Ca2+ is a ubiquitous intracellular messenger that is important for cell cycle progression. Genetic and biochemical evidence support a role for Ca2+ in mitosis. In contrast, there has been a long-standing debate as to whether Ca2+ signals are required for oocyte meiosis. Here, we show that cytoplasmic Ca2+ (Ca2+cyt) plays a dual role during Xenopus oocyte maturation. Ca2+ signals are dispensable for meiosis entry (germinal vesicle breakdown and chromosome condensation), but are required for the completion of meiosis I. Interestingly, in the absence of Ca2+cyt signals oocytes enter meiosis more rapidly due to faster activation of the MAPK-maturation promoting factor (MPF) kinase cascade. This Ca2+-dependent negative regulation of the cell cycle machinery (MAPK-MPF cascade) is due to Ca2+cyt acting downstream of protein kinase A but upstream of Mos (a MAPK kinase kinase). Therefore, high Ca2+cyt delays meiosis entry by negatively regulating the initiation of the MAPK-MPF cascade. These results show that Ca2+ modulates both the cell cycle machinery and nuclear maturation during meiosis.
Mammalian and amphibian oocytes arrest at the G2/M border of the cell cycle after oogenesis (Yamashita et al., 2000). Before these oocytes become fertilization competent, they undergo a so-called “oocyte maturation” period during which they acquire the ability to activate in response to sperm entry, and to support the early stages of embryonic development (Yamashita et al., 2000). During maturation, oocytes progress through meiosis and arrest at metaphase of meiosis II until fertilization.
Xenopus oocyte maturation provides a valuable model to elucidate the signal transduction cascade mediating meiosis entry and progression. In Xenopus, oocyte maturation is triggered by the hormone progesterone, which binds to a cell surface receptor and not the classical nuclear receptor/transcription factor. Progesterone leads to inhibition of cAMP-dependent PKA and translation of the proto-oncogene c-Mos, which induces the MAPK cascade culminating in the activation of maturation promoting factor (MPF; for review see Nebreda and Ferby, 2000). MPF is the central kinase that regulates meiotic progression, and consists of a catalytic p34cdc2 serine/threonine kinase subunit (Cdk 1), and a regulatory cyclin B subunit (Coleman and Dunphy, 1994). MPF is also activated by the removal of inhibitory phosphorylations by the Cdc25C phosphatase, which is induced by the polo-like kinase cascade (Nebreda and Ferby, 2000).
A variety of genetic and biochemical evidence support a role for Ca2+, and its downstream effectors CaM and Ca2+-CaM–dependent protein kinase II, in mitosis initiation and progression (Means, 1994; Whitaker, 1995). Ca2+ signals are required for nuclear envelope breakdown (NEBD), and chromosome condensation during mitosis (Steinhardt and Alderton, 1988; Twigg et al., 1988; Kao et al., 1990; Tombes et al., 1992; Ciapa et al., 1994). Furthermore, the cytoplasmic Ca2+ (Ca2+cyt) rise observed at fertilization is the universal signal for egg activation in all species investigated (Stricker, 2000). Ca2+ signals at fertilization release the metaphase II arrest by activating proteolytic degradation of cytostatic factor, thus inducing completion of meiosis II and entry into the mitotic cell cycle (Tunquist and Maller, 2003). In addition, Ca2+ release at fertilization induces both the fast and slow blocks to polyspermy in Xenopus eggs (Machaca et al., 2001). In contrast to the well-defined roles for Ca2+ signals in mitosis and after fertilization, the role of Ca2+ signals during oocyte maturation remains contentious.
There has been a long-standing debate in the literature as to whether Ca2+ signals are required for Xenopus oocyte maturation/meiosis (Duesbery and Masui, 1996). Early reports argued that a Ca2+cyt rise is sufficient to induce oocyte maturation (Wasserman and Masui, 1975; Moreau et al., 1976; Schorderet-Slatkine et al., 1976). Furthermore, oocytes injected with high concentrations of Ca2+ buffers were unable to mature (Moreau et al., 1976; Duesbery and Masui, 1996). However, injection of IP3, which induces Ca2+ release, did not stimulate meiotic maturation (Picard et al., 1985). Additional support for a Ca2+ role in oocyte maturation comes from reports that measured a Ca2+cyt rise after progesterone addition using 45Ca2+ as a tracer, Ca2+ imaging, or Ca2+-sensitive electrodes (O'Connor et al., 1977; Moreau et al., 1980; Wasserman et al., 1980). In contrast, others could not detect Ca2+cyt changes downstream of progesterone addition using similar techniques (Robinson, 1985; Cork et al., 1987). A role for CaM in Xenopus oocyte maturation has also been postulated (Wasserman and Smith, 1981) and challenged (Cicirelli and Smith, 1987). These conflicting reports argue that the relationship between Ca2+ and oocyte maturation is complex.
