Human Reproduction, Vol. 14, No. 12, 3060-3068,
December 1999
© 1999 European Society of Human Reproduction and Embryology
Epidermal growth factor enhances preimplantation developmental competence of maturing mouse oocytes
The Jackson Laboratory, Bar Harbor, ME 04609, USA
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Abstract |
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The objective of this study was to determine whether epidermal growth factor (EGF) promotes nuclear and cytoplasmic maturation of mouse oocytes grown in vivo or in vitro. In-vivo-grown oocytes were isolated at the germinal vesicle (GV) stage from gonadotrophin-primed (PR) or -unprimed (UPR) 22-day-old mice before in-vitro maturation (IVM). In-vitro-grown (IVG) oocytes were isolated from preantral follicles of 12-day-old mice and grown in vitro without gonadotrophins for 10 days before maturation (IVG/IVM oocytes). IVM and IVG/IVM oocytes were matured in medium supplemented with either EGF (10 ng/ml), follicle stimulating hormone (FSH) (100 ng/ml), EGF plus FSH, or with neither ligand (control). When oocyte–cumulus cell complexes were isolated from PR and UPR mice, IVM with EGF (10 ng/ml), alone or in combination with FSH (100 ng/ml), increased (P < 0.05) the incidence of nuclear maturation to metaphase II. Cytoplasmic maturation of oocytes from PR females, manifested as increased frequency of cleavage to the 2-cell stage and development to the blastocyst stage, was also enhanced with EGF (P < 0.05). Moreover, EGF increased the number of cells per blastocyst, but only in the absence of FSH (P < 0.01). In contrast, EGF, FSH, or EGF plus FSH did not affect the percentage of oocytes from UPR mice completing preimplantation development, but did increase the number of cells per blastocyst. These ligands also increased the proportion of IVG oocytes reaching metaphase II (53–57%) compared with controls (25%; P < 0.05). EGF alone or in combination with FSH increased (P < 0.05) the frequency of blastocyst formation (23% and 28%, respectively) compared with controls (13%). EGF treatment of maturing IVG oocytes produced blastocysts with more cells than other IVG groups (P < 0.05). It is concluded that gonadotrophins in vivo increase the sensitivity or responsiveness of cumulus cell-enclosed oocytes to EGF, thereby promoting both nuclear and cytoplasmic maturation. However, oocyte–granulosa cell complexes grown in vitro become responsive to EGF without gonadotrophin treatment. Thus, nuclear and cytoplasmic maturation of IVG oocytes is promoted by EGF treatment during meiotic maturation.
Key words: in-vitro-grown oocyte/mouse oocytes/nuclear and cytoplasmic maturation/preimplantation embryo
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Introduction |
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Oocytes from primordial follicles are arrested at prophase of the first meiotic division. During the growth and differentiation of diplotene stage oocytes, competence to resume meiosis is acquired following the progressive accumulation (Christmann et al., 1994
; Chesnel and Eppig, 1995; de Vantery et al., 1996), nuclear localization (Mitra and Schultz, 1996) and translational and post-translational modifications of cell cycle regulatory proteins (de Vantery et al., 1997). Nevertheless, meiotically competent oocytes are maintained in arrest by follicular somatic cells until after the preovulatory surge of gonadotrophins. Oocytes prepare for fertilization and embryogenesis by accumulating essential maternal factors and by undergoing genomic modifications during oocyte growth; final preparations are made during oocyte maturation. Oocyte maturation is often conceptually divided into nuclear and cytoplasmic processes. Nuclear maturation is a term that refers to the resumption of meiosis and progression to metaphase II. Cytoplasmic maturation is a more general term that refers to other maturational events not directly related to meiotic progression that prepare the oocyte for fertilization and preimplantation development (Eppig et al., 1994; Eppig, 1996).
Meiotically competent oocytes in most mammalian species undergo spontaneous maturation when released from their follicular environment and cultured (Pincus and Enzmann, 1935; Edwards, 1965). The mechanisms involved in germinal vesicle breakdown (GVB) as well as the cell signalling pathways driving the oocyte into metaphase II in response to the preovulatory gonadotrophin surge are not fully understood. However, both epidermal growth factor (EGF) and follicle stimulating hormone (FSH) induce nuclear maturation in mouse oocyte–cumulus complexes in vitro through a mechanism mediated by cumulus cells (Downs et al., 1988).
There are reports that EGF promotes nuclear maturation of human (Das et al., 1991), bovine (Kobayashi et al., 1994; Lorenzo et al., 1994; Lonergan et al., 1996; Rieger et al., 1998), and porcine (Singh et al., 1993; Ding and Foxcroft, 1994; Grupen et al., 1997) oocytes, as well as cytoplasmic maturation of mouse (Das et al., 1991), bovine (Kobayashi et al., 1994; Lonergan et al., 1996; Rieger et al., 1998), porcine (Ding and Foxcroft, 1994; Wang and Niwa, 1995; Grupen et al., 1997; Abeydeera et al., 1998a) and human (Goud et al., 1998) oocytes. However, effects of EGF are still not clear since there are other reports that EGF does not improve the cytoplasmic maturation of mouse oocytes (Merriman et al., 1998).
