Ionic currents induced by cell swelling were characterized in primary cultures of rabbit distal bright convoluted tubule (DCTb) by the whole cell patch-clamp technique. Cl− currents were produced spontaneously by whole cell recording with an isotonic pipette solution or by exposure to a hypotonic stress. Initial Cl− currents exhibited outwardly rectifying current-voltage relationship, whereas steady-state currents showed strong decay with depolarizing pulses. The ion selectivity sequence was I−= Br− > Cl− ≫ glutamate. Currents were inhibited by 0.1 mM 5-nitro-2-(3-phenylpropylamino)benzoic acid and 1 mM 4,4′-diisothiocyanostilbene-2,2′-disulfonic acid and strongly blocked by 1 mM diphenylamine-2-carboxylate. Currents were insensitive to intracellular Ca2+but required the presence of extracellular Ca2+. They were not activated in cells pretreated with 200 nM staurosporine, 50 μM LaCl3, 10 μM nifedipine, 100 μM verapamil, 5 μM tamoxifen, and 50 μM dideoxyforskolin. Staurosporine, tamoxifen, verapamil, or the absence of external Ca2+ was without effect on the fully developed Cl−currents. Osmotic shock also activated K+ currents in Cl−-free conditions. These currents were time independent, activated at depolarized potentials, and inhibited by 5 mM BaCl2. The activation of Cl− and K+ currents by an osmotic shock may be implicated in regulatory volume decrease in DCTb cells.
- whole cell
- cell volume
- ion conductance
in many epithelial cells, Cl− channels are essential for the transport of salt and water across the membrane bilayer, for stabilization of the resting membrane potential, and for regulation of cell volume. At least three distinct Cl− currents, regulated by adenosine 3′,5′-cyclic monophosphate (cAMP), cytosolic Ca2+, and osmotic pressure, have been found in several tissues, including airway epithelial cells, sweat gland, pancreas, and T84 intestinal cells (14). In the kidney, Cl− channels activated by cell swelling have been observed in proximal tubule, in the loop of Henle, and in collecting tubules (39). As in other tissues, these channels are one of the membrane transport pathways that mediate an efflux of osmolytes to bring about regulatory volume decrease (RVD). The activation of Cl−condutance during RVD is regularly associated with an increase in K+ conductance, leading to a net loss of KCl and a concomitant efflux of water. In the proximal tubule, cell swelling is mainly the result of the transport of osmotically active solutes into the cells via cotransporters coupled to the Na+ gradient. In these cells, mechanisms of RVD are well developed and have been extensively studied (19, 22, 23, 25, 41, 44). Presently, very few studies have examined swelling-activated conductances in the distal convoluted tubule, although, with the low osmolarity of the fluid delivered to this segment, the distal cells would be expected to require volume regulatory capabilities. In view of these indications, we carried out whole cell experiments to investigate further the hypotonically activated Cl− and K+ conductances in distal convoluted tubule. For this purpose, we used primary cultures of the bright part of rabbit distal convoluted tubule (DCTb). These cultures are now well characterized (26). They exhibit apical Cl− channels activated by cAMP and basolateral Ca2+-dependent Cl− conductance (3, 35, 43). This study describes hypotonically activated Cl− currents with pharmacological properties similar to those of swelling-activated Cl− current in epithelial cells undergoing RVD on exposure to hyposmotic medium. The activation of this Cl− conductance requires extracellular Ca2+ and is associated with the phosphorylation of P-glycoprotein. The hypotonic shock also activates K+conductance sensitive to Ba2+. It is therefore possible that the observed K+ and Cl− currents may be responsible for RVD.
MATERIAL AND METHODS
The primary cell culture technique used in this study has been described in detail in previous papers (26, 35). Briefly, the bright parts of the rabbit distal tubules were microdissected under sterile conditions from kidneys obtained from 4- to 5-wk-old male New Zealand rabbits. Kidneys were perfused with Hanks’ solution (GIBCO) containing 600–700 kU/l collagenase (Worthington) and cut into small pyramids, which were incubated in medium containing 150 kU/l collagenase. The tubules were seeded in collagen-coated 35-mm petri dishes filled with a culture medium composed of equal quantities of Dulbecco’s modified Eagle’s medium and Ham’s F-12 (GIBCO), containing 15 mM NaHCO3, 20 mMN-2-hydroxyethylpiperazine-N′-2-ethanesulfonic acid (HEPES), pH 7.5, 2 mM glutamine, 5 mg/l insulin, 50 nM dexamethasone, 10 μg/l epidermal growth factor, 5 mg/l transferrin, 30 nM sodium selenite, and 10 nM triiodothyronine. Cultures were maintained at 37°C in a 5% CO2-95% air water-saturated atmosphere. The medium was changed 4 days after seeding and then every 2 days.
Whole Cell Experiments
Whole cell currents were recorded from 12- to 22-day-old cultured cells grown on collagen-coated supports maintained at 33°C throughout the experiments. The ruptured-patch whole cell configuration of the patch-clamp technique was used. Patch pipettes were made from borosilicate capillary tubes (1.5 mm OD, 1.1 mm ID; Clay Adams) using a two-stage vertical puller (PP 83; Narishige, Tokyo, Japan). When filled with N-methyl-d-glucamine chloride (NMDG-Cl) solution the pipettes had a resistance ranging from 2 to 3 MΩ in a NMDG-Cl buffer. An Ag-AgCl pellet was used as the reference electrode. To reduce junction potentials, this electrode was bathed in an identical solution to that contained in the pipette and connected to the bath via a 3 M KCl-agar bridge. Cells were observed by using an inverted microscope (Zeiss IM 35), the stage of which was equipped with a water robot micromanipulator (MHW30, Narishige). The patch pipette was connected via an Ag-AgCl wire to the head stage of a RK 400 patch amplifier (Biologic). After the formation of a gigaohm seal, the fast compensation system of the amplifier was used to compensate for the head-stage intrinsic input capacitance (≈8 pF). The membrane was ruptured by additional suction to achieve the conventional whole cell configuration. At this stage, the cell capacitance (C m) was compensated for with a facility provided on the RK 400 amplifier. With NMDG-Cl in the bath and in the pipette, theC m of 146 cells was 27.2 ± 0.8 pF (mean ± SE), and the series resistance (R s) averaged 9.8 ± 0.8 MΩ. With potassium glutamate in the bath and in the pipette, theR s averaged 6.5 ± 0.3 MΩ (n = 32).R s was compensated for in K+ currents but not in Cl−currents. For this purpose, theR s compensation circuitry of the RK 400 was used. This facility allowed a 90% compensation, reducing the capacitive time constant in the same proportion. However, in all cases, experiments whereR s > 20 MΩ were discarded. Cell membrane potentials were measured at zero membrane current in the current-clamp mode of the amplifier. Extracellular test solutions were perfused into the bath using a four-channel glass pipette, the tip of which was placed as near as possible to the clamped cell.
