ca2+ homeostasis regulates xenopus oocyte maturation

Ca2+ Homeostasis Regulates Xenopus Oocyte Maturation1

Lu Sun3, Rawad Hodeify3, Shirley Haun3, Amanda Charlesworth4, Angus M. MacNicol3,4,7,Subramaniam Ponnappan5, Usha Ponnappan5,6,7, Claude Prigent8, and KhaledMachaca2,3,7

3Department of Physiology & Biophysics, University of Arkansas for Medical Sciences, Little Rock, Arkansas72205

4Department of Neurobiology & Developmental Sciences, University of Arkansas for Medical Sciences, LittleRock, Arkansas 72205

5Department of Geriatrics, University of Arkansas for Medical Sciences, Little Rock, Arkansas 72205

6Department of Immunology & Microbiology, University of Arkansas for Medical Sciences, Little Rock,Arkansas 72205

7Arkansas Cancer Research Center, University of Arkansas for Medical Sciences, Little Rock, Arkansas72205

8CNRS UMR6061, Genetique et Developpement, Universite de Rennes 1, 35043 Rennes, France

AbstractIn contrast to the well-defined role of Ca2+ signals during mitosis, the contribution of Ca2+ signalingto meiosis progression is controversial, despite several decades of investigating the role of Ca2+ andits effectors in vertebrate oocyte maturation. We have previously shown that during Xenopus oocytematuration, 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 firstpolar body. In this study, we test the contribution of several downstream effectors in mediating theCa2+ effects during oocyte maturation. We show that calmodulin and calcium-calmodulin-dependentprotein kinase II (CAMK2) are not critical downstream Ca2+ effectors during meiotic maturation. Incontrast, 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 maycontribute to the defective spindle phenotype following Ca2+ deprivation. These findings argue thatCa2+ homeostasis is important in establishing the oocyte’s competence to undergo maturation inpreparation for fertilization and embryonic development.

KeywordsAurora kinase A; calcium; gamete biology; meiosis; oocyte development

INTRODUCTIONIntracellular Ca2+ signals are ubiquitous and regulate a plethora of cell physiological anddevelopmental processes, including contraction, secretion, transcription, cell proliferation, and

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).2Correspondence: Khaled Machaca, Department of Physiology & Biophysics, 4301 West Markham St., Slot 505, University of Arkansasfor Medical Sciences (UAMS), Little Rock, AR 72205. FAX: 501 686 8167; e-mail: [email protected].

NIH Public AccessAuthor ManuscriptBiol Reprod. Author manuscript; available in PMC 2008 November 25.

Published in final edited form as:Biol Reprod. 2008 April ; 78(4): 726–735. doi:10.1095/biolreprod.107.063693.

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cell death. The ultimate cellular response to environmental and intrinsic cues integrates amultitude of signaling cascades. Therefore, cellular and developmental decisions are theoutcome of often competing signaling processes that include significant crosstalk and feedbackmechanisms, allowing tighter regulation of critical differentiation pathways. A fundamentaldevelopmental decision that initiates multicellular organism development is oocyte maturation.Oocyte maturation is a complex cellular differentiation that prepares the egg for fertilizationand early embryonic development [1]. In vertebrates, oocyte maturation includes transitionthrough meiosis and arrest at metaphase II until fertilization. This nuclear maturation is coupledto cytoplasmic maturation, including dramatic morphological and mechanistic alterations thatare essential for the egg-to-embryo transition [1,2]. As might be expected for such a complexcellular differentiation, several signaling cascades regulate oocyte maturation, culminating inthe activation of maturation-promoting factor (MPF) [1,3].

The role of Ca2+ signaling in regulating oocytematuration has been of wide interest, given thatCa2+ signals have been convincingly shown to play a role in mitosis and during egg activationat 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 evidencesupports a role for Ca2+ and its downstream effectors calmodulin and CAMK2 in mitosis andfollowing fertilization [7–9]. In contrast, the role of Ca2+ signals during vertebrate oocytemeiosis remains contentious [10–12]. In Xenopus, early reports argued that a Ca2+ rise issufficient 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 oocytematuration is also controversial [16–20]. These conflicting reports argue that in Xenopus therelationship between Ca2+ and oocyte maturation is complex. Studies regarding the role ofCa2+ during mammalian oocyte maturation also produce conflicting results [11,21].

