1 Laboratoire de Physiologie et Physiopathologie de la Signalisation Cellulaire, CNRS UMR 5543, Université de Bordeaux 2, 33076 Cedex Bordeaux; and 2 Laboratoire d'Immunologie et Cytométrie, Institut Bergonié, 33000 Bordeaux, France
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ABSTRACT |
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Previously, we showed that the peak density of the transient outward K+ current (Ito) expressed in GH3 cells was different in the S phase than in other phases of the cell cycle. Using cell synchronization, we show here that Ito drops precisely at the quiescent (G0 phase)/proliferating transition. This change is not due to a modification in the voltage dependence of Ito, but rather to a modification in its inactivation kinetics. Molecular determination of K+ channel subunits showed that Ito required the expression of Kv1.4, Kv4.1, and Kv4.3. We found that the increase in Ito density during the quiescent state was accompanied by an increase in Kv1.4 protein expression, whereas Kv4.3 expression remained unchanged. We further demonstrate that the link between Ito expression and cell proliferation is not mediated by variations in cell excitability. These results provide new evidence for the cell cycle dependence of Ito expression, which could be relevant in understanding the mechanisms leading to pituitary adenomas.
Kv1.4; cell growth; excitable cell; transient outward K+ current
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INTRODUCTION |
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A GREAT NUMBER OF STUDIES have suggested that K+ channels are required for cell proliferation. On the one hand, authors have shown that K+ channel blockers inhibit the proliferation of various cell types (40, 51, 60). This inhibition was sometimes associated with a blockade of the cell cycle in specific phases (57, 64). On the other hand, it has been observed that the expression and/or activity of K+ channels varies according to the cell cycle phases (2, 9, 14, 35), suggesting that transient changes in K+ channel activity play a key role in the transition from the quiescent state (G0 phase) or the early G1 phase to the DNA replication phase (S). It is noteworthy that most of these studies have been performed in nonexcitable cells.
Pituitary cells are excitable because they generate spontaneous action potentials. The GH3 pituitary cell line consists of mammosomatotroph cells secreting both prolactin and growth hormone. In the basal state, pituitary cells are most often quiescent. However, during postnatal life, pituitary cells and, more particularly, the lactotroph cell population, are subject to proliferation. Lactotroph hyperplasia is known to occur during pregnancy and lactation in humans (15, 48) and rats (45). In humans, pathological growth of lactotrophs gives rise to prolactinomas, one of the most frequent types of pituitary tumors (7). Lactotrophs thus represent a relevant model for investigating growth regulatory mechanisms at the pituitary level.
Using the GH3 cell line as a model, we have previously shown that
1) tetraethylammonium (TEA), a broad-spectrum K+
channel blocker, inhibits GH3 cell proliferation by inducing a cell
cycle arrest at the G1/S transition (55); and
2) the expression of the transient outward K+
current (Ito) varies according to the cell cycle
phase (12). Thus, by combining electrophysiology and the
incorporation of bromodeoxyuridine (BrdU), we demonstrated that the
peak current density of Ito was lower during the
S phase (BrdU+) than during the other cell cycle phases (BrdU)
(G0, G1, G2, and M). These studies
were the first to suggest a role for K+ channels in the
proliferation of excitable cells.
The aims of the present study were to 1) determine the precise phase in which the cell cycle alterations of Ito expression occurred in GH3 cells, 2) identify the K+ channel subunits underlying the observed increase in the functional expression of Ito, and 3) investigate the mechanisms responsible for this modification of Ito expression.
We show that Ito expression is upregulated in
quiescent cells (G0 phase) compared with proliferating
cells. We also show that the Kv1.4 and Kv4 -subunits are responsible
for Ito in GH3 cells and that the increase in
Ito expression during the quiescent state is
accompanied by an increase in the Kv1.4
-subunit at the protein level. Putative links between K+ channel expression and the
proliferation process are discussed.
