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BOR - Papers in Press, published online ahead of print December 19, 2007.
Biol Reprod 2007, 10.1095/biolreprod.107.063693
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BIOLOGY OF REPRODUCTION 78, 726–735 (2008)
DOI: 10.1095/biolreprod.107.063693
© 2008 by the Society for the Study of Reproduction, Inc.

Gamete Biology; 

Ca2+ Homeostasis Regulates Xenopus Oocyte Maturation1

Lu Sun 3, Rawad Hodeify 3, Shirley Haun 3, Amanda Charlesworth 4, Angus M. MacNicol 3 4 7, Subramaniam Ponnappan 5, Usha Ponnappan 5 6 7, Claude Prigent 8, and Khaled Machaca 2 3 7

Departments of Physiology & Biophysics,3 Neurobiology & Developmental Sciences,4 Geriatrics5 and Immunology & Microbiology,6 and the Arkansas Cancer Research Center,7 University of Arkansas for Medical Sciences, Little Rock, Arkansas 72205 CNRS UMR6061,8 Genetique et Developpement, Universite de Rennes 1, 35043 Rennes, France

ABSTRACT

In contrast to the well-defined role of Ca2+ signals during mitosis, the contribution of Ca2+ signaling to meiosis progression is controversial, despite several decades of investigating the role of Ca2+ and its effectors in vertebrate oocyte maturation. We have previously shown that during Xenopus oocyte maturation, Ca2+ signals are dispensable for entry into meiosis and for germinal vesicle breakdown. However, normal Ca2+ homeostasis is essential for completion of meiosis I and extrusion of the first polar body. In this study, we test the contribution of several downstream effectors in mediating the Ca2+ effects during oocyte maturation. We show that calmodulin and calcium-calmodulin-dependent protein kinase II (CAMK2) are not critical downstream Ca2+ effectors during meiotic maturation. In contrast, accumulation of Aurora kinase A (AURKA) protein is disrupted in cells deprived of Ca2+ signals. Since AURKA is required for bipolar spindle formation, failure to accumulate AURKA may contribute to the defective spindle phenotype following Ca2+ deprivation. These findings argue that Ca2+ homeostasis is important in establishing the oocyte's competence to undergo maturation in preparation for fertilization and embryonic development.

Aurora kinase A, calcium, gamete biology, meiosis, oocyte development

INTRODUCTION

Intracellular Ca2+ signals are ubiquitous and regulate a plethora of cell physiological and developmental processes, including contraction, secretion, transcription, cell proliferation, and cell death. The ultimate cellular response to environmental and intrinsic cues integrates a multitude of signaling cascades. Therefore, cellular and developmental decisions are the outcome of often competing signaling processes that include significant crosstalk and feedback mechanisms, allowing tighter regulation of critical differentiation pathways. A fundamental developmental decision that initiates multicellular organism development is oocyte maturation. Oocyte maturation is a complex cellular differentiation that prepares the egg for fertilization and early embryonic development [1]. In vertebrates, oocyte maturation includes transition through meiosis and arrest at metaphase II until fertilization. This nuclear maturation is coupled to cytoplasmic maturation, including dramatic morphological and mechanistic alterations that are essential for the egg-to-embryo transition [1, 2]. As might be expected for such a complex cellular differentiation, several signaling cascades regulate oocyte maturation, culminating in the activation of maturation-promoting factor (MPF) [1, 3].

The role of Ca2+ signaling in regulating oocyte maturation has been of wide interest, given that Ca2+ signals have been convincingly shown to play a role in mitosis and during egg activation at fertilization. During mitosis, Ca2+ signals are involved in nuclear envelope breakdown [4], anaphase onset [5], and cell cleavage [6]. A variety of genetic and biochemical evidence supports a role for Ca2+ and its downstream effectors calmodulin and CAMK2 in mitosis and following fertilization [7–9]. In contrast, the role of Ca2+ signals during vertebrate oocyte meiosis remains contentious [10–12]. In Xenopus, early reports argued that a Ca2+ rise is sufficient to induce oocyte maturation [13–15], and injection of Ca2+ buffers block maturation [10, 14]. Whether Ca2+ signals are induced downstream of progesterone during Xenopus oocyte maturation is also controversial [16–20]. These conflicting reports argue that in Xenopus the relationship between Ca2+ and oocyte maturation is complex. Studies regarding the role of Ca2+ during mammalian oocyte maturation also produce conflicting results [11, 21].

We have previously shown that in Xenopus, cytoplasmic Ca2+ negatively regulates meiosis entry, yet it is required for the completion of meiosis I [22]. This dual role of Ca2+ signals during oocyte maturation partly explains the controversy surrounding this topic. The goal of this study was to better define the signaling cascade(s) downstream of Ca2+ involved in regulating oocyte maturation.

MATERIALS AND METHODS

Gamete Preparation and Treatments

Xenopus oocytes were obtained as previously described in Machaca and Haun [23]. Frogs were purchased from Xenopus Express. Oocyte maturation was induced with 5 µg/ml progesterone. Germinal vesicle breakdown (GVBD) was detected visually by the appearance of a white spot at the animal pole of the cell and was confirmed by bisecting the cell into halves and detecting the nucleus after methanol fixation. The control L-15 solution contained 0.63 mM Ca2+. Ca2+ was buffered at 50 µmol/L in the low solution as calculated using the MaxChelator program (http://www.stanford.edu/~cpatton/maxc.html) by the addition of 0.58 mM EGTA. Maintenance and care for Xenopus frogs have been reviewed and approved by the UAMS Animal Care and Use Committee (IACUC).

Reagents

Lactacystin, MG-132, AIP, MLCK, MLCK control peptide, and calmidazolium were purchased from Calbiochem; thapsigargin was purchased from Molecular Probes; and fluphenazine dihydrochloride was purchased from Sigma. Messenger RNA for injection into oocytes was transcribed in vitro using the mMESSAGE mMACHINE T7 transcription kit (Ambion). The anti-Eg2 mAb was previously described [24].

Imaging Spindle Structure

Cells were fixed 3 h after GVBD in 100% methanol and stored at –20°C overnight. After rehydration in TBS/methanol (1:1) for 20 min, oocytes were washed twice with TBS for 15 min each and blocked for 3 h in TBS containing 2% BSA. Oocytes then were immunolabeled with an anti-{alpha}-tubulin monoclonal antibody (DM1A; Sigma) in TBS containing 2% BSA, followed by a Cy2-conjugated donkey anti-mouse secondary (Jackson) for 24 h each. The oocytes were washed five times in TBS for 24 h and stained with 1 µM Sytox Orange (Molecular Probes). After staining, cells were washed in TBS for 1 h, dehydrated in 100% methanol for 30 min, and cleared in benzyl alcohol-benzyl benzoate (1:2). Spindle structure images were collected on a Zeiss LSM510 confocal or on an Olympus Fluoview confocal. In some experiments, chromosome structure and polar body emission were visualized using Sytox Orange labeling without staining the spindle.

