Role of TASK2 in the Control of Apoptotic Volume Decrease in Proximal Kidney Cells
2007; Elsevier BV; Volume: 282; Issue: 50 Linguagem: Inglês
10.1074/jbc.m703933200
ISSN1083-351X
AutoresSébastien L’Hoste, Mallorie Poët, Christophe Duranton, Radia Belfodil, H. Barrière, Isabelle Rubera, Michel Tauc, Chantal Poujeol, Jacques Barhanin, Phillipe Poujeol,
Tópico(s)Phagocytosis and Immune Regulation
ResumoApoptotic volume decrease (AVD) is prerequisite to apoptotic events that lead to cell death. In a previous study, we demonstrated in kidney proximal cells that the TASK2 channel was involved in the K+ efflux that occurred during regulatory volume decrease. The aim of the present study was to determine the role of the TASK2 channel in the regulation of AVD and apoptosis phenomenon. For this purpose renal cells were immortalized from primary cultures of proximal convoluted tubules (PCT) from wild type and TASK2 knock-out mice (task2-/-). Apoptosis was induced by staurosporine, cyclosporin A, or tumor necrosis factor α. Cell volume, K+ conductance, caspase-3, and intracellular reactive oxygen species (ROS) levels were monitored during AVD. In wild type PCT cells the K+ conductance activated during AVD exhibited characteristics of TASK2 currents. In task2-/- PCT cells, AVD and caspase activation were reduced by 59%. Whole cell recordings indicated that large conductance calcium-activated K+ currents inhibited by iberiotoxin (BK channels) partially compensated for the deletion of TASK2 K+ currents in the task2-/- PCT cells. This result explained the residual AVD measured in these cells. In both cell lines, apoptosis was mediated via intracellular ROS increase. Moreover AVD, K+ conductances, and caspase-3 were strongly impaired by ROS scavenger N-acetylcysteine. In conclusion, the main K+ channels involved in staurosporine, cyclosporin A, and tumor necrosis factor-α-induced AVD are TASK2 K+ channels in proximal wild type cells and iberiotoxin-sensitive BK channels in proximal task2-/- cells. Both K+ channels could be activated by ROS production. Apoptotic volume decrease (AVD) is prerequisite to apoptotic events that lead to cell death. In a previous study, we demonstrated in kidney proximal cells that the TASK2 channel was involved in the K+ efflux that occurred during regulatory volume decrease. The aim of the present study was to determine the role of the TASK2 channel in the regulation of AVD and apoptosis phenomenon. For this purpose renal cells were immortalized from primary cultures of proximal convoluted tubules (PCT) from wild type and TASK2 knock-out mice (task2-/-). Apoptosis was induced by staurosporine, cyclosporin A, or tumor necrosis factor α. Cell volume, K+ conductance, caspase-3, and intracellular reactive oxygen species (ROS) levels were monitored during AVD. In wild type PCT cells the K+ conductance activated during AVD exhibited characteristics of TASK2 currents. In task2-/- PCT cells, AVD and caspase activation were reduced by 59%. Whole cell recordings indicated that large conductance calcium-activated K+ currents inhibited by iberiotoxin (BK channels) partially compensated for the deletion of TASK2 K+ currents in the task2-/- PCT cells. This result explained the residual AVD measured in these cells. In both cell lines, apoptosis was mediated via intracellular ROS increase. Moreover AVD, K+ conductances, and caspase-3 were strongly impaired by ROS scavenger N-acetylcysteine. In conclusion, the main K+ channels involved in staurosporine, cyclosporin A, and tumor necrosis factor-α-induced AVD are TASK2 K+ channels in proximal wild type cells and iberiotoxin-sensitive BK channels in proximal task2-/- cells. Both K+ channels could be activated by ROS production. Like many epithelial cells, renal cells are capable of regulating their volume in response to variations in external osmotic pressure (1Coca-Prados M. Anguita J. Chalfant M.L. Civan M.M. Am. J. Physiol. 