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MATERIALS AND METHODS |
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Cell culture. GH3 cells were cultured in Dulbecco's modified Eagle's medium (DMEM)/F12 (50/50) (Seromed, Strasbourg, France), containing 2 mM L-glutamine and 1 mM sodium pyruvate, supplemented with 15% heat-inactivated horse serum (Eurobio, Les Ullis, France) and 2.5% fetal bovine serum (Seromed). The cells were routinely grown as stocks in 75-cm2 flasks (Nunc, Polylabo, Strasbourg , France) at 37°C in a humidified atmosphere (95% air-5% CO2). The medium was changed twice a week, and the cells were trypsinized every 8-10 days.
For flow cytometry experiments, 6 × 105 cells were grown in 75-cm2 flasks. For electrophysiological recordings, 15 × 103 cells were seeded on glass coverslips pretreated with polyornithine (5 g/l). For Western blot analysis, cells were subcultured in 10-cm petri dishes and plated at 6 × 105 cells per dish, and four dishes were treated for each condition. No antibiotics were added to the cultures.Cell cycle synchronization. Cells in the G0 phase (quiescent state) were obtained by serum deprivation, commonly used to synchronize cell lines in this phase of the cell cycle (13, 30, 37). Twenty-four hours after plating, the serum-supplemented medium (SSM) was removed and the cells were rinsed twice and incubated with serum-free DMEM/F12 for 5 days. To check whether cells were able to resume proliferation and test the reversibility of the effects induced by serum deprivation, serum was added to the culture medium again for at least 20 h after the starvation period.
Cells in the G1 phase were obtained by chemical synchronization using lovastatin (21, 29, 62). Because GH3 cells proliferate asynchronously under standard culture conditions, presynchronization by serum deprivation was chosen to minimize the incubation time with lovastatin. Twenty-four hours after plating, cells were cultured in serum-free medium for 4 days. After this starvation period, serum was added to the culture medium in the presence of lovastatin (10 µM) for 20 h. The efficacy of these protocols was checked by DNA content measurement using flow cytometry.DNA content measurement by flow cytometry. For DNA content measurement, cells were stained as described by Vindelov et al. (56). Briefly, cells were incubated in PBS containing trypsin (30 µg/ml) and Nonidet P40 (0.1%) for 10 min at room temperature. After treatment with a trypsin inhibitor (0.5 mg/ml) and ribonuclease A (0.1 mg/ml) for 10 min, nuclear DNA was stained with propidium iodide (0.4 mg/ml) for 10 min. The stained samples were passed through a 48-µm nylon mesh before flow cytometric analysis. Cellular DNA content was measured with a FACScan (Becton Dickinson, San Jose, CA) equipped with a 488-nm, 15-mW argon ion laser and a double discrimination module to remove G2 doublets. Instrument performance was checked using DNA-QC particles from Becton Dickinson. Twenty thousand nuclei were acquired at a rate of 100-150 nuclei/s. Forward and side light scatter, as well as width and area of linear red fluorescence, were recorded in the list mode file. Forward and side light scatter were plotted as bivariant parameters to exclude debris. The width and area of linear red fluorescence were plotted to remove clumped nuclei from the analysis. Cell populations of interest were selected by gating, and DNA histograms of these cell populations were plotted. DNA content was proportional to the pulse area of red fluorescence. The fraction of cells in G0/G1, S, and G2-M phases was evaluated by means of ModFIT LT software (Verity Software House, Topsham, ME).
Apoptosis.
Apoptotic cells were identified in proliferating and quiescent cell
populations using two complementary fluorescent stainings. Depolarization of the mitochondrial potential (m), an early event
in the apoptotic mechanism, was visualized by
tetramethylrhodamine methyl ester (TMRM) dye (Sigma, France)
(27), and the apoptotic nuclei were observed after
Hoescht 33342 incorporation.
RT-PCR.
