1 Centre for Early Human Development, Institute of Reproduction and Development, Level 5, Monash University, Monash Medical Centre, 246 Clayton Road, Clayton, Victoria, Australia 3168 and 2 Department of Obstetrics and Gynaecology, National University of Singapore
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Abstract |
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Key words: 6-dimethylaminopurine/histone H1 kinase/human oocytes/in-vitro maturation
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Introduction |
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Meiotic and developmental competence are established during the period of oocyte growth preceding nuclear maturation. This phase of oocyte development is characterized by the synthesis and storage of RNA and translational products (Rossant and Pederson, 1986) which are subsequently used for meiotic and early embryonic developmental events (Kastrop et al., 1991
; Wickramasinghe and Albertini, 1993
; Fair et al., 1995
).
In vivo, each human menstrual cycle is characterized by the recruitment and growth of numerous follicles. One or two selected follicles continue growth until the day of ovulation, whilst most follicles undergo atresia, terminating their growth. By the time of ovulation the dominant follicle measures approximately 20 mmol/l in diameter and may have been growing for up to 14 days (Calderon and Healy, 1993). Human oocytes obtained for in-vitro maturation are aspirated from 212 mmol/l antral follicles early in the follicular phase of the menstrual cycle and are matured for only 48 h in culture.
Hence, at the time of aspiration for in-vitro maturation, human oocytes have not fully completed their phase of oocyte growth and may be retrieved during the process of atresia. Therefore, all human oocytes retrieved for in-vitro maturation have an artificially truncated growth phase. Consequently, it is possible that the maturational and developmental anomalies observed in in-vitro matured oocytes are attributable to their shortened growth phase and thus the inability to complete all the necessary transcriptional and translational changes required for complete maturation and developmental competence. Thus, extending the pre-maturation growth phase in vitro may benefit the embryonic development of in-vitro matured oocytes (Longeran et al., 1997). Since mammalian oocytes spontaneously mature upon liberation from the follicle, extending the growth phase requires manipulating the onset of meiotic maturation. The molecular control of meiotic maturation has been extensively studied in a variety of different species and it is believed to be controlled by M-phase promoting factor (MPF). MPF is a proteinaceous factor in the oocyte cytoplasm. Purification of MPF has shown that it is composed of p34cdc2 and cyclin B (Gautier et al., 1988
) and in its active form this heterodimer has p34cdc2 kinase activity that may be measured as histone H1 kinase activity (Arion et al., 1988
). Histone H1 kinase activity is responsible for entry into M-phase and the cascade of events associated with meiotic maturation. 6-Dimethylaminopurine (DMAP), a serine threonine protein kinase inhibitor, prevents spontaneous germinal vesicle breakdown by inhibiting the activation of histone H1 kinase, but does not interfere with protein synthesis (Rime et al., 1989
; Fulka et al., 1991
). Previous studies in rodents and larger domestic species has revealed that 6-DMAP can prevent the resumption of meiotic maturation in immature oocytes (Rime et al., 1989
; Motlík and Kubelka, 1990
; Fulka et al., 1991
; Szöllösi et al., 1991b
). However, few reports have described the fertilization and embryonic development of oocytes following DMAP treatment.
The present study examined whether DMAP treatment of immature oocytes could improve subsequent embryonic development. It was postulated that DMAP treatment might improve oocyte developmental competence by allowing additional transcription and translation to occur in the oocyte during the extended pre-maturation period.
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Materials and methods |
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Immature oocytes were also retrieved from 20-day-old pre-pubertal F1 female mice that had either received, or not received, priming with 5 IU PMSG 48 h prior to oocyte retrieval. The immature oocytes from the 20-day-old pre-pubertal mice were also retrieved according to previously described methods (Anderiesz and Trounson, 1995).
Ovulated metaphase II mouse oocytes were used as controls and were obtained by administering 5 IU human chorionic gonadotrophin (HCG; Chorulon, Intervet, Castle Hill, NSW, Australia) 48 h after PMSG. The animals were killed 1214 h after HCG injection and the mature metaphase II OCC were released from the oviducts into M2 medium (Quinn et al., 1982).
Immature human oocytes were aspirated from the ovaries of women that had not received exogenous gonadotrophin stimulation. The details and methods of recovery of human immature oocytes by ultrasound-guided aspiration of 212 mmol/l follicles has been described previously (Trounson et al., 1994).
Inhibition of meiotic resumption with DMAP
The medium used for mouse oocyte culture was Eagle's Minimal Essential Medium alpha modification (-EMEM; Sigma, St Louis, MO, USA) with 10% fetal calf serum (FCS; Commonwealth Serum Laboratories, Melbourne, VIC, Australia) and 0.2 IU ovine follicle stimulating hormone [FSH (o-FSH; Horizon Technology, Roseville, NSW, Australia)]. Immediately after removal from the ovarian follicle, groups of up to 15 mouse oocyte cumulus complexes (OCC) were cultured in 20 µl drops of
-EMEM medium supplemented with 2 mmol/l DMAP (Sigma) under light white mineral oil (Sigma) at 37°C in an atmosphere of 5% CO2 in air for 7 h. The effect of DMAP on the meiotic inhibition of mouse oocytes was assessed after 3.5 and 7 h. At these time points random samples of DMAP-treated mouse oocytes were collected and placed in M2 medium containing 40 IU hyaluronidase (type IV-S, Sigma) and the cumulus and corona radiata cells were removed by gentle micropipetting. Mouse oocytes were examined for an intact germinal vesicle (GV) or indications of germinal vesicle breakdown (GVBD) under interference contrast microscopy. Those mouse oocytes with an intact GV were washed several times in fresh culture medium and transferred to either histone kinase lysis buffer (Collas et al., 1993
) for histone H1 kinase analysis or Laemmli sample buffer (Laemmli, 1970
) for either the identification of proteins by SDSPAGE or Western blot analysis of cyclin B and p34cdc2. To confirm the persistence of DMAP inhibition, a third group of mouse oocytes was maintained in DMAP-supplemented medium for 18 h and subsequently examined for the presence of a GV by interference contrast microscopy.