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

MATERIALS AND METHODSGamete Preparation and Treatments

Xenopus oocytes were obtained as previously described in Machaca and Haun [23]. Frogs werepurchased 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 spotat the animal pole of the cell and was confirmed by bisecting the cell into halves and detectingthe 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 MaxChelatorprogram (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 UAMSAnimal Care and Use Committee (IACUC).

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

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http://www.stanford.edu/~cpatton/maxc.html

Imaging Spindle StructureCells were fixed 3 h after GVBD in 100% methanol and stored at −20°C overnight. Afterrehydration in TBS/methanol (1:1) for 20 min, oocytes were washed twice with TBS for 15min each and blocked for 3 h in TBS containing 2% BSA. Oocytes then were immunolabeledwith an anti-α-tubulin monoclonal antibody (DM1A; Sigma) in TBS containing 2% BSA,followed by a Cy2-conjugated donkey anti-mouse secondary (Jackson) for 24 h each. Theoocytes 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 structureimages were collected on a Zeiss LSM510 confocal or on an Olympus Fluoview confocal. Insome experiments, chromosome structure and polar body emission were visualized using SytoxOrange labeling without staining the spindle.

Polyadenylation AssayPolyadenylation 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 reactionusing T4 RNA ligase (New England Biolabs) according to manufacturer’s directions. Thewhole 10-µl RNA ligation reaction was used in a 50-µl reverse transcription reaction usingSuperscript III (Invitrogen), according to manufacturer’s directions using 0.4 µg P1′ as thereverse primer [25]. A total of 1 µl of this cDNA preparation was used in each 50-µl PCRreaction using Platinum Taq (Invitrogen), according to manufacturer’s directions. Polymerasechain reaction was performed for 40 cycles using a 56°C annealing temperature and 1.5 mMfinal concentration of Mg2+. For optimal resolution, specific primers used were designed to be70–90 nucleotides from the poly(A) addition site [26] according to sequence in GenBank: theaccession numbers are Z17206 (Eg2, AURKA), X13311 (MOS), and J03166 (cyclin B1). Theprimers used were Cyclin B1: GTG GCA TTC CAA TTG TGT ATT GTT; AURKA: GTTTCA ATC TTG TAT GTC CTT TTA; and MOS: GTT GCA TTG CTG TTT AAG TGG TAA.

PlasmidsWild-type (wt) and mutant CAMK2A clones were obtained from H. Schulman [27]. They weresubcloned 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 wereprovided by the Adelman Lab. Both wt and mutant calmodulin (with a D-to-A mutation in allfour EF hand motifs) were cloned into pBF, as previously described [28]. RNA fromcalmodulin (CALM) clones was generated by linearizing them with MluI and transcribing withSP6 polymerase.

Proteasome AssayProteasome-specific activity was evaluated in oocyte lysates employing a biotinylatedproteasome 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 mMsucrose, and 2 mM ATP. Lysates equalized for protein (30 µg) were labeled with 5 µMbiotinylated probe in a reaction buffer containing 50 mM Tris-HCl (pH 7.4), 5 mM MgCl2, 2mM ATP, and 2 mM dithiothreitol for 2 h at 37°C. Labeling reactions were quenched by theaddition of SDS sample buffer and boiling for 5 min. Samples were resolved by 12% SDS-PAGE, transferred to nitrocellulose membranes, immunoblotted with streptavidin conjugatedto horseradish peroxidase and detected using Enhanced Chemiluminescence (AmershamBiosciences). For the studies using proteasomal inhibitors, the inhibitors were present only

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during the maturation phase of the oocytes and were not added to the proteasome activity assayafter oocyte lysis.

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

Progesterone-dependent Xenopus oocyte maturation is believed to be initiated by a cell surfaceG 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 throughlargely unknown steps, mRNA translation of MOS and other regulators is induced throughpolyadenylation [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 andmeiosis, and it consists of a catalytic p34cdc2 Ser/Thr kinase subunit (CDK1) and a regulatorycyclin B subunit [35].