RNAs were prepared from GH3 cells and rat heart by acid guanidium
thiocyanate-phenol-chloroform extraction, as described by Chomczynski
and Sacchi (10). Total RNA (10 µg) was reverse
transcribed using random hexanucleotides (0.5 µl, 0.1 µg/µl) and
Superscript II reverse transcriptase (2 µl, 200 U/µl) (Invitrogen,
Cergy-Pontoise, France). The cDNAs obtained were amplified by hot-start
PCR. The PCR was performed in a final volume of 20 µl containing 500 ng cDNA, 200 µM of the four deoxyribonucleotides, 2 mM MgCl2, 2 µl 10 × PCR buffer, and 1.25 U Taq DNA polymerase
(Invitrogen). The mix was heated to 94°C for 1 min to seal the tube
with paraffin and then kept at 4°C for at least 2 min. Fifty
picomoles of each primer (Table 1) were
added, and the PCR was run for 35 cycles (94°C for 1 min, 58°C for
1 min, and 72°C for 45 s), followed by a final extension step at
72°C for 2 min (Mastercycler Personal, Eppendorf, Hambourg, Germany).
PCR product (10 µl) was mixed with loading buffer and run on a 2%
agarose gel electrophoresis stained with ethidium bromide. All RT-PCR
were performed at least twice. Each PCR included a negative control
consisting of an RT product in which reverse transcriptase was omitted
(RT).
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Western blot.
Cell homogenates (50 µg protein/lane), obtained as previously
described (5), were separated on 8% (for Kv1.4) or 10%
(for Kv4.3 and Kv1.5) SDS-PAGE and transferred to polyvinylidene
difluoride membrane by semidry blotting using a Bio-Rad Transblot SD.
The membrane was blocked for 1 h 30 min at room temperature with
5% milk powder in TBS containing 0.1% (vol/vol) Tween 20 (TTBS), followed by primary antibody incubation overnight at 4°C. Kv1.4 was
detected with a mouse monoclonal anti-Kv1.4 -subunit antibody (clone
K13/31; Upstate Biotechnology, Euromedex, Mundolsheim, France) diluted
1:500. Kv4.3 and Kv1.5 were detected with rabbit anti-Kv4.3 and rabbit
anti Kv1.5
-subunits (Alomone Laboratories, Euromedex),
respectively, both diluted 1:300. The membranes were then washed, and
primary antibodies were detected using goat anti-rabbit IgG
(Transduction Laboratories, Tebu, France) and goat anti-mouse IgG
(Santa Cruz Biotechnology, Tebu, France) conjugated to horseradish peroxidase, and the bands were visualized with enhanced
chemiluminescence (Cell Signaling Technology, Ozyme, Montigny le
Bretonneux, France). The photographic film was scanned, and the bands
were quantified using NIHimage version 1.62 software.
Electrophysiology and data analysis.
The whole cell mode of the patch-clamp technique was used. Membrane
voltage or current was recorded through an Axopatch one-dimensional amplifier (Axon Instruments, Foster City, CA). Stimulus control, data
acquisition, and processing were carried out on a PC computer, fitted
with a Digidata 1320A 16-bit data acquisition system (Axon Instruments)
using pCLAMP 8 software (Axon). Seal resistances were typically in the
10-30 G range. Recordings where series resistance resulted in a
5 mV or greater error in voltage commands were discarded. Currents were
low-pass filtered at 2 kHz and digitized at 10 kHz for storage and analysis.
Statistics. The results are expressed as means ± SE. Statistical analysis was performed using ANOVA with a Fisher PLSD as posttest or Mann-Whitney or Kruskal-Wallis two-tailed tests where appropriate. Differences of P < 0.05 or 0.01 were considered significant or extremely significant, respectively.
Chemicals. Phrixotoxin 2 (PaTx2) was purchased from Alomone Labs (Euromedex, France), and the other chemicals were from Sigma (L'Isle d'Abeau, France) unless otherwise specified.
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RESULTS |
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In a previous report, we showed that in GH3 cells the peak density
of the transient outward K+ current
(Ito) was 33% lower in the S phase than in
other phases of the cell cycle (G0, G1,
G2/M) (12). To further analyze this phenomenon, we sought to determine the precise stage of the cell cycle
in which Ito expression changed. Given that the
transition of cells from the resting stage, G0, into
G1, or from G1 to S after stimulation by growth
factors often involves modifications in K+ conductances
(17, 60), we postulated that the decrease in Ito observed in GH3 cells occurred either when
cells left the quiescent state (G0 phase) to resume
proliferation and reenter the cell cycle or at the G1/S
transition. If this hypothesis was correct, we should observe a larger
variation in Ito between cells in the
G0 or G1 phases and proliferating cells (PC)
than that previously measured between BrdU+ and BrdU cells.
