Causes of developmental failure of in-vitro matured rhesus monkey oocytes: impairments in embryonic genome activation

R.Dee Schramm1,3, Ann Marie Paprocki1 and Catherine A. VandeVoort2

1 Wisconsin National Primate Research Center, University of Wisconsin, Madison, WI 53715 and 2 California National Primate Research Center, Davis, CA, USA

3 To whom correspondence should be addressed at: Wisconsin National Primate Research Center, 1223 Capitol Court, Madison, WI 53715, USA. e-mail: schramm{at}primate.wisc.edu


    Abstract
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
BACKGROUND: Understanding the causes of developmental failure of in-vitro matured primate oocytes may lead to viable strategies for improving their developmental competence. The aims of this study were to determine whether the timely onset of embryonic genome activation among individual blastomeres of preimplantation macaque embryos is impaired by in-vitro maturation (IVM) of oocytes and whether these impairments are associated with developmental failure during the embryonically controlled period of preimplantation development. METHODS: Genome activation among individual blastomeres was assessed using expression of fibrillarin as a marker of nucleolar transcription. Immature oocytes were obtained from rhesus monkeys following treatment with recombinant human FSH and matured in-vitro in one of two IVM media (CMRLa or CMRLb). In-vivo matured oocytes were obtained from FSH treated monkeys following administration of hCG. Oocytes were fertilized in vitro and either cultured for developmental studies or processed at the 8–12-cell stage for expression of fibrillarin. RESULTS: Developmental competence of embryos derived from in-vitro matured CMRLa oocytes was markedly (P < 0.05) impaired compared with those derived from in-vivo matured or in-vitro matured CMRLb oocytes. Developmental profiles were similar among the groups prior to the 8-cell stage. However, in embryos derived from in-vitro matured CMRLa oocytes, developmental failure increased significantly (P < 0.05) after the time of genome activation compared with those derived from in-vivo matured or in-vitro matured CMRLb oocytes. The mean percentages of non-activated blastomeres per embryo, as well as the proportions of embryos with at least one non-activated blastomere, or with no activated blastomeres, were all significantly (P < 0.05) greater in embryos derived from in-vitro matured CMRLa oocytes than in those derived from in-vivo matured or CMRLb oocytes. CONCLUSIONS: The relatively poor developmental competence of in-vitro matured primate oocytes is likely caused in part by failure in the timely onset of embryonic genome activation resulting from incomplete cytoplasmic maturation.

Key words: fibrillarin/genome activation/in-vitro maturation/macaque/oocyte


    Introduction
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Development of the fertilized ovum through cleavage, blastocyst formation, implantation and term development is dependent upon the developmental programme contained within the oocyte itself. The acquisition of developmental competence by oocytes, which is commonly referred to as ‘cytoplasmic maturation’, is poorly understood in any species. To date, our inadequate understanding of the biochemical and molecular processes involved in cytoplasmic maturation of primate oocytes has hindered development of successful techniques for in-vitro maturation (IVM) of human and non-human primate oocytes. Information on the nature and timing of essential molecular and biochemical events during oocyte maturation and how they are subsequently involved in regulation of embryonic development is limited.

Mammalian preimplantation embryogenesis is initially dependent upon maternally-inherited molecules during the early cleavage stages (Bacharova and De Leon, 1980Go; McLaren, 1981Go; Telford et al., 1990Go). As development proceeds, and maternally-inherited molecules diminish, the process of embryogenesis becomes dependent upon the expression of genetic information derived from the embryonic genome (Telford et al., 1990Go; Tesarik, 1990Go). Developmental failure may thus result from insufficient accumulation of developmentally important maternally-derived molecules during development or maturation of oocytes, subsequently leading to cleavage arrest during the maternally-controlled period of development, impairments in the transition from maternal to embryonic control of development, or abnormal pre- or post-implantation gene expression. It has been proposed that failure of timely onset of embryonic transcription may be a common cause of developmental failure of in-vitro produced human embryos (Tesarik, 1987Go, 1989a,b; Tesarik et al., 1986bGo; Braude et al., 1988Go). Because genome activation is thought to be under the control of maternal genome products (Tesarik, 1987Go, 1994; Wang and Latham, 1997Go), it has been suggested that impairments in genome activation may likely result from incomplete or inadequate cytoplasmic maturation of oocytes (Tesarik, 1987Go, 1994; Winston et al., 1991Go; Hardy et al., 1993Go). Understanding the causes of developmental failure of in-vitro matured primate oocytes may lead to viable strategies for improving their cytoplasmic maturation and subsequent developmental competence.

