Heart and Stroke Richard Lewar Centre of Excellence, University of Toronto, Division of Cell and Molecular Biology, Toronto General Hospital Research Institute, and Department of Medicine, University of Toronto, Toronto, Ontario, Canada M5G-2C4
Submitted 7 November 2002 ; accepted in final form 17 March 2003
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ABSTRACT |
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calcineurin; c-Myb; plasma membrane Ca2+-ATPase-4; cell cycle
We previously showed that the cell cycle-associated repression of PMCA1
expression during G0 to G1/S progression in rat VSMC is
mediated by the c-Myb transcription factor
(1). However, the mechanism(s)
underlying the cell cycle-associated repression of PMCA4 had not been
elucidated. Guerini et al.
(18) demonstrated in mouse
neurons that PMCA4 expression can be repressed by a calcineurin-dependent
pathway. Given this result, we hypothesized that G1/S-associated
repression of PMCA4 expression in VSMC may also be mediated by calcineurin. To
explore these mechanisms in cell culture, we generated a clonal, immortalized
mouse VSMC line (MOVAS). Immuno-staining for smooth muscle-specific proteins
such as SM22, calponin, smooth muscle-specific
-actin and
desmin, as well as SM22
promoter-driven enhanced green fluorescent
protein (EGFP) expression, confirm the lineage and phenotype of these cells.
45Ca efflux assays and fura 2-based ratiometric
Ca2+ imaging reveal regulated Ca2+
efflux and [Ca2+]i at the G1/S
transition point. Western blot and real time RT-PCR reveal cell
cycle-regulated repression of mouse PMCA1 and PMCA4. Drugs inhibiting
calcineurin activity, such as the Ca2+-chelating agent
BAPTA and the calmodulin-dependent protein kinase-II (CaMK-II) inhibitor
KN-93, and retroviruses encoding either a calcineurin-inhibitory peptide
(CAIN), a peptide inhibitor (VIVIT) of the calcineurin-dependent nuclear
factor of activated T cells [NFAT; a transcription factor inhibited by the
peptide sequence VIVIT (please see Fig.
4A)], or a doxycycline (Dox)-inducible c-Myb neutralizing
antibody (ANTI-MYB), along with assays for CaMK-II activity and reporter
constructs responsive to calcineurin/NFAT and c-Myb, were employed to document
the effects of these modulators on calcineurin-dependent transcription,
[Ca2+]i, and PMCA4 expression during the
MOVAS cell cycle.
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MATERIALS AND METHODS |
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Cell migration. The migration rate of MOVAS cells was determined in a wound closure assay as previously described (27).
Immunofluorescence. Cells were stained with the following
antibodies (Ab): FITC-conjugated anti-smooth muscle specific -actin
(1:200), anti-calponin (1:100), and anti-desmin (1:10) (all from Sigma,
Mississauga, ON, Canada); anti-SM22
(1:100; rabbit polyclonal; Rolf
Ryseck, Squibb, Princeton, NJ), Oregon Green-conjugated anti-Rabbit IgG (1:50;
Molecular Probes, Eugene, OR), and FITC-conjugated anti-mouse IgG (1:50
dilution; Pierce, Rockford, IL). Slides were mounted and examined with a
confocal microscope (RTS2000, Bio-Rad, Hercules, CA).
SM22 promoter-driven EGFP. The enhanced green
fluorescent protein (EGFP) cDNA fragment of pCX-EGFP (courtesy of Dr. A Nagy,
Toronto, Canada) was inserted into pXU6 (courtesy of Dr. J. Miano, Rochester,
NY) downstream of the SM22
promoter
(29,
42) to obtain the plasmid
pSE12. MOVAS cells were transfected with pSE12 and Effectene as
per the manufacturer's specifications (Qiagen, Mississauga, ON, Canada). Cells
were imaged using fluorescent microscopy. Transfected Chinese hamster ovarian
(CHO)-luc cells (BD Biosciences, Mississauga, ON, Canada) served as a
negative control.