Cell cycle synchronization of GH3 cells.
To determine the effects of serum deprivation on the cell cycle, the
cellular DNA content was measured by flow cytometry analysis and the
relative percentages of cells in the G0 or G1
(2N DNA content), S (between 2N and 4N), and G2/M (4N DNA
content) stages were calculated. Flow cytometry measurements showed
that the relative percentage of cells in G0/G1
increased to 82 ± 2% after 5 days in serum-deprived cultures,
compared with 60 ± 1% in control cultures in serum-supplemented
medium (SSM) (Fig. 1; P < 0.01). Concomitantly, the relative
percentage of cells in the S phase decreased to 58% in serum-free
culture, whereas a constant percentage (~5% of the cells) showed the
4N DNA content characteristic of the G2/M phase. It is well
known that cultured cells deprived of growth factors withdraw from the
cell cycle with a 2N complement of DNA. The 2N stage into which these
noncycling cells (quiescent cells) withdraw is referred to as
G0 to distinguish it from the G1 stage of
actively cycling cells (13, 37). This cell proliferation
arrest was fully reversed by adding serum for 20 h to serum-free
medium, which resulted in a cell cycle distribution similar to that
observed in SSM (Fig. 1).
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Comparison of Ito density in cells in the
G0 or G1 phase to that in proliferating cells.
Ito was elicited by a 100 mV depolarizing step
from a holding potential of 80 mV. To prevent activation of the
delayed outward K+ current, ChTX (20 nM), apamin (100 nM),
and TEA (2 mM) were systematically added to the external recording
solution (12). Under these conditions, Ito was revealed and could be more easily
studied (Fig.
2C). Total peak current amplitudes were divided by cell capacitance to avoid differences due to cell size and thus expressed as current densities. Ito density measured in cells in the
G1 phase was not significantly different from that observed
in PC (53 ± 3 pA/pF, n = 46 in cells in
G1 phase vs. 47 ± 3 pA/pF, n = 127 in
PC, P > 0.05). In contrast, the mean
Ito density was significantly higher in cells in
G0 phase (78 ± 3 pA/pF, n = 151) than
in PC (P < 0.01). The quiescent cells that reentered
the cell cycle after exposure to serum for 20 h showed a mean
Ito density similar to that observed in PC
(43 ± 3 pA/pF, n = 33; P > 0.05)
(Fig. 2A).
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Electrophysiological characteristics of Ito in
quiescent and proliferating cells.
Differences in Ito amplitude measured in a
single-voltage test could be due to differences in the voltage
dependence of activation and/or inactivation in the two cell groups. We
thus studied the voltage activation of Ito by
measuring the current triggered by 10 mV-increment potential steps from
80 to + 60 mV and the voltage inactivation by measuring the peak
amplitude of current response evoked by a 500 ms test pulse to
20 mV,
following a 2-s conditioning prepulse to potentials from
100 mV to
10 mV (10 mV increments). No difference was observed between
quiescent and proliferating cells in the activation and inactivation
midpoint (V1/2), or in the activation or
inactivation rate (k) (Table
2). Besides voltage dependence,
inactivation kinetics could also account for differences in
Ito amplitude measured during a single voltage
test. We then investigated these parameters in both cell groups.
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Electrical activity in quiescent and proliferating cells.
It has been shown that Ito is involved in the
electrical activity of various cell types. We sought to determine
whether the increase in Ito amplitude observed
in quiescent cells affected their excitability, compared with
proliferating cells. Current-clamp recordings were performed to test
this hypothesis. No significant difference was observed between the two
cell groups in terms of membrane potential, action potential (AP)
frequency, or amplitude (Table 3). As
these results raised the issue of the role of
Ito in the electrical activity of GH3 cells, we
studied the effect of the blockade of Ito by
4-aminopyridine (4-AP) (46, 47) on the excitability of
this cell type. When applied to GH3 cells, 4-AP reduced the peak
current in a dose-dependent manner. The IC50 approximated
0.1 mM, and 0.5 mM 4-AP was sufficient to completely block
Ito (Fig.
