Regulation of ICl,swell in neuroblastoma cells by G protein signaling pathways

Ana Y. Estevez, Tamara Bond, and Kevin Strange

Departments of Anesthesiology and Pharmacology, Anesthesiology Research Division, Laboratories of Cellular and Molecular Physiology, Vanderbilt University Medical Center, Nashville, Tennessee 37232


    ABSTRACT
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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Guanosine 5'-O-(3-thiotriphosphate) (GTPgamma S) activated the ICl,swell anion channel in N1E115 neuroblastoma cells in a swelling-independent manner. GTPgamma S-induced current was unaffected by ATP removal and broadly selective tyrosine kinase inhibitors, demonstrating that phosphorylation events do not regulate G protein-dependent channel activation. Pertussis toxin had no effect on GTPgamma S-induced current. However, cholera toxin inhibited the current ~70%. Exposure of cells to 8-bromoadenosine 3',5'-cyclic monophosphate did not mimic the effect of cholera toxin, and its inhibitory action was not prevented by treatment of cells with an inhibitor of adenylyl cyclase. These results demonstrate that GTPgamma S does not act through Galpha i/o GTPases and that Galpha s/Gbeta gamma G proteins inhibit the channel and/or channel regulatory mechanisms through cAMP-independent mechanisms. Swelling-induced activation of ICl,swell was stimulated two- to threefold by GTPgamma S and inhibited by 10 mM guanosine 5'-O-(2-thiodiphosphate). The Rho GTPase inhibitor Clostridium difficile toxin B inhibited both GTPgamma S- and swelling-induced activation of ICl,swell. Taken together, these findings indicate that Rho GTPase signaling pathways regulate the ICl,swell channel via phosphorylation-independent mechanisms.

cell volume regulation; Rho GTPase; anion channel


    INTRODUCTION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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THE ABILITY TO SENSE AND RESPOND to changes in volume is an essential and fundamental property of all cells (32, 47). In response to cell swelling, most vertebrate cells activate an outwardly rectifying anion current termed ICl,swell. The ICl,swell channel appears to play an important role in regulating cell volume (reviewed in Refs. 43, 50, 57).

In addition to its role in volume homeostasis, the ICl,swell channel may function in other physiological processes and may contribute to organ system pathophysiology. For example, it has been suggested that the ICl,swell channel is a pathway for excitotoxic amino acid release in the central nervous system during cerebral ischemia and trauma (4, 29, 51). ICl,swell is constitutively active in ventricular myocytes isolated from dogs with tachycardia-induced congestive heart failure (CHF), suggesting that the channel contributes to electrophysiological and contractile abnormalities of CHF (11). Swelling-induced taurine release via the ICl,swell channel has been proposed to play a role in controlling osmotic regulation of vasopressin secretion in magnocellular neurons (13). The transformation of microglia from an ameboid to a ramified shape is modulated by a stretch-activated anion channel with biophysical characteristics similar to ICl,swell (20). Changes in cell volume are postulated to play important signaling roles in cell metabolism, excitability, contraction, growth, proliferation, and apoptosis (32, 38, 47). Volume-induced signaling may be mediated in part by changes in ICl,swell activity.

Although volume-sensitive ion channels are expressed ubiquitously and likely play important roles in cellular physiology and pathophysiology, the molecular identity of the channel responsible for ICl,swell is still unknown and the field is fraught with controversy (21, 50, 56). In addition, the signaling mechanisms by which cell swelling is transduced into channel activation are incompletely understood and may vary between different cell types. For example, some studies suggest a requirement for serine/threonine or tyrosine kinase phosphorylation (9, 12, 64) in ICl,swell activation, whereas others have demonstrated that ATP hydrolysis or phosphorylation events are not required (6, 49, 59). In contrast, it has also been suggested that dephosphorylation events mediate activation of the ICl,swell channel (14, 61). The apparent variation in signaling pathways that regulate the channel suggests three possibilities: 1) ICl,swell is due to the activity of more than a single channel type, 2) channel regulation varies between cell types, and/or 3) pharmacological and molecular disruption of signaling pathways has indirect effects on channel activity.

The uncertainty that exists over the signaling mechanisms that regulate ICl,swell and the molecular identity of the channel(s) underscores the need for extensive additional characterization of channel function and regulation. At present, it is known that ICl,swell can be activated by cell swelling (50, 57) and reduced intracellular ionic strength (8, 22, 44). Doroshenko and colleagues (15, 16) demonstrated that guanosine 5'-O-(3-thiotriphosphate) (GTPgamma S) activates an outwardly rectifying anion current with many of the properties of the ICl,swell channel in bovine chromaffin cells. More recently, Nilius and co-workers (46, 64) have shown that activation of G proteins activates ICl,swell in endothelial cells.

