Cell cycle-related changes in transient K+ current density in the GH3 pituitary cell line

A. Czarnecki, S. Vaur, L. Dufy-Barbe, B. Dufy, and L. Bresson-Bepoldin

Laboratoire de Neurophysiologie, Centre National de la Recherche Scientifique UMR 5543, Université de Bordeaux 2, 33076 Bordeaux Cedex, France


    ABSTRACT
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Our aim was to determine whether the expression of K+ currents is related to the cell cycle in the excitable GH3 pituitary cell line. K+ currents were studied by electrophysiology, and bromodeoxyuridine (BrdU) labeling was used to compare their expression in cells thereafter identified as being in the S or non-S phase of the cell cycle. We show that the peak density of the transient outward K+ current (Ito) was 33% lower in cells in S phase (BrdU+) than in cells in other phases of the cell cycle (BrdU-). The voltage-dependence of Ito was not modified. However, of the two kinetic components of Ito inactivation, the characteristics of the fast component differed significantly between BrdU+ and BrdU- cells. Recovery from inactivation of Ito showed biexponential and monoexponential function in BrdU- and BrdU+ cells, respectively. This suggests that the molecular basis of this current varies during the cell cycle. We further demonstrated that 4-aminopyridine, which blocks Ito, inhibited GH3 cell proliferation without altering the membrane potential. These data suggest that Ito may play a role in GH3 cell proliferation processes.

potassium current; excitable cells; cell growth


    INTRODUCTION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

A NUMBER OF RECENT STUDIES suggested that ion channels, and more particularly, K+ channels, are required for cell proliferation (9, 49). Different K+ channels have been found to be involved in the proliferation process in various cell models. Blockade of voltage-dependent K+ channels (Kv) by pharmacological agents inhibits proliferation of brown fat cells (30), myeloblastic cells (52), human melanoma cells (28, 21), prostatic cells (40), and colorectal cancer cells (53). Similarly, inhibition of ATP-sensitive or Ca2+-dependent K+ channels significantly altered the proliferation of other cell types (47, 50).

Some of these studies showed that inhibition of cell proliferation by K+ channel blockers resulted from a cell cycle block at specific phases (47, 52). More recently, the expression and/or activity of K+ channels has been found to vary according to the proliferation state of the cells or to the phases in the cell cycle. For example, in human lymphocytes, both Kv1.3 (23) and hSK4, a Ca2+/calmodulin-activated K+ channel (18), are upregulated during the proliferation process (31). Recently, Kotecha and Schlichter (19) reported that the changes in K+ currents observed during the proliferation of microglial cells corresponded to a switch in the expression of Kv1.3 and Kv1.5 channel proteins at the cell surface. In mouse oocytes, a large-conductance voltage-activated K+ channel is active throughout the M and G1 phases but inactive during the G1/S transition (8). Similarly, it seems that eag-related K+ channel regulation is cell cycle dependent in neuroblastoma cells (1). Most of those working in the field have thus suggested that these transient changes in K+ channel activity may play a key role in the transition from the quiescent state (G0) or the early G1 phase to the DNA replication (S) phase. However, despite all these studies, the link between K+ channel activity and cell cycle progression remains elusive. Among the various mechanisms proposed to account for the role of K+ channels in cell proliferation, studies have emphasized membrane hyperpolarization as an essential event required for cell cycle progression (49). According to this hypothesis, changes in membrane potential resulting from K+ channel activity directly interfere with mitogenic activity (12) and/or modify the driving force for the electrogenic transport of Ca2+ ions (4). This, in turn, would affect cell proliferation. It should be mentioned that most of these studies were performed in nonexcitable cells in which the control of membrane resting potential by K+ conductances as well as the regulation of Ca2+ influx differs drastically from that of excitable cells.

Pituitary lactotroph cells are excitable because they generate spontaneous action potentials. The ion currents expressed in this cell type have been extensively studied. However, although lactotrophs are subject to dynamic physiological (34, 38) or physiopathological (6) growth during postnatal life, the putative role of K+ channel activity in proliferation has not yet been investigated.

