Cell cycle-dependent expression of a glioma-specific chloride current: proposed link to cytoskeletal changes

Nicole Ullrich and Harald Sontheimer

Department of Neurobiology, University of Alabama at Birmingham, Birmingham, Alabama 35294

    ABSTRACT
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Abstract
Introduction
Methods
Results
Discussion
References

We recently demonstrated expression of a novel, glioma-specific Cl- current in glial-derived tumor cells (gliomas), including stable cell lines such as STTG1, derived from a human anaplastic astrocytoma. We used STTG1 cells to study whether glioma Cl- channel (GCC) activity is regulated during cell cycle progression. Cells were arrested in defined stages of cell cycle (G0, G1, G1/S, S, and M phases) using serum starvation, mevastatin, hydroxyurea, demecolcine, and cytosine beta -D-arabinofuranoside. Cell cycle arrest was confirmed by measuring [3H]thymidine incorporation and by DNA flow cytometry. Using whole cell patch-clamp recordings, we demonstrate differential changes in GCC activity after cell proliferation and cell cycle progression was selectively altered; specifically, channel expression was low in serum-starved, G0-arrested cells, increased significantly in early G1, decreased during S phase, and increased after arrest in M phase. Although the link between the cell cycle and GCC activity is not yet clear, we speculate that GCCs are linked to the cytoskeleton and that cytoskeletal rearrangements associated with cell division lead to the observed changes in channel activity. Consistent with this hypothesis, we demonstrate the activation of GCC by disruption of F-actin using cytochalasin D or osmotic cell swelling.

ion channels; flow cytometry; proliferation; chlorotoxin; glioblastoma; astrocytoma

    INTRODUCTION
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Abstract
Introduction
Methods
Results
Discussion
References

RECENT STUDIES SUGGEST THAT changes in ion channel activity may be associated with cell proliferation (10, 21, 22, 24, 38), differentiation (26), migration (25), and biological activation (13). Possible sites of involvement of ion channels include transduction of mitogen-stimulated protein synthesis (18), rearrangements in cytoskeletal F-actin (4), generation of Ca2+ signals (7), and pH regulation (22). In a number of cell types, increased ion permeability is one of the earliest events after mitogen stimulation (11), and, in particular, K+ and Cl- channels have been implicated in the proliferative response. For example, in lymphocytes, K+ channels are involved in volume regulation, control of cellular toxicity, and the immune response (23); application of K+ channel blockers such as tetraethylammonium, 4-aminopyridine, and quinine to lymphocytes suppresses proliferation in a concentration-dependent manner (10). Blockade of voltage-activated K+ channels also leads to a dose-dependent decrease in the proliferation of melanoma cells (21), breast cancer cells (38), and several glial cell types including Schwann cells (6), retinal glial cells (24), and astrocytes (22). More recently, Cl- channels have been implicated in the proliferative response of Schwann cells (37), B lymphocytes (9), and glioma cells (35), suggesting that the link between channel activity and proliferation also extends to this class of channels.

Cell division can be operationally divided into defined stages of the cell cycle (see also Fig. 6). Interestingly, the activity of some ion channels has been shown to vary during the cell cycle. For example, in mouse oocytes, a large-conductance (241 pS), voltage-activated K+ channel is active in G1 and M phases but is inactive during the G1/S transition (8). Similarly, in lymphocytes, the delayed rectifier K+ channel is thought to be required for cell cycle progression as a signal for progression from G0 to G1, a transition that relies on membrane depolarization and requires a transmembrane flux of Ca2+ (23). In these cells, Cl- permeability varies with cell cycle, being low in G0 and S phases and increases in G1/S (3). Transient changes in ion membrane permeability and cell ion content are generally assumed to play a key role in the transition from the quiescent state G0 or early G1 phase to the S phase of DNA replication (23, 27). Although the exact role that channel activity plays in cell cycle progression is not clear, it has been proposed that changes in channel activity result in both long-term changes in gene expression and short-term modulation of preexisting channel proteins (23).

We recently reported the expression of whole cell Cl- currents with unique biophysical and pharmacological properties that selectively characterizes glioma-derived cells (35). The goal of the present work was to determine whether astrocytoma Cl- current activity varies as a function of cell cycle. Our results suggest a marked and differential upregulation of Cl- currents after cell cycle arrest, with highest glioma Cl- channel (GCC) activity in early G1 and lowest activity in G0/G1 and S phases. We also show that GCC can be activated by disruption of F-actin filaments. Taken together, we speculate that changes in GCC activity during the cell cycle result from rearrangements of the cells cytoskeleton.