Polyadenylation Assay

Polyadenylation of endogenous mRNA species was assessed by RNA ligation-coupled RT-PCR, modified slightly from a previously described technique [25]. Total oocyte RNA (4 µg) from pools of five or six oocytes was ligated to 0.4 µg P1 anchor primer [25] in a 10-µl reaction using T4 RNA ligase (New England Biolabs) according to manufacturer's directions. The whole 10-µl RNA ligation reaction was used in a 50-µl reverse transcription reaction using Superscript III (Invitrogen), according to manufacturer's directions using 0.4 µg P1' as the reverse primer [25]. A total of 1 µl of this cDNA preparation was used in each 50-µl PCR reaction using Platinum Taq (Invitrogen), according to manufacturer's directions. Polymerase chain reaction was performed for 40 cycles using a 56°C annealing temperature and 1.5 mM final concentration of Mg2+. For optimal resolution, specific primers used were designed to be 70–90 nucleotides from the poly(A) addition site [26] according to sequence in GenBank: the accession numbers are Z17206 (Eg2, AURKA), X13311 (MOS), and J03166 (cyclin B1). The primers used were Cyclin B1: GTG GCA TTC CAA TTG TGT ATT GTT; AURKA: GTT TCA ATC TTG TAT GTC CTT TTA; and MOS: GTT GCA TTG CTG TTT AAG TGG TAA.

Plasmids

Wild-type (wt) and mutant CAMK2A clones were obtained from H. Schulman [27]. They were subcloned into pSGEM vector by cutting with PstI and HindIII. pSGEM-CAMK2A (wt), CAMK2A dominant-negative (dn, K42M), and CAMK2A constitutively active (ca, T286D) were linearized with NheI and transcribed with T7 polymerase. Rat calmodulin clones were provided by the Adelman Lab. Both wt and mutant calmodulin (with a D-to-A mutation in all four EF hand motifs) were cloned into pBF, as previously described [28]. RNA from calmodulin (CALM) clones was generated by linearizing them with MluI and transcribing with SP6 polymerase.

Proteasome Assay

Proteasome-specific activity was evaluated in oocyte lysates employing a biotinylated proteasome activity profiling probe, Ada-Lys (biotinyl)-(Ahx) 3-(Leu) 3-vinyl sulphone (BIOMOL). This probe specifically targets the active catalytic sites of the proteasome [29]. Oocytes were lysed in buffer containing 50 mM Tris-HCl (pH 7.4), 5 mM MgCl2, 250 mM sucrose, and 2 mM ATP. Lysates equalized for protein (30 µg) were labeled with 5 µM biotinylated probe in a reaction buffer containing 50 mM Tris-HCl (pH 7.4), 5 mM MgCl2, 2 mM ATP, and 2 mM dithiothreitol for 2 h at 37°C. Labeling reactions were quenched by the addition of SDS sample buffer and boiling for 5 min. Samples were resolved by 12% SDS-PAGE, transferred to nitrocellulose membranes, immunoblotted with streptavidin conjugated to horseradish peroxidase and detected using Enhanced Chemiluminescence (Amersham Biosciences). For the studies using proteasomal inhibitors, the inhibitors were present only during the maturation phase of the oocytes and were not added to the proteasome activity assay after oocyte lysis.

RESULTS

As we have previously shown, maturing oocytes in low extracellular Ca2+ (L-Ca; Ca2+ buffered at 50 µmol/L) enhances the rate of entry into meiosis as marked by GVBD (Fig. 1A) [22]. This enhancement is significantly more pronounced when intracellular Ca2+ stores are depleted with thapsigargin (Thaps) in L-Ca medium, thus depriving oocytes of both intracellular and extracellular Ca2+ signals (Fig. 1A). This faster maturation rate is associated, as would be expected, with premature activation of the MAPK-MPF signaling cascade [22], which regulates progression through oocyte maturation.


Figure 01
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  		FIG. 1. Ca2+ deprivation disrupts meiotic maturation. A) Representative GVBD time course for oocytes incubated in normal Ca2+ (0.6 mM, Con) or low-Ca2+ media (50 µM, L-Ca), or treated with 2 µM thapsigargin for 3 h before progesterone stimulation in L-Ca medium (Thaps). Germinal vesicle breakdown was scored every half hour after progesterone stimulation. B) MOS and Cyclin B1 RNA polyadenylation in control (Con) and thapsigargin-treated (Thaps) cells. Lysates were collected at hourly intervals after progesterone addition as indicated, at first GVBD (G0) and at GVBD50 (G50), when 50% of the oocytes in the population reached GVBD. For the G0 and G50 time points we collected lysates from oocytes that had undergone GVBD. C) Percentage of polar body emission levels from 1 to 4 h after GVBD in control (N-Ca), low-Ca2+ (L-Ca), and thapsigargin (Thaps) treatments. The data are mean ± SEM; the n number is indicated above each panel. D) Oocytes were fixed at the indicated times after GVBD and stained for tubulin and DNA to visualize spindle structure. Bar = 10 µm.

Progesterone-dependent Xenopus oocyte maturation is believed to be initiated by a cell surface G protein-coupled receptor [30, 31], rapidly (5–10 min) leading to inhibition of cAMP-dependent protein kinase (PKA) [32]. After a significant time delay (>1–2 h) and through largely unknown steps, mRNA translation of MOS and other regulators is induced through polyadenylation [33, 34], ultimately leading to the activation of MPF [3]. Maturation-promoting factor is the primary activity that regulates G2/M transition in both mitosis and meiosis, and it consists of a catalytic p34cdc2 Ser/Thr kinase subunit (CDK1) and a regulatory cyclin B subunit [35].

One of the earliest known molecular events downstream of PKA inhibition is translational regulation of maternal mRNAs through polyadenylation. Maternal mRNA polyadenylation is regulated in a temporal fashion through distinct 3' UTR regulatory elements [26, 36]. Some early-class mRNAs, such as MOS, are polyadenylated and translated prior to GVBD under the control of musashi 3' UTR regulator elements, whereas late-class mRNAs, such as cyclin B1, are polyadenylated and translated after GVBD under the control of late-acting cytoplasmic polyadenylation elements [26, 37]. We were interested in testing the effect of Ca2+ deprivation on the timing of polyadenylation of early- and late-class mRNAs. As shown in Figure 1B, in Ca2+-deprived oocytes, polyadenylation of MOS mRNA occurs as early as 2–3 h after progesterone addition (observed as an increase in PCR product size), compared with 4–5 h for control oocytes matured in normal Ca2+ medium. In contrast, the pattern of the late cyclin B1 mRNA polyadenylation occurs coincidently with GVBD in control and Ca2+-deprived oocytes. In Figure 1B, G0 refers to the time point at which the first cells in the population undergo GVBD, and G50 refers to when 50% of the cells in the population reach GVBD. These data show that the premature activation of the MOS-MAPK signaling cascade in Ca2+-deprived cells is likely due to a more rapid activation of the polyadenylation machinery regulating early-class maternal mRNA translational activation.

The experiments in Figure 1, A and B, were performed on oocyte populations from different donor females. There typically are significant variations in the rate of oocyte maturation in the population between different females, as illustrated in Figure 1, A and B. Oocytes in Figure 1A matured at a faster rate than those from the female in Figure 1B. Nonetheless, despite these differences in the rate of maturation, qualitatively the differences described in this manuscript in terms of Ca2+ deprivation and polyadenylation are consistently observed.

Another interesting phenotype of Ca2+-deprived oocytes is their inability to complete meiosis I and extrude the first polar body (Fig. 1C) [22]. Analyzing polar body emission over time reveals that the majority of oocytes matured in control medium complete meiosis I and extrude a polar body 2 h after GVBD. In contrast, oocytes matured in L-Ca medium, treated with thapsigargin, or injected with BATPA (not shown) fail to extrude a polar body up to 4 h after GVBD.