The purpose of the present study was to investigate the role of G proteins in regulation of ICl,swell activation in N1E115 neuroblastoma cells. Our results demonstrate that GTPgamma S activates ICl,swell in the absence of swelling. Current activation does not require phosphorylation events and is insensitive to pertussis toxin. However, cholera toxin and Clostridium difficile toxin B significantly inhibited GTPgamma S-induced current activation. Swelling-induced activation of ICl,swell was stimulated by GTPgamma S and inhibited by guanosine 5'-O-(2-thiodiphosphate) (GDPbeta S) and toxin B. Taken together, these results demonstrate that Galpha s/Gbeta gamma and Rho G protein signaling pathways are important regulators of the ICl,swell channel.


    MATERIALS AND METHODS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
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Cell culture. N1E115 mouse neuroblastoma cells (American Type Culture Collection, Manassas, VA) were cultured in the presence of 5% CO2-95% air in high-glucose DMEM (GIBCO, Gaithersburg, MD) containing 25 mM HEPES, 10% fetal bovine serum, 50 U/ml penicillin and 50 µg/ml streptomycin. Cells were used between passages 13 and 34. The osmolality of the growth medium was 295-305 mosmol/kgH2O.

Patch-clamp recordings. N1E115 cells were grown in 35-mm culture dishes and dissociated by brief treatment with Ca2+- and Mg2+-free modified Hanks' solution. Dissociated cells were allowed to reattach to the poly-L-lysine-coated coverslip bottom of a bath chamber (model R-26G; Warner Instrument, Hamden, CT) that was mounted onto the stage of a Nikon TE300 inverted microscope. Patch electrodes were pulled from 1.5-mm-outer diameter borosilicate glass microhematocrit tubes (Fisher Scientific, St. Louis, MO) that had been silanized with dimethyl-dichloro silane (Sigma Chemical, St. Louis, MO). Electrodes were not fire polished before use.

The bath solution contained (in mM) 70 N-methyl-D-glucamine chloride, 5 MgSO4, 12 HEPES, 8 Tris, 5 glucose, 2 glutamine, 120 sucrose, and 0.4 or 1.3 CaCl2, (pH 7.4; osmolality = 300 mosmol/kgH2O). Bath osmolality was altered by increasing or reducing sucrose concentration.

Patch clamping was carried out using a pipette solution that contained (in mM) 125 CsCl, 10 HEPES, 10 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA; tetracesium salt; Molecular Probes, Eugene, OR), 1 MgSO4, 5 CsOH, 2 ATP, and 0.5 GTP or GTPgamma S (tetralithium salt; Sigma; pH 7.2). In some pilot studies, 1 mM EGTA (Sigma) was used instead of BAPTA. To prevent spontaneous cell swelling, the osmolality of the pipette solution was hypotonic (280 mosmol/kgH2O) with respect to the bath.

Experiments where the ATP requirement of channel activation was studied utilized a pipette solution containing 125 mM CsCl, 10 mM HEPES, 10 mM BAPTA, 1 mM EDTA (Sigma), 5 mM CsOH, 0.5 mM GTPgamma S, 40 µM oligomycin, 5 µM iodoacetate, and 20 µM rotenone. ATP or 5'-adenylylimidodiphosphate (AMP-PNP; Boehringer Mannheim, Germany) were added as sodium and lithium salts, respectively. Metabolic inhibitors were added from concentrated stock solutions dissolved in DMSO. Final DMSO concentration in the pipette solutions was 0.2%.

Electrodes had direct current resistances of 3-5 MOmega . Cells were used only if the series resistance was no greater than ~150% of the pipette resistance and the reversal potential was within ±4 mV of the calculated value of +14.7 mV for a perfectly anion-selective channel. Reversal potentials significantly below +14.7 mV were taken as an indication of loss of seal integrity.

An Axopatch 200A (Axon Instruments, Foster City, CA) patch-clamp amplifier was used to voltage clamp N1E115 cells following gigaseal formation and attainment of whole cell access. Command voltage generation, data digitization, and data analysis were carried out on a Pentium II computer using a DigiData 1200 AD/DA interface with pCLAMP 6 software (Axon Instruments). Data were digitized at 5 kHz and filtered at 0.5 kHz using an eight-pole Bessel filter (model 902; Frequency Devices, Haverhill, MA). Electrical connections to the amplifier were made using Ag-AgCl pellets and 3 M KCl-agar bridges. Whole cell currents were measured by varying membrane potential from -80 to +80 mV at 80 mV/s every 5 s.

Measurement of relative cell volume changes. Whole cell currents and volume changes were measured simultaneously in single patch-clamped cells. Cells attached to the coverslip bottom of the patch-clamp bath chamber were visualized by video-enhanced differential interference contrast microscopy. Optical sectioning (58) demonstrated that the cells maintained a spherical morphology for at least 60 min after attachment to the coverslip. Cells were routinely removed from the bath chamber and replaced with fresh cells every 30-45 min. Given that the cells have a spherical morphology, relative cell volume change was determined as
Relative cell volume<IT>=</IT>(experimental CSA/control CSA)<SUP><IT>3/2</IT></SUP> (1)
where CSA is the cell cross-sectional area measured at a single focal plane. In all CSA measurements described in this paper, we imaged cells at focal planes located at the point of maximum cell diameter.