    METHODS
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Abstract
Introduction
Methods
Results
Discussion
References

Cell culture. The STTG1 cell line (American Type Culture Collection, Rockville, MD) was grown in Dulbecco's modified Eagle's medium (DMEM, GIBCO) plus 10% fetal calf serum (FCS, HyClone) at 37°C in a humidified 10% CO2-90% air atmosphere. Cells attaining nearly confluent growth were harvested and plated onto uncoated 75-cm2 flasks or uncoated 12-mm circular glass coverslips for electrophysiology and were used 36-72 h after plating, unless otherwise noted. Viable cell counts were determined by trypan blue exclusion.

Electrophysiology. Current and voltage recordings were obtained using standard whole cell patch-clamp methods with an Axopatch-1D amplifier (Axon Instruments). Patch pipettes were made from thin-walled borosilicate glass (WPI, TW150F-40, 1.5 mm OD, 1.2 mm ID) and were filled with a solution containing (in mM) 145 KCl, 1 MgCl2, 0.2 CaCl2, 10 ethylene glycol-bis(beta -aminoethyl ether)-N,N,N',N'-tetraacetic acid, and 10 N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid [HEPES; pH adjusted to 7.4 with tris(hydroxymethyl)aminomethane]. Pipettes typically had resistances between 4 and 7 MOmega . Cells were continuously superperfused with a standard bath solution containing (in mM) 125 NaCl, 5 KCl, 1.2 MgCl2, 1.0 CaCl2, 1.6 Na2HPO4, 0.4 NaH2PO4, 32.5 HEPES (acid), and 10.5 glucose (pH adjusted to 7.4 with NaOH). Osmolarity was measured with a vapor pressure osmometer (Wescor, Logan, UT) and ranged between 308 and 313 mosM. Calculated equilibrium potentials for K+, Na+, and Cl- under the imposed ionic gradients in control solution were as follows: EK = -83.4 mV, ENa = +62.6 mV, and ECl = +2.8 mV.

For whole cell recordings, cell capacitance compensation and series resistance compensation were used to minimize voltage errors. The amplifier reading of capacitance was used as the value for the whole cell membrane capacitance. Series resistances, monitored at regular intervals throughout each experiment, were usually 5-10 MOmega , and series resistance compensation was typically set to ~80%. Entrance potential, read from the amplifier at the time of entering the whole cell configuration, was used to determine each cell's resting potential. Images were taken of each recorded cell using a charge-coupled device camera (Sony, NY) and a video printer for cataloging of cell size, location, and morphology. Recordings were made at room temperature, typically 20-25°C.

Proliferation assay. Proliferation was studied quantitatively by determining incorporation of [3H]thymidine, as we have previously described (35). In brief, cells were plated and, after 1 day in culture, were treated for 48 h in the continuous presence or absence of the agent of interest. The growth effects of these agents were tested by the dilution of concentrated stock solutions into the medium. Cells were incubated with 1 µCi/ml radiolabeled thymidine ([methyl-3H]thymidine) for the final 4 h (at 37°C). Culture dishes were rinsed three times with ice-cold phosphate-buffered saline (PBS) and solublized with 0.3 N NaOH for 30 min at 37°C. One aliquot (50 µl) was used for cell protein determination using the bicinchoninic assay (Pierce, Rockford, IL). The remaining cell suspension was mixed with Ultima Gold, and radioactivity was determined with a scintillation counter. The results were expressed as counts per minute per milligram of protein.

DNA flow cytometric analysis. Cells from sister cultures were plated in parallel to those described above for proliferation assays at a density of 106 cells/well. Cells were incubated in the different drugs 48 h after seeding and were harvested by trypsinization after a 48-h incubation period. Cells were rinsed three times with cold PBS, fixed in 70% ethanol for 1 h at 4°C, rinsed with PBS, and incubated with propidium iodide (Boehringer Mannheim) for 1 h at 4°C in the dark. Flow cytometric analysis for cell cycle distribution was done on a FACScan cytofluorometer (Becton Dickinson) using CellFit software, which analyzes cells for their different DNA content based on propidium iodide staining. The SOBR model, which fits Gaussian distribution to the fluorescence peaks, was used to calculate the percentage of cells falling in the G0/G1, S, and G2+M populations.

Immunohistochemical staining. The cultures were fixed for at least 20 min at 4°C in 4% paraformaldehyde and then washed in PBS. Cells were incubated with rhodamine-conjugated phalloidin (Molecular Probes) for 30 min at room temperature. Coverslips were washed in PBS, mounted in Fluoromount (Fisher) on glass slides, and viewed under an epifluorescence microscope using standard procedures.

Data analysis. For all experiments, mean values, SD, and SE were computed from raw values entered into a spreadsheet (Excel, Microsoft). These data were exported to a scientific graphing and data analysis program (ORIGIN, MicroCal). Data were graphed as means ± SE. All statistical analysis was obtained using analysis of variance (ANOVA) test for multiple comparisons, and P values given represent Bonferroni-corrected values.

