currents in primary cultures of mouse nephron - American Journal of ...

Am J Physiol Renal Physiol 284: F796–F811, 2003. First published December 10, 2002; 10.1152/ajprenal.00237.2002.

CFTR null mutation altered cAMP-sensitive and swellingactivated Cl⫺ currents in primary cultures of mouse nephron Herve´ Barrie`re, Radia Belfodil, Isabelle Rubera, Michel Tauc, Chantal Poujeol, Michel Bidet, and Philippe Poujeol Unite´ Mixte de Recherche Centre National de la Recherche Scientifique 6548, Universite´ de Nice-Sophia Antipolis, 06108 Nice Cedex 2, France Submitted 26 June 2002; accepted in final form 3 December 2002

Barrie`re, Herve´, Radia Belfodil, Isabelle Rubera, Michel Tauc, Chantal Poujeol, Michel Bidet, and Philippe Poujeol. CFTR null mutation altered cAMP-sensitive and swelling-activated Cl⫺ currents in primary cultures of mouse nephron. Am J Physiol Renal Physiol 284: F796–F811, 2003. First published December 10, 2002; 10.1152/ajprenal.00237.2002.—The role of cystic fibrosis transmembrane conductance regulator (CFTR) in the control of Cl⫺ currents was studied in mouse kidney. Whole cell clamp was used to analyze Cl⫺ currents in primary cultures of proximal and distal convoluted and cortical collecting tubules from wild-type (WT) and cftr knockout (KO) mice. In WT mice, forskolin activated a linear Cl⫺ current only in distal convoluted and cortical collecting tubule cells. This current was not recorded in KO mice. In both mice, Ca2⫹dependent Cl⫺ currents were recorded in all segments. In WT mice, volume-sensitive Cl⫺ currents were implicated in regulatory volume decrease during hypotonicity. In KO mice, regulatory volume decrease and swelling-activated Cl⫺ current were impaired but were restored by adenosine perfusion. Extracellular ATP also restored swelling-activated Cl⫺ currents. The effect of ATP or adenosine was blocked by 8-cyclopentyl-1,3-diproxylxanthine. The ecto-ATPase inhibitor ARL-67156 inhibited the effect of hypotonicity and ATP. Finally, in KO mice, volume-sensitive Cl⫺ currents are potentially functional, but the absence of CFTR precludes their activation by extracellular nucleosides. This observation strengthens the hypothesis that CFTR is a modulator of ATP release in epithelia.

(CFTR) protein has been detected by electrophysiological techniques in a variety of cultured cells of the renal tubule, such as distal convoluted tubule (DCT) (21), cortical collecting tubule (CCT) (2, 30), and inner medullary collecting duct (10). In these segments, the presence of CFTR is correlated with activation of a cAMP-activated Cl⫺ current. However, along the nephron, CFTR is not always associated with these Cl⫺ currents. For instance, despite the presence of CFTR transcripts, CFTR expression, along with forskolininduced conductance, was not detected in rabbit prox-

imal tubule in primary culture (21). This observation highlights the fact that CFTR could play an important role in the control of different channels in kidney tissue. Such control is now well established in secretory epithelia. In these structures, besides the cAMP-sensitive Cl⫺ secretion, CFTR controls the epithelial Na⫹ channel (12, 14, 18, 28) and the outwardly rectifying Cl⫺ channel (24). Moreover, CFTR is also needed for an effective volume regulation in airway and intestinal epithelia (31, 32), suggesting that it could modulate K⫹ and Cl⫺ channels implicated in regulatory volume decrease (RVD). Indeed, these multiple functions of CFTR could explain the different phenotypes induced by cystic fibrosis (CF) in secretory epithelia. In contrast, the role of CFTR in the kidney remains uncertain, inasmuch as there is no major disruption of renal function in CF patients (27). Nevertheless, the reduced renal excretion of NaCl observed in CF indicates that Cl⫺ and Na⫹ channels could be dependent on CFTR expression, suggesting that mutation of CFTR could induce a primary defect in renal function. A better understanding of the function of CFTR in the kidney, therefore, seems to be necessary. For this reason, we chose to investigate the role of CFTR along the nephron using primary cultures of proximal convoluted tubules (PCT), DCT, and cortical collecting ducts microdissected from the kidney of cftr⫺/⫺ and cftr⫹/⫹ mice. The cftr⫺/⫺ mice lack cAMP-activated Cl⫺ currents in the colon, airways, and exocrine pancreas cells (6) and represent a useful model for studying the different ion channel defects due to CF. In the present study, using patch-clamp methodology, we confirmed that cAMP-sensitive Cl⫺ conductances measured in primary cultures of DCT and cortical collecting tubule (CCT) cells are linked to CFTR integrity. Moreover, in contrast to the data reported in the literature on airways and endothelial cells (33), an increase in Ca2⫹dependent Cl⫺ channels does not compensate for the lack of CFTR Cl⫺ channels in renal tissue. The PCT, DCT, and CCT cells from cftr⫺/⫺ mice lost their capacity to regulate their volume after a hypotonic shock because of the impairment of swelling-activated Cl⫺ channels. In cftr⫺/⫺ cells, the activity of these

Address for reprint requests and other correspondence: P. Poujeol, UMR CNRS 6548, Baˆtiment Sciences Naturelles Universite´ de NiceSophia Antipolis, Parc Valrose, 06108 Nice Cedex 2, France (E-mail: [email protected]).

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.

kidney; cystic fibrosis; cell volume; regulatory volume decrease

THE CYSTIC FIBROSIS TRANSMEMBRANE CONDUCTANCE REGULATOR

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0363-6127/03 $5.00 Copyright © 2003 the American Physiological Society

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CFTR AND RENAL CL⫺ CONDUCTANCE

channels could be restored by external application of adenosine. This suggests that CFTR controls the swelling-activated Cl⫺ channels by modulating adenosine autocrine production in renal cells. MATERIALS AND METHODS

Animals Knockout CFTR mice were generated with the genetargeting methodology previously described (26) at Centre de De´ veloppement des Techniques Avance´ es pour l’Expe´ rimentation Animale (Orle´ ans, France). This strain of mice was originally derived from ES129/Sv cells injected into C57BL/6 embryos. They were backcrossed with C57BL/6 mice for three generations and then intercrossed. Mice were allowed free access to food and water in a facility at 25 ⫾ 1°C with a 12:12-h light-dark cycle. The 4- to 6-wk-old wild-type cftr⫹/⫹ mice and cftr⫺/⫺ mice homozygous for the disrupted cftr gene were killed by cervical dislocation, and the kidneys were removed. All experiments were performed in accordance with the guidelines of the French Agricultural Office and the legislation governing animal studies. Primary Cell Cultures PCT, DCT, and collecting tubules were microdissected under sterile conditions. Kidneys were perfused with Hanks’ solution (GIBCO) containing 700 kU/l collagenase (Worthington), cut into small pyramids that were incubated for 1 h at room temperature in perfusion buffer (160 kU/l collagenase, 1% Nuserum, and 1 mM CaCl2), and continuously aerated. The pyramids were then rinsed thoroughly in the same buffer devoid of collagenase. The individual nephrons were dissected by hand in this buffer under binoculars using stainless steel needles mounted on Pasteur pipettes. The criteria used to identify the nephron segments have been described elsewhere (4). Briefly, PCT corresponded to the 1to 1.5-mm segment of tissue located immediately following the glomerulus. The DCT portion was the segment between the macula densa and the first branching with another tubule [i.e., connecting tubule (CNT)]. The CNT segment was discarded. The CCT was identified as the straight, poorly branched portion that followed the CNT segment. After they were rinsed in dissecting medium, tubules were transferred to collagen-coated 35-mm petri dishes filled with culture medium composed of equal quantities of DMEM and Ham’s F-12 (GIBCO) containing 15 mM NaHCO3, 20 mM HEPES, pH 7.4, 1% serum, 2 mM glutamine, 5 mg/l insulin, 50 nM dexamethasone, 10 ␮g/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 removed 4 days after seeding and then every 2 days. Electrophysiological Studies Whole cell currents were recorded from 6- to 20-day-old cultured cells grown on collagen-coated supports maintained at 33°C for the duration of the experiments. The rupturedpatch whole cell configuration of the patch-clamp technique was used. Patch pipettes (2- to 3-M⍀ resistance) were made from borosilicate capillary tubes (1.5 mm OD, 1.1 mm ID; Popper Manufacturing) using a two-stage vertical puller (model PP 83, Narishige, Tokyo, Japan) and filled with a solution containing (in mM) 140 N-methyl-D-glucamine (NMDG) chloride, 1 or 5 EGTA, 5 MgATP, and 10 HEPES, AJP-Renal Physiol • VOL