Voltage-clamp commands, data acquisition, and data analysis were controlled by an IBM-AT-compatible computer equipped with a Digi Data 1200 interface (Axon Instruments, Foster City, CA). Commercially available pCLAMP software (version 6.0, Axon Instruments) was used to generate whole cell current-voltage (I-V) relationships. Membrane currents resulting from voltage stimuli were filtered at 1 kHz, sampled at a rate of 2,560/s, and stored directly onto the hard disk. For the measurement of chloride currents, cells were held at a holding potential (V hold) of −50 mV, and 400-ms pulses from −100 to +120 mV were applied with increments of 20 mV every 2 s. K+ currents were measured by applying pulses ranging from −100 to +80 mV from aV hold of −50 mV.
Image analysis. The optical system was composed of a Zeiss ICM-405 inverted microscope and a Zeiss ×40 objective, which was used for epifluorescent measurements with a 75-W xenon lamp. The excitation beam was filtered through narrow-band filters (340, 360, and 380 nm; Oriel) mounted in a motorized wheel (Lambda 10–2; Sutter Instrument) equipped with a shutter to control the exposure times. The incident and the emitted fluorescence radiation were separated through a Zeiss chromatic beam splitter. Fluorescence emission was selected through a 510-nm narrow-band filter (Oriel). The transmitted light images were viewed by an intensified camera (Extended ISIS; Photonic Science, Sussex, UK). The 8-bit extended-ISIS camera was equipped with an integration module to maximize signal-to-noise ratio. The video signal from the the camera proceeded to an image processor integrated in a DT2867 image card (Data Translation) installed in a Pentium 100 PC. The processor converts the video signal into 512 lines by 768 square pixels per line by 8 bits per pixel. The 8-bit information for each pixel represents one of the 256 possible gray levels, ranging from 0 (for black) to 255 (for white). Image acquisition and analysis were performed by the 2.0 version of AIW software (Axon Instruments). The final calculations were made using the Excel software (Microsoft).
Intracellular Ca2+measurements. Fifteen- to twenty-day-old confluent DCTb monolayers grown on petri dish were loaded with a solution of 2 μM fura 2 containing 0.01% pluronic acid for 30 min at 37°C and were then washed with a NaCl solution. The cells were successively excited at 340 and 380 nm; the images were digitized and stored on the hard disk of the computer. Each raw image was the result of an integration of six frames averaged four times. The acquisition rate was 1 image/4 s. The intracellular Ca2+concentration was calculated from the dual-wavelength fluorescence ratio by using the Grynkiewicz equation (16).
Mn2+influx measurements. Fifteen- to twenty-day-old confluent DCTb monolayers were loaded for 45 min with a solution of 5 μM of the acetoxymethyl ester of fura 2 (fura 2-AM) containing 0.01% pluronic acid, and the cells were then washed with NaCl solution. Fluorescence quenching experiments were carried out by addition of 50 μM of MnCl2 in the NaCl medium. Fluorescent measurements were performed at 360 nm (isobestic point of fura 2) where the fluorescence signal is independent of free calcium concentration. The manganese influx was quantified by the slope of the quenching kinetics. Each raw image was the result of an integration of six frames averaged four times. The acquisition rate was 1 image/4 s.
Cell volume estimation. The relative cell volume was monitored by measuring trapped dye concentrations. This technique was derived from that previously reported (44), but fura 2 was used instead of 2′,7′-bis(carboxyethyl)-5(6)-carboxyfluorescein. The cells were loaded with fura 2, as described above, and the fluorescence was monitored with 360-nm excitation wavelength. At 360 nm, the variations of the signal emitted by the probe are directly proportional to the variations of the cell volume. In a typical experiment, the cells were first perfused with an isotonic NaCl solution containing (in mM) 110 NaCl, 5 KCl, 1 CaCl2, 90 mannitol, and 10N-2-hydroxyethylpiperazine-N′-2-ethanesulfonic acid [pH 7.4, osmotic pressure (Posm) = 320 mosmol/kgH2O at 30 ml/min], and images were averaged 30 times and recorded every 5 s for 4 min. Once the fluorescence stabilized, a hypotonic shock was induced by perfusing the NaCl solution without mannitol (Posm = 227 mosmol/kgH2O). The estimation of relative change in cell volume from the fluorescent signal was made, assuming that a 30% decrease in the osmolarity caused a decrease of the fluorescent signal corresponding to a maximum swelling of 30% compared with the initial volume. The means of relative volume changes were obtained by the analysis of 5–10 zones in each ofn number of cultures chosen with the software. Each zone delimited a cytoplasmic area chosen in individual cells.
A stock solution of 5-nitro-2-(3-phenylpropylamino)benzoic acid (NPPB) from Calbiochem was prepared at 100 mM in dimethyl sulfoxide (DMSO) and used at 0.1 mM in final solutions. Diphenylamine-2-carboxylate (DPC) from Aldrich was prepared as 1 mol/l stock solution in DMSO and dissolved at 1 mM in incubation medium. 4,4′-Diisothiocyanostilbene-2,2′-disulfonic acid (DIDS) from Sigma was directly dissolved at a final concentration of 1 mM. Fura 2-AM from Molecular Probes was dissolved at 3 mM in DMSO. Staurosporine and protein kinase inhibitor were from Calbiochem. Nifedipine, verapamil, dideoxyforskolin, and tamoxifen were obtained from Sigma.