We decided to revisit the role of Ca2+ during oocyte maturation/meiosis by framing the problem in terms of the spatially distinct sources of Ca2+ signals. Ca2+cyt signals can be generated either due to Ca2+ release from intracellular Ca2+ stores (ER) or Ca2+ influx from the extracellular space. In fact, these two Ca2+ sources are mechanistically linked through the store-operated Ca2+ entry (SOCE) pathway, which is activated in response to intracellular Ca2+ stores depletion. Therefore, Ca2+cyt is regulated by the balance between Ca2+ release and Ca2+ influx. By manipulating Ca2+ store load and the extent of Ca2+ influx through SOCE, we show here that Ca2+ signals are not required for meiosis entry. On the contrary, high Ca2+cyt delays meiosis entry. However, in the absence of Ca2+cyt signals oocytes arrest in meiosis I, form abnormal spindles, and do not extrude a polar body. Surprisingly, MAPK and MPF kinetics in oocytes deprived of Ca2+ signals are normal. We further mapped the site of action of Ca2+cyt on meiosis entry and show that Ca2+cyt negatively regulates the cell cycle machinery by acting downstream of PKA and upstream of Mos. These data argue that Ca2+ signals regulate the timing of meiosis entry, and that they are required for the completion of meiosis I. The dual role of Ca2+cyt revealed by these studies help explain some of the controversy surrounding the role of Ca2+ in oocyte maturation, and provides a framework to explore the role of Ca2+-dependent signaling cascades in meiosis.
Depleting intracellular Ca2+ stores accelerates entry into meiosis
Maturing oocytes in media with different Ca2+ concentrations ([Ca2+]) does not affect the time course or extent of germinal vesicle (nucleus) breakdown (GVBD; Fig. 1, A and C), which is indicative of meiosis entry. The rate of maturation in the population was quantified as the time required for 50% of the oocytes to undergo GVBD (GVBD50). Because the rate and extent of GVBD were unaffected in low Ca2+ (L-Ca) medium, this shows that Ca2+ influx is not required for entry into meiosis (Fig. 1, C and D).
To test whether intracellular Ca2+ levels affect oocyte maturation, we emptied Ca2+ stores either by treating cells with thapsigargin, an inhibitor of the ER Ca2+ ATPase (SERCA), or with the Ca2+ ionophore ionomycin. Thapsigargin leads to Ca2+ store depletion because of a poorly defined Ca2+ leak pathway from the ER (Camello et al., 2002). Emptying Ca2+ stores activates Ca2+ influx through SOCE (Parekh and Penner, 1997). Because the extent of Ca2+ influx through SOCE depends on [Ca2+] in the medium, oocytes incubated in high Ca2+ (H-Ca) medium will have more Ca2+ influx than those in normal Ca2+ (N-Ca) medium, and no Ca2+ influx in expected in L-Ca medium (see Fig. 4).
Emptying Ca2+ stores with either thapsigargin or ionomycin in N-Ca does not affect the time to GVBD50 (Fig. 1, B and C, Thap-N-Ca and Ion-N-Ca), but decreases maximal levels of GVBD (Fig. 1 D, Thap-N-Ca and Ion-N-Ca). In H-Ca, emptying Ca2+ stores results in high percentage of cellular degeneration due to excessive Ca2+ influx, thus prohibiting analysis of the rate of meiosis entry because GVBD50 is rarely reached under these conditions (Fig. 1 B, Thap-H-Ca and Ion-H-Ca). In contrast, emptying Ca2+ stores in L-Ca medium accelerates the rate of maturation (Fig. 1, B and C, Thap-L-Ca and Ion-L-Ca), without affecting maximal maturation levels (Fig. 1 D, Thap-L-Ca and Ion-L-Ca). In L-Ca medium with Ca2+ stores depleted, oocytes are unable to generate Ca2+ signals after progesterone addition because Ca2+ stores are depleted and Ca2+ influx is prevented in L-Ca medium (see Fig. 4); nonetheless they enter meiosis at an accelerated rate. These data show that Ca2+ signals are not required for GVBD and argue that Ca2+cyt negatively regulates meiosis entry.
Although Ca2+ signals after progesterone addition are dispensable for GVBD, it is conceivable that Ca2+cyt signals generated before progesterone addition are required for meiosis entry. To determine whether this is the case, we depleted Ca2+ stores with thapsigargin and waited for extended periods of time before inducing maturation with progesterone. We reasoned that if Ca2+ signals are required for GVBD, the longer we wait after depriving oocytes of Ca2+ signals the less effective progesterone will be in inducing maturation. Depleting stores for as long as 48 h does not affect the extent of oocyte maturation (Fig. 1 F), but still enhances oocyte maturation rate (Fig. 1 E). These results support the conclusion that Ca2+ signals are not required for entry into meiosis.
Lowering Ca2+cyt levels accelerates meiosis entry
The more rapid maturation observed in L-Ca medium when Ca2+ stores are depleted, argues that Ca2+cyt negatively regulates meiosis entry. It follows then that buffering Ca2+cyt at low levels should also accelerate meiosis entry. This is indeed the case as injection of 500 μM 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA) alone or in combination with thapsigargin in varying orders, results in a more rapid maturation (Fig. 2, A and B). BAPTA and thapsigargin treatments accelerate maturation to a similar extent with no significant additive effect. However, the longer the interval between BAPTA injection and progesterone addition the more rapid the maturation rate (Fig. 2 B). In all treatments the extent of maturation was comparable (Fig. 2 C). Similar results were obtained when BAPTA was injected at 1 mM (unpublished data). These data argue that at resting Ca2+cyt levels some Ca2+-dependent pathways are active and negatively regulate meiosis entry.
