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Am J Physiol Cell Physiol 292: C1510-C1518, 2007. First published November 8, 2006; doi:10.1152/ajpcell.00181.2006
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GROWTH, DIFFERENTIATION, AND APOPTOSIS

Cell cycle-dependent calcium oscillations in mouse embryonic stem cells

Nidhi Kapur, Gregory A. Mignery, and Kathrin Banach

Department of Physiology, Stritch School of Medicine, Loyola University Chicago, Maywood, Illinois

Submitted 12 April 2006 ; accepted in final form 4 November 2006


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
During cell cycle progression, somatic cells exhibit different patterns of intracellular Ca2+ signals during the G0 phase, the transition from G1 to S, and from G2 to M. Because pluripotent embryonic stem (ES) cells progress through cell cycle without the gap phases G1 and G2, we aimed to determine whether mouse ES (mES) cells still exhibit characteristic changes of intracellular Ca2+ concentration during cell cycle progression. With confocal imaging of the Ca2+-sensitive dye fluo-4 AM, we identified that undifferentiated mES cells exhibit spontaneous Ca2+ oscillations. In control cultures where 50.4% of the cells reside in the S phase of the cell cycle, oscillations appeared in 36% of the cells within a colony. Oscillations were not initiated by Ca2+ influx but depended on inositol 1,4,5-trisphosphate (IP3)-mediated Ca2+ release and the refilling of intracellular stores by a store-operated Ca2+ influx (SOC) mechanism. Using cell cycle synchronization, we determined that Ca2+ oscillations were confined to the G1/S phase (~70% oscillating cells vs. G2/M with ~15% oscillating cells) of the cell cycle. ATP induced Ca2+ oscillations, and activation of SOC could be induced in G1/S and G2/M synchronized cells. Intracellular Ca2+ stores were not depleted, and all three IP3 receptor isoforms were present throughout the cell cycle. Cell cycle analysis after EGTA, BAPTA-AM, 2-aminoethoxydiphenyl borate, thapsigargin, or U-73122 treatment emphasized that IP3-mediated Ca2+ release is necessary for cell cycle progression through G1/S. Because the IP3 receptor sensitizer thimerosal induced Ca2+ oscillations only in G1/S, we propose that changes in IP3 receptor sensitivity or basal levels of IP3 could be the basis for the G1/S-confined Ca2+ oscillations.

pluripotent; IP3; store operated Ca entry; IP3 receptor

PLURIPOTENT MOUSE EMBRYONIC stem cells have a high rate of proliferation (6, 23, 40) that does not depend on the addition of serum to the culture medium (41). Mouse ES (mES) cells further remain uninfluenced by contact inhibition. Their cell cycle is characterized by a virtual lack of the G1 and G2 gap phases that in somatic cells accompany and control cell cycle progression. As a consequence, ~50% of the ES cells sojourn in the S phase at any given point of culture (11, 23, 41). In ES cells and cells of the inner cell mass of the blastocyst, an active phosphatidylinositol (PtdIns) signaling system is described (9, 13, 23, 34); thereby, the phosphatidylinositol-3 kinase (PI3K) appears as a key regulator for cell survival as well as cell cycle progression. Blocking PI3K activity results in increased DNA fragmentation and induces a reversible cell cycle arrest during G1/S transition (23). Apoptosis as well as proliferation seem to involve the Akt pathway, whereas the MAP kinase-ERK pathway played no significant role in either process (23). The other important branch of the PtdIns signal transduction cascade involves the activation of phospholipase C (PLC), where the hydrolysis of phosphatidylinositol 4,5-bisphospate results in the production of the second messengers inositol 1,4,5-trisphosphate (IP3) and diacylglycerol (DAG). IP3 and DAG mediate the release of Ca2+ from intracellular stores and the activation of PKC, respectively. In the mouse, inhibition or knockout of PLC prevents blastocyst formation as well as proliferation of ES cells in vitro (34, 42). Although experimental results point to a relevance of the intracellular Ca2+ release in proliferation, the mechanisms of Ca2+ signaling during cell cycle progression are not yet described.

In mammalian somatic cells, the importance of intracellular Ca2+ signaling during cell cycle progression is well established (3, 37). Spontaneous Ca2+ oscillations can be observed in the G0/G1 phase, where they correlate with the activation of immediate-early genes that initiate reentry into the cell cycle. After a quiescent phase, oscillations resume in late G1, before the S phase initiation, and promote DNA synthesis. During G2/M transition, the increase in intracellular Ca2+ concentration ([Ca2+]i), which has been shown to depend on an increase in IP3 production, is further related to the breakdown of the nuclear envelope (7, 37). The cell cycle-dependent Ca2+ signaling relies on the presence of Ca2+ in the extracellular solution as well as on the presence of intact intracellular Ca2+ stores. Removal of Ca2+ from the solution or inhibition of Ca2+ reuptake into the ER results in most cells in cell cycle arrest in the G1/S phase (19, 30). The store-operated Ca2+ (SOC) entry as well as T-type Ca2+ channels have been described as Ca2+ entry mechanisms relevant for cell proliferation (12, 30, 32, 35, 44). The Ca2+ signals that can be recorded during the cell cycle vary from continuous increases in the basal Ca2+ level to individual transient increases and continuous Ca2+ oscillations (30). Recent studies have shown that mES cells are comparable to mesenchymal stem cells, having functional IP3 receptor (IP3R)-regulated intracellular Ca2+ stores. They are sensitive to stimulation with ATP, histamine, and platelet-derived growth factor and depend on SOC entry for the refilling of their intracellular Ca2+ stores. However, in contrast to mesenchymal stem cells, no Ca2+ oscillations were described (26, 36, 47).