Systems for oocyte development in vitro provide an opportunity to characterize critical somatic and germ cell interactions that confer the mammalian oocyte with full developmental competence (Eppig and Schroeder, 1989; Spears et al., 1994; Cortvrindt et al., 1996; Eppig and O'Brien, 1996; Eppig et al., 1996). Nevertheless, the maturation of in-vitro-grown (IVG) oocytes is deficient compared with that of oocytes grown in vivo. For example, the onset of GVB is delayed in oocyte–granulosa cell complexes cultured from the preantral follicle stage (Eppig et al., 1996). Furthermore, a lower percentage of IVG oocytes reach the blastocyst stage after in-vitro fertilization compared with oocytes grown in vivo (Eppig and O'Brien, 1998).
In combination with gonadotrophins, EGF promotes nuclear maturation in oocytes obtained from intact preantral follicles cultured to the antral follicle stage (Boland and Gosden, 1994; Smitz et al., 1998). However, effects of EGF on cytoplasmic maturation of IVG mouse oocytes have not been evaluated. The objective of the present study was to determine the effects of EGF, alone or in combination with FSH, during in-vitro-maturation (IVM) on nuclear and cytoplasmic maturation of mouse oocytes. The effects of EGF were evaluated using maturing oocytes from three different sources; namely, in-vivo-grown oocytes obtained from large antral follicles of gonadotrophin-primed (PR) or unprimed (UPR) females, as well as IVG oocytes.
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Materials and methods |
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In-vivo-grown oocytes
In-vivo-grown oocyte–cumulus cell complexes were obtained from 22-day-old (C57BL/6JxSJL/J)F1 PR and UPR female mice. Gonadotrophin priming was performed by i.p. injection of 5 IU equine chorionic gonadotrophin (Gestyl; Diosynth, B.V. Oss, Holland) of 20-day-old females. Mice were killed 48 h later by cervical dislocation and ovaries collected in 2.5 ml Waymouth medium (MB752/1; GIBCO, Life Technologies, Grand Island, NY, USA) supplemented with 0.23 mM pyruvic acid, 50 mg/l streptomycin sulphate, 75 mg/l penicillin-G (Sigma, St Louis, MO, USA) and 1 mg/ml bovine fetuin purified according to Spiro (1960) at 37°C. Oocyte–cumulus complexes were obtained by puncturing large antral follicles from PR and UPR females with sterile needles. Cumulus cell-enclosed oocytes were selected and washed three times with fresh bovine serum albumin (BSA)-free Waymouth medium supplemented with bovine fetuin as described above, before allocation to IVM groups.
In-vitro-grown (IVG) oocytes
Oocyte–granulosa cell complexes of preantral follicles were isolated from 12-day-old mice and cultured as previously described (Eppig et al., 1992; Eppig and O'Brien, 1996). Briefly, oocyte–granulosa cell complexes were obtained from preantral follicles of 12-day-old females after collagenase digestion. Oocyte–granulosa cell complexes were washed three times in collagenase-free Waymouth medium supplemented as described above, and with 3 mg/ml BSA (crystallized; ICN Biochemicals, Aurora, OH, USA), 5 µg/ml transferrin, 5 ng/ml selenium (ITS; Collaborative Research Inc., Bedford, MA, USA) and 5 µg/ml insulin (Collaborative Research Inc.). Approximately 300–350 oocyte–granulosa cell complexes were cultured per well in a 6-well dish on collagen-coated membranes (Biocoat Collagen I Inserts, Becton Dickinson, Bedford, MA, USA) in 4 ml of medium. IVG medium was supplemented with 1 mg/ml fetuin, to prevent hardening of the zona pellucida (Eppig et al., 1996), but not with gonadotrophins or EGF. Cultures were maintained for 10 days at 37°C under a humidified atmosphere of 5% O2, 5% CO2 and 90% N2 in modular incubation chambers (Billups-Rothenberg, Del Mar, CA, USA). At the end of the culture period, oocyte–granulosa cell complexes were detached from the collagen membranes and collected in BSA-free Waymouth medium supplemented with 1 mg/ml fetuin. ITS mixture was omitted from the maturation medium. Complexes were washed three times in 2.5 ml of fresh medium and groups of 150–200 oocyte–granulosa cell complexes were allocated randomly to different maturation conditions.
In-vitro oocyte maturation (IVM)
Oocyte–cumulus complexes obtained from PR and UPR females as well as IVG complexes at the germinal vesicle (GV) stage were cultured in BSA-free Waymouth medium supplemented with either 10 ng/ml EGF (Collaborative Research Inc.), 10 ng/ml EGF plus 100 ng/ml FSH, or 100 ng/ml FSH. Control oocyte–cumulus and/or granulosa cell complexes were cultured in BSA-free Waymouth medium without any growth factor or hormonal supplementation. FSH (oFSH-20) was generously provided by The National Hormone and Pituitary Program of the National Institute of Diabetes and Diseases of the Kidney (NIDDK). At the end of 17–18 h culture, the stage of nuclear maturation was assessed after removing surrounding granulosa cells by continuous pipetting of oocyte–granulosa cell complexes in IVM medium. The percentage of oocytes undergoing GVB and polar body extrusion (metaphase II oocytes) in all groups was determined by observation under a dissecting microscope.
