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Am J Physiol Renal Physiol 292: F628-F638, 2007. First published September 26, 2006; doi:10.1152/ajprenal.00132.2006
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Extracellular pH alkalinization by Cl–/HCO3– exchanger is crucial for TASK2 activation by hypotonic shock in proximal cell lines from mouse kidney

S. L'Hoste,1 H. Barriere,1 R. Belfodil,1 I. Rubera,1 C. Duranton,1 M. Tauc,1 C. Poujeol,1 J. Barhanin,2 and P. Poujeol1

1UMR Centre National de la Recherche Scientifique 6548, Université de Nice-Sophia Antipolis, Nice, and 2Institut de Pharmacologie du Centre National de la Recherche Scientifique, Valbonne Sophia-Antipolis, France

Submitted 18 April 2006 ; accepted in final form 20 September 2006


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
We have previously shown that K+-selective TASK2 channels and swelling-activated Cl– currents are involved in a regulatory volume decrease (RVD; Barriere H, Belfodil R, Rubera I, Tauc M, Lesage F, Poujeol C, Guy N, Barhanin J, Poujeol P. J Gen Physiol 122: 177–190, 2003; Belfodil R, Barriere H, Rubera I, Tauc M, Poujeol C, Bidet M, Poujeol P. Am J Physiol Renal Physiol 284: F812–F828, 2003). The aim of this study was to determine the mechanism responsible for the activation of TASK2 channels during RVD in proximal cell lines from mouse kidney. For this purpose, the patch-clamp whole-cell technique was used to test the effect of pH and the buffering capacity of external bath on Cl– and K+ currents during hypotonic shock. In the presence of a high buffer concentration (30 mM HEPES), the cells did not undergo RVD and did not develop outward K+ currents (TASK2). Interestingly, the hypotonic shock reduced the cytosolic pH (pHi) and increased the external pH (pHe) in wild-type but not in cftr –/– cells. The inhibitory effect of DIDS suggests that the acidification of pHi and the alkalinization of pHe induced by hypotonicity in wild-type cells could be due to an exit of HCO3–. In conclusion, these results indicate that Cl– influx will be the driving force for HCO3– exit through the activation of the Cl–/HCO3– exchanger. This efflux of HCO3– then alkalinizes pHe, which in turn activates TASK2 channels.

regulatory volume decrease; CFTR; KCNE5; potassium and chloride channels; external and internal pH

IT IS NOW WELL ESTABLISHED that the control of cell volume is an essential phenomenon, which maintains the homeostasis of numerous cell functions. Most animal cells ensure maintenance of their volume by controlling ion movements across their plasma membrane (19), in response to osmolarity changes in the extracellular medium. This is achieved through the activation of specific channels and transporters. Epithelial cells, such as those that constitute the different segments of the nephron, are particularly exposed to variations in extracellular osmolarity (11). These cells are subjected to an osmotic shock, either by accumulation of active osmolytes inside their cytoplasm (proximal tubule) or by dilution of the tubular fluid (distal tubule). In response to osmotic stress, these cells undergo a regulatory volume decrease (RVD) process characterized by the exit of Cl– and K+ ions, which finally drives water efflux (20). However, despite an impressive quantity of data in the literature, the molecular identity of the Cl– and K+ channels activated to achieve this regulation remains unclear (22). Moreover, the mechanisms precluding the activation of these channels are still controversial. This is probably due to the fact that, although RVD is an ubiquitous phenomenon, the nature of the channels involved in this process could vary depending on the tissue under study (25). In the mouse proximal tubule, we previously demonstrated that during a hypotonic shock CFTR modulated the swelling-activated Cl– currents by controlling a cascade involving apical ATP release, adenosine production, and Ca2+ entry (3). Concomitantly, the decrease in tonicity activated a K+ conductance through TASK2 channels (2, 5). Further work provided evidence that TASK2 channels could be involved in cell RVD. Moreover, these channels are known to be sensitive to external pH (pHe) (13, 26, 29, 30, 32). TASK2 channels belong to the large family of two-P domain K+ channels, some members of which are activated by cell swelling. Nevertheless, the exact mechanism responsible for the activation of TASK2 channels during RVD is unknown. Therefore, it was of interest to determine why hypotonicity could increase TASK2 K+ conductance. The high sensitivity of these currents to pHe suggests that the increase in pHe induced by HCO3– efflux activates TASK2. The related K+ permeability of TASK2 is concomitantly increased by hypotonic shock. In the proximal tubule, an increase in basolateral pHe could be due to HCO3– transport (42). Such transport is achieved by the Na+-3HCO3– cotransporter and by the Cl–/HCO3– exchanger. A specific coupling of TASK2 activity to HCO3– transport through external alkalinization was demonstrated by Warth et al. (42). Using isotonic conditions, these authors proposed a model of TASK2 function in the proximal tubule in which Na+ and HCO3– leave the cell by Na+-3HCO3– cotransport. Under hypotonic conditions, the involvement of HCO3– in RVD was also clearly established in different epithelial-derived cells (28, 36). However, although most of the authors agree with the observation that the presence of HCO3– is absolutely required for ensuring RVD, the relative participation of the different HCO3– transporters is still being discussed. In the present study, we examined the mechanism involved in the exit of HCO3– during a hypotonic shock in cell lines originating from mouse proximal tubule.

Here, we show that hypotonicity-induced cell swelling is followed by a rapid decrease in internal pH (pHi) and a concomitant increase in pHe due to an efflux of HCO3–. This Cl–-dependent and Na+-independent efflux could be due to selective activation of the Cl–/HCO3– exchanger. The increase in pHe activates TASK2 K+ channels, allowing the development of swelling-activated K+ currents.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Transformation of Primary Cultures with pSV3 neo

The primary cell culture technique has been described in detail in previous studies (4, 5). Ten-day-old primary cultures of S1 and S2 segments of proximal tubules from wild-type, cftr–/– (35), and task2–/– mice were transfected with pSV3 neo using lipofectin (Invitrogen). After 48 h, selection of the clones was performed by the addition of 500 µg/ml G418. Culture medium M1 containing G418 was changed every day. Resistant clones were isolated, subcultured, and used after 10 trypsinization steps.