In the present study, we tested the hypothesis that in mES cells, IP3-mediated Ca2+ oscillations are present in the different phases of the cell cycle and are a key element for cell cycle progression. Our results demonstrate that ES cells exhibit spontaneous Ca2+ oscillations that depend on IP3-mediated Ca2+ release. Although the principal mechanisms for Ca2+ signaling are maintained throughout the cell cycle, oscillations are confined to the transition from the G1 to the S phase of the cell cycle. The data reveal new information about the regulation of Ca2+ signaling in mES cells and its contribution to cell cycle progression.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Cell culture. Mouse ES cells of the cell line CRL (Specialty Media; Chemicon International) were propagated in culture as previously described (1, 18). The cells were maintained on a layer of mitomycin-inactivated primary mouse fibroblasts (Specialty Media; Chemicon International) in DMEM supplemented with 15% fetal calf serum (FCS; Invitrogen) and leukemia-inhibiting factor (LIF: ESGRO; 1,000 U/ml) to prevent differentiation. For the experiments, ES cells were trypsinized and plated onto gelatin-coated coverslips (diameter 25 mm) in a density of 0.1 x 106 cells/ml. No specific measures were taken to remove the fibroblasts from the culture; on average, an individual coverslip therefore contained 7 x 104 fibroblasts/ml. The proliferation of fibroblasts was inhibited by mitomycin treatment before coculture; therefore, they were outnumbered within 24 h of culture. Their distinct morphology allowed their identification, and Ca2+ measurements were only obtained within ES cell colonies. All cultures were maintained in LIF. Experiments were performed on control cultures the first day after plating. Pharmacological synchronization was started 24 h after plating, and cells were incubated for a minimum of 12 h before experimental use.

Cell cycle synchronization and fluorescence-activated cell sorting. Cell cycle synchronization of mES cells was achieved by 12-h incubation of the cells in hydroxyurea (HU; 2 mM), aphidicoline (APC; 20 mg/ml), or mimosine (MIM; 500 µM) for synchronization in the transition from G1 to S phase and in nocodazole (NOC; 100 ng/ml) or demecolcine (DC; 20 ng/ml) for synchronization in the transition from G2 to M phase (21). After 12 h, the cells were washed with Tyrode and used for Ca2+ imaging experiments within 2 h. Cell cycle distribution of control and synchronized cultures was determined using fluorescence-activated cell sorting (FACS; Becton Dickinson FACStar Plus). Cells were washed with PBS, fixed with 10-min incubation in ice-cold ethanol (70%), and, after washing, treated with RNase (100 mg/million cells in PBS; 30 min at 37°C). After washing, the cells were stained with propidium iodide (PI; 50 µg/ml) for 1 h at 4°C. PI intercalates into the cell's double-stranded nucleic acid and results in fluorescence that can be detected ~600 nm when excited at 488 nm. The FACS analysis then allows cell sorting for fluorescence intensity and therefore DNA content. Data were acquired with Cell Quest software, and the percentages of G1, S, and G2 phase cells were calculated with MODFIT software.

Microsomes. Cells were harvested and microsomes were prepared as described previously (30, 31). Briefly, cells were harvested and transferred into 50 mM Tris·HCl, pH 8.3, 1 mM EDTA, 1 mM beta-mercaptoethanol, 1 mM PMSF, 10 µM leupeptin, 10 µM pepstatin, and 100 µg/ml trypsin inhibitor and then lysed by 40 passages through a 27-gauge needle. Membranes were pelleted by a 20-min centrifugation (289,000 gav), resuspended in buffer, and either used immediately or frozen at –80°C.

SDS-PAGE and immunoblotting. Microsomes were analyzed using 5% SDS-PAGE followed by electrotransfer to nitrocellulose membranes and subjected to immunodetection with IP3R isoform-specific antibodies, described previously (38, 39), using chemiluminescence reagents (Amersham Pharmacia Biotech).

Fluorescence measurements. Spatially averaged measurements of the change in [Ca2+]i were performed on individual cells within an ES cell colony by using the Ca2+-sensitive dye fluo-4 AM. Fluo-4 AM (50 µg) was dissolved in DMSO (50 µl), Pluronic (50 µl of 20% stock in DMSO), and FCS (87.5 µl). We achieved a final dye concentration of 2.5 µM in standard extracellular solution. The coverslips with ES cells were incubated with the dye solution for 15 min, and deesterification of the dye was allowed for another 15 min.