In-vitro fertilization and embryo culture
Oocytes at metaphase I or metaphase II were washed three times in Minimum Essential Medium (MEM; GIBCO, Life Technologies) supplemented with 3 mg/ml BSA. In-vitro fertilization and culture were performed as described previously (Ho et al., 1995; Eppig, 1999). Eggs were removed from fertilization drops after 4–6 h, rinsed twice in 2.5 ml MEM and cultured overnight in 500 µl droplets of fresh medium under washed mineral oil in modular incubation chambers as described above. At 30 h post fertilization, cleavage-stage embryos were rinsed twice with KSOM medium supplemented with essential and non-essential amino acids (KSOM/AA) and cultured to the blastocyst stage at 37°C in 1 ml KSOM/AA medium in borosilicate tubes (Ho et al., 1995; Eppig, 1999). Blastocyst-stage embryos obtained on day 5 post fertilization were fixed for cell number determination as described previously (Van de Sandt et al., 1990).
Statistical analysis
Data are presented as the mean percentage of at least three independent experiments, variation among replicates is presented as the standard error of the mean. The percentage of oocytes undergoing GVB and polar body extrusion as well as the proportion of 2-cell and blastocyst-stage embryos obtained after different maturation conditions were analysed using arcsin-transformed data and compared by analysis of variance (ANOVA) using StatView (SAS Institute Inc., Cary, NC, USA). When a significant F-ratio was defined by ANOVA, groups were compared using the Fisher's Protected Least Significant Difference (PLSD) post-hoc test using StatView software; when P 
0.05, the difference was considered significant. The mean number of nuclei at the blastocyst stage on day 5 post fertilization is presented and groups compared using notched box and whisker plots (StatView). Non-overlapping notches between box plots indicates significant differences (P < 0.05) (Kafadar, 1985).
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Results |
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Nuclear maturation of in-vivo-grown oocyte–cumulus complexes in response to EGF or FSH
IVM under serum-free conditions produced morphological changes in oocyte-associated granulosa cells, resulting in loosening of cell contacts and a progressive dispersion of cells into the culture medium. Maximal effects were observed after a combined stimulus provided by EGF and FSH. The nuclear maturation of oocyte–cumulus complexes obtained from PR females in controls (n = 193) or after treatment with EGF alone (n = 180), EGF in combination with FSH (n = 175) or FSH (n = 199) were evaluated in four independent experiments. The proportion of oocyte–cumulus complexes undergoing spontaneous meiotic maturation and polar body formation is illustrated in Figure 1A
. A high percentage of oocytes (87–94%) underwent GVB in all groups evaluated. However, 57% of control oocytes (P < 0.0001) remained at metaphase I after 17–18 h of maturation. The proportion of oocytes reaching metaphase II, as indicated by the production of a polar body, increased after maturation with EGF alone (73%; P < 0.001) or EGF in combination with FSH (67%; P < 0.05) relative to controls. Treatment of maturing oocytes with EGF promoted a higher frequency of progression to metaphase II (P = 0.03) than treatment with FSH alone.
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The response of oocyte–cumulus complexes obtained from UPR females in controls (n = 157) or after exposure to EGF (n = 170), EGF + FSH (n = 162) or FSH (n = 162) was evaluated after three independent experimental replicates and is illustrated in Figure 1B
. EGF as well as FSH alone or in combination increased the percentage of oocytes undergoing GVB (92–98%; P < 0.05) compared with controls (85%). However, a high percentage of oocytes (46–56%) remained at metaphase I in all groups evaluated. The percentage of oocytes reaching metaphase II increased (P < 0.05) when complexes were stimulated with EGF (48%) or EGF + FSH (51.5%) compared with controls (31%). Although experiments using PR and UPR females were not conducted simultaneously, it was noted that the frequency of progression of meiosis to metaphase II in response to EGF was always greater in the complexes from PR mice (73% versus 48%).
Effect of maturation conditions on the cytoplasmic maturation of in-vivo grown oocyte–cumulus complexes
EGF alone, or in combination with FSH, increased the proportion of oocytes that cleaved to the 2-cell stage after in-vitro fertilization (72%; P = 0.05) and (74%; P < 0.05) respectively (Figure 2A) compared with untreated oocytes. Maturation with FSH alone had no effect (P = 0.16) on the proportion of oocytes that cleaved to the 2-cell stage (65%) compared with controls (52%). Furthermore, EGF as a sole stimulus, or EGF + FSH, during oocyte maturation increased the proportion of oocytes reaching the blastocyst stage (64%; P < 0.05) and (60%; P = 0.05) respectively. No significant differences were observed in the frequency of blastocyst formation between control (38%) or FSH-treated oocytes (46%; P = 0.3).
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The proportion of oocytes from UPR females (59–69%) that cleaved to the 2-cell stage by 30 h after fertilization did not show any significant differences among controls or any of the treatment groups (Figure 2B
). Similar rates of blastocyst formation were also found amongst controls and all treatment groups (42–45%).
Comparison of the number of cell nuclei per blastocyst on day 5 after fertilization of oocytes from PR females (Figure 3A), demonstrated significant differences after treatment with EGF + FSH (mean = 121.6 ± 4.1) and FSH (121.6 ± 5.0) compared with controls (97.2 ± 5; P < 0.05). Furthermore, EGF significantly increased the total number of nuclei (145.2 ± 5.0) compared with oocytes exposed to EGF + FSH (P < 0.0001) or FSH as an only stimulus (P = 0.0004). The number of nuclei in blastocysts obtained from UPR females (Figure 3B) was higher after oocyte maturation with EGF (103.2 ± 4.0) or FSH alone (95.1 ± 3.5) or in combination (98.2 ± 3.7) compared with controls (83.6 ± 4.0; P < 0.05). However, no differences were observed among the different treatment groups.