The medium used for human oocyte culture was Chang's medium (Irvine Scientific, Santa Ana, CA, USA) supplemented with 0.1 IU of human recombinant FSH/ml (hr-FSH) (Gonal-F; Serono, Geneva, Switzerland) and 0.1 IU human recombinant luteinizing hormone (LH)/ml (hr-LH; Serono, Geneva, Switzerland). Directly after aspiration, groups of up to three human OCC were cultured for 24 h in Chang's medium supplemented with 2 mmol/l DMAP in 10 µl drops under light white mineral oil in bacteriological grade sterile Petri dishes (Nunclon, Roskilda, Denmark) at 37°C in humidified 5% CO2 in air. After 24 h, random samples of human oocytes were collected and the adherent cumulus and coronal cells removed as previously described. Human oocytes were examined for an intact GV or GVBD after first staining with Hoechst 33342 (50 µg/ml; Sigma) for 5 min and examination by interference contrast microscopy and epifluorescence to confirm interphase chromatin. Those human oocytes displaying an intact germinal vesicle were washed in fresh culture medium and transferred to histone kinase lysis buffer for histone H1 kinase analysis.
Reversibility of inhibition and kinetics of meiotic maturation
The reversibility of meiotic inhibition and the subsequent timing of meiotic maturation of mouse oocytes were examined after 7 h of culture in DMAP. DMAP-treated mouse oocytes were washed thoroughly in -EMEM and cultured in 20 µl drops of
-EMEM supplemented with 10% FCS and 0.2 IU o-FSH under light white mineral oil for a further 18 h at 37°C in a humidified atmosphere of 5% CO2 in air. Mouse OCC not exposed to DMAP were used as in-vitro matured controls and were concurrently cultured in the
-EMEM maturation medium supplemented with 10% FCS and 0.2 IU o-FSH for 18 h. Progression of meiotic maturation in both DMAP-treated and untreated oocytes was monitored at 6, 8, 10 and 12 h of culture. At these time points, mouse oocytes were randomly selected and denuded of follicle cells in hyaluronidase solution, as previously described. Cumulus denuded oocytes were then placed in a 1% sodium citrate solution for 5 min, transferred to a glass microscope slide, and then fixed using a 3:1 solution of methanol and acetic acid. Oocytes were then stained using 10% Giemsa stain (British Drug House, Kilsyth, Vic, Australia) to visualize the chromosomes and determine the progression of nuclear maturation. Additionally, following the 18 h of in-vitro maturation, random samples of mouse oocytes were denuded of cumulus and coronal cells and examined by interference contrast microscopy for meiotic progression. Those oocytes that had progressed to metaphase II were transferred to either histone kinase lysis buffer or Laemmli sample buffer for either histone H1 kinase analysis or the identification of proteins by SDSPAGE.
The reversibility and timing of meiotic maturation in human oocytes exposed to DMAP for 24 h was tested by culturing DMAP treated oocytes for a further 48 h in vitro. Human OCC not exposed to DMAP were concurrently matured in vitro for 48 h as an in-vitro matured control. DMAP treated and control human oocytes were thoroughly washed in Chang's medium and transferred to 10 µl Chang's medium supplemented with 0.1 IU hr-FSH and 0.1 IU hr-LH. Oocytes were cultured at 37°C in a humidified atmosphere of 5% CO2 in air under mineral oil. Oocytes were examined for the presence of a germinal vesicle or extruded first polar body at 24, 30, 36 and 48 h of culture. At the end of 48 h culture, mature metaphase II oocytes were denuded of all follicle cells in hyaluronidase, as previously described and then transferred to histone kinase lysis buffer for histone H1 kinase analysis.
In-vitro fertilization of mouse and human oocytes
Mouse oocytes were fertilized in vitro (IVF) using spermatozoa retrieved from the cauda epididymis of two mature F1 (C57 BL/6 J WEHI femalexCBA/CaH WEHI male) male mice. The details and methods of mouse IVF have been previously described by Anderiesz and Trounson (1995). Successful fertilization was assessed by the appearance of two pronuclei and the extrusion of the second polar body 810 h after insemination.
In-vitro matured human oocytes were fertilized by intracytoplasmic sperm microinjection (ICSI) (Ng et al., 1991). Oocytes were examined under interference contrast microscopy, 1618 h after ICSI, for the presence of two pronuclei and the second polar body.
Embryonic development following IVF
Mouse zygotes were cultured for 96 h in 20 µl drops of M16 (Quinn et al., 1982) supplemented with 4 mg/ml BSA (Pentex crystallized, Miles, Kankakee, IL, USA) under light white mineral oil in humidified 5% CO2 in air. Development to blastocysts was recorded on day 4 post-insemination.
