Ca2+ current activity decreases during meiotic progression in bovine oocytes

Elisabetta Tosti1, Raffaele Boni2, and Annunziata Cuomo1

1 Cell Biology Unit, Stazione Zoologica Anton Dohrn, 80121 Napoli; and 2 Department of Animal Science, University of Basilicata, 85100 Potenza, Italy


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

By using the whole cell voltage-clamp technique, we studied changes in plasma membrane permeability at different meiotic stages of bovine oocytes. Follicular oocytes were matured in vitro and activated by Ca2+ ionophore. Oocytes at germinal vesicle (GV), germinal vesicle breakdown (GVBD), metaphase I (MI), metaphase II (MII), and meiosis exit were used for electrophysiological recording. By clamping the oocytes at -30 mV, we found that the L-type voltage-dependent Ca2+ channels were active at the GV stage and that their activity decreased after the GVBD stage. Furthermore, the resting potential decreased from the GV to the MI stage and increased again at MII. A significant decrease of the steady-state conductance occurred from the GV to the MI stage, followed by a sharp increase at the MII stage. With the addition of organic L-type Ca2+ channel blockers (nifedipine and verapamil), we inhibited the Ca2+ currents. However, only in the case of verapamil was there a decrease of in vitro maturation efficiency. Our results suggest that, in addition to the cumulus-oocyte junctions, the plasma membrane channels provide another mode of Ca2+ entry into bovine oocytes during meiosis.

oocyte maturation; L-type calcium channels; meiosis


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

IN MAMMALS, FOLLICULAR OOCYTES are arrested in the diplotene stage of the first meiotic prophase [germinal vesicle (GV) stage] until the start of follicle maturation release from the follicle environment (16). cAMP is suspected to maintain the meiotic arrest when transmitted from cumulus cells to the oocyte through gap junctions (1, 6, 15, 21). In response to the luteinizing hormone (LH) surge, cumulus cells transmit a Ca2+ signal to the oocyte (17), leading to gap junction regression (50). Simultaneously, cAMP levels decrease, which in turn releases the oocyte from meiotic arrest (46). These consecutive events are preceded by a relatively long lag phase lasting from the GV stage to germinal vesicle breakdown (GVBD), which is characterized by high protein synthesis and transcriptional activity (26). During this lag phase, cumulus-oocyte communication is open and the intracytoplasmatic cAMP levels are high (11). The oocyte then completes meiosis I by extruding the first polar body and begins the second meiotic division. This is characterized by a cell cycle block at metaphase II (MII) that lasts until fertilization. Sperm-oocyte interaction, as well as pharmacological substances, e.g., Ca2+ ionophore or ethanol, may induce meiosis completion and trigger early embryo development (8, 49, 57).

Meiosis and mitosis are regulated by two enzymes, histone 1 (H1) and mitogen-activated protein (MAP) kinase. H1, or maturation-promoting factor (MPF), is composed of cylin B and p34cdc2 subunits, which display a cyclical activity peaking at the metaphase stage (18, 56). MAP kinase is part of a kinase cascade that is likely initiated by c-mos (45). This pathway seems to be apparently involved in meiotic spindle organization, extrusion of the first polar body, and meiotic arrest at the MII stage (7).

It is well known that Ca2+ is involved in oocyte maturation (Ref. 25 for review). In the hamster (19) and mouse (5), a series of spontaneous Ca2+ oscillations occur in the oocyte after isolation from the follicle up to the GVBD stage. After these oscillations have subsided, Ca2+ does not affect further meiotic progression. In bovine and pig, no Ca2+ oscillations occur during meiosis progression; however, Ca2+ is necessary for meiotic progression since 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA), a Ca2+ chelator, causes a delay or block of meiosis (22). At fertilization, a new series of oscillations related to meiotic completion occurs (36). The Ca2+ ionophore, A-23187, induces meiosis resumption in oocytes blocked either at the GV stage (55) or MII (30, 48, 53) stage. Because extracellular Ca2+ is required for in vitro GVBD (14) and for first meiotic division (41), it appears that Ca2+ ion transport throughout the plasma membrane plays a functional role in maturation.