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Effect of EGF or FSH on the nuclear and cytoplasmic maturation of IVG oocytes
The response of maturing IVG oocyte–granulosa cell complexes in controls (n = 385) or after treatment with EGF alone (n = 460), EFG in combination with FSH (n = 483) or FSH as a sole stimulus (n = 471) in three independent experiments is illustrated in Figure 4A
. After 17–18 h of maturation, 62% of IVG oocytes in the control group remained at the GV stage. EGF alone or in combination with FSH increased the percentage of oocytes (80–93%) undergoing GVB (P < 0.05). All treatment conditions resulted in a higher proportion (53–57%) of oocytes at metaphase II compared with controls (25%; P < 0.05). FSH treatment alone was not significantly different to the control. The proportion (38–49%; P < 0.05) of 2-cell-stage embryos (Figure 4B) obtained by 30 h after fertilization also increased compared with controls (23%). Moreover, EGF + FSH significantly increased the percentage of oocytes (49%) at the 2-cell stage compared with FSH-treated oocytes (38%; P < 0.05). EGF alone, or in combination with FSH, increased the percentage of oocytes that reached the blastocyst stage on day 5 after fertilization (23%; P = 0.05) and (28%; P < 0.05) respectively, compared with controls (13%). The number of nuclei in blastocysts is illustrated in Figure 5. Blastocysts derived from IVG control oocytes contained 69.9 ± 3.6 nuclei. EGF significantly increased the mean number of nuclei (97.1 ± 3.7; P < 0.01) per blastocyst compared with controls and compared with either EGF + FSH treatment (79.2 ± 3.0) or FSH used as a sole stimulus (77.3 ± 3.2; P < 0.0001).
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Developmental potential of IVG oocytes after fertilization at metaphase I or metaphase II
This experiment was designed to determine the developmental potential of IVG oocytes that remained at metaphase I or metaphase II after maturation with EGF and/or FSH. After 17–18 h IVM with EGF alone, or in combination with FSH, resulting metaphase I and metaphase II oocytes were separated based on the absence or presence of a polar body. Oocytes were fertilized and cultured separately as described above. The preimplantation developmental potential of IVG metaphase I stage oocytes is illustrated in Figure 6A
. Only a small percentage (6–8%) of oocytes that arrested at metaphase I reached the 2-cell stage by 30 h after in-vitro fertilization. No significant differences were observed in the percentage of cleaved or blastocyst-stage embryos (2–5%) obtained from controls or any of the treatment groups evaluated. In contrast, EGF during meiotic maturation significantly increased (P < 0.05) the cleavage rate observed (41%) after fertilization of metaphase II oocytes (Figure 6B) compared with controls (21%). Moreover, EGF also increased the proportion of blastocysts obtained (27%; P < 0.05) compared with controls (11%) and with FSH-treated maturing oocytes (12%).
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The mean numbers of nuclei per blastocyst obtained after fertilization of metaphase I-arrested oocytes (Figure 7A
) were not different among controls and any of the treatment groups (47 ± 4 to 60 ± 5). In contrast, metaphase II oocytes produced in the EGF-treated group (Figure 7B) had a higher number of blastomere nuclei per blastocyst (86.3 ± 3.4; P < 0.05) compared with controls (73.1 ± 3.7). Moreover, significant differences in blastomere number were also found between EGF- and FSH-treated oocytes (76.8 ± 3.3; P < 0.05). Blastocysts derived from metaphase II-arrested oocytes from all groups contained almost twice the number of nuclei per blastocyst (P < 0.05) as blastocysts derived from metaphase I-arrested oocytes.
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Discussion |
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The effects of EGF, EGF plus FSH, and FSH alone on nuclear and cytoplasmic maturation of mouse oocytes are summarized in Table I
. Clearly, EGF and FSH alone, as well as EGF and FSH together, promoted nuclear maturation whether oocytes were grown in vivo or in vitro when compared with control oocytes maturing without either of these ligands. The most consistent—and perhaps most important—of these effects was on the percentage of oocytes in which meiosis progressed to metaphase II, although this effect was not observed in FSH-treated oocytes from UPR females. EGF also promoted cytoplasmic maturation. There were notable differences, however, between the effects of EGF on the two groups of in-vivo-grown oocytes. In oocytes from PR females, EGF promoted increased percentages of oocytes that cleaved to the 2-cell stage and developed to blastocysts, and the number of cells per blastocyst. In contrast, in oocytes from UPR females, EGF raised only the number of nuclei in blastocysts but not the percentage of oocytes that cleaved to the 2-cell stage or developed to blastocysts. Surprisingly, the effect of EGF on the cytoplasmic maturation of IVG oocytes was more similar to in-vivo-grown oocytes from PR females; the percentages of oocytes competent to complete preimplantation development and the number of cells per blastocyst were increased.