Pronuclear human oocytes were cultured in 4-well culture dishes (Nunclon) in modified human tubal fluid medium (IVF-50, Scandinavian IVF Science AB, Gothenburg, Sweden) without somatic cell support for 2 days after ICSI. From day 2 (24 cell stage) to day 5/6 (blastocyst stage) of culture, embryos were co-cultured on a monolayer of Vero cells (Fong et al., 1997). On the morning of day 3, the medium was changed to a 1:1 combination of 50% IVF-50 and 50% Hatch-50 (Scandinavian IVF Science AB) and 10% patients' own serum. On the morning of day 4, the medium was changed to 100% Hatch-50 supplemented with 10% patients' own serum, and embryos were cultured for a further 2 days. All cultures were carried out at 37°C in sealed humidified chambers with 5% CO2, 5% O2 and 90% N2.
Measurement of histone H1 kinase
Histone H1 kinase activity in mouse and human oocytes was determined by methods already described (Collas et al., 1993) using reagents from Sigma unless otherwise stated. Briefly, groups of 10 follicular cell denuded mouse oocytes, or single denuded human oocytes, were lysed by repeated freezing in liquid nitrogen then thawing in a volume of 10 µl of histone kinase lysis buffer [80 mmol/l glycerophosphate, 20 mmol/l EGTA, 15 mmol/l MgCl2, 1 mmol/l dithiothreitol (DTT), 10 µg/ml aprotinin, 10 µg/ml leupeptin, 10 µl/ml pepstatin and 50 nmol/l cyclic adenosine monophosphate-dependent protein kinase inhibitor]. The kinase reaction was initiated by the addition of 2.5 mg/ml histone H1, 0.7 mmol/l ATP and 12.5µ Ci of (
-32 P) ATP (Amersham, Buckinghamshire, UK). The reaction was carried out for 15 min at ambient temperature (2025°C) and was stopped by the addition of 5 µl of four times concentrated sample buffer (Laemmli, 1970
). The samples were then boiled for 3 min. Proteins were separated on a 12% polyacrylamide gel (SDSPAGE) as described (Laemmli, 1970
). Subsequently the gels were exposed to a Fuji phosphor screen (Fuji Photofilm Co. Ltd, Japan) overnight at room temperature. Histone H1 kinase activity was assessed by scanning the screen using a Fuji Bas 1000 Phosphorimage analyser using Fuji Bio-imager analyser MacBas Version 2.X software.
Immunoblotting and immunodetection of cyclin B and P34cdc2
Immunoblotting was used to identify the two components of MPF, cyclin B and p34cdc2. Groups of 30 follicular cell denuded murine oocytes were collected: (i) immediately after liberation from the follicle (GV stage), (ii) after 7 h of DMAP treatment (GV stage), (iii) after completion of meiotic maturation in vitro (18 h of culture without DMAP, MII stage), (iv) after completion of meiotic maturation in vitro following 7 h of DMAP exposure (metaphase II stage), and (v) after ovulation (metaphase II stage, in vivo control). In addition, to identify the direct effect of priming on MPF activation, cyclin B and p34cdc2 were immunodetected in 30 GV stage oocytes retrieved from day 20 pre-pubertal mice that had either received, or did not receive, priming with 5 IU PMSG.
The immunoblotting procedure entailed the solubilization of the groups of 30 murine oocytes in 10 µl of Laemmli sample buffer followed by one-dimensional SDS gel electrophoresis of oocyte proteins (Laemmli, 1970). Total oocyte proteins were electrophoretically transferred using a tank transfer system (Bio Rad, Hercules, CA, USA) at 80 V for 1 h. Proteins were transferred to a polyvinylidenedifluoride (PVDF) membrane (Immobilon-P; Millipore, Bedford, MA, USA) and the membranes were probed using anti-human cyclin B1 mouse monoclonal IgG2 b antibody (Upstate Biotechnology, Lake Placid, NY, USA) for the detection of the cyclin B antigen and anti-human cdc2 kinase (PSTAIR) rabbit polyclonal IgG antibody (Upstate Biotechnology), for the detection of the p34cdc2 antigen. The membranes were subsequently washed and incubated with a secondary antibody conjugated with horseradish peroxidase (HRPO) (Silenus, Hawthorn, Victoria, Australia) and antigen visualization was performed using enhanced chemiluminescent Western blotting detection reagents (Amersham, Buckinghamshire, UK).
Determination of mouse oocyte proteins
Mouse oocyte proteins were separated by 1-dimensional SDSPAGE. Groups of five follicular cell denuded oocytes were collected: immediately after liberation from the follicle, after 3.5 and 7 h of DMAP treatment, after meiotic maturation to metaphase II with and without prior DMAP exposure and after ovulation (in-vivo control). The groups of oocytes were solubilized by boiling for 3 min in 10 µl Laemmli sample buffer.
SDSPAGE was performed using a 12% (w/v) polyacrylamide gel (Laemmli, 1970). The gels were run at 150 V for 1 h. After electrophoresis the gels were silver stained (Heukeshaven and Dernick, 1988
) and air-dried overnight between two sheets of acetate paper (Promega, Annandale, NSW, Australia) at room temperature.