Careful analyses of spindle structure in Ca2+-deprived cells reveal that the primary defect is the inability of the first meiotic spindle to elongate in the absence of Ca2+ signals (Fig. 1D). Simply maturing oocyte in L-Ca medium (Fig. 1D) or blocking Ca2+ signals by BAPTA injection or thapsigargin treatment (not shown) inhibits spindle elongation. Ca2+-deprived cells assemble a compact spindle around the chromosomes but never progress to the elongated bipolar spindle observed in control cells (Fig. 1D). The inability of the spindle to elongate explains the defect in polar body emission observed in meiosis I.

Role of Ca2+-Calmodulin-Dependent Protein Kinase 2 (CAMK2)

We then tested the involvement of Ca2+-calmodulin-dependent protein kinase II (CAMK2) in the defects observed in Ca2+-deprived oocytes during maturation. This is because CAMK2 is a prominent downstream effector of Ca2+ signals and because it has been functionally implicated in meiosis. For example, CAMK2 has been shown to physically associate with the meiotic spindle in mouse eggs and to be important for the transition into anaphase during meiosis II [38, 39]. Furthermore, inhibition of CAMK2 activity in mouse oocytes blocks polar body emission [40], arguing that CAMK2 inhibition may be responsible for the defects observed following Ca2+ deprivation during Xenopus oocyte maturation. To test whether this is the case, we modulated CAMK2 activity by injecting oocytes with AIP, a potent and specific CAMK2 inhibitory peptide [41]. AIP injection (10 µM) did not affect the extent of GVBD in the population, but did significantly (P = 0.004) enhance the rate at which cells reach GVBD (Fig. 2A). Although this enhancement is consistent with a potential role for CAMK2 in effecting Ca2+ signals during oocyte maturation, it did not reach the levels of enhancement achieved by maturing oocytes in L-Ca medium or following Thaps treatment (Fig. 2A). Furthermore, AIP injection had no effect on the rate of GVBD when cells were pretreated with Thaps (Fig. 2A, T-AIP).


Figure 02
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  		FIG. 2. Effects of CAMK2 modulation on meiosis. A) Rates of oocyte maturation (top panel) and maximal GVBD levels reached (bottom panel). The rate of oocyte maturation was quantified as the time required for 50% of the oocytes in the population to undergo GVBD (G50). Rates were normalized to the time to G50 in normal Ca2+ medium (Con). AIP, a CAMK2 inhibitory peptide, was injected at 10 µM per oocyte 1 h before progesterone addition. Some oocytes were treated with Thaps before AIP injection (T-AIP), followed by progesterone stimulation. The L-Ca and Thaps treatments are as indicated in Figure 1 (n = 3). The letters above the bar graph indicate that means are statistically different (P < 0.02). The data are mean ± SEM. B) Oocytes were either matured in control (Con) or L-Ca medium (L-Ca), or injected with different CAMK2A mutant mRNAs and allowed to express for 1–2 days before progesterone addition. Oocytes were injected with wt CAMK2A (C-wt, 10 ng per oocyte), dn CAMK2A (C-dn, 10 ng per oocyte), and constitutively active CAMK2A (C-ca, 1 ng per oocyte). The times to G50 and maximal GVBD in the population are shown (n = 4–6 experiments). The letters above the bar graphs indicate statistical significance. The control and low groups are significantly different, P = 0.00015. The CAMK2A-injected groups were compared to each other. The CAMK2ca group was significantly different (P ≤ 0.034). C) Percentage of polar body (PB) emission in cells matured under control conditions (Con), low-Ca2+ medium (L-Ca), injected with AIP (10 or 20 µM), injected with wt CAMK2A (CAMK2-wt, 10 ng), or dn CAMK2A (CAMK2-dn, 10 ng). Oocytes were allowed to express CAMK2-wt for 1 day and CAMK2-dn for 3 days. Meiotic maturation was induced by progesterone (n = 20–23). The number is indicated above each bar. D) Western blot using anti-CAMK2A antibodies showing efficient expression of the different constructs. Lysates were from uninjected cells (Uninj.) or cells injected with different CAMK2A mutants as indicated.

We wanted to confirm the AIP result by expressing wt, dominant-negative (CAMK2dn), and constitutively active (CAMK2ca) CAMK2A mutants (Fig. 2B) [42]. Wild-type CAMK2A injection did not significantly (P = 0.083) affect the rate of maturation compared with the control (Fig. 2B). Injection of CAMK2dn tended to slow down the rate of maturation; however, the time to G50 did not reach statistical significance compared with wt CAMK2A injections (P = 0.064; Fig. 2B). CAMK2ca enhanced the rate of maturation (P = 0.034) compared with wt CAMK2A injection (Fig. 2B). None of the CAMK2A injections significantly affected the extent of maturation (Fig. 2B). These results argue that CAMK2 is not a downstream effector of Ca2+ signals in terms of setting the rate of maturation because, in contrast to Ca2+ deprivation, CAMK2dn slowed down the rate of maturation, and CAMK2ca enhanced it (Fig. 2B). If CAMK2 was indeed a downstream Ca2+ effector, one expects that CAMK2dn should enhance the rate of maturation in a fashion similar to Ca2+ deprivation. Given the fact that CAMK2 could impact a multitude of signaling cascades, the observed effects of overexpression of either the constitutively active or dn mutants of CAMK2 could be indirect. This is plausible, since AIP produces an opposite effect of CAMK2dn on the rate of maturation (Fig. 2B). Therefore, together these results argue that CAMK2 is not a major player in modulating the rate of oocyte maturation.

We then tested the role of CAMK2 in completion of meiosis I and first polar body emission. Maturing oocytes in L-Ca medium inhibit polar body emission (Fig. 2C). In contrast, AIP injection or injection of wt or dn CAMK2A did not inhibit the ability of oocytes to extrude a polar body (Fig. 2C). It was not possible to test the effect of the constitutively active CAMK2A mutant in these experiments, because cells expressing CAMK2ca did not survive long enough after GVBD to allow for first polar body extrusion. These results indicate that CAMK2 is not important for spindle formation in Xenopus oocytes.

Role of Calmodulin (CALM)

Calmodulin (CALM) is an excellent candidate for transmitting Ca2+ signals to downstream effectors during oocyte maturation, especially in the context of the spindle elongation defect. CALM contains four Ca2+-binding EF hands and acts as the Ca2+ sensor for many cellular functions [43]. CALM localizes to the spindle poles [44]. Furthermore, CALM confers Ca2+ sensitivity to dynein, where Ca2+-CALM binds to dynein and enhances its motor activity [45, 46]. This is relevant, since dynein localizes to the spindle [47, 48], regulates microtubule dynamics [49], and is critical for the formation of a normal spindle in Xenopus [49, 50]. In addition to its interactions with dynein, CALM interacts with other components at the spindle pole, primarily the 110-kDa spindle pole component, and this interaction is essential for normal spindle formation [51–53]. It is therefore possible that the spindle defect observed in meiosis I of Ca2+-deprived cells is due to disruption of CALM function/localization. In addition, CALM has been postulated to play a role in the progression of oocyte maturation in Xenopus, since CALM levels increase during maturation [54], and CALM injection has been reported to induce low levels of maturation, although this result was not reproducible [18, 55–57].