L-type Ca2+ channels are involved in numerous physiological processes (2, 24). These voltage-gated channels have been found in oocytes of the marine invertebrates (tunicates) (10) as well as in mammalian oocytes (39). In the mouse, Murnane and De Felice (38) showed a selective increase of these channels on the oocyte plasma membrane after puberty, corresponding to meiotic competence occurrence.

In this study, we have analyzed the electrical properties of the plasma membrane in bovine oocytes at different meiotic stages, focusing primarily on the activity of L-type voltage-dependent Ca2+ channels.


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

Materials. If not otherwise stated, all chemicals were purchased from Sigma Chemical (St. Louis, MO).

Oocyte source. Ovaries from slaughtered cows were collected from the abattoir and transported in a thermal bag at 30-35°C to the laboratory within 3-4 h of collection. The laboratory temperature was 30°C. Immature oocytes were collected from 2- to 8-mm follicles by an 18-gauge needle under controlled pressure (50-70 mmHg). Cumulus-oocyte complexes (COC) were isolated from the follicular fluid and washed three times with TCM199 supplemented with 5% FCS and 10 mM HEPES. The COC were then transferred into maturation medium (TCM199 supplemented with 10% FCS, 10 IU/ml LH, 0.1 IU/ml follicle-stimulating hormone, and 1 µg/ml 17beta -estradiol) (40 µl/COC) and left in an incubator at 39°C in 5% CO2 humidified air. Twenty-four hours later, the COC were freed from the cumulus cells by vortexing for 3 min and were parthenogenetically activated by 5-min exposure to 5 µM Ca2+ ionophore, A-23187, in Fert-TALP medium (42) as described by Liu et al. (30). The oocytes were then transferred in Fert-TALP medium and kept in the incubator until the electrical recording, scheduled 15-16 h after the ionophore treatment. At the time of the electrophysiological studies, batches of oocytes (control groups) were submitted to the same A-23187 treatment, followed by 3.5 h of incubation in culture dishes (Nunclon, Nunc, Denmark) that contained Fert-TALP supplemented with 2.5 mM 6-dimethylaminopurine. Finally, the oocytes were transferred in Fert-TALP medium covered with embryo-tested oil (Medicult, Denmark) and cultured in a gas mixture of 5% CO2, 7% O2, and 88% N2 for 1 day. Zygotes and embryos were cultured in SOF medium containing amino acids and BSA (51) in the previous gas mixture for 8 days postactivation for blastocyst development.

Electrophysiology. Electrical recording was performed at 37°C on oocytes at the following stages: GV, promptly isolated from follicles; GVBD, after 8 h of maturation; metaphase I (MI), after 12 h of maturation; and MII, after 24 h of maturation (47). Meiosis exit occurred 15-16 h after Ca2+ ionophore treatment, corresponding to the time span related to the decrease of MPF and MAP kinases (29) and the extrusion of the second polar body. Before micromanipulation, the oocytes at all stages were freed from the cumulus as described above, and the zona pellucida was removed by incubating the oocytes in 0.5% pronase for 1.5-2 min at 37°C.

The zona-free oocytes were subsequently placed in a recording chamber that contained 2 ml of Ham's F-10 (Mascia Brunelli, Italy). Oocytes were voltage clamped by standard techniques (4). Patch pipettes of 10-MOmega resistance and 1- to 2-µm tip diameter were filled with an intracellular-like solution that contained 70 mM KCl, 7 mM NaCl, 10 mM EGTA, and 10 mM HEPES, pH 7.4 and 280 mosmol/kgH2O. After obtaining a giga-seal, we set the pipette voltage to the desired negative potential and ruptured the patch. Observation of a stable negative resting potential signaled access to the cytosol. Depolarizing and hyperpolarizing voltage steps of 10 mV and 500 ms were applied to generate the voltage-dependent currents. External Ca2+ concentration was altered by adding 10 mM final concentration CaCl2/2H2O to the bath solution. Inhibitors of the L-type Ca2+ currents (verapamil and nifedipine) were also added to the bath as required. Oocytes at the GV stage were also in vitro matured in the presence of either 100 µM verapamil or 100 µM nifedipine. Currents were recorded on a List EP7 amplifier, and data were stored on a videocassette recorder tape for subsequent analysis. Data were analyzed using the ANOVA test of the General Linear Model procedure of Statistical Analysis Systems (46).