Drugs used. Mevastatin was purchased from Biomol (Plymouth Meeting, PA). Unless otherwise noted, cytosine beta -D-arabinofuranoside (Ara-C), hydroxyurea, demecolcine, and all other chemicals were purchased from Sigma. Ara-C, hydroxyurea, and demecolcine were diluted in distilled H2O. Mevastatin was diluted in ethanol (final concentration 0.08%).

    RESULTS
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Abstract
Introduction
Methods
Results
Discussion
References

Whole cell voltage-clamp recordings were obtained from the human anaplastic astrocytoma cell line, STTG1. Most of the STTG1 cells (85%) were glial fibrillary acidic protein-positive (GFAP). Unsynchronized cells were used as the "control" cell population and varied in morphology from flat, polygonal cells with short or no processes to elongated, bipolar-shaped cells (see Fig. 3A). Recordings from these control cells consistently showed time- and voltage-dependent outward currents in all (n = 843, Fig. 1A) cells. We previously showed that these outward currents were mediated by the inward movement of Cl- through a glioma-specific class of Cl- channels (35). The resting potential, determined as the entrance potential with the KCl-containing pipette solution, was -14.1 ± 0.56 (SE) mV (n = 843). A representative example of whole cell recordings from an STTG1 human astrocytoma cell in response to depolarizing voltage steps is displayed in Fig. 1A. Recordings were obtained by stepping the cells from a holding potential of 0 mV to a series of test potentials between -105 and 195 mV in 25-mV increments. Potentials >45 mV resulted in fast activating, noninactivating outward currents. Cells showed large outward transients on termination of voltage steps (Fig. 1A). The current-voltage (I-V) relationship plotting peak current amplitude as a function of voltage showed pronounced voltage dependence and outward rectification for both the steady-state (Fig. 1B, ×) and outward transient currents (Fig. 1B, *). Cells from all studied STTG1 cells displayed such outwardly rectifying currents. We recently published a comprehensive biophysical and pharmacological characterization of these currents in STTG1 cells, suggesting that the underlying channels, which we have since termed glioma Cl- channels (GCCs), are unique in some of their biophysical and pharmacological features (35).


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Fig. 1.   Glioma Cl- channel (GCC) currents in unsynchronized cells. A: whole cell voltage-clamp recordings obtained from a representative STTG1 cell. Cells were stepped to test potentials between -105 and 195 mV in 25-mV increments from a holding potential of 0 mV (inset). Potentials >45 mV resulted in fast-activating, noninactivating outwardly rectifying currents (×). Cells showed large transients on termination of voltage steps (*). B: current-voltage relationship of steady-state and transient peak current amplitude is plotted as a function of applied voltage.

Dependence of Cl- channel expression on cell cycle. To test the hypothesis that GCC activity may change during cell cycle progression, STTG1 cells were arrested in defined stages of cell cycle. Methods used for cell cycle arrest were 1) serum starvation (to 0.1% FCS), a procedure commonly used to synchronize and arrest the cells in early G0/G1 (15); 2) treatment with mevastatin (2.5 µM), which arrests cells in early G1 (32); 3) treatment with hydroxyurea (1 mM), which arrests cells at the G1/S boundary (32); 4) treatment with Ara-C (10 µM), which arrests cells in S phase; and 5) treatment with demecolcine (0.05 µM), which arrests cells in M phase (33). The concentrations of each of these reagents were established empirically as the lowest required concentration to yield the desired effect. These are in good agreement with concentrations used by others (32).

To evaluate the effectiveness of the chosen cell cycle arrest procedures to arrest glioma proliferation, we determined the relative incorporation of [3H]thymidine as a quantitative marker of DNA synthesis. Cells were treated at 1 day in culture (DIC) with the appropriate agents (see above), and [3H]thymidine incorporation into DNA was assayed 48 h later, at 3 DIC, a period of intense cell proliferation of untreated control cultures. Experiments were performed on cells derived from at least four separate cell passages. As expected, all cycle arrest agents led to a marked decrease in proliferation (Fig. 2). Of these, Ara-C was the most effective [76.5 ± 1.8 (SD), n = 23]; serum starvation, mevastatin, hydroxyurea, and demecolcine reduced proliferation by 39.7 ± 3.8% (SE; n = 18), 49.4 ± 4.9% (n = 12), 59.6 ± 3.2% (n = 28), and 55.2 ± 3.6% (n = 27), respectively (Fig. 2). These inhibitory effects on proliferation were statistically significant from control (ANOVA test, Bonferroni P < 0.001). Sister cultures were also grown in the presence of 0.1% ethanol, which had no appreciable affect on cell proliferation.