One of the earliest known molecular events downstream of PKA inhibition is translationalregulation of maternal mRNAs through polyadenylation. Maternal mRNA polyadenylation isregulated in a temporal fashion through distinct 3′ UTR regulatory elements [26,36]. Someearly-class mRNAs, such as MOS, are polyadenylated and translated prior to GVBD under thecontrol 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 cytoplasmicpolyadenylation elements [26,37]. We were interested in testing the effect of Ca2+ deprivationon the timing of polyadenylation of early- and late-class mRNAs. As shown in Figure 1B, inCa2+-deprived oocytes, polyadenylation of MOS mRNA occurs as early as 2–3 h afterprogesterone addition (observed as an increase in PCR product size), compared with 4–5 h forcontrol oocytes matured in normal Ca2+ medium. In contrast, the pattern of the late cyclin B1mRNA 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 undergoGVBD, and G50 refers to when 50% of the cells in the population reach GVBD. These datashow that the premature activation of the MOS-MAPK signaling cascade in Ca2+-deprivedcells 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 differentdonor females. There typically are significant variations in the rate of oocyte maturation in thepopulation between different females, as illustrated in Figure 1, A and B. Oocytes in Figure1A matured at a faster rate than those from the female in Figure 1B. Nonetheless, despite thesedifferences in the rate of maturation, qualitatively the differences described in this manuscriptin terms of Ca2+ deprivation and polyadenylation are consistently observed.

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

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Careful analyses of spindle structure in Ca2+-deprived cells reveal that the primary defect isthe 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 BAPTAinjection or thapsigargin treatment (not shown) inhibits spindle elongation. Ca2+-deprived cellsassemble a compact spindle around the chromosomes but never progress to the elongatedbipolar spindle observed in control cells (Fig. 1D). The inability of the spindle to elongateexplains 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) inthe defects observed in Ca2+-deprived oocytes during maturation. This is because CAMK2 isa prominent downstream effector of Ca2+ signals and because it has been functionallyimplicated in meiosis. For example, CAMK2 has been shown to physically associate with themeiotic spindle in mouse eggs and to be important for the transition into anaphase duringmeiosis II [38,39]. Furthermore, inhibition of CAMK2 activity in mouse oocytes blocks polarbody emission [40], arguing that CAMK2 inhibition may be responsible for the defectsobserved following Ca2+ deprivation during Xenopus oocyte maturation. To test whether thisis the case, we modulated CAMK2 activity by injecting oocytes with AIP, a potent and specificCAMK2 inhibitory peptide [41]. AIP injection (10 µM) did not affect the extent of GVBD inthe 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 ineffecting Ca2+ signals during oocyte maturation, it did not reach the levels of enhancementachieved 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 withThaps (Fig. 2A, T-AIP).

We wanted to confirm the AIP result by expressing wt, dominant-negative (CAMK2dn), andconstitutively active (CAMK2ca) CAMK2A mutants (Fig. 2B) [42]. Wild-type CAMK2Ainjection did not significantly (P = 0.083) affect the rate of maturation compared with thecontrol (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 withwt CAMK2A injection (Fig. 2B). None of the CAMK2A injections significantly affected theextent of maturation (Fig. 2B). These results argue that CAMK2 is not a downstream effectorof 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 shouldenhance the rate of maturation in a fashion similar to Ca2+ deprivation. Given the fact thatCAMK2 could impact a multitude of signaling cascades, the observed effects of overexpressionof 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 therate 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, AIPinjection or injection of wt or dn CAMK2A did not inhibit the ability of oocytes to extrude apolar body (Fig. 2C). It was not possible to test the effect of the constitutively activeCAMK2A mutant in these experiments, because cells expressing CAMK2ca did not survive longenough after GVBD to allow for first polar body extrusion. These results indicate that CAMK2is not important for spindle formation in Xenopus oocytes.

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Role of Calmodulin (CALM)Calmodulin (CALM) is an excellent candidate for transmitting Ca2+ signals to downstreameffectors 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 cellularfunctions [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 microtubuledynamics [49], and is critical for the formation of a normal spindle in Xenopus [49,50]. Inaddition to its interactions with dynein, CALM interacts with other components at the spindlepole, primarily the 110-kDa spindle pole component, and this interaction is essential for normalspindle formation [51–53]. It is therefore possible that the spindle defect observed in meiosisI of Ca2+-deprived cells is due to disruption of CALM function/localization. In addition, CALMhas been postulated to play a role in the progression of oocyte maturation in Xenopus, sinceCALM levels increase during maturation [54], and CALM injection has been reported to inducelow 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 inhibitedCALM function with calmidazolium (CDZ) or fluphenazine (FLZ). Calmidazolium is animidazole compound that acts as a specific CALM antagonist by strongly binding to thehydrophobic surface of CALM, preventing the binding of Ca2+ and, thus, the correct foldingof active CALM [58]. Fluphenazine is an antipsychotic drug that blocks CALM interactionwith a variety of downstream effectors [59]. Calmidazolium did not affect the rate or extent ofentry 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), althoughthese effects were not nearly as dramatic as maturing oocytes in the absence of extracellularCa2+. 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+ deprivationwithout affecting maximal GVBD (Fig. 3B). Furthermore, FLZ was more likely to lead tospindle defects, although the effects were quite variable (Fig. 3B) yet, surprisingly, this did notaffect the ability of the oocytes to extrude a polar body.