Meiosis progression assessment. Just before the experiment, samples from each stage were fixed with acetic-ethanol (1:3) and stained with acetic-lacmoid (Aldrich) for assessing meiotic progression. Confocal analysis was also performed on batches of oocytes to obtain additional information on the examined meiotic stages. The oocytes were fixed for 1 h in 2% formaldehyde in PBS, transferred in 0.01% Triton X-100 in PBS supplemented with 0.01% sodium azide, and kept at 4°C for 48 h. After being washed three times with PBS, the oocytes were stained to identify DNA with 0.01% propidium iodide supplemented with 0.01% Triton X-100 and 0.1 mM EDTA. After being washed three times in PBS, the oocytes were double stained after a 20-min incubation with either FITC-conjugated wheat germ lectin at the GV and GVBD stages to visualize the nuclear membrane or with FITC-conjugated anti-alpha -tubulin at the MI and MII stages to visualize the meiotic spindle. Finally, after being washed twice with PBS, the oocytes were scanned with an Olympus Fluoview confocal microscope.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

A total of 335 oocytes (14 replications) were parthenogenetically activated and produced a cleavage rate of 81.5 ± 6.2% and an expanded blastocyst production of 22.7 ± 4.7%. Examination of the fixed oocytes showed that the majority reached the stages studied, i.e., GV, 92.6% (25/27); GVBD, 85.7% (30/35); MI, 71.4% (20/28); MII, 84.2% (32/38); and meiosis exit, 73.3% (22/30).

The resting potentials of the oocytes at particular meiotic stages are shown in Fig. 1A. These potentials did not differ from the GV stage to the GVBD stage, decreased significantly (P < 0.05) at the MI stage, increased (P < 0.05) again at the MII stage and, finally, decreased at the meiosis exit stage. By clamping the cells at -30 mV and applying ramps of 10-mV depolarizing and hyperpolarizing steps, a series of whole cell currents were generated. The outward currents suggested a rectifier K+ channel similar to that described in the human oocyte by De Felice et al. (13). To obtain steady-state conductance, we plotted the peak current amplitude against the tip potential. This resulted in a linear relationship (Fig. 1B). The steady-state conductance current-voltage (I-V) significantly decreased (P < 0.01) from the GV to the GVBD and MI stages; it increased (P < 0.01) at MII and, finally, decreased again at the meiosis exit stage (Fig. 1C).


View larger version (20K):
[in this window]
[in a new window]
 
Fig. 1.   Resting potential (means ± SE) (A), current-voltage (I-V) curves of whole cell currents (B), and steady-state conductance (means ± SE) (C) patterns calculated at -30-mV clamp value in bovine oocytes throughout meiosis. A, B and C, D and E, F (P < 0.01); a, b (P < 0.05). GV, germinal vesicle; GVBD, germinal vesicle breakdown; MI, metaphase I; MII, metaphase II.

From the voltage clamp of -30 mV to test potentials, we observed an inward component of current activating in 30 ms and slowly inactivating in 250 ms, reaching a plateau in 500 ms. Typical leak-subtracted currents from -30 mV and I-V curves for the leak-subtracted currents at -30-mV voltage clamp are shown in Fig. 2 for each stage. Their amplitude, calculated as the difference between the peak and the steady state, significantly (P < 0.01) decreased from GVBD to the subsequent stages.


View larger version (35K):
[in this window]
[in a new window]
 
Fig. 2.   Left: confocal fluorescence images of bovine oocytes at different meiotic stages. GV stage shows the nuclear membrane and decondensed chromatin. GVBD is characterized by the absence of nuclear membrane. MI and MII stages show the meiotic spindle and first polar body (arrow) in the case of MII. Meiosis exit (Exit) is assessed by the presence of two polar bodies (arrows) close to the organizing female pronucleus. Middle: leak-subtracted currents recorded at +30-mV step potential, from the voltage-clamp values of -80 mV (upper trace) and -30 mV (lower trace) at the corresponding meiotic stages. Right: I-V relationships of the L-type Ca2+ currents at -30-mV holding voltage under standard culture conditions (0.5 mM Ca2+; red trace) and by increasing external Ca2+ at 10 mM (blue trace).

