We have previously shown that despite the presence of mRNA encoding CFTR, renal proximal cells do not exhibit cAMP-sensitive Cl− conductance (Rubera I, Tauc M, Bidet M, Poujeol C, Cuiller B, Watrin A, Touret N, Poujeol P. Am J Physiol Renal Physiol 275: F651–F663, 1998). Nevertheless, in these cells, CFTR plays a crucial role in the control of the volume-sensitive outwardly rectifying (VSOR) activated Cl− currents during hypotonic shock. The aim of this study was to determine the role of CFTR in the regulation of apoptosis volume decrease (AVD) and the apoptosis phenomenon. For this purpose, renal cells were immortalized from primary cultures of proximal convoluted tubules from cftr+/+ and cftr−/− mice. Apoptosis was induced by staurosporine (STS; 1 μM). Cell volume, Cl− conductance, caspase-3 activity, intracellular level of reactive oxygen species (ROS), and glutathione content (GSH/GSSG) were monitored during AVD. In cftr+/+ cells, AVD and caspase-3 activation were strongly impaired by conventional Cl− channel blockers and by a specific CFTR inhibitor (CFTRinh-172; 5 μM). STS induced activation of CFTR conductance within 15 min, which was progressively replaced by VSOR Cl− currents after 60 min of exposure. In parallel, STS induced an increase in ROS content in the time course of VSOR Cl− current activation. This increase was impaired by CFTRinh-172 and was not observed in cftr−/− cells. Furthermore, the intracellular GSH/GSSG content decreased during STS exposure in cftr+/+ cells only. In conclusion, CFTR could play a key role in the cascade of events leading to apoptosis. This role probably involves control of the intracellular ROS balance by some CFTR-dependent modulation of GSH concentration.
cftr is known not only as a cAMP-dependent Cl− channel in the epithelium but also as a modulator of other ion channels and transporters. Notably, in renal cells, regulatory volume decrease (RVD) after hypotonic challenge is dependent on CFTR. Thus, in mouse proximal convoluted tubules (PCT), we previously demonstrated that, during hypotonic shock, CFTR modulated the swelling-activated Cl− current by controlling a cascade of events involving ATP release, adenosine production, and Ca2+ entry (3). Concomitantly, the decrease in tonicity activated K+ conductance through TASK2 channels (2, 6). Interestingly, it is now well established that ion channels responsible for RVD are also involved in the induction of apoptotic volume decrease (AVD), which is an early event in apoptotic cell death. Thus, by an examination of various cell lines exposed to different apoptotic stimuli, it has been demonstrated that the separate pharmacological blockade of volume-regulatory K+ and Cl− channels prevents AVD and subsequent apoptosis. In the mouse proximal tubule, we previously found that TASK2 K+ channels are also involved in the control of cell volume during staurosporine (STS)-induced AVD. Regarding the Cl− channel recorded during AVD, it exhibits volume sensitivity, and phenotypical properties of the VSOR Cl− channel and literature data converge toward the conclusion that this Cl− channel is probably ubiquitously expressed in animal cells. The assumption that the Cl− and the K+ channels involved in AVD in proximal tubules are the same as those implicated in RVD logically led us to study the putative role of CFTR in the control of AVD. This study has been further supported by the results described in different studies indicating that CFTR could be involved in the apoptosis phenomenon (4, 14, 17, 29). On the other hand, it has already been reported that cardiomyocyte apoptosis involves VSOR Cl− channel activity and intracellular reactive oxygen species (ROS) production (40). Furthermore, an interplay among AVD, ROS production, and TASK2 channels seems to be essential for permitting the cells to enter into apoptosis during STS exposure (22). Thus it was interesting to address the question of CFTR involvement in STS-induced AVD and whether apoptosis is related to the control of the VSOR channel by ROS production. Therefore, our study was conducted in proximal tubule cell lines originating from cftr+/+ and cftr−/− mice. As expected, we observed that STS-mediated AVD was caused by an increase in VSOR resulting from a CFTR-dependent ROS increase. To explain this dependency, we then investigated the handling of GSH during STS-induced AVD in cftr+/+ and cftr−/− proximal cell lines. GSH has a ubiquitous distribution within most tissue and plays a major role in protecting cells from oxidative stress, mostly by scavenging intracellular ROS (35). Moreover, depletion of GSH is also associated with the initiation of apoptosis (24, 25), probably because the lowering of GSH increases ROS production. There is evidence that the maintenance of the GSH pool is critical for maintaining normal cell function, especially in the proximal convoluted tubule cells, which are potentially exposed to a high concentration of nephrotoxic agents and/or their reactive intermediates. To ensure GSH homeostasis, the proximal tubule exhibited several processes involved in GSH metabolism and transport (23). Of the different putative transporters, the multidrug resistance proteins (MRP) could play a role in efflux of GSH across the brush-border membrane (16, 24). CFTR belongs to the same transporter family (ABC transporters) and has been shown to mediate GSH flux in kidney cell lines (26) as well as in lung tissue (18). In agreement with these results, we subsequently demonstrated that the role of CFTR in STS-induced AVD and apoptosis involves control of the intracellular ROS balance by CFTR-dependent modulation of GSH concentration. To complete our study, it was necessary to precisely identify the mechanism by which STS activates CFTR. Given that, we have demonstrated that in cultured proximal cells, CFTR Cl− conductance was not activated via the cAMP-PKA pathway (36). We have therefore investigated whether a MAP kinase pathway could be involved in the rapid development of CFTR Cl− conductance during acute STS application. Finally, our data suggest that the induction of ERK1/2 phosphorylation by STS was a prerequisite effect for activating CFTR and for initiating the cascade of events that ultimately triggers apoptosis. This CFTR-dependent cascade included GSH efflux, ROS production, activation of VSOR conductance, AVD, and an increase in caspase 3 activity.