The compositions of the different solutions used in these experiments are given in Table 1.
Cl− Currents in Unstimulated Cultured DCTb Cells
Whole cells currents were recorded with Ca2+-free pipette solutions containing NMDG-Cl (Table 1, solution f ) and in an extracellular solution containing NMDG-Cl (Table 1,solution c). Both solutions were isotonic (290 mosmol/kgH2O). After successful gigaohm seal formation, the whole cell configuration was obtained in 25% of the cases. Within 2–4 min after the mechanical rupture of the membrane, the cell depolarized as the pipette solution equilibrated with the cell interior. Voltage-clamp experiments were performed, holding the cell at −50 mV and applying voltage steps of 400-ms duration every 2 s from −100 to +120 mV in 20-mV increments. Once cell depolarization was complete, currents elicited by the voltage protocol were recorded every minute. Figure1,A–D, shows families of currents recorded immediately, 2, 4, or 6 min after the beginning of whole cell recording (recording time). Only 105 of 265 cells displayed these currents in isotonic conditions. In these cells, we have considered that the control currents (t = 0 min) were those recorded when membrane potential just reached 0 mV. In these conditions, the initial currents measured 6 ms after the onset of the voltage pulse, rectified slightly in the outward direction (Fig. 1 E). They reversed at −2.3 ± 1.4 mV (n = 20), and the total current at 100 mV was 1.9 times the current at −100 mV (100 mV = 257.4 ± 24.4 pA, −100 mV = −137.1 ± 14.9 pA;n = 20,P < 0.01). The initial currents then increased with time and stabilized after 6 min (Fig. 1,B–D).TheI-V relationships for initial and steady-state currents are illustrated in Fig. 1,E andF, respectively. Initial currents displayed an outward rectification that increased with time (Fig.1 E). The maximal current was reached 6 min after the beginning of the whole cell recording. At this time, the initial current recorded at 100 mV was 2.4 times the current at −100 mV (100 mV = 1,117 ± 77 pA, −100 mV = −474.1 ± 73.0 pA; n = 20,P < 0.001). These large, outwardly rectifying currents showed time-dependent inactivation at depolarizing step potentials >40 mV. Generally, the time course of this inactivation could be well fitted with a single exponential regardless of the recording time. In 15 of the 20 cells studied, the greater the voltage, the faster was the rate of the decay. At 120 mV, the currents inactivated with a time constant of 132.4 ± 6.50 ms (n = 15). To better illustrate the decay of the currents, the percentage of inactivation was calculated as the difference between the initial current at 6 ms and the steady current at 390 ms into the same voltage step divided by the current at 390 ms. At every imposed potential, the inactivation was independent of the recording time. Figure 2 shows that the inactivation during the 400-ms pulse >40 mV strongly increased with more depolarizing potentials. These data were obtained from experiments performed in symmetrical Cl−concentrations. The reversal potential (E rev) was very close to that of Cl−, and, in the absence of permeable cations in the pipette, the outward current was carried by Cl−.
To study the anion permeability of the cell membrane, all except 2 mM of the Cl− in the bath solution was replaced with I−, Br−, or glutamate. Figure3,A–C, gives typical recordings of the currents obtained at 6 min in the presence of the three different anions. Figure 3, D andE, showsI-V relations for initial and steady-state currents. Table 2summarizes E revvalues as well as the calculated permeability ratios obtained for a given anion. To minimize the effects of capacitance transients, the whole cell currents were fitted to an exponential curve over the interval of 20–390 ms. The amplitude of the instantaneous current was extrapolated to the time of the onset of the voltage step. In the presence of I− or Br−,E rev shifted toward the negative values and was independent from the duration of the stimulation. Replacing external chloride with glutamate markedly reduced the initial and steady-state outward currents. During this substitution, the inward currents carried by chloride remained unaffected, andE rev moved toward positive voltages. However,E rev values calculated for the initial I-V curves were significantly lower than those calculated for steady-state currents. The curve in Fig. 4 illustrates the variations ofE rev for glutamate as a function of the voltage pulse duration. Finally, the sequence for this conductance was I− = Br− > Cl− ≫ glutamate.
To further characterize the Cl− current, we tested three anion channel blockers added separately to the bathing solution. Figure5,A–D, gives typical traces of the current obtained 6 min after starting the whole cell recording. The addition of NPPB, DIDS, or DPC to the bathing solution inhibited the whole cell Cl− currents within 2 min. The effects of these blockers were reversible on washing (data not given). The I-Vrelationships for initial and steady-state currents are given in Fig.5, E andF, respectively. Overall, 0.1 mM NPPB or 1 mM DIDS inhibited reversibly both initial inward (%inhibition at −100 mV: NPPB, 44.3 ± 3.8, n= 8; DIDS, 34.6 ± 5.0, n = 18) and outward currents (%inhibition at +100 mV: NPPB, 71.4 ± 4.6,n = 8; DIDS, 77.2 ± 6.8,n = 18). The effects of NPPB and DIDS were therefore voltage dependent. By contrast, the blocking effects of DPC on initial currents were similar at −100 and +100 mV (%inhibition at −100 mV, 71.3 ± 2.8; %inhibition at +100 mV, 73.5 ± 4.6, n = 5).