In recent work in the GH3 mammosomatotroph pituitary cell line, we have shown that tetraethylammonium chloride (TEA), a wide-range blocker of K+ conductances, reduced cell proliferation by inducing a cell cycle block at the G1/S transition (46). This finding suggests that K+ channels are probably involved in the proliferation of these excitable cells. The GH3 cell line expresses several K+ channels, including voltage-dependent K+ channels (29), three types of Ca2+-dependent K+ channels (10, 36), and an inward rectifying K+ channel (3). The aim of the present report was to study the expression of K+ currents in GH3 cells by using electrophysiological techniques, distinguishing between the S phase and the other cell cycle phases. We show that the density of the transient outward K+ current (Ito) drops significantly during the S phase. Moreover, inhibition of the Ito by 4-aminopyridine (4-AP) reduced cell proliferation. The involvement of this current in the electrical activity of GH3 cells was investigated and the mechanism by which Ito may play a role in cell proliferation was studied.


    MATERIALS AND METHODS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
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Cell Culture

GH3 cells were cultured in DMEM-F-12 (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 an humidified atmosphere (95% air-5% CO2). The medium was changed twice a week and the cells were passaged every 8-10 days.

For cell proliferation assays, the cells were subcultured in 24-well plates (Nunc), seeded at 6 × 104 cells per well. For electrophysiological recordings, the cells were subcultured on microgrid glass coverslips (Eppendorf, Hamburg, Germany) pretreated with polyornithine (5 g/l). They were used 3-5 days after trypsinization.

No antibiotics were added to the cultures.

Cell Proliferation Measurement

Cell proliferation was estimated by [3H]thymidine incorporation as previously described (46). Briefly, the cells were treated for 72 h with various concentrations of 4-AP. During the last 12 h of treatment, [methyl-3H]thymidine (2 µCi/ml; ICN, Orsay, France; specific activity 60 Ci/mM) was added to the culture medium. The cultures were then washed twice in fresh culture medium and pulsed for 2 h with 5 µM unlabeled thymidine. The chase medium was discarded, and the cells were lysed in 0.2 M NaOH. After neutralization with 0.2 M HCl, the lysate was transferred into vials for scintillation counting on a Beckman LS6000 SC counter.

Cytotoxicity Assays

Cell viability was determined using a Cytotoxicity Detection Kit purchased from Boehringer Mannheim (Strasbourg, France), which measures lactate dehydrogenase (LDH) activity. LDH, a stable cytosolic enzyme, is rapidly released into the culture medium after disruption of the plasma membrane. LDH activity was assessed by measuring the optical density, at 500 nm, of the cell sample medium, some of which had been treated with 4-AP.

Electrophysiology

Patch-clamp recordings. The whole cell mode of the patch technique was employed. The electrodes were pulled on an L/U-3P (List-Medical, Darmstad, Germany) puller in two stages from borosilicate glass capillaries (1.5-mm diameter; Clarke, Pangbourne Readings, UK) to a tip diameter of 1.2-2 µm. The pipette resistance was 2-4 MOmega .

Membrane voltage or current was recorded through a Biologic RK300 amplifier (Grenoble, France). Stimulus control, data acquisition, and processing were carried out with an IBM personal computer fitted with a Labmaster TL1/DMA interface (Axon Instruments, Foster City, CA) using pCLAMP 5.5.1 software (Axon Instruments). Seal resistances were typically in the range of 10-30 GOmega . Recordings for which series resistance resulted in a 5-mV or greater error in voltage commands were discarded. Currents were low-pass filtered at 1 kHz with a 5-pole Tchebicheff filter and digitized at 10 kHz for storage and analysis.

Recording solutions. 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, pH 7.3 (osmolarity 300-310 mosmol/kg). Charybdotoxin (ChTX; 20 nM), tetrodotoxin (TTX; 0.5 µM), apamin (100 nM), and TEA (2 mM) were added systematically for study of the transient outward K+ current. The recording pipette was filled with a solution containing (in mM) 150 K-gluconate, 2 MgCl2, 1.1 EGTA, and 5 HEPES, pH 7.3 (osmolarity 290 mosmol/kg). GTP (40 µM) and MgATP (2 mM) were added to the pipette solution for current-clamp experiments.

An additional pipette with a tip opening of 5-10 µm was used to apply drugs by low-pressure ejection in the neighborhood of the investigated cell. All experiments were performed at 37 ± 1°C.

Data analysis. In spontaneously active cells, the most hyperpolarized value measured during the first minute of recording was chosen as the membrane potential value.