To test the role of CALM in oocyte maturation and meiosis I spindle formation, we inhibited CALM function with calmidazolium (CDZ) or fluphenazine (FLZ). Calmidazolium is an imidazole compound that acts as a specific CALM antagonist by strongly binding to the hydrophobic surface of CALM, preventing the binding of Ca2+ and, thus, the correct folding of active CALM [58]. Fluphenazine is an antipsychotic drug that blocks CALM interaction with a variety of downstream effectors [59]. Calmidazolium did not affect the rate or extent of entry into meiosis, nor did it significantly alter spindle structure (Fig. 3A). It did, however, have a significant effect on the ability of the oocyte to extrude a polar body (Fig. 3A), although these effects were not nearly as dramatic as maturing oocytes in the absence of extracellular Ca2+. In contrast, as previously reported for FLZ [60] and other CALM antagonists [61, 62], FLZ enhances the rate of oocyte maturation in a manner consistent with Ca2+ deprivation without affecting maximal GVBD (Fig. 3B). Furthermore, FLZ was more likely to lead to spindle defects, although the effects were quite variable (Fig. 3B) yet, surprisingly, this did not affect the ability of the oocytes to extrude a polar body.


Figure 03
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  		FIG. 3. Effect of Calmodulin inhibition on oocyte maturation. A) Oocytes were incubated in L-15 media containing dimethylsulfoxide (Con) or 1 µM calmidozolium (CDZ) for 3 h before progesterone addition. The upper panels in A show a representative GVBD time course and average data for the maximal GVBD reached and time to GVBD50 (n = 8); the lower panels show representative images of spindle structure at 3 h after GVBD. Percentages of cells with normal spindle structure and normalized polar body emission levels are shown (n = 4). Polar body emission rates were significantly decreased (P = 0.0034) following CDZ treatment. B) Oocytes were incubated in L-15 alone (Con) or L-15 containing 20 µM FLZ (FLZ) for 3 h before progesterone addition. A representative GVBD time course and averaged data for maximal GVBD and rate to GVBD50 are shown in B, upper panels (n = 4). Fluphenazine significantly (P = 10–5) enhanced the time to G50. The lower panels in B show spindle structure at 3 h after GVBD and the percentage of cells with normal spindles and normalized polar body emission levels (n = 4). Summary data are presented as the mean ± SEM. Bars = 10 µm.

The differential effect of FLZ and CDZ on the rate of maturation is not surprising, since CDZ has been previously reported not to effectively inhibit certain CALM-dependent processes where CALM is tightly associated with its downstream effector, as for L-type Ca2+ channels [63] and small-conductance Ca2+-activated K channels [64]. Therefore, the FLZ data argue that CALM is involved in regulating the rate of oocyte maturation. However, attempts to confirm the effects of a pharmacological block of CALM on the rate of maturation using either injection of a CALM inhibitory peptide (MLCK peptide) or a CALM mutant with all four EF hands mutated (CALMs 1, 2, 3, 4) and thus unable to bind Ca2+ were unsuccessful (Table 1). In either case, these treatments were ineffective at enhancing the rate of maturation, bringing into question the importance of CALM in regulating the rate of maturation. Therefore, as is the case with CAMK2, the fact that different approaches to interfere with CALM function produce disparate results in terms of controlling the rate of oocyte maturation argues that CALM is not a critical determinant of the processes by which Ca2+ regulates progression through maturation.


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  		TABLE 1. Effects of calmodulin (CALM) inhibition on oocyte maturation.

In contrast to the effects of CALM inhibition on the rate of maturation, consistent results were obtained on polar body emission. None of the experimental manipulations of CALM inhibited polar body emission to levels similar to Ca2+ deprivation (Fig. 3 and Table 1). This argues that the Ca2+ effects on spindle formation and polar body emission are not mediated through CALM.

Aurora Kinase A (AURKA)

The inability of Ca2+-deprived oocytes to complete meiosis normally and extrude a polar body is interesting because this nuclear defect occurs while the kinetics of the MAPK-MPF cascade proceed normally [22]. These observations suggest that Ca2+ deprivation uncouples the kinase signaling cascade regulating oocyte maturation from the nuclear meiotic events. Several other experimental manipulations, such as inhibition of the MAPK cascade [65], downregulation of MPF [66], and inhibition of protein synthesis [67], lead to a meiosis I arrest that is coupled to an interphaselike state. In contrast, Ca2+ deprivation does not affect the MAPK-MPF kinase signaling cascade, yet it disrupts spindle elongation in meiosis I and prevents polar body emission. Interestingly, manipulation of the Aurora kinases produces a similar phenotype. Downregulation of Aurora B in Caenorhabditis elegans inhibits polar body extrusion in meiosis I [68]. In Xenopus oocytes, injection of anti-AURKA antibodies inhibits polar body extrusion without affecting chromosome condensation or MPF activity [69]. In addition, expression of a constitutively active AURKA in Xenopus oocytes leads to a meiosis I arrest, with high MPF activity and condensed chromosomes but defective spindles [70]. Therefore, manipulating AURKA activity in Xenopus oocytes phenocopies Ca2+ deprivation, arguing that AURKA regulation may have a Ca2+-dependent component. To test this possibility, we measured the levels of AURKA in control and Ca2+-deprived oocytes throughout the maturation process (Fig. 4A). As previously shown [71], AURKA gradually accumulates during maturation, especially after GVBD (Fig. 4A). In contrast, AURKA accumulation is significantly attenuated during maturation of Ca2+-deprived, Thaps-treated oocytes (Fig. 4A). Although the tubulin loading control shows more protein loading toward the later time points in both the low Ca2+ and Thaps treatments (Fig. 4A), this still correlates with lower accumulated levels of AURKA protein, showing that the ability of cells deprived of Ca2+ signals to accumulate AURKA is compromised.


Figure 04
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  		FIG. 4. Kinetics of AURKA accumulation. A) Oocytes were incubated in normal calcium L-15 medium (Con) or low-Ca2+ medium and treated with 2 µM of thapsigargin for 3 h (Thaps) before progesterone stimulation. At each time point as indicated, lysates were assayed for AURKA levels using an anti-Eg2 monoclonal antibody (1C1). The time course is divided into two phases: hours after progesterone addition, and hours after GVBD. The relative intensity of every band normalized to the levels at the 1 h after GVBD time point. Below the bar graph, a Western blot for tubulin is shown as a loading control. These data are representative of three similar experiments. B) Oocytes were treated as described in A. Cell lysates were collected from immature oocytes (Ooc), at 3 h after GVBD (Egg), at first GVBD (G0), at GVBD50 (G50), and at GVBD100 (G100). For the GVBD50 time point we collected lysates from oocytes that had undergone GVBD (white spot, w) and those that had not (no white spot, nw). Tubulin was used as a loading control. C) AURKA RNA polyadenylation in Con and Thaps-treated cells. The same lysates as in Figure 1B were used for this experiment.

Supporting the time course data, when lysates are prepared at different time points during maturation, the inhibition of AURKA accumulation in Ca2+-deprived cells is apparent (Fig. 4B). For this experiment, we isolated lysates from: (1) oocytes (oocytes); (2) eggs, which have undergone GVBD >3 h earlier; (3) when cells in the population first reach GVBD (G0); (4) when 50% of the cells in the population reach GVBD (G50)—in this case, lysates from both cells with a white spot (w) and those without (nw) were collected—and (5) when 100% of the cells in the population reach GVBD (G100). Similar to the time course data (Fig. 4A), Ca2+-deprived cells are less efficient at accumulating AURKA compared with controls (Fig. 4B).