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pH 7.4. The bath solution contained (in mM) 140 NMDG chloride, 1 CaCl2, 60 mannitol, and 10 HEPES, pH 7.4. Cells were observed using an inverted microscope; the stage of the microscope was equipped with a water robot micromanipulator (model WR 89, Narishige). The patch pipette was connected via an Ag-AgCl wire to the head stage of a patch amplifier (model RK 400, Biologic). 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 (model RK 400) were used to compensate for cell capacitance. No series resistance compensation was applied, but experiments in which the series resistance was ⬎20 M⍀ were discarded. 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. Data acquisition and analysis. Voltage-clamp commands, data acquisition, and data analysis were controlled via a computer equipped with a Digidata 1200 interface (Axon Instruments). pCLAMP software (versions 5.51 and 6.0, Axon Instruments) was used to generate whole cell currentvoltage (I-V) relations, with the membrane currents resulting from voltage stimuli filtered at 1 kHz, sampled at 2.5 kHz, and stored directly on the computer hard disk. Cells were held at ⫺50 mV, and 400-ms pulses from ⫺100 to ⫹120 mV were applied in 20-mV increments every 2 s. Cell Volume Measurement The relative cell volume was monitored by image analysis with fura 2 as fluorescent volume indicator, as previously reported (22). Six- to 20-day-old cell monolayers grown on petri dishes were loaded with a solution of 2 ␮M fura 2 containing 0.01% pluronic acid for 20 min at 37°C and then washed with an NaCl solution. The fluorescence was monitored with 360-nm excitation wavelength. At 360 nm, the variations in the signal emitted by the probe are directly proportional to the variations in 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 10 HEPES, pH 7.4 [osmotic pressure (Posm) ⫽ 320 mosmol/kgH2O] at 30 ml/min, and images were averaged eight times and recorded every 5 s for 15 min. Once the fluorescence was stabilized, a hypotonic shock was induced by perfusing the NaCl solution without mannitol (Posm ⫽ 200 mosmol/kgH2O). The relative change in cell volume was estimated from the fluorescent signal by assuming that a 30% decrease in osmolarity caused a decrease in the fluorescent signal corresponding to a maximum swelling of 30% compared with the initial volume. The means of relative volume changes were obtained by analysis of 10–20 zones in each culture (n) chosen with the software. Each zone delimited a cytoplasmic area chosen in individual cells. Image analysis. The optical system was composed of a Zeiss ICM-405 inverted microscope and a Zeiss ⫻40 objective, which was used for epifluorescent measurement with a 75-W xenon lamp. The excitation beam was filtered through a narrow-band filter centered at 360 nm, mounted in a motorized wheel (model Lambda 10-2, Sutter Instrument), and 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

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intensified camera (Extended ISIS, Photonic Science, Sussex, UK). The eight-bit Extended ISIS camera was equipped with an integration module to maximize signal-to-noise ratio. The video signal from the camera proceeded to an image processor integrated in a DT2867 image card (Data Translation) installed in a Pentium 100 personal computer. The processor converts the video signal to 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 with the AIW software (version 2.0, Axon Instruments). The final calculations were made using Excel software (Microsoft). Calibration. We used the methods described by Tauc et al. (29) using 2⬘,7⬘-bis(2-carboxyethyl)-5(6)-carboxyfluorescein and improved more recently by Raat et al. (17) using fura 2. After cells were loaded with the fluorescent probe in the culture medium, they were perfused with a solution adjusted to various osmolarities (150–400 mosmol/kgH2O) by omitting mannitol. For each osmolarity, two images were stored, averaged, and subsequently corrected for fading after background subtraction. The mean fluorescence (360 nm) of five areas was plotted against the inverse of Posm (in mosmol/ kgH2O). Data showed that when the cells were exposed to a hyposmotic solution, fluorescence decreased linearly with Posm according to Boyle’s law. To verify that cells in culture behave as osmometers in a reversible manner, we performed experiments in which the cultures were perfused successively and randomly with 200–300 mosmol/kgH2O solutions. The fluorescent signal was related to Posm in a reversible way. In all calibration experiments, images were recorded 1–2 min after the beginning of perfusion, at which time the swelling in hypotonic solutions reached the maximum value. These methods measure variations in the relative volume as a function of Posm of the perfusion medium (29). Intracellular Ca2⫹ Measurements Intracellular Ca2⫹ concentration ([Ca2⫹]i) was measured in cells grown in petri dishes and loaded for 45 min at room temperature with a solution of 2 ␮M fura 2-AM containing 0.01% pluronic acid. The cells were washed with NaCl solution containing (in mM) 140 NaCl, 5 KCl, 1 MgSO4, 5 glucose, 20 HEPES, pH 7.40, and 1 Tris. Cells were successively excited at 350 and 380 nm, with images digitized and stored on the computer hard disk for later analysis. Each raw image was the result of an integration of four to five frames averaged four times. The acquisition rate was one image every 10 s. For each monolayer, [Ca2⫹]i was monitored in 18–20 random cells. The equation of Grynkiewicz et al. (9) was used to calculate [Ca2⫹]i from the dual wavelength-to-fluorescence ratio. Expression in Cultured Cells The cDNA encoding CFTR was introduced into a polycistronic expression vector derived from the pIRESneo plasmid (cytomegalovirus promoter; Clontech) in which the neomycin resistance gene had been replaced by cDNA encoding the chain of the human CD8 cell surface antigen. Cells were transfected using the DAC-30 method according to the manufacturer’s instructions (Eurogentec, Herstal, Belgium). Sixday-old cultured cells grown on 35-mm-diameter petri dishes were serum starved for 24 h before transfection. Transfected cells with 2 ␮g of CD8-CFTR coexpress CFTR and CD8 at their plasma membrane and can be visualized using antiCD8 antibody-coated beads (Dynabeads M-450, Dynal, Oslo, AJP-Renal Physiol • VOL