Cell images were recorded continuously throughout a patch-clamp experiment using a super VHS videocassette recorder (model SVO-2000; Sony Electronics, San Jose, CA) and a Hamamatsu charge-coupled device camera (model C2400; Hamamatsu Photonics, Hamamatsu City, Japan). CSAs of single cells were quantified by digitizing recorded video images with an image-processing computer board (MV-1000; MuTech, Woburn, MA) with 512 × 480 × 8-bit resolution and a Pentium II computer. Digitized images were displayed on the computer monitor, and cell borders were traced using a mouse and a computer-generated cursor. The CSA of a traced region was determined by image analysis software (Optimas; Bioscan, Edmonds, WA). This image acquisition and analysis system allows detection of changes in CSA with an accuracy of ±2-3%.

Data analysis. Whole cell currents were recorded within 15-20 s after membrane rupture. The mean resting or baseline current is defined as current measured before activation by GTPgamma S or cell swelling. Baseline current was subtracted from all data points within a given record to correct for variability in resting current levels between different cells. Because of culture-to-culture variability in the response to GTPgamma S, control measurements were performed in parallel with all experimental treatments.

Rates of GTPgamma S current activation and inactivation and peak current were measured. Current activation is defined as the point at which there is a continuous increase in current amplitude above the baseline current (6). Rates of current activation and inactivation were quantified by linear regression analysis.

Under control conditions, a small percentage (<10%) of cells treated with GTPgamma S showed no current activation. To facilitate comparison with experimental treatments that may have inhibited the GTPgamma S response, nonactivating cells were included when calculating the means ± SE rate of GTPgamma S-induced current activation.

Whole cell anion current was also activated by cell swelling. Rates of current activation and cell swelling were determined by linear regression analysis. For these studies, we also quantified the cell volume set point of the channel. Cell volume set point is defined as the relative cell volume at which current activation begins (6).

Throughout the course of the experiments, a small percentage of cells exhibited bleb formation during current recordings. These cells were excluded from our analyses.

Statistical analysis. Data are presented as means ± SE. Statistical significance was determined using Student's two-tailed t-test for unpaired, independent means. When comparing three or more groups, statistical significance was determined by one-way analysis of variance. Values of P < 0.05 were taken to indicate statistical significance.


    RESULTS
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RESULTS
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GTPgamma S activates ICl,swell in the absence of cell swelling. Dialysis of N1E115 cells with 0.5 mM GTPgamma S activated an outwardly rectifying whole cell anion current (Fig. 1A). Current activation began within 0.85 ± 0.08 min after the whole cell configuration was obtained and reached a plateau within 5.3 ± 0.8 min (n = 19; Fig. 1B). Activation was transient, and current levels returned to baseline 11.5 ± 1.2 min (n = 16; Fig. 1B) after the plateau was reached. Cell swelling was not observed during GTPgamma S-induced current activation (Fig. 1B). The mean relative cell volume at the peak GTPgamma S-induced current was 0.99 ± 0.02 (n = 26).


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Fig. 1.   Guanosine 5'-O-(3-thiotriphosphate) (GTPgamma S) activates an outwardly rectifying anion current in N1E115 neuroblastoma cells. A: steady-state current-voltage (I-V) relationship of current activated by GTPgamma S in a single neuroblastoma cell. Calculated reversal potential for a perfectly anion-selective channel is 14.7 mV. Measured reversal potential is 15.2 mV. B: example of simultaneous current and volume measurements performed on a single patch-clamped cell dialyzed with 0.5 mM GTPgamma S. Time 0 refers to the time at which recordings were initiated.

G proteins cycle between an active GTP-bound state and an inactive GDP-bound state. GDPbeta S is a nonhydrolyzable analog of GDP that competes with GTP or GTP analogs for the nucleotide binding sites on G proteins, rendering them inactive. To examine further the role of G proteins in whole cell anion current activation, 10 mM GDPbeta S was included in the pipette solution. In the presence of 10 mM GDPbeta S, only three of six cells (50%) activated spontaneously with GTPgamma S compared with six of six cells in the paired control group. GDPbeta S also inhibited the rate of GTPgamma S- induced current activation and decreased the amplitude of the peak current by 82% and 71%, respectively (Fig. 2).


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Fig. 2.   Guanosine 5'-O-(2-thiodiphosphate) (GDPbeta S) inhibits both the rate of activation and peak GTPgamma S-induced current. For these experiments, the concentration of CsCl in the pipette solution was reduced to 110 mM to osmotically compensate for the GDPbeta S that was added. The control pipette solution contained 110 mM CsCl and 10 mM Li3 citrate to control for the Li+ that was added with GDPbeta S. Values are means ± SE. *P < 0.05; ***P < 0.001. Number of observations (n) is shown in parentheses.