Primary Cell Cultures
The primary cell culture technique has been described in detail in previous studies (6). Briefly,. 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 the 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 segments of PCT located 1–1.5 mm after the glomerulus were microdissected by hand in this buffer under binoculars using stainless steel needles. After they were rinsed in the 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 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 removed 4 days after seeding and then every 2 days.
Transformation of Primary Cultures with pSV3 Neo and Culture Protocol
Ten-day-old primary cultures of S1 and S2 segments of proximal tubules from cftr+/+ mice were transfected with pSV3 neo using lipofectin (Invitrogen). After 48 h, selection of the clones was performed by the addition of 500 μg/ml G418. Culture medium (DMEM-F12, Sigma, St. Quentin Fallavier, France) containing G418, 15 mM NaHCO3, 20 mM HEPES (pH 7.4), and 1% FCS was changed every day (20) Resistant clones were isolated, subcultured, and used after 10 trypsinization steps. Immortalized proximal cftr+/+ and cftr−/− cell lines were grown on collagen-coated supports (35-mm petri dishes) in a 5% CO2 atmosphere at 37°C in the culture medium described above.
Identification of CFTR mRNA in PCT Cell Lines
Total RNA was prepared using a RNeasy Mini Kit (Qiagen) from immortalized proximal tubule cells. cDNA synthesis was carried out using Superscript First-Strand Synthesis System for RT-PCR (Invitrogen) according to the manufacturer's instructions. cDNA specific for CFTR was amplified using sequence-specific oligonucleotide primers giving rise to product of 550 bp (CFTR sense: 5′-GGATCAGGAAAGACATCACTCCTG -3′; CFTR antisense: 5′-GAACTGAAGCTCGGACGTAGACT-3′). GAPDH expression was evaluated as an internal control (GAPDH sense: 5′-ACCACAGTCCATGCCATCAC-3′; GAPDH antisense: 5′-TCCACCACCCTGTTGCTGTA-3′).
Immunofluorescence Staining of PCT Cell Lines
To detect CFTR localization, PCT cell lines grown on glass coverslips were washed in PBS, fixed in paraformaldehyde (3.7%) at room temperature for 15 min, and blocked using PBS NH4Cl5 mM for 15 min. Cells were then permeabilized with PBS, 0.3% Triton X-100 for 5 min, blocked 20 min with saturation buffer (0.03% Triton X-100, 2.5% goat serum, 1% bovine serum albumin, and 0.2% gelatin in PBS). Using this saturation buffer throughout the experiment, the cells were incubated overnight with anti-CFTR antibodies at 4°C (M3A7, UBI 05-583, 4 μg/ml), washed with PBS 0.3%, and revealed with fluorescein isothiocyanate-conjugated anti-mouse (1:500, Molecular Probes) antibodies and phalloidin-TRITC (1:1,000, Sigma) for 1 h at room temperature. Pictures were taken with a ×63-magnification lens using a confocal microscope (Zeiss). The cells incubated with a control rabbit IgG showed no staining.
Cftr−/− PCT Cell Lines Stably Transfected with CFTR
PCT cell lines stably expressing CFTR were generated by DAC 30-mediated transfection with constructs containing the full-length cDNA encoding human wild-type CFTR. These constructs were obtained by transferring the 4.5-kb CFTR cDNA excised from the pTG5960 plasmid (Transgène) in the polycloning site of the eukaryote expression vector pCB6. pCB6 is a 6.2-kb vector that possesses the hygromycine resistance gene and a cloning site that is under the control of the cytomegalovirus promoter. The resulting pCB6 CFTR plasmid was transfected into cftr−/− PCT cells using DAC 30 according to the protocol provided by the manufacturer (Life Technologies, Cergy Pontoise, France). CFTR transfectants were isolated by growth in media containing hygromycine.
Apoptosis was induced by either STS (1 μM) or concomitant addition of TNF-α (0.5 ng/ml) and cycloheximide (CHX; 1 μg/ml) in proximal cftr+/+ and cftr−/− cell lines. Cells were maintained in serum- and growth factor-free culture medium (DMEM-F12, Sigma) in a 5% CO2 atmosphere at 37°C.
Morphological Counting of Apoptotic Cells
STS-induced apoptosis was studied in cftr+/+ and cftr−/− cell lines. Cells were grown in 35-mm petri dishes. After the appropriate time of incubation with the apoptosis inductor (STS or TNF-α), living cells were carefully washed with fresh culture medium and incubated for 10 min in the presence of Hoechst-33258 (50 μM) and propidium iodide (1 μM). Digital micrographs were successively taken at 455 nm for Hoechst-33258 and 585 nm for propidium iodide. Afterward, the cell preparation was washed and stained with orcein solution [250 mg orcein, 2 ml ethanol (70%), 150 μl HCl (12 N)]. Micrographs of orcein-stained cells were then taken. Thus, in a given culture, the same zone was visualized after individual staining with Hoechst-33258, propidium iodide, and orcein. Apoptotic cells were counted by comparing the three stains. A cell was considered apoptotic only if the nucleus was not stained by propidium iodide and did present chromatin condensation with visible apoptotic bodies. The counts of apoptotic nuclei were performed directly on the digital micrographs. Between 100 and 200 cells were scored by 3 different observers who were blinded to the culture conditions. The numbers of cells with DNA condensation and with staining by propidium iodide were expressed as the percentage of total cells.
Cell Volume Measurements
Cell volume was measured by an electronic sizing technique using a CASY 1 cell counter (Schärfe System) (22). Briefly, proximal cftr+/+ and cftr−/− cells that were exposed to STS treatment were rapidly trypsinized (1×) and cell volume measurements were performed just after the cells were suspended in Casyton solution (NaCl isotonic solution).