In macaque, as well as human embryos, genome activation occurs at the 6- to 8-cell stage (Tesarik, 1987Go; Tesarik et al., 1986aGo,b, 1988; Artley et al., 1992Go; Weston and Wolf, 1994Go; Schramm and Bavister, 1999bGo), coincident with the onset of nucleolar rRNA synthesis (Tesarik et al., 1986aGo,b, 1987; Schramm and Bavister, 1999bGo), and expression of fibrillarin (Schramm and Bavister, 1999bGo), a nucleolar protein involved in methylation and processing of pre-rRNA (Kass et al., 1990Go; Calzergues-Ferrer et al., 1991Go; Kiss-Laszlo et al., 1996Go). We hypothesize that incomplete cytoplasmic maturation of oocytes during IVM leads to impairments in genome activation resulting in developmental failure during the embryonically-controlled period of preimplantation development. Specific aims of the this study were to determine whether the timely onset of embryonic genome activation among individual blastomeres of preimplantation embryos is impaired by IVM of oocytes and whether these impairments are associated with stage specific developmental failure in macaque embryos.


    Materials and methods
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Ovarian stimulation/oocyte recovery/oocyte maturation
The general care and housing of rhesus monkeys (Macaca mulatta) at the Wisconsin (WNPRC) and California (CNPRC) National Primate Research Centers have been described previously (Goy and Robinson, 1982Go; Goy and Kemnitz, 1983Go; VandeVoort and Tarantal, 2001Go). Both Centers are fully accredited by the Association for Assessment and Accreditation of Laboratory Animal Care (AAALAC), and animal protocols and experiments were approved by the respective Graduate School Animal Care and Use Committees. The animals were maintained according to recommendations of the Guide for the Care and Use of Laboratory Animals and the Animal Welfare Act with its subsequent amendments.

For collection of in-vivo matured oocytes, rhesus monkeys (Macaca mulatta; n = 13) received twice daily i.m. injections of 30 IU recombinant human FSH (rhFSH; Organon Inc., NJ, USA) for 7 days, beginning on days 1 to 3 of the menstrual cycle (day 1 = first day of menstruation). Recombinant hCG (1000 IU; Ares Advanced Technology, NJ, USA) was injected (i.m.) on treatment day 8 for induction of oocyte maturation. Oocytes were aspirated laparoscopically into Tyrode’s lactate (TL)-HEPES medium (37°C) containing 0.1 mg/ml polyvinyl alcohol (PVA) and 10 IU/ml heparin 27–32 h following injection of hCG. Oocytes were retrieved from aspirates using an EM Con filter (Veterinary Concepts, Spring Valley, WI, USA). Cumulus masses were treated with 0.1% hyaluronidase to facilitate recovery of oocytes. Oocytes were cultured in modified (Boatman, 1987Go) CMRL-1066 medium; (Connaught Medical Research Laboratories Medium-1066; Invitrogen, Carlsbad, CA, USA) containing 20% bovine calf serum (Hyclone, Logan, UT, USA) at 37°C in a humidified atmosphere of 5% CO2 in air for 4–8 h prior to insemination.

Immature germinal vesicle (GV) oocytes for IVM, were obtained from both the WNPRC and the CNPRC. Monkeys (n = 28) received rhFSH as described above, but did not receive hCG prior to follicular aspiration (Schramm and Bavister, 1994Go). Oocytes were retrieved from monkeys either laparoscopically (n = 9; WNPRC) or using ultrasonography (n = 19; CNPRC) on the morning following the last day of rhFSH treatment. Immature oocytes (enclosed by at least three layers of cumulus cells) from each monkey were cultured in one of two types of medium, but not in both. Thus, oocytes cultured in each of the two IVM treatments were obtained during different cycles, as were the in-vivo matured oocytes. All oocytes from the WNPRC were cultured in modified CMRL-1066 medium with or without human gonadotrophins (5 µg/ml hFSH and 10 µg/ml hLH) containing 20% bovine calf serum (CMRLa), as described previously (Morgan et al., 1991Go; Schramm et al., 1993Go, 1994; Schramm and Bavister, 1994Go, 1995, 1996a). At the CNPRC, oocytes from 16 monkeys were cultured in modified CMRL-1066 medium containing hFSH and hLH (0.03 IU/ml Pergonal, Ares-Serono), 10 µg/ml androstenedione (Steraloids, Wilton, NH, USA) and 10% bovine calf serum (CMRLb). Oocytes obtained from three monkeys at the CNPRC were cultured in CMRLa medium with gonadotrophins, as described above. All oocytes were cultured in microdrops under mineral oil at 37°C in a humidified atmosphere of 5% CO2 in air for 28–30 h prior to insemination.