Norway) (11a). Cells were electrophysiologically tested 48 h after transfection. Chemicals 5-Nitro-2-(3-phenylpropylamino)-benzoic acid (NPPB; Calbiochem) was prepared at 100 mM in DMSO and used at 0.1 mM in final solutions. 4–4⬘-Diisothiocyanostilbene-2,2⬘-disulfonic acid (DIDS) was directly dissolved at a final concentration of 1 mM. Forskolin and ionomycin were prepared at 10 and 2 mM, respectively, in ethanol and used at 10 and 2 ␮M, respectively, in bath medium. DIDS, forskolin, ARL67156 (6-N,N-diethyl-␤-␥-dibromomethylene-D-adenosine-5⬘triphosphate trisodium), apamin, and ionomycin were obtained from Sigma (Saint Quentin Fallavier, France). Fura 2-AM (Molecular Probes) was dissolved at 3 mM in DMSO and added to the loading solution at a final concentration of 2 ␮M, along with 0.01% pluronic acid. RESULTS

Cl⫺ Currents Activated by Forskolin Experiments were performed in a hyperosmotic extracellular solution (350 mosmol/kgH2O) to characterize Cl⫺ currents activated by forskolin in PCT, DCT, and CCT cells. Under these conditions, volume-activated Cl⫺ currents could not be detected. In the absence of forskolin, the voltage-step protocol elicited small currents that changed linearly with the membrane potential in PCT, DCT, and CCT cell cultures from kidneys of cftr⫹/⫹ and cftr⫺/⫺ mice (data not shown). In cftr⫹/⫹ mice, exposure of cultured DCT and CCT cells to 10 ␮M forskolin induced an increase in membrane current amplitudes (Fig. 1A) that reached a peak value 3–4 min after the beginning of the perfusion. These activated currents exhibited a linear I-V relationship, with a reversal potential (Erev) of ⫺1.3 ⫾ 2.3 mV and a conductance of 6.4 ⫾ 0.6 nS in DCT cells and an Erev of ⫺2.2 ⫾ 2.9 mV and a conductance of 6.0 ⫾ 1.2 nS in CCT cells (n ⫽ 8 monolayers from 4 mice). In contrast, application of forskolin did not modify the currents recorded in cultured PCT cells (Fig. 1A). The unstimulated whole cell current in these cells reversed at ⫺3.1 ⫾ 1.2 mV, with a slope conductance of 1.1 ⫾ 0.1 nS (n ⫽ 9 monolayers from 4 mice). As expected, in cftr⫺/⫺ mice, addition of forskolin did not stimulate Cl⫺ conductance in all the cultured segments studied. This observation clearly indicated that, in DCT and CCT cells from cftr⫹/⫹ mice, the Cl⫺ conductance stimulated by forskolin was related to CFTR. The forskolin-sensitive Cl⫺ currents measured at ⫹100 mV are compared in primary cultures of PCT, DCT, and CCT from cftr⫹/⫹ and cftr⫺/⫺ mice in Fig. 1B. Only DCT and CCT from wild-type mice exhibited forskolin-activated Cl⫺ currents that were blocked by 0.1 mM NPPB and insensitive to 1 mM DIDS in the extracellular bath. Moreover, in these segments, replacing external Cl⫺ with I⫺ strongly inhibited the Cl⫺ currents activated by forskolin and caused Erev to shift toward positive values: Erev for I⫺ ⫽ 37.5 ⫾ 1.4 and 17.0 ⫾ 6.8 mV for DCT and CCT, respectively (n ⫽ 4 monolayers from 4 different mice).

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Fig. 1. A: forskolin-induced whole cell Cl⫺ currents in proximal convoluted tubule (PCT), distal convoluted tubule (DCT), and cortical collecting tubule (CCT) cells in primary culture of cftr⫹/⫹ and cftr⫺/⫺ mice. Membrane voltage was held at ⫺50 mV and stepped to test potential of ⫺100 to ⫹120 mV in 20-mV increments. Whole cell currents were recorded after 3 min of extracellular perfusion of 10 ␮M forskolin in the presence of 1 mM EGTA and 5 mM MgATP in pipette solution and 1 mM CaCl2 in extracellular bath. B: effects of 1 mM DIDS, 0.1 mM 5-nitro-2-(3-phenylpropylamino)-benzoic acid (NPPB), and extracellular Cl⫺ substitution by I⫺ on forskolin-induced whole cell Cl⫺ currents measured at ⫹100 mV. Values are means ⫾ SE; n, number of monolayers from 4 different mice.

Ca2⫹-Induced Cl⫺ Currents Whole cell currents were recorded with Ca2⫹-free (1 mM EGTA) solutions containing NMDG chloride as the major cation in the pipette and with extracellular solutions containing NMDG chloride and 1 mM CaCl2. The extracellular solution was adjusted to 350 mosmol/ kgH2O with mannitol to avoid inducing volume-activated currents. The control macroscopic currents were recorded, and 2 ␮M ionomycin was added to the NMDG chloride bathing solution. Stimulated currents were recorded after 2 min. Figure 2A shows the currents recorded in PCT, DCT, and CCT monolayers from cftr⫹/⫹ and cftr⫺/⫺ mice. In all cultured segments from both types of mice, addition of ionomycin stimulated Cl⫺ currents, which increased during depolarizing voltage pulses. The kinetics of the macroscopic current were clearly time dependent for depolarizing AJP-Renal Physiol • VOL

potentials with a slowly developing component. In cultured PCT cells from cftr⫹/⫹ mice, currents reversed at ⫺2.6 ⫾ 0.3 mV (n ⫽ 16 monolayers from 4 different mice). Instantaneous currents measured 5 ms after the beginning of the stimulation were almost linear, with an inward current of 485 ⫾ 34 pA at ⫺100 mV and an outward current of 635 ⫾ 42 pA at ⫹100 mV. The steady-state current at 380 ms exhibited a marked outward rectification, with an inward current of 352 ⫾ 40 pA at ⫺100 mV and an outward current of 834 ⫾ 79 pA at ⫹100 mV (n ⫽ 16 monolayers from 4 different mice). When the steady-state current measurements were used to calculate the Cl⫺ conductance, the maximal outward conductance was significantly different from the maximal inward conductance: 9.6 ⫾ 0.8 and 2.4 ⫾ 0.4 nS, respectively (n ⫽ 16 monolayers from 4 different mice; P ⬍ 0.02). Finally, as shown in Fig. 2B,

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Fig. 2. A: Ca2⫹-induced whole cell Cl⫺ currents in PCT, DCT, and CCT cells in primary culture of cftr⫹/⫹ and cftr⫺/⫺ mice. Membrane voltage was held at ⫺50 mV and stepped to test potential of ⫺100 to ⫹120 mV in 20-mV increments. Whole cell currents were recorded after 2 min of extracellular perfusion of 2 ␮M ionomycin in the presence of 1 mM EGTA and 5 mM MgATP in pipette solution and 1 mM CaCl2 in extracellular bath. B: effects of 1 mM DIDS on Ca2⫹-induced whole cell Cl⫺ currents. Steady-state currents at ⫹100 mV were measured 380 ms after onset of pulse. Values are means ⫾ SE; n, number of monolayers from 4 different mice.