At -30-mV voltage clamp, the addition of 10 mM external Ca2+ to the bath increased the inward component at all stages. The I-V relationship of the peak amplitude at high Ca2+ concentration is shown in Fig. 2. The inward component increased at all the examined stages, with a maximum difference at the GV stage. Moreover, the pattern in the high Ca2+ regime was shifted toward more positive voltage values (Fig. 2). High Ca2+ also caused a transient hyperpolarization of the plasma membrane at both the GV and MII stages. Ca2+ currents were completely inhibited in the GV and GVBD stages by adding to the bath either nifedipine or verapamil at concentrations >5 µM. Moreover, maturing GV oocytes in the presence of either 100 µM verapamil or 100 µM nifedipine caused decreased (P < 0.01) cleavage efficiency (45 or 62% vs. 89%) and blastocyst development (10 or 31% vs. 32%) in the case of verapamil.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

In this study, we have shown that the plasma membrane of bovine oocytes undergoes profound electrical modification throughout meiosis. In particular, differences among meiotic stages were found in the resting potential as well as in steady-state conductance and voltage-gated Ca2+ channel activity. Little is known about the relationship between the resting potential of plasma membrane and the cell cycle. A change in plasma membrane polarization and ion permeability has already been described during maturation in invertebrate (37), amphibian (54), and mammalian (38, 43, 44) oocytes. We measured a stable negative resting potential and changes related to oocyte activation (12). It is likely that in the bovine oocyte, a low resting potential in both the GV and MII stages is associated with a "standby" status. In the case of GV, we suggest that the plasma membrane waits for a signal to resume meiosis, in which there is a large exchange of Ca2+ ions. As soon as cell cycle progression is resumed, the plasma membrane depolarizes. In such a case, however, it is difficult to explain the resting potential of GVBD, which represents the first stage of meiosis resumption. This may reflect meiosis vs. mitosis, bearing in mind that GV to GVBD is a period in which the metabolic oocyte activity is high (20, 26), cumulus-oocyte communication is intact (50), and cytoplasmic cAMP is elevated (11, 58).

The plasma membrane permeability, measured as steady-state conductance, is high at the GV stage and decreases during meiosis with little restoration at the MII stage. A similarity between these values at the GV and MII stages was reported in the mouse (38). Hence, the highest permeability corresponds to the two meiotic arrest phases. It is feasible that the high-ion exchange is related to the large metabolic activity of the GV stage or to the preparation of the plasma membrane for fertilization at the MII stage.

The I-V relationship of the leak-subtracted peak currents at different holding voltages, as well as the results obtained at high Ca2+ regime and the sensitivity at pharmacological agents, strongly suggest that these currents represent L-type Ca2+ channels. This is in agreement with previous findings in mouse (38) and invertebrate (10) oocytes. L-type Ca2+ channels have been demonstrated to underlay meiosis resumption in mussel (52), Pleurodeles (40), and in mammalian (39) oocytes. In bovine oocytes, the predominance of these channels at the GV and GVBD stages suggests a role for Ca2+ during the first meiotic resumption. Indeed, during maturation, the activity of plasma membrane Ca2+ channels decreases. This pattern may support the cytosolic Ca2+ rise at GV in addition to the LH and/or the growth factor-mediated Ca2+ surge via cumulus-oocyte communication (23, 31). In contrast, the low plasma membrane Ca2+ channel activity at the MII stage argues for a minor role of external Ca2+, whereas intracellular Ca2+ mobilization mechanisms appear to be more important for oocyte activation and fertilization.