The differential effect of FLZ and CDZ on the rate of maturation is not surprising, since CDZhas been previously reported not to effectively inhibit certain CALM-dependent processeswhere 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 arguethat CALM is involved in regulating the rate of oocyte maturation. However, attempts toconfirm the effects of a pharmacological block of CALM on the rate of maturation using eitherinjection of a CALM inhibitory peptide (MLCK peptide) or a CALM mutant with all four EFhands 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, bringinginto question the importance of CALM in regulating the rate of maturation. Therefore, as isthe case with CAMK2, the fact that different approaches to interfere with CALM functionproduce disparate results in terms of controlling the rate of oocyte maturation argues thatCALM is not a critical determinant of the processes by which Ca2+ regulates progressionthrough maturation.

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

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Aurora Kinase A (AURKA)The inability of Ca2+-deprived oocytes to complete meiosis normally and extrude a polar bodyis interesting because this nuclear defect occurs while the kinetics of the MAPK-MPF cascadeproceed normally [22]. These observations suggest that Ca2+ deprivation uncouples the kinasesignaling cascade regulating oocyte maturation from the nuclear meiotic events. Several otherexperimental manipulations, such as inhibition of the MAPK cascade [65], downregulation ofMPF [66], and inhibition of protein synthesis [67], lead to a meiosis I arrest that is coupled toan interphaselike state. In contrast, Ca2+ deprivation does not affect the MAPK-MPF kinasesignaling cascade, yet it disrupts spindle elongation in meiosis I and prevents polar bodyemission. Interestingly, manipulation of the Aurora kinases produces a similar phenotype.Downregulation of Aurora B in Caenorhabditis elegans inhibits polar body extrusion inmeiosis I [68]. In Xenopus oocytes, injection of anti-AURKA antibodies inhibits polar bodyextrusion 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 thatAURKA regulation may have a Ca2+-dependent component. To test this possibility, wemeasured the levels of AURKA in control and Ca2+-deprived oocytes throughout thematuration process (Fig. 4A). As previously shown [71], AURKA gradually accumulatesduring maturation, especially after GVBD (Fig. 4A). In contrast, AURKA accumulation issignificantly attenuated during maturation of Ca2+-deprived, Thaps-treated oocytes (Fig. 4A).Although the tubulin loading control shows more protein loading toward the later time pointsin both the low Ca2+ and Thaps treatments (Fig. 4A), this still correlates with loweraccumulated levels of AURKA protein, showing that the ability of cells deprived of Ca2+

signals to accumulate AURKA is compromised.

Supporting the time course data, when lysates are prepared at different time points duringmaturation, 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 haveundergone 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 bothcells with a white spot (w) and those without (nw) were collected—and (5) when 100% of thecells 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 initiationof endogenous AURKA mRNA polyadenylation (Fig. 4C). This argues that the lack of AURKAaccumulation following disruption of Ca2+ homeostasis is not due to differentialpolyadenylation, but rather either reflects a specific defect in AURKA mRNA translation orposttranslational stabilization of AURKA protein.

Proteasome ModulationOne possibility to explain the AURKA results is modulation of proteasome activity followingCa2+ deprivation. The 26S proteasome regulates protein degradation followingpolyubiquitination and has been implicated in oocyte meiosis in several species. Proteasomeactivity 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 proteasomeinhibitors block polar body extrusion [75]. Furthermore, the levels of certain critical regulatorsof 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

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regulating oocyte maturation following Ca2+ deprivation. Proteasome activity was not affectedwhen 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, asexplained in Materials and Methods (Fig. 5A). We further tested the ability of two proteasomeinhibitors, lactacystin and MG132, to block Xenopus oocyte proteasome activity. Lactacystin(at the doses employed) was ineffective at blocking the proteasome, whereas MG132 inhibitedproteasome activity in a dose-dependent fashion (Fig. 5A). Nevertheless, oocytes treated witheither inhibitor matured with kinetics similar to those in control cells (Fig. 5B), showing thatproteasome inhibition is not sufficient to enhance the rate of oocyte maturation. These dataargue that modulation of proteasome function is not responsible for the defective AURKAaccumulation, 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+ deprivationon proteasome activity.