Statistical analyses
Replicate data were analysed for homogeneity and results are presented as mean ± SEM. A minimum of three replicates was performed for every test and variability amongst replicates was examined. No variability amongst replicates existed (2 = 0.235 on 2 d.f). Resumption of maturation, progression of germinal vesicle breakdown, and kinetics of polar body extrusion were analysed using the
2 test for comparisons between DMAP-treated and untreated (control) groups. P < 0.05 was accepted as the level of statistical significance. Fertilization and embryonic data were analysed using linear logistic regression. Hypotheses were tested using the Log likelihood Ratio Statistic (LRS) which was
2 distributed.
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Results |
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After 24 h culture, 81% of immature human oocytes are known to progress to GVBD (Trounson et al., 1994, 1998
). The presence of 2 mmol/l DMAP effectively inhibited GVBD in 83% (n = 12) of human oocytes over a 24-h culture period (Table I
).
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The proportion of DMAP-treated human oocytes that completed meiotic maturation after withdrawal of DMAP was similar to that of in-vitro matured controls (Table I). Furthermore, the kinetics of polar body extrusion was similar in DMAP treated and untreated human oocytes (Table II
). Thus, DMAP can temporarily inhibit mouse and human meiotic maturation without effecting subsequent maturation to metaphase II.
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There was no significant difference between the fertilization rates for human oocytes after withdrawal of DMAP (77%) and control, untreated oocytes (75%) (Table I). Hence the temporary inhibition of germinal vesicle breakdown with DMAP had no effect on the ability of human or mouse oocytes to successfully fertilize in vitro.
Embryonic development
Embryonic development was assessed in mouse oocytes over a 4-day culture period post-insemination. The development of embryos to blastocysts was used as an indicator of developmental competence. Development to blastocyst was significantly higher (P < 0.001) when mouse oocytes were matured in vivo (65%, n = 114) than in vitro (19%, n = 118). Furthermore, the temporary inhibition of the resumption of meiosis with DMAP resulted in a significant reduction (P = 0.002) in development to the blastocyst stage in culture (6%, n = 108).
Human embryonic developmental competence was assessed by monitoring cleavage over a 5 day culture period. Table III shows the cell number of DMAP treated and control human embryos culture over a 5-day period. There was no apparent effect of DMAP treatment on subsequent embryonic cleavage rate or cell number of in-vitro matured and fertilized oocytes.
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Discussion |
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The ability to reversibly inhibit meiotic maturation in human oocytes has never previously been reported. These findings demonstrate that concentrations effective at preventing meiotic resumption in other species (Motlík and Rimkevicová, 1990; Fulka et al., 1991
) effectively prevent GVBD in 83% of human oocytes for 24 h. The temporary inhibition of nuclear maturation was fully reversible and DMAP-treated human oocytes matured to metaphase II in the same numbers and with the same maturation kinetics as non-treated oocytes.
Similarly, 2 mmol/l DMAP was shown to be capable of preventing meiotic maturation in mouse oocytes, confirming previous reports (Rime et al., 1989; Szöllösi et al., 1991b
) and subsequent maturation to metaphase II, after 18 h, was similar in DMAP-treated and untreated oocytes. Interestingly, the kinetics of meiotic resumption was altered in the DMAP-treated oocytes. Proteins with a rapid turnover rate are involved in the spontaneous maturation of rodent oocytes (Downs, 1990
) and, during prolonged periods of meiotic arrest, these short-lived proteins are degraded (Ekholm and Magnusson, 1979
). Furthermore, other studies (Motlík and Rimkevicová, 1990
) demonstrated that mouse oocytes preincubated with DMAP require active protein synthesis for the resumption of GVBD. Thus, the time taken for the resynthesis of proteins required for GVBD may be responsible for the slower resumption of meiotic maturation observed in DMAP treated oocytes.
Although meiotic resumption to the GVBD stage occurs spontaneously in rodent oocytes in vitro, the meiotic progression of oocytes from the GVBD to the metaphase II stage requires active protein synthesis (Shultz and Wassarman, 1977; Downs, 1990
). In the presence of DMAP, translationary activity continues in mouse oocytes (Rime et al., 1989
). Consequently, proteins required for the completion of meiotic maturation may be synthesized and stored in the ooplasm during DMAP treatment. In addition, the acceleration of morphological events such as chromatin condensation and microtubule formation following DMAP treatment has been previously demonstrated in activated mouse oocytes (Szöllösi et al., 1993
) and was attributable to the inhibition of specific kinases (Moreno and Nurse, 1990
). Thus, the immediate utilization of translated and stored proteins together with kinase inhibition may account for the acceleration of meiotic progression from GVBD to metaphase II observed in DMAP-treated mouse oocytes.
The failure of human oocytes to display accelerated maturation kinetics may be due to the inability of oocytes to generate and accumulate all the proteins required for meiotic maturation during the 24-h period of meiotic inhibition with DMAP. That is, a longer period may be required by human oocytes to translate all the proteins required for meiotic resumption and progression.