The mechanism of how Ca2+ affects meiosis progression is unclear. However, we know that 1) BAPTA delays kinase activity and inhibits maturation (22); 2) Ca2+ may influence protein synthesis that is essential in maturational processes (28); and 3) Ca2+ modulates gap junction functionality, allowing cumulus-mediated intracytoplasmatic cAMP levels (58). In bovine oocytes, it has been shown that Ca2+ participates in the progression of meiosis, although spontaneous Ca2+ oscillations do not occur as in hamster and mouse oocytes (19, 22). D-Myo-inositol 1,4,5-trisphosphate (IP3)-induced Ca2+ release has been suggested as a primary mechanism for maturation of bovine oocytes because the cytoplasmic IP3 receptors increase in number during the meiotic progression (22). A minor role has been attributed to Ca2+ release through ryanodine receptors, which are poorly expressed in bovine oocytes (22).

Differential mechanisms of Ca2+ release in bovine oocytes could explain the effect of Ca2+ channel inhibitors during in vitro maturation. Since verapamil inhibits L-type Ca2+ channels, whereas nifedipine inhibits only the dihydropyridine (DHP)-sensitive L-type Ca2+ channels (33), we suppose that either 1) non-DHP-sensitive L-type Ca2+ channels play a role in maturation or 2) lower inward flux of Ca2+ caused by verapamil negatively affects maturation. On the basis of these findings, it seems likely that cumulus cells mediate intracytoplasmatic Ca2+ influx and, notwithstanding Ca2+ channel block, support the outcome of maturation in at least some oocytes.

In summary, these results suggest that in bovine oocytes, at the start of meiosis, in addition to the LH-mediated Ca2+ surge, Ca2+ entry arises through Ca2+ channels on the oocyte plasma membrane other than via gap junction cumulus-oocyte communication. Because the oocyte plasma membrane does not contain LH receptors, the initial Ca2+ influx comes from cumulus cells. This may cause a change in membrane potential and gating of voltage-dependent Ca2+ channels. The intracytoplasmatic Ca2+ rise may undergo a self-amplifying mechanism (Ca2+-induced Ca2+ release, IP3-induced Ca2+ release, or Ca2+-induced IP3 release) (3). If such a mechanism exists, it could potentiate the cumulus-oocyte communication necessary for metabolic exchange and the high cAMP levels during early maturation. High Ca2+ would then close the gap junctions (27), causing a drop in cAMP.

We have also shown that the MII stage is characterized by an increase of steady-state conductance due to K+ channels that is not accompanied by Ca2+ channel activity. In mammals, sperm-mediated oocyte activation is accompanied by a hyperpolarization of the plasma membrane due to Ca2+-activated K+ channels (9, 34, 35). Because we parthenogenetically activate oocytes by using Ca2+ ionophore, thus simulating the sperm-mediated Ca2+ surge, our data support the idea that in bovine oocytes external Ca2+ is not involved in meiosis exit. Indeed, the intracytoplasmatic Ca2+ surge may activate K+ channels. The decrease of Ca2+ channels during maturation may be correlated with the maturation of Ca2+ release mechanisms occurring at MII (32). These findings suggest that whereas external Ca2+ influences sperm-mediated Ca2+ elevation at fertilization, it mainly depends on intracellular Ca2+ stores.

In conclusion, during meiosis the plasma membrane of bovine oocytes undergoes a progressive depolarization and Ca2+ channel depletion. These findings provide new information and insight into the mechanisms and dynamics of meiosis.


    ACKNOWLEDGEMENTS

We thank Prof. L. J. De Felice and Dr. E. Brown for helpful comments and critical revision of the manuscript. We also thank G. Gargiulo for computer acquisition and photography.


    FOOTNOTES

Address for reprint requests and other correspondence: E. Tosti, Stazione Zoologica, Villa Comunale, 80121 Napoli, Italy (E-mail tosti{at}alpha.szn.it).

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Received 26 May 2000; accepted in final form 18 July 2000.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

1.   Aktas, H, Wheeler MB, First NL, and Leibfried-Rutledge ML. Maintenance of meiotic arrest by increasing cAMP may have physiological relevance in bovine oocytes. J Reprod Fertil 105: 237-245, 1995[Abstract].

