (pH 7.3), 30 mM MgCl2, 2 mM dithiothreitol, 2 mM PMSF, 20 µg/ml pepstatin, 20 µg/ml leupeptin, and 20 µg/ml aprotinin. Oocytes were lysed by three successive freezing and thawing cycles. Kinase reaction was initiated by the addition of 10 µl of reaction buffer containing 7 mg/ml histone H1, 1 mg/ml ATP (Amersham, Les Ulis, France), and 30 µCi/ml [{gamma}-32P]ATP (Amersham). After 30 min of incubation at 37°C, the reaction was stopped by the addition of 20 µl of a buffer containing 10% w:v SDS, 160 mM trizma base (pH 6.8), 10 mM EDTA, 20% v:v glycerol, 10% v:v ß-mercaptoethanol, and bromophenol blue. The samples were boiled for 3 min. Proteins were separated by 15% SDS-PAGE according to Laemmli [27]. Prestained standards (range: Mr 6.9–202 x 10-3; Bio-Rad, Ivry sur Seine, France) and negative control samples containing 5 µl PBS without an oocyte were run simultaneously. After air drying, gels were exposed to Kodak Biomax x-ray films at -80°C. The films were digitalized with an Eikonix 1412 scanner camera (Eastman Kodak, Rochester, NY), and patterns were quantified using Kepler software (Large Scale Biology Corporation, Rockville, MD). In each assay, 19 individual oocytes were run simultaneously.

Statistical Analysis

The chi-square test was used to compare oocyte recovery and maturation rate in the three groups (groups DF, EF, and DL). Analysis of oocyte maturation after in vitro culture according to follicle diameter was performed by logistical regression analysis. Nonparametric test (G-test = 2I-test) was used for comparison of nuclear maturation between the different classes of follicle diameter. Analysis of H1 kinase activity was performed by unbalanced variance analysis (Statistical Analysis System software, GLM procedure, Cary, NC); each assay was considered as a block.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
For each of the 10 mares, 7 ultrasound-guided puncture attempts were performed alternately during the follicular phase (DF; n = 3), at the end of the follicular phase (EF; n = 2), and during the luteal phase (DL; n = 2).

Plasma Progesterone Levels

The oocytes collected during DF and EF puncture attempts came from follicles that grew in the presence of low levels of progesterone. However, for 3 of 20 EF attempts and 3 of 30 DF attempts, the levels of circulating progesterone were higher than 2 ng/ml. The oocytes collected during the DL puncture attempts came from follicles that grew in the presence of high levels of progesterone. However, for 2 of 20 attempts, the levels of progesterone were lower than 2 ng/ml. All the collected oocytes were evaluated, regardless of the plasma progesterone levels.

Oocyte Recovery

At the end of the follicular phase (EF), preovulatory (larger than 35 mm in diameter) and nonpreovulatory follicles were punctured. During the follicular phase (DF) and during the luteal phase (DL), all follicles were nonpreovulatory.

From the 20 EF puncture attempts, 27 preovulatory follicles were flushed, and 14 COCs were collected. Four ovulations occurred before puncture. Averages of 0.7 COCs per attempt and 0.5 COCs per preovulatory follicle were obtained. From the 70 puncture attempts (20 EF, 20 DL, and 30 DF), 647 nonpreovulatory follicles were flushed and 320 COCs were recovered. An average of 4.6 COCs per attempt was obtained (4.0 in DF, 5.4 in EF, and 4.5 in DL). The recovery rate per follicle (0.5 COCs on average) was not significantly different between the three groups or between the follicular diameters (data not shown).

The interval between EF and DL was 8.7 days, and the interval between DL and EF was 13.7 days. Puncturing once at the end of the follicular phase and once during the luteal phase yielded a mean of 10.6 COCs (0.7 COCs from preovulatory follicles + (5.4 + 4.5) COCs from nonpreovulatory follicles) per 22 days, the duration of a cycle. The interval between two DF was 7.5 days. In 22 days, 3 DF could be performed, and 12 COCs (4 COCs per attempt) could be obtained.

Cumulus Aspect at Recovery

All 14 COCs collected from preovulatory follicles (larger than 35 mm in diameter) had an expanded cumulus at collection.

In the nonpreovulatory follicles, 6% (19 of 320) of the COCs had an expanded cumulus, 16% (52 of 320) were surrounded by a compact corona radiata, and 78% (249 of 320) were surrounded by a compact cumulus and cells from the follicular wall. No clear relationship between follicle size and cumulus aspect could be established. No significant difference in the portion of expanded COCs was observed according to group (Table 1). The percentage of corona radiata was significantly higher in the EF group (p < 0.05). Consequently, the percentage of compact cumulus and cells from the follicular wall was significantly lower in the EF group (p < 0.05).