One possibility that we considered to explain the lack of AURKA accumulation in Ca2+-deprived cells was the failure to polyadenylate the early class AURKA mRNA [26, 72]. However, similar to the MOS mRNA (Fig. 1B), Thaps treatment led to an accelerated initiation of endogenous AURKA mRNA polyadenylation (Fig. 4C). This argues that the lack of AURKA accumulation following disruption of Ca2+ homeostasis is not due to differential polyadenylation, but rather either reflects a specific defect in AURKA mRNA translation or posttranslational stabilization of AURKA protein.

Proteasome Modulation

One possibility to explain the AURKA results is modulation of proteasome activity following Ca2+ deprivation. The 26S proteasome regulates protein degradation following polyubiquitination and has been implicated in oocyte meiosis in several species. Proteasome activity increases in a Ca2+-dependent fashion during ascidian and Xenopus egg activation [73, 74]; the proteasome associates with the spindle during rat oocyte meiosis; and proteasome inhibitors block polar body extrusion [75]. Furthermore, the levels of certain critical regulators of Xenopus oocyte maturation, such as CPEB and MOS, are dependent on the proteasome [76, 77]. Based on these findings, we were interested in testing the role of the proteasome in regulating oocyte maturation following Ca2+ deprivation. Proteasome activity was not affected when cells were matured in L-Ca medium or when they were treated with thapsigargin (Fig. 5A). Proteasome activity is reflected in band intensity of the different proteasomal subunits, given the specific labeling using a biotinylated proteasome activity profiling probe, as explained in Materials and Methods (Fig. 5A). We further tested the ability of two proteasome inhibitors, lactacystin and MG132, to block Xenopus oocyte proteasome activity. Lactacystin (at the doses employed) was ineffective at blocking the proteasome, whereas MG132 inhibited proteasome activity in a dose-dependent fashion (Fig. 5A). Nevertheless, oocytes treated with either inhibitor matured with kinetics similar to those in control cells (Fig. 5B), showing that proteasome inhibition is not sufficient to enhance the rate of oocyte maturation. These data argue that modulation of proteasome function is not responsible for the defective AURKA accumulation, since proteasome activity is not altered following Ca2+ deprivation. Furthermore, the activation of the MAPK cascade, which depends on MOS accumulation, occurs earlier in Ca2+-deprived cells [22], consistent with a lack of effect of Ca2+ deprivation on proteasome activity.


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  		FIG. 5. Proteasome activity. A) Lysates were collected from oocytes incubated in normal Ca2+ medium (Con), low-Ca2+ medium (L-Ca), treated with 2 µM Thaps for 3 h in L-Ca (Thaps), or treated with the indicated concentrations of lactacystin (Lact) or MG-132 (MG132) for 1 h before progesterone stimulation. The proteasome activity assay was performed as described in Materials and Methods. Molecular weight markers are indicated to the left of the gel (Mr = x 106). B) Oocytes were untreated (Con) or incubated with lactacystin (Lact) and MG-132 (MG132) for 1 h before progesterone stimulation. The rate of oocyte maturation was quantified as the time required for 50% of the oocytes to reach GVBD (G50). The G50 for each indicated treatment was normalized to G50 in the Con treatment. Summary data are presented as the mean ± SEM; the n number is indicated on each bar.

DISCUSSION

Calmodulin and CAMK2

Depriving Xenopus oocytes of Ca2+ signals does not affect their ability to enter meiosis or undergo GVBD, but it results in a dramatically disrupted spindle structure and a lack of polar body emission [22]. In Ca2+-deprived oocytes, the meiosis I spindle does not elongate into a bipolar spindle (Fig. 1D), which could form the basis for the lack of polar body emission. It is unclear why disrupting Ca2+ homeostasis leads to defective spindles. One possibility is that Ca2+ chelation directly affects microtubule assembly, as previously described [78, 79]. Another explanation for the Ca2+-dependent meiosis I defect would be faulty signaling directly downstream of Ca2+. This was the incentive for us to test two ubiquitous Ca2+-dependent downstream effectors, calmodulin and CAMK2. However, our results argue that neither of these effectors is involved in the defects observed following Ca2+ deprivation during oocyte maturation. Interfering with CALM or CAMK2 function using various approaches produces disparate effects that are inconsistent with the effect of Ca2+ deprivation on the rate of progression of oocyte maturation (Figs. 2 and 3). This argues that CALM-CAMK2 is not a critical effector of Ca2+ signals during oocyte maturation in terms of setting the rate of maturation. In a similar fashion, disruption of either CALM or CAMK2 function did not significantly and consistently affect the ability of maturing oocytes to extrude a polar body. We conclude from these results that the phenotypes observed following Ca2+ deprivation (i.e., faster maturation and lack of polar body emission) are not due to disruption of the CALM-CAMK2 signaling axis.

Aurora Kinase A

Ca2+ deprivation results in a faster maturation rate, raising the possibility that this could underlie the meiosis I defect. In that context, tight coordination of different signaling cascades is essential for timely progression through oocyte maturation [3]. We were therefore interested in testing whether depriving oocytes of Ca2+ signals somehow disrupts this tight coordination. AURKA was an obvious target to test, because altering AURKA activity phenocopies the Ca2+ deprivation-dependent spindle defect [69, 70]. Furthermore, the Aurora family encodes Ser/Thr kinases that localize to the spindle poles and are crucial for spindle assembly during both meiosis and mitosis [80–82].

AURKA protein gradually accumulates during oocyte maturation (Fig. 4A), and this increase is important for AURKA function during meiosis [71]. Although the ability of AURKA to accumulate in Ca2+-deprived oocytes is compromised (Fig. 4, A and B), this defect is not correlated with a failure to polyadenylate AURKA mRNA prior to GVBD (Fig. 4C). These findings indicate that unlike the translation of MOS and activation of MAP kinase signaling, AURKA mRNA translation is specifically attenuated, or the newly synthesized AURKA protein is destabilized. The failure to accumulate AURKA protein could contribute to the spindle elongation defect and the inability of Ca2+-deprived oocytes to extrude a polar body, because the levels of AURKA are critical for normal progression through meiosis [69, 70]. However, it is not clear at this point whether there exists a cause-effect relationship between the lack of AURKA accumulation and the defective spindle phenotype observed in Ca2+-deprived cells. Unfortunately, directly testing this relationship is technically difficult because the levels of AURKA have to be tightly controlled. Either interfering with AURKA function or increasing it leads to defective spindles and disrupts meiosis I completion [69, 70].

It remains unclear why AURKA accumulation is disrupted following Ca2+ deprivation, although we can rule out direct effects on the proteasome (Fig. 5). Interfering with Ca2+ signaling enhances the rate at which oocytes mature by activating the MAPK-MPF cascade earlier [22]. This is due to a more rapid polyadenylation of early-class mRNAs, such as MOS (Fig. 1B). AURKA mRNA is also polyadenylated earlier in Ca2+-deprived cells, but this is associated with defective accumulation of AURKA. The fact that AURKA accumulation is not necessarily dependent on polyadenylation [71] may partially explain the differential effect of Ca2+ deprivation on the MOS-MAPK cascade versus AURKA. Accumulation of AURKA is a balance between both mRNA translation and AURKA protein degradation rates. Since proteasome activity is not directly affected in Ca2+-deprived oocytes, Ca2+ deprivation could be inhibiting AURKA accumulation by affecting AURKA mRNA de-repression independently of polyadenylation and/or AURKA protein stability.