Solutions. During the experiment, the cells were superfused with a standard extracellular Tyrode solution containing (in mM) 140 NaCl, 5.0 KCl, 1.0 MgCl2, 5.5 glucose, 10 HEPES, and 2 CaCl2; pH 7.4. To analyze the contribution of Ca2+ influx to the spontaneous oscillations, in some experiments (Ca2+ free) we omitted Ca2+ from the extracellular solution. Further pharmacological tools used were 2-aminoethoxydiphenyl borate (2-APB; 10 µM), ATP (10 µM), BAPTA-AM (1 µM), caffeine (10 mM), lanthanum (0.1 mM), ryanodine (Ry; 10 µM), thapsigargin (1 µM), thimerosal (10 µM), and U-73122 (10 µM). All chemicals were purchased from Sigma.

Confocal imaging. [Ca2+]i was monitored using two-dimensional confocal imaging (Zeiss LSM-410 microscope). Fluo-4 AM was excited at a wavelength of 488 nm, and emission was detected at >515 nm. Changes in [Ca2+]i are expressed as R = F/F0, where R is the fluorescence (F) normalized to the resting fluorescence (F0). Because of the slow time course of spontaneous Ca2+ oscillations, two-dimensional confocal images were taken at a time interval of 5 s. Intracellular Ca2+ was analyzed in defined regions of interest that included one entire cell over the entire time series.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Spontaneous Ca2+ oscillations in ES cells. Undifferentiated ES cells in culture form multicellular colonies that exhibit a distinct round boundary without signs of outgrowth (8, 41). After 1 day in culture in the presence of LIF, the observed colony size ranged from 19.8 to 73 µm (n = 46, mean: 45.9 ± 15 µm) with a mean cell diameter of 10.3 µm. Following the loading of the Ca2+-sensitive dye Fluo-4 AM, we monitored the time-dependent change in [Ca2+]i in individual cells within a colony. Figure 1, A and B, shows a representative example of an ES cell colony in which individual cells exhibited spontaneous Ca2+ oscillations. The average amplitude of spontaneous oscillations was 1.57 ± 0.7 [extracellular Ca2+ concentration ([Ca2+]o) = 2 mM]. They occurred with an intertransient interval (ITI) of 76 ± 24 s and a mean duration of 38 ± 10 s (at 50% amplitude; n = 32 experiments). The rise in [Ca2+]i was not strongly synchronized among neighboring cells, although intercellular coupling of cell colonies could be confirmed by intercellular diffusion of the dye Lucifer yellow (data now shown). Overall, in control colonies, spontaneous oscillations occurred in 36% of the cells within the colony. The results demonstrate that undifferentiated ES cells exhibit spontaneous Ca2+ oscillations; however, in control cultures the cells are heterogeneous in their spontaneous Ca2+ signaling.


Figure 1
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  		Fig. 1. Mouse ES (mES) cells exhibit spontaneous Ca2+ oscillations. Two-dimensional confocal images were taken at 5-s intervals from a mES cell colony loaded with the Ca2+-sensitive dye fluo-4 AM. A: an image sequence visualizes the oscillatory increase of the intracellular Ca2+ concentration ([Ca2+]i) within individual cells of the colony. The sequence of images is displayed from left to right and from top to bottom at 25-s intervals. B: the fluorescence (F/F0) of individual cells plotted as a function of time displays oscillatory changes in [Ca2+]i that are not synchronized among neighboring cells. The numbers of individual traces 1–5 correspond to the cells numbered in the first image of A; asterisks mark the points of time that correspond to the images in A. Oscillations exhibited a mean interval of 76 ± 24 s, a duration of 38 ± 10 s (at 50% amplitude), and an amplitude of 1.57 ± 0.7 (n = 32 experiments).

 
Ca2+ oscillations do not depend on Ca2+ influx but on intracellular Ca2+ release. Spontaneous Ca2+ oscillations can be driven by either Ca2+ influx (L-type, T-type, or SOC entry) or Ca2+ release from intracellular Ca2+ stores like the ER (3, 4, 25). We tested whether Ca2+ influx induces spontaneous Ca2+ oscillations in ES cells by superfusing ES cell colonies with a Ca2+-free extracellular solution. As shown in Fig. 2A, spontaneous changes in [Ca2+]i continued even in the absence of Ca2+ from the extracellular solution. This result was confirmed in 5 independent experiments including 22 oscillating cells. However, as shown in Fig. 2C (n = 5), in most cells Ca2+ transient amplitudes decreased with time in Ca2+-free solution. The data underline that extracellular Ca2+ is not required for the induction of Ca2+ oscillations; however, it is required to maintain the filling of the intracellular stores.


Figure 2
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  		Fig. 2. Modulation of mES cell Ca2+ oscillations by extracellular Ca2+ ([Ca2+]o) or ryanodine receptor (RyR). A: representative trace of [Ca2+]i from 1 cell within an mES cell colony during superfusion with control (Ctrl; [Ca2+]o = 2 mM) and Ca2+-free solution. Ca2+ transient amplitude (TA) decreased slowly during continuous superfusion with Ca2+-free solution (C). TAs were normalized to the last transient before application of Ca2+-free solution (n = 5 experiments; means ± SE). B: characteristic recording of Ca2+ oscillations before and during superfusion with ryanodine (Ry; 10 µM). Intertransient interval (ITI; D) and amplitude (E) of Ca2+ oscillations remained unchanged in the presence of Ry (%control; n = 4). Data in D and E are presented as means (SD); in D and E, changes were not significant (Student's t-test: P > 0.05).