Flow cytometry. MOVAS cells were serum starved for 48 h, pulse labeled with bromodeoxyuridine (BrdU) at 0, 8, 16, and 24 h after serum stimulation as described (9), incubated with FITC-conjugated anti-BrdU antibody (BD Biosciences), and stained with propidium iodide. Cells were counted in a flow cytometer (FACScan, BD Biosciences), and G0, G1, S, and G2 cell percentages were calculated with Cell Quest software (BD Biosciences).
45Ca efflux. Calcium efflux assays were carried out as previously described (23). In experiments involving La3+, 1 mM LaCl3 was added to the efflux solution. Using plots of intracellular 45Ca vs. time, efflux rates were calculated. Each experiment was repeated in triplicate.
Calcium imaging. [Ca2+]i in cell cycle-synchronized MOVAS cells was measured using fura 2-AM as previously described (22, 23). Free [Ca2+]i was measured by fluorescence ratio imaging using an Image-Master DeltaRAM system (Photon Technology International, London, ON, Canada) with an Olympus IX70 inverted microscope and an IC-200 intensified charge-coupled device camera. Actual [Ca2+]i (nM) were calculated from experimental ratios using established formulas derived from in situ calibrations with ionophore and known [Ca2+]i after appropriate background subtraction (17).
Calcineurin- and c-Myb-responsive reporter assays. MOVAS cells were transduced with ANTI-MYB, CAIN (30), or VIVIT (2) for 24 h. Cells were then transfected with 2 µg of mim1-luc [a c-Myb-responsive luciferase reporter; courtesy of J. Lipsick, Palo Alto, CA (13)], 2 µg of mouse PMCA1-promoter-luciferase reporter (1), or 2 µg of a NFAT-IL-2 promoter-firefly luciferase reporter construct [NFAT-luc (37)] and 50 ng of a thymidine kinase promoter-renilla luciferase reporter (pRL-TK; Promega, Madison, WI) using Lipo-fectamine (Invitrogen, Burlington, ON). Cells were starved for 48 h posttransfection and harvested at 0 and 16 h postserum stimulation. Luciferase activity was measured with the DLR kit (Promega) with a luminometer (Berthold-Lumat; Wallac/Perkin-Elmer, Boston, MA). Raw relative light units (RLU) were corrected for transfection efficiency by subtracting the corresponding renilla luciferase-based RLU. The mean corrected RLU for nontransduced G0 cells was used to normalize corrected RLU for all other samples.
Retroviral constructs for CAIN, VIVIT, and anti-Myb. PA317 (ATCC no. CRL-9078, Manassas, VA) fibroblasts carrying a helper virus conferring retroviral packaging function [pPAM3 (33)] were employed. Empty-vector (MINV; courtesy of R. G. Hawley, Bethesda, MD), EGFP (BD Biosciences), CAIN [courtesy of J. Molkentin, Cincinnati, OH (10)] or VIVIT [courtesy A. Rao, Boston, MA (2)] encoding cDNAs were cloned into MINV while an anti-c-Myb antibody-encoding cDNA [courtesy of D. T. Curiel (28)] was cloned into a Dox-inducible retroviral expression vector (pTRE-Rev; BD Biosciences) to generate ANTI-MYB. These recombinant constructs were transfected into PA317 followed by drug selection to generate cell lines used for virion harvest.
CaMK-II activity assay. CaMK-II autophosphorylation activity
assays were performed in vitro as previously described
(46), with some modifications.
Cytosolic (S-100) protein extracts were prepared from G0 and
G1/S stage MOVAS cells treated or untreated with KN-93 (1 µM)
for 1624 h before harvest. For each sample, 100 µg of cytosolic
extract were hybridized with 500 ng of anti-CaMK-II polyclonal antibody
recognizing all CaMK-II isoforms (catalog no. SC9035; Santa Cruz
Biotechnology, Santa Cruz, CA) and immunoprecipitated with protein G-Sepharose
beads. The beads were exposed to 2.5 µCi [-32P]ATP (6000
Ci/mmol) and 0.1 µM calmodulin in kinase buffer
(34) without exogenous
substrate and with (Ca2+-dependent autophosphorylation)
or without Ca2+ (Ca2+-independent
autophosphorylation) at 30°C for 10 min, resolved on a 38% SDS-PAGE
gradient gel, blotted onto nitrocellulose, and autoradiographed. NIH Image
software was used to quantify intensities of the 53-kDa CaMK-II radiolabeled
bands. Values were background subtracted and normalized to the untreated
G0 value. In drug-treated samples, KN-93 (1 µM) was included
during the protein extraction and the phosphorylation step of the activity
assay. To measure Ca2+-independent CaMK-II activity,
EGTA (2 mM) was added during the kinase step.