As reported by others, injecting high BAPTA concentrations (2.5–5 mM) blocks GVBD, arguing that Ca2+ is required for GVBD (Moreau et al., 1976; Duesbery and Masui, 1996). However, at such high BAPTA concentrations we observe significant levels of oocyte degeneration, bringing into question the specificity of this treatment. As shown below, injecting BAPTA at 500 μM effectively buffers Ca2+cyt transients (Fig. 4). In addition, depleting Ca2+ stores in the absence of extracellular Ca2+, thus depriving oocytes of Ca2+ signals, does not block GVBD. Furthermore, treating oocytes with the heavy metal chelator TPEN blocks oocyte maturation (unpublished data). Based on these findings, it is possible that high BAPTA concentrations block GVBD in a Ca2+-independent manner, either due to a nonspecific effect of BAPTA, or chelation of other metal ions because BAPTA is a potent chelator of transition metals (Arslan et al., 1985).
High Ca2+cyt delays meiosis entry
If Ca2+cyt negatively regulates meiosis entry it is expected that raising Ca2+cyt levels would lead to a slower rate of oocyte maturation. To test whether this is the case we induced different levels of Ca2+ influx through SOCE by depleting Ca2+ stores with thapsigargin and incubating oocytes with solutions containing 5 mM, 3 mM, 1.5 mM, 0.6 mM, and 50 μM Ca2+ (H-5, H-3, H-1.5, N-Ca, and L-Ca, respectively). Inducing maturation in solutions with different [Ca2+] has no effect on the rate of oocyte maturation, except for L-Ca medium where maturation rate was more rapid (Fig. 3, A and C). Although this enhancement was more pronounced in this set of experiments, Fig. 1 shows a similar tendency toward a more rapid maturation in L-Ca medium. More importantly, after store depletion with thapsigargin, the higher the concentration of extracellular Ca2+ the slower the rate of maturation (Fig. 3, B and C). Furthermore, the extent of maturation was reduced in H-Ca containing solutions (H-5, H-3, and H1.5). At both 3 and 5 mM of extracellular Ca2+ (H-5 and H-3) some cellular degeneration was observed, but at 1.5 mM of extracellular Ca2+ the oocytes were healthy, but the rate of oocyte maturation was slower. These data show that the higher the level of Ca2+ influx the slower the rate of maturation, supporting the conclusion that high Ca2+cyt levels negatively regulate meiosis entry.
Ca2+-activated Cl−currents (ICa,Cl) as markers for Ca2+cyt levels
To confirm that the different treatments are modulating Ca2+cyt levels as predicted, we used endogenous Ca2+-activated Cl− current (ICl,Ca), as an in situ marker of Ca2+cyt levels (Fig. 4). We have shown previously that ICl,Ca provides an accurate measure of both Ca2+ release and influx (Machaca and Hartzell, 1999). During Ca2+ release ICl,Ca is activated as a sustained current (ICl1) at depolarized voltages (+40 mV; Fig. 4 A, left, trace t). ICl1 is sustained because during Ca2+ release Ca2+cyt levels remain high for the duration of the voltage pulse. In contrast, during Ca2+ influx ICl,Ca is activated as a transient current (IClT) only when the +40 mV pulse is preceded by a hyperpolarization step (−140 mV) to induce Ca2+ influx (Fig. 4 A, right, traces w–z). IClT is transient because Ca2+ flows into the cell during the preceding −140 mV pulse, and then dissipates rapidly resulting in current inactivation (Machaca and Hartzell, 1999). We have shown using simultaneous electrical recording and Ca2+ imaging that ICl,Ca faithfully reports the levels and kinetics of Ca2+ release and influx (Machaca and Hartzell, 1999). However, it is important to note that although ICl,Ca provides an accurate measure of Ca2+ release and Ca2+ influx, it does not directly reflect Ca2+ levels deep in the cytosol as these channels localize to the plasma membrane.
To determine store Ca2+ load in the different treatments, we incubated oocytes in Ca2+-free Ringer (F-Ca), and depleted Ca2+ stores with ionomycin. Because no Ca2+ influx is possible in Ca2+-free solution, the level of ICl1 activated in response to ionomycin provides a measure of the extent of store Ca2+ load (Fig. 4, A and B). After the dissipation of the Ca2+ release transient indicated by the return of ICl1 to baseline (Fig. 4 B, squares), oocytes were sequentially exposed to L-Ca, N-Ca, H1.5-Ca, H3-Ca, and H5-Ca to determine the extent of Ca2+ influx (Fig. 4, B, D, and F). This protocol was applied to control untreated oocytes (Fig. 4 B) or to oocytes incubated in thapsigargin to fully deplete Ca2+ stores (Fig. 4 D), or injected with 500 μM BAPTA (Fig. 4 F). The levels of Ca2+ release as indicated by ICl1 and the levels of Ca2+ influx as indicated by IClT were quantified in the different treatments (Fig. 4, C, E, and G). In control oocytes ionomycin activates a large ICl1, indicating that Ca2+ stores are fully loaded (Fig. 4 B, squares; Fig. 4 C, Ca Rel.). Ca2+ release leads to store depletion which activates SOCE. As expected, no IClT is detected in either Ca2+-free solution (F-Ca) or in L-Ca medium (L-Ca) confirming our prediction that at 50 μM of extracellular Ca2+ no Ca2+ influx occurs (Fig. 4 B, circles; Fig. 4 C). Increasing levels of Ca2+ influx (indicated by IClT) are observed in media with increasing [Ca2+] (Fig. 4 B, circles; Fig. 4 C; Fig. 4 A, right).