1995; 268: C572-C579Crossref PubMed Google Scholar, 2De Smet P. Simaels J. Van Driessche W. Pflugers Arch. 1995; 430: 936-944Crossref PubMed Scopus (33) Google Scholar, 3Lopes A.G. Amzel L.M. Markakis D. Guggino W.B. Proc. Natl. Acad. Sci. U. S. A. 1988; 85: 2873-2877Crossref PubMed Scopus (26) Google Scholar). Briefly, cells respond to an increase in medium osmolarity by a process referred to as regulatory volume increase, whereas cells respond to the dilution of external medium by a regulatory volume decrease (RVD) 2The abbreviations used are:RVDregulatory volume decreaseAVDapoptotic volume decreaseChTxcharybdotoxinIbTXiberiotoxinSTSstaurosporineCsAcyclosporin ATNF-αtumor necrosis factor-αROSreactive oxygen speciescarboxy-H2DCFDA(5-and-6)-carboxy-2′,7′-dichlorodihydrofluorescein diacetateTEAtetraethylammoniumNACN-acetylcysteine. (4Pedersen S.F. Mills J.W. Hoffmann E.K. Exp. Cell Res. 1999; 252: 63-74Crossref PubMed Scopus (98) Google Scholar). A variety of transport pathways have been implicated in both processes and result in rapid water flux across the plasma membrane, which causes cells to recuperate their initial volume correspondingly (5Okada Y. Maeno E. Shimizu T. Dezaki K. Wang J. Morishima S. J. Physiol. 2001; 532: 3-16Crossref PubMed Scopus (466) Google Scholar). Along the proximal tubule, the cells are submitted to hypotonic shock because water accompanies the transport of ions by membrane co-transports. In response to this osmotic stress, the proximal cells undergo a RVD process that is characterized by an exit of Cl- and K+ ions, which finally drives water efflux (6Knoblauch C. Montrose M.H. Murer H. Am. J. Physiol. 1989; 256: C252-C259Crossref PubMed Google Scholar). However, changes in cell volume are not only due to variation in medium osmolarity. It is now well established that the initial process leading toward apoptotic cell death is coupled to normotonic cell shrinkage (7Maeno E. Ishizaki Y. Kanaseki T. Hazama A. Okada Y. Proc. Natl. Acad. Sci. U. S. A. 2000; 97: 9487-9492Crossref PubMed Scopus (660) Google Scholar), called apoptotic volume decrease (AVD). The proximal tubule is a major site of agent-induced nephrotoxicity (drugs, heavy metals, hypoxia etc.), which can induce AVD and lead to cell death by apoptosis. It is therefore interesting to understand the mechanisms involved in this phenomenon. As in RVD, the changes in cell volume during AVD are the consequence of an exit of Cl- and K+ from the cells, and the question arises as to whether the Cl- and the K+ currents are driven by the same type of channels (8Okada Y. Maeno E. Comp. Biochem. Physiol. A Mol. Integr. Physiol. 2001; 130: 377-383Crossref PubMed Scopus (169) Google Scholar). The AVD-induced Cl- channel has been identified as a volume-sensitive outwardly rectifying Cl- channel in both HeLa cells and cardiomyocytes treated with staurosporine (9Shimizu T. Numata T. Okada Y. Proc. Natl. Acad. Sci. U. S. A. 2004; 101: 6770-6773Crossref PubMed Scopus (210) Google Scholar, 10Okada Y. Shimizu T. Maeno E. Tanabe S. Wang X. Takahashi N. J. Membr. Biol. 2006; 209: 21-29Crossref PubMed Scopus (215) Google Scholar). This channel shares many properties with the Cl- channel that are induced in RVD in mouse proximal cells (5Okada Y. Maeno E. Shimizu T. Dezaki K. Wang J. Morishima S. J. Physiol. 2001; 532: 3-16Crossref PubMed Scopus (466) Google Scholar). The molecular nature of this Cl- channel is not fully elucidated, but the literature data converges toward the conclusion that this Cl- channel type is probably ubiquitously expressed in animal cells (7Maeno E. Ishizaki Y. Kanaseki T. Hazama A. Okada Y. Proc. Natl. Acad. Sci. U. S. A. 2000; 97: 9487-9492Crossref PubMed Scopus (660) Google Scholar). By contrast, the molecular identity of the K+ channel involved in both AVD and RVD is still under discussion because different candidates have been proposed depending on the tissue under study (8Okada Y. Maeno E. Comp. Biochem. Physiol. A Mol. Integr. Physiol. 