BCECF Cell Volume Measurement

The relative cell volume was monitored by image analysis with BCECF-AM used as a fluorescent volume indicator, as previously reported (31, 37). At 450 nm (isobestic point), BCECF fluorescence is pH insensitive. The emitted fluorescent signal at 520 nm was therefore related to the intracellular dye concentration only and reflected the variations in cell volume (37). Briefly, proximal cell lines grown in 35-mm petri dishes were incubated in the presence of 4 µM BCECF-AM (final concentration) at 37°C for 15 min in a humidified atmosphere of 5% CO2-95% air. Cells were then incubated in isotonic HCO3–-buffered solutions containing (in mM) 80 NaCl, 15 NaHCO3, 5 KCl, 1 CaCl2, with low (1 HEPES) or high buffering capacity (30 HEPES), pH 7.4, in a 5% CO2 atmosphere (Fig. 1B). These solutions were adjusted to 290 or 230 mosmol/kgH2O by addition of mannitol. The relative change in cell volume was estimated from the fluorescent signal by assuming that a 30% decrease in osmolality caused a decrease in the fluorescent signal corresponding to a minimum swelling of 30% of the initial volume. The means of relative volume changes were obtained by analysis of 10–25 cells in each culture. Cell volume variations were calibrated according to the method described by Tauc et al. (37).


Figure 1
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  		Fig. 1. Effect of pH and buffering capacity of external bath solution on regulatory volume decrease (RVD). A: effect of external pH (pHe) on hypotonicity-induced RVD in cultured proximal convoluted tubule (PCT) cell line from wild-type mice. Cell volume was measured using BCECF-AM. After a control period in an isotonic bath solution (290 mosmol/kgH2O), a hypotonic shock was induced by perfusing the cells with a hypotonic bath solution (230 mosmol/kgH2O). Relative volume changes (±SE) as the percentage of initial volume were plotted against time. Experiments were performed at various extracellular pH, ranging from 6.0 to 8.0 as indicated. Measurements were performed on 8 different monolayers (19–25 random cells each) at each pH. B: effect of buffering capacity of external bath on RVD in proximal cell line from wild-type mice. After a control period with cells in an isotonic bath solution (290 mosmol/kgH2O), a hypotonic shock was induced by perfusing the cells with a hypotonic bath solution (230 mosmol/kgH2O) containing either 1 or 30 mM HEPES at pH 7.4 or 30 mM HEPES at pH 8.0. Relative volume changes (±SE) as the percentage of initial volume were plotted against time. Measurements were performed on 8 different monolayers (21–23 random cells each) at each HEPES concentration.

 
pHi Measurements

The fluorescent pH indicator BCECF-AM was also used to measure pHi, as described in detail previously (7). Proximal cell lines grown in 35-mm petri dishes were incubated in the presence of 4 µM BCECF-AM at 37°C for 15 min in a humidified atmosphere of 5% CO2-95% air. Loaded cells were carefully rinsed and placed on the stage of an inverted microscope. The cells were excited successively at 450 and 490 nm, and the emitted light was recorded at 520 nm. At the end of each experiment, the fluorescence signals related to pHi changes were calibrated using the K+/H+ exchange ionophore nigericin. For this purpose, the cells were perfused with KCl solutions containing (in mM) 140 KCl, 1 CaCl2, and 10 HEPES, the pH of which was adjusted to 8.0, 7.0, and 6.0, respectively, with Tris buffer. Finally, 10 µM nigericin was added to all of these solutions.

The cell-buffering capacity was determined by the NH4+ technique (15, 33). pHi was recorded after addition of 20 mM NH4Cl. The buffering capacity was calculated according to the method of Roos and Boron (33). From the above two parameters (pHi and buffering capacity), the H+ efflux as a function of time was calculated as follow: H+ efflux = buffering capacity x {Delta}pHi (in mmol·l–1·min–1). Under all experimental conditions, the fluorescence was recorded continuously and {Delta}pHi was calculated using the first 30-s initial rate of H+ efflux (6). The initial rate of the change in pHi ({Delta}pHi/min) was calculated using linear regression analysis. To ensure an adequate renewal of the medium, the solutions were perfused at a rate of 2 ml/min.

To determine the activity of the Cl–/HCO3– exchanger at physiological pH values, cells were first incubated in HCO3–-buffered NaCl solution containing (in mM) 125 NaCl, 15 NaHCO3, 5 KCl, 1 CaCl2, 5 glucose, and 10 HEPES, pH 7.4. The solution was then replaced by a Cl –-free solution containing (in mM) 125 sodium gluconate, 15 NaHCO3, 5 potassium gluconate, 3 calcium gluconate, 5 glucose, and 10 HEPES, pH 7.4. Under these conditions, intracellular Cl– is rapidly exchanged against extracellular HCO3–, and pHi is increased.

pHe Measurement

The nonesterified form of BCECF was used to assess pHe. Proximal cell lines from wild-type, task2 –/–, and cftr –/– mice were grown on 100-mm petri dishes. After trypsinization, cells were centrifuged and resuspended in 500 µl of either isotonic or hypotonic solutions containing (in mM) 105 NaCl, 5 KCl, 1 CaCl2, 5 glucose, and 1 HEPES, pH 7.0; when necessary, isotonicity was adjusted by adding 72 mM mannitol. The suspension was adjusted to a final number of 2 x 107 cells/ml. Cells were maintained in isotonic or hypotonic medium for 8 min in a humidified atmosphere of 5% CO2-95% air. Cells were then centrifuged at 1,300 g for 2 min, and BCECF (25 µM) was added to the supernatant. The fluorescence of the supernatant was rapidly measured using a spectrofluorimeter (Safas, Monaco). For this purpose, the supernatants were excited successively at 490 and 450 nm and the emitted fluorescence was recorded at 520 nm. At the end of each experiment, the fluorescence values were converted to pH units using calibration curves performed with solutions adjusted at pHe ranging from 6.0 to 8.0.