Pharmacological agents. For experiments employing induction of ANTI-MYB, Dox (0.5 µg/ml; Sigma) was added to the culture medium after the 24-h viral transduction step, and fresh aliquots of the drug were provided daily in fresh medium. BAPTA (5 µM) and the water-soluble form of KN-93 (1 µM) (Calbiochem, La Jolla, CA) were added to the culture medium for 1624 h before assay.
Real-time RT-PCR. Total RNA was extracted (Nucleospin RNAII kit; BD Biosciences) from G0 and G1/S stage MOVAS cells. Known amounts of DNase-treated total RNA were used to amplify mouse PMCA1 and PMCA4 in separate real-time PCR assays (SYBR Green kit; Applied Biosystems, Foster City, CA). Reaction conditions of each primer set were optimized to generate a single PCR product of expected size and melting temperature, and relative standard calibration curves were generated for each gene as per Protocol 4304965 (Applied Biosystems, Foster City, CA) and used to quantify the mRNA level of the PMCA4 and PMCA1 genes in MOVAS cells. Gene-specific real-time PCR primers employed were as follows: mouse PMCA1: 5'-TTA-GTC-TGG-GAA-GCA-TTA-CAA-GAT-GTC-AC-3', 5'-CTT-CTT-CCC-CAA-CAG-AAA-CTT-CTC-C-3'; mouse PMCA4: 5'-ACG-TCT-TCC-CAC-CCA-AGG-TTC-3', 5'-CCA-GCA-GCC-CAC-ACT-CTG-TC-3'.
Western blot. G0- and G1/S-synchronized MOVAS cells were swollen in hypotonic buffer (10 mM Tris, pH 8.0; 1 mM DTT; 1 mM PMSF), lysed with 20 strokes of the loose pestle in a Dounce homogenizer, and spun at 900 g for 10 min, and the supernatant was spun again at 120,000 g for 1 h. The resulting microsomal pellet was resuspended in 10 mM MOPS, pH 7.4, and equivalent amounts (50 µg) of each sample were used for Western blotting with a monoclonal anti-PMCA Ab (5F10; 1:1,000 dilution; Sigma) and an anti-Actin Ab (Sigma Cat.# A-4700; 1: 500 dilution). Antibody binding was visualized with an HRP-conjugated anti-mouse IgG secondary Ab followed by DAB staining (Vector Elite kit; Vector Labs Canada, Burlington, ON, Canada). The processed blot was scanned with a densitometer, and band intensities were quantified using NIH Image software and normalized as described above.
Statistical analysis. Flow cytometry, 45Ca efflux
rates, Ca2+ imaging data, luciferase assays, and
quantitative real time RT-PCR data are shown as means ± SE and
represent results from at least two separate experiments (n 2).
Flow cytometry data were log-transformed. Analysis of variance, comparison of
proportions, and the Student's t-test were employed for statistical
comparisons. Statistical significance was defined as P
0.05.
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RESULTS |
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S-phase entry of MOVAS. Flow cytometry of BrdU-labeled MOVAS cells
was conducted to assess our ability to growth arrest and synchronize entry
into S phase. Serum deprivation for 48 h forced 63 ± 4% of cells into
G0 phase, and only 16 ± 1% were in S phase under these
conditions (n = 4 experiments; G0-0 h vs. S-0 h;
P < 0.0001). The presence of some residual S-phase cells in the
putative G0 samples is consistent with the known difficulty of
perfectly synchronizing cells immortalized with large T antigen
(32). However, upon serum
stimulation, the proportion of cells in S phase increased steadily to reach a
peak of 60 ± 4% after 16 h (n = 4; S-0 h vs. S-16 h;
P < 0.0001), whereas the proportion of G0 cells
decreased to 34 ± 4% (n = 4; G0-0 h vs.