2001; 130: 377-383Crossref PubMed Scopus (169) Google Scholar, 11Burg E.D. Remillard C.V. Yuan J.X. J. Membr. Biol. 2006; 209: 3-20Crossref PubMed Scopus (130) Google Scholar, 12Remillard C.V. Yuan J.X. Am. J. 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Physiol. 2002; 282: C588-C594Crossref PubMed Scopus (77) Google Scholar) have provided evidence that two-pore domain K+ channels underlie K+ efflux during AVD in mouse embryos. Based on these observations, it was reasonable to postulate that TASK2 K+ channels could also be involved in the regulation of AVD and apoptosis in the proximal tubules. Thus, the present study addresses the role of TASK2 in apoptosis induced by staurosporine. In a large variety of cells, staurosporine is known to induce apoptosis through a mitochondria-mediated pathway and to increase oxidative stress by the production of ROS generated by the mitochondria (9Shimizu T. Numata T. Okada Y. Proc. Natl. Acad. Sci. U. S. A. 2004; 101: 6770-6773Crossref PubMed Scopus (210) Google Scholar, 18Gil J. Almeida S. Oliveira C.R. Rego A.C. Free Radic. Biol. Med. 2003; 35: 1500-1514Crossref PubMed Scopus (93) Google Scholar). In proximal cells, this mitochondrial mechanism may be the predominant mode of inducing apoptosis in the presence of many nephrotoxic agents (19Thevenod F. Nephron Physiol. 2003; 93: 87-93Crossref Scopus (225) Google Scholar, 20Baek S.M. Kwon C.H. Kim J.H. Woo J.S. Jung J.S. Kim Y.K. J. Lab. Clin. Med. 2003; 142: 178-186Abstract Full Text Full Text PDF PubMed Scopus (157) Google Scholar, 21Erkan E. Devarajan P. Schwartz G.J. J. Am. Soc. Nephrol. 2007; 18: 1199-1208Crossref PubMed Scopus (80) Google Scholar). Therefore, the staurosporine-induced apoptosis is a useful model to assess the role of ion channels in controlling apoptosis in renal cells. The present study was conducted on proximal tubule cell lines originating from wild type and task2-/- mice. In wild type mice, we demonstrated that staurosporine-induced AVD was mainly associated with the activity of TASK2 K+ channels. Surprisingly, staurosporine-induced AVD persisted in the task2-/- proximal cell line and could be controlled by a Ca2+-activated K+ channel that is sensitive to iberiotoxin. regulatory volume decrease apoptotic volume decrease charybdotoxin iberiotoxin staurosporine cyclosporin A tumor necrosis factor-α reactive oxygen species (5-and-6)-carboxy-2′,7′-dichlorodihydrofluorescein diacetate tetraethylammonium N-acetylcysteine Transformation of Primary Cultures with pSV3 neo and Culture Protocol—The primary cell culture technique has been described in detail in previous studies (22Belfodil R. Barriere H. Rubera I. Tauc M. Poujeol C. Bidet M. Poujeol P. Am. J. Physiol. 2003; 284: F812-F828Google Scholar). Briefly, 10-day-old primary cultures of S1 and S2 segments of proximal tubules from wild type and task2-/- mice were transfected with pSV3 neo using Lipofectin (Invitrogen). After 48 h, selection of the clones was performed by the addition of G418 (500 μg/ml). Culture medium (Dulbecco's modified Eagle's medium-F12, Sigma, Saint Quentin Fallavier, France) containing 250 μg/ml G418, 15 mm NaHCO3, 20 mm HEPES (pH 7.4), growth factors (23Barriere H. Rubera I. Belfodil R. Tauc M. Tonnerieux N. Poujeol C. Barhanin J. Poujeol P. J. Membr. Biol. 2003; 193: 153-170Crossref PubMed Scopus (35) Google Scholar), and 1% FCS was changed every day. Resistant clones were isolated, subcultured, and used after 10 trypsinization steps. Immortalized proximal wild type and task2-/- 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. Apoptosis Induction—Apoptosis was induced by staurosporine (STS, 1 μm), cyclosporin A (CsA, 25 μm), or tumor necrosis factor-α (TNF-α, 0.5 ng/ml) in proximal wild type and task2-/- cell lines that were maintained in serum and growth factor-free culture medium (Dulbecco's modified Eagle's medium/F-12, Sigma) in a 5% CO2 atmosphere at 37 °C. Staurosporine (STS) and cyclosporin A (CsA) were dissolved in Me2SO. The quantity of Me2SO added to the incubation solutions never exceeded 0.1%. Control experiments were performed by incubating the cells with 0.1% Me2SO only. Measurement of Caspase-3 Activity—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 wild type and task2-/- cells. 