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The high rates of GVB observed in oocyte–cumulus complexes obtained from PR and UPR females suggest that removal from the intrafollicular environment is sufficient to release the oocyte from the inhibitory signals maintaining meiotic arrest. However, further stimulation is required to drive oocytes into completion of the first meiotic division. In contrast, IVG oocytes required stimulation with either EGF or FSH to undergo GVB with high frequency. This is due to a greater ability of the oocyte-associated granulosa cells (OAGCs) enclosing the IVG oocytes to sustain meiotic arrest than the cumulus cells enclosing in-vivo-grown oocytes, since removal of OAGCs from IVG oocytes results in maturation of all competent oocytes without EGF or FSH stimulation (Eppig et al., 1996
). The OAGCs enclosing IVG oocytes have clear differences from cumulus cells, as demonstrated by qualitative and quantitative differences in protein synthetic patterns shown by high-resolution, two-dimensional gel electrophoresis (Latham et al., 1999). It has been suggested that the default pathway of granulosa cell differentiation is that of mural granulosa cells, and that this is abrogated by paracrine factors secreted by oocytes (Eppig et al., 1997a,b). However, in this culture system, factors from oocytes may undergo dilution to the point where they are unable to prevent some aspects of the default pathway from occurring (Eppig et al., 1998). Mural granulosa cells apparently play an important role in sustaining oocyte meiotic arrest, since separation of cumulus cell-enclosed oocytes from the mural granulosa results in GVB (Racowsky and Baldwin, 1989). It is therefore possible that some OAGCs enclosing IVG oocytes display similar enhanced activity as mural granulosa cells in sustaining meiotic arrest because of the inability of the oocytes to alter relevant aspects of the mural granulosa cell's default pathway. EGF and FSH, however, are able either to terminate the meiosis-arresting activity of these cells or to promote the generation of a positive signal to overcome it, or both.
Culture conditions during meiotic maturation can also have a significant impact on the progression to metaphase II (Downs and Mastropolo, 1997) and developmental potential (Van de Sandt et al., 1990). For example, several culture media support the completion of meiosis to a different extent according to the concentration and type of energy substrates contained. Moreover, variations in the culture environment (pH) may limit the availability of substrates required for proper metabolism in the oocyte–cumulus complex (Downs and Mastropolo, 1997). The frequency of polar body extrusion after maturation in Waymouth medium is lower compared with other oocyte maturation media (Downs and Mastropolo, 1997). However, maximal rates of blastocyst formation were found after in-vitro maturation of mouse oocytes in Waymouth medium (Van de Sandt et al., 1990). Culture conditions may explain the differences in the rates of polar body extrusion observed after EGF and/or FSH stimulation of mouse oocytes in the present study compared with those of others (Merriman et al., 1998). In the latter study, 84–96% of oocytes extruded a polar body when cultured in serum-supplemented MEM medium, albeit with a lower preimplantation development in vitro (Merriman et al., 1998).
In combination with gonadotrophins, EGF increased cumulus expansion and the rates of polar body extrusion in oocytes obtained using a whole follicle culture system (Boland and Gosden, 1994; Cortvrindt et al., 1998; Smitz et al., 1998). Maturation medium supplemented with EGF plus FSH and 5% serum supported polar body formation in 90% of oocytes (Boland and Gosden, 1994), with between 9% and 41% of oocytes reaching the blastocyst stage after in-vitro fertilization (Spears et al., 1994). Although recombinant meiosis-stimulating ligands were used in these experiments, oocyte maturation was conducted in the presence of serum, which could confound interpretation. Culture and maturation of oocyte–granulosa cell complexes in the presence of serum results in similar rates of cleavage and blastocyst formation after in-vitro fertilization compared with in-vivo-grown oocytes (Eppig and O'Brien, 1998). However, serum- or BSA-supplementation during meiotic maturation remain a source of undefined components that may `mask' the effects of growth factor or hormonal supplementation on cumulus expansion and meiotic progression. For example, stimulation with EGF alone had a comparable effect on nuclear maturation of pig oocytes as treatment with porcine follicular fluid (PFF) and supplementation of PFF-containing medium with EGF affords no additional improvements in the rates of oocyte maturation (Reed et al., 1993). Moreover, EGF supplementation to medium containing fetal calf serum or PFF had no effect on meiotic maturation or cleavage rates in bovine (Lonergan et al., 1996) or porcine oocytes (Abeydeera et al., 1998b) respectively.
EGF treatment of maturing oocytes from UPR females did not affect the percentage of embryos that completed preimplantation development, in contrast to the beneficial effect of EGF on the development of embryos derived from oocytes of PR females. EGF promoted higher cell numbers in the blastocysts of both groups. Thus, effects of EGF during oocyte maturation on blastocyst quality are separable from effects on frequency of blastocyst development. In addition, these results suggest that gonadotrophins in vivo increase the sensitivity or responsiveness of complexes to EGF, thereby promoting both nuclear and cytoplasmic maturation of in-vivo-grown cumulus cell-enclosed oocytes cultured with EGF. However, IVG oocytes behaved more similarly to oocytes from PR females than UPR females with regard to their response to the maturation-stimulating ligands; the frequency of completion of preimplantation development was increased, as well as the number of cells per blastocyst. This common response to these ligands during maturation occurred despite the development of oocyte–granulosa cell complexes in vitro in the absence of gonadotrophins. The implications of this observation are not clear, but it has also been noted that more blastocysts are produced per ovary after IVG than development in vivo when calculated on a per animal basis (Eppig and O'Brien, 1998). Perhaps both the responses of maturing oocytes in vitro to EGF noted above and the greater production of blastocysts per animal when oocytes were grown in vitro reflect suppressive or selective follicular regulatory processes occurring in vivo that are not functional under the conditions used in vitro.
