Potassium channel expression level is dependent on the proliferation state in the GH3 pituitary cell line

Antonny Czarnecki1, Luce Dufy-Barbe1, Sylvie Huet2, Marie-Françoise Odessa1, and Laurence Bresson-Bepoldin1

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


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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 alpha -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 alpha -subunit at the protein level. Putative links between K+ channel expression and the proliferation process are discussed.


    MATERIALS AND METHODS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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 (Delta Psi 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.

The percentage of apoptotic cells was assessed as (the number of cells exhibiting apoptotic nuclei and/or Delta Psi m depolarization)/(the total number of cells) × 100. Over 600 cells were counted in at least 3 independent experiments.

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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Table 1.   Oligonucleotides used for the RT-PCR

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 alpha -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 alpha -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 GOmega 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.

The standard external solution comprised (in mM) 140 NaCl, 5 KCl, 2 CaCl2, 2 MgCl2, 0.3 Na2HPO4, 0.4 KH2PO4, 4 NaHCO3, 5 glucose, and 10 HEPES; osmolarity: 300-310 mosmol/kgH20, pH 7.3. To study the transient outward K+ current, charybdotoxin (ChTX; 20 nM), tetrodotoxin (TTX; 0.5 µM), apamin (100 nM), and 2 mM TEA were added systematically. The recording pipette was filled with a solution containing (in mM) 150 K-gluconate, 2 MgCl2, 1.1 EGTA, and 5 HEPES. GTP (40 µM) and ATP-Mg (2 mM) were added systematically; pH 7.3; osmolarity: 290 mosmol/kgH2O. Pipette resistance was 2-4 MOmega . All experiments were performed at 37 ± 1°C.

Normalized peak current values (I/Imax) and normalized Ito conductance (G/Gmax) were plotted vs. membrane potential for individual cells, and the resulting inactivation and activation curves were fitted to the Boltzmann equation: I/Imax = [1 + exp(V - V1/2)/k]-1, where V1/2 is the half-maximal activation or inactivation voltage and k is the slope factor. The conductance underlying Ito (G) was calculated as G = I/(V - Vrev) (under our experimental conditions, Vrev = -79 mV). Time constants characterizing the decay of Ito components were estimated using a fitting program which provided an estimate of current amplitude (I) as a function of time (t) according to the equation: I(t) = A0 + A1 exp(-t/tau 1) + A2 exp(-t/tau 2) + ... An exp(-t/tau n), where An is the current amplitude of the time constant tau n.

The time courses of recovery from inactivation of Ito were fitted by a biexponential function (% of recovery = Afast*[1 - exp(-t/tau fast] + Aslow*[1 - exp(-t/tau slow)]), where Afast and Aslow were the amplitude of the fast and slow components of recovery, t is the time spent at the recovery potential, and tau fast and tau slow are the time constants. Microcal Origin 5.0 software was used for data fitting.

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.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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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Fig. 1.   Flow cytometry analysis of cell synchronization in the G0 and G1 phases. A: cell cycle distribution when the cells were cultured in the continuous presence of serum (+serum/PC) (PC, proliferating cells), in serum-free medium for 5 days (-serum/G0), in serum-free medium for 4 days followed by the addition of serum and lovastatin for 20 h (lovastatin/G1), or in serum free-medium for 5 days followed by the addition of serum for 20 h (20 h serum/PC20). The first peak represents the cells in G0 or G1 and the second peak the cells in G2/M. The area between the 2 peaks corresponds to the cells in the S phase. B: histogram of the percentage of cells in the 3 phases under different culture conditions (+serum, -serum, 20 h serum + lovastatin, 20 h serum). Results were obtained in 5 independent experiments. **P < 0.01.

For cell synchronization in the G1 phase, GH3 cells were cultured in serum-free medium for 5 days and then returned to SSM containing 10 µM lovastatin for 20 h. In the first step of this protocol, the cells were growth-arrested by the serum deprivation period and, in the second step, the presence of serum in the culture medium resumed cell proliferation (cells left the G0 phase and reentered the cell cycle in the G1 phase), but the presence of lovastatin stopped cell cycle progression by blocking cells in the G1 phase (21, 62). Flow cytometry measurements revealed that 86 ± 3% of the cells remained in the G1 phase, even in the presence of serum (Fig. 1).

A possible induction of apoptosis in cells cultured for 5 days in serum-free medium was also examined by counting cells exhibiting a picnotic nucleus after Hoescht staining and/or depolarization of the mitochondrial potential visualized after TMRM loading (see MATERIALS AND METHODS). There were fewer than 4% apoptotic cells in SSM (n = 900), serum-free medium (n = 1,092), or lovastatin (n = 600) (data not shown).