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Fig. 2.   Specific cell cycle arrest. Effects of presence or absence of fetal calf serum (FCS; serum starvation conditions), mevastatin, hydroxyurea, cytosine beta -D-arabinofuranoside (Ara-C), and demecolcine on proliferation of astrocytoma cells, assessed by [3H]thymidine incorporation after 24-h incubation with reagent of interest. Mean effects (expressed as cpm/mg protein; error bars = SE) plotted for each agent tested were result of a minimum of 4 experiments each (see text for details). These effects were statistically significant (ANOVA test, Bonferroni P < 0.001).

DNA flow cytometric analysis. DNA flow cytometric analysis was used to confirm that the cells chosen for physiological evaluation were arrested at the predicted state of the cell cycle by the different reagents (5). Experiments were performed on sister cultures in parallel and electrophysiological recordings. In the absence of any reagents, most of the unsynchronized cells were in G0/G1 phase of cell cycle (60.6%), with a subpopulation of cells dividing so that 20.2% of the cells were in S phase and 19.2% of the cells were in G2+M phases. By contrast, with each of the arrest agents, we observed a selective accumulation of cells in defined stages of the cell cycle; the relative percentages of cells in the various cell cycle phases under the experimental conditions are given in Table 1. Because flow cytometry only distinguishes three groups of phases of cell cycle, namely, G0+G1, S, and G2+M, arrests by serum starvation (G0/G1) and hydroxyurea (G1/S) are virtually indistinguishable, since no clear G0/G1 distinction is possible. However, a large literature exists that suggests that serum starvation arrests at an earlier cell cycle stage than hydroxyurea.

                              
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Table 1.   Cell cycle stage distribution

Effect of cell cycle on Cl- channel expression and cellular morphology in astrocytoma cells. Patch-clamp recordings were obtained from sister cultures of STTG1 astrocytoma cells that had been incubated for 44-52 h in the agents described above, and GCC conductances were determined to allow comparison of GCC expression between these cell populations. Because cell shape and cellular surface area change during cell division (31), conductances were normalized to cell size as conductance densities to allow for an independent comparison. Representative phase images of cells treated with cell cycle arrest agents are shown in Fig. 3A, and mean conductance densities are plotted in Fig. 3B. Marked changes in cell morphology and GCC conductance density values were observed. Conductance densities were highest in early G1, lowest in S phase, and intermediate in M phase. Table 2 lists mean values of resting potential, cell capacitance, peak current amplitude, and conductance densities for each condition. Conductance density did not differ significantly (P = 0.46) when serum-starved cells (442 ± 58 pS/pF, n = 19) and control cells (497 ± 55 pS/pF, n = 40) were compared. Serum starvation changed cell morphology to a more triangular, flat cell body with elongated, bipolar processes, and cell resting potential was consistently more hyperpolarized (-30.1 mV compared with -13.6 mV for control cells). A second population of cells was allowed to progress to early G1 phase and arrested with mevastatin. Cell processes were almost exclusively bipolar (Fig. 3A), and conductance density was significantly (P < 0.05) higher with values of 798 ± 100 pS/pF (n = 19). Arrest at the G1/S transition with hydroxyurea (Fig. 3B) led to a small decrease in GCC conductance density (590 ± 78 pS/pF, n = 21) that was not significantly different from control or serum-starved cells and a change to more polygonal cellular morphology. Conductance density was lowest after arrest in S phase by Ara-C (318 ± 44 pS/pF, n = 20, P < 0.01; Fig. 3B). Arrest by demecolcine in M phase yielded conductance density values of 595 ± 71 pS/pF (n = 21, P < 0.01). Cells treated with demecolcine were visibly arrested in the process of cell division, had rounded cell bodies, and had retracted all processes (Fig. 3A). In all cell cycle arrest conditions, both steady-state and transient components of GCC currents were affected about equally (Table 2).


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Fig. 3.   Effects of cell cycle arrest on cellular morphology and GCC expression. Resting STTG1 cells were incubated for 48 h in continued presence or absence of 0.1% FCS or cell cycle arrest agents mevastatin, hydroxyurea, Ara-C, or demecolcine. Mean GCC conductance densities were determined for each condition and compared with control. Cellular morphology (A) and GCC conductance densities (B) varied depending on cell cycle phase at which cells were arrested.