Together, our data argue that the effects of Ca2+ deprivation on oocyte maturation are mediated through Ca2+-dependent regulation of mRNA translation. CPEB is a central player in regulating translation during oocyte maturation, and recent evidence argues that CPEB phosphorylation is regulated by the Xenopus Rho-family guanine nucleotide exchange factor [83, 84], raising the possibility that some of the Ca2+-dependent effects on mRNA translation could be mediated through this pathway.

Ca2+ Homeostasis

Taken together, our data argue that when Ca2+ homeostasis is disrupted, this breaks down the coordination between the MAPK-MPF signaling cascade and AURKA accumulation, potentially contributing to the spindle formation defects. Ca2+ appears to mediate these effects through regulation of maternal mRNA translation, which is crucial for oocyte maturation. Therefore, Ca2+ homeostasis is a critical determinant of the oocyte's competence to undergo maturation in preparation for fertilization. If Ca2+ homeostasis is disrupted, this signals a compromised oocyte that is unlikely to achieve normal developmental competence. Therefore, normal Ca2+ homeostasis is an important factor in coordinating biochemical and nuclear meiotic events during oocyte maturation.

ACKNOWLEDGMENTS

We are grateful to Howard Schulman for providing the wild-type, dominant-negative, and constitutively active CAMK2A clones, and to Kevin Foskett and John Adelman for the wild-type and mutant CALM clones.

FOOTNOTES

1Supported by National Institutes of Health grants GM-61829 (K.M.), HD35688 (A.M.M.), AG13081 (U.P.), and RR20146 (A.C.). C.P. is funded by grants from the CNRS and LNCC (équipe labellisée). Back

Correspondence: 2Khaled Machaca, Department of Physiology & Biophysics, 4301 West Markham St., Slot 505, University of Arkansas for Medical Sciences (UAMS), Little Rock, AR 72205. FAX: 501 686 8167; e-mail: kamachaca{at}uams.edu.

Received: 26 June 2007.

First decision: 7 August 2007.

Accepted: 30 November 2007.