G0-16 h; P = 0.0038). Hence, most MOVAS cells enter S
phase at 16 h post-serum stimulation.
Intracellular Ca2+ and Ca2+ efflux. The Ca2+ efflux rate from MOVAS cells progressing through the cell cycle was assayed, and the contribution to this rate by PMCA activity was quantified. Inhibiting PMCA activity with La3+ yielded a 40% reduction in the total rate of Ca2+ efflux at G0 (Fig. 2A). The total Ca2+ efflux rate also dropped 50% as cell cycle-synchronized MOVAS cells moved from G0 to G1/S (P < 0.005), at which point further reductions with La3+ were not significant (Fig. 2A). These data demonstrate a decrease in La3+-sensitive 45Ca efflux activity during MOVAS cell cycle progression, a result consistent with previous observations in other cell lines (3, 22, 23).
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Ratiometric fluorescence imaging was used to measure [Ca2+]i in MOVAS cells during G0-to-G1/S cell cycle progression. In nontransduced and untreated MOVAS, [Ca2+]i increased from 114 ± 5 nM at G0 to 192 ± 6 nM at G1/S (P < 0.001). Control transductions with empty virions or virions encoding EGFP reduced the G1/S [Ca2+]i to 118 ± 4 nM. However, retroviral transduction with CAIN or VIVIT (Fig. 3) caused significant further reductions in [Ca2+]i: [G1/S [Ca2+]i with CAIN = 75 ± 8 nM (P < 0.001) and with VIVIT = 95 ± 5 nM (P < 0.001)] compared with control-transduced cells. Similarly, transduction with Dox-inducible ANTI-MYB reduced G1/S [Ca2+]i level to 75 ± 3 nM (P < 0.001) upon addition of Dox (Fig. 3), whereas addition of Dox had no effect on [Ca2+]i of nontransduced cells (G0 + Dox = 113 ± 11 nM, and G1/S + Dox = 195 ± 22 nM) or the [Ca2+]i of control-transduced cells (data not shown). Thus inhibitors of calcineurin, NFAT, or c-Myb significantly reduced G1/S-associated [Ca2+]i. Of note, treatment of nontransduced MOVAS with BAPTA or KN-93 also significantly reduced cytosolic Ca2+ levels at G1/S (Fig. 3). These data demonstrate a cell cycle-dependent increase in the [Ca2+]i of MOVAS that can be blocked by peptide inhibitors of calcineurin and NFAT, a Ca2+-chelating agent, an inhibitor of CaMK-II, and the inducible expression of a neutralizing antibody to c-Myb.
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Inhibiting calcineurin activity with CAIN, VIVIT, and BAPTA. To monitor calcineurin-mediated transcriptional activity, we employed a NFAT promoter-luciferase reporter construct (NFAT-luc; see MATERIALS AND METHODS and Fig. 4A). Transient transfection assays showed that calcineurin activity rises approximately fivefold as MOVAS cells progress from G0 to the G1/S transition (Fig. 4B). Both CAIN and VIVIT encoding retroviral transduction suppressed this rise in calcineurin/NFAT activity, as did treatment with BAPTA (Fig. 4B). Of note, the CAIN- and VIVIT-mediated reductions in NFAT-dependent luciferase activity at G0, a time point where CAIN and VIVIT had no effect on MOVAS [Ca2+]i, suggest that these agents were acting as direct inhibitors of calcineurin/NFAT and not via indirect effects on [Ca2+]i. By contrast, the reduction in calcineurin/NFAT-responsive reporter activity noted with BAPTA is likely dependent on its ability to chelate and lower [Ca2+]i.
Inhibiting calcineurin and CaMK-II activity with KN-93. The effect
of the CaMK-II inhibitor KN-93 on calcineurin/NFAT-responsive reporter
activity (Fig. 4B) may
be due to both indirect effects on [Ca2+]i
and direct effects on reduced CaMK-II-dependent calcineurin activation.