IVF/embryo culture
Sperm was collected from adult males by electroejaculation as described previously (Bavister et al., 1983Go; Sarason et al., 1991Go). Sperm capacitation and IVF were done as described previously (Bavister et al., 1983Go) with a few minor modifications. Briefly, 10 x 106 washed sperm/ml were resuspended in 2 ml TALP medium and incubated at 37°C in 5 % CO2 in air for 1–10 h. Sperm were treated with dbcAMP (1mmol/l) and caffeine (1mmol/l) to induce hyperactivation either during (WNPRC) or for 30 min before (CNPRC) co-incubation with oocytes. Sperm (300 000/ml) were co-incubated with oocytes for 12–16 h at 37°C in a humidified atmosphere of 5% CO2 in air. Sperm and remaining cumulus cells were then removed manually by pipetting through a finely pulled glass pipette, and oocytes were examined for evidence of fertilization. Embryos used for developmental studies were cultured in HECM-9 medium (McKiernan and Bavister, 2000Go) for 48 h, then switched into HECM-9 containing 5% bovine calf serum. Embryos were cultured in 5% CO2, 5% O2 and 90% N2 at 37°C in microdrops under mineral oil and placed into fresh media every other day until zona escape or developmental arrest. Embryos used for genome activation studies were cultured similarly and processed for immunocytochemistry at the 8- to 12-cell stage. Embryos obtained from the CNPRC to be used for genome activation studies were inseminated and cultured to the pronucleate stage, and then loaded into cryovials in equilibrated HECM-9 medium and shipped overnight to the WNPRC in a portable incubator (Minitube, Inc., Madison, WI, USA) at 37°C. Upon arriving at the WNPRC, embryos that had progressed to the 4-cell stage were cultured an additional 24 h (8- to 12-cell stage) before processing for immunocytochemistry. The authors have used this method of shipping embryos extensively and have never observed impairments in subsequent development (R.D.Schramm, unpublished data).

Assessment of nucleolar transcriptional activation in individual blastomeres
Immunocytochemistry for expression of fibrillarin was done as described previously for macaque embryos (Schramm and Bavister, 1999bGo). Embryos were attached to poly-L-lysine-coated slides, fixed in methanol-free formaldehyde for 1 h and then permeabilized at 37°C in 0.1 mol/l phosphate buffered saline containing 1.0% Triton X-100 (PBS-triton). Following a glycine rinse for reducing free aldehydes, embryos were blocked in PBS-triton containing 3 mg/ml non-fat dry milk (NFDM) and then incubated for 1 h at 37°C with the anti-fibrillarin antibody purchased as the nucleolar pattern component of the Anti-Nuclear Antibody kit (ANA-N; Sigma). After rinsing in PBS-triton-NFDM, embryos were incubated as above in fluorescein isothiocyanate (FITC)-conjugated goat anti-human immunoglobulin diluted in PBS with Evans blue (Sigma). After a rinse in PBS-triton, containing 20 µg/ml Hoechst 33342 (Sigma), specimens were mounted in Vectashield mountant (Vector Laboratories), and stored at 4°C in the dark. Negative controls for background fluorescence were treated as described above, but the primary antibody (ANA-N) was omitted. Whole mounts were examined at 100–200x using a Nikon Eclipse TE 300 microscope. Ultra violet and FITC signals were detected using appropriate filter combinations, and photographed with a Nikon 2000 camera mounted on the microscope.

Statistical analyses
Percentages for developmental data, mean regression lines (slopes) for various developmental periods, and numbers of activated cells per embryo were compared using general linear models. Percentage data was arcsine transformed to increase linearity (Sokal and Rohlf, 1995Go). Post-hoc treatment differences were determined using Fisher’s least significant difference (LSD) test. Percentages of embryos having no activated cells and >=1 non-activated cell were compared using protected Pearson {chi}2 analyses.