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  		TABLE 1. Cumulus aspect of COCs from non-preovulatory follicles (< 35 mm) according to the group.

Nuclear Stage of Oocytes

All 33 expanded COCs were analyzed immediately at recovery. Of the 14 oocytes with an expanded cumulus that were collected from preovulatory follicles, 50% had completed meiosis (7 of 14 in metaphase II, 2 in metaphase I, 1 with dense chromatin, and 4 degenerated). Of the 19 oocytes with an expanded cumulus that were collected from nonpreovulatory follicles, 2 were in metaphase II, 3 were in metaphase I, 2 had dense chromatin, 6 were in germinal vesicle stage, and 6 were degenerated.

All 301 oocytes with a compact cumulus at recovery were cultured for 30 h. After in vitro culture, 100% of the COCs had an expanded cumulus; 299 oocytes were analyzed for nuclear maturation (two were lost during analysis), and 33% (99 of 299) had reached metaphase II. The maturation rate (oocytes in metaphase II) and the rate of oocytes able to resume or complete meiosis (oocytes with dense chromatin, in metaphase I, or in metaphase II) significantly increased with follicle diameter (p < 0.05 and p < 0.01, respectively); the latter was significantly lower in follicles 5–9 mm (38%, 32 of 84) than in follicles 10–34 mm (76%, 164 of 215; p < 0.01). Oocyte competence for in vitro maturation was influenced by the estrous cycle stage (Fig. 1). That is, the maturation rate was higher for the oocytes recovered at the end of the follicular phase than for the oocytes recovered during the luteal phase (51% [52 of 101] vs. 34% [30 of 88]; p < 0.05); the maturation rate of oocytes collected during the follicular phase was lower than for those from the EF and DL groups (15% [17 of 110]; p < 0.01). The percentage of degenerated oocytes did not vary with the diameter of the follicle of origin and was not significantly different between the 3 groups: 15% (17 of 110) in group DF, 9% (9 of 101) in group EF, and 10% (9 of 88) in group DL.



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  		FIG. 1. Maturation rate after in vitro culture of COCs from nonpreovulatory follicles according to stage of the estrous cycle. The maturation rates in group DF vs. EF and DF vs. DL are significantly different (p < 0.01). The maturation rates in group EF vs. DL are significantly different (p < 0.05). Values with different superscripts differ significantly: a,b and a,cp < 0.01; b,cp < 0.05.

Histone H1 Kinase Activity

A total of 190 oocytes were assayed individually for histone H1 kinase activity (171 oocytes in various nuclear stages after in vitro culture, 7 preovulatory oocytes in metaphase II, and 12 oocytes as immature controls). The expected band of phosphorylated histone H1 at 31 kDa was visualized for each oocyte analyzed (Fig. 2, A–C). The intensity of the band varied markedly between oocytes in different nuclear maturation stages and with the maturation conditions (in vitro vs. in vivo).



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  		FIG. 2. Representative profiles of histone H1 kinase activity in equine oocytes in different nuclear maturation stages and conditions. Only the Mr 32 000 prestained standard is visualized. A) Lane 1: one oocyte that remained in germinal vesicle stage after in vitro culture. Lane 2: One oocyte with dense chromatin after in vitro culture. Lane 3: one oocyte in metaphase I after in vitro culture. Lane 4: one oocyte in metaphase II after in vitro culture. B) Lane 1: one immature oocyte that was not cultured in vitro (negative control). Lane 2: one oocyte that remained in germinal vesicle stage after in vitro culture. C) Lane 1: one preovulatory oocyte that reached metaphase II in vivo. Lane 2: one oocyte in metaphase II after in vitro culture.

Quantitative analysis is illustrated in Figure 3. After in vitro culture, the histone H1 kinase activity was significantly lower in oocytes that remained in germinal vesicle or dense chromatin stages than in oocytes that reached metaphase I or metaphase II (Fig. 2A, lanes 1, 2, 3, and 4, respectively, and Fig. 3) (p < 0.05); the activity was not significantly different between oocytes with germinal vesicle and oocytes with dense chromatin or between metaphase I and metaphase II oocytes. No significant difference in histone H1 kinase activity was observed between the oocytes that remained in germinal vesicle stage after in vitro culture and the immature control, i.e. the immature oocytes that were not cultured in vitro (Fig. 2B, lanes 1 and 2, and Fig. 3). The histone H1 kinase activity was significantly higher in the preovulatory oocytes that reached metaphase II in vivo than in those that reached metaphase II after in vitro culture (Fig. 2C, lanes 1 and 2, and Fig. 3) (p < 0.001). Among the oocytes that reached a given nuclear stage, no significant difference in histone H1 kinase activity was observed between the follicular diameters or between the stages of the estrous cycle.