3A). However, modifications in
the electrical activity were only observed for 4-AP concentrations 1
mM, whereas application of 0.5 mM 4-AP had no significant effect (Fig.
3B, Table 3).
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Molecular basis of Ito in GH3 cells.
One of the main issues to be solved in understanding the modification
of Ito expression during the
quiescent/proliferating transition is to determine the molecular basis
of this current in GH3 cells. We used two complementary approaches. The
first approach was to measure the kinetics of recovery from
inactivation of Ito by electrophysiology. These
kinetics are known to differ markedly according to the K+
channel -subunits responsible for the current. The second approach was to use RT-PCR for a direct identification of the principal K+ channel
-subunits mRNA expressed in these cells,
which are likely to account for Ito.
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Involvement of Kv4.3 K+ channels in Ito expression in quiescent and proliferating cells. Until now, no pharmacological tools were available to distinguish between Kv1.4 and Kv4 K+ channels. Only recently, Diochot et al. (16) have isolated toxins, so-called phrixotoxins (PaTx), from a spider venom that specifically block the Kv4.2 and Kv4.3 K+ channels. Thus we have tested the effect of the PaTx2 on the expression of Ito in GH3 cells.
Ito evoked by a depolarizing pulse from
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Expression of Kv channel proteins in quiescent and proliferating cells. Among the various mechanisms that could account for the increase in Ito density observed in quiescent cells, we studied the quantitative expression of the K+ channel subunits in both cell groups by Western blot. Our study focused on Kv1.4 and Kv4.3 channel expression, because antibodies for Kv4.1 are not yet available.
The results in Fig. 6A show that anti-Kv1.4 recognizes a main band at ~96 kDa and sometimes a faint band at ~80 kDa. These two bands have been previously described in GH3 cells, and it has been shown that the difference between the two molecular masses was due to glycosylation of the channel proteins (53). The analysis of the Western blot profile shows that the expression of Kv1.4 increased significantly in quiescent cells (209 ± 29% that of proliferating cells, n = 5, P < 0.01). This phenomenon could be reversed, because serum addition for 20 h induced Kv1.4 expression to return to the level observed in cells cultured in SSM (Fig. 6). The expression of the Kv4.3 channel subunit, visualized as a single band of ~70 kDa revealed with anti Kv4.3 antibody, remained similar under the three different conditions (Fig. 6). Kv1.4 can be expressed as a homomeric or as heteromeric channel. In GH3 cells, it has been suggested that Kv1.4 was essentially expressed as a heteromeric channel with Kv1.5 subunits (33, 53). We thus studied the expression of Kv1.5
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DISCUSSION |
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Until now, the involvement of K+ channels in
controlling cell proliferation had been mainly studied in unexcitable
cell types (32, 41, 59). Using the excitable GH3 cell
line, we had previously shown that the peak current density of
Ito was lower in bromodeoxyuridine-positive
cells (i.e., DNA-synthesizing cells or S phase cells) than in
BrDU cells (non-S phase cells, i.e., cells in
G0, G1, G2, or M)
(12). However, this previous study did not enable us to
determine the specific stage in the cycle when this modification took
place. Because the G0/G1 and G1/S
transitions are the major checkpoints in the cell cycle
(67) and modifications in K+ channel
expression often occur at these stages, we compared
Ito expression and characteristics in GH3 cells
synchronized in G0 (by serum deprivation) or G1
(by treatment with lovastatin) with those of a proliferating cell
population. We found that Ito peak density
measured in quiescent cells (G0) was 166% that of
proliferating cells, whereas no change was observed in cells
synchronized in G1. Ito peak density
dropped back to values observed in proliferating cells when serum was
added to the medium again, i.e., as soon as cells reentered the cell
cycle. These results suggest that a decrease in
Ito expression occurs at the
G0/G1 (quiescent/proliferating) transition.