Cl− Currents Induced by a Hypotonic Shock
Currents recorded in these DCTb cells were produced spontaneously when the monolayers were bathed with a solution having the same osmotic pressure as the pipette solution. The overall characteristics of this Cl− conductance are similar to the swelling-induced currents described in several tissues (8, 9,21, 40, 49, 52). To study the effects of changes in osmotic pressure on the development of this conductance, currents were then induced by osmotic shock. In these experiments, the pipette solution was maintained at 290 mosmol/kgH2O. Moreover, to eliminate any participation of cations in the inward current, experiments were carried out after replacing Na+ in the bath solution by NMDG+. Figure6 Aillustrates the time course of the initial currents measured at +100 mV as a function of the osmolarity of the bath solution. At an extracellular solution osmolarity of 350 mosmol/kgH2O (Table 1,solution c + 60 mM mannitol), the voltage-step protocol elicited small, time-independent currents that changed linearly with the membrane voltage, with a slope conductance of 0.97 ± 0.04 nS andE rev of −0.69 ± 0.03 mV (n = 21). Because of their small amplitude, the nature of these currents was not analyzed further. The monolayer was then perfused with a 290 mosmol/kgH2O solution. In >95% of the cells, an increase in the whole cell current was observed within 1 min. The currents reached a maximal after 5–6 min and remained stable for 5 min. When the cells were reexposed to hyperosmotic solution, the currents returned to the control level within 2–3 min. The currents induced by hypotonicity were analyzed at their maximal values. It was found that they were very similar to those developed spontaneously in isotonic solutions. This is illustrated by the I-V relationships for initial currents reported in Fig. 6 B. The substitution of external Cl−by I− increased the outward currents and movedE rev toward more negative values, giving aE rev of −21.6 ± 0.94 mV and a calculated relative I−-to-Cl−permeability ratio of 2.40 ± 0.11 (n = 5). Furthermore, glutamate causedE rev to shift by 38.5 ± 1.2 mV such that the glutamate-to-chloride permeability ratio was 0.20 ± 0.01 (n = 5). Figure 6 B also shows the effect of DIDS and NPPB on swelling-activated chloride currents. The blocking effect of both drugs was voltage dependent. However, the potency of NPPB to decrease the inward current was found to be greater than that of DIDS (%inhibition at −100 mV: NPPB, 68.0 ± 7.1; DIDS, 17.9 ± 3.6; %inhibition at +100 mV: NPPB, 80.1 ± 8.2, n = 5; DIDS, 78.1 ± 6.4,n = 5).
Regulation of the Cl− Conductance Induced by Hypotonic Shock
Calcium. To eliminate the implication of cytosolic Ca2+ in the development of hypotonicity-induced Cl− currents, experiments were generally performed using pipette solutions containing 5 mM ethylene glycol-bis(β-aminoethyl ether)-N,N,N′,N′-tetraacetic acid (EGTA) without additional Ca2+. However, some experiments were carried out by increasing EGTA concentration up to 10 mM. In 100% of the cells tested, this maneuver did not modify the Cl− currents on exposure to a hypotonic medium (data not given). In further experiments, the putative variations of cytosolic Ca2+ induced by hypotonic shock were followed by using fura 2 as a Ca2+ probe. The data clearly indicated that no significant variation of intracellular Ca2+ could be detected even with a hypotonic shock of 100 mosmol/kgH2O (isotonicity: intracellular Ca2+ concentration, 83.7 ± 6.0 nM; hypotonicity: intracellular Ca2+ concentration, 79.3 ± 6.3 nM; n = 13).
The effects of extracellular Ca2+on the development of hypotonicity-induced Cl− currents were also tested. The histogram of Fig. 7 shows that when the hypotonic shock was carried out in the absence of bath Ca2+, the development of the Cl− current was significantly impaired. In fact, in the absence of extracellular Ca2+, 7 of the 13 recorded cells did not exhibit Cl− currents in response to a hypotonic shock, whereas the remaining six cells elicited a response, which was reduced by 46.2 ± 8.1% compared with the control value. Conversely, removal of bath Ca2+ after the current had been activated did not modify significantly the amplitude of the Cl− currents in any of the cells tested (Fig. 8). These experiments indicate that extracellular Ca2+was required to activate the swelling-activated Cl− conductance. To determine whether this activation was related to Ca2+ influx across the cell membrane, we studied the role of ionomycin in the development of the Cl− conductance. To provide evidence that ionomycin induced an increase of divalent cation entry, we took advantage of the property of fura 2 fluorescence being quenched by Mn2+, a commonly used substitute for Ca2+. The fluorescence at 360 nm (isobestic point for fura 2) is independent of cytosolic Ca2+. Thus only fura 2 quenching due to Mn2+ uptake by the cells modified the fluorescence signal. When NaCl solution containing MnCl2 was perfused, an immediate fluorescence decrease (0.30 ± 0.01 fluorescence arbitrary units/s, n = 3) was observed. As shown in Fig. 9, extracellular application of ionomycin accelerated the rate of quenching (0.73 ± 0.03 fluorescence arbitrary units/s, n = 3), indicating an enhanced influx of divalent cations. This uptake mechanism was blocked by 10 μM lanthanum (data not given). In a second series of experiments, the effects of ionomycin were tested on whole cell Cl− currents developed in the presence or the absence of intracellular free Ca2+. To avoid the apparition of swelling-induced Cl−conductance, the currents were recorded in monolayers continuously perfused with hypertonic NMDG solution (Table 1,solution c + 60 mM mannitol). In the experiments shown in Fig.10 A, whole cell currents were recorded in the absence of EGTA in the pipette solution (Table 1, solution f without EGTA). After control macroscopic currents were recorded, 2 μM ionomycin was added to the bathing NMDG solution, and the stimulated currents were recorded after 1 min. In these conditions, the cytoplasmic free Ca2+ rose at 1.00 ± 0.19 μM (n = 13). Figure10 A shows that in the presence of ionomycin, the currents increased during depolarizing voltage pulses. The kinetics of the macroscopic current were clearly time dependent for depolarizing potentials with a slow-developing component. The corresponding I-V relationships for early and steady-state activated currents are given in Fig.10 A. Currents reversed at −0.25 ± 0.2 mV (n = 6). Instantaneous currents showed a slight outward rectification: the inward current at −100 mV was 263.7 ± 41.0 pA, and the outward current at +100 mV was 353.7 ± 34.0 pA. The steady-state current presented marked outward rectification with an inward current at −100 mV of 214.0 ± 74.0 pA and an outward current at +100 mV of 524.0 ± 78.0 pA (n = 6). In the experiments of Fig.10 B, whole cell currents were recorded in the presence of 5 mM EGTA in the pipette solution (Table 1,solution f). In 12 of the 20 cells analyzed, the addition of 2 μM ionomycin induced the activation of Cl− currents within 3–4 min. Under these conditions, ionomycin also enhanced the divalent-cation influx across the membrane (see above), but this increase was not accompanied by visible variations of intracellular Ca2+ (data not given). The iononycin-activated Cl−currents showed time-dependent inactivation at depolarizing step potentials of >60 mV and displayed an outwardly rectified instantaneous I-V plot (Fig.10 B) with anE rev of −0.35 ± 0.01 mV (n = 12). Overall, these currents were quite similar to those induced by hypotonic shock.