Electrophysiological Studies

Whole-cell currents were recorded from cultured cells grown on collagen-coated supports (35-mm petri dishes) maintained at 37°C for the duration of the experiments. The ruptured whole-cell configuration of the patch-clamp technique was used. After formation of a gigaseal, the fast-compensation system of the amplifier was used to compensate for the intrinsic input capacitance of the head stage and the pipette capacitance. The membrane was ruptured by additional suction to achieve the conventional whole-cell configuration. Settings available on the amplifier were used to compensate for cell capacitance. The series resistances were not compensated, but experiments in which the series resistance was >20 M{Omega} were discarded. The offset potentials between both electrodes were zeroed before sealing, and the liquid junction potentials were measured experimentally before each experiment and corrected accordingly (measured junction potentials were negligible for Cl– conductance experiments and were 9.96 ± 0.91 mV for K+ conductance experiments). Solutions were perfused in the extracellular bath using a four-channel glass pipette, with the tip placed as close as possible to the clamped cell. Voltage-clamp commands, data acquisition, and data analysis were controlled via a VP 500 amplifier (Biologic) connected to a computer. The whole-cell currents resulting from voltage stimuli were sampled at 2.5 kHz and filtered at 1 kHz. Cells were held at –50 mV, and 400-ms pulses from –100 to +100 or +120 mV were applied in 20-mV increments.

The composition of the pipette and bath solutions is described in Table 1.


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  		Table 1. Composition of solutions used in whole-cell clamp experiments

 
Chemical Compounds

5-Nitro-2-(3-phenylpropylamino)-benzoic acid (NPPB; Calbiochem) was prepared at 100 mM in DMSO. DIDS or 4–4'-dinitrostilbene-2, 2-disulfonic acid (DNDS) was directly dissolved in the medium at a final concentration of 1 mM. Diphenylamine-2-carboxylate (DPC) was prepared at 1 M stock solution in DMSO.

DIDS, DPC, and forskolin were obtained from Sigma-Aldrich (Saint Quentin Fallavier, France), while DNDS, BCECF-AM, and BCECF were obtained from Molecular Probes (Leiden, The Netherlands). Clofilium was prepared at 10 mM in 50% DMSO-50% water. Clofilium was a gift from Dr. Barhanin (UMR CNRS 6097).


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Effect of pH and Buffering Capacity of External Bath Solution on RVD

Relative cell volume variation during a hypotonic shock was measured using fluorescence-videomicroscopy in proximal cell lines from wild-type mice. The cell monolayers were bathed in an isotonic solution (290 mosmol/kgH2O) and then perfused continuously with a hypotonic solution (230 mosmol/kgH2O). As illustrated in Fig. 1A, RVD was recorded at different pHe, varying from 6.0 to 8.0. Interestingly, the RVD phenomenon was minimal at pHe = 6.0 and maximal at pHe = 8.0.

Based on this result, we investigated whether an increase in pHe was necessary to ensure RVD. For this purpose, RVD was recorded in cells bathed in weakly (1 mM HEPES) or highly (30 mM HEPES) buffered solutions (Fig. 1B). In the presence of a highly buffered solution, the cells did not undergo RVD, indicating that pHe variations influenced the RVD response. To exclude the possibility that high concentrations of HEPES inhibited the RVD phenomenon, relative cell volume variation was recorded in a highly buffered bath solution (30 mM), with pH adjusted to 8.0. Under this condition, cells underwent classic RVD, as illustrated in Fig. 1B. Taken together, these results confirmed that external alkalinization was required to ensure RVD during a hypotonic shock. Experiments were then performed to identify the precise mechanism involved in this alkalinization.

Effect of Cl– and Na+ Removal on RVD

In the proximal tubule, pHe is modulated by the exit of HCO3– mainly through Na+-3HCO3– cotransport and/or Cl–/HCO3– exchangers. To test the requirement of Na+ and Cl– during the RVD phenomenon, relative cell volume variations during a hypotonic shock were determined in the presence or absence of Na+ or Cl–, respectively. In the experiments described in Fig. 2A, proximal monolayers from wild-type mice were first incubated for 60 min in a Na+-free bath solution (pH = 7.4) and the hypotonic shock was then performed in the absence of Na+ in the bath solution. The removal of Na+ did not significantly modify the RVD process. In a second set of experiments (Fig. 2B), the hypotonic shock was performed in the absence of external Cl– (pH 7.4). Under these conditions, the cells became swollen but did not exhibit RVD, indicating that Cl– contributed to the RVD. However, the absence of RVD in Cl–-free solutions could be the consequence of two processes. 1) RVD depended on the efflux of Cl– through Cl– channels only. In this case, the massive loss of intracellular Cl– due to the acute removal of external Cl– could decrease the intracellular Cl– ion pool and strongly reduced the activity of volume-sensitive Cl– channels. 2) RVD depended on both an efflux of Cl– through the volume-sensitive Cl– channel and on the efflux of HCO3– through the Cl–/HCO3– exchanger, which could increase pHe. In this case, the removal of external Cl– could decrease this HCO3– efflux and could reverse the driving force of Cl–. Consequently, HCO3– entry precluded pHe alkalinization.


Figure 2
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  		Fig. 2. Effect of Cl– and Na+ removal on RVD. A: effect of Na+ substitution on hypotonicity-induced RVD in cultured PCT cell line from wild-type mice. After a control period (isotonic bath solution, 290 mosmol/kgH2O), a hypotonic shock was induced by perfusing the cells with a hypotonic bath solution (230 mosmol/kgH2O) in the presence or absence of Na+. Measurements were performed on 7 different monolayers with or without Na+ (20–23 random cells from each monolayer). B: effect of Cl– substitution on hypotonicity-induced RVD in cultured PCT cell line from wild-type mice. After a control period (isotonic bath solution, 290 mosmol/kgH2O), a hypotonic shock was induced by perfusing the cells with a hypotonic bath solution (230 mosmol/kgH2O) in the presence or absence of Cl–. Measurements were performed on 7 different monolayers with or without Cl– (19–22 random cells from each monolayer). C: effect of extracellular alkalinization on hypotonicity-induced RVD in cultured PCT cell line from wild-type mice. Experiments were performed as in B using an external bath solution adjusted to pH 8.0 instead of 7.4. Measurements were performed on 6 different monolayers with or without Cl– (10–20 random cells from each monolayer).

 
To verify these hypotheses, the RVD phenomenon was assessed in cells suspended in a Cl–-free solution with pH adjusted to 8.0. As illustrated in Fig. 2C, the hypotonic shock was followed by a RVD. This observation indicated that when the external bath pH was maintained at alkaline pH, the acute Cl– removal did not inhibit the RVD process. Thus, in the experiments described in Fig. 2B, the absence of RVD was probably the consequence of an alkalinization defect of pHe due to a dysfunction of the Cl–/HCO3– exchanger (see hypothesis 2, above).