Cell capacitance was measured by integrating the area of the capacitive transient elicited by a 10-mV hyperpolarizing step from the holding potential. These values were used as a measure of cell size, so voltage-clamp currents were expressed as densities (i.e., pA/pF).

Normalized peak current values (I/Imax) and normalized Ito conductance (G/Gmax) were plotted against membrane potential for individual cells, and the resulting inactivation and activation curves were fitted to the Boltzmann equation: I/Imax = [1 + exp(V - V[1/2])/k]-1, where V is membrane potential, V[1/2] is midpoint of activation, and k is the slope factor. The means ± SE of these curves were plotted and fitted to the Boltzmann equation. The conductance underlying Ito (G) was calculated as G = I/(V - Vrev), where the reversal potential, Vrev, was -79 mV under our experimental conditions.

Time constants characterizing the decay of Ito components were estimated with the use of a fitting program that provides 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 amplitude of current of the time constant of inactivation, tau n.

The time courses of recovery from inactivation of Ito were fitted by either a monoexponential {%recovery = A · [1 - exp(-t/tau )]} or biexponential {%recovery = Afast · [1 - exp(-t/tau fast)] + Aslow · [1 - exp(-t/tau slow)]} function, where Afast and Aslow are the amplitude of the fast and slow components of recovery, respectively, t is the time spent at the recovery potential, and tau fast and tau slow the time constants of the fast and slow components of recovery.

Microcal Origin 5.0 software was used for data fitting.

BrdU Labeling

We used BrdU labeling to distinguish proliferating from nonproliferating cells. BrdU was incorporated instead of thymidine into dividing cells during the S phase, without affecting the cell cycle progress. BrdU (10 µM) was added to the culture medium of GH3 cells 3 h before electrophysiological recordings were made. To prevent aspecific BrdU labeling, we added fluorodeoxyuridine (100 µM) to the external recording medium during the electrophysiological experimentation period, which lasted <1.5 h. At the end of the recording session, the cells were fixed in 70% ethanol diluted in 50 mM glycine buffer (pH 2) for at least 16 h. The incorporation of BrdU in cells was detected with a mouse anti-BrdU monoclonal primary antibody followed by an alkaline phosphatase-conjugated secondary antibody (Boehringer Mannheim). The recorded cells were located on the coverslips by their etched grid coordinates and could be identified as BrdU+ (S phase) or BrdU- (non-S phase).

Chemicals

ChTX was purchased from Alomone Labs (Jerusalem, Israel). Other chemicals were supplied by Sigma (L'Isle d'Abeau, Chesne, France).

Statistical Analysis

Results are expressed as means ± SE. The unpaired t-test was used for statistical comparison of mean and differences, with P < 0.05 considered significant.


    RESULTS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
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To determine the putative involvement of K+ channels in GH3 cell proliferation, we compared the expression of the outward K+ currents activated by depolarization according to the cell cycle phases.

K+ Currents Elicited by Depolarization in GH3 Cells

In GH3 cells, a 100-mV depolarizing step, from a holding potential of -80 mV, activated outward K+ currents that consisted of a rapidly activating and inactivating current (Ito) followed by a delayed slowly inactivating current (IKD) (Fig. 1A). These two currents have already been extensively described by Ritchie (36) and Oxford and Wagoner (29). The IKD current has been characterized as a voltage- and Ca2+-activated K+ current sensitive to ChTX, apamin, and TEA (20), while Ito seems to be mostly voltage dependent. The electrophysiological protocols used to differentiate Ito and IKD in many other cell types (e.g., recordings at different holding potentials, -80 and -40 mV) do not apply to GH3 cells because Ito is not totally inactivated at -40 mV (see Voltage Dependence of Ito in BrdU+ and BrdU- GH3 Cells). To study Ito we thus systematically blocked IKD by adding ChTX (20 nM), apamin (100 nM), and TEA (2 mM) to the external recording solution. Under these conditions, Ito was revealed and could be more easily studied (Fig. 1B). As already described (36), Ito was completely blocked by an acute application of 1 mM 4-AP (Fig. 1C), while IKD was only slightly inhibited. These effects of 4-AP were partly reversible.