REFERENCES

  1. The Ovary . : Eppig JJ, Viveiros MM, Marin-Bivens C, De La Fuente R. Regulation of mammalian oocyte maturation. 2004San Diego: Elsevier Press;113–129. In:
  2. ; : Voronina E and Wessel GM. The regulation of oocyte maturation. Curr Top Dev Biol 2003 5853–110[Medline]
  3. ; : Nebreda AR and Ferby I. Regulation of the meiotic cell cycle in oocytes. Curr Opin Cell Biol 2000 12666–675[CrossRef][Medline]
  4. ; : Baitinger C, Alderton J, Poenie M, Schulman H, Steinhardt RA. Multifunctional Ca2+/calmodulin-dependent protein kinase is necessary for nuclear envelope breakdown. J Cell Biol 1990 1111763–1773[Abstract/Free Full Text]
  5. ; : Morin N, Abrieu A, Lorca T, Martin F, Doree M. The proteolysis-dependent metaphase to anaphase transition: calcium/calmodulin-dependent protein kinase II mediates onset of anaphase in extracts prepared from unfertilized Xenopus eggs. EMBO J 1994 134343–4352[Medline]
  6. ; : Poenie M, Alderton J, Tsien RY, Steinhardt RA. Changes of free calcium levels with stages of the cell division cycle. Nature 1985 315147–149[CrossRef][Medline]
  7. ; : Means AR. Calcium, calmodulin and cell cycle regulation. FEBS Lett 1994 3471–4[CrossRef][Medline]
  8. ; : Whitaker M. Regulation of the cell division cycle by inositol trisphosphate and the calcium signaling pathway. Adv Sec Mess Phosph Res 1995 30299–310
  9. ; : Stricker SA. Comparative biology of calcium signaling during fertilization and egg activation in animals. Dev Biol 2000 211157–176[CrossRef]
  10. ; : Duesbery NS and Masui Y. The role of Ca2+ in progesterone-induced germinal vesicle breakdown of Xenopus laevis oocytes: the synergic effects of microtubule depolymerization and Ca2+. Dev Genes Evol 1996 206110–124[CrossRef]
  11. ; : Homa ST, Carroll J, Swann K. The role of calcium in mammalian oocyte maturation and egg activation. Hum Reprod 1993 81274–1281[Abstract/Free Full Text]
  12. ; : Tombes RM, Simerly C, Borisy GG, Schatten G. Meiosis, egg activation, and nuclear envelope breakdown are differentially reliant on Ca2+, whereas germinal vesicle breakdown is Ca2+ independent in the mouse oocyte. J Cell Biol 1992 117799–811[Abstract/Free Full Text]
  13. ; : 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 1975 193369–375[CrossRef][Medline]
  14. ; : Moreau M, Doree M, Guerrier P. Electrophoretic introduction of calcium into the cortex of Xenopus laevis oocytes triggers meiosis reinitiation. J Exp Zool 1976 197443–449[CrossRef][Medline]
  15. ; : Schorderet-Slatkine S, Schorderet M, Baulieu EE. Initiation of meiotic maturation in Xenopus laevis oocytes by lanthanum. Nature 1976 262289–290[CrossRef][Medline]
  16. ; : O'Connor CM, Robinson KR, Smith LD. Calcium, potassium, and sodium exchange by full-grown and maturing Xenopus laevis oocytes. Dev Biol 1977 6128–40[CrossRef][Medline]
  17. ; : Wasserman WJ, Pinto LH, O'Connor CM, Smith LD. Progesterone induces a rapid increase in [Ca2+]in of Xenopus laevis oocytes. Proc Natl Acad Sci U S A 1980 771534–1536[Abstract/Free Full Text]
  18. ; : Moreau M, Vilain JP, Guerrier P. Free calcium changes associated with hormone action in amphibian oocytes. Dev Biol 1980 78201–214[CrossRef][Medline]
  19. ; : Robinson KR. Maturation of Xenopus oocytes is not accompanied by electrode-detectable calcium changes. Dev Biol 1985 109504–508[CrossRef][Medline]
  20. ; : Cork RJ, Cicirelli MF, Robinson KR. A rise in cytosolic calcium is not necessary for maturation of Xenopus laevis oocytes. Dev Biol 1987 12141–47[CrossRef][Medline]
  21. ; : Carroll J and Swann K. Spontaneous cytosolic calcium oscillations driven by inositol trisphosphate occur during in vitro maturation of mouse oocytes. J Biol Chem 1992 26711196–11201[Abstract/Free Full Text]
  22. ; : Sun L and Machaca K. Ca2+cyt negatively regulates the initiation of oocyte maturation. J Cell Biol 2004 16563–75[Abstract/Free Full Text]
  23. ; : Machaca K and Haun S. Induction of maturation-promoting factor during Xenopus oocyte maturation uncouples Ca2+ store depletion from store-operated Ca2+ entry. J Cell Biol 2002 15675–85[Abstract/Free Full Text]
  24. ; : Roghi C, Giet R, Uzbekov R, Morin N, Chartrain I, Le Guellec R, Couturier A, Doree M, Philippe M, Prigent C. The Xenopus protein kinase pEg2 associates with the centrosome in a cell cycle-dependent manner, binds to the spindle microtubules and is involved in bipolar mitotic spindle assembly. J Cell Sci 1998 111557–572[Abstract]
  25. ; : Rassa JC, Wilson GM, Brewer GA, Parks GD. Spacing constraints on reinitiation of paramyxovirus transcription: the gene end U tract acts as a spacer to separate gene end from gene start sites. Virology 2000 274438–449[CrossRef][Medline]
  26. ; : Charlesworth A, Cox LL, MacNicol AM. Cytoplasmic polyadenylation element (CPE)- and CPE-binding protein (CPEB)-independent mechanisms regulate early class maternal mRNA translational activation in Xenopus oocytes. J Biol Chem 2004 27917650–17659[Abstract/Free Full Text]
  27. ; : Hudmon A and Schulman H. Structure-function of the multifunctional Ca2+/calmodulin-dependent protein kinase II. Biochem J 2002 364593–611[CrossRef][Medline]
  28. ; : Mak DO, McBride SM, Petrenko NB, Foskett JK. Novel regulation of calcium inhibition of the inositol 1,4,5-trisphosphate receptor calcium-release channel. J Gen Physiol 2003 122569–581[Abstract/Free Full Text]
  29. ; : Kessler BM, Tortorella D, Altun M, Kisselev AF, Fiebiger E, Hekking BG, Ploegh HL, Overkleeft HS. Extended peptide-based inhibitors efficiently target the proteasome and reveal overlapping specificities of the catalytic beta-subunits. Chem Biol 2001 8913–929[CrossRef][Medline]
  30. ; : Zhu Y, Bond J, Thomas P. Identification, classification, and partial characterization of genes in humans and other vertebrates homologous to a fish membrane progestin receptor. Proc Natl Acad Sci U S A 2003 1002237–2242[Abstract/Free Full Text]
  31. ; : Sadler SE and Maller JL. Identification of a steroid receptor on the surface of Xenopus oocytes by photoaffinity labeling. J Biol Chem 1982 257355–361[Abstract/Free Full Text]
  32. ; : Maller JL, Butcher FR, Krebs EG. Early effect of progesterone on levels of cyclic adenosine 3':5'-monophosphate in Xenopus oocytes. J Biol Chem 1979 254579–582[Abstract/Free Full Text]
  33. ; : Sagata N, Oskarsson M, Copeland T, Brumbaugh J, Vande Woude GF. Function of the c-mos proto-oncogene product in meiotic maturation in Xenopus oocytes. Nature 1988 335519–525[CrossRef][Medline]
  34. ; : Barkoff A, Ballantyne S, Wickens M. Meiotic maturation in Xenopus requires polyadenylation of multiple mRNAs. EMBO J 1998 173168–3175[CrossRef][Medline]
  35. ; : Coleman TR and Dunphy WG. Cdc2 regulatory factors. Curr Opin Cell Biol 1994 6877–882[CrossRef][Medline]
  36. ; : Mendez R and Richter JD. Translational control by CPEB: a means to the end. Nat Rev Mol Cell Biol 2001 2521–529[CrossRef][Medline]
  37. ; : Charlesworth A, Wilczynska A, Thampi P, Cox LL, MacNicol AM. Musashi regulates the temporal order of mRNA translation during Xenopus oocyte maturation. EMBO J 2006 252792–2801[CrossRef][Medline]
  38. ; : Johnson J, Bierle BM, Gallicano GI, Capco DG. Calcium/calmodulin-dependent protein kinase II and calmodulin: regulators of the meiotic spindle in mouse eggs. Dev Biol 1998 204464–477[CrossRef][Medline]
  39. ; : Madgwick S, Levasseur M, Jones KT. Calmodulin-dependent protein kinase II, and not protein kinase C, is sufficient for triggering cell-cycle resumption in mammalian eggs. J Cell Sci 2005 1183849–3859[Abstract/Free Full Text]
  40. ; : Su YQ and Eppig JJ. Evidence that multifunctional calcium/calmodulin-dependent protein kinase II (CaM KII) participates in the meiotic maturation of mouse oocytes. Mol Reprod Dev 2002 61560–569[CrossRef][Medline]
  41. ; : Ishida A, Kameshita I, Okuno S, Kitani T, Fujisawa H. A novel highly specific and potent inhibitor of calmodulin-dependent protein kinase II. Biochem Biophys Res Commun 1995 212806–812[CrossRef][Medline]
  42. ; : Waldmann R, Hanson PI, Schulman H. Multifunctional Ca2+/calmodulin-dependent protein kinase made Ca2+ independent for functional studies. Biochem 1990 291679–1684[CrossRef][Medline]
  43. ; : Chin D and Means AR. Calmodulin: a prototypical calcium sensor. Trends Cell Biol 2000 10322–328[CrossRef][Medline]
  44. ; : Welsh MJ, Dedman JR, Brinkley BR, Means AR. Calcium-dependent regulator protein: localization in mitotic apparatus of eukaryotic cells. Proc Natl Acad Sci U S A 1978 751867–1871[Abstract/Free Full Text]