In our previous studies on the preimplantation developmental competence of in-vivo-grown and in-vitro-matured oocytes, or oocytes both grown and matured in vitro, FSH has been used as the standard promoter of maturation. It is worthwhile, therefore, to consider whether EGF alone or EGF plus FSH offer benefits compared with FSH alone for promoting nuclear and cytoplasmic maturation in vitro. These comparisons are summarized in Table II. EGF or EGF plus FSH are always at least the equal of FSH alone for promoting nuclear and cytoplasmic maturation. There were no benefits of EGF, compared with FSH, detected for oocytes from UPR females. However, there were significant benefits noted for using EGF alone, or EGF and FSH together, for the nuclear and cytoplasmic maturation of oocytes from PR females and IVG oocytes. In fact, EGF treatment of IVG oocytes resulted in the production of blastocysts having a cell number equivalent to that of oocytes from UPR females. However, the frequency at which these IVG oocytes are able to complete preimplantation development is still below that for both groups of in-vivo-grown oocytes. Nevertheless, the use of EGF for the maturation of IVG oocytes still represents a significant advance in the technology for producing oocytes in vitro that are competent of undergoing preimplantation development.
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The effects of EGF and FSH on meiotic maturation of in-vivo-produced oocyte–cumulus complexes are mediated by cumulus cells (Downs et al., 1988
). The receptor for EGF (EGFr) is found in cumulus cells and expressed at low levels in the mouse oocyte (Wiley et al., 1992). Both EGF and transforming growth factor (TGF
) bind to the EGFr (Hollenberg and Gregory, 1977). However, their physiological role, if any, on meiotic maturation after the gonadotrophin surge in vivo is not clear. In the present study, a combined stimulus provided by EGF and FSH increased the frequency of blastocyst formation from in-vitro-grown oocyte–granulosa cell complexes as well as in-vivo-produced oocytes. However, an effect on blastomere proliferation was consistently observed only after treatment with EGF as a sole stimulus. It is possible that at the concentration used in this study, FSH may have interfered with the mitogenic stimulus provided by EGF. For example, recent experiments suggest that both EGF and FSH can effectively activate the mitogen-activated protein kinase (MAPK) cascade in granulosa cells (Maizels et al., 1998). However, increases in cyclic adenine monophosphate (cAMP) synthesis induced by FSH interfered with both activation of the MAPK signalling pathway in response to EGF (Wu et al., 1993) and its mitogenic effects in rat fibroblasts (Cook and McCormick, 1993). Further studies are thus required to understand the cellular and molecular processes associated with cytoplasmic maturation, as well as the possible physiological role of EGF and its interactions with gonadotrophins in preparing the mammalian egg for fertilization and embryogenesis.
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Acknowledgments |
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This research was performed as part of the National Cooperative Program on Non-Human In Vitro Fertilization and Preimplantation Development, and was funded by the National Institute of Child Health and Human Development (NICHD), NIH, through Cooperative Agreement HD21970. The scientific services of the Jackson Laboratory receive support from a Cancer Center Core Grant (CA34196) from the National Cancer Institute. We thank Drs Wes Beamer, Keith Latham, Andy Watson, and Randy Prather for their helpful comments in the preparation of this manuscript.
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Notes |
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1 To whom correspondence should be addressed
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References |
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Abeydeera, L.R., Wang, W.H., Cantley, T.C. et al. (1998a) Presence of epidermal growth factor during in vitro maturation of pig oocytes and embryo culture can modulate blastocyst development after in vitro fertilization. Mol. Reprod. Dev., 51, 395–401.[Web of Science][Medline]
Abeydeera, L.R., Wang, W.H., Prather, R.S. and Day, B.N. (1998b) Maturation in vitro of pig oocytes in protein-free culture media: fertilization and subsequent embryo development in vitro. Biol. Reprod., 58, 1316–1320.
Boland, N.I. and Gosden, R.G. (1994) Effects of epidermal growth factor on the growth and differentiation of cultured mouse ovarian follicles. J. Reprod. Fertil., 101, 369–374.
Chesnel, F. and Eppig, J.J. (1995) Synthesis and accumulation of p34cdc2 and cyclin B in mouse oocytes during acquisition of competence to resume meiosis. Mol. Reprod. Dev., 40, 503–508.[Web of Science][Medline]
Christmann, L., Jung, T. and Moor, R.M. (1994) MPF components and meiotic competence in growing pig oocytes. Mol. Reprod. Dev., 38, 85–90.[Web of Science][Medline]
Cook, S.J. and McCormick, F. (1993) Inhibition by cAMP of Ras-dependent activation of Raf. Science, 262, 1069–1072.
Cortvrindt, R., Smitz, J. and Van Steirteghem, A.C. (1996) In-vitro maturation, fertilization and embryo development of immature oocytes from early preantral follicles from prepubertal mice in a simplified culture system. Hum. Reprod., 11, 2656–2666.