The outwardly rectifying current could be due to activation of ICl,swell or Ca2+-dependent Cl- channels (ICl,Ca). G protein stimulation has been reported to activate ICl,Ca in several (28, 36), but not all (30, 45), cell types. All experiments presented in this paper, however, were carried out using nominal Ca2+ in the bath solution (0.4 mM) and a pipette solution containing the highly selective fast Ca2+ buffer BAPTA (10 mM), demonstrating that ICl,Ca are not responsible for the GTPgamma S-induced current.

Intracellular Ca2+ actually appeared to exert an inhibitory effect on the GTPgamma S current activation. During pilot studies, cells were patch clamped in the presence of 1.3 mM bath Ca2+ and a pipette solution containing 1 mM EGTA instead of 10 mM BAPTA. Peak GTPgamma S-induced currents and rates of current activation and inactivation were unaffected by reduced Ca2+ buffering (data not shown). However, in the presence of 1 mM EGTA, GTPgamma S-induced current activation occurred in <50% of patch-clamped cells. A much more consistent activation of the GTPgamma S current was observed when 10 mM BAPTA was used to buffer intracellular Ca2+. Current activation was detected in 93% (n = 92) of control cells dialyzed with BAPTA-buffered pipette solutions. The reason increased Ca2+ buffering increases the frequency of current activation is unknown. It is conceivable that a Ca2+-dependent process antagonizes the stimulatory effect of GTPgamma S.

To determine whether the GTPgamma S-induced current is due to the activity of the ICl,swell channel, we examined its biophysical characteristics and volume sensitivity. As shown in Table 1, the rectification ratio and relative anion permeability of the channel responsible for the GTPgamma S current are the same as those observed for ICl,swell. Furthermore, neither current exhibited significant voltage-dependent activation or inactivation (Fig. 3A).

                              
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Table 1.   Characteristics of Cl- currents activated by GTPgamma S and cell swelling



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Fig. 3.   Characteristics of GTPgamma S- and swelling-induced anion currents. A: whole cell currents elicited by stepping membrane voltage from -100 to +100 mV in 20-mV steps from a holding potential of 0 mV. ICl,swell was activated by exposing cells to 0.5 mM GTPgamma S or by swelling with a hypotonic bath solution (200 mosmol/kgH2O). The arrows indicate zero-current levels. B: cell shrinkage inhibits the GTPgamma S-induced current. Current and volume measurements shown are for a single patch-clamped cell. Exposure of the cell to a hypertonic bath solution (400 mosmol/kgH2O or 400 mOsm) caused the GTPgamma S-induced current to rapidly inactivate. The mean rate of current inactivation for all experiments was ~8 times faster than spontaneous inactivation observed in the absence of cell shrinkage (see Fig. 1A).

The rate of spontaneous current inactivation (see Fig. 1A) during GTPgamma S stimulation was -0.7 ± 0.2 pA · pF-1 · min-1 (n = 18). To determine whether the GTPgamma S-induced current was volume sensitive, the current was allowed to reach a stable plateau level and cells were then shrunken by exposure to a hypertonic bath solution (400 mosmol/kgH2O). Cell shrinkage inhibited the peak GTPgamma S-induced current by 68 ± 6% (n = 5) at a rate of -5.5 ± 2.3 pA · pF-1 · min-1 (n = 5; see Fig. 3B). The rate of shrinkage-induced inactivation is nearly eight times faster than the rate of spontaneous current inactivation, demonstrating that the GTPgamma S-activated channel is sensitive to cell volume. On the basis of results shown in Figs. 1-3 and Table 1, we conclude that stimulation of G proteins with GTPgamma S activates ICl,swell in the absence of cell swelling.

GTPgamma S-induced ICl,swell activation is not modulated by ATP or phosphorylation reactions. Intracellular ATP and nonhydrolyzable ATP analogs modulate but are not essential for swelling-induced activation of ICl,swell in N1E115 cells (6). In a variety of cell types, nonhydrolyzable ATP analogs support normal ICl,swell activity, indicating that phosphorylation events are not involved in channel activation (6, 49). However, the results of a number of studies have also suggested that protein kinases and phosphatases modulate channel activity (14, 61, 64). Recently, Voets et al. (64) proposed that the stimulatory effect of GTPgamma S on ICl,swell in endothelial cells is mediated by tyrosine phosphorylation.

Given these findings, we examined the effect of tyrosine kinase inhibitors on GTPgamma S-induced activation of ICl,swell in N1E115 cells. Cells were treated with the broadly selective tyrosine kinase inhibitors genistein (100 µM) or tyrphostin A51 (100 µM). Two experimental protocols were used. Cells were patch clamped with inhibitors present only in the pipette solution or with the inhibitors present in both the pipette and bath solutions. In the latter case, cells were preincubated for 7-21 min with inhibitors before patch clamping. Pipette solutions were kept on ice and bath and pipette solutions were remade every hour to minimize problems associated with breakdown of the inhibitors (64). Results using the two protocols were not significantly different for either genistein (P > 0.7) or tyrphostin A51 (P > 0.3), and the data were therefore averaged and are presented in Fig. 4. GTPgamma S activated ICl,swell in all cells treated with tyrphostin A51 and DMSO and in 9 of 10 cells treated with genistein. Neither inhibitor significantly altered the rate of current activation or peak current (Fig. 4).