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 (Bio-Tek Instruments, Inc.). Apoptotic Cell Counts—STS-induced apoptosis was studied in wild type and task2-/- cell lines. Cells were grown in 35-mm Petri dishes. After an appropriate incubation with the apoptosis inductor (STS), living cells were carefully washed with fresh culture medium and incubated 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 of orcein, 2 ml of ethanol 70%, 150 μl of 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 presented 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 three different observers who were blinded to the culture conditions. The numbers of cells with DNA condensation and propidium iodide staining were expressed as the percentage of total cells. Cell Volume Measurement—Cell volume was measured by an electronic sizing technique using a CASY 1 cell counter (SCHÁRFE SYSTEM®). Briefly, proximal wild type and task2-/- cells that were exposed to different treatments (STS, CsA, or TNF-α) were rapidly trypsinized (1 times for 45 s), and cell volume measurement was performed just after suspending the cells in Casyton® solution (NaCl isotonic solution). Electrophysiological Studies—Whole cell currents were performed on cultured proximal wild type and task2-/- 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-megaohm resistance) were made from borosilicate capillary tubes (1.5 mm outer diameter, 1.1 mm inner diameter; 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 higher than 20 megaohms were discarded. The offset potentials between both electrodes were zeroed before sealing, and the liquid junction potential was measured experimentally prior to each experiment and corrected accordingly (measured junctions potentials were 11.34 ± 0.79 mV for K+ conductance experiments). Solutions were perfused in the extracellular bath using a four-channel glass pipette, with the tip placed as close as possible to the clamped cell. Voltage-clamp commands, data acquisition, and data analysis were controlled via 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): 100 K-gluconate, 25 KHCO3, 20 KCl, 10 HEPES (pH 7.4 adjusted with 1 n KOH), 5 MgATP, and 0 or 30 EGTA (Pos = 300 milliosmole/kg of H2O). To avoid spontaneous activation of volume-sensitive K+ currents, the bath solution was slightly hyperosmotic and contained (in mm): 110 NMDG-Cl, 5 glucose, 5 potassium gluconate, 1 CaCl2, 1 HEPES (pH 7.4 adjusted with 1 n HCl), and 100 mannitol (Pos = 330-340 milliosmole/kg of H2O). Measurement of Reactive Oxygen Species (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 wild type and task2-/- cells were incubated in Petri dishes at 37 °C for 30 min in the presence of carboxy-H2DCFDA (10 μm) and gently washed in serum-free culture medium. Two experimental protocols were developed to measure the fluorescence increase according to the time kinetic leading to ROS increase. For rapid kinetic (less than 1 h), cells were rapidly trypsinized (×10 for 45 s) and incubated in the absence or presence of either STS (1 μm) or NAC (N-acetylcysteine, 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. For long time kinetic (experiments performed in the presence of TNF-α), the ROS production was monitored by using fluorescent video microscopy. Briefly, proximal cell lines grown in 35-mm Petri dishes were incubated in the presence of