Cortvrindt, R.G., Hu, Y., Liu, J. and Smitz, J.E. (1998) Timed analysis of the nuclear maturation of oocytes in early preantral mouse follicle culture supplemented with recombinant gonadotropin. Fertil. Steril., 70, 1114–1125.[Web of Science][Medline]
Das, K., Stout, L.E., Hensleigh, H.C. et al. (1991) Direct positive effect of epidermal growth factor on the cytoplasmic maturation of mouse and human oocytes. Fertil. Steril., 55, 1000–1004.[Web of Science][Medline]
de Vantery, C., Gavin, A.C., Vassalli, J.D. and Schorderet-Slatkine, S. (1996) An accumulation of p34cdc2 at the end of mouse oocyte growth correlates with the acquisition of meiotic competence. Dev. Biol., 174, 335–344.[Web of Science][Medline]
de Vantery, C., Stutz, A., Vassalli, J.D. and Schorderet-Slatkine, S. (1997) Acquisition of meiotic competence in growing mouse oocytes is controlled at both translational and posttranslational levels. Dev. Biol., 187, 43–54.[Web of Science][Medline]
Ding, J.C. and Foxcroft, G.R. (1994) Epidermal growth factor enhances oocyte maturation in pigs. Mol. Reprod. Dev., 39, 30–40.[Web of Science][Medline]
Downs, S.M. and Mastropolo, A.M. (1997) Culture conditions affect meiotic regulation in cumulus cell-enclosed mouse oocytes. Mol. Reprod. Dev., 46, 551–566.[Web of Science][Medline]
Downs, S.M., Daniel, S.A.J. and Eppig, J.J. (1988) Induction of maturation in cumulus cell-enclosed mouse oocytes by follicle-stimulating hormone and epidermal growth factor: evidence for a positive stimulus of somatic cell origin. J. Exp. Zool., 245, 86–96.[Web of Science][Medline]
Edwards, R.G. (1965) Maturation in vitro of mouse, sheep, cow, pig, rhesus monkey and human ovarian oocytes. Nature, 208, 349–351.[Medline]
Eppig, J.J. (1996) Coordination of nuclear and cytoplasmic oocyte maturation in eutherian mammals. Reprod. Fert. Dev., 8, 485–489.[Medline]
Eppig, J.J. (1999) Mouse oocyte maturation, fertilization, and preimplantation development in vitro. In Richter, J.D. (ed.), A Comparative Methods Approach to the Study of Oocytes and Embryos. Oxford University Press, Oxford, pp. 3–9.
Eppig, J.J. and O'Brien, M.J. (1996) Development in vitro of mouse oocytes from primordial follicles. Biol. Reprod., 54, 197–207.[Abstract]
Eppig, J.J. and O'Brien, M.J. (1998) Comparison of preimplantation developmental competence after mouse oocyte growth and development in vitro and in vivo. Theriogenology, 49, 415–422.[Web of Science][Medline]
Eppig, J.J. and Schroeder, A.C. (1989) Capacity of mouse oocytes from preantral follicles to undergo embryogenesis and development to live young after growth, maturation and fertilization in vitro. Biol. Reprod., 41, 268–276.[Abstract]
Eppig, J.J., Wigglesworth, K. and O'Brien, M.J. (1992) Comparison of embryonic developmental competence of mouse oocytes grown with and without serum. Mol. Reprod. Dev., 32, 33–40.[Web of Science][Medline]
Eppig, J.J., Schultz, R.M., O'Brien, M. and Chesnel, F. (1994) Relationship between the developmental programs controlling nuclear and cytoplasmic maturation of mouse oocytes. Dev. Biol., 164, 1–9.[Web of Science][Medline]
Eppig, J.J., O'Brien, M. and Wigglesworth, K. (1996) Mammalian oocyte growth and development in vitro. Mol. Reprod. Dev., 44, 260–273.[Web of Science][Medline]
Eppig, J.J., Chesnel, F., Hirao, Y. et al. (1997a) Oocyte control of granulosa cell development: how and why. Hum. Reprod., 12, Natl Suppl. JBFS, 2, 127–132.
Eppig, J.J., Wigglesworth, K., Pendola, F.L. and Hirao, Y. (1997b) Murine oocytes suppress expression of luteinizing hormone receptor messenger ribonucleic acid by granulosa cells. Biol. Reprod., 56, 976–984.[Abstract]
Eppig, J.J., O'Brien, M.J., Pendola, F.L. and Watanabe, S. (1998) Factors affecting the developmental competence of mouse oocytes grown in vitro: follicle-stimulating hormone and insulin. Biol. Reprod., 59, 1445–1453.
Goud, P.T., Goud, A.P., Qian, C. et al. (1998) In-vitro maturation of human germinal vesicle stage oocytes: role of cumulus cells and epidermal growth factor in the culture medium. Hum. Reprod., 13, 1638–1644.
Grupen, C.G., Nagashima, H. and Nottle, M.B. (1997) Role of epidermal growth factor and insulin-like growth factor-I on porcine oocyte maturation and embryonic development in vitro. Reprod. Fert. Dev., 9, 571–575.[Medline]
Ho, Y., Wigglesworth, K., Eppig, J.J. and Schultz, R.M. (1995) Preimplantation development of mouse embryos in KSOM: augmentation by amino acids and analysis of gene expression. Mol. Reprod. Dev., 41, 232–238.[Web of Science][Medline]
Hollenberg, M.D. and Gregory, H. (1977) Human urogastrone and mouse epidermal growth factor share a common receptor site in cultured human fibroblasts. Life Sci., 20, 267–274.[Web of Science][Medline]
Kafadar, K. (1985) Notched box-and-whisker plot. Encyclopedia of Statistical Sciences, 6, 367–370.