2.   Bean, B. Two kinds of Ca2+ channels in canine atrial cells. J Gen Physiol 86: 1-30, 1985[Abstract].

3.   Berridge, MJ. Inositol trisphosphate and Ca2+ signaling. Nature 361: 315-325, 1993[ISI][Medline].

4.   Boni, R, Tosti E, Roviello S, and Dale B. Intercellular communication in in vivo- and in vitro-produced bovine embryos. Biol Reprod 61: 1050-1055, 1999[Abstract/Free Full Text].

5.   Carroll, J, Swann K, Whittingham DJ, and Whitaker M. Spatiotemporal dynamics of intracellular [Ca2+]i oscillations during the growth and meiotic maturation of mouse oocytes. Development 120: 3507-3517, 1995[Abstract/Free Full Text].

6.   Cho, WK, Stern S, and Biggers JD. Inhibitor effect of dibutyril cAMP on mouse oocyte maturation in vitro. J Exp Zool 187: 383-386, 1974[ISI][Medline].

7.   Choi, T, Rulong S, Resau J, Fukasawa K, Matten W, Kuryama M, Mansour S, Ahn N, and Vande Woude GF. Mos/mitogen-activated protein kinase can induce early phenotypes in the absence of maturation-promoting factor: a novel system for analyzing spindle formation during meiosis I. Proc Natl Acad Sci USA 93: 4730-4735, 1996[Abstract/Free Full Text].

8.   Dale, B. Oocyte activation in invertebrates and humans. Zygote 2: 373-377, 1994[Medline].

9.   Dale, B, Fortunato A, Monfrecola V, and Tosti E. A soluble sperm factor gates Ca2+-activated K+ channels in human oocytes. J Assist Reprod Genet 13: 573-577, 1996[ISI][Medline].

10.   Dale, B, Talevi R, and De Felice LJ. L-type Ca2+ currents in Ascidian eggs. Exp Cell Res 192: 302-306, 1991[ISI][Medline].

11.   Davis, JS, Weakland LL, West LA, and Farese LV. Luteinizing hormone stimulates the formation of inositol trisphosphate and cyclic AMP in rat granulosa cells: evidence for phospholipase C generated second messenger in the action of luteinizing hormone. Biochem J 238: 597-604, 1986[ISI][Medline].

12.   De Felice, LJ. Electrical properties of cells. New York: Plenum, 1997, p. 58-59.

13.   De Felice, LJ, Mazzanti M, Murnane M, and Cohen J. Patch-clamp and whole-cell recording from human oocytes (Abstract). Biophys J 53: 547, 1988.

14.   DeFelici, M, and Siracusa G. Survival of isolated, fully grown mouse ovarian oocytes is strictly dependent on external Ca2+. Dev Biol 92: 539-543, 1982[ISI][Medline].

15.   Dekel, N, and Beers WH. Rat oocyte maturation in vitro relief of cAMP inhibition by gonadotropins. Proc Natl Acad Sci USA 75: 4369-4373, 1978[Abstract].

16.   Edwards, RG. Maturation in vitro in mouse, sheep, cow, pig, rhesus monkey and human ovarian oocytes. Nature 208: 349-351, 1965[ISI][Medline].

17.   Eppig, JJ, and Downs SM. Chemical signals that regulate mammalian oocyte maturation. Biol Reprod 30: 1-11, 1984[Abstract].

18.   Fissore, RA, He CL, and Vande Woude GF. Potential role of mitogen-activated protein (MAP) kinase during meiosis resumption in bovine oocytes. Biol Reprod 55: 1261-1270, 1996[Abstract].

19.   Fujiwara, T, Nakada K, Shirakawa H, and Miyazaki S. Development of inositol trisphosphate-induced calcium release mechanism during maturation in hamster oocytes. Dev Biol 156: 69-79, 1993[ISI][Medline].

20.   Fulka, J, Jr, Flechon JE, Motlik J, Fulka J, and Jilck F. Effect of cycloheximide on nuclear maturation of pig and mouse oocytes. J Reprod Fertil 77: 281-286, 1986[Abstract].

21.   Gilula, NB, Epstein ML, and Beers WH. Cell-to-cell communication and ovulation. J Cell Biol 78: 58-75, 1978[Abstract].