The development of hypotonicity-induced Cl− currents was further studied in the presence of several factors that could interfere with Cl− conductance.
Lanthanum. In a series of experiments, hypotonic shock was carried out in the presence of 50 μM La3+ in the bath solution. This trivalent cation irreversibily suppressed the Cl− currents (Fig. 7).
P-glycoprotein-related inhibitors. To investigate a possible involvement of P-glycoprotein in the generation of the hypotonicity-induced Cl− currents, inhibitors for this protein were tested. As shown in Fig. 7, verapamil (100 μM) and nifedipine (10 μM) completely prevented activation of Cl− currents on exposure of the cells to a hypotonic solution. Similar inhibition was also induced by the extracellular application of tamoxifen (5 μM). These inhibitors have been known to reverse multidrug resistance by inhibiting P-glycoprotein-mediated drug efflux and to block the P-glycoprotein-associated Cl− channel (53). The effects of these drugs were partially reversed while rinsing the monolayers with a solution without inhibitors. Verapamil and tamoxifen were also applied separately following preactivation of Cl− currents by hypotonic shock. As illustrated in Fig. 8, neither drug significantly affected the fully developed Cl−currents. Dideoxyforskolin has also been reported to inhibit swelling Cl− conductance thought to be related to the expression of P-glycoprotein. As shown in Fig. 7, 50 μM dideoxyforskolin prevented activation of the current on hypotonic swelling. Moreover, the drug also produced a marked irreversible inhibition (68.4 ± 4.8%, n = 3) of the preactivated swelling-induced Cl− current (Fig.8).
Intracellular ATP. All the experiments were performed with 5 mM MgATP in the pipette solution. However, to check the influence of cytosolic ATP on the generation of swelling-induced Cl−currents, a series of experiments was carried out in the absence of ATP in the pipette medium. Under these conditions, exposure of the cells to hypotonic solutions induced small Cl− currents, the amplitude of which slowly decreased and stabilized after 6–8 min (initial currents recorded at 100 mV 6–8 min after hypotonic shock in the presence of ATP, 1,937.0 ± 328.9 pA,n = 5; in the absence of ATP, 541.7 ± 34.7 pA, n = 5).
Protein kinases. To check whether activation of the Cl− current by hypotonic stress could be regulated by protein kinases, the effects of inhibitors of protein kinase A and protein kinase C were studied. When the hypotonic shock was carried out in the presence of 10 μM of protein kinase A inhibitor (PKI) in the pipette solution, the whole cell Cl− currents developed normally in all cells tested (Fig. 7). When the shock was performed in the presence of 200 nM staurosporine in the bath medium, no significant increase in Cl− currents was recorded (Fig. 7). However, bath staurosporine did not affect the magnitude of the Cl−currents once they were preactivated by the hypotonic solution (Fig.8).
K+ Currents Induced by Hypotonic Shock
The effect of hypotonic swelling was tested under Cl−-free conditions. The first experimental series was carried out with 140 mM potassium glutamate and EGTA in the pipette solution and 140 mM sodium glutamate in the bath medium. In each experimental condition, only positive current curves were observed. Figure11 Aillustrates the family of current recordings made in a hypertonic bath medium with test potential that ranged from −100 mV to +80 mV in increments of 20 mV. Outward currents were significantly different from 0 at −60 mV (23.9 ± 11.0 pA;n = 25,P < 0.05) and increased during more positive voltage pulses. These currents showed virtually no inactivation during the 400-ms pulse. The correspondingI-V curve in Fig.11 E confirms that the channels involved in this conductance were activated at depolarized potentials. The conductance measured by the maximal slope of theI-V curve averaged 24.0 ± 2.2 nS (n = 25). As illustrated in Fig.11 B, outward currents were activated when the hypertonic bath solution was replaced by an isotonic solution of identical ionic composition. Maximal current activation was reached 2 min after the onset of the osmotic shock. TheI-V curve of Fig.11 E shows that the activation of outward currents became significantly different from control from −40 mV (control current, 53.6 ± 16 pA; volume-activated current, 111.0 ± 10 pA; P < 0.01, n = 25) and increased with depolarizing potentials. The calculated maximal slope conductance increased significantly (34.5 ± 2.7 nS;P < 0.01,n = 25) compared with control values (see above). Whole cell conductance was then analyzed following the addition of 5 mM Ba2+ to the isotonic bath solution. Figure 11, Cand E, clearly indicate that, atV hold between −40 mV and +20 mV, the currents recorded in the presence of Ba2+ were significantly lower than those recorded in isotonic bath solution without Ba2+. Conversely, the currents determined for depolarizing voltage steps were not significantly altered. Experiments in the control bathing solution following this intervention showed that the effect of Ba2+ could be completely reversed (data not given). Finally, reexposing the same cells to hypertonic solution induced a decrease in the amplitude of the outward currents within 1 min (Fig. 11 C). The current amplitude returned to values slightly lower than that initially measured (Fig. 11 D), suggesting that currents generated by osmotic shock were associated with cell swelling. At negative imposed potentials, block of the outward conductance by Ba2+ strongly indicated that the currents induced by the osmotic shock were mainly the result of an efflux of K+ from the cell. Moreover, in the absence of permeable anions in the solutions, it is very likely that the outward current measured at positive potentials was also carried by K+. To further analyze the ionic nature of these conductances, a series of experiments was performed whereby all Na+ in the bathing medium was replaced by K+. Under these conditions, osmotic shock increased both inward and outward currents (Fig.12 A) with a E rev close to 0 mV. The initial current recorded at 80 mV was measured to be 3.6 times the current at −80 mV (80 mV = 2,600 ± 232 pA; −80 mV = −718 ± 94 pA;n = 9,P < 0.001). These large, outwardly rectifying currents were time independent and had a maximal slope conductance of 40.2 ± 2.6 nS (n = 9). They were carried by K+, which was the only permeable ion into pipette and bath solutions. To confirm the implication of a K+conductance, the effect of Ba2+ in the bathing medium was then tested. The recording of Fig.12 B and theI-V plot of Fig.12 E show that 5 mM BaCl2 almost completely blocked the inward current without significantly modifying the outward current. After the washout of Ba2+, the inward conductance recovered completely (Fig.12 C). Finally, reexposure of cells to a hypertonic solution inhibited both outward and inward currents (Fig. 12, D andE).