Studies performed in nonexcitable cells have shown that mitogens generally stimulate the activity level of K+ channels (20, 34, 41). Growth factors have also been shown to have a modulating effect, usually inhibitory, on Ito in excitable cells. For example, Ito expression in myocytes is downregulated by nerve growth factor (25) or paracrine hypertrophic factor (24). The glial cell-derived neurotrophic factor (GDNF) inhibits IA (Ito-like) in midbrain neurons in culture (66). The reduced expression of Ito observed in GH3 cells after serum addition to the medium may thus result from a direct action of the numerous growth factors contained in the serum. These growth factors could act via proteins involved in cell cycle progression, as was the case with mitosis-promoting factor (MPF), which was found to modulate R-eag K+ channel expression in Xenopus oocytes (6). Alternatively, reorganization of the cytoskeleton occurring during cell cycle progression could also modify current amplitude (8, 54).
A central point in understanding the mechanisms of variations in
Ito expression is the elucidation of the
molecular basis of this current in GH3 cells. Because the kinetics of
recovery from inactivation are specific to the K+ channel
subunit responsible for the current, we determined the recovery
kinetics from inactivation of Ito. Our data
revealed two kinetically distinct components, suggesting that at least two different K+ channel subunits are responsible for
Ito in GH3 cells. The fast component had very
similar recovery kinetics to those of shal-related channels
(i.e., Kv4) expressed in mammalian cells (58) whereas the
slow component value was compatible with the expression of the Kv 1.4 K+ channel subunit (44). These kinetic
constants are on the same order of magnitude in quiescent and
proliferating cells, suggesting that the molecular composition of
Ito is the same in both populations. These data
were corroborated by the results of RT-PCR experiments, which showed
the expression of Kv1.4, Kv4.1, and Kv4.3 -subunits, all known to
induce Ito-type K+ currents after
heterologous expression (49, 50, 52). Moreover, the use of
PaTx2, a specific blocker of Kv4.2 and Kv4.3 K+ channels
(16), confirms for the first time the functional
expression of Kv4.3 in GH3 cells. Thus Ito
probably results from the expression of at least Kv1.4, Kv4.1, and
Kv4.3 K+ channel
-subunits.
The regulation mechanism(s) of K+ channel activity during the cell cycle have yet to be determined. It is not known, for instance, whether K+ channel activity is modulated via control of K+ channel expression or modulation of the activity of existing K+ channels. Electrophysiological experiments performed in presence of PaTx2 showed that the difference in Ito peak density between quiescent and proliferating cells was maintained, suggesting that Kv4.3 K+ channels were not involved in the increase in Ito observed in quiescent cells. This result was confirmed by Western blot experiments, which showed no modification of the expression level of Kv4.3 protein in both cell populations, whereas the expression level of Kv1.4 protein in quiescent cells was twice that in proliferating cells. It is thus likely that the increase in Ito expression results from an increase in Kv1.4 subunit expression level, leading to the insertion of newly functional Kv1.4 K+ channels into the cell membrane.
K+ channels can consist of homomeric or heteromeric
combinations of subunits encoded by distinct, yet closely related
genes. It has been suggested that, in GH3 cells, Kv1.4 could combine with Kv1.5 subunits to produce an inactivating A-type K+
channel (33, 53). We thus investigated Kv1.5 expression by Western blot and found that it was reduced by half in quiescent compared with proliferating cells. It is thus possible that some of the
Kv1.4 -subunits are expressed in homomeric form in quiescent cells,
whereas combined Kv1.4/Kv1.5 subunits are predominant in proliferating
cells. This hypothesis is consistent with the fact that the time
constant of Ito inactivation (
1)
is faster in quiescent than proliferating cells, as would be expected
from the expression of homomeric Kv1.4 channels. These data also
suggest that Kv1.4 and Kv1.5 are regulated in opposite ways during the
cell cycle. A similar inverse regulation of Kv1.4 and Kv1.5 subunits
has also been described in hypertrophic cardiomyocytes
(36).