22.   He, CL, Damiani P, Parys JB, and Fissore RA. Calcium, calcium release receptors, and meiotic resumption in bovine oocytes. Biol Reprod 57: 1245-1255, 1997[Abstract].

23.   Hill, JL, Hammar K, Smith PJS, and Gross DJ. Stage-dependent effects of epidermal growth factor on Ca2+ efflux in mouse oocytes. Mol Reprod Dev 53: 244-253, 1999[ISI][Medline].

24.   Hirano, Y, Fozzard HA, and January CT. Characteristics of L- and T-type Ca2+ currents in canine cardiac Purkinje cells. Am J Physiol Heart Circ Physiol 256: H1478-H1492, 1989[Abstract/Free Full Text].

25.   Homa, ST. Calcium and meiotic maturation of the mammalian oocyte. Mol Reprod Dev 40: 122-134, 1995[ISI][Medline].

26.   Hunter, AG, and Moor RM. Stage dependent effects of inhibiting RNA and protein synthesis on meiotic maturation of bovine oocytes in-vitro. J Dairy Sci 70: 1646-1651, 1987[ISI][Medline].

27.   Johnston, MF, Simon SA, and Ramon F. Interaction of anesthetics with electrical synapses. Nature 286: 498-500, 1980[ISI][Medline].

28.   Levesque, JT, and Sirard MA. Resumption of meiosis is initiated by the accumulation of cyclin B in bovine oocytes. Biol Reprod 55: 1427-1436, 1996[Abstract].

29.   Liu, L, Ju JC, and Yang X. Differential inactivation of maturation-promoting factor and mitogen-activated protein kinase following parthenogenetic activation of bovine oocytes. Biol Reprod 59: 537-545, 1998[Abstract/Free Full Text].

30.   Liu, L, Ju JC, and Yang X. Parthenogenetic development and protein patterns of newly matured bovine oocytes after chemical activation. Mol Reprod Dev 49: 298-307, 1998[ISI][Medline].

31.   Mattioli, M, Gioia L, and Barboni B. Calcium elevation in sheep cumulus-oocyte complexes after luteinizing hormone stimulation. Mol Reprod Dev 50: 361-369, 1998[ISI][Medline].

32.   Mehlman, LM, and Kline D. Regulation of intracellular calcium in the mouse egg: calcium release in response to sperm or inositol trisphosphate is enhanced after meiotic maturation. Biol Reprod 51: 1088-1098, 1994[Abstract].

33.   Miller, RJ. Multiple calcium channels and neuronal function. Science 235: 46-52, 1987[ISI][Medline].

34.   Miyazaki, S. Fertilization potential and calcium transient in mammalian eggs. Dev Growth Diff 30: 603-610, 1988[ISI].

35.   Miyazaki, S, Hashimoto N, Yoshimoto Y, Kishimoto T, and Igusa Y. Temporal and spatial dynamics of the period increase in intracellular free calcium at fertilization of golden hamster eggs. Dev Biol 118: 259-267, 1986[ISI][Medline].

36.   Miyazaki, S, and Igusa Y. Fertilization potential in golden hamster eggs consists of recurring hyperpolarizations. Nature 290: 706-707, 1981[ISI][Medline].

37.   Moreau, M, Leclerc C, and Guerrier P. Meiosis reinitiation in Ruditapes philippinarum (Mollusca): involvement of L-calcium channels in the release of metaphase I block. Zygote 4: 151-157, 1996[ISI][Medline].

38.   Murnane, J, and De Felice LJ. Electrical maturation of murine oocytes: an increase in calcium current coincides with acquisition of meiotic competence. Zygote 1: 49-60, 1993[Medline].

39.   Murnane, JM, De Felice LJ, and Cohen J. Development of ionic currents in mouse oocyte (Abstract). J Cell Biol 107: 4664, 1988.

40.   Ouadid-Ahidouch, H. Voltage gated calcium channels in Pleurodeles oocytes: classification, modulation and functional roles. Zygote 6: 85-95, 1998[ISI][Medline].