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  		FIG. 3. Quantitative analysis of histone H1 kinase activity in in vitro- and in vivo-matured equine oocytes at different stages of nuclear maturation. GV: Oocytes in germinal vesicle stage. DC: Oocytes with dense chromatin. MI: Oocytes in metaphase I. MII: Oocytes in metaphase II. IC: Immature control (immature oocytes that were not cultured in vitro). Results are presented as mean ± SEM. After in vitro culture, the histone H1 kinase activity was significantly lower in GV and DC than in MI and MII (p < 0.05); the activity was not significantly different between GV and DC, MI and MII, or GV and IC. The histone H1 kinase activity was significantly higher in preovulatory MII at recovery than in MII after in vitro culture (p < 0.001). Values with different superscripts differ significantly: a,bp < 0.05; b,cp < 0.01.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
The aim of this study was to investigate the meiotic competence of equine oocytes collected in different physiological stages and to analyze the MPF activity in oocytes after in vitro culture.

Oocytes were collected by using in vivo ultrasound-guided transvaginal puncture, which allows collection of a well-characterized population of oocytes and comparison of different populations within the same animal. In the present study, we collected 0.5 COCs per nonpreovulatory follicle punctured. There was no effect of the hormonal environment on recovery rate, which supports our previous work [4]. The two rhythms of successive punctures tested, one puncture at the end of the follicular phase plus one during the luteal phase, and repeated punctures during the follicular phase, provided 10–12 COCs per 22 days. Moreover, only 8 of 70 puncture attempts were not performed in the expected conditions. For 2 DL puncture attempts, the levels of progesterone were lower than 2 ng/ml, because no secretion of progesterone occurred after the previous EF puncture attempts. The puncture of preovulatory follicles usually results in an active corpus luteum [28]. However, the scraping of the follicular wall during the puncture may have led to damage in the granulosa cells. On the other hand, 6 puncture attempts during (DF) or at the end (EF) of the follicular phase were performed in the presence of high levels of progesterone. They all succeeded puncture attempts performed during the follicular phase, which can lead to luteinization of some follicles [25].

All COCs recovered from preovulatory follicles had an expanded cumulus at collection, and half of the enclosed oocytes had completed meiosis. Some COCs recovered from nonpreovulatory follicles had an expanded cumulus at collection, but only a few of the enclosed oocytes had completed meiosis. Similarly, all the compact COCs underwent cumulus expansion during in vitro culture, but not all oocytes completed meiosis, as previously described in mice [29], pigs [30], cattle [31], and horses [4]. Thus, in the horse, cumulus expansion in vivo as well as in vitro is not linked to oocyte nuclear maturation. Moreover, most COCs recovered from nonpreovulatory follicles had a compact cumulus; the cumulus aspect was not linked to the follicle size, in agreement with previous results [4, 32], but was influenced by stage of the estrous cycle. There was a higher percentage of compact corona radiata in group EF than in groups DL and DF that was not observed in our previous study [4]. No physiological reasons seem to explain this difference. Any comparison with findings in other reports is not reliable, as the classification according to morphology of the cumulus is highly variable from one report to another. Moreover, the morphology of the cumulus may be influenced by the differences in collection techniques.

In the present study, oocyte competence for in vitro maturation was influenced by the diameter of the follicle of origin and was higher at the end of the follicular phase than in luteal phase, again supporting our previous observations [4]. The majority of oocytes collected during the follicular phase before the emergence of the dominant follicle were not meiotically competent, in spite of the injection of exogenous gonadotropin. The growth of the dominant follicle induces atresia in subordinate ones but seems to increase the percentage of competent oocytes. A higher percentage of competent oocytes in follicles in early atresia has been suggested in cattle [33, 34] and has been recently shown in the horse [5]. These authors suggest that acquisition of meiotic competence is related to a loss of inhibiting activity by the degenerating follicle. On the other hand, the percentage of competent oocytes for in vitro maturation was higher for oocytes collected during luteal phase (group DL) than during follicular phase (group DF). Some factors secreted during luteal phase, such as progesterone, could promote acquisition of oocyte competence for in vitro maturation. As a result, puncturing once at the end of the follicular phase (EF) and once during the luteal phase (DL) yielded 10.6 COCs per 22 days; 4.3 of them were competent for in vitro maturation, and 0.4 of them (3.8% of total collected) reached metaphase II in preovulatory follicles. This latter pattern of repeated punctures provided the opportunity to increase the number of well-characterized oocytes collected per cycle and per mare.