    Results
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
The developmental profiles of in-vitro matured (CMRLa or CMRLb medium) and in-vivo matured oocytes are shown in Figure 1. No differences were detected at any developmental stage among in-vitro matured oocytes cultured with or without gonadotrophins in CMRLa culture medium (not shown), so data were pooled for analyses. The developmental competence (% blastocysts) of embryos derived from in-vitro matured CMRLa oocytes (15.3%) was markedly (P < 0.05) impaired compared with that of embryos derived from in-vivo matured oocytes (60.8%) or in-vitro matured CMRLb oocytes (50.9%). The incidence of developmental failure of embryos derived from in-vitro matured CMRLa oocytes, compared with in-vivo matured or in-vitro matured CMRLb oocytes, was significantly (P < 0.05) greater after the time of genome activation, but was similar (P > 0.05) among treatment groups prior to the 8-cell stage. Analyses of the developmental regression lines indicate that the rates of developmental failure through both the 9- to 16-cell stage (mean slope = –9.9) and between the morula and blastocyst stages (mean slope = –7.2) were similar (P > 0.05, non-significant) among treatment groups. However, the rate of developmental failure increased (P < 0.01) dramatically between the 9- to 16-cell and morula stages in embryos derived from in-vitro matured CMRLa oocytes (slope = –46.5), compared with in-vivo matured or in-vitro matured CMRLb oocytes (slopes = –7.3 and –7.9 respectively).



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Figure 1. Developmental profiles of embryos derived from in-vivo matured (n = 52 zygotes) or in-vitro matured oocytes cultured during maturation in CMRLa (n = 48 zygotes) or CMRLb (n = 69 zygotes) medium. Values are percentages of zygotes progressing to or beyond each developmental stage. *Significantly different (P <= 0.05) from the other two treatment groups; §slope of regression line (9–16 cell to morula stages) differed significantly (P <= 0.01) from other two treatment groups. Slopes were –7.3, –7.9 and –46.5 for in-vivo, in-vitro matured CMRLa and in-vitro matured CMRLb oocytes respectively.

 
Figure 2 illustrates a representative 8-cell stage embryo derived from an in-vivo matured oocyte after dual labelling with Hoechst 33342 (a) and with an anti-fibrillarin antibody (ANA-N) (b). Fibrillarin was typically detected in all blastomeres of embryos derived from in-vivo matured oocytes. Figure 3 illustrates representative embryos (8- to 9-cell stage) derived from in-vitro matured oocytes matured in CMRLa medium after dual labelling with Hoechst 33342 (a,c,e) and with ANA-N (b,d,f). Fibrillarin was typically either not detected in any blastomeres (a,b) or limited to only some blastomeres (c–f). Unlike in embryos derived from oocytes matured in CMRLa medium, in those derived from oocytes matured in CMRLb medium (Figure 4), fibrillarin was typically expressed in all blastomeres (a,b), similar to embryos derived from in-vivo matured oocytes.



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Figure 2. Representative 8-cell stage embryo derived from an in-vivo matured oocyte after dual labelling with Hoechst 33342 (a) and with an anti-fibrillarin antibody (ANA-N) (b). Fibrillarin was typically detected in all blastomeres of embryos derived from in-vivo matured oocytes.

 


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Figure 3. Representative embryos (8- to 9-cell stage) derived from in-vitro matured oocytes cultured during maturation in CMRLa medium after dual labelling with Hoechst 33342 (a,c,e) and with an anti-fibrillarin antibody (ANA-N) (b,d,f). Expression of fibrillarin was typically either not detected in any blastomeres (a,b) or limited to only some blastomeres (cf).

 


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Figure 4. Representative 8-cell stage embryo derived from an in-vitro matured oocyte cultured during maturation in CMRLb medium after dual labelling with Hoechst 33342 (a) and with an anti-fibrillarin antibody (ANA-N) (b). Fibrillarin was typically expressed in all blastomeres, similar to that of embryos derived from in-vivo matured oocytes.