Oocytes treated with thapsigargin did not release Ca2+ in response to ionomycin as no ICl1 was activated (Fig. 4 D, squares; Fig. 4 E, Ca Rel.), showing that Ca2+ stores were depleted. Because thapsigargin depletes Ca2+ stores, it activates Ca2+ influx through SOCE. As for control oocytes no Ca2+ influx is observed in F-Ca or L-Ca solutions, and higher levels of Ca2+ influx, as indicated by IClT, are detected in solutions containing increasing Ca2+ (Fig. 4 D, circles; Fig. 4 E). Ca2+ influx levels in thapsigargin-treated cells were similar to those in control cells (Fig. 4, C and E).
BAPTA injection dramatically reduces both Ca2+ release (ICl1) and Ca2+ influx (IClT) transients (Fig. 4, F and G). Small levels of Ca2+ release are observed in BAPTA-injected cells (Fig. 4 F, squares; Fig. 4 G, Ca Rel.), indicating that Ca2+ stores still contain Ca2+, but that as Ca2+ is released it is chelated by BAPTA, thus drastically reducing the levels of free Ca2+ available to activate ICa,Cl. The same is true during the Ca2+ influx phase in different [Ca2+] (Fig. 4, F and G). No Ca2+ influx can be detected in L-Ca solution, and small levels of IClT are observed in N-Ca through H3-Ca. Only H5-Ca produced evident, but small IClT consistently (Fig. 4 G). This indicates that the primary effect of BAPTA injection is to buffer Ca2+cyt at low levels. The fact that BAPTA injection enhances the rate of meiosis entry in a similar fashion to store depletion argues that this enhancement is due to a reduction of Ca2+cyt levels. It is noteworthy that Ca2+ store depletion has been shown to alter ER protein expression (Soboloff and Berger, 2002), however, based on the BAPTA data and the delayed maturation rate with high Ca2+cyt, it is unlikely that this is affecting meiosis entry.
Kinetics of MAPK and MPF activation
We assayed the rate and extent of oocyte maturation above based on the GVBD time course. GVBD marks entry into meiosis but does not provide any information about meiosis progression. Oocyte maturation is considered complete once oocytes reach metaphase of meiosis II. Although, the data presented so far show that Ca2+ signals are not required for entry into meiosis, they do not address whether meiosis/oocyte maturation can progress normally in the absence of Ca2+ signals. To determine whether interfering with Ca2+ signaling pathways affects meiosis progression, we tested the activation kinetics of the MAPK-MPF kinase cascade, which regulates meiosis transitions (Nebreda and Ferby, 2000). As described above (Figs. 1 and 2), treating cells with either thapsigargin or injecting BAPTA accelerates meiosis entry (Fig. 5 A). In N-Ca medium, MAPK phosphorylation is first detected 2 h before GVBD, peaks at GVBD, and remains high for the remainder of maturation (Fig. 4 B, N-Ca). MAPK activates with similar kinetics in L-Ca, or after the thapsigargin (Thaps) or BAPTA treatments (Fig. 5 B). However, consistent with the GVBD time course, MAPK activated 1 h earlier in L-Ca medium and 2 h earlier after thapsigargin or BAPTA treatments (Fig. 5 B). These data show that in the absence of Ca2+ signals MAPK is induced earlier, but has normal kinetics after GVBD.
MPF activates in a characteristic fashion during oocyte maturation with a sharp peak at GVBD followed by a decline to an intermediate level that is indicative of the meiosis I to meiosis II transition, and rises again as oocytes progress through meiosis II (Fig. 4 C, N-Ca). MPF activity in oocytes matured in L-Ca medium follows the same kinetics except that peak MPF activity (GVBD) occurs ∼1 h earlier than in N-Ca. This is consistent with the MAPK activation kinetics (Fig. 5 B) and the rate of GVBD (Fig. 5 A). MPF kinetics in the thapsigargin and BAPTA treatments are normal, except that, as for GVBD and MAPK, MPF activity peaks 2 h earlier than in the N-Ca treatment (Fig. 5 C, Thaps and BAPTA). Interestingly, an increase in MPF activity is detected as early as 1 h before GVBD (Fig. 5 C, Thaps and BAPTA, arrows). Such a premature activation of MPF is never observed in N-Ca or L-Ca. Therefore, MAPK and MPF activation kinetics (Fig. 5, B and C) correlate well with each other and with the rate of GVBD (Fig. 5 A), and show that reducing Ca2+cyt transients leads to premature activation of the MAPK-MPF kinase cascade. This premature activation explains the accelerated maturation rate in treatments that reduce Ca2+cyt transients. Therefore, Ca2+cyt modulates meiosis entry by negatively regulating the MAPK-MPF cascade.
Spindle formation and nuclear maturation
Kinase data in oocytes deprived of Ca2+ signals (Thaps and BAPTA) suggest that in the absence of Ca2+cyt signals meiosis proceeds normally after GVBD. To determine whether this is the case we imaged meiotic spindle structure and chromosome dynamics in oocytes matured in N-Ca, L-Ca, and oocytes treated with thapsigargin or BAPTA (Fig. 6). This allowed us to assess the progression of nuclear maturation, and directly compare it to MPF, MAPK, and GVBD kinetics because all three experiments were performed on the same batch of oocytes.
Control oocytes matured in N-Ca medium progress normally through meiosis (Fig. 6 A, Table I, N-Ca). At GVBD oocytes were at the late prophase I stage (Fig. 6 A, N-Ca, P), which refers to oocytes that have undergone GVBD, have condensed chromosomes, and organized microtubules around the chromosomes, but have not yet formed a bipolar spindle (Fig. 6 A, N-Ca, P). At 0.5 h after GVBD prometaphase I structures (30%; Table I) are observed with a typical bipolar spindle and associated chromosomes (Fig. 6 A, N-Ca, PM I). This is followed by metaphase I with chromosome lined up at the metaphase plate (Fig. 6 A, N-Ca, M I) at ∼1 h after GVBD (Table I). Between 2–4 h after GVBD oocytes progress from M I to metaphase II (Table I) at which stage they arrest. Examples of anaphase I (A), prometaphase II (PM II), and metaphase II spindles are shown in Fig. 6 A (N-Ca).