MPF activity was analyzed in equine oocytes in relation to their stage of nuclear maturation and the maturation conditions. After in vitro culture, MPF activity was significantly lower in oocytes that remained in the germinal vesicle or dense chromatin stages than in oocytes that reached metaphase I or metaphase II. This is consistent with findings in goat [6] and pig [35] oocytes. Moreover, we observed that the oocytes that remained in germinal vesicle stage after in vitro culture displayed H1 kinase activity levels comparable to those of immature oocytes that were not cultured in vitro. Similar results were obtained in the pig [35]. During oocyte maturation, in mammalian species studied so far, MPF activity displays a low level at the germinal vesicle stage, increases progressively at the germinal vesicle breakdown stage to reach maximum level in metaphase I oocytes, decreases sharply during the anaphase-telophase I transition, and increases thereafter to a high level in metaphase II oocytes (mice [14, 15], rabbits [16], pigs [17], goats [18], and cattle [19]). Histone H1 kinase activity has been suggested to cause nuclear lamina disassembly [36], nucleolar disassembly [37], chromosome condensation [38], microfilament rearrangement [39], and reorganization of the intermediate filament network [40]. Thus, one can speculate that, in incompetent oocytes, a lasting low MPF activity may prevent occurrence of the events that constitute meiotic maturation, i.e., nuclear envelope breakdown, chromosome condensation, and spindle assembly. One could notice that the oocytes with dense chromatin after in vitro culture displayed H1 kinase activity levels comparable to those of oocytes with germinal vesicle. Though the dense chromatin is the beginning of the chromosome condensation, no increase in the H1 kinase activity was yet detectable. The nuclear lamina disassembly occurred around the time of chromatin condensation, but was not detectable with the technique we used.

After in vitro culture, the MPF activity was not significantly different in equine oocytes that reached metaphase I than in oocytes in metaphase II. This is consistent with findings in mice [14, 15], rabbits [16], pigs [17], goats [18], and cattle [19]. Moreover, in these species, as noticed above, the transition between metaphase I and metaphase II is correlated with a transient drop in MPF activity. Therefore, the incompetence of equine oocytes to go beyond metaphase I could be explained by a persistent high MPF activity. Further studies will be necessary to determine the factor(s) responsible for the blocking in metaphase I of the equine oocytes: factor(s) that regulate synthesis and degradation of the MPF components, MPF phosphorylation regulators such as the phosphatase cdc25 [41], or other kinases involved in the regulation of meiotic events such as mitogen-activated protein kinase [42].

In this study, the histone H1 kinase activity was significantly higher in the preovulatory oocytes that reached metaphase II in vivo than in the oocytes that reached metaphase II after in vitro culture. To our knowledge, this is the first comparison of histone H1 kinase activity between in vivo- and in vitro-matured oocytes from mammalian species. The lower activity could be a sign of incomplete cytoplasmic maturation after in vitro culture. Actually, in porcine oocytes, a lower male pronucleus formation ability is associated with a lower histone H1 kinase activity [43]. Moreover, we have showed previously that meiotic spindles obtained after in vitro culture are significantly wider and longer than spindles obtained from oocytes at recovery [4]; this could be due to a change in concentration of ooplasmic proteins, such as tubulin.

In conclusion, this study confirms that the acquisition of meiotic competence in equine oocytes occurs progressively during antral follicle growth and demonstrates that oocyte competence is influenced by stage of the estrous cycle. We established an assay system for MPF activity in equine oocytes and showed that oocyte competence is associated with histone H1 kinase activity. This is the first report on biochemical analysis of cell cycle regulation in equine oocytes.


   ACKNOWLEDGMENTS
 
We wish to thank Dr. Laurence Gall and Véronique de Smedt (Institut National de la Recherche Agronomique, Jouy-en-Josas, France) for their kind advice for the histone H1 kinase activity assay, and Monique Ottogalli and Dr. Christine Briant for generous supplies of crude equine gonadotropin. We are grateful to Isabelle Couty and the staff of the experimental stud for technical assistance and Alain Beguey and Odile Moulin for photographic work.


   FOOTNOTES
 
1 This work was supported by grants from the Institut National de la Recherche Agronomique, France, and the Haras Nationaux, France. G.G. was supported by a fellowship from the Institut National de la Recherche Agronomique, France, and the Région Centre, France. Back

2 Correspondence. FAX: 33.2.47.42.77.43; gerard{at}tours.inra.fr Back

Accepted: April 10, 1998.

Received: January 6, 1998.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

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