                              
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Table 2.   Effects of cell cycle arrest on glioma Cl- currents

Changes in astrocytoma morphology and GCC activity are associated with cytoskeletal changes. Cell proliferation involves dramatic rearrangements of a cell's cytoskeleton. It is now evident that ion channels including Cl- channels can be functionally linked to the cytoskeleton, and disruptions of these contacts have been shown to lead to changes in the functional state of ion channels (20). Consequently, we explored whether the observed changes in GCC amplitudes after cell cycle arrest could be explained by possible changes in the GCC-cytoskeleton relationship. Similar studies in nontumor astrocytes (16) suggest that rearrangements in the cell cytoskeleton can activate Cl- channels. To establish that GCC channels are linked to the cytoskeleton, we experimentally disrupted the cytoskeleton by swelling cells with hyposmotic solution. To visualize changes in the cytoskeleton, cells were stained with a rhodamine conjugate of phalloidin. In control cells exposed to normosmolar bath solution (310 mosM), the F-actin appeared in a well-organized pattern of stress fibers that traversed the entire cell and intersected with the cell membrane (Fig. 4A). When the cells were swelled after change to a hypotonic bath solution (200 mosM), the organized pattern of actin staining was disrupted and stress fibers were greatly reduced (Fig. 4B). These data suggest that shape and volume changes are indeed associated with cytoskeletal changes in glioma cells and could be used as a model to study channel-cytoskeletal interactions.


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Fig. 4.   Osmotic swelling leads to disruption of actin cytoskeleton associated with increased GCC currents. Osmotic challenges greatly reduced stress fibers as shown by staining cells with rhodamine-conjugated phalloidin. Labeling of control cells (A) was compared with cells exposed for 10 min to hypotonic solutions (B). C: family of voltage step from -105 to 195 mV is shown before and after osmotic challenge (310right-arrow200 mosM). Note that no currents (I) were elicited with voltages <0 mV and individual traces are difficult to see as they superimpose. D: peak currents from C were plotted as a function of voltage before and after osmotic challenge for voltages ranging from -105 to 195 mV.

Disruptions of cytoskeletal F-actin such as those induced by reduction of bath osmolarity also led to pronounced changes in GCC activity. Figure 4C shows whole cell currents recorded before and after the application of hypotonic bathing solution. The cell was initially bathed in standard NaCl solution (310 mosM), and hypotonic solutions were applied from a perfusion system. After a delay of ~30 s, the amplitude of GCC currents reversibly increased and subsequently diminished to control values on return to the standard bath solution within 5 min. The I-V curves of the steady-state current component (Fig. 4D) show that, under hypotonic conditions, current amplitudes were increased. The mean values for this increase determined at 145 mV in eight cells was 64.1 ± 17.6% (SE). In hypotonic bathing solution, the outward current activated at potentials less depolarized than 45 mV, suggesting that the threshold for activation can be altered to more physiological ranges.

Because the hypotonic bath solution was created by removal of 50 mM NaCl, we ensured that the observed effects were not secondary to a shift in ECl. Therefore cells were exposed to hypotonic bath solution made normosmolar by the addition of 100 mosM mannitol (100 mM mannitol). Under such bath conditions, currents did not change appreciably from control, suggesting that the increase in current amplitude was the result of osmotic changes and not due to changes in extracellular Cl- concentration (not shown).

To test the hypothesis that changes in actin cytoskeleton accompany the swelling-induced changes in GCC currents, phalloidin, which binds and stabilizes F-actin, was included in the pipette. Cell dialysis with 1 µM phalloidin before exposing cells to the hypotonic solutions inhibited the osmotically induced increases in GCC [93.1 ± 1.35% (SE), n = 4]. Such effects of phalloidin on osmotically activated whole cell Cl- currents have previously been observed in renal cells (29). By contrast, inclusion of 5 µM cytochalasin D, which depolymerizes actin by augmenting actin-ATP hydrolysis (28), elicited increased GCC amplitudes in the absence of osmotic changes [123.6 ± 2.6% (SE), n = 5; Fig. 5]. In most cells, this increase was evident within 3 min of achieving whole cell configuration and was maximal after 10 min. Differences in the time course may reflect cell-cell variability in access resistance as well as cell size. A summary of the changes in GCC after osmotic or cytoskeletal disruption is shown in Fig. 5. As seen, hypotonic challenges lead to increased GCC amplitudes that are inhibited by cell dialysis with phalloidin. In contrast, isochloric normosmotic solutions failed to induce changes in GCC. In addition, actin disruption with cytochalasin was sufficient to cause increases in GCC. These data suggest a functional interaction between GCC and the cytoskeleton.


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Fig. 5.   Changes in GCC currents in response to osmotic challenges can be overcome by phalloidin. Osmotically induced current increases can be prevented by stabilizing cytoskeleton with phalloidin. Increases in whole cell currents induced by osmotic challenges can be mimicked by disruption of cytoskeleton using cytochalasin D in absence of any osmotic changes. Osmotically induced increases and cytochalasin D effects were statistically significant (** P < 0.01; * P < 0.05).