Involvement of Cl–/HCO3– Exchanger in RVD

The above experiments suggested the involvement of the Cl–/HCO3– exchanger in the RVD phenomenon. To confirm this hypothesis, relative cell volume variations during a hypotonic shock were measured in a proximal tubule cell line in the presence of DIDS or DNDS (DIDS analog). Two experimental conditions were used. In a first set of experiments, the hypotonic shock was performed with an acute application of DIDS or DNDS. As expected, each inhibitor prevented RVD during the hypotonic shock (Fig. 3). In a second set of experiments, cells were preincubated for 60 min with DIDS or DNDS and thoroughly rinsed for 10 min to remove these drugs. Afterward, a hypotonic shock was applied with an inhibitor-free solution. Preincubation with DIDS prevented the RVD (Fig. 3). In contrast, cells preincubated with DNDS returned to their initial volume in response to the hypotonic shock. As expected, the absence of RVD during the acute application of DIDS or DNDS was mainly due to an inhibition of the volume-sensitive Cl– channels since these drugs are potent inhibitors of these channels in most cells. By contrast, the absence of RVD in cells preincubated with DIDS and rinsed out is probably due to a specific inhibition of the Cl–/HCO3– exchanger because sustained DIDS application could irreversibly block this exchanger.


Figure 3
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  		Fig. 3. Involvement of Cl –/HCO3– exchanger activity on RVD in cultured PCT cell lines from wild-type mice. Cell volume was measured by an electronic sizing technique with a CASY 1 cell counter (Schärfe System). Cells were suspended in casyton solution (NaCl isotonic solution) for control condition (bullet) and then in 30% diluted casyton solution in the absence or presence of DIDS or 4–4'-dinitrostilbene-2, 2-disulfonic acid (DNDS). Values are expressed as percentage of cell volume variations measured during hypotonic shock in the absence of inhibitors ({circ}), in the presence of DIDS (1 mM, {square}), or DNDS (1 mM, {blacksquare}). Cell volume measurements were also performed with cells preincubated for 1 h with DIDS (1 mM, {triangleup}) or DNDS (1 mM, {blacktriangledown}) and then thoroughly washed to remove the inhibitors. Values are means ± SE of 5–7 different experiments.

 
Cl–/HCO3– Exchanger Activity in Proximal Cell Lines

To further confirm this possibility, we estimated the activity of this exchanger by recording pHi variations during external Cl– removal. For this purpose, a proximal cell line from wild-type mice loaded with BCECF was maintained in NaCl solutions containing HCO3– and continuously gassed with 5% CO2. The pHi was recorded by fluorescence microscopy. Figure 4A gives an example of the time course of the pHi variations. In a first step, external Cl– was replaced by gluconate, causing pHi to increase over a 6-min period. This effect was reversed when the gluconate solution was replaced with the NaCl solution. The initial rates of pHi increase are reported in Fig. 4B. Compared with the control condition, the acute application of DIDS or DNDS strongly blocked the pHi increase induced by Cl– removal. In the second set of experiments, the cells were treated with DIDS or DNDS for 60 min. Inhibitors were then removed from the solution, and the pHi increase was recorded. Figure 4B clearly shows that sustained incubation with DIDS precluded the pHi increase, whereas incubation with DNDS did not modify the pHi increase induced by external Cl– removal. Taken together, these data suggested that during chronic application, DIDS bound covalently to the Cl–/HCO3– exchanger and irreversibly inhibited its activity. This inhibition resulted in a concomitant inhibition of the RVD, confirming the involvement of this exchanger in the control of cell volume during a hypotonic shock.


Figure 4
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  		Fig. 4. Measurement of internal (cytosolic) pH (pHi) variations in proximal cell lines from wild-type, task2 –/–, and cftr –/– mice during extracellular Cl– removal. A: cells were loaded for 15 min with BCECF, and the fluorescence was recorded during repetitive replacements of the bath by Na gluconate solution (Cl– free) or NaCl solution in the absence or presence of DIDS (1 mM). The sequence of bath solution substitution is indicated (top). All solutions were buffered with NaHCO3 (15 mM), and experiments were performed at constant PCO2 (5%).Values are means ± SE of 6 monolayers. B: initial rate of pHi increment ({Delta}pH/min) was measured after replacement of external NaCl solution by a Cl–-free solution (Na-gluconate) in the absence (control) or presence of DIDS (1 mM) or DNDS (1 mM). Additional experiments were performed with cells preincubated for 1 h with DIDS and DNDS and then thoroughly washed to remove the inhibitors. Values are means ± SE of 6 different experiments for each experimental conditions. C: {Delta}pH/min was measured after replacement of external NaCl solution by a Cl–-free solution (Na-gluconate) in proximal cell lines from wild-type, task2 –/–, and cftr –/– mice. Values are means ± SE of 6 monolayers/each mouse strain.

 
Further experiments were also performed to identify whether the Cl–/HCO3– exchanger was active in two different cell lines already described as being unable to regulate their volume during hypotonic shock (task2 –/– and cftr –/–) (2, 3). As illustrated in Fig. 4C, the rate of rise in pHi following Cl– removal was identical in cells from wild-type, task2 –/–, and cftr –/– mice. These data clearly indicated that the Cl–/HCO3– exchanger was functional in the different cell lines independently of the presence or absence of CFTR or TASK2 channels.

Measurement of pHi and pHe Variations During RVD

To further prove that the Cl–/HCO3– exchanger was involved in the pHe increment during RVD, we monitored pHi and pHe changes during a hypotonic shock.

pHi. As reported in Table 2, the buffering capacity was not significantly different from one cell line to another. The pHi behavior of cells bathed in HCO3– medium is illustrated in Fig. 5A. In proximal tubule cells from wild-type mice, the hypotonic shock reduced pHi from 7.51 ± 0.01 to 7.01 ± 0.05 (n = 7) within 4–5 min. Afterward, pHi increased to stabilize at 7.48 ± 0.03, a value not significantly different from the value recorded just before the onset of the shock. In cells from task2 –/– mice which did not undergo RVD (2), the hypotonic shock induced a similar decrease in pHi (7.42 ± 0.02 to 6.81 ± 0.03, n = 7), which was also followed by pHi recovery (pHi = 7.44 ± 0.05). In both cell lines, the application of 1 mM DIDS completely prevented the pHi changes during the hypotonic shock.