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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Fig. 2.   Ito expression and characteristics in proliferating and synchronized GH3 cells in the G0 and G1 phases. A: Ito peak density in proliferating (PC and PC20) and synchronized cells in the G0 and G1 phases. Ito density was measured as the total peak amplitude evoked by a 500-ms depolarizing pulse from -80 to +20 mV recorded in Hanks' balanced salt solution (HBSS) containing tetrodotoxin (TTX; 0.5 µM), charybdotoxin (ChTX; 20 nM), tetraethylammonium ester (TEA; 2 mM), and apamin (100 nM). Results are expressed as means ± SE (**P < 0.01). B: voltage-clamp protocol used to determine the voltage dependence of Ito activation. Current was activated by 10-mV incremental potential steps from holding potentials of -80 to +60 mV for 500 ms. Inset: I-V curves obtained in quiescent (G0) (; n = 11) and proliferating cells (, n = 13). C: Ito kinetics in quiescent (G0) and proliferating cells. The currents expressed in quiescent and proliferating cells have been scaled to each other and superimposed. Inset: magnification of the time to peak observed in G0 and PC.

These results confirm that the expression of Ito in GH3 cells is closely related to the cell cycle and suggest that the variations in Ito density probably occur at the quiescent/proliferating transition. We then determined which mechanisms were responsible for the Ito density increase in quiescent (G0) cells.

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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Table 2.   Electrophysiological characteristics of Ito in quiescent and proliferating cells

As previously described by Oxford and Wagoner (39), Ito inactivation proceeds as the sum of two exponentials with distinct time constants (tau 1 and tau 2). We thus studied the kinetics of Ito inactivation occurring during a 500-ms voltage step from -80 to +20 mV (Fig. 2C). Our data show that the time constant of the fast (tau 1) inactivation components was significantly faster in quiescent than proliferating cells (Table 2, Fig. 2D), whereas the slow (tau 2) was unchanged (Table. 2). Moreover, we also observed that the time to peak of Ito (tpeak) was significantly faster in quiescent than proliferating cells (Table 2).

These results suggest that the change in Ito density between quiescent and proliferating cells is probably due to a modification in the expression of the fast component of this current.

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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Table 3.   Comparison of the electrical activity characteristics of GH3 cells



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Fig. 3.   4-Aminopyridine (4-AP) blocked Ito in GH3 cells: effects on electrical activity. A: plot of average peak current ± SE as a function of 4-AP concentration (n = 4). Inset: Ito evoked by a step from -80 to +20 mV in the presence of increasing concentrations of 4-AP. B: the electrical activity of quiescent cells, recorded under current-clamp conditions (I = 0), was studied before and after addition of 4-AP (0.5 mM or 1 mM) to the recording bath.

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 alpha -subunits responsible for the current. The second approach was to use RT-PCR for a direct identification of the principal K+ channel alpha -subunits mRNA expressed in these cells, which are likely to account for Ito.

We thus measured recovery from inactivation of Ito using a double pulse protocol. Cells were depolarized from the holding potential (Vh = -80 mV) to +20 mV for 400 ms (test pulse) and then returned to -80 V for 25 ms to 5 s (interpulse interval). A second test pulse was then applied. The magnitude of Ito induced by the second pulse was expressed as a percentage of that induced by the first pulse (% recovery) and plotted as a function of the interpulse interval (Fig. 4A). Recovery in quiescent and proliferating cells was best fitted by a biexponential function showing constant reactivation times on the same order of magnitude in both cell groups (tau fast = 62 ms and 35 ms, tau slow = 1,070 ms and 1,101 ms for proliferating and quiescent cells, respectively). These results suggest that two different K+ channels are responsible for Ito in GH3 cells. We then used RT-PCR to study the expression of the four main K+ channel subunits known to form homomeric channels with Ito-like properties: Kv1.4, Kv4.1, Kv4.2, and Kv4.3. These experiments revealed that Kv1.4, Kv4.1, and Kv4.3 mRNA (Fig. 4Bb) were expressed in both quiescent and proliferating cells (data not shown). The Kv4.2 subunit mRNA detected in rat heart extracts as shown in Fig. 4Ba was not found in GH3 extracts, suggesting that it is not expressed in this cell line.