Histone H1 kinase activity was qualitatively assessed in DMAP-treated and control, GV and metaphase II-stage oocytes. Cyclin B and p34cdc2, the two components of MPF, exist in a dimerized, fully phosphorylated and non-active form in the cytoplasm of growing oocytes as pre-MPF (Gautier and Maller, 1991; Christmann et al., 1994
). However, the association of p34cdc2 with cyclin B is insufficient for activation of kinase activity (Jeffrey et al., 1995
). The activation of H1 kinase activity is associated with the p34cdc2 component of the MPF heterodimer (Choi et al., 1991
; de Vantéry et al., 1996
) and the activation of kinase activity is dependent on the association of p34cdc2 with cyclin B and the subsequent dephosphorylation of its two major residues, tyrosine 14 and threonine 15, by a cdc 25 phosphatase (Maller, 1994
). Histone H1 kinase activity has previously been shown to exist in GV stage porcine and murine oocytes (Naito and Toyoda, 1991
; Choi et al., 1991
; Naito et al., 1992
; Christmann et al., 1994
; Gavin et al., 1994
). Therefore, histone H1 kinase activity can be detected in the cytoplasm of oocytes prior to the breakdown of the GV, and then increases to high concentrations at metaphase I. The activity decreases during anaphase and telophase and increases again to a high concentration at metaphase II (Doree et al., 1983
; Gerhart et al., 1984
). In the current study, histone H1 kinase activity was present in mouse GV stage oocytes retrieved directly from the ovaries of PMSG-treated mice and the kinase activity was maintained in the GV stage oocytes for the duration of DMAP treatment. Immunodetection of cyclin B and p34cdc2 in GV stage DMAP-treated and freshly isolated control, PMSG primed mouse oocytes revealed the presence of both cyclin B and p34cdc2. Furthermore, p34cdc2 was present in its most dephosphorylated and active form, confirming the existence of histone H1 kinase activity in this population of oocytes. The observation of premature chromosome condensation is also consistent with the presence of histone H1 kinase activity and has been previously observed in DMAP-treated oocytes (Szöllösi et al., 1991b
). Priming with PMSG appears to be responsible for activating histone H1 kinase activity in GV stage oocytes, as evidenced from the appearance of histone H1 kinase activity, increased concentrations of cyclin B, and the stimulated generation of p34cdc2 in its dephosphorylated and active form in the GV oocytes of pre-pubertal mice. Interestingly, GV stage mouse oocytes that expressed histone H1 kinase activity did not undergo subsequent meiotic maturation. It has been suggested (de Vantéry et al. 1996
) that a critical threshold concentration of p34cdc2 has to be reached for meiotic maturation to be spontaneously triggered. Thus, the inability of these mouse oocytes to undergo GVBD may be due to sub-threshold concentrations of p34cdc2.
Interestingly, at the time of retrieval, GV stage oocytes from patients 1 and 2 exhibited histone H1 kinase activity. Histone H1 kinase activity was not expected in the GV stage oocytes as these populations of oocytes were retrieved from women that were not exposed to any exogenous gonadotrophin treatment prior to oocyte aspiration. Oocyte aspiration is a non-selective retrieval process that results in the collection of oocytes from the entire cohort of follicles. Considering that only one follicle reaches dominance during the human menstrual cycle, whilst the other follicles become atretic, the majority of oocytes collected from these patients would have been rescued from follicles either destined for or in the process of follicular atresia. Whilst incapable of supporting complete nuclear maturation (Gougeon and Testart, 1986), follicular atresia has been implicated in modifying GV chromatin and eliciting the resumption of meiotic maturation in oocytes (Lefevre et al., 1989
). The maturational changes associated with atresia are believed to be due, in part, to a decline or removal of inhibitory factors (Channing et al., 1982
; Gougeon and Testart, 1986
) resulting from atresia related follicular cell death (Channing et al., 1982
). Since all the oocytes assessed from patients 1 and 2 were homogeneous in their expression of histone H1 kinase activity, at the time of oocyte retrieval, it is possible that these oocytes were derived from follicular cohorts in the process of atresia. Consequently, the kinase activity would be due to follicular atresia.
In contrast, the consistent lack of kinase activity in the freshly aspirated oocytes from patient 3 may be due to the retrieval of oocytes from a cohort of viable, growing follicles, prior to the onset of atretic changes.
In keeping with previous findings (Rime et al., 1989; Szöllösi et al., 1993
) we have reported that once histone H1 kinase activity is present, DMAP is unable to inactivate the histone H1 kinase activity. The inability of DMAP to inactivate the active kinase activity is attributable to the prevention of cyclin degradation and the protection of the cdc2 kinase from inactivation (Felix et al., 1990
). However, in the presence of MPF activity, DMAP prevented the germinal vesicle breakdown and progression of meiotic maturation. Hunt proposed that, once activated by dephosphorylation, the kinase activity of p34cdc2 could control its own subsequent dephosphorylation, leading to the auto-amplification of MPF (Hunt, 1989
). Furthermore, it has been suggested that DMAP prevents the auto-amplification of MPF (Jessus et al., 1991
). Thus, the inhibition of complete meiotic maturation by DMAP in the presence of histone H1 kinase activity may be due to the prevention of auto-amplification of MPF.
In comparison, if histone H1 kinase activity is not present in the oocyte, DMAP can successfully prevent the appearance of the kinase by preventing the dephosphorylation of p34cdc2 (Jessus et al., 1991) for the duration of DMAP exposure.
All the DMAP-treated human and mouse oocytes that matured to metaphase II expressed histone H1 kinase activity, demonstrating that short-term meiotic inhibition and DMAP exposure does not effect cell cycle regulation.