41.   Paleos, GA, and Powers RD. The effect of calcium on the first meiotic division of the mammalian oocyte. J Exp Zool 217: 409-416, 1981[ISI][Medline].

42.   Parrish, JJ, Susko-Parrish JL, Liebfried-Ruthledge ML, Critser ES, Eyestone WH, and First NL. Bovine in vitro fertilization with frozen/thawed semen. Theriogenology 25: 591-600, 1986[ISI].

43.   Powers, RD. Change in mouse oocyte membrane potential and permeability during meiotic maturation. J Exp Zool 221: 365-371, 1982[ISI][Medline].

44.   Powers, RD, and Biggers JD. Inhibition of mouse oocyte maturation by cell membrane potential hyperpolarization (Abstract). J Cell Biol 70: 352, 1976.

45.   Sagata, N, Daar I, Oskarsson M, Showalter SD, and Vande Woude GF. The product of mos-oncogene as a candidate "initiator" for oocyte maturation. Science 245: 643-646, 1989[ISI][Medline].

46.   SAS User's Guide/STAT (Release 6.03 Edition). Cary, NC: Statistical Analysis System Institute, 1988.

47.   Schultz, RM, Montgomery RR, and Belanof JR. Regulation of mouse oocyte maturation: implication of a decrease in oocyte cAMP and protein phosphorylation in commitment to resume meiosis. Dev Biol 97: 264-273, 1983[ISI][Medline].

48.   Sirard, MA, Florman HM, Leibfried-Rutledge ML, Barnes FL, Sims ML, and First NL. Time of nuclear progression and protein synthesis necessary for meiotic maturation of bovine oocytes. Biol Reprod 40: 1257-1263, 1989[Abstract].

49.   Steinhardt, RA, Epel D, Carrol EF, and Yanagimachi R. Is calcium ionophore a universal activator for unfertilized eggs? Nature 252: 41-43, 1974[ISI][Medline].

50.   Sutovsky, P, Flechon JE, Flechon B, Motlik J, Peynot N, Chesne P, and Heyman Y. Dynamic change of gap junction and cytoskeleton during in vitro culture of cattle oocyte-cumulus complexes. Biol Reprod 49: 1277-1287, 1993[Abstract].

51.   Thompson, JG. Defining the requirements for bovine embryo culture. Theriogenology 45: 27-40, 1996[ISI].

52.   Tomkoviak, M, Guerrier P, and Krantic S. Meiosis reinitiation of mussel oocytes involves L-type voltage gated calcium channels. J Cell Biochem 64: 152-160, 1997[ISI][Medline].

53.   Vincent, C, Cheek TR, and Johnson MH. Cell cycle progression of parthenogenetically activated mouse oocytes to interphase is dependent on the level of internal calcium. J Cell Sci 103: 389-396, 1992[Abstract/Free Full Text].

54.   Wallace, RA, and Steinhardt RA. Membrane potential of the amphibian oocyte. Dev Biol 57: 305-316, 1977[ISI][Medline].

55.   Wasserman, WJ, and Masui Y. Initiation of meiotic maturation in Xenopus laevis oocytes by the combination of divalent cations and ionophore A23187. J Exp Zool 193: 369-375, 1975[ISI][Medline].

56.   Wu, B, Ignotz G, Currie B, and Yang X. Dynamics of maturation promoting factor and its constituent proteins during in vitro maturation of bovine oocytes. Biol Reprod 56: 253-259, 1997[Abstract].

57.   Yanagimachi, R. Mammalian fertilization. In: The Physiology of Reproduction, edited by Knobil E, and Neill JD. New York: Raven, 1994, p. 189-317.

58.   Yoshimura, Y, Nakamura Y, Oda T, Ando M, Ubukata Y, Karube M, Koyama N, and Yamada H. Induction of meiotic maturation of follicle-enclosed oocytes of rabbits by a transient increase followed by an abrupt decrease in cyclic AMP concentration. J Reprod Fertil 95: 803-812, 1992[Abstract].


Am J Physiol Cell Physiol 279(6):C1795-C1800
0363-6143/00 $5.00 Copyright © 2000 the American Physiological Society