Surprisingly, oocytes matured in L-Ca medium alone or treated with thapsigargin or BAPTA formed abnormal spindles. The progression through meiosis was not significantly different between the three treatments which will be discuss as a group. At GVBD a large percentage of these oocytes (≥57%; Table I) had condensed chromosomes, but the microtubule were still dispersed over a large area. We refer to this stage as early prophase (EP; Fig. 6 A), because eventually these oocytes do progress to the late prophase stage (P) as described for the control group (N-Ca) above (Table I). However, oocytes matured in L-Ca medium or treated with BAPTA or thapsigargin rarely progress to prometaphase I and never reach metaphase I (Fig. 6 A; Table I). Instead, they form abnormal structures at different rates depending on the treatment as detailed in Table I. Based on the severity of defects we divided abnormal spindles into three groups: (1) We refer to small and/or slightly disorganized spindles as prometaphase like (PM-L; Fig. 6 A). These spindles are the least disorganized and are observed throughout the time period studied (Table I). (2) The second group represents completely disrupted spindles (Ab) with no clear structure and with the microtubules highly condensed and/or spread over a large area. In most but not all instances condensed chromosomes were still associated with the disrupted spindle (Fig. 6 A, Ab). (3) The last and most interesting group we refer to as the double spindle (DS) group (see Fig. 6 A). These double spindles were most common in the L-Ca group (Table I), but were also observed in the thapsigargin treatment (Fig. 6 A, Thaps, DS).
It is interesting that the highest percentage of disrupted spindles (Ab) in the experimental groups (L-Ca, Thaps, and BAPTA) is observed at 2–3 h after GVBD which corresponds to the meiosis I to meiosis II transition in control oocytes. At later time points, spindle structures are improved (PM-L), as if the cells are attempting to go through meiosis II. This is supported by the appearance of DS structures at 3–4 h after GVBD arguing that although these oocytes do not progress normally through meiosis I, they attempt to form a meiosis II spindle, that in some cases lead to the observed double-spindle structures. This might be expected with normal MAPK and MPF kinetics past GVBD in all the treatments (Fig. 5), which will drive these cells into meiosis II despite the abnormal progression through meiosis I. The fact that MPF and MAPK kinetics are normal past GVBD in the L-Ca, thapsigargin, and BAPTA treatments (Fig. 5), but that meiotic spindles are disorganized show that interfering with Ca2+ signaling during meiosis uncouples the MAPK-MPF cascade from spindle structure regulation. Furthermore, these results show that although Ca2+ signals are not required for meiosis entry (GVBD and chromosome condensation), they are necessary for progression through meiosis I and for bipolar spindle formation.
Polar body emission
Spindle structure data show that interfering with Ca2+ signaling leads to abnormal spindles, arguing that oocytes do not complete meiosis I. To directly confirm this, we assessed polar body emission from 1–4 h after GVBD in the different treatments (Fig. 6, B and C). In N-Ca medium, polar bodies were observed in the majority of the cells, but as expected from the spindle structure experiments, oocytes matured in L-Ca medium or treated with thapsigargin or BAPTA rarely extrude a polar body (Fig. 6 C). This shows that interfering with Ca2+ signaling inhibits the completion of meiosis I.
The fact that oocytes matured in L-Ca medium had abnormal spindle structure and could not finish meiosis I argues that either Ca2+ influx from the extracellular space is required during the early stages of oocytes maturation for normal meiosis progression, or that oocytes are unable to form a polar body at low extracellular Ca2+. To differentiate between these possibilities we assessed polar body emission in oocytes incubated in N-Ca solution until GVBD and then switched to L-Ca solution (N-L); or in oocytes incubated in L-Ca solution until GVBD and then switched to N-Ca medium (L-N). Oocytes in the N-L group, but not the L-N group, emitted polar bodies to the same extent as the N-Ca control group (Fig. 6, B and C), arguing that Ca2+ influx in the early stages of oocyte maturation is required for meiosis I progression.