Caspase-3 Activity Measurements
Caspase-3 activity was measured using colorimetric assays (CaspACE assay system, colorimetric, Promega). The activity was assayed in triplicate or quadruplicate on protein extracts obtained after lysis of transformed proximal cftr+/+ and cftr−/− cell lines. As indicated by the supplier, the involvement of other related proteases was excluded by observing the difference between color intensity in the absence and presence of a specific caspase-3 inhibitor (Z-VAD). The absorbance was measured at 405 nm using an Automated Microplate Reader ELX-800, and the given values represent the quantity of colored pNA (μmol·min−1·μg protein−1) measured for each experimental condition (Bio-Tek Instruments).
Whole-cell currents were studied using cultured proximal cftr+/+ and cftr−/− cells grown on 35-mm petri dishes maintained at 37°C for the duration of the experiments. The ruptured whole-cell configuration of the patch-clamp technique was used. Patch pipettes (2- to 4-MΩ resistance) were made from borosilicate capillary tubes (1.5-mm OD, 1.1-mm ID, Fisher Manufacturing) using a two-stage vertical puller (model PP 830, Narishige, Tokyo, Japan). 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 VP 500, Biologic). The membrane was ruptured by additional suction to achieve the conventional whole-cell configuration. Settings available on the amplifier were used to compensate for cell capacitance. The series resistances were not compensated, but experiments in which the series resistance was >20 MΩ were discarded. The offset potentials between both electrodes were zeroed before sealing, and the liquid junction potential was measured experimentally before each experiment and corrected accordingly. Solutions were perfused in the extracellular bath using a four-channel glass pipette, with the tip placed as close as possible to the clamped cell. Voltage-clamp commands, data acquisition, and data analysis were controlled via the VP 500 amplifier connected to a computer. The whole-cell currents resulting from voltage stimuli were sampled at 2.5 kHz and filtered at 1 kHz. Cells were held at −50 mV, and 400-ms pulses from −100 to +120 mV were applied in 20-mV increments.
The pipette solution contained (in mM) 140 NMDG-Cl, 10 HEPES (pH 7.4 adjusted with 1 N KOH), 5 MgATP, and 5 EGTA (osmolality = 300 mosmol/kgH2O). To avoid spontaneous activation of volume-sensitive Cl− currents, the bath solution was slightly hyperosmotic and contained (in mM) 140 NMDGCl, 5 glucose, 1 CaCl2, 1 HEPES (pH 7.4 adjusted with 1 N HCl), and 40 mM mannitol (osmolality = 330–340 mosmol/kgH2O).
Cell lysates were prepared in ice-cold lysis buffer containing (in mM) 50 HEPES, pH 7.4, 150 NaCl, 100 NaF, 10 EDTA, 10 Na4P2O7, and 2 Na3VO4 with 1% Triton X-100 and supplemented with protease inhibitors [aprotinin (2 μg/ml), leupeptin (10 μM), and 4-(2-aminoethyl)benzenesulfonyl fluoride (1 mM)]. Equivalent amounts of protein (20 μg) were separated by SDS-PAGE on 10% acrylamide gels. Proteins were electrically transferred to Hybond-C Extra membranes (Amersham) and stained with amido black to verify an even transfer, and the blots were then incubated in blocking buffer [1× Tris-buffered saline (TBS), 0.1% Tween 20, 5% nonfat dry milk] for 1 h at room temperature. The membranes were washed in washing buffer (1×TBS, 0.1% Tween 20) three times, for 5 min per wash, and probed first with the primary antibody [anti-phosphospecific ERK1/2 (dilution 1:10,000, Sigma)] and then with the horseradish peroxidase-conjugated secondary antibody for 1 h in 1×TBS, 0.1% Tween 20, 1% nonfat dry milk. After incubation, membranes were washed three times for 10 min in washing buffer. Proteins were then visualized by the Amersham ECL system. After stripping, equal loadings of proteins were verified by reprobing the blots with total anti-ERK1/2 (1:10,000, Sigma). Band intensity was quantified using PCBas software.
Measurement of ROS
Levels of cellular oxidative stress were measured using the fluorescent probe [5-(and -6)-carboxy-2′,7′-dichlorodihydrofluorescein diacetate] (carboxy-H2DCFDA). Carboxy-H2DCFDA is a cell-permeable indicator for ROS that becomes spontaneously fluorescent when the acetate groups are removed by intracellular esterases and cell oxidation. This probe is trapped mainly in the cytoplasm and is oxidized by several ROS, most notably hydrogen peroxide. Briefly, proximal cftr+/+ and cftr−/− cells were incubated in petri dishes for 30 min in the presence of carboxy-H2DCFDA (10 μM) and gently washed in serum-free culture medium. Cells were then rapidly trypsinized (1×) and incubated in the absence or presence of STS (1 μM) or N-acetylcysteine (NAC; 10 mM) or both substances. Variations of fluorescence of the cell suspension were measured every 2 min using a Genius Spectrofluorimeter (SAFAS, Monaco) at 538 nm.
Measurements of Intracellular GSH-GSSG Content
Variations of intracellular GSH-GSSG content in cftr+/+ and cftr−/− cell lines were quantified using an enzymatic kit (glutathione assay kit, Sigma) (15). Cells were incubated with STS (1 μM) for 10, 20, or 30 min at 37°C and rapidly lysed (liquid nitrogen treatment). The cell lysate was analyzed for GSH content (GSH+GSSG) by quantifying the formation of the 5 thio-bis (2-nitrobenzoic) acid (TNB) amount obtained by reduction of DTNB in the presence of glutathione reductase. The quantity of TNB was assayed at 412 nm using Automated Microplate Reader ELX-800 and reflected the total amount of GSH/GSSG content of the sample.