1 mM DIDS inhibited the ionomycin conductance by 85.6 ⫾ 5.1% (n ⫽ 16 monolayers from 4 different mice). In cultured DCT and CCT cells from cftr⫹/⫹ mice, the Cl⫺ currents induced by ionomycin strongly resembled those induced in cultured PCT cells: the steady-state currents were outwardly rectifying and strongly blocked by 1 mM DIDS (Fig. 2B). Moreover, as illustrated in Fig. 2, the ionomycin-sensitive Cl⫺ conductances measured in cftr⫺/⫺ mice exhibited characteristics roughly similar to those measured in cftr⫹/⫹ mice, indicating that CFTR does not participate in the Ca2⫹-sensitive Cl⫺ conductance along the mouse nephron. Cl⫺ Currents Induced by a Hypotonic Shock To study the effects of changes in Posm on the development of Cl⫺ conductance, currents were induced by AJP-Renal Physiol • VOL

osmotic shock. Whole cell currents were recorded with Ca2⫹-free (5 mM EGTA) pipette solutions containing NMDG chloride and maintained at 290 mosmol/ kgH2O. Moreover, to eliminate any participation of cations in the inward current, experiments were carried out after Na⫹ in the bath solution was replaced with NMDG chloride and in the presence of 1 mM CaCl2. In cftr⫹/⫹ mice, the control currents were first measured in cultured PCT, DCT, and CCT cells with an extracellular solution osmolarity of 350 mosmol/ kgH2O. Under this condition, the voltage-step protocol elicited small time-independent currents that changed linearly with the membrane voltage and had Erev of ⫺6.2 ⫾ 1.5, ⫺1.5 ⫾ 1.6, and ⫹1.9 ⫾ 0.8 mV for PCT, DCT, and CCT cells, respectively (n ⫽ 4 monolayers from 4 different mice). Because of their small amplitude, the nature of these currents was not analyzed further.

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The monolayers were then perfused with a 290 mosmol/kgH2O solution. Figure 3A gives the currents recorded in PCT, DCT, and CCT cells. In ⬎95% of the cftr⫹/⫹ cells, an increase in the whole cell current was observed within 1 min. In all epithelial cell types, the currents reached a maximum after 4–5 min. Under these conditions, the initial currents recorded at ⫹100 mV were ⬃2.5 times the amplitude of the currents recorded at ⫺100 mV. These large, outwardly rectifying currents showed a small time-dependent inactivation at depolarizing potentials ⱖ60 mV in cultured PCT and CCT cells and ⱖ40 mV in cultured DCT cells. In most cases, the time course of this inactivation could be well fitted with a single exponential irrespective of the recording time. When the cells were reexposed to the hyperosmotic solution, the currents returned to the control level within 2–3 min (Fig. 3B). In the three

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cultured segments, the currents induced by hypotonicity were strongly blocked by 1 mM DIDS (Fig. 3B). In cftr⫺/⫺ mice, hypotonic shock was completely inefficient for increasing Cl⫺ conductance in the three different cultured segments (Fig. 3, A and B). In all nephron segments studied, an absence of response to hypotonic shock was observed in 100% of the recorded cells. This result implicates CFTR in the control of the swelling-activated Cl⫺ conductance in renal epithelium. The results reported above clearly show that Cl⫺ conductances developed in the presence of forskolin, Ca2⫹, or hypotonic shock in DCT cells were roughly similar to those recorded in CCT cells under the same experimental conditions. Therefore, in the following experimental series, no distinction was made between DCT and CCT cells.

Fig. 3. A: characteristics of swellinginduced whole cell Cl⫺ currents in PCT, DCT, and CCT cells in primary culture of cftr⫹/⫹ and cftr⫺/⫺ mice. Membrane voltage was held at ⫺50 mV and stepped to test potential of ⫺100 to ⫹120 mV in 20-mV increments. Whole cell currents were recorded after 4–5 min of extracellular perfusion of a 30% hypotonic solution in the presence of 5 mM EGTA and 5 mM MgATP in pipette solution and 1 mM CaCl2 in extracellular bath. B: effects of 1 mM DIDS and hyperosmotic solution (350 mosmol/kgH2O) on swelling-induced whole cell Cl⫺ currents. Steady-state currents at ⫹100 mV were measured 20 ms after onset of pulse. Values are means ⫾ SE; n, number of monolayers from 4 different mice.

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Cl⫺ Currents in Cultured PCT and DCT Cells From cftr⫺/⫺ Mice Transfected With CFTR cDNA The cftr⫺/⫺ PCT and DCT cells in primary culture were transfected with CD8-CFTR plasmid, which allows visualization of transfected cells using anti-CD8 antibody-coated beads. After 48 h of transfection, whole cell currents of coated cells were recorded and compared with whole cell currents of control unlabeled cells. Figures 4 and 5 illustrate the currents recorded in cftr⫺/⫺-transfected PCT and DCT cells in the presence of 10 ␮M forskolin. As expected, addition of forskolin induced an increase in membrane current ampli-

tudes in PCT (Fig. 4Ab) and DCT (Fig. 5Ab) cells coated with beads only. Figures 4Ae and 5Ae show that the forskolin-activated currents exhibited a linear I-V relation, with an Erev of 0.2 ⫾ 0.1 mV and a conductance of 7.6 ⫾ 0.4 nS for PCT cells (n ⫽ 5 cells) and an Erev of 0.16 ⫾ 0.5 mV and a conductance of 8.3 ⫾ 0.5 nS for DCT cells (n ⫽ 7 cells). Currents in both cell types were insensitive to 1 mM DIDS (Figs. 4Ac and 5Ac) and blocked by 77 ⫾ 3 and 85 ⫾ 2% for PCT and DCT, respectively, when Cl⫺ was replaced by I⫺ (Figs. 4Ad and 5Ad). Moreover, this substitution shifted the Erev toward the more positive value: Erev for I⫺ ⫽ 36.4 ⫾ 8

Fig. 4. Restoration of CFTR currents and swelling-activated Cl⫺ currents by transitory transfection of pIRES-CD8cftr in PCT cells from cftr⫺/⫺ mice. Transfected cells were visualized using anti-CD8 antibody-coated beads. Membrane potential was held at ⫺50 mV and stepped to test potential of ⫺100 to ⫹120 mV in 20-mV increments. A: CFTR currents in cells labeled with anti-CD8-coated beads. a: Control; b: 10 ␮M forskolin in bath solution; c: 10 ␮M forskolin ⫹ 1 mM DIDS; d: 10 ␮M forskolin with extracellular substitution of Cl⫺ by I⫺. e: Average currentvoltage (I-V) relationships measured 200 ms after onset of pulse, obtained from the same cell at rest, during forskolin stimulation alone and after Cl⫺ substitution by I⫺. Values are means ⫾ SE of 5 cells from 3 transfected monolayers. B: swelling-activated Cl⫺ currents recorded after 4–5 min of extracellular perfusion of a 30% hypotonic solution in the presence of 5 mM EGTA and 5 mM MgATP in pipette solution and 1 mM CaCl2 in extracellular bath. a–c: Whole cell currents in cells labeled with anti-CD8-coated beads. d: Average I-V relationships measured 20 ms after onset of pulse, obtained from the same cell at rest (B), during perfusion with hyposmotic solution, and after perfusion with hyperosmotic solution. Values are means ⫾ SE of 4 cells obtained from 3 transfected monolayers.