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Fig. 1.   Electrophysiological recordings of outward K+ currents in GH3 cells. A: a 2-s depolarizing step from -80 to +20 mV (top) evoked a transient outward current (Ito; *) followed by a delayed slowly inactivating current (IKD; **) (bottom). B: recording of K+ currents in control condition () or in the presence of charybdotoxin (ChTX; 20 nM), tetraethylammonium (TEA; 2 mM), and apamin (Apa; 100 nM) (), which inhibited IKD and revealed Ito. C: effect of 4-aminopyridine (4-AP; 1 mM) on Ito. Inset: peak current amplitude vs. membrane potential before () and during application of 4-AP () and after washout (black-triangle).

Ito and IKD Current Densities in BrdU+ and BrdU- GH3 Cells

If Ito and/or IKD are important for cell proliferation, their electrophysiological characteristics should vary according to the phases in the cell cycle. We thus studied these currents in cells undergoing DNA synthesis (S phase) or in other phases. To this end, cells were preincubated with BrdU for 3 h before electrophysiological recordings were made and immunoreacted for BrdU after recordings were completed (see MATERIALS AND METHODS). This protocol establishes a correlation between the current profile of the recorded cell and its cycle phase. All current amplitudes were divided by cell capacitance to avoid differences due to the cell size. There was no difference in cell capacitance between BrdU+ and BrdU- cells (11.8 ± 0.4 pF, n = 80, for BrdU- cells vs. 12.0 ± 0.3 pF, n = 52, for BrdU+ cells; P > 0.05). The value chosen for calculation of the IKD current density was the plateau phase amplitude reached at the end of the step potential. IKD was studied in 31 cells, 18 of which were BrdU- and 13 BrdU+. There was no significant difference between the two groups in the mean IKD density evoked by a 2-s voltage step from -80 to +20 mV (93 ± 17 pA/pF in BrdU- cells vs. 97 ± 22 pA/pF in BrdU+ cells; P > 0.05). In contrast, the Ito density measured at the peak of the current elicited by a 500-ms depolarizing step from -80 to +20 mV was significantly higher in BrdU- cells (63 ± 4 pA/pF, n = 53) than in BrdU+ cells (42 ± 3 pA/pF, n = 40; P < 0.05) (Fig. 2). This result led us to study and compare the electrophysiological characteristics of Ito in the two cell groups.


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Fig. 2.   Current density of Ito and IKD in GH3 cells in S phase (BrdU+) and other cell cycle phases (BrdU-). Current density of Ito was measured as peak amplitude evoked by a 400-ms depolarizing pulse from -80 to +20 mV recorded in Hanks' balanced salt solution (HBSS) containing ChTX (20 nM), TEA (2 mM), and Apa (100 nM). Current density of IKD was measured as steady-state amplitude evoked by a 2-s depolarizing pulse from -80 to +20 mV. Results are expressed as means ± SE (*P < 0.05).

Voltage Dependence of Ito in BrdU+ and BrdU- GH3 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 incremental potential steps from -80 to + 70 mV. An example of currents observed in response to this protocol is shown in Fig. 3A. The mean activation of Ito in BrdU+ and BrdU- cells as normalized conductance amplitude (G/Gmax) was plotted as a function of test voltage and fitted to the Boltzmann equation (Fig. 3C). No difference in activation midpoint was observed between BrdU+ and BrdU- cells (V[1/2] = 16 ± 2 mV in BrdU+ cells, n = 8, vs. 14 ± 1 mV in BrdU- cells, n = 17; P > 0.05), and in all cells tested the activation threshold was observed between -20 and -10 mV. The voltage dependence of Ito inactivation was studied 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 to -10 mV (10-mV increments). The currents observed under these conditions (Fig. 3B), normalized as I/Imax, were expressed as a function of the conditioning prepulse voltage. The mean value of Ito inactivation was plotted and fitted to the Boltzmann equation for both cell groups (Fig. 3C). There was no significant difference in Ito inactivation mid-point between BrdU+ (V[1/2] = -42 ± 1 mV, n = 10) and BrdU- cells (V[1/2] = -42 ± 1 mV, n = 11; P > 0.05). Because no difference was observed between BrdU+ and BrdU- GH3 cells in terms of the voltage activation and inactivation dependence of Ito, we then investigated the Ito inactivation kinetics in both cell groups.