Flux of GSH and Conjugates
The flux of different forms of GSH was monitored by image analysis with 5-chloro-methyfluorescein diacetate (CMFDA). This fluorescent probe measures changes in the level of intracellular GSH and derivates. Cell tracker CMFDA freely diffuses into the cell, where cytosolic esterases cleave the acetate groups and release the fluorescent product, and has been shown to be suitable for long-term cell labeling. The excitation wavelength for the CMFDA fluorochrome was 490 nm. The emission aperture for fluorescence detection was 510–540 nm for CMFDA. Briefly, proximal cell lines grown in 35-mm petri dishes (1 wk old) were incubated in the presence of 1 μM CMFDA (final concentration) at 37°C for 2 min in a humidified atmosphere of 5% CO2-95% air. Cells were incubated in serum- and growth factor-free culture medium (DMEM-F12). After incubation with the fluorochrome, the staining solution was replaced with fresh prewarmed cell culture medium and cells were analyzed. The mean of GSH and conjugate flux was obtained by analysis of 15–25 cells in each culture.
STS and CFTRinh-172 (Sigma-Aldrich) were prepared in DMSO and used at final concentrations of 1 and 5 μM, respectively. TNF-α (Sigma-Aldrich) was prepared in distilled water and used at a final concentration of 0.5 ng/ml. 5-Nitro-2-(3-phenylpropylamino)-benzoic acid (NPPB; Calbiochem) was prepared at 100 mM in DMSO. DIDS was directly dissolved in the medium at a final concentration of 1 mM. The quantity of DMSO added to the incubation solutions never exceeded 0.1%. Control experiments were performed by incubating the cells with 0.1% DMSO only. The fluorescent probes carboxy-H2DCFDA (Molecular Probes) and CMFDA were used at 10 and 1 μM, respectively. NAC (10 mM) and l-buthionine-S,R-sulfoximine (BSO; 1 mM) were obtained from Sigma-Aldrich (France).
CFTR Expression in cftr+/+ and cftr−/− PCT Cell Lines
Identification of transcripts encoding the mouse CFTR sequence by RT-PCR.
PCT cell mRNA was reverse transcribed and amplified by PCR using primers amplying a product of 550 bp encoding for a fragment located between exon 10 and exon 13 of the mouse CFTR. The analysis of the RT-PCR products by electrophoresis on agarose gels stained with ethidium bromide revealed one product of 550 bp in the cftr+/+ cell line only (Fig. 1A). Identical analysis without prior reverse transcriptase of the RNA sample revealed no amplification of any product.
Labeling of cftr+/+ cell line with M3A7.
The indirect immunofluorescence technique was used to localize CFTR in the cftr+/+ cell line. M3A7 (monoclonal antibody) was used on cells grown on collagen-coated glass coverslips. As shown in Fig. 1B, the antibody revealed CFTR staining at the plasma membrane surface of the cftr+/+ cell line. By contrast, no CFTR staining was observed in the cftr−/− cell line.
Role of CFTR and Volume-Regulated Cl− Channels in STS-Induced Caspase-3 Activation and Apoptosis
First, we compared the activation of caspase-3 after STS (1 μM) exposure in both proximal cftr−/− and cftr+/+ cell lines. As shown in Fig. 2A, STS exposure increased caspase-3 activity within 6 h in the cftr+/+ cell line (n = 7). The Cl− channel inhibitors (DIDS and NPPB) significantly inhibited this activation. Furthermore, the inhibition of caspase-3 activity by the specific inhibitor of CFTR (CFTRinh-172) indicated the involvement of CFTR in the STS-induced caspase-3 activation of these cells. This involvement of CFTR was subsequently confirmed by the use of a cftr−/− cell line in which the STS had no effect. These initial results suggest that both Cl− channels (the DIDS-sensitive channel and the CFTR protein) might be involved in apoptosis induced by STS.
To verify these observations, the apoptosis phenomenon was also assessed on the basis of morphological criteria. cftr+/+ and cftr−/− cell lines were stained with Hoescht-33258 and propidium iodide to selectively distinguish between apoptotic and necrotic cells. Under control conditions, the nuclei of wild-type cells excluded propidium iodide and exhibited a normal morphology, with Hoescht-33258 diffusely labeling the normal chromatin (Fig. 2B). In sharp contrast, after STS exposure (1 μM, 6–8 h), Hoescht-33258 staining revealed that several cells exhibited very intense staining of condensed and fragmented chromatin and were not stained with propidium iodide, indicating preservation of plasma membrane integrity (Fig. 2, C and D). The condensation and fragmentation of DNA clearly show that STS induced apoptosis. These morphological characteristics could also be observed independently with orcein staining: control cells (without STS) did not exhibit chromatin condensation. In contrast, a dense and thin crown of nuclear coloration, typical of chromatin condensation, could be observed after STS exposure (data not shown).
Hoescht-33258 staining on STS-treated cftr−/− cells revealed an absence of condensed and fragmented chromatin compared with cftr+/+ cells (Fig. 2, E and F) By contrast these cells were largely necrotic, as indicated by propidium iodide staining (Fig. 2G). On the basis of these morphological criteria, the number of apoptotic cells was determined in both cell lines. Figure 2C shows the percentage of apoptotic cells determined after 4, 6, or 8 h in the presence of STS (1 μM). In cftr+/+ cells, the percentage of apoptotic cells increased significantly with the time of incubation. At each time point (4, 6, and 8 h), it is interesting to note that the percentage of apoptotic cells is significantly higher in the cftr+/+ cells compared with the cftr−/− cell line. At 8 h, the percentage of apoptotic cells reached 42.5 ± 6.3% in the cftr+/+ cell line but only 3.9 ± 0.5% in the cftr−/− cell line (Fig. 2H). At the same time, the percentage of necrotic cells was determined in both cell lines: At 8 h, the percentage of necrotic cells reached 44.8 ± 6.3% in cftr−/− cells but only 10.4 ± 4.2% in cftr+/+ cells (Fig. 2I).