 
Ca2+ oscillations depend on IP3-mediated Ca2+ release. In most electrically excitable cells, Ca2+ is released from the intracellular stores through the ryanodine receptor (RyR; Ref. 10). The signal is mediated by a Ca2+-induced Ca2+ release mechanism (CICR). In electrically unexcitable cells like endothelial cells and the mouse oocyte, Ca2+ is mostly released through IP3Rs. To determine whether the spontaneous oscillations in ES cells are mediated by CICR, we superfused the cultures with Ry (10 µM), an antagonist of RyR that, at this concentration, blocks the RyR in its open configuration (10, 29). Figure 2B shows a representative recording from an ES cell exhibiting spontaneous Ca2+ oscillations. In all spontaneously oscillating ES cells (n = 5), superfusion with Ry did not block spontaneous oscillations or change the ITI (Fig. 2D) or the amplitude of the oscillations (Fig. 2E). To confirm the result, we superfused cells with the RyR agonist caffeine (10 mM; not shown); however, no caffeine-induced Ca2+ release could be observed. Caffeine exposure was reduced to <5 s. The results indicate that spontaneous oscillations of Ca2+ do not depend on RyR-mediated Ca2+ release or CICR.

To determine whether spontaneous oscillations of [Ca2+]i depend on the presence of IP3-sensitive intracellular stores, we superfused ES cell colonies with the IP3R antagonist 2-APB (10 µM) (46). Application of 2-APB resulted in a reversible inhibition of Ca2+ oscillations, indicating the contribution of Ca2+ release from IP3-sensitive stores (Fig. 3A; n = 5; 15 oscillating cells). 2-APB is a rather unspecific inhibitor of the IP3R and has further inhibitory action on gap junction channels and SOC entry (16, 33). In an alternative approach, we tested the aminosteroid U-73122 {1-[6-({(17beta)-3-methoxyestra-1,3,5[10]-trien-17-yl}amino)hexyl]-1H-pyrrole-2,5-dione; 10 µM}, an inhibitor of PLC that was previously used to examine the signaling mechanisms in ES cells (15, 34). Also in this case, as shown in Fig. 3B, spontaneous oscillations were suppressed in four independent experiments. Our data indicate that spontaneous oscillations in undifferentiated ES cells depend on the release of Ca2+ from IP3R-regulated intracellular stores and are in accordance with previous reports that demonstrate the presence of all three IP3R isoforms in mES cells (45) (see also GoGoFig. 6D).


Figure 3
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  		Fig. 3. Ca2+ oscillations depend on inositol 1,4,5-trisphosphate receptor (IP3R)-mediated Ca2+ release. Representative recordings of [Ca2+]i within a single cell during superfusion with the IP3R antagonist 2-aminoethoxydiphenyl borate (2-APB; A) and the PLC inhibitor U-73122 (B). 2-APB and U-73122 both inhibited spontaneous oscillations of [Ca2+]i in undifferentiated ES cells.

 

Figure 4
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  		Fig. 4. Store-operated Ca2+ (SOC) entry in mES cells. A: changes in [Ca2+]i in a single mES cell during store depletion with the endoplasmic reticulum Ca2+-ATPase inhibitor thapsigargin (Thaps) in the absence and presence of [Ca2+]o and during superfusion with La3+. B: La3+ reversibly blocked 75% of the Ca2+ entry induced by depletion of the intracellular Ca2+ stores. Data are presented as means (SD).

 

Figure 5
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  		Fig. 5. Pharmacologically induced changes of cell cycle distribution. A: histogram showing cell cycle distribution of individual experiments analyzed using fluorescence-activated cell sorting (FACS). Histograms of control, mimosine (MIM), and demecolcine (DC) synchronized cultures are superimposed. B: bar graph allows the direct comparison of the percentage of cells that reside in the G0/G1, S, or G2/M phases of the cell cycle in control conditions and after MIM or DC treatment. Data are presented as means (SD) obtained from 3 independent experiments.

 

Figure 6
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  		Fig. 6. Oscillations of [Ca2+]i in synchronized mES cells. Bar graphs present the percentage of oscillating cells within individual mES cell colonies (A), the interval of the Ca2+ transients (B), and their transient duration in control cultures and cultures that were synchronized with DC, NOC, MIM, aphidicoline (APC), or hydroxyurea (HU) (C). D: immunoblots demonstrate the presence of the IP3R isoforms T1, T2, and T3 in control cultures and cultures that were serum starved (0%) or synchronized with HU or NOC, respectively. Data are expressed as means (SD). (Student's t-test: *P > 0.05; {dagger}P > 0.05).

 
SOC entry in ES cells. Recent data demonstrated the presence of a SOC entry mechanism in mesenchymal as well as mES cells (26, 47). To determine whether this mechanism is present in our cell system and whether it contributes to the refilling of the intracellular Ca2+ stores, we superfused mES cells with the Ca2+-ATPase inhibitor thapsigargin (1 µM) in a Ca2+-free solution (47). In the experiment shown in Fig. 4A, thapsigargin induced a transient increase in Ca2+ due to the uncompensated release of Ca2+ from the intracellular stores. Readdition of Ca2+ (4 mM) to the culture resulted in a reversible increase of Ca2+. We found that 75% of this Ca2+ influx could be blocked by lanthanum (0.1 mM), an inhibitor of SOC (Fig. 4B; n = 9 cells). The data indicate that refilling of intracellular Ca2+ stores in ES cells is mediated by a SOC influx mechanism.