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Fig. 5. Restoration of CFTR currents and swelling-activated Cl⫺ currents by transitory transfection of pIRES-CD8cftr in DCT cells from cftr⫺/⫺ mice. Transfected cells were visualized using anti-CD8 antibody-coated beads. Membrane potential was held at ⫺50 mV and stepped to test potential of ⫺100 to ⫹120 mV in 20-mV increments. A: CFTR currents in cells labeled with anti-CD8-coated beads. a: Control; b: 10 ␮M forskolin in bath solution; c: 10 ␮M forskolin ⫹ 1 mM DIDS; d: 10 ␮M forskolin with extracellular substitution of Cl⫺ by I⫺. e: Average I-V relations measured 200 ms after onset of pulse, obtained from the same cell at rest, during forskolin stimulation alone and after Cl⫺ substitution by I⫺. Values are means ⫾ SE of 7 cells from 3 transfected monolayers. B and C: swelling-activated Cl⫺ currents recorded after 4–5 min of extracellular perfusion of a 30% hypotonic solution in the presence of 5 mM EGTA and 5 mM MgATP in pipette solution and 1 mM CaCl2 in extracellular bath. Ba, Bb, and Bc: whole cell currents in cells labeled with anti-CD8-coated beads. Bd: average I-V relationships measured 20 ms after onset of pulse, obtained from the same cell at rest (B), during perfusion with hyposmotic solution, and after perfusion with hyperosmotic solution. Values are means ⫾ SE of 4 cells from 3 transfected monolayers. C: whole cell currents in cells not labeled with anti-CD8-coated beads.

and 25.7 ⫾ 9 mV in PCT and DCT, respectively. Overall, these forskolin-sensitive Cl⫺ currents were identical to those measured in cftr⫹/⫹ DCT cells, indicating that transfection with CFTR plasmid could restore the normal CFTR currents in PCT and DCT cftr⫺/⫺ cells. In another experimental series, the effect of a hypotonic shock was studied in cftr⫺/⫺ PCT and DCT cells transfected with the cftr plasmid. In both cell types, after the hypotonic shock, the coated cells developed Cl⫺ currents within 3 min (Figs. 4B and 5B). The initial currents measured 20 ms after the onset of the voltage pulse rectified in the outward direction (Figs. 4Bd and 5Bd). For PCT cells, they reversed at ⫹0.6 ⫾ 0.4 mV (n ⫽ 4 cells), and the total current at ⫹100 mV AJP-Renal Physiol • VOL

was 3.8 times that at ⫺100 mV: 1,555 ⫾ 150 vs. ⫺405 ⫾ 18 pA (n ⫽ 4 cells). For DCT cells, they reversed at ⫹0.9 ⫾ 0.3 mV (n ⫽ 4 cells), and the total current at ⫹100 mV was 2.2 times that at ⫺100 mV: 1,123 ⫾ 155 vs. ⫺508 ⫾ 86 pA (n ⫽ 4 cells). These large outwardly rectifying currents showed time-dependent inactivation at depolarizing step potential ⬎40 mV. Finally, replacement of the hypotonic bath solution by a hypertonic solution inhibited the Cl⫺ currents by 74 ⫾ 3 and 80 ⫾ 4% for PCT and DCT cells, respectively (n ⫽ 4). As expected, the uncoated cells remained insensitive to the hypotonic shock (Fig. 5B). Therefore, transfection of CFTR also restores the swelling-activated Cl⫺ conductance in cftr⫺/⫺ PCT and DCT cells.

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Regulation of the Cl⫺ Conductance Induced by Hypotonic Shock in cftr⫹/⫹ and cftr⫺/⫺ DCT and CCT Cells Role of extracellular Ca2⫹ in the presence of high EGTA concentration in the pipette solution. In cftr⫹/⫹ cells, 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 EGTA without additional Ca2⫹. The effects of extracellular Ca2⫹ on the development of hypotonicity-induced Cl⫺ currents were also tested in cftr⫹/⫹ DCT and CCT cells. When the hypotonic shock was carried out in the absence of bath Ca2⫹, development of the Cl⫺ current was significantly

impaired (Fig. 6A). As previously reported in rabbit distal bright convoluted tubule (DCTb) in primary culture (20), these experiments confirm that extracellular Ca2⫹ was required to activate the swelling-activated Cl⫺ conductance in DCT and CCT cells cultured from cftr⫹/⫹ mice. Using this information, we therefore decided to study the effect of an influx of Ca2⫹ on swelling-activated Cl⫺ conductance in cftr⫺/⫺ DCT and CCT cells. For this purpose, the effects of ionomycin were tested on whole cell Cl⫺ currents recorded in the absence of intracellular free Ca2⫹. Whole cell currents were recorded in the presence of 20 mM EGTA in the pipette solution and 1 mM free Ca2⫹ in the bath (Fig. 6B). In the absence of ionomycin in the bath

Fig. 6. A: effect of extracellular Ca2⫹ on development of hypotonicity-induced Cl⫺ currents in cultured DCT cells from cftr⫹/⫹ mice. Membrane voltage was held at ⫺50 mV and stepped to test potential of ⫺100 to ⫹120 mV in 20-mV increments. Whole cell currents were recorded after 4–5 min of extracellular perfusion of a 30% hypotonic solution in the presence of 5 mM EGTA in pipette solution and in the absence of extracellular Ca2⫹ in bath solution. B: effects of ionomycin on development of Cl⫺ currents in cultured DCT cells from cftr⫺/⫺ mice. Membrane voltage was held at ⫺50 mV and stepped to test potential of ⫺100 to ⫹120 mV in 20-mV increments. Whole cell currents were recorded after 4–5 min of extracellular perfusion of a 30% hypotonic solution in the presence of 20 mM EGTA and 5 mM MgATP in pipette solution and 1 mM CaCl2 in extracellular bath. Whole cell currents were measured during hypotonic shock. a: Control cells; b: 2 ␮M ionomycin; c: 2 min of replacement of extracellular solution with a hypertonic solution; d: 1 mM DIDS. e: Average I-V relationships measured 20 ms after onset of pulse, obtained from the same cell at rest. Values are means ⫾ SE; n, number of cells from 4 monolayers.