DISCUSSIONCalmodulin and CAMK2

Depriving Xenopus oocytes of Ca2+ signals does not affect their ability to enter meiosis orundergo GVBD, but it results in a dramatically disrupted spindle structure and a lack of polarbody emission [22]. In Ca2+-deprived oocytes, the meiosis I spindle does not elongate into abipolar spindle (Fig. 1D), which could form the basis for the lack of polar body emission. It isunclear why disrupting Ca2+ homeostasis leads to defective spindles. One possibility is thatCa2+ chelation directly affects microtubule assembly, as previously described [78,79]. Anotherexplanation for the Ca2+-dependent meiosis I defect would be faulty signaling directlydownstream of Ca2+. This was the incentive for us to test two ubiquitous Ca2+-dependentdownstream effectors, calmodulin and CAMK2. However, our results argue that neither ofthese effectors is involved in the defects observed following Ca2+ deprivation during oocytematuration. Interfering with CALM or CAMK2 function using various approaches producesdisparate effects that are inconsistent with the effect of Ca2+ deprivation on the rate ofprogression of oocyte maturation (Fig. 2 and Fig. 3). This argues that CALM-CAMK2 is nota critical effector of Ca2+ signals during oocyte maturation in terms of setting the rate ofmaturation. In a similar fashion, disruption of either CALM or CAMK2 function did notsignificantly 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 ACa2+ deprivation results in a faster maturation rate, raising the possibility that this couldunderlie the meiosis I defect. In that context, tight coordination of different signaling cascadesis essential for timely progression through oocyte maturation [3]. We were therefore interestedin testing whether depriving oocytes of Ca2+ signals somehow disrupts this tight coordination.AURKA was an obvious target to test, because altering AURKA activity phenocopies theCa2+ deprivation-dependent spindle defect [69,70]. Furthermore, the Aurora family encodesSer/Thr kinases that localize to the spindle poles and are crucial for spindle assembly duringboth meiosis and mitosis [80–82].

AURKA protein gradually accumulates during oocyte maturation (Fig. 4A), and this increaseis important for AURKA function during meiosis [71]. Although the ability of AURKA toaccumulate in Ca2+-deprived oocytes is compromised (Fig. 4, A and B), this defect is not

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correlated with a failure to polyadenylate AURKA mRNA prior to GVBD (Fig. 4C). Thesefindings indicate that unlike the translation of MOS and activation of MAP kinase signaling,AURKA mRNA translation is specifically attenuated, or the newly synthesized AURKA proteinis destabilized. The failure to accumulate AURKA protein could contribute to the spindleelongation defect and the inability of Ca2+-deprived oocytes to extrude a polar body, becausethe 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 ofAURKA accumulation and the defective spindle phenotype observed in Ca2+-deprived cells.Unfortunately, directly testing this relationship is technically difficult because the levels ofAURKA have to be tightly controlled. Either interfering with AURKA function or increasingit 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 cascadeearlier [22]. This is due to a more rapid polyadenylation of early-class mRNAs, such asMOS (Fig. 1B). AURKA mRNA is also polyadenylated earlier in Ca2+-deprived cells, but thisis associated with defective accumulation of AURKA. The fact that AURKA accumulation isnot necessarily dependent on polyadenylation [71] may partially explain the differential effectof Ca2+ deprivation on the MOS-MAPK cascade versus AURKA. Accumulation of AURKAis a balance between both mRNA translation and AURKA protein degradation rates. Sinceproteasome activity is not directly affected in Ca2+-deprived oocytes, Ca2+ deprivation couldbe inhibiting AURKA accumulation by affecting AURKA mRNA de-repression independentlyof polyadenylation and/or AURKA protein stability.

Together, our data argue that the effects of Ca2+ deprivation on oocyte maturation are mediatedthrough Ca2+-dependent regulation of mRNA translation. CPEB is a central player inregulating translation during oocyte maturation, and recent evidence argues that CPEBphosphorylation 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 translationcould be mediated through this pathway.