Ca2+cyt acts between PKA and Mos to negatively regulate entry into meiosis
To better define the mechanism by which Ca2+cyt negatively regulates meiosis entry, we mapped the site of action of Ca2+cyt on the cell cycle machinery using an epistatic approach. For these experiments thapsigargin-treated oocytes represented the experimental group deprived of Ca2+ signals. Control oocytes were activated in N-Ca solution. Our approach was to activate the cell cycle machinery at different points along the MAPK-MPF signal transduction cascade, and determine whether the enhancing effect of Ca2+ depravation on the rate of maturation is still observed. Oocytes were activated with progesterone, or by injection of an inhibitor of PKA inhibitor (PKI), Mos RNA (Mos), cyclin B1 RNA (Cy) and Δ87cyclin B1 protein (CyP; Fig. 7 A). Thapsigargin treatment enhanced the rate of oocyte maturation to a similar extent in oocytes activated with progesterone or PKI (Fig. 7 A, PKI). Thapsigargin-treated oocytes activate faster than controls after Mos RNA injection, but the effect on Ca2+ deprivation on the rate of maturation is smaller than in the case of progesterone (Fig. 7 A, Mos). Because the kinetics of the kinase cascade downstream of Mos are similar in control and thapsigargin-treated oocytes (Fig. 7 C; Fig. 5), we wondered whether the faster maturation rate in thapsigargin-treated Mos-injected oocytes is due to an effect of Ca2+ deprivation on RNA translation. To determine whether this is the case we induced meiosis by directly activating MPF through the expression of cyclin B1 to activate the free cdc2 pool in the oocyte. Similar to Mos RNA injection, thapsigargin-treated oocytes activated faster than controls after cyclin B1 RNA injection (Fig. 7 A, Cy). In contrast, injecting oocytes with cyclin B1 protein induces GVBD with a similar time course in both thapsigargin-treated and control oocytes (Fig. 7 A, CyP). The rate of oocyte maturation with the different activators is summarized in Fig. 7 B. Because the rate of maturation varies between activators we normalized maturation rate for each activator to the rate of activation in N-Ca (Fig. 7 B) to allow a better visualization of the relative effect of Ca2+ deprivation on maturation. For example, although cyclin B activates maturation faster than Mos (Fig. 7 A), the relative enhancement in the rate of maturation (∼20% faster) is similar between the two activators in the absence of Ca2+ signals (Fig. 7 B). This argues that the more rapid maturation in oocytes deprived of Ca2+ signals after both Mos and cyclin RNA injections is due to an effect of Ca2+cyt on translation of the injected RNAs. This conclusion is supported by the fact that control and thapsigargin-treated oocytes mature at similar rates when activated with Δ87cyclin B1 protein. However, the different responses after cyclin RNA or protein injections could be due to an effect of Ca2+cyt on protein turnover because we injected full-length cyclin B1 RNA and Δ87cyclin B1 protein, which is missing the first 87 aa and is thus nondegradable because it lacks the destruction box (Kumagai and Dunphy, 1995). Nonetheless, we favor an effect of Ca2+cyt on RNA translation because maturation is enhanced to a similar level when oocytes are activated with Mos or Cyclin B1, two activators that induce the cell cycle kinase cascade at different points.
These data show that Ca2+cyt negatively regulates meiosis entry by acting on at least two sites between PKA inhibition and Mos activation. One site is downstream of PKA inhibition and the other site appears to be mRNA translation, which is required for the induction of the cell cycle machinery (Fig. 7 D). Furthermore, the fact that the rate of maturation is enhanced to a similar extent in oocytes activated with progesterone and PKI (Fig. 7, A and B, PKI) argues that Ca2+cyt acts downstream of PKI (Fig. 7 D).
Ca2+cyt negatively regulates the initiation of the MAPK-MPF cascade
To confirm that Ca2+cyt acts upstream of Mos we analyzed in more details the steps of the cell cycle machinery downstream of Mos in both control and thapsigargin-treated oocytes (Fig. 7 C). As described above MAPK activates significantly earlier in thapsigargin-treated oocytes consistent with the GVBD time course (Fig. 7 C). p90RSK, the downstream substrate of MAPK is activated with a similar time course to MAPK. For these experiments we analyzed lysates form oocytes at GVBD and at GVBD50. For the latter time point lysates from oocytes that have undergone GVBD (w) or not (nw) were collected. Interestingly, p90RSK was phosphorylated to higher levels in the GVBD50-nw group in thapsigargin-treated oocytes as compared with controls, thus confirming the earlier activation of the MAPK cascade in thapsigargin-treated oocytes.
Xenopus oocytes contain two pools of cdc2, the catalytic subunit of MPF: the preMPF and the free cdc2 pool. The preMPF pool which is activated at GVBD, contains cdc2 associated with cyclin B. Pre-MPF is kept inactive by phosphorylation on Tyr15 of cdc2 (Nebreda and Ferby, 2000). The free cdc2 pool is activated after association with B-type cyclins synthesized during meiosis I (Hochegger et al., 2001). To determine which pool of cdc2 is activated in thapsigargin-treated oocytes we probed Western blots with a phosphospecific antibody against Tyr15 of cdc2 (Fig. 7 C, P-Y15-cdc2), and determined MPF activity as the H1-kinase activity from p13suc1 pulldowns (Fig. 7 C, MPF). In both control and thapsigargin-treated oocytes the disappearance of P-Y15 immunoreactivity coincides with an increase in MPF activity indicating that as in control oocytes the preMPF pool is activated in thapsigargin-treated oocytes. Interestingly, a small level of MPF activity is detected at the GVBD50 time point in thapsigargin-treated oocytes that have not undergone GVBD (Fig. 7, Thaps, G50 nw), confirming the early activation of MPF before GVBD observed in Fig. 5.
These data show that in the absence of Ca2+cyt signals the cell cycle kinase cascade downstream of Mos activates normally. Therefore, the more rapid entry into meiosis (GVBD) observed in oocytes deprived of Ca2+ signals is due to a negative regulation of Ca2+cyt on the initiation of this kinase cascade upstream of Mos. Blocking Ca2+cyt signals relieves this negative regulation thus allowing more rapid induction of the MAPK-MPF cascade and GVBD.