Indeed, KN-93 treatment reduced Ca2+-dependent CaMK-II
activity by 33% at G0 (1.00 ± 0.04 vs. 0.67 ± 0.11;
P < 0.05) and 52% at G1/S (1.19 ± 0.03 vs. 0.57
± 0.05; P < 0.01) (Fig.
5) at a drug concentration and time of exposure identical to those
used to inhibit repression of PMCA4 expression
(Fig. 6). While total (i.e.,
Ca2+-dependent and
Ca2+-independent) CaMK-II activity in MOVAS increased by
20% from G0 to G1/S, EGTA (2 mM)-defined
Ca2+-independent CaMK-II activity [a measure of
Ca2+/calmodulin-mediated CaMK-II autophosphorylation
(20,
21)] increased eightfold
(Fig. 5). These data suggest
significant cell cycle-dependent activation of CaMK-II that mediates, at least
in part, activation of the calcineurin/NFAT pathway and repression of PMCA4
mRNA expression in MOVAS.
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PMCA expression. PMCA4 mRNA levels were measured with quantitative
real-time RT-PCR in cell cycle-synchronized MOVAS
(Fig. 6). Control reactions in
which the reverse transcriptase had been omitted did not produce PCR products
(data not shown), confirming that RNA samples were free of genomic DNA
contamination. PMCA4 mRNA levels decreased by 40% as cells progressed
from G0 to G1/S (Fig.
6; G1/S value = 0.62 ± 0.01; P <
0.001 vs. nontransduced G0 cells). However, expression of CAIN
(G1/S value = 0.66 ± 0.01) or VIVIT (G1/S value =
0.59 ± 0.01) had no significant effect on the
G1/S-associated repression of PMCA4 expression (P = NS vs.
empty vector). Although BAPTA had no significant effect on PMCA4 expression
levels at G1/S, the CaMK-II inhibitor KN93 did appear to increase
the expression of PMCA4 at this point in the cell cycle (0.87 ± 0.01;
P < 0.001 vs. nontransduced cells). Together, these data suggest
that calcineurin does not play a role in repressing cell cycle-dependent PMCA4
expression but that other targets of CaMK-II may play a role.
To exclude possible calcineurin-mediated effects on PMCA1, real-time RT-PCR was also performed for this pump. G0-normalized G1/S mRNA levels of PMCA1 were not significantly de-repressed with CAIN (0.82 ± 0.01; P = 0.17 vs. empty vector) or VIVIT (0.85 ± 0.03; P = 0.16 vs. empty vector), although the effect of these inhibitors was not as neutral on PMCA1 as it was on PMCA4.
The lack of a mouse-specific anti-PMCA4 antibody restricted our ability to specifically measure PMCA4 protein in the MOVAS cell cycle. However, Western blots with the isoform and species-nonspecific anti-PMCA antibody 5F10 detected a mean 40% decrease in the density of both putative PMCA1 and PMCA4 protein bands (as defined by molecular weight and relative abundance) at G1/S vs. G0 (Fig. 2B). These results are consistent with previous data in rat VSMC (23) and suggest that the transcriptional regulation of PMCA4 is translated into an effect at the protein level.
c-Myb activity and effects of ANTI-MYB on PMCA4 expression. Having previously demonstrated cell cycle-associated repression of PMCA mRNA expression in rat VSMC (23) and c-Myb-mediated repression of the mouse PMCA1 promoter (1), we sought to examine whether the observed repression of PMCA4 expression in MOVAS was also c-Myb-dependent. Using a c-Myb responsive reporter (mim-1-luc), we found that c-Myb-dependent reporter activity increased three- to sixfold as MOVAS moved from G0 to G1/S, which is consistent with data in other VSMC (22, 40, 41). Transduction with a Dox-inducible ANTI-MYB followed by transfection with a mouse PMCA1 promoter-luciferase reporter (1) revealed a G1/S-specific repression of PMCA1 promoter activity (G0 = 1.0 ± 0.15 vs. G1/S = 0.49 ± 0.01; P < 0.05). Upon Dox-mediated induction of the anti-Myb antibody, the PMCA1 promoter was significantly de-repressed (G1/S without Dox = 0.49 ± 0.01 vs. G1/S + Dox = 0.94 ± 0.02; P < 0.001). We then explored the effect of ANTI-MYB on PMCA4 levels in proliferating MOVAS cells. Expression of the ANTI-MYB construct was observed to alleviate the G1/S-specific repression of PMCA4 (Fig. 6, G1/S: ANTI-MYB + Dox vs. ANTI-MYB without Dox = 0.85 ± 0.01 vs. 0.67 ± 0.01; P < 0.01).