  45. ; : Hisanaga S and Pratt MM. Calmodulin interaction with cytoplasmic and flagellar dynein: calcium-dependent binding and stimulation of adenosinetriphosphatase activity. Biochem 1984 233032–3037[CrossRef][Medline]
  46. ; : Blum JJ, Hayes A, Jamieson GA Jr, Vanaman TC. Calmodulin confers calcium sensitivity on ciliary dynein ATPase. J Cell Biol 1980 87386–397[Abstract/Free Full Text]
  47. ; : Pfarr CM, Coue M, Grissom PM, Hays TS, Porter ME, McIntosh JR. Cytoplasmic dynein is localized to kinetochores during mitosis. Nature 1990 345263–265[CrossRef][Medline]
  48. ; : Steuer ER, Wordeman L, Schroer TA, Sheetz MP. Localization of cytoplasmic dynein to mitotic spindles and kinetochores. Nature 1990 345266–268[CrossRef][Medline]
  49. ; : Verde F, Berrez JM, Antony C, Karsenti E. Taxol-induced microtubule asters in mitotic extracts of Xenopus eggs: requirement for phosphorylated factors and cytoplasmic dynein. J Cell Biol 1991 1121177–1187[Abstract/Free Full Text]
  50. ; : Becker BE, Romney SJ, Gard DL. XMAP215, XKCM1, NuMA, and cytoplasmic dynein are required for the assembly and organization of the transient microtubule array during the maturation of Xenopus oocytes. Dev Biol 2003 261488–505[CrossRef][Medline]
  51. ; : Geiser JR, Sundberg HA, Chang BH, Muller EG, Davis TN. The essential mitotic target of calmodulin is the 110-kilodalton component of the spindle pole body in Saccharomyces cerevisiae. Mol Cell Biol 1993 137913–7924[Abstract/Free Full Text]
  52. ; : Stirling DA, Rayner TF, Prescott AR, Stark MJ. Mutations which block the binding of calmodulin to Spc110p cause multiple mitotic defects. J Cell Sci 1996 1091297–1310[Abstract]
  53. ; : Sundberg HA, Goetsch L, Byers B, Davis TN. Role of calmodulin and Spc110p interaction in the proper assembly of spindle pole body compenents. J Cell Biol 1996 133111–124[Abstract/Free Full Text]
  54. ; : Cartaud A, Ozon R, Walsh MP, Haiech J, Demaille JG. Xenopus laevis oocyte calmodulin in the process of meiotic maturation. J Biol Chem 1980 2559404–9408[Abstract/Free Full Text]
  55. ; : Cicirelli MF and Smith LD. Do calcium and calmodulin trigger maturation in amphibian oocytes? Dev Biol 1987 12148–57[CrossRef][Medline]
  56. ; : Wasserman WJ and Smith LD. Calmodulin triggers the resumption of meiosis in amphibian oocytes. J Cell Biol 1981 89389–394[Abstract/Free Full Text]
  57. ; : Huchon D, Ozon R, Fischer EH, Demaille JG. The pure inhibitor of cAMP-dependent protein kinase initiates Xenopus laevis meiotic maturation. A 4-step scheme for meiotic maturation. Mol Cell Endocrinol 1981 22211–222[CrossRef][Medline]
  58. ; : el Saleh SC and Solaro RJ. Calmidazolium, a calmodulin antagonist, stimulates calcium-troponin C and calcium-calmodulin-dependent activation of striated muscle myofilaments. J Biol Chem 1987 26217240–17246[Abstract/Free Full Text]
  59. ; : Weiss B and Levin RM. Mechanism for selectively inhibiting the activation of cyclic nucleotide phosphodiesterase and adenylate cyclase by antipsychotic agents. Adv Cyclic Nucleotide Res 1978 9285–303[Medline]
  60. ; : Cartaud A, Huchon D, Marot J, Ozon R, Demaille JG. Calmodulin is involved in the first step of oocyte maturation: effects of the antipsychotic drug fluphenazine and of anticalmodulin antibodies on the progesterone-induced maturation of xenopus laevis oocyte. Cell Differ 1981 10357–363[CrossRef][Medline]
  61. ; : Baulieu EE, Godeau F, Schorderet M, Schorderet-Slatkine S. Steroid-induced meiotic division in Xenopus laevis oocytes: surface and calcium. Nature 1978 275593–598[CrossRef][Medline]
  62. ; : Hollinger TG and Alvarez IM. The effect of trifluoperazine on maturation of Xenopus laevis oocytes. J Exp Zool 1984 230427–441[CrossRef][Medline]
  63. ; : Zuhlke RD, Pitt GS, Deisseroth K, Tsien RW, Reuter H. Calmodulin supports both inactivation and facilitation of L-type calcium channels. Nature 1999 399159–162[CrossRef][Medline]
  64. ; : Xia XM, Fakler B, Rivard A, Wayman G, Johnson-Pais T, Keen JE, Ishii T, Hirschberg B, Bond CT, Lutsenko S, Maylie J, Adelman JP. Mechanism of calcium gating in small-conductance calcium-activated potassium channels. Nature 1998 395503–507[CrossRef][Medline]
  65. ; : Gross SD, Schwab MS, Taieb FE, Lewellyn AL, Qian YW, Maller JL. The critical role of the MAP kinase pathway in meiosis II in Xenopus oocytes is mediated by p90(Rsk). Curr Biol 2000 10430–438[CrossRef][Medline]
  66. ; : Nakajo N, Yoshitome S, Iwashita J, Iida M, Uto K, Ueno S, Okamoto K, Sagata N. Absence of Wee1 ensures the meiotic cell cycle in Xenopus oocytes. Genes Dev 2000 14328–338[Abstract/Free Full Text]
  67. ; : Kanki JP and Donoghue DJ. Progression from meiosis I to meiosis II in Xenopus oocytes requires de novo translation of the mosxe protooncogene. Proc Natl Acad Sci U S A 1991 885794–5798[Abstract/Free Full Text]
  68. ; : Schumacher JM, Golden A, Donovan PJ. AIR-2: An Aurora/Ipl1-related protein kinase associated with chromosomes and midbody microtubules is required for polar body extrusion and cytokinesis in Caenorhabditis elegans embryos. J Cell Biol 1998 1431635–1646[Abstract/Free Full Text]
  69. ; : Castro A, Mandart E, Lorca T, Galas S. Involvement of Aurora A kinase during meiosis I-II transition in Xenopus oocytes. J Biol Chem 2003 2782236–2241[Abstract/Free Full Text]
  70. ; : Ma C, Cummings C, Liu XJ. Biphasic activation of Aurora-A kinase during the meiosis I- meiosis II transition in Xenopus oocytes. Mol Cell Biol 2003 231703–1716[Abstract/Free Full Text]
  71. ; : Frank-Vaillant M, Haccard O, Thibier C, Ozon R, Arlot-Bonnemains Y, Prigent C, Jessus C. Progesterone regulates the accumulation and the activation of Eg2 kinase in Xenopus oocytes. J Cell Sci 2000 1131127–1138[Abstract]
  72. ; : Paris J, Swenson K, Piwnica-Worms H, Richter JD. Maturation-specific polyadenylation: in vitro activation by p34cdc2 and phosphorylation of a 58-kD CPE-binding protein. Genes Dev 1991 51697–1708[Abstract/Free Full Text]
  73. ; : Kawahara H and Yokosawa H. Intracellular calcium mobilization regulates the activity of 26 S proteasome during the metaphase-anaphase transition in the ascidian meiotic cell cycle. Dev Biol 1994 166623–633[CrossRef][Medline]
  74. ; : Aizawa H, Kawahara H, Tanaka K, Yokosawa H. Activation of the proteasome during Xenopus egg activation implies a link between proteasome activation and intracellular calcium release. Biochem Biophys Res Commun 1996 218224–228[CrossRef][Medline]
  75. ; : Josefsberg LB, Galiani D, Dantes A, Amsterdam A, Dekel N. The proteasome is involved in the first metaphase-to-anaphase transition of meiosis in rat oocytes. Biol Reprod 2000 621270–1277[Abstract/Free Full Text]
  76. ; : Reverte CG, Ahearn MD, Hake LE. CPEB degradation during Xenopus oocyte maturation requires a PEST domain and the 26S proteasome. Dev Biol 2001 231447–458[CrossRef][Medline]
  77. ; : Ishida N, Tanaka K, Tamura T, Nishizawa M, Okazaki K, Sagata N, Ichihara A. Mos is degraded by the 26S proteasome in a ubiquitin-dependent fashion. FEBS Lett 1993 324345–348[CrossRef][Medline]
  78. ; : Weisenberg RC. Microtubule formation in vitro in solutions containing low calcium concentrations. Science 1972 1771104–1105[Abstract/Free Full Text]
  79. ; : Kirschner MW, Williams RC, Weingarten M, Gerhart JC. Microtubules from mammalian brain: some properties of their depolymerization products and a proposed mechanism of assembly and disassembly. Proc Natl Acad Sci U S A 1974 711159–1163[Abstract/Free Full Text]
  80. ; : Eyers PA and Maller JL. Regulating the regulators: Aurora A activation and mitosis. Cell Cycle 2003 2287–289[Medline]
  81. ; : Nigg EA. Mitotic kinases as regulators of cell division and its checkpoints. Nat Rev Mol Cell Biol 2001 221–32[CrossRef][Medline]
  82. ; : Gruss OJ and Vernos I. The mechanism of spindle assembly: functions of Ran and its target TPX2. J Cell Biol 2004 166949–955[Abstract/Free Full Text]
  83. ; : Martinez SE, Yuan L, Lacza C, Ransom H, Mahon GM, Whitehead IP, Hake LE. XGef mediates early CPEB phosphorylation during Xenopus oocyte meiotic maturation. Mol Biol Cell 2005 161152–1164[Abstract/Free Full Text]
  84. ; : Keady BT, Kuo P, Martinez SE, Yuan L, Hake LE. MAPK interacts with XGef and is required for CPEB activation during meiosis in Xenopus oocytes. J Cell Sci 2007 1201093–1103[Abstract/Free Full Text]




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