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Fig. 4.   Tyrosine kinase inhibitors have no significant (P > 0.5) effect on GTPgamma S-induced current activation. Genistein (100 µM) and tyrphostin A51 (100 µM) were dissolved in DMSO and added to the patch pipette and bath solutions at a final DMSO concentration of 0.1%. Control solutions contained 0.1% DMSO. Values are means ± SE. Number of observations (n) is shown in parentheses.

To corroborate the inhibitor studies and to further test for the involvement of phosphorylation events in the GTPgamma S-induced activation of ICl,swell, cells were metabolically poisoned and patch clamped with Mg2+-free pipette solutions containing 1 mM EDTA. Cellular ATP production was blocked by incubating cells for 10-30 min in bath solution containing 5 mM 2-deoxyglucose and 100 nM rotenone. In addition, the pipette solution contained 40 µM oligomycin, 20 µM rotenone, and 5 µM iodoacetate.

In metabolically poisoned cells, removal of ATP from the pipette solution or replacement with 2 mM AMP-PNP had no significant effect on the rate of GTPgamma S-induced ICl,swell activation or peak current (Fig. 5). These results demonstrate clearly that ATP hydrolysis does not play a role in the GTPgamma S signaling pathway. The number of metabolically poisoned cells in which GTPgamma S triggered current activation was 9 of 9 in the presence of 2 mM ATP, 17 of 18 with 0 mM ATP, and 16 of 16 cells with 2 mM AMP-PNP in the pipette solution.


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Fig. 5.   GTPgamma S-induced current activation is not modulated by intracellular ATP or phosphorylation reactions. All cells were patch clamped with a Mg2+-free pipette solution containing the metabolic inhibitors oligomycin (40 µM), rotenone (20 µM), and iodoacetate (5 µM) and were also preincubated for 10-30 min in a bath solution containing 2-deoxyglucose (5 mM) and rotenone (100 nM). ATP removal or substitution with 5'-adenylylimidodiphosphate (AMP-PNP) had no significant (P > 0.1) effect on the rate of activation and peak GTPgamma S-induced current. Values are means ± SE. Number of observations (n) is shown in parentheses.

Cholera toxin inhibits GTPgamma S-induced activation of ICl,swell via cAMP-independent mechanisms. G proteins are categorized into three families: heterotrimeric, low-molecular-weight (small) monomeric, and high-molecular-weight (large) monomeric (3). The heterotrimeric G proteins are composed of three subunits termed alpha , beta , and gamma . Pertussis and cholera toxins are commonly used to determine whether a heterotrimeric G protein family is involved in a specific signaling pathway.

Pertussis toxin catalyzes the ADP-ribosylation and inactivation of members of the Galpha i/o subfamily. To test for the involvement of Galpha i/o G proteins in ICl,swell regulation, N1E115 cells were preincubated with 100 ng/ml pertussis toxin for 6-10 h before patch-clamp measurements were taken. Pretreatment with 1 µg/ml of pertussis toxin for >4 h is sufficient to completely ADP-ribosylate the alpha -subunit of Gi in N1E115 cells (7). Pertussis toxin had no significant (P > 0.5) effect on ICl,swell activation. Rates of GTPgamma S-induced current activation and peak currents (means ± SE) in the presence and absence of pertussis toxin were 1.8 ± 0.4 pA · pF-1 · min-1 (n = 8) and 4.9 ± 0.8 pA/pF (n = 7), and 1.4 ± 0.4 pA · pF-1 · min-1 (n = 10) and 5.0 ± 0.9 pA/pF (n = 5), respectively. These results indicate that that Galpha i/o proteins do not mediate the effect of GTPgamma S.

Cholera toxin catalyzes the ADP-ribosylation of the Galpha s subfamily of G proteins, rendering them constitutively active. Overnight incubation with 100 ng/ml cholera toxin reduced GTPgamma S-induced ICl,swell activation and peak current by ~70% (Fig. 6).


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Fig. 6.   Galpha s signaling pathways inhibit GTPgamma S-induced activation of ICl,swell via cAMP-independent mechanisms. Incubation of cells overnight with 100 ng/ml cholera toxin (CTX) inhibited GTPgamma S-induced current activation ~70%. Exposure of cells to the adenylyl cyclase inhibitor 2'-5'-dideoxyadenosine (DDA; 100 µM) had no effect on the inhibitory action of CTX. Overnight incubation of cells with 500 µM 8-bromoadenosine 3',5'-cyclic monophosphate (8-BrcAMP) did not mimic the effects of CTX treatment. Values are means ± SE. *P < 0.05; **P < 0.01; ***P < 0.001. Number of observations (n) is shown in parentheses.

Activation of Galpha s activates adenylyl cyclase and elevates intracellular cAMP. This suggests that cAMP might mediate the inhibitory effect of cholera toxin. To test whether inhibition occurred via a cAMP-dependent mechanism, cells were incubated overnight with cholera toxin and 2'-5'-dideoxyadenosine (DDA; 100 µM), an inhibitor of adenylyl cyclase. As shown in Fig. 6, the inhibitory effect of cholera toxin was unaltered by DDA.