Role of CFTR and Volume-Regulated Cl− Channels in STS-Induced AVD
A specific cell volume decrease (AVD) has been generally observed during apoptosis (7, 22, 28). Therefore, the time course of relative cell volume variation during STS treatment was measured in proximal cell lines from cftr+/+ and cftr−/− mice. In cftr+/+ cells, a 20% volume decrease was measured 1 h after the beginning of STS exposure (Fig. 3A). The maximal cell volume reduction (39.3 ± 2.8%, n = 8) was reached 6 h after the application of STS. After this time, the volume remained almost constant for at least 12 h (data not shown). In contrast, cftr−/− cells did not undergo STS-induced AVD. The STS-mediated AVD was then studied in the presence of DIDS or NPPB. As illustrated in Fig. 3B, in the cftr+/+ cell line, the STS-induced AVD was strongly inhibited by DIDS and NPPB. The absence of AVD during acute application of DIDS was mainly due to an inhibition of the VISOR Cl− channels, since this drug is a potent inhibitor of these channels in most cells.
The previous experiments (Fig. 3A) strongly suggested the involvement of CFTR in the AVD phenomenon. To further support this involvement, relative cell volume variations during STS exposure were measured in the presence of CFTRinh-172 (Fig. 3C). This inhibitor was found to markedly suppress STS-induced AVD. Moreover, CFTRinh-172 alone did not significantly affect the relative volume.
Cl− Conductances Activated During the AVD Process
Whole-cell experiments were then performed to further analyze the nature of the Cl− conductance triggered by STS exposure. Figure 4A illustrates the Cl− currents recorded in cftr+/+ cells 30 min before the addition of STS (control). The voltage-step protocol elicited linear currents. The corresponding current-voltage (I/V) curve (Fig. 4B) indicated a reversal potential (Erev) of 0.3 ± 0.1 mV and a maximal slope conductance (calculated between +80 and +120 mV) of 5.3 ± 0.8 nS (n = 8). STS exposure induced a large increase in the linear currents within 10 min. These currents were maximal after 30 min, with a maximal slope conductance of 23.4 ± 1.6 nS (n = 8) and a reversal potential of 0.2 ± 0.2 mV. As illustrated in Fig. 4, A–C, the STS-induced currents were not affected by DIDS (1 mM) but were strongly reduced by CFTRinh-172 and by isosmotic replacement of Cl− ions in the bath by I− ions. Moreover, substitution of extracellular Cl− by I− shifted the Erev toward a more positive value (Erev for I− = 28.8 ± 8.3 mV). These characteristics tightly correspond to CFTR Cl− conductance.
Figure 4D illustrates the Cl− currents recorded 1 h after the beginning of STS exposure in cftr+/+ cells. Under these conditions, the developing current exhibited an outwardly rectifying I/V relationship (Fig. 4E) and reached a maximal conductance of 19.6 ± 2.6 nS (n = 8). As illustrated in Fig. 4, D—F, this current was inhibited by the addition of DIDS or NPPB and was not significantly modified either by iodide substitution or by CFTRinh-172 application. In another experimental series, the effect of STS was studied in the cftr−/− cell line. No STS-activated Cl− currents were observed after 0.5 or 1 h of treatment (Fig. 4, G and H).
Mechanism of CFTR Activation During STS Exposure
To determine whether the activation of CFTR by STS could be due to cAMP, experiments were performed in the presence of H89 (an inhibitor of PKA). The preincubation of cftr+/+ cells with H89 for 30 min did not modify the development of STS-activated CFTR Cl− currents (Fig. 5, A and B). The implication of the MAP kinases (ERK1/2) was also checked by using an inhibitor of ERK1/2 phosphorylation (PD98059). As illustrated in Fig. 5, A and B, the preincubation of the cells with PD98059 completely prevented the development of STS-induced CFTR Cl− currents. This result indicates that ERK phosphorylation could be involved in the stimulation of CFTR Cl− currents by STS. The Western blot technique has been used to determine whether the ERK1/2 pathway is activated by STS in the cftr−/− and cftr+/+ cell lines. The results of the Western blotting were expressed in arbitrary units and represent the level of ERK phosphorylation of the two cell lines stimulated by STS for 8, 30, and 60 min (Fig. 5, C and D, n = 3). STS (1 μM) enhanced a rapid and sustained phosphorylation of ERK1/2 within 8 min, which reached a maximum after 60 min in both cell lines. The increased phosphorylation of ERK1/2 was completely prevented by pretreatment with PD98059 (10 μM). ERK1/2 was still stimulated by STS in cftr−/− proximal cells, indicating CFTR-independent ERK1/2 activation (Fig. 5D). It should be noted that the phosphorylation of the two forms of ERK (p42/ERK2 and p44/ERK1) exhibited sensitivity to PD98059 but remained insensitive to H89.
Role of ROS in AVD and Apoptosis
It has been already proposed that ROS production induced by STS could be an early process for inducing AVD. To test this possibility, cftr+/+ and cftr−/− cells were loaded with a membrane-permeable fluorescent probe, carboxy-H2DCFDA, to determine the intracellular level of ROS. In cftr+/+ cells (Fig. 6A), the intracellular level of ROS rapidly increased with time after STS addition. The STS-induced ROS increase was completely abolished in the presence of PD98059, CFTRinh-172, and NAC (a ROS scavenger). NAC alone did not significantly affect the ROS level in the absence of STS (data not shown). Moreover, in the cftr−/− cell line, STS exposure did not increase ROS production (Fig. 6B). CFTR could therefore be involved in STS-induced ROS generation.
Based on these results, we have investigated the relationship among a ROS increase, AVD, and the apoptotic process. For this, we followed the cell volume variation (Fig. 6C) and measured caspase-3 activity (Fig. 6D) during STS exposure in the presence of NAC, CFTRinh-172, PD98059, or H89. Interestingly, all inhibitors (NAC, CFTR 172, and PD 98059), which prevented both the increase in ROS level and the increase in CFTR Cl− conductance, also abolished STS-induced AVD (Fig. 6C) and caspase-3 stimulation (Fig. 6D). By contrast, H89, which had no effect on CFTR Cl− conductance, did not block STS-induced AVD (Fig. 6C) or caspase-3 (Fig. 6D).