Kobayashi, K., Yamashita, S. and Hoshi, H. (1994) Influence of epidermal growth factor and transforming growth factor- on in vitro maturation of cumulus cell-enclosed bovine oocytes in a defined medium. J. Reprod. Fertil., 100, 439–446.
Latham, K.E., Bautista, D.M., Hirao, Y. et al. (1999) Comparison of protein synthesis patterns in mouse cumulus cells and mural granulosa cells: effects of follicle-stimulating hormone and insulin of granulosa cell differentiation in vitro. Biol. Reprod., 61, 482–492.
Lonergan, P., Carolan, C., Van Langendonckt, A. et al. (1996) Role of epidermal growth factor in bovine oocyte maturation and preimplantation embryo development in vitro. Biol. Reprod., 54, 1420–1429.[Abstract]
Lorenzo, P.L., Illera, M.J., Illera, J.C. and Illera, M. (1994) Enhancement of cumulus expansion and nuclear maturation during bovine oocyte maturation in vitro by the addition of epidermal growth factor and insulin-like growth factor I. J. Reprod. Fertil., 101, 697–701.
Maizels, E.T., Cottom, J., Jones, J.C. and Hunzicker-Dunn, M. (1998) Follicle stimulating hormone (FSH) activates the p38 mitogen-activated protein kinase pathway, inducing small heat shock protein phosphorylation and cell rounding in immature rat ovarian granulosa cells. Endocrinology, 139, 3353–3356.
Merriman, J.A., Whittingham, D.G. and Carroll, J. (1998) The effect of follicle stimulating hormone and epidermal growth factor on the developmental capacity of in-vitro matured mouse oocytes. Hum. Reprod., 13, 690–695.
Mitra, J. and Schultz, R.M. (1996) Regulation of the acquisition of meiotic competence in the mouse: changes in the subcellular localization of cdc2, cyclin b1, cdc25C and wee1, and in the concentration of these proteins and their transcripts. J. Cell Sci., 109, 2407–2415.[Abstract]
Pincus, G. and Enzmann, E.V. (1935) The comparative behavior of mammalian eggs in vivo and in vitro. I. The activation of ovarian eggs. J. Exp. Med., 62, 655–675.
Racowsky, C. and Baldwin, K.V. (1989) In vitro and in vivo studies reveal that hamster oocyte meiotic arrest is maintained only transiently by follicular fluid, but persistently by membrana/cumulus granulosa cell contact. Dev. Biol., 134, 297–306.[Web of Science][Medline]
Reed, M.L., Estrada, J.L., Illera, M.J. and Petters, R.M. (1993) Effects of epidermal growth factor, insulin-like growth factor-I, and dialyzed porcine follicular fluid on porcine oocyte maturation in vitro. J. Exp. Zool., 266, 74–78.[Web of Science][Medline]
Rieger, D., Luciano, A.M., Modina, S. et al. (1998) The effects of epidermal growth factor and insulin-like growth factor I on the metabolic activity, nuclear maturation and subsequent development of cattle oocytes in vitro. J. Reprod. Fertil., 112, 123–130.
Singh, B., Barbe, G.J. and Armstrong, D.T. (1993) Factors influencing resumption of meiotic maturation and cumulus expansion of porcine oocyte–cumulus cell complexes in vitro. Mol. Reprod. Dev., 36, 113–119.[Web of Science][Medline]
Smitz, J., Cortvrindt, R. and Hu, Y.X. (1998) Epidermal growth factor combined with recombinant human chorionic gonadotrophin improves meiotic progression in mouse follicle-enclosed oocyte culture. Hum. Reprod., 13, 664–669.
Spears, N., Boland, N.I., Murray, A.A. and Gosden, R.G. (1994) Mouse oocytes derived from in vitro grown primary ovarian follicles are fertile. Hum. Reprod., 9, 527–532.
Spiro, R.G. (1960) Studies on fetuin, a glycoprotein of fetal serum 1. Isolation, chemical composition, and physicochemical properties. J. Biol. Chem., 235, 2860–2869.
Van de Sandt, J.J.M., Schroeder, A.C. and Eppig, J.J. (1990) Culture media for mouse oocyte maturation affect subsequent embryonic development. Mol. Reprod. Dev., 25, 164–171.[Web of Science][Medline]
Wang, W.H. and Niwa, K. (1995) Synergetic effects of epidermal growth factor and gonadotropins on the cytoplasmic maturation of pig oocytes in a serum-free medium. Zygote, 3, 345–350.[Web of Science][Medline]
Wiley, L.M., Wu, J.X., Harari, I. and Adamson, E.D. (1992) Epidermal growth factor receptor mRNA and protein increase after the four-cell preimplantation stage in murine development. Dev. Biol., 149, 247–260.[Web of Science][Medline]
Wu, J., Dent, P., Jelinek, T. et al. (1993) Inhibition of the EGF-activated MAP kinase signaling pathway by adenosine 3',5'-monophosphate. Science, 262, 1065–1069.
Submitted on May 27, 1999; accepted on September 20, 1999.
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