In an effort to mimic the inhibitory action of cholera toxin, we incubated cells overnight with 500 µM 8-bromoadenosine 3',5'-cyclic monophosphate (8-BrcAMP) and included it in the patch pipette solution. Current activation was not significantly different in 8-BrcAMP-treated cells (Fig. 6). We conclude that Galpha s inhibits GTPgamma S-induced activation of ICl,swell by directly inhibiting the channel and/or channel regulatory mechanisms.

GTPgamma S-induced activation of ICl,swell is mediated by Rho GTPases. The low-molecular-weight monomeric G proteins include the Ras, Rho, Rab, Arf, and Ran families (3). Ten classes of mammalian Rho GTPases have been identified (5): Rho (A, B, C isoforms), Rac, Cdc42, Rnd1/Rho6, Rnd2/Rho7, Rnd3/RhoE, Rho D, RhoG, TC10, and TTF. To test for the involvement of Rho G proteins in ICl,swell regulation, N1E115 cells were incubated with 1 ng/ml C. difficile toxin B for 19-24 h. Toxin B catalyzes the UDP-glucosylation of the Rho subfamily of monomeric G proteins including Rho, Rac, and Cdc42 (1). Incubation with toxin B inhibited the rate of GTPgamma S-induced current activation and peak current ~70% (Fig. 7). These results demonstrate that Rho GTPase signaling pathways regulate GTPgamma S-induced activation of ICl,swell.


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Fig. 7.   GTPgamma S-induced activation of ICl,swell is regulated by Rho GTPases. Incubation of cells overnight with 1 ng/ml C. difficile toxin B inhibited the rate of current activation and peak current. Values are means ± SE. *P < 0.05. Number of observations (n) is shown in parentheses.

Swelling-induced activation of ICl,swell is modulated by Rho signaling pathways. To determine whether G protein signaling pathways modulate swelling-induced activation of ICl,swell, cells were dialyzed with GTPgamma S and exposed to a hypotonic bath (100 mosmol/kgH2O reduction in bath osmolality) after the GTPgamma S-induced current had inactivated (Fig. 8A). Spontaneous inactivation of the GTPgamma S-induced current did not preclude further activation of ICl,swell with a swelling stimulus. The mean ± SE volume set points for current activation in the presence and absence of GTPgamma S were 1.13 ± 0.02 (n = 22) and 1.18 ± 0.03 (n = 9), respectively, and were not significantly (P > 0.1) different. However, GTPgamma S stimulated the rate of swelling-induced current activation two- to threefold (Fig. 8A).


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Fig. 8.   G protein signaling pathways regulate swelling-induced activation of ICl,swell. A: effect of GTPgamma S treatment on swelling-induced current activation in a single N1E115 cell. After decay of the GTPgamma S current, cell was swollen by exposure to a hypotonic (200 mosmol/kgH2O) bath solution. Inset: mean rates of swelling-induced current activation in cells dialyzed with 0.5 mM GTP or GTPgamma S. GTPgamma S increased rate of swelling-induced current activation 2- to 3-fold. B: dialysis of cells with 10 mM GDPbeta S or overnight exposure to 10 ng/ml toxin B inhibited swelling-induced ICl,swell activation. For experiments with GDPbeta S, CsCl concentration in the pipette solution was reduced to 110 mM to maintain osmolality. The control pipette solution contained 110 mM CsCl and 10 mM Li3 citrate to control for the Li+ that was added with GDPbeta S. Values are means ± SE. *P < 0.03; ***P < 0.001. Number of observations (n) is shown in parentheses.

To determine whether G proteins are required for swelling-induced current activation, 10 mM GDPbeta S was included in the pipette solution in the absence of GTPgamma S. GDPbeta S inhibited swelling-induced ICl,swell activation by ~80% (Fig. 8B) and significantly (P < 0.002) increased mean ± SE channel volume set point from 1.1 ± 0.02 (n = 4) to 1.22 ± 0.01 (n = 4). Overnight exposure of cells to 10 ng/ml toxin B inhibited current activation ~70% (Fig. 8B) without altering channel volume set point (control = 1.1 ± 0.02, n = 7; toxin B = 1.1 ± 0.03, n = 5). Taken together, these data suggest strongly that Rho GTPase signaling pathways regulate swelling-induced activation of ICl,swell.


    DISCUSSION
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ABSTRACT
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ICl,swell is a swelling-activated anion current that appears to be expressed ubiquitously in mammalian cells (43, 50, 57). This current is outwardly rectifying, exhibits an Eisenman type I anion permeability sequence, and is inhibited by a wide variety of pharmacological agents. The ICl,swell channel plays an important role in cell volume regulation (reviewed in Refs. 43, 50, 57) and may participate in the control of other physiological processes such as cell metabolism, membrane excitability, and cell growth, proliferation, and apoptosis (32, 38, 47).