STS-Induced Cl− Currents are Activated by ROS Production
Based on the literature, we hypothesized that ROS could activate the VSOR Cl− channels (37). We therefore tested the effect of H89, NAC, and PD98059 on the enablement of Cl− VSOR conductances induced by STS. The results reported in Fig. 7, A and B, show that the inhibitors that prevented both STS-induced ROS production and STS-induced CFTR Cl− currents (PD98059 and NAC) also abolished STS-induced VSOR Cl− conductance. Furthermore, as expected, H89 did not modify the VSOR Cl− conductance activated by STS. To confirm that ROS production could be responsible for the activation of VSOR Cl− channels, whole-cell experiments were performed to test the effect of oxidative stress (500 μM H2O2) on the Cl− currents in cftr+/+ and cftr−/− cell lines. As illustrated in Fig. 7, C and D, in the cftr+/+ cell line addition of H2O2 to the bath solution induced a rapid increase (<15 min) in Cl− currents. These currents were significantly inhibited by addition of DIDS or NAC, suggesting an activation of VSOR Cl− channels by H2O2. Similarly, in the cftr−/− cell line, the addition of H2O2 also increased Cl− currents, which were largely blocked by DIDS and NAC (Fig. 7, E and F).
STS-Induced GSH/GSSG Content Decrease is Mediated by a CFTR-Dependent GSH Efflux
GSH is present in high amounts in proximal tubule cells and reacts to neutralize ROS. Because STS increases ROS production, it was interesting to measure the variation of intracellular GSH concentration during STS exposure in cftr+/+ and cftr−/− cells. For this purpose, the total GSH content (GSH+GSSG) was determined. Figure 8A demonstrates that STS induced, within 30 min, a 44% decrease in intracellular GSH content in the cftr+/+ line. This decrease was inhibited in the presence of PD98059 or CFTRinh-172 (Fig. 8A). In contrast, in the cftr−/− cell line, STS remained without effect (Fig. 8B). This result suggested that the CFTR protein was involved in the reduction of intracellular GSH content induced by STS. We therefore hypothesized that this reduction might be due to the release of GSH either directly through CFTR or by a mechanism controlled by CFTR. To confirm this hypothesis, we have determined whether the flow of various forms of GSH may depend on CFTR. For this purpose, cftr+/+ and cftr−/− proximal cells were incubated for 5 min with the fluorescent probe CMFDA (1 μM). Thus, by analyzing the fluorescence over time, and therefore the concentration of GSH in the cell population, we were able to estimate the GSH efflux. As illustrated in Fig. 8C, addition of STS (1 μM) to CMFDA-loaded cftr+/+ cells accelerated the intracellular decrease in the fluorescent CMFDA conjugate (control: 3.61 ± 0.83%/min, n = 5; STS: 7.92 ± 0.0.72%/min, n = 4). This effect was inhibited in the presence of CFTRinh-172 (STS+CFTRinh-172: 3.84 ± 0.38%/min, n = 3). The histogram shown in Fig. 8E summarizes the slopes of the intracellular fluorescence variations measured under the different experimental conditions. These slopes indirectly represented the rate of GSH efflux from the cells. It clearly appears that STS increased the rate of GSH efflux in the cftr+/+ but had no effect on the cft−− cell line (Fig. 8, D and E). Moreover, this increase was inhibited by CFTRinh-172 and PD98059 and remained insensitive to DIDS. Finally, the decrease in intracellular GSH content following the application of STS is very likely due to an efflux of this tripeptide linked to the activation of CFTR. It was also interesting to know whether the reduction in the intracellular GSH content could induce apoptosis. We therefore tested the effect of BSO (an inhibitor of glutathione biosynthesis) on caspase-3 activity in the presence or absence of STS in the cftr+/+ cell line. As illustrated in Fig. 8F, the addition of BSO (1 mM) induced an increase in caspase-3 activity and enhanced STS-induced caspase-3 activation. These results confirm that the decrease in intracellular GSH content is important for apoptosis. In contrast, BSO application had no effect on caspase-3 activation in the cftr−/− cell line (data not shown).
TNF-α-Induced AVD and Caspase-3 Activation are Mediated by ROS
To confirm the difference obtained between the cftr+/+ and cftr−/− cell lines in STS-induced AVD and caspase-3 activity, experiments were performed by replacing the apoptotic inducer STS with TNF-α+CHX. CHX sensitizes cells to TNF-α-induced apoptosis (28, 33) Under these conditions, TNF-α/CHX also induced AVD (Fig. 9A), ROS production (Fig. 9C), and significantly increased caspase-3 activity (Fig. 10A) in cftr+/+ cell lines only. AVD and caspase-3 activity were inhibited in the presence of DIDS, CFTRinh-172, PD98059, and NAC. Moreover, in the cftr−/− cell line, TNF-α induced neither AVD (Fig. 9B), ROS production (Fig. 9D), nor caspase-3 activation (Fig. 10A). As observed with STS, TNF-α decreased the intracellular GSH content in the cftr+/+ cell line (Fig. 9B). PD98059 and CFTRinh-172 inhibited this decrease. Finally, as expected, no decrease in the intracellular GSH was detected in the cftr−/− cell line (Fig. 10C). In addition, we showed that the application of TNF-α+CHX enhanced a rapid and sustained phosphorylation of ERK1/2 within 30 min in both cell lines (Fig. 10, D and E). ERK1/2 phosphorylation was insensitive to NAC. It seems that the TNF-α+CHX induced the same cascade of events leading to apoptosis as did STS.
Effect of STS on Apoptosis, Cl− Currents, and ROS Production in cftr−/− PCT Cell Line Transfected with hCFTR cDNA
The cftr−/− PCT cell line were stably transfected with PCB6 CFTR plasmid. First, apoptosis was assessed using annexin V-labeling experiments. The results reported in Fig. 11, A and B, clearly show that the percentage of cells stained by annexin V was significantly increased in CFTR-transfected cells after 6 h of incubation with 1 μM STS. Moreover, STS exposure also increased caspase-3 activity within 6 h (Fig. 11C, n = 3).