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  		Table 2. Cell-buffering capacity

 

Figure 5
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  		Fig. 5. Measurement of pHi and pHe variations during RVD in proximal cell lines from wild-type, task2 –/–, and cftr –/– mice. A: for pHi measurement, the cells were loaded for 15 min with BCECF-AM and washed in isotonic NaCl solution. pHi variations were then recorded during hypotonic shock induced by perfusion of a hypotonic NaCl bath solution (230 mosmol/kgH2O). A pH calibration protocol was performed at the end of each experiment by the perfusion of nigericin containing solutions adjusted to various pH values as indicated. Values are means ± SE of n different cell cultures. B: for pHe measurement, the cells were incubated for 8 min in isotonic or hypotonic NaCl solutions. After centrifugation, BCECF (25 µM) was added to the supernatant. The fluorescence of the supernatant was rapidly measured using a spectrofluorimeter. Fluorescence units were converted in pH units using a pH calibration curve. Values are means ± SE of 6 different cell cultures. NPPB, 5-Nitro-2-(3-phenylpropylamino)-benzoic acid; DPC, diphenylamine-2-carboxylate;

 
The pHi variation during the hypotonic shock was also measured in a proximal tubule cell line from cftr –/– mice. In these cells, the hypotonic shock did not induce significant pHi change (Fig. 5A) and did not activate volume-sensitive Cl– channels (3). Therefore, the pHi decrement induced by hypotonicity in wild-type cells was probably related to the presence of functional volume-sensitive Cl– channels.

pHe. Experiments were performed to test whether a hypotonic shock could also increase pHe during the time course of the RVD. pHe was determined in cells obtained by trypsinization of proximal monolayers from wild-type, cftr –/–, and task2 –/– mice and resuspended in a weakly buffered solution (1 mM HEPES) maintained at pH = 7.0. The pHe changes were measured 8 min after the onset of the hypotonic shock. As illustrated in the histogram in Fig. 5B, the hypotonic shock performed on cells from wild-type and task2 –/– mice induced a significant increment of pHe (control: 7.01 ± 0.02; wild-type cells: 7.35 ± 0.03, and task2 –/– cells: 7.32 ± 0.05, for n = 6 for all experimental conditions). This increment of pHe was not detected in the presence of 1 mM DIDS, 100 µM NPPB, or 1 mM DPC. Moreover, the hypotonic shock was unable to promote pHe variations in experiments performed with cftr –/– cells (Fig. 5B).

The inhibitory effects of DIDS suggested that the acidification of pHi and the alkalinization of pHe induced by hypotonicity in wild-type and task2 –/– cell lines could be due to an exit of HCO3–. Moreover, the inhibitory effect of NPPB and DPC in wild-type cells indicated also the involvement of Cl– channels in the pHe alkalinization mediated by the Cl–/HCO3– exchanger. This hypothesis is supported by the inability of the cftr –/– cell line to alkalinize the pHe or to acidify the pHi during the hypotonic shock. These experiments suggested that alkalinization of pHe by the Cl–/HCO3– exchanger was a prerequisite to generate the RVD during a hypotonic shock.

In the following study, whole-cell experiments were performed to assess the effect of external pH variations on the volume-sensitive Cl– and K+ conductances during RVD.

Effect of Buffering Capacity of External Bath Solution on Cl– and K+ Currents Activated During RVD

During these experiments the mean cell membrane capacitances were 20.1 ± 1.4, 18.6 ± 1.3, and 19.2 ± 1.1 (n = 20) for wild-type, task2 –/–, and cftr –/– proximal cells, respectively, and they did not differ significantly between the cell lines (paired t-test).

Cl– currents. To test the effect of buffering capacity on swelling-activated Cl– currents, whole-cell Cl– conductance was recorded during a hypotonic shock. Whole-cell currents were recorded with Ca2+-free (5 mM EGTA) NMDGCl pipette solution (osmolality of 290 mosmol/kgH2O). In Fig. 6A, control currents were measured with an extracellular weakly buffered solution (1 mM HEPES, osmolality of 350 mosmol/kgH2O). The monolayers were then perfused with a 290 mosmol/kgH2O solution. In more than 95% of the cells, whole-cell currents increased within 1 min and reached maximum amplitude after 4–5 min. These large outwardly rectifying currents showed a time-dependent inactivation at depolarizing potentials ≥60 mV. When the cells were exposed to 100 µM NPPB, the currents returned to the control value within 2–3 min. Increasing the buffering capacity of the bath solution (30 mM HEPES) did not modify the development of the swelling-activated Cl– currents (Fig. 6B). Finally, these swelling-activated Cl– currents did not depend on the buffering capacity of the extracellular bath. Proximal cells from task2–/– mice exhibited swelling-activated Cl– currents sharing similar characteristics (data not given).


Figure 6
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  		Fig. 6. Effect of buffering capacity on development of hypotonicity-induced Cl– currents in proximal cell lines from wild-type mice. A: whole-cell currents in PCT cell line from wild-type mice were recorded in weakly buffered bath solutions (1 mM HEPES). Cl– currents were measured in control hypertonic solution (350 mosmol/kgH2O), 4–5 min after replacement of the bath by an isotonic solution (290 mosmol/kgH2O), and finally in the presence of NPPB (100 µM). Membrane voltage was held at –50 mV and stepped to test potential of –100 to +120 mV in 20-mV increments. Shown are whole-cell recordings and corresponding current-voltage (I-V) relationships. Values are means ± SE of 5 cells from 5 different monolayers. B: whole-cell currents in PCT cell line from wild-type mice were recorded in highly buffered bath solutions (30 mM HEPES). Cl– currents were measured in control hypertonic solutions (350 mosmol/kgH2O), 4–5 min after replacement of the bath by an isotonic solution (290 mosmol/kgH2O), and finally in the presence of NPPB (100 µM). Shown are whole-cell recordings and corresponding I-V relationships. Values are means ± SE of 5 cells from 4 different monolayers.