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Fig. 3.   Voltage dependence of Ito in BrdU+ and BrdU- cells. Currents were recorded in HBSS containing ChTX (20 nM), TEA (2 mM), and Apa (100 nM). A: voltage-clamp protocol used to determine Ito voltage-dependence activation. Current was activated by 10-mV incremental potential steps from holding potentials of -80 to +70 mV for 400 ms. B: voltage-clamp protocol used to determine Ito voltage-dependence inactivation. Current was elicited by a test pulse to +20 mV (400 ms long) after a 2-s conditioning prepulse from -100 to -10 mV in 10-mV increments. C: activation (circles) and inactivation (triangles) plots for Ito in BrdU+ (filled symbols) and BrdU- (open symbols) cells. Activation and inactivation plots show normalized conductance (G/Gmax) and current (I/Imax) as a function of voltage, respectively. Continuous and dotted lines are fits of the BrdU+ and BrdU- cell data by Boltzmann equation, respectively (see MATERIALS AND METHODS).

Ito Inactivation Kinetics in BrdU+ and BrdU- GH3 Cells

Differences in Ito inactivation kinetics might account for differences in Ito amplitude measured during a single-voltage test. As previously described by Oxford and Wagoner (29), the Ito inactivation proceeds as the sum of two exponentials with distinct amplitude and time constants. We studied the kinetics of Ito inactivation values occurring during a 500-ms voltage step from -80 to +20 mV (Fig. 4A). Our data show that the time constant of the fast component of inactivation (tau 1) was faster in BrdU- than in BrdU+ cells (tau 1 = 16.2 ± 1.4 ms in BrdU+ cells, n = 16, vs. tau 1 = 12.4 ± 0.7 ms in BrdU- cells, n = 19; P < 0.05), while no significant difference was observed in the time constant of the slow component of inactivation (tau 2) (tau 2 = 189.3 ± 36 ms in BrdU+ cells, n = 16, and tau 2 = 170.8 ± 24.1 ms in BrdU- cells, n = 19; P > 0.05) (Fig. 4B). Moreover, we found that the current density of the fast component (A1) was significantly higher in the BrdU- cells (A1 = 51.5 ± 7.1 pA/pF) than in the BrdU+ cells (A1 = 31.1 ± 4.4 pA/pF; P < 0.05), while no significant difference was observed in the current density of the slow component (A2) (Fig. 4C).


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Fig. 4.   Ito inactivation kinetics in BrdU+ and BrdU- cells. A: current evoked by a step to +20 mV from a holding potential of -80 mV. Curve fitting of the decay shows that the time course of inactivation is best fitted as a sum of two exponentials (fit superimposed on plot). B: duration of fast (tau 1) and slow (tau 2) components of Ito in BrdU+ (n = 16) and BrdU- cells (n = 19). C: amplitude of fast (A1) and slow (A2) components of Ito in BrdU+ and BrdU- cells (*P < 0.05).

These data suggest that the change in Ito current density observed between BrdU+ and BrdU- cells is probably due to a modification in the expression of the fast component of this current.

Recovery From Inactivation of Ito

The kinetics of recovery from inactivation of Ito differ markedly according to the K+ channel alpha -subunits responsible for this current. To determine the molecular basis of Ito in GH3 cells, we thus measured recovery from inactivation of Ito by using a double-pulse protocol. Cells were depolarized from the holding potential (VH = -80 mV) to +20 mV for 400 ms (test pulse); they were returned to -80 mV for 25 ms to 5 s (interpulse interval), and then a second test pulse was applied. Figure 5A shows an example of currents elicited in response to this protocol. The magnitude of Ito induced by the second pulse was expressed as a percentage of the Ito induced by the first pulse (%recovery) and then plotted as a function of the interpulse interval. Recovery of BrdU+ cells was best fitted by a monoexponential function (n = 4 of 6) (Fig. 5B), whereas, in BrdU- cells, recovery was best fitted by a biexponential function (n = 6 of 9) (Fig. 5B). No significant difference was found between the time constant tau  of BrdU+ cells and the tau slow of BrdU- cells (tau  = 649 ± 138 ms; tau slow = 1,055 ± 227 ms; P > 0.05), whereas the tau fast of BrdU- was significantly different from the tau  of BrdU+ cells (tau fast = 151 ± 34 ms; P < 0.05).