ES cells exhibit heterogeneity in their spontaneous Ca2+ oscillations. Mouse ES cells are derived from the inner cell mass of the mouse blastocyst; they are pluripotent and can be propagated indefinitely (8). Although they should represent a uniform population of cells, spontaneous Ca2+ oscillations could only be observed in 36% of the cells within individual colonies. To determine whether the heterogeneity depends on the ES cells' cell cycle progression, we synchronized ES cell cultures at different stages of the cell cycle before examining their Ca2+ oscillations. To evaluate whether the Ca2+ oscillations coincide with the cells' transition from G1 to S phase, we synchronized the cell cultures 24 h after plating by 12-h incubation in MIM (500 µM) (21, 31, 43). For synchronization of the cells in the transition from G2 to M phase, cells were incubated for 12 h with DC (20 ng/ml). FACS analysis of PI-stained MIM- and DC-treated cultures revealed a shift of the cell cycle distribution from control conditions. Whereas in control cultures, 50.4 ± 1.3% of the cells resided in the S phase, that number decreased to 38.6 ± 0.7 and 14.8 ± 1.9% in MIM- and DC-treated cultures, respectively (see Fig. 5, A and B). In MIM-treated cultures, the majority of the cells resided in the G1 phase (control: 19.6 ± 4.1%; MIM: 50.1 ± 11.1%), whereas in DC-treated cultures, the cells accumulated in G2/M (control: 32.1 ± 7.4%; DC: 72.8 ± 11.9%), demonstrating a successful shift in cell cycle distribution.

Analysis of the spontaneous Ca2+ oscillations in the synchronized cultures revealed that incubation of the cells with MIM resulted in an increase in the number of cells exhibiting Ca2+ oscillations (control: 36 ± 23%; MIM: 58 ± 23%; Fig. 6A). This increase correlates well with the accumulation of the cells in the G1 phase of the cell cycle (control: 19.6%; MIM: 50.1%; Fig. 5A). In contrast, in DC-treated cultures, only 8 ± 8% oscillating cells were found within individual ES cell colonies. Also in this case, the significant decrease in oscillating cells correlates with the low number of cells residing in the G1 phase of the cell cycle after DC synchronization (4.0 ± 3%). To rule out the possibility that the change in the number of oscillating cells was due to the cell cycle drugs themselves, rather than to cell cycle synchronization, we repeated the experiment with other cell cycle inhibitors. To block the transition from the G1 to the S phase, we additionally used either APC (20 µg/ml) or HU (2 mM) to block the transition from G2 to M, we further used NOC (100 ng/ml) (21, 43). Analysis of the Ca2+ oscillations in the APC- and HU- or NOC-treated cells revealed results comparable to those obtained with MIM and DC, respectively. The number of cells with Ca2+ oscillations increased to 72 ± 13 and 76 ± 22% in APC- and HU-treated colonies, respectively, whereas Ca2+ oscillations decreased to 15 ± 11% of the cells in NOC-treated cultures (Fig. 6A). The data show that Ca2+ oscillations within ES cell colonies occur during the cells' progression through the cell cycle and that those in synchronized colonies correlate with the percentage of cells residing in the G1 phase.

We measured and compared the frequency and the duration of the spontaneous oscillations of [Ca2+]i that we recorded in the synchronized cultures. The analysis revealed that neither ITI nor the transient duration (TD) were significantly different in control cultures and cultures treated with DC, NOC, MIM, APC, or HU. Mean ITI values ranged from 70 to 99 s, whereas mean TD values ranged from 38 to 47 s (Fig. 6, B and C). The comparability of the Ca2+ transients observed in the control and synchronized cultures further underlines that we are looking at a defined Ca2+ oscillation pattern that marks the transition from the G1 to the S phase of the cell cycle.

Ca2+ signaling mechanisms during different phases of the cell cycle. To understand the mechanism of the time-restricted presence of Ca2+ oscillations in the transition from G1 to S, we determined the presence of cell cycle-dependent changes in the Ca2+ handling mechanisms (e.g., IP3R expression, IP3-mediated Ca2+ release, SOC, and the basal activity of PLC). In Fig. 6D, immunoblots are shown that were obtained from control cultures, cultures maintained in 0% FCS for 12 h, and cultures synchronized with either HU in the G1/S transition or NOC in the G2/M transition. In all cultures examined, we identified the IP3R isoforms 1, 2, and 3. The data indicate that the confinement of the Ca2+ oscillations to the G1/S phase most likely does not depend on a change in IP3R isoform expression. To exclude the possibility that the reduction of Ca2+ oscillations in the G2/M phase of the cell cycle is due to a depletion of the intracellular Ca2+ stores and a reduction of the SOC entry mechanism, we superfused HU- and DC-synchronized cultures with thapsigargin. Representative experiments for a HU (Fig. 7A, n = 4) and a DC (Fig. 7B, n = 3) treated culture are shown. In both cultures, thapsigargin induced a transient increase in [Ca2+]i due to the leak of Ca2+ from the intracellular stores. Readdition of Ca2+ (2 mM) to the extracellular solution resulted in the previously described Ca2+ influx mediated by SOC (see also Fig. 4A). The data point out that the lack of Ca2+ oscillations in G2/M synchronized ES cells is not based on a depletion of the intracellular Ca stores or the lack of Ca2+ influx through SOC.