Regarding Cl− conductance, the application of STS induced a large increase in the linear currents within 10 min (Fig. 11D). In control conditions, the currents exhibited a maximal slope conductance (calculated between +80 and +100 mV) of 2.1 ± 0.5 nS (n = 5) and a Erev of −3 ± 1.8 mV. The application of STS induced a significant increase in Cl− currents and the maximal slope conductance reached 4.5 ± 0.5 nS (n = 7) without modification of the Erev. The STS-induced Cl− conductance were strongly reduced by CFTRinh-172 (Fig. 11D). Additionally, the intracellular level of ROS rapidly increased with time after STS addition (Fig. 11E). Taken as a whole, these results clearly indicates that the exogenous expression of hCFTR in the cftr−/− cell line rescued the ability of STS to activate the main steps of the cascade described in the cftr+/+ cell line, i.e., activation of CFTR Cl− conductance, increase in ROS, and apoptosis.
Effect of STS in Primary Cultures of PCT from Wild-Type and CFTR Knockout Mice
The cell lines used in this study were obtained from primary cultures of microdissected PCT from wild-type and CFTR knockout mice. To eliminate further the possibility that these cells line have drifted from their properties, an experimental series was performed using the original primary cultures. Figures 12 and 13 show that the results were qualitatively identical to those obtained with the derived cell line. In cftr+/+ PCT cells, STS induced early CFTR Cl− currents blocked by CFTRinh-172 and I− and insensitive to DIDS (Fig. 12, A and B). One hour after the onset of STS application, DIDS-sensitive VSOR Cl− currents were recorded. Moreover, NAC completely prevented the development of these currents (Fig. 12, C and D). Both currents were never detected in cftr−/− primary cultures (Fig. 12, E and F). As illustrated in Fig. 13, STS also triggered AVD (Fig. 13A), ROS production.(Fig. 13C), and increased caspase 3 activity (Fig. 13E) These effects were completely inhibited by CFTRinh-172 and were not observed in cftr−/− PCT cells (Fig. 13, B, D, and E). Finally, STS increased ERK phosphorylation in both cell cultures (Fig. 13, G and H). These results strongly strengthen the statement that the involvement of CFTR in the process of STS- induced apoptosis was not due to a difference in the phenotypic properties of each cell lines.
The aim of this work was to investigate the role of CFTR in the control of AVD during STS-induced apoptosis. This objective has been guided by our preceding works demonstrating that CFTR is involved in lovastatin-induced apoptosis (4) and also in the control of RVD (3). Because similarities exist between RVD and AVD, it has been postulated that Cl− and K+ channels involved in both processes could be the same (31). Interestingly, in cultured mouse proximal cells, we have previously found that the TASK2 K+ channel could be the main K+ channel involved in STS-induced AVD (22). Moreover, CFTR and TASK2 are also implicated in the RVD process (3, 5). Based on these observations, we have raised the hypothesis that the involvement of CFTR in apoptosis could be due to the control of AVD. For this purpose, we used two different apoptosis inducers, STS and TNF-α. STS has severe toxic effects on renal cells and primarily mediates mitochondrial apoptosis, whereas TNF-α is an inflammatory cytokine that induces apoptosis mediated by a membrane death receptor. As expected, both substances increased caspase-3 activity and apoptosis. These increases were preceded by a reduction in cell volume corresponding to the AVD phenomenon. It is now well established that this normotonic cell shrinkage during apoptosis is a common mechanism in all cell types (7). However, the present study shows that in cultured mouse proximal cells, AVD and apoptosis depend on the presence of CFTR. This observation was supported by the fact that STS- or TNF-α-induced AVD and caspase-3 activation were not observed in proximal cells lacking CFTR. Consequently, apoptotic death was strongly reduced and necrotic death became predominant.
To understand the role of CFTR in the AVD and apoptotic process, we have analyzed Cl− conductance during AVD. Surprisingly, our results demonstrate that in proximal cells from wild-type mice (cftr+/+), application of STS or TNF-α first enhanced Cl− conductance that exhibited properties consistent with CFTR Cl− channels. Thus Cl− currents were blocked by I− substitution and by CFTRinh-172, a very specific inhibitor of CFTR (27, 30). Interestingly, this CFTR Cl− conductance was progressively replaced (45–60 min after the beginning of STS application) by another type of Cl− conductance that shared similar biophysical and pharmacological features with the VSOR Cl− channel currents. To further implicate CFTR in these Cl− currents, experiments were performed in proximal cells from cftr−/− mice. As expected, these cells did not exhibit any STS- or TNF-α-dependent Cl− currents.
The AVD-induced Cl− channel has already been functionally identified to be the VSOR Cl− channel in different cell types stimulated with STS or TNF-α (32). However, before the present study, the relationship between the activation of this channel and the presence of CFTR during AVD had never been described. Nevertheless, in the RVD phenomenon, it has been postulated that CFTR controls the VSOR channels by modulating autocrine adenosine production in renal cells. The resulting exit of Cl− would provide the driving force to activate the Cl−/HCO3− exchanger, allowing an efflux of HCO3−, which in turn would activate the TASK2 channels (3, 6, 22). In contrast, in the present study, AVD was insensitive to adenosine and to the activity of the Cl−/HCO3− (data not shown), suggesting that CFTR controls AVD by a different mechanism.