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solution, the hypotonic shock remained inefficient for triggering Cl⫺ currents in cftr⫺/⫺ cells (Fig. 6Ba). In contrast, when the hypotonic shock was performed in the presence of 2 ␮M ionomycin, Cl⫺ currents were activated within 5 min (Fig. 6Bb). These currents showed time-dependent inactivation at depolarizing step potentials ⬎60 mV and displayed an outwardly rectified instantaneous I-V plot (Fig. 6Be) with an Erev of ⫹ 1.1 ⫾ 0.3 mV (n ⫽ 7). When the cells were reexposed to the hyperosmotic solution, the currents returned toward control level within 2–3 min (Fig. 6, Bc and Be). Alternatively, addition of DIDS rapidly reduced the Cl⫺ currents (89.7 ⫾ 4% inhibition at ⫹100 mV, n ⫽ 5; Fig. 6Bd). Overall, the ionomycin-induced Cl⫺ currents developed during hypotonicity in DCT and CCT cells from cftr⫺/⫺ mice were quite similar to the swelling-activated Cl⫺ currents measured in cftr⫹/⫹ mice. Very similar results were obtained with PCT cells in primary culture. Briefly, in cftr⫹/⫹ cells, the swellingsensitive Cl⫺ conductance depended on external Ca2⫹, and in cftr⫺/⫺ cells, this conductance could be reactivated by addition of ionomycin to the bath solution (data not shown). Role of extracellular Ca2⫹ in the absence of EGTA in the pipette solution. The experiments described above indicate that Ca2⫹ influx induced by ionomycin could restore the swelling-activated Cl⫺ currents in cftr⫺/⫺ cells. To further analyze this phenomenon, the effect of ionomycin was tested in the absence of EGTA in the pipette solution. Two successive, increased external Ca2⫹ concentrations were applied to the same cftr⫺/⫺ DCT cells. The results are reported in Fig. 7A. Control currents were recorded, and the cells were perfused with a Ca2⫹-free solution containing 2 ␮M ionomycin. After 2 min, raising the Ca2⫹ concentration to 0.1 ␮M induced Cl⫺ currents that were identical to the swelling-activated Cl⫺ currents (Fig. 7Ab). A further new increase in Ca2⫹ concentration to 1 ␮M enhanced the currents (Fig. 7Ac). These currents showed virtually no inactivation during the 400-ms voltage pulse. Currents obtained by subtracting the current recorded at 0.1 ␮M external Ca2⫹ from that recorded at 1 ␮M Ca2⫹ are shown in Fig. 7Ad. The resulting currents exhibited the characteristic profile of the Ca2⫹-sensitive Cl⫺ currents. On the basis of these results, it appears that Ca2⫹ entry is an important step in the development of swelling-activated Cl⫺ conductance. Fluorescence experiments using fura 2-loaded DCT cells were therefore carried out to follow cytosolic Ca2⫹ variations during hypotonic shock. The effect of hypotonic solution on [Ca2⫹]i in DCT cells from cftr⫹/⫹ and cftr⫺/⫺ mice is shown in Fig. 7B. In both types of mice, when the cells were bathed with an isotonic NaCl solution (300 mosmol/kgH2O) containing 1 mM CaCl2, the resting [Ca2⫹]i averaged 30.8 ⫾ 5.2 nM (n ⫽ 20). Swelling the cftr⫹/⫹ DCT cells with hypotonic NaCl solution (200 mosmol/kgH2O) induced a transient increase of [Ca2⫹]i that reached a maximum value of 120.1 ⫾ 25.1 nM and returned close to the control value within 5 min (Fig. AJP-Renal Physiol • VOL

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7B). In contrast, swelling the cftr⫺/⫺ DCT cells did not significantly modify [Ca2⫹]i. Role of extracellular adenosine. We previously demonstrated that stimulation of A1 adenosine receptors could be implicated in the control of swelling-induced Cl⫺ currents in rabbit DCT (20), and experiments were therefore performed to determine the role of adenosine in Cl⫺ permeability of PCT and DCT cells from cftr⫹/⫹ and cftr⫺/⫺ mice. Results of whole cell experiments performed in cftr⫺/⫺ PCT and DCT cells are illustrated in Fig. 8. These results were strictly identical to those obtained with cftr⫹/⫹ PCT and DCT cells. In both types of primary cultures, 10 ␮M adenosine activated an outwardly rectifying Cl⫺ conductance with a time-dependent inactivation at depolarizing potentials and with a maximal effect at 3–4 min (Fig. 8A). Erev of the stimulated current were 3.8 ⫾ 3.7 mV (n ⫽ 5 monolayers) and 0.3 ⫾ 2.9 mV (n ⫽ 4) for PCT and DCT cells, respectively. In the presence of adenosine, the maximal slope conductances reached 19 ⫾ 9 nS (n ⫽ 5) and 11 ⫾ 4 nS (n ⫽ 4) in PCT and DCT cells, respectively. These adenosine-sensitive Cl⫺ currents were decreased in the presence of 1 mM DIDS by 90 and 78% in PCT and DCT cells, respectively. To determine whether the response to adenosine occurred via receptor-mediated mechanisms, we examined the effect of a P1-selective receptor antagonist, 8-cyclopentyl-1,3diproxylzanthine (DPCPX). Treatment of DCT cells with 10 ␮M DPCPX completely inhibited the development of outward Cl⫺ currents first induced by 10 ␮M adenosine in PCT and DCT cells (Fig. 8). The effect of adenosine was concentration dependent. The dose-response curve in cultured DCT cells from cftr⫹/⫹ mice is shown in Fig. 9. The half-maximal effect from this curve occurred at 5.0 ⫻ 10⫺7 M adenosine and the maximal effect at ⬎10⫺5 M. Role of extracellular ATP. In addition to adenosine, it has been postulated that ATP could activate a volumesensitive-like Cl⫺ conductance in immortalized rabbit distal cells (20). To check this possibility in PCT and DCT cells from cftr⫹/⫹ and cftr⫺/⫺ mice, we studied the role of ATP in the control of whole cell Cl⫺ currents in the presence of 5 mM EGTA in the pipette solution. In PCT and DCT monolayers, addition of 10 ␮M ATP to the bath solution induced activation of Cl⫺ currents within 4–5 min. This ATP-activated Cl⫺ current showed time-dependent inactivation at depolarizing step potentials ⬎60 mV (Fig. 10, A and B) and displayed an outwardly rectified instantaneous I-V plot (data not given) with Erev close to 0 mV. DIDS (1 mM) strongly decreased ATP-activated currents in both types of monolayers. Overall, these currents were quite similar to those induced by adenosine. Moreover, the effect of ATP was completely blocked by 10 ␮M DPCPX, indicating that the action was triggered via P1, rather than P2, receptors. Such results suggested that stimulation of Cl⫺ currents in the presence of ATP was most probably due to an action of adenosine generated by degradation of ATP. Experiments were therefore carried out to check this hypothesis. For this

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Fig. 7. A: effect of extracellular Ca2⫹ concentration ([Ca2⫹]ext) on development of Cl⫺ currents in cultured DCT cells from cftr⫹/⫹ mice. Whole cell currents were recorded in the absence of EGTA in pipette solution. a: Control; b: 2 min of extracellular perfusion of 0.1 ␮M Ca2⫹ in the presence of ionomycin; c: 2 min of extracellular perfusion of 1 ␮M Ca2⫹ in the presence of ionomycin; d: currents obtained by subtraction of c from b using pCLAMPFIT 6.0 software. e: Average I-V relationships measured 20 ms after onset of pulse, obtained from the same cell at rest, during perfusion of ionomycin in the presence of different Ca2⫹ concentrations. Values are means ⫾ SE of 6 cells obtained from 6 monolayers. B: effect of hypotonic shock on intracellular free Ca2⫹ concentration ([Ca2⫹]i) in fura 2-loaded DCT cells from cftr⫹/⫹ and cftr⫺/⫺ mice. Fura 2 fluorescence was monitored and converted to [Ca2⫹]i as described in MATERIALS AND METHODS. Hypotonic shock was induced by perfusion of a hypotonic NaCl solution (200 mosmol/kgH2O). Values are means ⫾ SE of 20 random cells from 3 monolayers.

purpose, DCT cells from cftr⫹/⫹ mice were subjected to a hypotonic shock in the presence of the selective ectoATPase inhibitor ARL-67156. ARL-67156 (100 ␮M) completely blocked the swelling-activated Cl⫺ currents (Fig. 10C). Adenosine (10 ␮M) restored a swellingactivated Cl⫺ conductance, which displayed an outwardly rectified instantaneous I-V plot with Erev close to 0 mV (Fig. 10Cd) and was strongly inhibited by DIDS (Fig. 10Cb). AJP-Renal Physiol • VOL