 
K+ currents. It has previously been suggested (2, 5) that TASK2 could be the K+ channel involved in cell volume regulation. Since TASK2 is activated at alkaline pH, we investigated the effect of the buffering capacity of the bath solution on the activation of K+ conductances during a hypotonic shock in proximal cell lines from wild-type and task2 –/– mice. Whole-cell recordings were performed on confluent monolayers bathed in a HCO3–-free, weakly (1 mM HEPES, pH = 7.4) or highly (30 mM HEPES, pH = 7.4 or 8.0) buffered solutions. The currents were recorded with Ca2+-free pipette solutions containing 25 mM HCO3– (osmolality of 290 mosmol/kgH2O). Experiments were performed in the presence of 100 µM NPPB to avoid the development of volume-activated Cl– currents. Figure 7A shows K+ currents recorded in proximal cells from wild-type mice bathed in weakly buffered solution (1 mM HEPES, pH 7.4, osmolality of 290 mosmol/kgH2O). The voltage-step protocol elicited time-independent outwardly rectifying currents, with a reversal potential of –61.2 ± 4.0 mV and a maximal slope conductance of 3.6 ± 0.6 nS (n = 5, Fig. 7A). The monolayers were then perfused with a 230 mosmol/kgH2O solution. This hypotonic shock induced the development of K+ currents within 4–5 min. These large outwardly rectifying currents reversed at –62.3 ± 4.1 mV with a maximum slope conductance of 26.6 ± 1.6 nS (n = 5, Fig. 7A). These currents were strongly inhibited by the application of 10 µM clofilium (Fig. 7A).


Figure 7
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  		Fig. 7. Effect of buffering capacity of external bath solution on K+ currents activated during RVD in proximal cell lines from wild-type and task2 –/– mice. A: whole-cell currents in PCT cells line from wild-type mice were recorded in weakly buffered bath solutions (1 mM HEPES, pHe = 7.4). K+ currents were measured in control isotonic solutions (290 mosmol/kgH2O), 4–5 min after replacement of the bath by a hypotonic solution (230 mosmol/kgH2O), and finally in the presence of clofilium (10 µM). The solutions contained NPPB (100 µM). Membrane voltage was held at –50 mV and stepped to test potential of –100 to +120 mV in 20-mV increments. Shown are whole-cell recordings and corresponding I-V relationships. Values are means ± SE of 5 cells from 5 different monolayers. B: whole-cell currents in PCT cells line from wild-type mice were recorded in highly buffered bath solutions (30 mM HEPES). K+ currents were measured in control isotonic solution (290 mosmol/kgH2O, pHe = 7.4), 4–5 min after replacement of the bath by a hypotonic solution (230 mosmol/kgH2O, pHe = 7.4), 4–5 min after replacement of the bath by a hypotonic solution (230 mosmol/kgH2O, pHe = 8.0), and finally in the presence of clofilium (10 µM). The solutions contained NPPB (100 µM). Shown are whole-cell recordings and corresponding I-V relationships. Values are means ± SE of 5 cells from 4 monolayers. C: whole-cell currents in PCT cell line from task2 –/– mice were recorded in weakly buffered bath solutions (1 mM HEPES, pHe = 7.4). K+ currents were measured in control isotonic solution (290 mosmol/kgH2O) and 4–5 min after replacement of the bath by a hypotonic solution (230 mosmol/kgH2O).The solutions contained NPPB (100 µM). Shown are whole-cell recordings and corresponding I-V relationships. Values are means ± SE of 5 cells from 6 different monolayers.

 
When similar hypotonic shock experiments were performed in a highly buffered bath solution (30 mM HEPES, pH 7.4, 230 mosmol/kgH2O), no significant increase in the outward K+ currents was observed (Fig. 7B). To test whether a high HEPES concentration modified the swelling-activated K+ currents, the monolayers were perfused with a highly buffered hypotonic bath solution (30 mM HEPES, pH 8.0). Under this condition (Fig. 7B), large outwardly rectifying K+ currents were recorded with a reversal potential of –57.3 ± 3.4 mV and a slope conductance of 30.2 ± 3.1 pS (n = 5). These data indicated that high HEPES concentrations did not inhibit TASK2 K+ currents.

As expected for a K+ current flowing through TASK2 channels, no swelling-activated K+ currents were observed in task2 –/– cells (Fig. 7C).

Finally, the data strongly suggest that cell swelling induced an increase in pHe, which activated TASK2 K+ currents, leading finally to the RVD process. The increase in pHe is probably due to the extrusion of HCO3– through the Cl–/HCO3– exchanger.

Involvement of Cl–/HCO3– Exchanger in Activation of K+ Currents During RVD

To test whether the Cl–/HCO3– exchanger was involved in the activation of K+ conductance, whole-cell currents were recorded during a hypotonic shock in a weakly buffered (1 mM HEPES) bath solution. The experimental solutions were chosen to promote a Cl–/HCO3– exchange by increasing the outward gradient of HCO3– and the inward gradient of Cl–. Under these control conditions, the hypotonic shock activated TASK2 K+ currents (see Fig. 7A). In contrast, preincubation of proximal cells with 1 mM DIDS for 60 min followed by intensive washout impaired the development of outwardly rectifying K+ conductance during the hypotonic shock (Fig. 8A). Thus the irreversible inhibition of the Cl–/HCO3– exchanger due to covalent DIDS binding strongly decreased TASK2 K+ conductance. To discard the possibility that this inhibition of TASK2 K+ conductance could be indirectly due to a blockade of Cl– conductance by DIDS, further experiments were performed with another Cl– channel blocker. The presence of 100 µM NPPB during the hypotonic shock did not modify the development of K+ conductance (Fig. 8B).