Besides an increase in Kv1.4 channel expression levels, several other
mechanisms may be involved in regulating Ito
between the quiescent and proliferating states. First, a
posttranslational modification by kinases could regulate K+
channel activity (26, 42, 63). However, various kinases (tyrosine kinases, phosphatidylinositol 3-kinase, and protein kinase A
and C) seem unable to regulate Ito in GH3 cells
(personal observations, data not shown). Second, various auxiliary
subunits (Kv, KCHaP, and Kchip) could heteromerize with Kv
-subunits, thereby modifying the amplitude, gating, and/or
expression of K+ currents (31, 43, 65).
The precise nature of the link between K+ channel activity and cell cycle progression has not yet been elucidated. The main mechanism evoked to account for the role of K+ channels in the proliferation of nonexcitable cells is the regulation of the membrane potential. Membrane hyperpolarization resulting from increased K+ channel activity would interfere directly with mitogenic activity by increasing the driving force for electrogenic entry of Ca2+ ions, a condition necessary for cell cycle progression (3, 61).
Ito is involved in the excitability of myocytes
(19) and neurons (11), in which it modulates
interspike latency and action potential repolarization (22,
28). A modification in cell excitability caused by variations in
Ito amplitude could, therefore, constitute a
messenger for cell growth (38, 66). Our data show that the
electrophysiological parameters, including membrane resting potential,
AP frequency, and AP amplitude, were similar, irrespective of the
proliferation state of the cells. It is particularly interesting that
complete inhibition of Ito by 0.5 mM 4-AP had no
significant effect on membrane excitability, because the acceleration of AP firing rate at concentrations 0.5 mM 4-AP probably results from
the inhibition of 4-AP-sensitive K+ currents other than
Ito (23).
Ito is thus unlikely to play a significant, if
any, role in regulating GH3 cell excitability, and the link between
Ito expression and cell cycle progression is
still open to question. Besides the well-known role of
Ito and, more particularly, of Kv1.4 in cell
excitability, it has been recently described that this channel could
play a role in the proliferation process of glial cells (1,
18). An Ito-like current has been
described in differentiated astrocytes but not in glioma cells. The
disappearance of this current seems to be an early feature accompanying
the transformation of a normal astrocyte into a tumor glial cell
(4). The Kv1.4 channel subunit has also been found to be
upregulated in oligodendrocytes induced to proliferate after chronic
spinal cord injury (18). However, the action mechanism of
Kv1.4 in these proliferation processes could not be assessed because no
selective pharmacological agents are available.
In conclusion, we show here in a pituitary cell model that the expression of the transient outward K+ current is cell cycle dependent. We found that upregulation of Ito density in quiescent cells was probably due to a selective increase in Kv1.4 channel subunit expression at the protein level. Moreover, we demonstrate that the link between Ito and cell proliferation is apparently not mediated by variations in cell excitability. The etiology of pituitary adenomas is imperfectly known at the present time. Thus these findings are particularly relevant in understanding proliferative mechanisms of pituitary cells in a physiological context (lactation) or during tumorigenesis.
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ACKNOWLEDGEMENTS |
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We thank Marie-Claude Audy for help in setting up the Western blot technique in our laboratory and François Ichas for advice concerning apoptosis evaluation. We are also very grateful to Bernard Dufy for comments on an earlier draft of this manuscript.
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FOOTNOTES |
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This work was supported by Centre National de la Recherche Scientifique, University of Bordeaux 2, and Etablissement Public Régional (Aquitaine Région). A. Czarnecki was financed by Association pour la Recherche contre le Cancer.
Address for reprint requests and other correspondence: L. Bresson-Bepoldin, Laboratoire de Physiologie et Physiopathologie de la Signalisation Cellulaire, CNRS UMR 5543, Université de Bordeaux 2, 146 rue Léo Saignat, 33076 Bordeaux cedex, France (E-mail: laurence.bepoldin{at}umr5543.u-bordeaux2.fr).
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.
10.1152/ajpcell.00446.2002
Received 26 September 2002; accepted in final form 16 December 2002.
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