 
The incidences of genome activation failure in embryos derived from in-vitro matured (CMRLa or CMRLb medium) and in-vivo matured oocytes are shown in Figure 5. Genome activation was assessed based upon expression of fibrillarin in individual blastomeres of 8- to 12-cell stage embryos. The mean proportion of activated blastomeres per embryo was significantly (P < 0.001) reduced in embryos derived from in-vitro matured CMRLa oocytes (44.4%) compared with those derived from in-vivo matured oocytes (91.9%) or in-vitro matured CMRLb oocytes (74.8%), with no differences detected between the latter two. Proportions of embryos with at least one non-activated blastomere were significantly (P < 0.05) greater for embryos derived from in-vitro matured CMRLa oocytes (74.1%) compared with those from in-vivo matured (25.9%) or in-vitro matured CMRLb oocytes (46.4%), with no differences detected between the latter two. The proportion of embryos that had none of their blastomeres stained, indicative of complete genome activation failure, was significantly (P < 0.01) greater for embryos derived from in-vitro matured CMRLa oocytes (44.4%) than for embryos derived from in-vivo matured oocytes (3.7%) or in-vitro matured CMRLb oocytes (10.7%), with no differences detected between the latter two.



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Figure 5. Incidences of genome activation failure, as assessed by expression of fibrillarin, in individual blastomeres in 8- to 12-cell stage embryos derived from in-vivo matured (n = 27 embryos) or in-vitro matured (IVM) oocytes cultured during maturation in CMRLa (n = 27 embryos) or CMRLb (n = 28 embryos) medium. *denotes significant (P <= 0.05) differences from other treatments.

 

    Discussion
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Although a limited number of in-vitro matured/IVF oocytes have resulted in the birth of normal offspring following transfer in women (Barnes et al., 1995Go; Russell et al., 1997Go) and monkeys (Schramm and Paprocki, 2000aGo), IVM is far from being a successful procedure for the production of developmentally competent primate oocytes. The developmental competence of primate oocytes matured in vitro is markedly inferior to that of their in-vivo matured counterparts (Bavister et al., 1983Go; Boatman, 1987Go; Wolf et al., 1989Go; Lanzendorf et al., 1990Go; Cha et al., 1991Go, 1992; Morgan et al., 1991Go; Zhang et al., 1993Go; Barnes et al., 1995Go; Schramm and Bavister 1996aGo,b, 1999a). Although some progress has been made in developing culture conditions that support cytoplasmic maturation of oocytes from rodents (Eppig and Schroeder, 1989Go; Hirao et al., 1990Go; Vanderhyden and Armstrong, 1990Go; Eppig et al., 1992Go) and domestic species (Liebfried-Rutledge et al., 1987Go; Mattioli et al., 1988Go; Frei et al., 1989Go; Mochizuki et al., 1991Go; Funahashi et al., 1994Go; Hirao et al., 1994Go; Kobayahi et al., 1994Go), attempts to improve the developmental competence of primate (human or non-human) oocytes matured in vitro have met with little success (Schramm and Bavister, 1995Go, 1996a, 1999a; Trounson et al., 1998Go). To date, our inadequate understanding of the biochemical and molecular processes involved in cytoplasmic maturation of primate oocytes has hindered development of successful techniques for IVM of primate oocytes. Previous studies using both non-stimulated and gonadotrophin-primed monkeys indicate that developmental competence is acquired relatively late during the follicular growth process, and that cytoplasmic maturation is acquired both during oocyte development, prior to meiotic resumption (Schramm and Bavister, 1994Go, 1999a), as well as following the ovulatory surge of LH (or injection of hCG; Schramm and Bavister, 1995Go, 1996a,b, 1999a). Information on the specific molecular processes involved in cytoplasmic maturation of oocytes and how they are subsequently involved in regulating the maternally and embryonically controlled periods of development are largely unknown.

In this study, relatively little developmental failure occurred prior to the time of genome activation in in-vivo or in-vitro matured oocytes, similar to that reported previously for in-vivo matured rhesus oocytes (Schramm and Bavister, 1996bGo, 1999a). However, earlier studies have shown that significant developmental failure occurred prior to the 8-cell stage in in-vitro matured oocytes obtained from non-stimulated, as opposed to FSH-primed, monkeys (Schramm and Bavister, 1994Go, 1995, 1996a, 1999a). Thus, the ability to develop through the maternally controlled period of development may likely be acquired during the later stages of follicular development, even in fully grown oocytes.