    DISCUSSION
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Abstract
Introduction
Methods
Results
Discussion
References

GCC expression in STTG1 human anaplastic astrocytoma cells was altered if cell proliferation and cell cycle progression were selectively inhibited in defined phases of cell cycle by chemical reagents. STTG1 cells are well-differentiated, GFAP-positive cells widely used as an in vitro model for astrocytoma cell growth. These cells show other hallmarks of astrocytoma cells including responsiveness to epidermal growth factor (unpublished observations) and, as we have previously shown, consistent expression of GCC currents (35).

Striking changes in cellular morphology and GCC activity were associated with each phase of cell cycle. GCC conductance density was approximately twofold higher in G1-arrested cells than in quiescent cells and, overall, was lowest in cells arrested in S phase. Thus, if one views the cell cycle as a continuum, there appear to be cyclical changes in GCC activity (Fig. 6). These changes affected both steady-state and transient current components. Changes in whole cell GCC currents could have resulted from changes in channel expression, recruitment of quiescent channels, or changes in the biophysical properties of the underlying channels. Our electrophysiological approach cannot discriminate between these possibilities. Consequently, we do not at present understand the underlying mechanisms; however, examples exist in the literature to support each of these possibilities. For example, in mouse early embryos, a voltage-activated K+ channel is active throughout M and G1 phases and switches off during the G1/S transition (8). These changes in channel activity are accompanied by shifts in resting potential. K+ channels are also dependent on the cell cycle in breast cancer cells, where inhibition of K+ channels by channel blockers with different mechanisms of actions led to inhibition of proliferation in the G0/G1 phase of cell cycle (38). In HeLa cells (34) and neuroblastoma cells (1), the activation kinetics of the inwardly rectifying K+ channels and the resting potential itself were linked to the G1/S transition, and it is thought that the changes in channel biophysics were responsible for a depolarized resting potential that may be permissive for DNA synthesis. Because we observed reversible changes in GCC conductance of similar magnitude by experimental disruption of the cytoskeleton, we are compelled to speculate that conductance changes during the cell cycle were likewise due to changes in channel activity.


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Fig. 6.   A: schematic diagram of cell cycle. B: changes in Cl- channel conductance density at defined stages of cell cycle. Conductance density was highest in early G1, lowest in S phase, and intermediate in M phase.

Cl- channels have also been shown to be regulated by cell cycle in lymphocytes (3), cultured myoblasts (14) and Boltenia embryos (36). In Boltenia embryos and in mouse early embryos, changes in current amplitude are thought to be due to cyclical insertion of the channel protein into and removal from the plasma membrane. The incidence and current amplitudes of the large-conductance Cl- channel's cultured myoblasts were substantially higher in proliferating vs. nonproliferating cells (14).

Our data do not distinguish between changes in ion channel numbers and channel open probability. However, channels can be modulated on a short-term basis by both changes in intracellular Ca2+ concentration and second messenger systems. The primary physiological regulator of Cl- conductance in lymphocyte cycling is probably intracellular Ca2+ (2). In these cells, cytosolic Ca2+ is low in G0 and S and increases threefold in G1 (30). We are currently investigating whether such cyclical changes in Ca2+ also occur in glioma cells. Other regulatory mechanisms of ion channels can involve their phosphorylation and dephosphorylation (36). These properties are integral to the functioning of cyclins and cyclin-dependent kinases, the operators of the cell-cycle clock (17). It is possible that the channel's voltage sensitivity or activation properties are changed by phosphorylation. Clearly, further studies are needed to address these important issues.

The role of GCC channels in glioma cell function and their possible relationship to the cell cycle is unclear. However, Cl- channels are commonly implicated in cell volume regulation and water transport, and increased Cl- conductances have been demonstrated to be associated with volume changes in a number of cell types (12, 19). Consistent with a possible involvement of GCCs in volume regulation of glioma cells, GCC activity was enhanced in response to osmotic swelling associated with hypotonic challenges (66% osmolarity). Moreover, our experiments showing changes in GCC activity after perturbation of the cytoskeleton led us to hypothesize that GCC activity may be functionally linked to changes in cell shape, occurring during cell swelling or cell migration. Any change in cell shape that results in a net gain or loss of H2O requires movements of Cl- and K+ across the cell membrane. It is thus conceivable that changes in GCC activity are associated with H2O movements in conjunction with cell shape changes.

A unique characteristic of glial tumors is their tendency to migrate along anatomic pathways throughout the brain. To migrate, glioma cells must be able to readily change cell shape, requiring significant rearrangements in their cytoskeleton. Moreover, migrating glioma cells have to squeeze through narrow extracellular spaces, probably requiring them to transiently decrease their cell volume. As discussed above, any decrease in cell volume requires extrusion of H2O in conjunction with Cl- and K+. It is thus conceivable that GCC channels play an important role in facilitating volume and shape changes associated with glioma migration. Such an involvement of Cl- channels in shape changes has been demonstrated for nontumor astrocytes in which the transition from a flat to a stellate cell leads to the activation of Cl- currents, a process that can be prevented by stabilization of F-actin with phalloidin (16). Indeed, a link of ion channels and particularly Cl- channels to cytoskeletal rearrangements has been documented for a number of other preparations (20). On the basis of these findings, we hypothesize that glioma Cl- channels may represent an adaptive feature facilitating shape and volume changes related to glioma cell proliferation and migration. Further studies are needed to clarify the role of GCC in these processes.