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Fig. 5.   Recovery from inactivation of Ito. A: example of superimposed Ito currents used to measure the time course of recovery from inactivation. B: recovery of Ito from inactivation was measured by using 2 depolarizing pulses to +20 mV (inset) separated by intervals of increasing duration (from 25 to 5,000 ms). Data were normalized to maximum current value and plotted as a function of recovery time to show the recovery from inactivation of a BrdU- cell (open circle ) and a BrdU+ cell (). The lines through the data represent the best fit of a second-order exponential function for the BrdU- cell and a first-order exponential function for the BrdU+ cell, respectively.

These results suggest that the molecular basis of K+ channels responsible for Ito in the S-phase cells differs from that in the non-S phase. Ito may be due to the activation of a single type of K+ channel in BrdU+ cells, whereas, in BrdU- cells, two types of K+ channels may be required.

Effect of K+ Channel Blocker 4-AP on [3H]Thymidine Incorporation

These results show a modification in the expression of Ito during the cell cycle. We wondered whether this modulation of Ito amplitude could be the cause or consequence of progression through the cell cycle. We therefore tested the effect of the inhibition of Ito by 4-AP on GH3 cell proliferation.

Incubation of GH3 cells with increasing concentrations of 4-AP (0.1 to 0.4 mM) for 72-h induced a dose-dependent inhibition of [3H]thymidine incorporation compared with controls (Fig. 6). This inhibition was not due to a cytotoxic effect of 4-AP, as shown by measurement of LDH activity (Fig. 6, inset). This result suggests that Ito current may be involved in the proliferation process of GH3 cells.


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Fig. 6.   Effect of 4-AP on GH3 cell proliferation and viability. Various concentrations of 4-AP were added to the culture medium for 72 h. Cell proliferation was then evaluated by the [3H]thymidine incorporation assay, as described in MATERIALS AND METHODS. Data are means ± SE obtained in 3 independent experiments (*P < 0.05). Inset: effect of 4-AP (0.4 mM) on cell viability as determined by the lactate dehydrogenase (LDH) assay. Cont, control; OD, optical density.

We then wondered whether the relationship between the variation in Ito and proliferation could be due to a modification of cell excitability.

Involvement of Ito in GH3 Cell Excitability

It has been shown that Ito is involved in the electrical activity of various cell types. We sought to determine whether the decrease in Ito amplitude observed in BrdU+ GH3 cells affected their excitability, compared with BrdU- cells. Current-clamp recordings were performed to test this hypothesis. A comparison of the electrical activity of BrdU+ and BrdU- GH3 cells showed no significant modification in membrane potential or action potential frequency and amplitude (Table 1).

                              
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Table 1.   Characteristics of electrical activity of BrdU+ and BrdU- GH3 cells or after treatment with 0.5 mM 4-AP for 24-30 h

We then studied the effect of chronic exposure of GH3 cells to 4-AP on electrical activity. These experiments 1) investigated the role of Ito in cell excitability and 2) were intended to determine whether the effect of 4-AP on cell proliferation could be linked to putative long-term modifications in electrical activity. Cells treated for 24-30 h with 0.5 mM 4-AP showed a complete inhibition of Ito in all tested cells (n = 9), as shown by voltage-clamp experiments. In current-clamp experiments, no significant difference was observed between controls (n = 11) and 4-AP-treated cells (n = 8) in terms of membrane potential or action potential amplitude and frequency (Table 1).