Figure 7
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  		Fig. 7. Presence of SOC in HU- and DC-synchronized mES cells. Representative recordings of changes in [Ca2+]i in individual mES cells from cultures that were synchronized by treatment with either HU (A) or DC (B). In both stages of the cell cycle, thapsigargin induced depletion of the intracellular Ca2+ stores that resulted in the activation of SOC.

 
It was reported that ATP induces a transient elevation of [Ca2+]i in undifferentiated ES cells (47). The ATP-induced Ca2+ release is mediated by PLC-dependent IP3 production, stimulated by a G protein-coupled purinergic receptor. Cells that were blocked in their cell cycle progression by DC or NOC between the G2 and M phase or by MIM, APC, or HU between the G1 and S phase all responded with a transient increase in [Ca2+]i upon stimulation with ATP (G1/S: n = 4; G2/M: n = 5). Representative recordings of a culture synchronized with HU and DC are shown in Fig. 8, Aa and Ba, respectively. The data underline that receptor-mediated IP3 production and IP3-mediated Ca2+ release are not impaired in either phase of the cell cycle.


Figure 8
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  		Fig. 8. IP3-mediated Ca2+ oscillations in synchronized mES cells. Representative recordings of ATP- or thimerosal-induced changes in [Ca2+]i. Responses shown are from individual mES cells that were synchronized by treatment with either HU (A) or DC (B). C: comparison of the mean change of ATP-induced Ca2+ transients in control cultures and cultures synchronized at either G1/S (APC, HU) or G2/M (NOC, DC). Data are expressed as means (SD). D: cell cycle progression of control cultures and cultures that were maintained in the presence of either U-73122 (10 µM) or EGTA (3 mM).

 
Spontaneous changes in IP3 production as they occur in sea urchin eggs (7, 20, 29) have been discussed to induce spontaneous Ca2+ oscillations during cell cycle progression. We tested for potential differences in IP3 levels or IP3R sensitivity in cultures synchronized with either DC or NOC (block between G2 and M) or by MIM, APC, or HU (block between G1 and S). We superfused the synchronized cultures with thimerosal, a thiol reagent described to increase the sensitivity of IP3R for its agonist (5). In control, APC-, and HU-synchronized cells, thimerosal (10 µM) rapidly increased the frequency of oscillations, which ultimately resulted in an increase of the basal Ca2+ levels (n = 4). However, in DC- or NOC-treated cultures, thimerosal remained without effect (n = 3). Since IP3-mediated Ca2+ release is present at all stages of the cell cycle, the transition from the G1 to the S phase seems to coincide with changes of IP3R open probability or IP3 production.

Relevance of Ca2+ oscillations for cell cycle progression. To determine the relevance of the Ca2+ oscillations observed for cell cycle progression, we determined the growth rate of undifferentiated ES cells in control medium and in medium that was supplemented with either the PLC inhibitor U-73122 (10 µM) or EGTA (3 mM). A semilogarithmic plot of the cell numbers counted in one experiment is shown in Fig. 8D. The reduced slope in the cultures treated with U-73122 and EGTA indicates a decreased rate of proliferation. The mean values of five independent experiments reveal that under control conditions, the cells had an average doubling time of 10.2 ± 0.16 h, whereas in the presence of U-73122 or EGTA, cultures exhibited an increased doubling time of 15.3 ± 2.3 and 20.9 ± 1.27 h, respectively (n = 5). A similar effect was obtained after incubation of proliferating ES cell cultures with 2-APB (10 µM), thapsigargin (1 µM), or the membrane-permeable Ca2+ buffer BAPTA-AM (1 µM) to block an increase in [Ca2+]i. In all cases, a significant slowing in cell cycle progression could be detected compared with control cultures (Fig. 9A). FACS analysis of U-73122-, 2-APB-, thapsigargin-, and BATA-AM-treated cultures after 24 h of incubation revealed in all cases a reduced number of cells residing in the G2/M phase, whereas the number of cells in the G1 phase of the cell cycle was increased (Fig. 9B). U-73122-, 2-APB-, and BAPTA-AM-treated cultures, in contrast to thapsigargin, also had increased numbers of cells in the S phase. The data underline the relevance of PLC-mediated Ca2+ oscillations for cell cycle progression through the G1/S phase in mES cells.


Figure 9
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  		Fig. 9. Delay of cell cycle progression by inhibitors of IP3-mediated Ca2+ release. A: cell cycle progression of control, 2-APB (n = 6)-, BAPTA-AM (n = 4)-, U-73122 (n = 2)-, and thapsigargin (n = 6)-treated cultures normalized to the number of control cells. B: FACS analysis of the same culture treatments after 12-h incubation reveals a shift of the cell cycle distribution toward the G1/S phase of the cell cycle. Data are presented as normalized changes from control cultures. (Significance of the change in cell cycle distribution was confirmed using the {chi}2 test: P < 0.005).