The molecular identity of the channel responsible for ICl,swell is uncertain and controversial. P-glycoprotein and pICln have both been suggested to function as the ICl,swell channel. However, most workers in the field no longer consider these proteins to be viable channel candidates (21, 50, 56). More recently, ICl,swell was proposed to be due to the activity of ClC-3, a member of the ClC superfamily of voltage-gated anion channels (19). The findings of Duan and co-workers (19) on ClC-3 have not yet been reproduced by other laboratories, and a variety of recent observations have begun to raise doubts about a widespread role for this channel in ICl,swell function (35, 48, 65).

Regulation of ICl,swell. Cell swelling and reduced intracellular ionic strength activate ICl,swell (8, 22, 44, 50, 57). The signal transduction mechanisms responsible for channel activation are unclear. Recently, Nilius and co-workers (46) demonstrated that ICl,swell in endothelial cells is activated transiently by GTPgamma S in a pertussis toxin-insensitive manner. A similar outwardly rectifying anion current exhibiting many of the basic properties of ICl,swell was originally shown by Doroshenko and colleagues (15, 16) to be triggered by GTPgamma S in bovine chromaffin cells.

ICl,swell in N1E115 neuroblastoma cells is also activated transiently by GTPgamma S (Fig. 1). Activation of ICl,swell by swelling is dramatically stimulated by GTPgamma S (Fig. 8A) and inhibited by GDPbeta S (Fig. 8B). Taken together, these results indicate that cell swelling is transduced into channel activation at least in part via G protein signaling pathways.

GTPgamma S current activation occurs via a pertussis toxin-insensitive mechanism. However, cholera toxin significantly inhibited GTPgamma S-induced current development, suggesting that Galpha s regulates ICl,swell (Fig. 6). Activation of Galpha s stimulates adenylyl cyclase. Several studies have recently demonstrated that ICl,swell is modulated by cAMP. Du and Sorota (17) observed both inhibitory and stimulatory effects of cAMP in dog atrial cells. They showed that inhibition of ICl,swell is due to cAMP-induced activation of protein kinase A (PKA) and increased protein phosphorylation, whereas the stimulatory effect of cAMP occurs in a phosphorylation-independent fashion. More recently, Shimizu et al. (55) demonstrated that cAMP enhances ICl,swell activation in Intestine 407 cells by a PKA-independent mechanism.

We tested for the involvement of adenylyl cyclase and cAMP in mediating the effect of cholera toxin on GTPgamma S-induced activation of ICl,swell. The inhibitory action of cholera toxin was not mimicked by overnight exposure to 8-BrcAMP and was not blocked by the adenylyl cyclase inhibitor DDA (Fig. 6), indicating that Galpha s functions through cAMP-independent pathways.

Recent studies have demonstrated that Galpha s is capable of modulating ion channel activity in the absence of adenylyl cyclase and PKA function (e.g., Refs. 31, 37). Galpha s may directly inhibit the ICl,swell channel and/or may act on signaling pathways that regulate channel activation. Gbeta gamma may also inhibit ICl,swell in a cAMP-independent manner. It has been demonstrated that Gbeta gamma subunits can directly modulate ion channel activity in a stimulatory or inhibitory fashion (10, 52). It is interesting to speculate that spontaneous inactivation of ICl,swell (Fig. 1) may be mediated by GTPgamma S stimulation of Galpha s or Gbeta gamma signaling mechanisms. Extensive molecular biological studies are required to fully determine which heterotrimeric subunit inhibits ICl,swell activity and to delineate the mechanism by which this inhibition occurs.

Regulation of ICl,swell is mediated at least in part by small monomeric Rho GTPases. C. difficile toxin B inhibited both GTPgamma S- and swelling-induced channel activation (Figs. 7 and 8). Toxin B inhibits Rho, Rac, and Cdc42 Rho GTPases (1). Recently, Nilius et al. (46) demonstrated that ICl,swell in endothelial cells is inhibited by Clostridium C3 exoenzyme. The C3 exoenzyme is a selective inhibitor of Rho A, B, and C (1). C3 exoenzyme does not readily permeate plasma membranes, and, in our study, we were unable to ensure that it was loaded effectively into N1E115 cells. However, assuming that ICl,swell in endothelial and N1E115 cells are controlled by similar mechanisms, our findings in conjunction with those of Nilius et al. (46) argue that Rho A, B, and/or C are important regulators of this current.

The molecular details of how Rho GTPase signaling pathways regulate ICl,swell are unknown. Rho GTPases have been implicated in a variety of cellular processes including actin cytoskeletal organization, membrane trafficking, transcriptional activation, cell growth, motility, and morphogenesis (24, 53, 63). Alterations in cytoskeletal organization regulated by Rho GTPases have been studied extensively and include the formation of focal adhesions, actin stress fibers, lamellipodia, membrane ruffles, and filopodia (24, 42, 53, 60).