RVD in PCT and DCT Cells From cftr⫹/⫹ and cftr⫺/⫺ Mice To confirm the role of swelling-activated Cl⫺ currents in cell volume regulation, we measured relative cell volume in monolayers by fluorescence-video microscopy in PCT and DCT cells from cftr⫹/⫹ and cftr⫺/⫺ mice. As expected, PCT and DCT cells swelled in response to a hypotonic shock (Fig. 11). This cell

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Fig. 8. Effects of adenosine on development of Cl⫺ currents in cultured PCT (A) and DCT (B) cells from cftr⫺/⫺ mice. Membrane voltage was held at ⫺50 mV and stepped to test potential of ⫺100 to ⫹120 mV in 20-mV increments. Whole cell currents were recorded after 4–5 min of extracellular perfusion of a 30% hypotonic solution in the presence of 5 mM EGTA and 5 mM MgATP in pipette solution and 1 mM CaCl2 ⫹ 10 ␮M adenosine in extracellular bath. DIDS (1 mM) was perfused after development of Cl⫺ currents. Cells were treated with antagonist 8-cyclopentyl-1,3-diproxylxanthine (DPCPX) before exposure to adenosine.

swelling was followed by an RVD in PCT and DCT cells from cftr⫹/⫹ mice. At 2 min after the hypotonic shock, the relative cell volume reached 130 ⫾ 2% (n ⫽ 3 monolayers) and 120 ⫾ 1% (n ⫽ 8 monolayers) of the initial volume in PCT and DCT cells, respectively. PCT cells returned to 105 ⫾ 1% of their original volume within 4 min (Fig. 11A), whereas DCT cells recovered 104 ⫾ 1% of their volume within only 2 min (Fig. 11B). The RVD phenomenon in both cell types was inhibited in the presence of 100 ␮M NPPB (Fig. 11). Contrary to observations in cells from cftr⫹/⫹ mice, the RVD mechanism was completely impaired in PCT and DCT cells from cftr⫺/⫺ mice. When perfused with the hypotonic solutions, these cells never returned to their initial volume. Addition of 10 ␮M adenosine during hypotonic

Fig. 9. Adenosine dose-response curve. Membrane voltage was held at ⫺50 mV and stepped to test potential of ⫺100 to ⫹120 mV in 20-mV increments. Whole cell currents were recorded after 3 min of extracellular perfusion of adenosine at different concentrations in isotonic N-methyl-D-glucamine solutions in the presence of 5 mM EGTA and 5 mM MgATP in pipette solution and 1 mM CaCl2 in extracellular bath. Values at ⫹100 mV were converted to percent activation. Values are means ⫾ SE of 6 cells from 3 monolayers. AJP-Renal Physiol • VOL

shock restored RVD in DCT cells from cftr⫺/⫺ mice (Fig. 11B). DISCUSSION

The aim of the present study was to investigate the putative role of CFTR in the control of Cl⫺ conductances along the different segments of the mouse nephron. Using the patch-clamp technique to measure whole cell conductance, we analyzed three distinct types of Cl⫺ currents in primary cultures of PCT, DCT, and CCT segments obtained by microdissection of kidney cortex from wild-type cftr⫹/⫹ and cftr⫺/⫺ mice. These Cl⫺ conductances consisted of forskolin-activated, volumesensitive, and Ca2⫹-activated Cl⫺ currents. In the first series of experiments, the effect of forskolin on Cl⫺ conductance was tested in PCT, DCT, and CCT cells. For this purpose, swelling-activated currents were blocked by exposing the cells to a hyperosmotic solution, and Ca2⫹-activated conductances were impaired by the use of high EGTA concentrations in the pipette solution. In cftr⫹/⫹ mice, external application of forskolin activated a linear Cl⫺ current in DCT and CCT, but not PCT, cells. The halide selectivity was consistent with low relative I⫺ permeability and with an inhibitory effect of I⫺. Moreover, this forskolinstimulated conductance was blocked by NPPB but was quite insensitive to DIDS. These characteristics are very similar to those reported previously in rabbit distal bright convoluted tubule (DCTb) in primary culture (21). In contrast, in cultured DCT and CCT cells from cftr⫺/⫺ mice, addition of forskolin remained completely inefficient for increasing Cl⫺ conductances. Taken together, the results obtained in primary cultures from cftr⫹/⫹ and cftr⫺/⫺ mice clearly demonstrate that, at least in DCT and CCT cells, the activity of forskolin-activated Cl⫺ channels is consistent with CFTR. In other words, as we concluded in a previous study (21), the channel involved in the Cl⫺ currents activated by forskolin in DCT and CCT is the smallconductance CFTR Cl⫺ channel.

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Fig. 10. Effects of ATP on development of Cl⫺ currents in cultured PCT (A) and DCT (B) cells from cftr⫺/⫺ mice. Membrane voltage was held at ⫺50 mV and stepped to test potential of ⫺100 to ⫹120 mV in 20-mV increments. Whole cell currents were recorded after 4–5 min of extracellular perfusion of a 30% hypotonic solution in the presence of 5 mM EGTA and 5 mM MgATP in pipette solution and 1 mM CaCl2 ⫹ 10 ␮M ATP in extracellular bath. DIDS (1 mM) was perfused after development of Cl⫺ currents. Cells were treated with DPCPX before exposure to adenosine. Values are means ⫾ SE; n, number of cells from 3 monolayers. C: effects of ecto-ATPase antagonist ARL-67156 on swelling-activated Cl⫺ currents in DCT cells from cftr⫹/⫹ mice. Whole cell currents were recorded after 4–5 min of extracellular perfusion of a hypotonic solution in the presence of 100 ␮M ARL-67156 (a), 10 ␮M adenosine (b), or 1 mM DIDS (c). d: Average I-V relationships measured 20 ms after onset of pulse obtained from the same cell at rest. Values are means ⫾ SE of 5 cells from 3 monolayers.

Interestingly, application of forskolin did not stimulate any Cl⫺ current in primary cultures of PCT cells from cftr⫹/⫹ mice. Such an observation was reported in primary culture of rabbit PCT, in which no CFTR expression and no forskolin-activated Cl⫺ currents were detected in the apical membrane, despite the presence of CFTR mRNA (21). The presence of CFTR in the mammalian kidney is now well documented (2, 15, 16), but the absence of detectable renal disease in CF patients led several authors to postulate that an increase of another type of Cl⫺ channel might compensate for the lack of cAMPactivated Cl⫺ channels in renal tissue (6, 11). On the other hand, the cftr⫺/⫺ mice used in the present study did not present significant pulmonary disease. Moreover, in these mice, it has been shown that the Ca2⫹activated Cl⫺ channels could be candidates for compenAJP-Renal Physiol • VOL

sation of the missing CFTR Cl⫺ channels (6). To determine whether this possibility could arise in the renal epithelium, we studied the Ca2⫹-activated conductance in PCT, DCT, and CCT cells. As expected, in cftr⫹/⫹ mice, extracellular application of ionomycin rapidly activated currents in all types of monolayers. This Ca2⫹-sensitive conductance was similar to that previously described in rabbit PCT and DCTb cells under identical experimental conditions (21, 22). In cftr⫺/⫺ mice, the increase of Cl⫺ conductance triggered by ionomycin was strikingly identical to that observed in wild-type mice, eliminating the hypothesis that Ca2⫹-activated conductance could substitute for CFTR Cl⫺ conductance in renal epithelium. In cftr⫹/⫹ mice, cultured PCT, DCT, and CCT cells developed a volume-sensitive Cl⫺ current when exposed to a hypotonic shock. The biophysical and phar-