Influence of Hypotonic Shock on Relative Cell Volume
Figure13 Ashows that the reduction of the osmolarity of the perfused solution caused a rapid increase in relative cell volume. This cell swelling was followed by a RVD. One minute after the hypotonic shock, the relative cell volume reached 128.9 ± 0.8% (n = 9 cells from 3 monolayers) of the initial volume and returned to 106.1 ± 0.3% of the original volume within 2 min. To test the involvement of K+ conductance in the RVD process, experiments were carried out in the presence of 5 mM external Ba2+. When Ba2+ was perfused with the hypotonic solution, the cells never returned to their initial volume (Fig. 13 B). To demonstrate that Cl− conductance was involved in RVD of distal cells, the effect of the Cl− channel blocker NPPB was then tested. Figure 13 C shows that, when 0.1 mM NPPB was added to the hyposmotic solution, RVD was completely abolished.
In the present study, we have demonstrated that cultured DCTb cells exhibit swelling-activated whole cell Cl− currents. These currents are clearly distinct from the Ca2+-activated Cl− currents and cAMP-dependent Cl− currents we have previously described in these primary cultures (3, 35, 43). In the first series of experiments presented here, spontaneously developing Cl− currents were measured when isotonic bathing and pipette solution were used. The currents reached a maximum 6–7 min after establishment of the whole cell configuration. According to Worrell et al. (51), this increase of Cl− conductance could be the result of cell swelling in isosmotic medium. Under these conditions, the presence of a diffusion barrier within the cytoplasm could impair the complete equilibration between the cell interior and the pipette solution. Consequently, the intracellular medium would maintain a significant oncotic pressure due to the conservation of high-molecular-weight proteins. In a second series of experiments, Cl− currents also developed in cells exposed to hypotonic shock. Control currents were obtained by maintaining the cells in a bath, which was 50 mosmol/kgH2O hypertonic to the pipette solution. In 95% of cells tested, the currents were of very small amplitude and time independent. The measuredE rev was close to 0 mV, indicating that most of the control current was carried by Cl− ions. The remaining 5% of cells developed swelling Cl− currents immediately after the whole cell configuration was obtained. This was probably because the hypertonicity of the bath medium was not sufficient to prevent cell swelling. In the hypertonic solution, the basal Cl− currents remained small and almost constant for at least 8 min. When the osmolarity of the bath solution was lowered to 290 mosmol/kgH2O, whole cell conductance increased ∼10-fold within 6–8 min. Once maximally developed, the current remained very stable and could be rapidly inhibited by reexposing the cells to the hypertonic bath solution.
The biophysical and the pharmacological characteristics of the Cl− conductances induced either spontaneously in isotonic medium or induced by hypotonic stress show strong similarities with the properties of swelling-activated Cl− currents described in many other epithelial cells (8, 9, 21, 40, 49, 52). The peak currents exhibited an outwardly rectifying I-Vrelationship, whereas the steady-state currents showed a strong decay at depolarizing pulses. This decay was time dependent and increased with increasing cell depolarization. As previously suggested (47), this phenomenon may well correspond to the closure of channels already activated at the holding potential. Nevertheless, the physiological relevance of this inactivation is questionable, since the current inactivation was obtained at voltages far removed from the Cl− equilibrium potential. The swelling-induced currents were carried mainly by Cl−. This was confirmed by the removal of extracellular Cl−, which strongly reduced the amplitude of the outward currents. Moreover, when the major cations in the pipette and bath solutions were replaced with NMDG,E rev remained close to zero, as in symmetrical chloride solutions. This finding clearly demonstrates that the major part of the swelling-induced current was Cl− selective. However, when glutamate was substituted for Cl− in the bathing solution, the E rev was shifted to just 60 mV. Because the theoreticalE rev for glutamate substitution was >100 mV, the Cl− channel could be slightly permeable to glutamate. Low levels of organic anion permeability have already been reported in kidney cells (48). Interestingly,E rev for glutamate was dependent on the duration of the stimulation. Thus the relative permeability for glutamate was higher when calculated using instantaneous currents than using steady-state conductance. Two hypotheses could be proposed to support this observation:1) Cl− and glutamate could permeate the cell membrane through separate channels both being activated by hypotonicity. The channels activate at depolarizing potential, but the conductance deactivation kinetic could be faster for glutamate than for Cl−currents. Volume-sensitive anion channels with a relative high permeability to glutamate, aspartate, and taurine were already found in Madin-Darby canine kidney (MDCK) cells (2). However, according to the recent findings of Boese et al. (4), the hypotonic stress would activate a common pathway for conductive Cl− and amino acids (as glutamate or taurine). 2) Cl− and glutamate fluxes process through the same channel. It is possible that large depolarizing potentials modify the relative permeability at the onset of the voltage pulse. For example, depolarization could induce important changes in channel conformation (54), resulting in transitory modifications in the channel selectivity.