In contrast to the established role of Ca2+ signaling in mitosis (Whitaker and Larman, 2001), the requirement for Ca2+ in both Xenopus and mammalian oocyte meiotic maturation has been difficult to define (Homa et al., 1993; Duesbery and Masui, 1996). To delineate the function of Ca2+ during Xenopus oocyte meiosis we manipulated Ca2+cyt, extracellular Ca2+, and store Ca2+ load and tested the effect on nuclear maturation and the cell cycle machinery. Our data show that Ca2+ has two opposing roles during Xenopus oocyte maturation: It negatively regulates meiosis entry by delaying the activation of the cell cycle machinery, and it is required for completion of meiosis I (Fig. 7 D).
Ca2+cyt negatively regulates the activation of the cell cycle kinase cascade
Progesterone leads to lower cAMP levels and PKA inhibition within 10 min (Sadler and Maller, 1981), but the next known step in the pathway, that is polyadenylation of maternal RNAs to induce their translation, does not occur until much later (Sheets et al., 1995). The molecular steps during this time lag are not known. Our data show that Ca2+cyt is an important regulator of the transition between PKA inhibition and mRNA translation. Ca2+cyt negatively regulates the activation of the cell cycle machinery by acting on at least two sites between PKA and Mos (Fig. 7 D). One site of action appears to be mRNA translation. Therefore, the level of Ca2+cyt provides a timing mechanism for entry into meiosis by regulating the initiation of the MAPK-MPF cascade downstream of PKA inhibition (Fig. 7 D). It is tempting to propose that by acting in this capacity Ca2+cyt could synchronize morphological and biochemical changes during oocyte maturation. Under such a scenario, which is completely speculative at this point, Ca2+cyt levels could signal the physiological preparedness of the oocyte to begin maturation. Relatively low Ca2+cyt levels would be indicative of proper functioning of the Ca2+ signaling machinery, and thus a healthy oocyte that is ready to mature. In contrast, relatively high Ca2+cyt levels would indicate a compromised oocyte where Ca2+cyt would negatively regulate initiation of maturation. It is interesting in that context that Ca2+cyt acts upstream of Mos, that is before the oocyte activates the cell cycle machinery and commits to maturation. The proposed role of Ca2+cyt in synchronizing morphological and biochemical changes in the oocyte during maturation is further supported by the fact that disrupting Ca2+cyt signaling uncouples the nuclear cell cycle from the MAPK-MPF kinase cascade (Fig. 6).
Ca2+cyt is required for completion of meiosis I
Oocytes deprived of Ca2+ signals do not complete meiosis I as they do not extrude a polar body. Rather, they form abnormal spindles early in meiosis I despite normal MAPK and MPF kinetics. This shows that progression through meiosis I requires Ca2+, possibly Ca2+ influx before GVBD because oocytes are dependent on extracellular Ca2+ only before GVBD (Fig. 6). A Ca2+ influx requirement before GVBD fits nicely with the regulation of SOCE during oocyte maturation because SOCE inactivates at the GVBD stage due to MPF activation (Machaca and Haun, 2000, 2002).
Interestingly, others have shown that inhibition of the MAPK cascade (Gross et al., 2000), down-regulation of MPF (Nakajo et al., 2000), or inhibition of protein synthesis (Kanki and Donoghue, 1991) block completion of meiosis I. These treatments lead to a decrease in MPF activity, and induce an interphase-like state that is usually absent between meiosis I and II. In contrast, interfering with Ca2+ signaling blocks meiosis I completion, but is not associated with an interphase-like state. Rather, spindle structure is disrupted and polar body formation inhibited, but the chromosomes remain condensed. Furthermore, MPF activity cycles normally with a dip in activity between 1–2 h after GVBD, the expected time for meiosis I to meiosis II transition. These data show that disruption of Ca2+ signaling uncouples the cell cycle machinery (MAPK-MPF) from nuclear maturation (i.e., bipolar spindle formation and completion of meiosis I).
It is interesting that the disrupted meiosis I spindle does not activate a spindle checkpoint to arrest the cell cycle. However, there is good evidence against the existence of a meiosis I spindle checkpoint in Xenopus oocytes. Blocking the activity of the APC/C or the checkpoint protein Mad2 does not affect progression through meiosis I (Peter et al., 2001; Taieb et al., 2001). This is consistent with our observation of a lack of cell cycle arrest in the absence of Ca2+ signals despite disrupted meiosis I spindles.
Recently, Castro et al. (2003) described a similar block of meiosis I after inhibition of Aurora A kinase or its substrate Eg5 (a kinesin-like protein) in Xenopus oocytes. Unfortunately, these authors did not assess spindle morphology, but they show that blocking Aurora A inhibits polar body formation with chromosomes maintaining their condensed state long after GVBD (Castro et al., 2003). Members of the Aurora kinase family associate with the spindle and have been shown to be important for both meiosis and mitosis transitions. Therefore, it is possible that Ca2+-dependent pathways somehow modulate Aurora A kinase activity which in turn regulates spindle structure. This possibility remains to be explored.