    DISCUSSION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Until now, the involvement of K+ channels in the control of cell proliferation had been mainly studied in nonexcitable cell types (49). In a previous study, we showed for the first time that K+ channels were likely to be involved in the proliferation of excitable GH3 cells, because the broad-spectrum K+ channel blocker TEA reduced cell proliferation by blocking the cell cycle at the G1/S boundary (46). In the present work, we investigated whether K+ currents per se could be involved in the proliferation mechanism of these excitable cells. We therefore studied the expression of K+ currents during different cell cycle phases by combining the incorporation of BrdU and electrophysiological recordings (see MATERIALS AND METHODS). Although the different cell cycle phases could not be precisely distinguished with the use of this protocol, K+ conductance expression activated by depolarization could be easily compared in cells in the S phase (BrdU+) and in other cell cycle phases (BrdU-) without any pretreatment that could alter cell physiology. In voltage-clamp experiments, a depolarizing pulse from -80 to +20 mV triggered an Ito and an IKD, which were mainly due to voltage- and Ca2+-dependent K+ conductances. These two currents were suppressed by the application of 4-AP and a cocktail of K+ channel inhibitors, TEA-ChTX-apamin, as previously described by Ritchie (36), respectively. Comparison of both currents in BrdU+ and BrdU- cells showed a 33% lower peak density for Ito in BrdU+ than in BrdU- GH3 cells, while the density of IKD was unchanged. Several studies have described modification in ion current amplitude during cell cycle progression (1, 8, 45, 24). These changes have mainly been observed at the G1/S and G2/M transitions. The G1 and G2 phases of the cell cycle are the functional periods during which cells prepare for DNA replication (S phase) and mitosis (M phase), respectively. Passage beyond the restriction point in G1 is controlled by a number of complex transcription factors as well as the expression of various cell cycle-related proteins. Our results suggest that changes in Ito density between BrdU- and BrdU+ cells may occur during the G1 or G2/M cell cycle phases. To determine whether Ito could be involved in proliferation processes, we then studied the effect of 4-AP on GH3 cell proliferation. Although 4-AP is the main Ito blocker, its inhibitory spectrum is fairly broad. To prevent possible nonspecific effects, we used the lowest 4-AP concentrations inducing Ito inhibition of GH3 cell proliferation. Our results show that 4-AP inhibits GH3 cell proliferation, suggesting that a change in Ito density may be required for cell cycle progress. From a more physiological point of view, it is noteworthy that thyrotropin releasing hormone, which decreases Ito amplitude (2, 41), induces cell cycle block in the G1 and G2/M phases in GH4 cells, a cell line closely related to GH3 (35).

Among the voltage-dependent K+ channel family (Kv), the Shal (Kv4) and members of the Shaker (Kv1 and particularly Kv1.4) subfamily have been shown to be responsible for Ito in various cell types (25, 48). The GH3 cell line expresses Kv1.4 and several types of Kv4 K+ channels, including Kv4.1 (51) and Kv4.3 (42). To determine the molecular basis for the K+ channel(s) responsible for the Ito expressed in BrdU+ and BrdU- cells, we studied the rate of recovery from inactivation of this current. Recovery from inactivation revealed two kinetically distinct components of Ito in BrdU- cells, suggesting that two different K+ channels are responsible for this current in S-phase cells. The kinetics of the rapid component (tau fast) were very similar to the kinetics of recovery of Kv4 expressed in other mammalian cells (48), which suggests that Shal-related K+ channels (i.e., Kv4.1 or Kv4.3) contribute to Ito in BrdU- cells. In addition to the rapidly recovering component in BrdU- cells, a slowly recovering component was also observed with a time constant that did not differ significantly from the single time constant of BrdU+ cells. Given that Kv1.4 is expressed in GH3 cells, it is possible that this channel contributes to Ito, although the values of the time constants observed here (tau  and tau slow) were faster than those usually described for Kv1.4 in mammalian cells. Kv1.4 may act as an homomeric or heteromeric alpha -subunit in GH3 cells along with Kv1.5 (22), and we cannot exclude the possible participation of another K+ channel, such as Kv3.4, in Ito. Our findings indicate that an additional Kv4 K+ channel alpha -subunit is expressed or activated in BrdU- but not in BrdU+ cells. Further molecular biology experiments will be necessary to confirm these electrophysiological data.

In addition to recovery from inactivation, comparison of the characteristics of Ito in BrdU+ and BrdU- cells showed a difference in inactivation kinetics of the current, whereas activation and inactivation voltage dependencies were unchanged. Indeed, characterization of the decaying phase of Ito revealed that the density of the faster component of the current (A1) was 40% higher in BrdU- than in BrdU+ cells. This was associated with an inactivation time constant (tau 1) faster in BrdU- than in BrdU+ cells. These results may explain the difference in Ito density observed between the two cell groups. Two hypotheses could account for these results. 1) Distinct channel types mediate the different inactivating components of the macroscopic Ito and, by a transcriptional regulation or translocation phenomenon (27, 19), their number in the plasma membrane may vary during the cell cycle. 2) A posttranslational regulation of one (or more) channel types may be responsible for the different current densities and time constants of the decaying phase of Ito. Indeed, Ito has been shown to be regulated by protein kinase A and/or protein kinase C in hippocampal pyramidal neurons (13) and myocytes (33). More recently, it was found that Kv1.4 and Kv1.5 K+ channels are regulated by tyrosine kinase phosphorylation (14, 32). This last observation is of particular interest because tyrosine kinase transduction pathways are involved in proliferation processes (44).