The halide selectivity sequence permits the different types of Cl− channels to be identified. In DCTb cells, the sequence for the hypotonicity-induced Cl− current was I− = Br− > Cl−. In many cell types studied, swelling-induced Cl− channels have been reported to be more permeable to I− than to Br− (6, 49). However, this is not a constant finding, because, in some tissues, the anion conductivity sequence was found identical to that we reported in the present study (8). Sensitivity to various anion channel blockers also helps to distinguish the type of Cl− channel under investigation. For this purpose, we examined the effects of NPPB, DPC, and DIDS. NPPB and DIDS modified mainly the outward current, whereas DPC strongly blocked both outward and inward currents. In the literature, the voltage dependence of the blocking effect of DIDS and the voltage independence of the inhibitory effect of both DPC and NPPB are common characteristics of swelling-activated Cl− currents. In cultured DCTb cells, NPPB acted in a similar fashion to DIDS, suggesting that their blocking mechanisms may be identical. However, it is necessary to note than the effect of DIDS on the inward current was much more constant from one cell to another than the effect of NPPB.
Despite abundant literature, the precise mechanism underlying the activation of swelling-induced Cl− current remains unclear. In several studies, an increase in cytosolic Ca2+ has been reported during the hypotonic stress (24, 25). In the present study, we found that the activation of a swelling-induced Cl− conductance took place in the presence of high concentrations of EGTA in the pipette solution. Moreover, we also demonstrated that no significant cytosolic Ca2+ variation could be detected during the hypotonic shock. These results demonstrate that DCTb Cl− conductance during swelling is probably insensitive to intracellular Ca2+. This finding agrees with that reported for a number of cells, including human sweat gland (11), endothelial (31), epididymal (8), intestinal 407 (21), and T84 cells (51). Moreover, in proximal convoluted tubules (PCT), the changes in cytosolic Ca2+ were shown to play no role in RVD and, therefore, on swelling-induced activation of ion conductances (5). Nevertheless, other studies have underlined a positive role of cytoplasmic Ca2+ in the control of the volume-sensitive Cl−currents (12, 24). The question of whether Ca2+ is involved in the activation of these currents is not so clear cut. Interestingly, we have demonstrated that the removal of external Ca2+ just before the hypotonic shock completely impaired Cl− current activation, suggesting that Ca2+ influx could participate in activating the Cl− channels. The observation that the Ca2+ channel blocker La3+ also abolished the Cl− current activation corroborates this hypothesis. However, to better support this hypothesis, we studied the effects of ionomycin in the activation of Cl− channels in hypertonic bathing medium. The ionophore induced two types of Cl− conductances, depending on the presence of high EGTA concentrations in the pipette solution. In the absence of chelator, the Cl− currents induced by ionomycin were roughly identical to the Ca2+-dependent Cl− currents previously found in cultured DCTb (3). These currents were likely stimulated by an increase in cytoplasmic Ca2+ due to Ca2+ release from intracellular stores. By contrast, in the presence of EGTA, the application of ionomycin increased the uptake of Mn2+. This increase is evidence of Ca2+ uptake across the membrane (7). This maneuver induced large Cl− currents, which inactivated during depolarizing voltage steps. Although they are not fully characterized, these currents closely resemble the swelling-activated Cl−currents described in the present study. Finally, Ca2+ entry across the plasma membrane could be one of the mediators of the activation of Cl− currents by hypotonicity. The role of external Ca2+ was already postulated by McCarty and O’Neil (25). They showed that RVD in proximal straight tubules was highly dependent on the extracellular Ca2+ concentration. Moreover, they concluded that Ca2+ channels may be responsible for a swelling-activated Ca2+ entry. In previous papers, we have shown that Ca2+ channels are present in the apical membrane of DCTb cells (34) and may represent the apical influx pathway for transepithelilal Ca2+ transport. These Ca2+ channels are distinct from the nonselective cation channels and are blocked by nifedipine, verapamil, and La3+. In the present study, the problem has been to better characterize the involvement of the calcium channels in the control of swelling-induced conductance. Nifedipine and verapamil are the only drugs that block the Ca2+ channel in DCTb (36), and these drugs also directly interfere with the swelling Cl− channels (see below). While the response to hypotonic shock was developing, removal of external Ca2+ did not affect the Cl− current. Very similar results were previously found in pancreatic duct cells (49), whereon the authors concluded that Ca2+ is only involved in events occurring early in the mechanism. Their conclusions also support our data.
In most of the experiments described here, ATP was included in the pipette solutions. When ATP was omitted from the pipette, hypotonic stress always triggered Cl−conductance, but the currents never reached their maximum amplitude and rapidly decreased with time. It is therefore probable that this decrease began as the endogenous ATP was washed out after the whole cell configuration was established. This characteristic of the swelling-induced Cl− current is similar to that reported previously in several tissues (11, 31, 49). However, divergent results were also obtained in other cells (18, 40). One of the hypotheses advanced to explain the action of ATP is that swelling-induced Cl−currents are associated with the multidrug-resistance P-glycoprotein. Thus the gradual depletion of endogenous ATP would decrease the activity of this protein.
To determine whether P-glycoprotein is implicated in the swelling-induced Cl−conductance of DCTb cells, we have tested the effect of different drugs that have been reported to inhibit Cl− currents thought to be related to the expression of P-glycoprotein (46, 47). Verapamil, nifedipine, tamoxifen, and dideoxyforskolin strongly impaired the development of swelling currents. However, tamoxifen and verapamil failed to produce significant inhibition of preactivated Cl− current, whereas dideoxyforskolin almost completely supressed it. Taken together, these findings indicate that P-glycoprotein could participate in the regulation of the swelling-induced Cl− current but that this protein is probably not itself the actual Cl− channel. This observation supports recent literature reports of different cell types (17, 45).