 

   DISCUSSION
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
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In the present study, we have demonstrated that cell cycle progression of mES cells through the G1/S phase is accompanied by spontaneous oscillations of [Ca2+]i. Because intracellular Ca2+ signaling mechanisms like the presence of IP3R isoforms, loading of Ca2+ stores, PLC-mediated IP3 production, and SOC seem to remain unchanged during the cell cycle, we propose that these Ca2+ oscillations depend on an increased sensitivity of IP3R or increased levels of IP3 synthesis.

Ca2+ oscillations in mES cells. In mammalian cells, Ca2+ is used as an ubiquitous second messenger (4), and in mES cells that were differentiated into endodermal cells, Ca2+ oscillations mediate exo- and endocytotic vesicle shuttling. There is substantial evidence that Ca2+ oscillations also play an important role in cell cycle progression (38). In fibroblasts, HeLa, and smooth muscle cells, Ca2+ oscillations accompany the transition from the G0 to the G1 phase (2), and in the transition from the G1 to the S phase, a basal elevation of [Ca2+]i is described. The ES cell cycle is characterized by a virtual lack of G1 and G2 gap phases, and the majority of the cells (50%; see Fig. 4, A and B) sojourn in the S phase at any given point of culture (11, 23, 29, 41). We identified Ca2+ oscillations in 36% of the cells within an ES cell colony under control conditions. The lack of observation in previous studies may be related to the low incidence of oscillating cells (26, 47). Ca2+ oscillations in ES cell preparations are described in primitive endodermal cells after induction of differentiation (absence of LIF, culture in an embryoid body) (39). Although the oscillatory pattern is comparable, the progressed differentiation may explain their dependence on extracellular Ca2+ and sensitivity to nickel (39). We determined that the oscillations in mES cells coincide with the G1 phase of the cell cycle by pharmacologically blocking cell cycle progression. The substances MIM, APC, and HU induced arrest at the transition from the G1 to the S phase by inhibiting the formation of replication forks and by DNA polymerase or ribonucleotide diphosphate reductase inhibition, respectively (21, 31, 43). Although the effects of the drugs we used were different, all resulted in an increased number of oscillating cells (58, 72, and 76%, respectively for MIM, APC, and HU). The consistent outcome excludes the possibility that the increase in oscillations was due to an unspecific effect of the individual substances. We confirmed the change in cell cycle distribution by FACS analysis of MIM-treated cultures. The number of cells residing in G1 increased from 19 ± 4 to 50 ± 11% (n = 2) after 12 h of MIM treatment. The data underline that in MIM-treated cultures, the G1 phase is the dominating cell cycle stage. The effectiveness of cell cycle synchronization is in good agreement with previously published data (11, 43). The percentage of oscillating cells correlates well with the number of cells in G1. Nevertheless, since we observed 8 and 15% oscillating cells in mES cell cultures synchronized with either DC or NOC, we cannot rule out the possibility that oscillations occur in other stages of the cell cycle. In other cell types, Ca2+ oscillations are described in the G2 as well as the M phase of the cell cycle (29, 38). However, transient duration and frequency of the Ca2+ oscillations in DC- or NOC-treated cultures were comparable to those observed in G1/S synchronized cultures. Because FACS analysis of DC-treated cultures revealed that 5% of the cells still ranked in the G1 and 14% in the S phase of the cell cycle, we hypothesize that the observed Ca2+ oscillations in ES cells are characteristic of the transition from the G1 to the S phase of the cell cycle.

IP3R-mediated Ca2+ release. The regulation of intracellular Ca2+ signals depends on the interplay of intracellular and extracellular Ca2+ sources (4). Entry of Ca2+ into the cell occurs through voltage-dependent, receptor-operated, and store-operated Ca2+ channels, whereas the ER functions as an intracellular Ca2+ source. Ca2+ release from the ER is controlled by IP3R and/or RyR (4). To maintain Ca2+ oscillations, cells need a source for the immediate increase of [Ca2+]i. Our experimental data indicate that not Ca2+ influx (Fig. 2A) but the release of Ca2+ from intracellular stores is the primary source for the oscillatory rise in [Ca2+]i. The identification of all three IP3R isoforms (Fig. 6D; Ref. 47), the sensitivity of the oscillations to 2-APB (Fig. 3A), and their insensitivity to Ry (Fig. 3B) support the hypothesis that IP3R-operated intracellular stores contribute to the oscillatory rise in [Ca2+]i.

Ca2+ influx and extrusion mechanisms. After the release of Ca2+ from the intracellular stores, Ca2+ removal from the cytoplasm can occur through either reuptake of Ca2+ into the intracellular stores or extrusion over the plasma membrane by the Na+/Ca2+ exchange (NCX) mechanism or the plasma membrane Ca2+-ATPase (PMCA). NCX and PMCA are expressed in undifferentiated ES cells (47); however, the continuation of Ca2+ oscillations for up to 15 min (Fig. 2A) indicates reuptake of Ca2+ into intracellular stores. However, average data show a decrease of the transient amplitude in the presence of Ca2+-free solution (Fig. 2C), supporting the hypothesis that Ca2+ influx is necessary to maintain the amplitude of the oscillations. Overall, our data support the hypothesis that SOC entry is a major Ca2+ influx mechanism in mES cells. It can maintain the filling of the intracellular Ca2+ stores after depletion (Figs. 4A and 7, A and B). This hypothesis is also supported by previous reports that demonstrate the presence of SOC but that failed to identify other influx mechanisms via voltage-dependent T- or L-type Ca2+ channels (47, 48).