Interestingly, one-third of the in-vitro matured metaphase II control oocytes from patient 3 did not exhibit kinase activity. High concentrations of MPF activity are maintained by cytostatic factor (CSF) in vivo (Masui and Market, 1971). Therefore, the lack of kinase activity in these metaphase II stage oocytes may be due to a defect in CSF or may indicate the presence of a parallel pathway for meiotic maturation, possibly via the activation of p42 MAP kinase (Gavin et al., 1994
).
The reduced development to blastocyst of DMAP-treated mouse oocytes was obviously not related to MPF activation at metaphase II or the protein content of mature oocytes. DMAP is a non-physiological inhibitor involved in kinase and phosphorylation inhibition. Phosphorylation plays an important role in the post-translational modification of proteins and the action of kinases is similarly associated with a plethora of subcellular responses. Thus, the reduction in mouse blastocyst development may be due to the inhibition of different kinase activities and the modification of proteins that participate in embryogenesis and normally function in a phosphorylated state. Furthermore, these changes in phosphorylation may be below the detection limits of silver nitrate staining and consequently not observed on our gels. In addition, some disorganization of meiotic spindles, abnormalities in microtubule organizing centres, and damage to kinetochores and microtubules have been observed in DMAP-treated oocytes (Rime et al., 1989; Szöllösi et al., 1991b
). Cell cycle progression involves cyclin degradation and this occurs by a ubiquitin-dependent pathway requiring intact microtubules (Glotzer et al., 1991
; Kubiak et al., 1993
). It is therefore possible that the reduced development to blastocyst is attributable to slowing and eventual cessation of cell cycle progression due to inhibited cyclin degradation.
Artificially extending the prematuration period with DMAP neither positively nor negatively effected either the fertilization or early cleavage of human embryos. The developmental retardation and blockage of human embryos in the present experiments was similar to that observed for oocytes under a wide range of conditions in vitro (Barnes et al., 1996; Trounson et al., 1996
, 1998
). It is evident that in-vitro matured human oocytes are developmentally compromised and the nature of the developmental deficiencies is still speculative. Despite the lack of improvement in embryonic development following DMAP exposure, the extension of the prematuration period of oocyte growth may still represent a feasible technique for the improvement of embryonic developmental competence. However, a more specific or physiological meiotic inhibitor would be required.
The results of this study demonstrate for the first time that it is possible to reversibly inhibit meiotic maturation in human oocytes with DMAP with no apparent effect on embryonic development. Furthermore, the ability to regulate human oocyte maturation provides us with a powerful tool to further investigate the mechanisms of human oocyte maturation and the acquisition of developmental competence.
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Acknowledgments |
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Notes |
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References |
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Arion, D., Meijer, L., Brizuela, L. et al. (1988) cdc2 is a component of the M phase specific histone H1 kinase: evidence for identity with MPF. Cell, 55, 371378.[ISI][Medline]
Barnes, F.L., Kausche, A., Tiglias, J. et al. (1996) Production of embryos from in vitro-matured primary human oocytes. Fertil. Steril., 65, 11511156.[ISI][Medline]
Calderon, I. and Healy, D. (1993) Endocrinology of IVF. In Trounson, A. and Gardner, D.K. (eds), Handbook of In vitro Fertilisation. CRC Press, Boca Raton, Florida, pp. 116.
Cha, Y.K., Koo, J.J., Ko, J.J. et al. (1991) Pregnancy after in vitro fertilisation of human follicular oocytes collected from nonstimulated cycles, their culture in vitro and their transfer in a donor oocyte program. Fertil. Steril., 55, 109113.[ISI][Medline]
Channing, C.P., Pomerantz, S.H., Bae, I.H. et al. (1982) Actions of hormone and other factors upon oocyte maturation. Adv. Exp. Med. Biol., 147, 189210.[ISI][Medline]
Choi, T., Aoki, F., Mori, M. et al. (1991) Activation of p34 cdc2 protein kinase activity in meiotic and mitotic cell cycles in mouse oocytes and embryos. Development, 113, 789795.[Abstract]
Christmann, L., Jung, T., and Moor, R. (1994). MPF components and meiotic competence in growing pig oocytes. Mol. Reprod. Dev., 38, 8590.[ISI][Medline]
Collas, P., Sullivan, E.J. and Barnes, F.L. (1993) Histone H1 kinase activity in bovine oocytes following calcium stimulation. Mol. Reprod. Dev., 34, 224231.[ISI][Medline]