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

ACKNOWLEDGMENTSWe are grateful to Howard Schulman for providing the wild-type, dominant-negative, and constitutively activeCAMK2A clones, and to Kevin Foskett and John Adelman for the wild-type and mutant CALM clones.

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FIG. 1.Ca2+ deprivation disrupts meiotic maturation. A) Representative GVBD time course foroocytes incubated in normal Ca2+ (0.6 mM, Con) or low-Ca2+ media (50 µM, L-Ca), or treatedwith 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 firstGVBD (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 isindicated above each panel. D) Oocytes were fixed at the indicated times after GVBD andstained for tubulin and DNA to visualize spindle structure. Bar = 10 µm.

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FIG. 2.Effects of CAMK2 modulation on meiosis. A) Rates of oocyte maturation (top panel) andmaximal GVBD levels reached (bottom panel). The rate of oocyte maturation was quantifiedas the time required for 50% of the oocytes in the population to undergo GVBD (G50). Rateswere normalized to the time to G50 in normal Ca2+ medium (Con). AIP, a CAMK2 inhibitorypeptide, was injected at 10 µM per oocyte 1 h before progesterone addition. Some oocyteswere 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 bargraph 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 differentCAMK2A 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 ngper oocyte), and constitutively active CAMK2A (C-ca, 1 ng per oocyte). The times to G50 andmaximal GVBD in the population are shown (n = 4–6 experiments). The letters above the bargraphs 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

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was significantly different (P ≤ 0.034). C) Percentage of polar body (PB) emission in cellsmatured under control conditions (Con), low-Ca2+ medium (L-Ca), injected with AIP (10 or20 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. Meioticmaturation 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 differentconstructs. Lysates were from uninjected cells (Uninj.) or cells injected with differentCAMK2A mutants as indicated.

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FIG. 3.Effect of Calmodulin inhibition on oocyte maturation. A) Oocytes were incubated in L-15media containing dimethylsulfoxide (Con) or 1 µM calmidozolium (CDZ) for 3 h beforeprogesterone addition. The upper panels in A show a representative GVBD time course andaverage data for the maximal GVBD reached and time to GVBD50 (n = 8); the lower panelsshow representative images of spindle structure at 3 h after GVBD. Percentages of cells withnormal spindle structure and normalized polar body emission levels are shown (n = 4). Polarbody 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 hbefore progesterone addition. A representative GVBD time course and averaged data formaximal GVBD and rate to GVBD50 are shown in B, upper panels (n = 4). Fluphenazinesignificantly (P = 10−5) enhanced the time to G50. The lower panels in B show spindle structure

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at 3 h after GVBD and the percentage of cells with normal spindles and normalized polar bodyemission levels (n = 4). Summary data are presented as the mean ± SEM. Bars = 10 µm.

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FIG. 4.Kinetics of AURKA accumulation. A) Oocytes were incubated in normal calcium L-15medium (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 forAURKA levels using an anti-Eg2 monoclonal antibody (1C1). The time course is divided intotwo phases: hours after progesterone addition, and hours after GVBD. The relative intensityof 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 threesimilar experiments. B) Oocytes were treated as described in A. Cell lysates were collectedfrom 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 thathad undergone GVBD (white spot, w) and those that had not (no white spot, nw). Tubulin wasused 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.

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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 treatedwith the indicated concentrations of lactacystin (Lact) or MG-132 (MG132) for 1 h beforeprogesterone stimulation. The proteasome activity assay was performed as described inMaterials and Methods. Molecular weight markers are indicated to the left of the gel (Mr = ×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 quantifiedas the time required for 50% of the oocytes to reach GVBD (G50). The G50 for each indicatedtreatment was normalized to G50 in the Con treatment. Summary data are presented as the mean± SEM; the n number is indicated on each bar.

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

Treatment Polar body Max GVBD Normalized time to G50

Controla 7/7 (100%) 84% ± 8% 1CALM 5/5 (100%) 79.9% ± 4.6% 1.2 ± 0.1CALM1,2,3,4 4/4 (100%) 70.6% ± 6.1% 1.4 ± 0.1Controla 11/12 (91.7%) 86.9% ± 13.2% 1MLCK-control 16/25 (64%) 81.9% ± 14.5% 1.04 ± 0.003MLCK-peptide 14/27 (51.9%) 84.2% ± 15.8% 0.93 ± 0.04

aUninjected cells.


ca2+ homeostasis regulates xenopus oocyte maturation

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