    ACKNOWLEDGEMENTS

This work was supported by National Institute of Neurological Disorders and Stroke Grant R01-31234.

    FOOTNOTES

Present address of N. Ullrich: Interdepartmental Neuroscience Program, Yale University School of Medicine, New Haven, CT 06510.

Address for reprint requests: H. Sontheimer, Dept. of Neurobiology, University of Alabama at Birmingham, 1719 6th Ave S., CIRC Rm 545, Birmingham, AL 35294.

Received 6 November 1996; accepted in final form 27 May 1997.

    REFERENCES
Top
Abstract
Introduction
Methods
Results
Discussion
References

1.   Arcangeli, A., L. Bianchi, A. Becchetti, L. Faravelli, M. Coronnello, E. Mini, M. Olivotto, and E. Wanke. A novel inward-rectifying K+ current with a cell-cycle dependence governs the resting potential of mammalian neuroblastoma cells. J. Physiol. (Lond.) 489: 455-471, 1995[Abstract].

2.   Baran, I. Calcium and cell cycle progression: possible effects of external perturbations on cell proliferation. Biophys. J. 70: 1198-1213, 1996[Abstract].

3.   Bubien, J. K., K. L. Kirk, T. A. Rado, and R. A. Frizzell. Cell cycle dependence of chloride permeability in normal and cystic fibrosis lymphocytes. Science 248: 1416-1419, 1990[Medline].

4.   Cantiello, H. F. Role of the actin cytoskeleton on epithelial Na+ channel regulation. Kidney Int. 48: 970-984, 1995[Medline].

5.   Carey, F. A. Measurement of nuclear DNA content in histological and cytological specimens: principles and applications. J. Pathol. 172: 307-312, 1994[Medline].

6.   Chiu, S. Y., and G. F. Wilson. The role of potassium channels in Schwann cell proliferation in Wallerian degeneration of explant rabbit sciatic nerves. J. Physiol. (Lond.) 408: 199-222, 1989[Abstract].

7.   Cornell-Bell, A. H., S. M. Finkbeiner, M. S. Cooper, and S. J. Smith. Glutamate induced calcium waves in cultured astrocytes: long-range glial signaling. Science 247: 470-473, 1990[Medline].

8.   Day, M. L., S. J. Pickering, M. H. Johnson, and D. I. Cook. Cell-cycle control of a large-conductance K+ channel in mouse early embryos. Nature 365: 560-562, 1993[Medline].

9.   Deane, K. H., and M. D. Mannie. An alternative pathway of B cell activation: stilbene disulfonates interact with a Cl- binding motif on AEN-related proteins to stimulate mitogenesis. Eur. J. Immunol. 22: 1165-1171, 1992[Medline].

10.   DeCoursey, T. E., G. Chandy, S. Gupta, and M. D. Cahalan. Voltage-gated K+ channels in human T lymphocytes: a role in mitogenesis? Nature 307: 465-468, 1984[Medline].

11.   Dubois, J. M., and B. Rouzaire-Dubois. Role of potassium channels in mitogenesis. Prog. Biophys. Mol. Biol. 59: 1-21, 1993[Medline].

12.   Garber, S. S. Outwardly rectifying chloride channels in lymphocytes. J. Membr. Biol. 127: 49-56, 1992[Medline].

13.   Gardner, P. Patch clamp studies of lymphocyte activation. Annu. Rev. Immunol. 8: 231-252, 1990[Medline].

14.   Hurnak, O., and J. Zachar. Conductance-voltage relations in large-conductance chloride channels in proliferating L6 myoblasts. Gen. Physiol. Biophys. 13: 171-192, 1994[Medline].

15.   Langan, T. J., M. C. Slater, and K. Kelly. Novel relationships of growth factors to the G1/S transition in cultured astrocytes from rat forebrain. Glia 10: 30-39, 1994[Medline].

16.   Lascola, C. D., and R. P. Kraig. Whole-cell chloride currents in rat astrocytes accompany changes in cell morphology. J. Neurosci. 16: 2532-2545, 1996[Abstract].

17.   Leake, R. The cell cycle and regulation of cancer cell growth. Ann. NY Acad. Sci. 784: 252-262, 1996[Abstract].

18.   Lewis, R. S., and M. D. Cahalan. Potassium and calcium channels in lymphocytes. Annu. Rev. Immunol. 13: 623-653, 1995[Medline].