de Vantéry, C., Gavin, A.C., Vassalli, J.D. and Schorderet-Slatkine, S. (1996) An accumulation of p34 cdc2 at the end of mouse oocyte growth correlates with the acquisition of meiotic competence. Dev. Biol., 174, 335344.[ISI][Medline]
Doree, M., Peaucellier, G. and Picard, A. (1983) Activity of maturation-promoting factor and the extent of protein phosphorylation oscillate simultaneously during meiotic maturation of starfish oocytes. Dev. Biol., 99, 489501.[ISI][Medline]
Downs, S.M. (1990) Protein synthesis inhibitors prevent both spontaneous and hormone dependent maturation of isolated mouse oocytes. Mol. Reprod. Dev., 27, 235243.[ISI][Medline]
Edwards, R.G. (1965) Maturation in vitro of mouse, sheep, cow, pig, rhesus monkey and human ovarian oocytes. Nature, 208, 349351.[ISI][Medline]
Ekholm, C. and Magnusson, C. (1979) Rat oocyte maturation: effects of protein synthesis inhibitors. Biol. Reprod., 21, 12871293.[ISI][Medline]
Fair, T., Hyttel, P. and Greve, T. (1995) Bovine oocyte diameter in relation to maturational competence and transcriptional activity. Mol. Reprod. Dev., 42, 437442.[ISI][Medline]
Felix, M.-A, Labbé, J.-C,. Dorée, M. et al. (1990) Triggering of cyclic degradation in interphase extracts of amphibian eggs by cdc2 kinase. Nature, 346, 379382.[ISI][Medline]
Fong, C.Y., Bongso, A., Ng, S.C. et al., (1997) Ongoing pregnancy after transfer of zona-free blastocysts: implications for embryo transfer in the human. Hum. Reprod., 12, 557560.[ISI][Medline]
Fulka, A, Jr. Leibfried-Rutledge, M.L. and First, N.L. (1991) Effect of 6-dimethylaminopurine on germinal vesicle breakdown of bovine oocytes. Mol. Reprod. Dev., 29, 379384.[ISI][Medline]
Gautier, J., Norbury, C., Lohka, M. et al. (1988) Purified maturation-promoting factor contains the product of a Xenopus homolog of the fission yeast cell cycle control gene cdc2+. Cell, 54, 433439.[ISI][Medline]
Gautier, J. and Maller, J.L.(1991). Cyclin B in Xenopus oocytes: implications for the mechanisms of pre-MPF activation. EMBO J., 10, 177182.[Abstract]
Gavin, A.C., Cavadore, J.C. and Schorderet-Slatkine, S. (1994) Histone H1 kinase activity, germinal vesicle breakdown and M phase entry in mouse oocytes. J. Cell Sci., 107, 275283.
Gerhart, J.C., Wu, M. and Kirschner, M.W. (1984) Cell cycle dynamics of an M-phase-specific cytoplasmic factor in Xenopus laevis oocytes and eggs. J. Cell Biol., 98, 12471255.[Abstract]
Glotzer, M., Murray, A.W. and Kirschner, M.W. (1991) Cyclin is degraded by the ubiquitin pathway. Nature (London), 349, 132138.[ISI][Medline]
Gougeon, A. and Testart, J. (1986) Germinal vesicle breakdown in oocytes of human atretic follicles during the menstrual cycle. J. Reprod. Fertil., 78, 389401.[Abstract]
Haider, S. and Chaube, S.K. (1996) The in vitro effects of forskolin, IBMX and cyanoketone on meiotic maturation in follicle-enclosed catfish (Clarias batrachus) oocytes. Endocrinology, 115, 117123.
Heukeshaven, J. and Dernick, R. (1988) Improved silver staining procedure for fast staining in PhastSystem Development Unit I staining of sodium dodecyl sulfate gels. Electrophoresis, 9, 2832[ISI][Medline]
Hunt, T. (1989) Under arrest in the cell cycle. Nature, 342, 483484.[ISI][Medline]
Jeffrey, P.D., Russo, A.A., Polyak, K. et al. (1995). Mechanism of CDK activation revealed by the structure of a cyclin A-CDK2 complex. Nature, 376, 313320.[ISI][Medline]
Jessus, C., Rime, H., Haccard, O. et al. (1991) Tyrosine phosphorylation of p34cdc2 and p42 during meiotic maturation of Xenopus oocyte. Development, 111, 813820.[Abstract]
Kastrop, P.M.M., Hulshof, S.C.J., Bevers, M.M. et al. (1991) The effects of -amatin and cycloheximide on nuclear progression, protein synthesis, and phosphorylation during bovine oocyte maturation in vitro. Mol. Reprod. Dev., 28, 249254.[ISI][Medline]
Kubiak, J.Z., Weber, M., de Pennart, H. et al. (1993) The metaphase II arrest in mouse oocytes is controlled through microtubule-dependent destruction of cyclin B in the presence of CSF. EMBO J., 12, 37733778.[Abstract]
Laemmli, U.K. (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature, 227, 680685.[ISI][Medline]
Lefevre, B., Gougeon, A., Nome, F. et al. (1989) In vivo changes in oocytes germinal vesicle related to follicular quality and size at mid-follicular phase during stimulated cycles in the Cynomolgus monkey. Reprod. Nutr. Dev., 29, 523531.[ISI][Medline]
Longeran, P., Khatir, H., Carolan, C. et al. (1997) Bovine blastocyst production in vitro after inhibition of oocyte meiotic resumption for 24 h. J. Reprod. Fertil., 109, 355365.[Abstract]
Maller, J.L. (1994) Biochemistry of cell cycle check points at the G2/M and metaphase/anaphase transitions. Semin. Dev. Biol., 5, 183190.