Role of Ca2+ signaling in GVBD
During mitosis NEBD has been shown to be dependent on Ca2+ (Poenie et al., 1985; Steinhardt and Alderton, 1988; Twigg et al., 1988; Kao et al., 1990; Wilding et al., 1996), and studies in sea urchin embryos suggest that Ca2+ exerts its effect through CaMKII activation (Baitinger et al., 1990). In contrast, during meiosis GVBD is Ca2+-independent as shown here for Xenopus oocytes, and in both mouse (Carroll and Swann, 1992; Tombes et al., 1992) and starfish oocytes (Witchel and Steinhardt, 1990). One exception to this rule are some bivalve molluscs where GVBD has been shown to require Ca2+ (Deguchi and Osanai, 1994), but unlike amphibian and mammalian oocytes, in this case oocyte maturation and GVBD occur after fertilization which invariably induces a Ca2+cyt rise. Nonetheless, the differential requirement of NEBD on Ca2+ signals during meiosis and mitosis is surprising because both NEBD and GVBD require the activation of MPF (Lenart and Ellenberg, 2003), and it is reasonable to assume that the basic structural properties of the nuclear envelope are similar in mitotic and meiotic cells. It has been argued that a Ca2+ signal is still required for GVBD but occurs very early or even before the initiation of oocyte maturation in some species (Tombes et al., 1992; Homa et al., 1993). This does not seem to be the case for Xenopus, because as shown in Fig. 1 F, eliminating Ca2+ signals for as long as 48 h before inducing oocyte maturation has no effect on GVBD, strongly arguing that GVBD is Ca2+ independent. Therefore, the differential requirement for Ca2+ during the breakdown of the nuclear envelope suggests that NEBD and GVBD are mechanistically distinct.
In conclusion, our results show that Ca2+ signals are dispensable for GVBD and chromosome condensation, that Ca2+cyt controls the timing of meiosis entry by negatively regulating the initiation of cell cycle machinery, and that N-Ca2+ homeostasis is important for bipolar spindle formation and completion of meiosis I. These results provide a framework to further explore and better define the role of Ca2+-dependent signaling pathways in meiosis and oocyte maturation.
Materials And Methods
Xenopus oocytes were obtained as described previously (Machaca and Haun, 2002). The control l-15 solution contains 0.63 mM Ca2+. Ca2+ was buffered at 50 μM 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. For the H-Ca l-15 solutions Ca2+ was added to the indicated concentration as CaCl2. In all experiments, GVBD was visually confirmed by fixing oocytes in methanol and bisecting them in half.
Recording of the ICa,Cl was performed as described previously (Machaca and Haun, 2000). ICa,Cl were recorded in the following solutions: F-Ca contains in micromolars: 96 NaCl, 2.5 KCl, 5 MgCl2.6H2O, 0.1 EGTA, 10 Hepes, pH 7.4. Low Ca2+ Ringer solution (50 μM free Ca2+) contains in micromolars: 96 NaCl, 2.5 KCl, 4.37 MgCl2.6H2O, 0.63 CaCl2.2H2O, 0.58 EGTA, 10 Hepes, pH 7.4. Normal Ringer (N-Ca, 0.63 mM Ca2+) and H-Ca solutions (1.5, 3, and 5 mM Ca2+) had the following composition: 96 NaCl, 2.5 KCl, 10 Hepes, pH 7.4; with Ca2+ and Mg2+ concentrations adding up to 5 mM.
Western blots and MPF kinase assays
MPF kinase activity assay and phospho-MAPK Western blots were prepared as described previously (Machaca and Haun, 2002), except that α-tubulin was used as the loading control for MAPK Western blots. The activation of p90RSK and preMPF was assessed using phosphospecific antibodies against phospho-Thr573 of p90RSK and phospho-Tyr15 of cdc2 (Cell Signaling). MPF activity was also assayed from lysates affinity purified on p13suc1 beads using histone-H1 kinase as a substrate essentially as described previously (Howard et al., 1999).
Plasmids and reagents
Heat stable PKI was purchased from Calbiochem. Cyclin B1 and Mos RNAs were synthesized from a pXen-GST-Mos and pSP64-cyclinB1xen plasmids provided by A. Macnicol (University of Arkansas for Medical Sciences; Freeman et al., 1991; Howard et al., 1999) using the mMessage Mmachine transcription kit (Ambion). The His6-tagged Δ87cyclin B1 protein was used as described previously (Machaca and Haun, 2002).
Spindle and polar body staining and image acquisition
Oocytes were fixed in 100% methanol, bisected in half, and incubated in DM1A an antitubulin mAb (Sigma-Aldrich) in TBS containing 2% BSA, followed by a Cy2-conjugated donkey anti–mouse secondary (Jackson ImmunoResearch Laboratory) for 24 h each. The oocytes were washed, dehydrated, stained in 1 μM Sytox® orange (Molecular Probes), and cleared in benzyl alcohol/benzyl benzoate (1:2). Images were collected using a Fluoview confocal (Olympus) coupled to a microscope (model IX70; Olympus) at RT, using a UPlanApo 40× oil objective with an NA of 1.00. The acquisition software was Fluoview 2.1 and figures were compiled using Adobe Photoshop 7.0. For each spindle a z section was obtained and projected onto a single plane to visualize the entire spindle. For polar body emission studies oocytes were fixed in methanol, stained with Sytox® orange, and visualized by confocal microscopy as described for the spindle staining.
We thank members of the Machaca Lab for critical reading of the manuscript, A. Macnicol for providing plasmid constructs, and A. Charlesworth for help with the p13suc1 MPF assay.
This work was supported by grant GM-61829 from the National Institutes of Health.
Abbreviations used in this paper: BAPTA, 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid; CA2+cyt, cytoplasmic Ca2+; F-Ca, Ca2+-free Ringer; GVBD, germinal vesicle breakdown; H-Ca, high Ca2+; ICa,Cl, Ca2+-activated Cl− currents; L-Ca, low Ca2+; MPF, maturation promoting factor; N-Ca, normal Ca2+; NEBD, nuclear envelope breakdown; PKI, PKA inhibitor; SOCE, store-operated Ca2+ entry.