Figure 8
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  		Fig. 8. Involvement of Cl–/HCO3– exchanger activity in activation of K+ currents during RVD in proximal cell lines from wild-type mice. A: whole-cell currents in DIDS (1 mM)-preincubated PCT cells from wild-type mice were recorded in control condition (isotonic solution, 290 mosmol/kgH2O) and after 4–5 min of extracellular perfusion of a hypotonic solution (230 mosmol/kgH2O). Before the patch-clamp experiments were performed, the preincubated cells were thoroughly washed to eliminate unbound DIDS. Membrane voltage was held at –50 mV and stepped to test potential of –100 to +120 mV in 20-mV increments. Shown are whole-cell recordings and corresponding I-V relationships. Values are means ± SE of 6 cells from 4 different monolayers. B: whole-cell currents in a PCT cell line from wild-type mice were recorded in the continuous presence of NPPB (100 µM) in control condition (isotonic solution, 290 mosmol/kgH2O) or after 4–5 min of extracellular perfusion of a hypotonic solution (230 mosmol/kgH2O). At the end of the experiment, a hypotonic solution containing DIDS (1 mM) was perfused. Shown are whole-cell recordings and corresponding I-V relationships. Values are means ± SE of 6 cells from 4 different monolayers.

 
Role of pHe in Activation of K+ Currents by Hypotonicity

To further demonstrate the effect of pHe on the activation of K+ currents by a hypotonic shock, whole-cell currents were recorded in weakly buffered bath solutions (1 mM HEPES) containing 1 mM DIDS. The results are reported in Fig. 9. In an isotonic bath solution (290 mosmol/kgH2O), increasing pHe from 7.4 to 8.0 enhanced outward K+ conductance (maximal slope conductance = 3.9 ± 0.7 nS at pHe = 7.4 and 18.1 ± 1.2 nS at pHe = 8.0, n = 5, Fig. 9). Interestingly, the perfusion of a hypotonic solution (230 mosmol/kgH2O) at pHe = 8.0 induced a further increase in the outward K+ current with a maximal slope conductance of 29.9 ± 2.8 nS (n = 5). This current was markedly blocked by clofilium (10 µM).


Figure 9
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  		Fig. 9. Role of pHe in the activation of K+ currents by hypotonicity. Whole-cell currents in PCT cell line from wild-type mice were recorded in the continuous presence of DIDS (1 mM) in an isotonic solution (control pH 7.4, 290 mosmol/kgH2O), 4–5 min after extracellular perfusion of an isotonic solution at pH 8.0, 4–5 min after perfusion of a hypotonic solution at pH 8.0 (230 mosmol/kgH2O), and finally in the presence of clofilium (10 µM). Shown are whole-cell recordings and corresponding I-V relationships. Values are means ± SE of 5 cells from 5 different monolayers.

 
Therefore, these results strongly suggested that alkalinization of the extracellular medium is a prerequisite to trigger the activation of TASK2 during a hypotonic shock.


   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
In the different segments of the nephron, cells are subjected to osmotic shocks, either by accumulation of active osmolytes in their cytoplasm (proximal tubule) or by dilution of the tubular fluid (distal tubule). In response to such osmotic stress, these cells undergo a RVD process by activating swelling-sensitive Cl– and K+ conductances (5). Of the K+ channels involved in the osmotic response, TASK2 K+ channels play an essential role in the proximal tubule (2). However, the mechanisms responsible for TASK2 activation during a hypotonic shock are not yet established. Interestingly, the TASK2 K+ current shows a strong dependence on pHe, being activated at alkaline pH within physiological ranges of variation (21). This property leads us, first, to investigate whether the activation of TASK2 by a hypotonic shock could result from an increase in pHe. For this purpose, we have developed immortalized cell lines from primary cultures of proximal tubules from wild-type and task2 –/– mice. All the cell lines formed monolayers, and the wild-type cells were capable of RVD mediated by Cl– and K+ conductances. These conductances shared similar biophysical and pharmacological features with those described in primary cultures of mouse proximal tubules (2, 5). Several findings in the present study indicate that a pHe increment could mediate the RVD. For example, we demonstrated that the RVD phenomenon was strongly dependent on pHe, with an inhibition at acidic pH and activation at alkaline pH. Interestingly, an effect of pH on RVD has already been reported in Ehrlich cells submitted to a hypotonic shock (18). If we link this finding to the observation that a high buffering capacity also blocks RVD, then we could assume that the hypotonic shock triggered an increase in pHe. Direct measurement of pHe during RVD confirmed this assumption since the hypotonic shock induced an increase in pHe by >0.3 pH units.

In the present study, it was demonstrated that swelling-activated Cl– channels were insensitive to pHe. Therefore, the pHe sensitivity of the RVD phenomenon is probably conferred by swelling-activated TASK2 K+ conductance. The blockade of TASK2 currents by highly buffered concentrations during hypotonicity corroborated the essential role of this channel.

In isotonic conditions, the activation of TASK2 channels was demonstrated to be mediated by the rise in pHe induced by HCO3– efflux (42). It is probably also the case in hypotonic conditions since the TASK2 current was not observed in the presence of DIDS. However, the increase pHe alone was not sufficient to account for the increase in K+ conductance during the hypotonic shock. Reciprocally, the hypotonic shock was inefficient to enhance TASK2-mediated K+ currents when the increase in pHe was prevented. Thus alkalinization of extracellular medium is a prerequisite to trigger the activation of TASK2 during a hypotonic shock. Moreover, the observation that RVD occurred concomitantly with an intracellular DIDS-sensitive acidification suggested that the hypotonic shock induced an exit of HCO3– (14, 28). The role of HCO3– in RVD (8) and the involvement of the Cl–/HCO3– exchanger in HCO3– transport during RVD (9, 16, 28, 39) have already been suggested in previous experiments. This is shown to be the case in the present study because RVD was insensitive to Na+ suppression and was inhibited by external Cl– removal. Indeed DIDS completely prevented the RVD, but its action could be due to a simultaneous effect on both the Cl–/HCO3– exchanger and swelling-activated Cl– conductance. The observation that RVD was inhibited in cells preincubated with DIDS and not with DNDS corroborates the central role of Cl–/HCO3–. In fact, it has been demonstrated that DIDS but not its analog DNDS covalently binds the AE1 exchanger when incubated for 60 min or more (8, 34, 38). Therefore, in the present study the Cl–/HCO3– exchanger remained blocked once DIDS was removed and the cells were unable to undergo RVD, although Cl– conductance was still functional.