Masui, Y. and Market, C.L. (1971) Cytoplasmic control of nuclear behaviour during meiotic maturation of frog oocytes. J. Exp. Zool., 177, 129145.[ISI][Medline]
Mattioli, M., Galeati, M.L., Bacci, M.L. et al. (1991) Changes in maturation-promoting activity in the cytoplasm of pig oocyte throughout maturation. Mol. Reprod. Dev., 30, 119125.[ISI][Medline]
Moor, R.M. and Trounson, A.O. (1977) Hormonal and follicular factors and their subsequent developmental capacity. J. Reprod. Fertil., 49, 101109.[Abstract]
Moor, R.M., Osborn, J.C. and Crosby, I.M. (1985) Gonadotrophin induced abnormalities in sheep oocytes after superovulation. J. Reprod. Fertil., 74, 167172.[Abstract]
Moreno, S. and Nurse, P. (1990) Substrates for p34cdc2: in vivo veritas? Cell, 61, 549551.[ISI][Medline]
Motlík, J. and Kubelka, M. (1990) Cell cycle aspects of growth and maturation of mammalian oocytes. Mol. Reprod. Dev., 27, 366375.[ISI][Medline]
Motlík, J. and Rimkevicová, Z. (1990) Combined effects of protein synthesis and phosphorylation inhibitors on maturation of mouse oocytes in vitro. Mol. Reprod. Dev., 27, 230234.[ISI][Medline]
Naito, K. and Toyoda, Y. (1991) Fluctuation of histone H1 kinase activity during meiotic maturation in porcine oocytes. J. Reprod. Fertil., 93, 467473.[Abstract]
Naito, K., Dean, F.P. and Toyoda, Y. (1992) Comparison of histone H1 kinase activity during meiotic maturation between two types of porcine oocytes matured in different media in vitro. Biol. Reprod., 47, 4347.[Abstract]
Ng, S.C., Bongso, A. and Ratnam, S.S. (1991) Microinjection of human oocytes: a technique for severe oligoasthenoteratozoospermia. Fertil. Steril., 56, 11171123.[ISI][Medline]
Pincus, G. and Enzmann, E.V. (1935) The comparative behaviour of mammalian eggs in vivo and in vitro. J. Exp. Med., 62, 665675.
Quinn, P., Barros, C. and Wittingham, D.G. (1982) Preservation of hamster oocytes to assay fertilisation capacity of human spermatozoa. J. Reprod. Fertil., 66, 161168.[Abstract]
Rime, H., Neant, I., Guerrier, P. et al. (1989) 6-Dimethylaminopurine (6-DMAP), a reversible inhibitor of the transition to metaphase during the first meiotic division of the mouse oocyte. Dev. Biol., 133, 169179.[ISI][Medline]
Rossant, J. and Pederson, R.A. (eds) (1986) Experimental Approaches to Mammalian Embryonic Development. Cambridge University Press, New York, pp 195.
Russell, J.B., Knezevich, K.M., Fabian, K.F. et al. (1997) Unstimulated immature oocyte retrieval: early versus midfollicular endometrial priming. Fertil. Steril., 67, 616620.[ISI][Medline]
Segaloff, D.L. and Ascoli, M. (1993) The lutropin/choriogonadotropin receptor ...4 years later. Endocr Rev., 14, 324347.[Abstract]
Shultz, R.M. and Wassarman, P.M. (1977) Specific changes in the pattern of protein synthesis during meiotic maturation of mammalian oocytes in vitro. Proc. Natl. Acad. Sci. USA, 74, 538541.[Abstract]
Sirard, M.A. and First, N.L. (1988) In vitro inhibition of oocyte nuclear maturation in the bovine. Biol. Reprod., 39, 229234.[Abstract]
Süss, U., Wüthrich, K. and Stranzinger, G. (1988) Chromosome configurations and the time sequence of the first meiotic division in bovine oocytes matured in vitro. Biol. Reprod., 38, 871880.[Abstract]
Szöllösi, M.S., Debey, D., Szöllösi. D. et al. (1991a) Effects of puromycin and 6-DMAP on mouse oocyte maturation. Bull. Assoc. Anat. (Nancy), 75, 99103.[Medline]
Szöllösi, M.S., Debey, D., Szöllösi. D. et al. (1991b) Chromatin behaviour under influence of puromycin and 6-DMAP at different stages of mouse oocyte maturation. Chromosoma, 100, 339354.[ISI][Medline]
Szöllösi, M.S., Kubiak, J.Z., Debey, P. et al. (1993) Inhibition of protein kinases by 6-dimethylaminopurine accelerates the transition to interphase in activated mouse oocytes. J. Cell Sci., 104, 861872.
Trounson, A., Wood, C, and Kausche, A. (1994) In vitro maturation and the fertilisation and developmental competence of oocytes recovered from untreated polycystic ovarian patients. Fertil. Steril., 62, 353362.[ISI][Medline]
Trounson, A.O., Bongso, A., Szell, A. et al. (1996) Maturation of human and bovine primary oocytes in vitro for fertilisation and embryo production. Sing. J. Obstet. Gynaecol., 27, 7884.
Trounson, A., Anderiesz, C., Jones, G.M. et al. (1998) Oocyte maturation. Hum. Reprod., 13 (Suppl. 3), 5262.[Medline]
Wickramasinghe, D. and Albertini, D.F. (1993) Cell cycle control during mammalian oogenesis. Curr. Top. Dev. Biol., 28, 125153.[ISI][Medline]
Wynn, P., Picton, H.M., Krapez, J.A. et al. (1998) Pretreatment with follicle stimulating hormone promotes the numbers of human oocytes reaching metaphase II by in vitro maturation. Hum Reprod., 13 31333138.
Submitted on April 30, 1999; accepted on October 22, 1999.