P-glycoprotein is a drug transporter modulated by protein kinase C (PKC). We therefore examined whether PKC might also play a role in the control of the swelling-activated Cl− current. Bath application of staurosporine, which is a membrane-permeable inhibitor of PKC, completely inhibited the development of the swelling-induced conductance. However, staurosporine never suppressed the current when applied after the commencement of the hypotonic chock. These results agree with those obtained for pancreatic duct cells (49) and suggest that PKC regulates swelling-induced Cl− currents in cultured DCTb. If it is generally accepted that PKC is implicated in the control of this conductance, the exact nature of this control is under discussion. In fact, according to some authors, activation of PKC inhibits the channel by phosphorylation of P-glycoprotein (17), whereas others have shown that PKC increases the channel activity (49) or is not involved in the mechanism of regulation (42, 45). On the basis of data obtained in pancreatic duct cells, Verdon et al. (49) have proposed a very attractive hypothesis. They suggest that cell swelling leads to an influx of Ca2+ and that the concomittant increase of intracellular Ca2+ will be sufficient to activate PKC. The PKC in turn phophorylates a regulatory protein. This protein could be the P-glycoprotein. Such an intracellular signaling pathway was proposed by Nishizuka (32) in a recent review, at least up to the point activation of the PKC.
It remains to be shown how intracellular Ca2+ can increase in the presence of a high EGTA concentration. The observations of Evans and Marty (10) shed light on this problem by indicating that, with EGTA as a buffer, a whole region of the cell could escape control by the Ca2+ buffer. Because this region could extend to a large part of the plasma membrane (10), a local transient increase of Ca2+ could arise without being detected by fluorescence methods.
Because activation of the PKA pathway is known to regulate Cl− channels in epithelia (14), we therefore investigated this possibility on the swelling-induced Cl−conductance of DCTb cells. The use of the specific PKA inhibitor peptide, PKI (5, 24), clearly indicates that PKA is not involved in activating the channel. This finding is strengthened by experiments in which we demonstrated that the application of forskolin did not modify the activation of the Cl− conductance during the hypotonic shock (data not given). In epithelial cells, the activation of the swelling-induced Cl−conductance generally appears to be independent of cAMP (31, 49).
In many cells, swelling induced by exposure to hyposmotic solutions is followed by RVD mediated by KCl loss via K+ and Cl− channels. Concerning the mammalian kidney, RVD has been primarily studied in PCT (19, 22, 23,25, 41, 44), in medullary thick ascending limb cells (30), and in the cell line MDCK (1, 20). In a previous study (22), we demonstrated that primary cultures of PCT responded to hypotonic shock by activating K+ and Cl− conductance. Further experiments on rabbit proximal straight tubules brought evidence for a swelling-sensitive Cl−conductance in the basolateral membrane (23, 37). Swelling-induced Cl− channels were also described in the distal nephron, notably in RCCT-28A cells, an immortalized cell line derived from rabbit collecting duct (39) and recently in primary culture of rat inner medullary collecting duct cells (50) and in M-1 cell line obtained from mouse collecting duct (29). Interestingly, the Cl−channel recorded in RCCT-28A cells is regulated by a signaling pathway that involves PKC and could mediate Cl− efflux during RVD. To determine whether cultured DCTb cells develop RVD after a hypotonic shock, we used a simple fluorescence method for studying relative cell volume variations (44). The findings indicate that DCTb cells are sensitive to osmolarity changes in the bathing medium and that they are capable of RVD after a hypotonic shock. The RVD process was impaired by NPPB, confirming the implication of a Cl−-conductive pathway. Moreover, the observation that Ba2+ completely blocked RVD strongly indicates the involvement of K+ conductance. The first experiments that we have performed to characterize cation conductance during osmotic challenge indicate that K+ currents also increased during the hypotonic shock. To characterize this K+ conductance, it was necessary to use Cl−-free solutions, because, in the presence of KCl, the swelling-activated current was mainly due to Cl− (data not given). In the presence of symmetrical K+ solutions, the whole cell conductance activated by the osmotic shock was Ba2+ sensitive. Ba2+ blocked mainly the inward currents, indicating a strong voltage dependence as is expected for Ba2+. Ba2+-sensitive Ca2+-dependent maxi K+ channels have been found in the apical membrane of cultured proximal tubules (28), cultured cortical ascending limb of the loop of Henle (27), and cortical collecting duct cells (13, 15). However, in the present experiments, the pipette solution contained 5 mM EGTA, and it is unlikely that Ca2+-activated maxi K+ currents participated in the resting K+ conductance. As discussed above for Cl−channels, the osmotic shock could modulate Ca2+ influx through Ca2+ channels, and a local increase of cytosolic Ca2+ could activate the Ca2+-sensitive maxi K+ channel. A similar mechanism has already been postulated to explain the RVD in primary cultures of renal proximal tubule (19, 22, 44).
It is of interest to examine the effect of osmotic stress on Cl− and K+ conductances determined close to the resting membrane potential. With consideration to an imposed potential of −60 mV 2 min after the beginning of the osmotic shock, the outward Cl−current was increased by 56% (initial current att = 0 min, −88.2 ± 9.0 pA; initial current at t = 2 min, −157.8 ± 17.0 pA; n= 19, P < 0.01), whereas the outward K+ current was not significantly modified. An increase in outward K+ current became significant at −40 mV (see results). Therefore, cell swelling induced an increase predominantly in Cl− current. Such increase has also been demonstrated in rat colonic crypt (9) and rat mesangial cells (33). According to these studies, hypotonic cell swelling could induce a depolarization of membrane potential. If it is also the case in our experiments, the activation of a swelling-dependent Cl− conductance might depolarize the membrane potential toward a value where the efflux of K+ from the cell becomes significant. Finally, in cultured DCTb, a hyposmotic shock could well induce a net loss of KCl, thus allowing the cells to regulate their volume.
In summary, DCTb cells in primary culture exhibit hypotonic shock-induced Cl− and K+ conductances and RVD process. The Cl− currents described here share characteristics with the P-glycoprotein Cl− currents described in other systems (31, 46, 52), although P-glycoprotein is more likely to be a regulator of the channel than the chloride channel itself (17). The accompanying K+ currents are strongly voltage dependent and blocked by external Ba2+. Thus the activation of Cl− and K+ currents by an osmotic shock may be implicated in regulatory volume decrease in DCTb cells.
Address for reprint requests: P. Poujeol, UMR-CNRS 6548, Bâtiment Sciences Naturelles, Université de Nice-Sophia Antipolis, Parc Valrose, 06108 Nice Cedex 2, France.
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