The transition from maternal to embryonic control of development occurs at species specific stages and has been commonly referred to as ‘genome activation’. It is coincident with this critical stage that preimplantation embryos experience species–specific blocks to development in the presence of {alpha}-amanitin and under various culture conditions in vitro. Since genome activation is thought to be under the control of maternal genome products (Tesarik, 1987Go, 1994; Wang and Latham, 1997Go), it has been suggested that impairments in genome activation may result from incomplete or inadequate cytoplasmic maturation of oocytes (Tesarik, 1987Go, 1994; Winston et al., 1991Go; Hardy et al., 1993Go). In the present study, we have shown that developmental failure of embryos derived from in vitro matured oocytes (cultured in CMRLa medium) occurred predominantly during the embryonically controlled period of development. Analyses of the developmental regression lines, or slopes, indicate that the rate of developmental failure was similar among treatments through the maternally controlled period of development, but increased dramatically beginning shortly after the time of genome activation in embryos derived from oocytes matured in vitro in CMRLa medium. A similar pattern of developmental failure was recently reported for in-vivo matured oocytes obtained from prenatally androgenized rhesus monkeys, in conjunction with abnormal serum and follicular fluid hormone concentrations both before and after hCG (Dumesic et al., 2002Go). Taken together, these data suggest that cytoplasmic impairments in oocytes, incurred during oocyte development or maturation, may subsequently lead to impairments in the transition from maternal to embryonic control of development. Interestingly, although the percentages of oocytes developing into morulae and blastocysts were markedly reduced when matured in CMRLa medium, the developmental slope between the morula and blastocyst stage was similar to that of the other two treatment groups. This implies that IVM derived embryos that succeed in completing the transition from maternal to embryonic control of development are not further impaired in their ability to complete the morula to blastocyst stage transition. This does not however, imply that resulting blastocysts are of equal quality.

The increased incidence of developmental failure after the time of genome activation was not evident in IVM derived oocytes cultured in CMRLb medium, indicating that this medium may provide factors, such as androstenedione, necessary for development through the embryonically driven period of development. Differential effects of in-vitro culture and culture media on gene expression in preimplantation embryos have previously been demonstrated in other species and have been related to developmental abnormalities (Reik et al., 1993Go; Ho et al., 1994Go, 1995; Behboodi et al., 1995Go; Doherty et al., 2000Go). In order to determine whether developmental failure during the embryonically driven period of development was caused by impairments in embryonic genome activation, we used the expression of fibrillarin in individual blastomeres as a marker for genome activation, as described previously (Pinto-Correia et al., 1995Go). In previous studies in rhesus monkeys, the onset of fibrillarin expression mirrored that of tritiated uridine incorporation into the nucleolus, first appearing in 6- to 8-cell stage embryos (Schramm and Bavister, 1999bGo). In similar studies in mice (Baran et al., 1995Go; Cuadros-Fernandez and Esponda, 1996Go), rabbits (Pinto-Correia et al., 1995Go; Baran et al., 1997Go) and cows (Schramm and Paprocki, 2000bGo), nucleolar expression of fibrillarin was first detected at the species-specific time of genome activation, coincident with that of nucleolin, protein B23, RNA polymerase I and the onset of nucleolar transcription (Baran et al., 1995Go, 1996; Cuadros-Fernandez and Esponda, 1996Go). Results of this study demonstrate that the mean percentages of non-activated blastomeres per embryo, as well as the proportions of embryos with at least one blastomere that had failed to undergo genome activation or with no blastomeres that had undergone genome activation were all significantly greater in embryos derived from oocytes matured in CMRLa than in those matured in CMRLb medium or embryos derived from in-vivo matured oocytes. In fact, 12/27 IVM-derived embryos matured in CMRLa had none of their blastomeres stained, indicative of complete genome activation failure. There were no significant differences in any of these endpoints between embryos derived from in-vivo matured and in-vitro matured CMRLb oocytes. It should be noted that while expression of fibrillarin requires genome activation, it does not neccessarily indicate that activation was complete. Likewise, the absence of fibrillarin expression does not necessarily imply that no aspect of genome activation has occurred. It is also possible that genome activation may have been delayed, rather than failing completely in blastomeres failing to express fibrillarin. Nevertheless, these findings are compatible with the developmental findings above, suggesting that impairments in genome activation may be a prevalent cause of developmental failure in IVM derived primate embryos.