19.   Lohr, J. W., and L. A. Yohe. Mechanisms of hypoosmotic volume regulation in glioma cells. Brain Res. 667: 263-268, 1994[Medline].

20.   Mills, J. W., E. M. Schwiebert, and B. A. Stanton. The cytoskeleton and membrane transport. Curr. Opin. Nephrol. Hypertens. 3: 529-534, 1994[Medline].

21.   Nilius, B., and W. Wohlrab. Potassium channels and regulation of proliferation of human melanoma cells. J. Physiol. (Lond.) 445: 537-548, 1992[Abstract].

22.   Pappas, C. A., N. Ullrich, and H. Sontheimer. Reduction of glial proliferation by K+ channel blockers is mediated by changes in pHi. Neuroreport 6: 193-196, 1994[Medline].

23.   Premack, B. A., and P. Gardner. Role of ion channels in lymphocytes. J. Clin. Immunol. 11: 225-238, 1991[Medline].

24.   Puro, D. G., F. Roberge, and C. C. Chan. Retinal glial cell proliferation and ion channels: a possible link. Invest. Ophathal. Vis. Sci. 30: 521-529, 1989[Abstract].

25.   Rakic, P., R. S. Cameron, and H. Komuro. Recognition, adhesion, transmembrane signaling and cell motility in guided neuronal migration. Curr. Opin. Neurobiol. 4: 63-69, 1994[Medline].

26.   Reuter, H., A. Bouron, R. Neuhaus, C. Becker, and B. Reber. Inhibition of protein kinases in rat pheochromocytoma (pc12) cells promotes morphological differentiation and down-regulates ion channel expression. Proc. R. Soc. Lond. B Biol. Sci. 249: 211-216, 1992[Medline].

27.   Rozengurt, E., and S. Mendoza. Monovalent ion fluxes and the control of cell proliferation in cultured fibroblasts. Ann. NY Acad. Sci. 339: 175-190, 1980[Medline].

28.   Sampath, P., and T. D. Pollard. Effects of cytochalasin, phalloidin, and pH on the elongation of actin filaments. Biochemistry 30: 1973-1980, 1991[Medline].

29.   Schwiebert, E. M., J. W. Mills, and B. A. Stanton. Actin-based cytoskeleton regulates a chloride channel and cell volume in a renal cortical collecting duct cell line. J. Biol. Chem. 269: 7081-7089, 1994[Abstract/Free Full Text].

30.   Silver, R. B. Calcium and cellular clocks orchestrate cell division. Ann. NY Acad. Sci. 582: 207-221, 1990[Abstract].

31.   Sit, K. H., R. Paramanantham, B. H. Bay, and K. P. Wong. Reduced surface area in apoptotic rounding of human chang liver cells from serum deprivation. Anat. Rec. 240: 456-468, 1994[Medline].

32.   Soma, M. R., R. Baetta, S. Bergamaschi, M. R. De Renzis, C. Davegna, F. Battaini, R. Fumagalli, and S. Govoni. PKC activity in rat C6 glioma cells: changes associated with cell cycle and simvastatin treatment. Biochem. Biophys. Res. Commun. 200: 1143-1149, 1994[Medline].

33.   Takagi, K., Y. Isobe, K. Yasukawa, E. Okouchi, and Y. Suketa. Nitric oxide blocks the cell cycle of mouse macrophage-like cells in the early G2+M phase. FEBS Lett. 340: 159-162, 1994[Medline].

34.  Takahashi, A., H. Yamaguchi, and H. Miyamoto. Change in density of K+ current of HeLa cells during the cell cycle. Jpn. J. Physiol. 44, Suppl. 2: S321-S324, 1994.

35.   Ullrich, N., and H. Sontheimer. Biophysical and pharmacological characterization of chloride currents in human astrocytoma cells. Am. J. Physiol. 270 (Cell Physiol. 39): C1511-C1521, 1996[Abstract/Free Full Text].

36.   Villaz, M., J. C. Cinniger, and W. J. Moody. A voltage-gated chloride channel in ascidian embryos modulated by both the cell cycle clock and cell volume. J. Physiol. (Lond.) 488: 689-699, 1995[Abstract].

37.   Wilson, G. F., and S. Y. Chiu. Mitogenic factors regulate ion channels in Schwann cells cultured from newborn rat sciatic nerve. J. Physiol. (Lond.) 470: 501-520, 1993[Abstract].

38.   Woodfork, K. A., W. F. Wonderlin, V. A. Peterson, and J. S. Strobl. Inhibition of ATP-sensitive potassium channels causes reversible cell-cycle arrest of human breast cancer cells in tissue culture. J. Cell. Physiol. 162: 163-171, 1995[Medline].


AJP Cell Physiol 273(4):C1290-C1297
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