Although the involvement of K+ channels in the proliferation of various cell types has been demonstrated, the action mechanism involved in this effect has not yet been elucidated. A number of studies have emphasized that membrane hyperpolarization is an essential event required to trigger cell proliferation (49). Membrane hyperpolarization, by increasing the driving force for Ca2+, induces a rise in intracellular Ca2+ concentration, a necessary signal for cell cycle progression (4). We wondered whether in excitable cells, in which the control of membrane potential by K+ currents and the regulation of Ca2+ homeostasis are different from those of nonexcitable cells, the coupling between the K+ conductances and the proliferation may involve a variation of membrane potential or cell excitability.

The role of Ito in cell excitability has been extensively studied in neurons (7) and myocytes (11), in which it is mainly involved in interspike latency and action potential repolarization. In these cells, a variation in Ito amplitude causes a modification in cell excitability, which may be a messenger for cell proliferation, as suggested for neurons during corticogenesis (26). In the present work, the fact that there is no difference between the characteristics of the electrical activity of BrdU+ and BrdU- cells or between 4-AP-treated and control cells suggests that Ito may not be involved in regulating the excitability of GH3 cells. Although these results are surprising, they could be explained by the biophysical characteristics of the Ito expressed in GH3 cells. Indeed, the voltage activation and inactivation of Ito is more depolarized in GH3 cells than that usually described in neurons. In neurons, the potential values for the half-inactivation and the activation threshold are around -70 and -50 mV, respectively (39). These characteristics are compatible with a role of Ito in the excitability of neurons (action potential repolarization, interspike latency, and, to a lesser extent, action potential duration) (17, 37). In GH3 cells, we found the Ito inactivation midpoint at -40 mV and the activation threshold at -10 mV. These results are in agreement with those previously reported by Oxford and Wagoner (29). However, because the electrical activity of GH3 cells is between -40 and 0 mV, these biophysical characteristics make Ito inappropriate to play a significant role in excitability of these cells. A transient outward K+ current has been described in normal astrocytes but not in glioma cells. The role of this current in nonexcitable cells of this type is not known. However, the absence of this current in glioma cells appears to be an early feature accompanying the transformation of a normal astrocyte into a tumor cell (5).

However, the lack of long-term effect of 4-AP on membrane potential, while inhibiting cell proliferation, and the absence of difference between the membrane potential of BrdU+ and BrdU- cells suggest that variations of membrane potential are probably not involved in the transduction pathway by which K+ channels play a role in cell cycle progression in this cell type. Other mechanisms linking K+ channel activity and cell proliferation have also been proposed, including local variations in intracellular K+ concentration and/or osmolarity, which may activate several enzymes necessary for cell cycle progression (16). Another interesting possibility is the interaction between K+ channels and oncogenes or tumor-suppressor genes. It has been shown that transfection of fibroblasts with ras21 or treatment of fibroblasts with growth factors leads to the induction of a Ca2+-activated K+ channel that is essential for cell cycle progression (15). It is also noteworthy that, in Drosophila, the product of the tumor-suppressor gene dlg interacts with Kv1.4 K+ channels, leading to channel clustering (43). It is clear that further experiments will be required to determine how changes in K+ channel expression alter cell proliferation.

In conclusion, our results show that Ito is probably involved in the proliferation processes of the GH3 cell line because its inhibition by 4-AP reduces cell proliferation. Its expression level as well as its molecular composition are also clearly cell cycle dependent. Moreover, the link between Ito and cell proliferation is apparently not mediated by variations in membrane potential. It will be necessary to determine whether these findings are specific to this particular transformed cell model or whether they can be extended to other excitable pituitary and/or nonpituitary cells.


    ACKNOWLEDGEMENTS

This work was supported by Centre National de la Recherche Scientifique, University of Bordeaux 2, and Etablissement Public Regional Aquitaine.


    FOOTNOTES

Address for reprint requests and other correspondence: L. Bresson-Bepoldin, Laboratoire de Neurophysiologie, CNRS UMR 5543, Université de Bordeaux 2, 146 rue Leo 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.

Received 12 March 2000; accepted in final form 24 July 2000.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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