Similar studies have not been done on IVM oocytes in other species. However, autoradiographic studies of tritiated uridine incorporation into IVF embryos obtained from in-vivo matured human oocytes have shown that transcription failure is not uncommon in blastomeres of 8-cell and morula stage embryos (Camous et al., 1986Go). These blastomeres express very low levels of extranucleolar RNA synthesis and a complete absence of nucleolar RNA synthesis, with up to 30% of blastomeres exhibiting this impairment. The absence of rRNA synthesis in these blastomeres is of particular importance since the embryo must support its demand on protein synthesis using maternally inherited ribosomes, which are rapidly exhausted during the first three cleavage divisions (Tesarik et al., 1986aGo). Some of these embryos in which the switch from maternal to embryonic gene activity has failed in a large proportion of blastomeres can progress to the morula stage (Tesarik, 1987Go, 1989a,b; Tesarik et al., 1987Go) but fail to develop into blastocysts (Tesarik, 1989a,Go 1994). Similar studies in bovine embryos (Pavlok et al., 1993Go) have shown that unlike in 8-cell embryos derived from oocytes from large antral follicles, those derived from oocytes from small (1–2mm) antral follicles exhibited very low levels of extranucleolar RNA synthesis and the absence of nucleolar RNA synthesis, indicative of a delayed onset of genome activation. This was observed not only among embryos, but also among blastomeres within the same embryo (Pavlok et al., 1993Go), and was associated with a high incidence of developmental failure. Taken together, these developmental and molecular findings indicate that the relatively poor developmental competence typical of in-vitro matured human (Cha et al., 1991Go, 1992; Barnes et al., 1995Go, 1996; Trounson et al., 1998Go) and non-human primate (Morgan et al., 1991Go; Schramm and Bavister, 1994Go, 1995, 1996a, 1999a) oocytes is likely caused, in part, by impairments in the timely onset of embryonic transcription, resulting in developmental failure.

Although the precise mechanisms and components necessary for initiation of embryonic transcription are not known, genome activation may be under the control of maternally inherited factors, such as Oct 4 (Rosner et al., 1990Go; Abdel-Rahman et al., 1995Go), eukaryotic transcription initiation factor (Scholer et al., 1991Go; Rosner et al., 1990Go; De Sousa et al., 1998Go), or the maternal gene factor MATER (Dean, 2002Go). Impairments in the transition from maternal to embryonic control of development in IVM derived embryos may have their origins in aberrant expression of genes for these or other maternal factors resulting from incomplete cytoplasmic maturation. Although genome activation failure may contribute to developmental failure in IVM derived embryos, other molecular impairments may also contribute to developmental failure. Some embryos may be impaired in their ability to transcribe some, but not all, embryonically encoded genes (Artley et al., 1992Go). In addition, some maternally inherited messages may be involved in the control of cellular events in relatively late stages of human preimplantation development, after genome activation has occurred (Tesarik, 1989aGo), and insufficient accumulation of these messages may lead to impairments in the morula to blastocyst stage transition (Renard et al., 1994Go; Moor et al., 1998Go). The effects of in-vitro culture of oocytes on expression of maternally derived messages and subsequent activation of specific embryonically encoded genes has not been examined in any species.

In conclusion, we have demonstrated that the relatively poor developmental competence of in-vitro matured oocytes is likely caused, in part, by failure in the timely onset of embryonic transcription, resulting from incomplete cytoplasmic maturation during IVM. Identification of specific maternal transcripts and cytoplasmic components acquired during oocyte development and maturation that are essential for normal pre- and post-implantation embryogenesis, will vastly improve our understanding of oocyte cytoplasmic maturation on a molecular level, and how cytoplasmic changes incurred during oocyte development and maturation are subsequently linked to the regulation of embryonic gene expression and preimplantation embryogenesis in primates. Such information will be of tremendous value in formulation of strategies for production of developmentally competent human oocytes by IVM techniques.


    Acknowledgements
 
The authors thank Steve Eisele, Michele Shotzko, Jennifer Lambert-Newman and Eric Peterson of Reproductive Services for menstrual cycle monitoring, hormone injections, oocyte retrievals and semen collection. We also thank Denny Mohr for surgical assistance with oocyte retrievals and gratefully acknowledge Dr David Abbott for assistance with statistical analyses. We are grateful to Organon Inc., West Orange, NJ, USA for the generous supply of recombinant human FSH and Serono laboratories (Ares Advanced Technology), Randolph, MA, USA for the gift of recombinant hCG. This research was supported by research grants NIH RR00167, RR14093, RR00169 and RR13439. This is publication number #42-018 of the WNPRC.


    References
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
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Submitted on August 30, 2002; resubmitted on November 21, 2002; accepted on December 4, 2002.