Changes in the organization of the actin cytoskeleton have long been implicated in regulating volume-sensitive transport pathways (32, 41, 50, 57), including the ICl,swell channel (34, 54, 62). Mechanical forces have been shown to directly modulate G protein activity (23) as well as cytoskeletal architecture (25, 26). It is attractive to postulate then that swelling-induced alterations in Rho GTPase activity may alter cytoskeletal structure, which in turn triggers ICl,swell activation. Alternatively, cell swelling may directly alter the organization of the cytoskeleton. Cytoskeletal changes could conceivably activate Rho G protein signaling pathways that regulate ICl,swell.

Role of protein phosphorylation in ICl,swell regulation. Phosphorylation has emerged as an extremely confounding variable in understanding how ICl,swell is regulated. In numerous cell types, nonhydrolyzable ATP analogs support normal swelling-induced current activation (6, 49, 50, 57), an observation that argues strongly against a role for phosphorylation events in channel regulation. However, serine/threonine phosphorylation (e.g., Refs. 9, 40), serine/threonine dephosphorylation (18, 19), tyrosine phosphorylation (e.g., Refs. 12, 33), and tyrosine dephosphorylation (14, 61) have been proposed to play roles in channel activation.

Even experiments from the same laboratory have generated confounding results. Szücs et al. (59) failed to detect an inhibitory effect of the tyrosine kinase inhibitor genistein on swelling-induced ICl,swell activation in bovine endothelial cells. However, more recently, Voets et al. (64) demonstrated that this compound inhibited activation of the current by both GTPgamma S and swelling. Voets et al. (64) suggested that the discrepant findings may have been due to low solubility and stability of genistein.

In our study, we were unable to detect any inhibitory action of genistein or tyrphostin A51 on GTPgamma S-induced activation of ICl,swell (Fig. 4). Because of potential problems associated with the use of these drugs (see Ref. 64), we examined the combined effects of intracellular ATP and Mg2+ removal on current activation. As shown in Fig. 5, ATP and Mg2+ removal or replacement of ATP with the nonhydrolyzable analog AMP-PNP in metabolically poisoned cells has no effect on GTPgamma S-induced current activation. Similarly, we have shown previously that swelling-induced ICl,swell activation in neuroblastoma cells occurs normally in the absence of hydrolyzable ATP (6). Indeed, even dialysis of metabolically poisoned cells with Mg2+- and ATP-free pipette solutions containing AMP-PNP and alkaline phosphatase to dephosphorylate proteins has no effect on current activation (6). On the basis of these results, we conclude that phosphorylation signaling pathways do not regulate ICl,swell in N1E115 neuroblastoma cells during GTPgamma S- or swelling-induced activation.

The requirement for phosphorylation observed in other cell types may reflect the existence of distinct channel types. Alternatively, it may reflect the existence of multiple signaling/regulatory pathways involved in channel activation. These pathways could be cell specific, they may reflect the physiological status of the cell, and/or they may be sensitive to experimental parameters such as the mechanism or rate of cell swelling (see Ref. 6). It is also distinctly possible that pharmacological agents used to inhibit kinases and phosphatases may directly block the channel, such as has been shown for inhibitors of arachidonic acid metabolism (39), or they may have other nonspecific effects. Conclusions drawn from pharmacological studies of phosphorylation-dependent regulation of ICl,swell should be corroborated where possible by metabolic inhibition and Mg2+ and ATP removal experiments such as those shown in Fig. 5 and described by Bond et al. (6).

Downstream effectors of Rho GTPases include various protein and lipid kinases (2, 5), a finding consistent with the postulated role of Rho kinases in regulating ICl,swell in endothelial cells (46). However, Rho GTPases can also regulate cellular processes in a phosphorylation-independent manner. For example, Rho GTPase-regulated actin polymerization and cross-linking in vitro occur in the absence of ATP and phosphorylation reactions (27). The nonkinase effectors of Rho GTPases include various scaffolding proteins that play important roles in actin cytoskeletal organization (2, 5). Phosphorylation-independent activation of ICl,swell in N1E115 cells may be mediated by changes in the interaction of Rho-regulated scaffolding proteins with the ICl,swell channel and/or associated regulatory machinery.

To conclude, we have demonstrated that ICl,swell in neuroblastoma cells is regulated by G protein signaling pathways. Swelling- and GTPgamma S-induced channel activation are mediated at least in part by Rho GTPases. Extensive additional studies utilizing molecular and electrophysiological approaches are required to fully elucidate the mechanisms by which G protein-dependent regulation occurs.


    ACKNOWLEDGEMENTS

This work was supported by National Institutes of Health Grants NS-30591 and DK-51610. A.Y. Estevez was supported by a National Science Foundation postdoctoral fellowship. T. Bond was supported by a Stroke Investigator Award from the Heart and Stroke Foundation of Ontario.


    FOOTNOTES

Address for reprint requests and other correspondence: K. Strange, Vanderbilt Univ. Medical Center, Anesthesiology Research Division, T-4202 Medical Center North, Nashville, TN 37232-2520 (E-mail: kevin.strange{at}mcmail.vanderbilt.edu).

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 10 October 2000; accepted in final form 31 January 2001.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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