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DISCUSSION |
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Both c-Myb and calcineurin influence Ca2+-mediated
signals of cell cycle (1,
22,
47,
48). A marked rise in
[Ca2+]i is critically required for the
G1/S transition of VSMC and is brought about by an increase in
Ca2+ influx and a c-Myb-dependent reduction in
Ca2+ efflux mediated via repression of PMCA1
(1,
22,
23,
40,
41). Indeed, 60% of the
c-Myb effect on VSMC proliferation is exerted via PMCA1 repression
(22). However, PMCA4 also
appeared to be regulated during the cell cycle progression of rat VSMC
(23), and overexpression of
PMCA4 had been shown to induce terminal differentiation of skeletal myoblasts
(19). Given that PMCA4
expression levels in mouse neurons were repressed via calcineurin
(18), we sought to examine
whether this pathway participated in the cell cycle-regulated
[Ca2+]i of mouse VSMC.
Primary VSMC cultures exhibit slow growth and cannot be used beyond a limited number of passages due to senescence and phenotypic changes occurring late in culture. Moreover, the use of primary cells is labor intensive, expensive, and time consuming. Furthermore, an immortalized mouse VSMC line would allow complementary in vitro studies of mouse models of cardiovascular disease in which, for example, transgenic manipulations and/or their effects have been restricted to VSMC in vivo (26, 49). For these reasons, we developed and characterized a novel mouse VSMC cell line (MOVAS) and employed it in our study.
We showed that PMCA4 mRNA levels decreased twofold as MOVAS cells moved from G0 to the G1/S transition, and this correlated with a 50% reduction in Ca2+ efflux and a twofold increase in [Ca2+]i. We found a three- to sixfold rise in c-Myb activity at the G1/S transition of MOVAS cells and inhibiting c-Myb with the ANTI-MYB antibody de-repressed PMCA4 expression. Importantly, these data add to several lines of evidence supporting dynamic regulation of PMCA isoform expression in development, differentiation, proliferation, and in response to a variety of signaling pathways (reviewed in Ref. 43).
Our data show a fivefold rise in calcineurin activity as MOVAS progress from G0 to G1/S. Blocking the G1/S-specific increase in calcineurin activity with BAPTA, CAIN, or VIVIT reduced [Ca2+]i but did not de-repress PMCA4. By contrast, inhibiting CaMK-II (with KN-93) or c-Myb (with ANTI-MYB) decreased [Ca2+]i and also de-repressed PMCA4 at the G1/S transition. Although we cannot completely rule out a role of calcineurin in mediating some of the effect of KN-93, it appears more likely that the KN-93 mediated de-repression of PMCA4 occurs via a calcineurin-independent pathway. We speculate that calcineurin acts on regulators of cell cycle-associated [Ca2+]i other than PMCA1 or PMCA4. For example, it may modulate the expression of the sarcoendoplasmic reticulum Ca2+-ATPase-2 (SERCA2) or the inositol 1,4,5-triphosphate receptor type-1 (IP3R1). Indeed, calcineurin is known to increase transcription of IP3R1 in neurons (14, 16), and calcineurin has been shown to control calcium homeostasis in yeast by repressing SERCA activity (45). Further studies are needed to dissect the role of calcineurin in regulating cell cycle-associated [Ca2+]i in VSMC.
A particularly novel finding of our study is the potentially important role played by CaMK-II in cell cycle-dependent regulation of [Ca2+]i and PMCA4 expression. Indeed, the eightfold increase in Ca2+-independent CaMK-II activity at G1/S, a measure of Ca2+/calmodulin-induced constitutive CaMK-II activity (20, 21), suggests significant cell cycle-dependent activation of CaMK-II. Further studies are needed to address the mechanisms through which CaMK-II subsequently mediates its effects on PMCA4 expression.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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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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