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Fig. 11. Effects of hypotonic shock on cell volume in PCT and DCT cells from cftr⫹/⫹ and cftr⫺/⫺ mice. Cultures were loaded with 2 ␮M fura 2 and rinsed in an isotonic NaCl solution (300 mosmol/kgH2O) for 3 min. A hypotonic shock was induced by reducing osmolarity of NaCl solution to 200 mosmol/kgH2O. Images were recorded every 15 s. After analysis, relative volume change as percentage of initial volume was plotted against time. A: regulatory volume decrease (RVD) in 3 monolayers from PCT cells (25 random cells each) in the presence or absence of 100 ␮M 5-nitro-2-(3-phenylpropylamino)benzoic acid (NPPB). B: RVD in 10 monolayers from DCT cells (25 random cells each) from cftr⫺/⫺ mice and 8 monolayers (25 random cells each) from DCT cells from cftr⫹/⫹ mice in the presence or absence of 100 ␮M NPPB or adenosine.

macological characteristics of this Cl⫺ conductance show strong similarities to the properties of swellingactivated Cl⫺ currents described in many other epithelial cells, including rabbit PCT and DCTb in primary AJP-Renal Physiol • VOL

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culture (7, 25). Null mutation of the cftr gene strongly impaired the swelling-activated Cl⫺ currents in the three different nephron segments. Moreover, cftr⫺/⫺ DCT or CCT cells transfected with cftr cDNA displayed complete restoration of cAMP-dependent and swellingactivated Cl⫺ currents. Transfection of PCT cells with cftr cDNA also restored both conductances. These observations indicate that PCT cells have maintained their ability to insert exogenous CFTR into the apical membrane. Therefore, the lack of forskolin-induced Cl⫺ conductance in wild-type PCT cells is probably due to a difference in the protein function, rather than a modification of the intracellular trafficking leading to protein retention in intracellular membranes. It is well established that the swelling-activated Cl⫺ channels participate in the RVD phenomenon, which is induced by exposure of cells to hyposmotic solutions. In the present study, to determine whether cultured PCT, DCT, and CCT cells develop RVD after a hypotonic shock, we used a simple fluorescence method for studying relative cell volume variations (29). The findings indicate that cultured cells from cftr⫹/⫹ mice are sensitive to osmolarity changes in the bathing medium and that they are capable of RVD after hypotonic shock. RVD was also examined in cultured cells from cftr⫺/⫺ mice. These cells exhibited a defective volume regulation after a hyposmotic shock. This observation confirms the results in the literature (31) and indicates that CFTR could play a role in the RVD of epithelia. Obviously, this defective RVD is due to the fact that the hypotonic shock is completely inefficient for increasing Cl⫺ conductances. The intervention of CFTR in the control of swelling-activated Cl⫺ conductances has been proposed by Chan et al. (5), who demonstrated, in the human colonic cell line T84, that an antibody against CFTR inhibited the cAMP- as well as the swelling-induced whole cell Cl⫺ conductances but did not affect the Ca2⫹-activated Cl⫺ channel. In previous studies, we found that development of Cl⫺ conductance after a hypotonic shock in rabbit DCT cells was related to an influx of external Ca2⫹ through Ca2⫹ channels (21, 22). Such a hypothesis could also apply to the data obtained in DCT cells from cftr⫹/⫹ mice, because removal of external Ca2⫹ just before the hypotonic shock completely impaired the increase in Cl⫺ current, suggesting that Ca2⫹ influx could participate in activation of the Cl⫺ channels in mouse kidney. It is now proposed that CFTR could regulate other ion channel proteins (23) and could also be implicated in different cell functions such as apoptosis (1, 8, 13) or cytosolic Ca2⫹ regulation (19). Moreover, a recent study by Braunstein et al. (3) clearly demonstrated that CFTR participates in cell volume regulation via control of ATP release. Taken together, these observations led us to propose the hypothesis that the absence of swellingactivated Cl⫺ conductance in DCT cells from cftr⫺/⫺ could be related to an alteration of the Ca2⫹ entry. This hypothesis is strengthened by two main results: 1) The hypotonic shock induced an increase of [Ca2]i in cftr⫹/⫹ DCT cells, but not in cftr⫺/⫺ cells. 2) In cftr⫺/⫺ cells, addition of ionomycin in the presence of

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high intracellular EGTA concentration restored the ability of the cells to respond to the hypotonic shock by increasing swelling-sensitive Cl⫺ conductance. It remains to be shown how intracellular Ca2⫹ can increase in the presence of a high EGTA concentration. The observations of Evans and Marty (6a) 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 in the presence of EGTA. In accordance with the model proposed by Braunstein et al. (3), the defect in cell volume regulation that we observed in cftr⫺/⫺ renal cells could be due to a defect in the ATP release pathway. We have proposed (20) that hypotonic shock stimulates ATP release from rabbit DCT cells. A1 receptors are then activated by adenosine generated by the degradation of ATP by membrane ectoenzymes, and this stimulation of A1 receptors induces an influx of extracellular Ca2⫹. Finally, this Ca2⫹ influx activates the Cl⫺ channel. The observation that adenosine restores swelling-activated Cl⫺ conductance and RVD in cftr⫺/⫺ cells confirms that adenosine is a mediator of RVD, at least in renal epithelium. Therefore, in cftr⫺/⫺ cells, there is no volume-sensitive ATP release, and the cascade of events that triggers the final increase in Cl⫺ conductance no longer occurs. The present results confirm that autocrine ATP release is probably an essential step in the cell volume regulation phenomenon. However, it could be questioned why adenosine or ATP did not activate a current consistent with a Ca2⫹-activated Cl⫺ current during a hypotonic shock. We have demonstrated that swellingactivated and Ca2⫹-sensitive Cl⫺ currents could be additive but also that their thresholds of activation by Ca2⫹ were quite different. Thus the former was activated at 0.1 ␮M Ca2⫹, whereas the latter was activated at 1 ␮M Ca2⫹. The cytosolic Ca2⫹ concentration induced by hypotonic shock never exceeded 0.15 ␮M. This small increase is consistent with the fact that CFTR control of ATP release during swelling involved probably low ATP concentration and, consequently, low adenosine production. In the present study, the effect of adenosine in increasing swelling Cl⫺ currents was concentration dependent, with a half-maximal effect at 5.0 ⫻ 10⫺7 M. As previously reported in rabbit DCT cells, this adenosine concentration raised cell Ca2⫹ to 0.11 nM, which was sufficient to trigger swelling-activated Cl⫺ currents but too low to induce Ca2⫹dependent Cl⫺ currents. The RVD process involves Cl⫺ and K⫹ efflux. Previous data indicate that impairment of RVD in jejunal crypts of cftr⫺/⫺ mice was due to a defective K⫹ channel (31, 32). However, the nature of the K⫹ channels stimulated during hypotonic shock remains very uncertain and appears to depend on the tissue under investigation (32). Therefore, in the companion article (1a), we investigate the K⫹ conductances along the different nephron segments of cftr⫺/⫺ and cftr⫹/⫹ mice. AJP-Renal Physiol • VOL

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