In proximal tubule cell lines from wild-type, task2 –/–, and cftr –/– mice, the strong increase in pHi induced by external Cl– removal in a HCO3– medium could well be mediated by the activation of the Cl–/HCO3– exchanger. However, it is interesting to note that in cftr –/– cells the hypotonic shock was insufficient to decrease pHi while the Cl–/HCO3– exchanger remained functional. Moreover, these cells were unable to undergo RVD upon the hypotonic shock due to a loss of the signaling cascade that controls swelling-activated Cl– channels (3). Therefore, it is clear that an increase in Cl– conductance induced by a hypotonic shock is required to activate the Cl–/HCO3– exchanger. This observation was confirmed in wild-type and task2 –/– cells in which the application of the Cl– channel blocker NPPB prevented the pHi decrease upon hypotonicity.

In the original study of Dellasega and Grantham (12), in vitro experiments clearly indicated that a hypotonic bath influenced cell volume in nonperfused proximal tubules. In response to this shock, the proximal tubule underwent RVD. It is clear that the model and the experimental conditions were quite different in this study. A 50% hypotonic shock was applied in the basolateral compartment, and the extracellular volume was one million times larger than the volume of the tubule (12). Under these conditions, it was difficult to measure a pHe change. However, on the basis of our experiments it is possible that the pHe variation was restricted to the immediate vicinity of the cells. Such a local phenomenon could be sufficient to increase the local pHe and activate TASK2. This latter hypothesis could partially explain the activation of TASK2 K+ conductance recorded in whole-cell clamp experiments during a hypotonic shock. However, in vitro studies performed in intact proximal tubules (43) showed that intracellular HCO3– depletion did not alter RVD. Such an observation is at variance with our present data since we attributed a crucial role to HCO3– in the alkalinization of the extracellular compartment. This discrepancy could probably be explained by the huge difference between the HCO3– concentrations released by the cells compared with the external bath buffering capacity. Interestingly, the HCO3– requirement in the RVD process has already been described in cells of proximal tubules of Necturus (24) and mouse (41), cells of the thin descending limb of Henle’s loop in the rabbit (23), and in other cell types such as single osteosarcoma UMR-106–01 cells (36) and the epithelial-derived human breast cancer cell line ZR-75–1 (28).

The relationship between CFTR and HCO3– secretion has been well documented in secretory epithelia. Different mechanisms have been identified such as a Cl–-dependent secretion associated with apical membrane Cl–/HCO3– exchange, or a cAMP-induced secretion through the Cl–-permeable pore of CFTR (40). In primary cultures of mouse proximal tubules, it has been demonstrated that the application of forskolin did not stimulate any Cl– currents (3). This feature has also been found in the mouse proximal cell line used in the present study (data not shown). The absence of cAMP-stimulated Cl– channels suggests that CFTR does not conduct HCO3– directly.

The data from the present study are summarized in Fig. 10. In proximal tubule cells, a hypotonic shock induces the activation of swelling-activated Cl– channels in the presence of CFTR. The molecular nature of these channels is not yet known, but CFTR controls them by modulating autocrine adenosine production (5). The resulting exit of Cl– would decrease cytosolic Cl– activity, providing the driving force for the activation of the Cl–/HCO3–exchanger. The efflux of HCO3– then alkalinizes pHe, which activates the TASK2 channels, allowing for a further increment of the K+ conductance induced by the hypotonicity. This sensitivity to hypotonic shock has already been described in several studies which demonstrated that TASK2 could be a swelling-activated K+ channel (2, 17, 29).


Figure 10
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  		Fig. 10. Working model of TASK2 and CFTR during RVD in proximal cell lines. Putative mechanisms for cell volume regulation during a hypotonic shock are shown.

 
Finally, the RVD process is achieved by an exit of KCl controlled indirectly by CFTR (3). It is clear that further experiments will be necessary to confirm this hypothesis. Notably, the membrane localization of the different transporters and channels involved in the RVD is not fully established. Even if TASK2 channels are located mainly in the proximal tubule (2), their basolateral localization has been determined in functional studies only (2, 42). Numerous lines of evidence suggest that the Cl–/HCO3– exchanger could contribute to basal efflux-mediating HCO3– reabsorption across the basolateral membrane (1). Concerning the swelling-activated Cl– channel, the fact that it could be modulated by CFTR-dependent autocrine control suggested a common localization with CFTR. At present, CFTR has been detected in the apical membrane by immunofluorescence study but patch-clamp recordings localize its activity to the basolateral membrane of the proximal tubule (10).

In conclusion, the present study attributes a central role to HCO3– in the control of RVD in proximal tubule cells. The originality of the model proposed here is based on the assumption that TASK2-mediated K+ permeability is stimulated by hypotonic shock and the concomitant increase in pHe is induced by HCO3– efflux. This mechanism is physiologically relevant in the proximal tubule because the active apical absorption of ions and organic solutes increases osmolytes inside the cytoplasm. The proximal tubule exhibits high water permeability of both cell membranes (apical and basolateral) due to the presence of aquaporin-1 (27). Therefore, the solute entries are accompanied by water fluxes in the same direction and RVD must occur to continuously maintain cell volume. The RVD in proximal tubules cells is the result of a KCl efflux via Cl– and K+ channels. However, in the proximal tubule, the reabsorption of HCO3– in excess of water alkalinizes the blood in the peritubular capillaries, allowing the activation of TASK2 channels. By using an in vitro system, we differentiated 1) the exit of HCO3– induced by cell swelling from 2) the exit of HCO3– resulting from its transcellular transport. The first is driven by a Cl–/HCO3– exchanger, whereas the second is driven by a Na+-3HCO3– cotransporter (42). In the proximal tubule in vivo, both systems work together to ensure HCO3– reabsorption and cell volume regulation. TASK2 K+ channels are continuously activated by cell swelling and basolateral HCO3– accumulation, allowing the cell to maintain its volume and its membrane potential, which is depolarized by Cl– exit (through swelling-activated Cl– channels) and electrogenic exit of Na+ and HCO3– (through the Na+-3HCO3– cotransporter).


   ACKNOWLEDGMENTS
 
We thank Dr. K. Mitchell and Prof. Dr. W. Skarnes for generously providing task2 –/– mice.


   FOOTNOTES
 

Address for reprint requests and other correspondence: P. Poujeol, UMR CNRS 6548, Université de Nice-Sophia Antipolis, 06108 Nice Cedex 2, France (e-mail: philippe.poujeol{at}unice.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.


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00132.2006v1
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