It is known that mitochondria play an important role in the regulation of apoptosis (39, 41). This role could be due to production of ROS during the mitochondrial damage induced by apoptotic agents (1, 31, 40). Our results clearly demonstrated that STS and TNF-α induced a rapid increase in ROS in wild-type cells (22). This increase was not observed in cftr−/− cells or in cftr+/+ cells treated with CFTRinh-172. As a result of these observations, we have proposed the hypothesis that the involvement of CFTR in AVD and apoptosis could be linked to the capability of this protein to modulate the level of ROS. Actually, the literature data converge toward the conclusion that ROS serve as an upstream signal to activate K+ and Cl− channels involved in the AVD of cells exposed to mitochondrion-mediated apoptosis inducers. In the present study, the interaction of ROS with ion channels was further supported by the observation that H2O2 application induced swelling-activated Cl− currents in cftr+/+ and cftr−/− cells. Moreover, we have recently demonstrated that, in cultured proximal cells, ROS production activated the TASK2 channel, which is the main K+ conductance involved in STS- and TNF-α-induced AVD and apoptosis (22). Therefore, all our data support the fact that activation of CFTR by an apoptosis inducer is the starting point of the cascade of events that leads proximal cells to undergo apoptosis. It was then interesting to try to explain how CFTR could control the ROS level following exposure to STS or TNF-α.
According to the literature data, GSH, normally present in high amounts in tubular cells, can react with and neutralize ROS (for a review, see Ref. 23). Thus GSH is an important determinant of intracellular redox status. It has been demonstrated that GSH is depleted before the onset of apoptosis induced by various drugs (13) and that reduction of cytosolic GSH renders cells more sensitive to apoptotic agents (12). Moreover, it has been reported that early GSH decrease causes ROS generation before induction of apoptosis. In our present study, we have observed that STS and TNF-α provoked a decrease in the intracellular content of GSH, which correlated well with an increase in GSH efflux and preceded the rise in ROS. These phenomena were strictly dependent on CFTR, since they were strongly blocked by CFTRinh-172 and were not observed in cftr−/− cells.
Hence, it is possible that this CFTR-dependent GSH efflux depletes intracellular GSH, leading to an increase in the ROS level. As discussed above, this ROS increase triggers AVD and apoptosis. Many studies ascertain that CFTR, in addition to its role as a Cl− channel, may function as a permeation pathway by which GSH and GSSG exit the cell (11, 19, 26). Moreover, a direct link between GSH transport by CFTR and apoptosis has already been suggested by measuring H2O2-induced apoptosis in epithelial cells expressing mutant and normal CFTR (17). In the proximal tubule cells, our previous study has indicated that the decrease in total GSH content induced by Cd2+ correlated with an increased GSH efflux and is strictly related to the presence of CFTR (21). Moreover, a relationship between this process and the CFTR has already been suggested.
It is recognized that the depletion of GSH is associated with the initiation of apoptosis. Thus using colonic epithelial cells, Circu et al. (10) reported that STS causes apoptosis that is preceded by significant GSH and GSSG efflux and is independent of changes in cellular GSH/GSSG redox status. Our present study is in accordance with these results and establishes the crucial role of CFTR in apoptosis via the control of GSH level and ROS activation during STS or TNF-α exposure of proximal tubule cells. Another notable observation is the greater sensibility of proximal tubule cells, compared with HT29 cells, to STS challenge. In fact, 1 μM STS elicited a rapid apoptosis in proximal tubule cells compared with HT29 cells, in which 2 μM STS induced apoptosis only 24 h after its addition. These observations are consistent with the notion that STS induces differential susceptibility in proximal tubule and colonic cells (10).
The last issue in our work was how apoptotic agents could activate CFTR Cl− currents and/or GSH fluxes in proximal cells. One possibility would be that STS and TNF-α directly activate CFTR, but this appears unlikely because the activation required minutes instead of seconds to increase Cl− conductance. Alternatively, STS and TNF-α could indirectly regulate CFTR by activating signaling pathways, as suggested by the critical role played by PKA in CFTR opening (8). In cftr+/+ cells, H89 did not inhibit the activation of CFTR Cl− currents by apoptotic agents. This result confirms our previous observations, which indicated that forskolin and cAMP-permeant derivatives did not directly stimulate CFTR Cl− conductance in primary cultures of rabbit (36) or mouse (3) proximal tubules, despite the presence of CFTR mRNA. Therefore, it was important to investigate the possibility that another signal pathway was involved in the activation of CFTR Cl− currents by STS and TNF-α. ERK and MAPKK are candidates known to activate ion channels. Treatment of cftr+/+ cells with STS and TNF-α induced a rapid and sustained activation of ERK1/2. Furthermore, preincubation of the cells with PD98059 drastically reduced the ability of STS and TNF-α to activate ERK1/2, to increase the CFTR Cl− currents, and to accelerate GSH efflux. These data provide evidence that the ERK1/2 pathway could participate in CFTR activation. A relationship between ERK1/2/MAPKK activation and butyrate-induced expression of CFTR was previously demonstrated by Sugita et al. (38). According to their results, the integrity of the R domain of CFTR could be important in this regulation, involving active ERK1/2/MAPK in the biogenesis of the protein. Moreover, ERK1/2 might phosphorylate proteins that interact and regulate CFTR. For example, emerging data implicate MAPK signaling in the control of F-actin (34), and it is also postulated that actin must be directly associated with CFTR to elicit its activation (9). It is evident that the mechanism of CFTR activation by ERK1/2 remains to be further explored. However, whatever the exact mechanism, our data suggest that STS or TNF-α could induce the phosphorylation of ERK1/2 with a kinetic activity compatible with the activation of CFTR.
In conclusion, the present paper reports the role of CFTR in the control of apoptosis in proximal cells. As summarized in Fig. 14, the application of apoptosis inducers activates the CFTR Cl− current. This activation depends on ERK1/2 phosphorylation. Once activated, CFTR might also transport GSH. As suggested in the literature, it is likely that Cl− and GSH share common permeation through CFTR (19, 26). In proximal cells, the increase in GSH permeation through CFTR could deplete the intracellular pool of GSH and decrease the capacity of the cell to scavenge ROS produced by apoptotic inducers. The resulting ROS increase stimulates both VSOR Cl− and TASK2 K+ channels, leading to AVD and apoptosis.
No conflicts of interest are declared by the authors.
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