Maintenance of intracellular Ca2+ oscillations. Different paradigms exist to explain sustained intracellular Ca2+ oscillations. In undifferentiated ES cells, the increase of Ca2+ depends on its release from IP3R-operated stores. That this increase also coincides with the generation of IP3 by a PLC-dependent mechanism is supported by the block of the oscillations by the PLC inhibitor U-73122 (Fig. 3B) and their amplification in the presence of thimerosal (Fig. 8A). The presence of IP3-mediated Ca2+ signaling mechanisms is further supported by the induction of Ca2+ transients after receptor-mediated stimulation of the PLC signaling pathways by either ATP (Fig. 8, Aa and Ba) or histamine (47). The critical question is why Ca2+ oscillations dominate in the G1/S phase of the cell cycle. Our experimental results indicate that the SOC entry is present in G1/S as well as G2/M synchronized cells (Fig. 7), and as a result, Ca2+ stores are also filled in both phases of the cell cycle (Fig. 7). These data exclude depletion of the stores as a reason for the lack of Ca2+ oscillations in G2/M. We observed Ca2+ transients as a result of receptor-stimulated IP3 production (Fig. 8, Aa and Ba) and detected all three IP3R isoforms in G1/S and G2/M synchronized cells (Fig. 6D). Therefore, lack of IP3Rs or IP3-generating second messenger pathways cannot account for the lack of Ca2+ oscillations. In sea urchin eggs, changes in the basal concentration of IP3 during the cell cycle were described as the basis for changes in [Ca2+]i (7). A similar mechanism could be present in undifferentiated ES cells. The lack of thimerosal-induced Ca2+ oscillations in G2/M synchronized cells could indicate that the basal second messenger concentration is not high enough to induce IP3-mediated Ca2+ release or that the sensitivity of IP3R is reduced (Fig. 8, Ab and Bb) at this stage. These changes could be brought about by subtle changes in the basal level of [Ca2+]i and the activity of CaM or CaMKII (45). Other regulators of Ca2+ homoeostasis have been related to cell cycle regulation, and the expression levels of proteins such as PMCAs (22), CaM, or CaMK have yet to be determined (38) in ES cells.

IP3R-mediated Ca2+ release and cell cycle progression. A role of the PtdIns signaling pathway in mES cell cycle progression has been proposed previously, but its relation to IP3-mediated Ca2+ release has not yet been demonstrated (9, 14, 23). Our data show ES cell proliferation significantly decreases in the presence of drugs that prevent a rise in [Ca2+]i, or more specifically, PLC- and IP3-mediated Ca2+ release (Figs. 8D and 9A). They are in good agreement with a recent report that EGF stimulates proliferation of mES cells via PLC-dependent changes in [Ca2+]i (17). Although changes in [Ca2+]i also might be important for other phases of the cell cycle, a clear shift of the cell cycle distribution to the G1/S phase could be observed, further supporting the relevance of this signaling mechanism in this phase of the cell cycle. The significant decrease of cells in the S phase after thapsigargin treatment could be based on the apoptotic action of thapsigargin (28); however, it remains to be determined whether prolonged suppression of IP3-mediated signaling could promote a change in the cellular phenotype by induction of differentiation. The IP3-mediated Ca2+ release may not be the only mechanism involved in the ES cells cell cycle regulation. Other potential Ca2+ entry pathways such as SOC or the voltage-operated L- and T-type Ca2+ channels have been attributed to regulate cell proliferation in pulmonary smooth muscle and rat neonatal cardiomyocytes, as well as tumor cells (27, 32, 35). Although we have not excluded their influence individually, analysis of mRNA does not indicate the presence of voltage-dependent Ca2+ channels in these cells (47, 48).

In conclusion, in the present study we have demonstrated that mES cells exhibit IP3-mediated oscillations of [Ca2+]i that are confined to the transition from the G1 phase to the S phase of the cell cycle and that play a role in cell cycle progression. We propose that changes in IP3R sensitivity or changes of the basal level of IP3 could be the basis for the oscillations, and we plan to address this in future experiments. The results reveal further insight into cell cycle progression in ES cells and describe a signaling mechanism that could promote the cells' rapid transition out of G1 and therefore support the preservation of their pluripotent state.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was supported by American Heart Association Grant 0330393Z (to K. Banach), Potts Foundation Loyola University Chicago Grant RFC 11086 (to K. Banach), and National Institute of Mental Health Grant MH-53367 (to G. A. Mignery).


   FOOTNOTES
 

Address for reprint requests and other correspondence: K. Banach, Dept. of Physiology, Stritch School of Medicine, Loyola Univ. Chicago, 2160 South First Ave., Maywood, IL 60153 (e-mail: kbanac1{at}lumc.edu)

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


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 GRANTS
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