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Fig. 4.   Molecular basis of Ito in GH3 cells. A: recovery from Ito inactivation was measured using 2 depolarizing pulses to +20 mV, separated by intervals of increasing duration (from 25 to 5,000 ms) (inset). Data were normalized to maximum current value and plotted as a function of recovery time to show the recovery from inactivation of proliferating (open circle , n = 8) and quiescent (, n = 6) cells. The lines represent the best fit of a second-order exponential function for both cell populations (line, proliferating cells; dots, quiescent cells). The time constants obtained suggest the expression of Kv1.4 and Kv4 alpha -subunit channels. B: RT-PCR amplification of selected Kv alpha -subunit mRNA (see Table 1) isolated from rat heart (a) or the GH3 cell line (b). Amplified products were displayed in 2% agarose gel stained with ethidium bromide. RT+: PCR amplification of RNA reverse transcribed in the presence of superscript II. RT-: PCR amplification of RNA reverse transcribed in absence of Superscript II. Lane 1: molecular weight markers.

These data suggest that, in GH3 cells, Ito results from the expression of Kv1.4, Kv4.1, and Kv4.3 K+ channel subunits.

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 -80 to +20 mV was partly blocked by application of PaTx2. The maximal inhibition was obtained with PaTx2 at 500 nM and reached 26.3 ± 4.0% (Fig. 5, A and B), suggesting that Kv4.3 channels participate in functional Ito expression in GH3 cells. To determine whether the regulation of Ito expression observed between quiescent and proliferating cells was due to a regulation of functional Kv4.3 K+ channel expression, we have compared Ito densities in quiescent and proliferating cells in the presence of PaTx2 in the bath recording. Under these conditions, the increase in Ito density observed in quiescent cells was maintained and was very similar to that described in absence of PaTx2 (%Ito density in quiescent/proliferating cells = 164 ± 7.7% and 166 ± 3.8% in the presence or absence of PaTx2 500 nM, respectively; P > 0.05, Fig. 5C).


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Fig. 5.   Effects of phrixotoxin 2 (PaTx2) on Ito expression in GH3 cells. A: PaTx2 partly blocked Ito evoked by a depolarizing pulse from -80 to +20 mV. Inset: dose response curve for PaTx2. B, left: Ito was activated by 20 mV incremental potential steps from -80 to +60 mV for 500 ms in absence (solid line) or in the presence (dotted line) of PaTx2 (500 nM) in the bath recording. Right: difference currents, corresponding to PaTx2-sensitive current, was obtained by subtracting Ito in the presence (dotted line) and in the absence (solid line) of PaTx2 (500 nM). Inset: peak current amplitude vs. membrane potential before () and after (open circle ) PaTx2 (500 nM) application; black-triangle, PaTx2-sensitive current. C: percentage of Ito density in quiescent cells (-Se) compared with proliferating cells measured in the presence or absence of PaTx2 (500 nM).

These data show for the first time that Kv4.3 K+ channels are functional in GH3 cells and participate in Ito expression but are probably not involved in the increase in Ito density observed during in quiescent state in this cell line.

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 alpha -subunit in quiescent and proliferating GH3 cells. Our data show that, in contrast to Kv1.4, the expression of Kv1.5 visualized as a single band of 76 kDa was lower in quiescent (46 ± 8%) than proliferating cells (Fig. 6). This effect was partially reversed by the addition of serum for 20 h (Fig. 6).


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Fig. 6.   Western blot analysis of the expression of Kv1.4, Kv1.5, and Kv4.3 in quiescent and proliferating GH3 cells. A: whole cell extracts of GH3 cells (50 µg of protein/lane) cultured in the presence of serum (PC), in serum-free medium for 5 days (G0), or in serum-free medium for 5 days followed by the addition of serum for 20 h (PC20), were subjected to immunoblot analysis with anti-Kv1.4 (top), anti-Kv1.5 (middle) and anti-Kv4.3 (bottom). B: graphic representation of the relative expression of Kv1.4, Kv1.5, and Kv4.3 alpha -subunit K+ channels in proliferating (PC and PC20) and quiescent cells (G0) obtained after quantification of at least 3 experiments. Results are expressed as means ± SE (**P < 0.01).

These data confirm that Kv4.3 is probably not responsible for the increase in Ito peak density in quiescent cells and suggest that this is likely due to an increase in Kv1.4 protein expression level.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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 alpha -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 alpha -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 alpha -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 (tau 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 (Kvbeta , KCHaP, and Kchip) could heteromerize with Kv alpha -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.


    ACKNOWLEDGEMENTS

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.


    FOOTNOTES

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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