carboxy-H2DCFDA (10 μm) at 37 °C for 30 min in a humidified atmosphere of 5% CO2, 95% air. Cells were gently washed and incubated in an isotonic serum-free medium containing 30 mm HEPES in the absence or presence of TNF-α. The variation of fluorescence was measured every 15 min (during 24 h) at 538 nm. Data are expressed as the fluorescence ratio F/F0, where F is the relative fluorescence intensity measured every 15 min and F0 the relative fluorescence intensity measured at t = 0. The mean values of ROS production were obtained from analysis of 18-25 cells in each culture. Intracellular Ca2+ Measurements—The intracellular Ca2+ concentration ([Ca2+]i) was measured in cells grown in Petri dishes and loaded for 45 min at room temperature in the presence of fura 2-AM (2 μm) and pluronic acid (0.01%). The cells were washed with a NaCl solution containing in mm, 140 NaCl, 5 KCl, 1 MgSO4, 1 CaCl2, 5 glucose, and 20 HEPES (pH 7.4). Cells were successively excited at 350 and 380 nm and the paired images were digitized. 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. (24Grynkiewicz G. Poenie M. Tsien R.Y. J. Biol. Chem. 1985; 260: 3440-3450Abstract Full Text PDF PubMed Scopus (80) Google Scholar) was used to calculate [Ca2+]i from the dual wavelength-to-fluorescence ratio. Chemical Compounds—STS (Sigma) and CsA (Sigma) were prepared in Me2SO and used at final concentrations of 1 and 25 μm, respectively. TNF-α (Sigma) was prepared in distilled water and used at a final concentration of 0.5 ng/ml. Clofilium was prepared at 10 mm in a solution containing 50% Me2SO, 50% water. Clofilium and CsA were a gift from Dr. Barhanin (UMR CNRS 6097). Stock solutions of xanthine (50 mm) and xanthine oxidase (50 milliunits ml-1) were prepared and kept at +5 °C and -20 °C, respectively. The fluorescent probe carboxy-H2DCFDA (Molecular Probes) was used at 10 μm (stock solution at 10 mm in Me2SO). Fura 2-AM (Molecular Probes) was dissolved at 3 mm in Me2SO and added to the loading solution at a final concentration of 2 μm, along with 0.01% pluronic acid. NAC (10 mm), tetraethylammonium (TEA, 1 mm), charybdotoxin (ChTX, 10 nm), and iberiotoxin (IbTX, 100 nm) were obtained from Sigma. Induction of Caspase-3 Activation and Chromatin Condensation by STS in Wild Type and task2-/- Proximal Cell Lines—Previous experiments performed on proximal cell lines have established a crucial role for the TASK2 K+ channel in the regulatory volume decrease during hypotonic shock. Furthermore, a specific cell volume decrease (AVD) is generally observed during apoptosis process. In the present study, the involvement of TASK2 channels during chemically induced apoptosis was investigated. For this purpose, proximal cell lines from wild type and task2-/- mice were exposed to STS (1 μm) for 6 h, and the caspase-3 activity was determined. As illustrated in Fig. 1A, STS exposure induced a strong increase in caspase-3 activity in wild type cells. Interestingly, this increase was ∼2-fold higher than that observed in task2-/- cells (Fig. 1A). This suggests that TASK2 channels were involved in the STS-induced apoptotic process. The STS-induced caspase-3 activation was significantly inhibited by clofilium, high [K+]e, or external acidic pH confirming the involvement of TASK2 (these effectors have already been shown to inhibit TASK2 K+ currents). In these cells, the potent blocker of Ca2+-dependent K+ channels, IbTX, did not affect the STS-induced caspase-3 activation. Surprisingly, the addition of STS still enhanced the level of caspase-3 in task2-/- cells. As expected, this moderate increase was not modified by the addition of clofilium or by the acidification of the external pH. However, the application of IbTX or high [K+]e abolished STS-induced caspase-3 activation. These results suggested that, in task2-/- cells, the STS-induced caspase-3 activation could be driven by an IbTX-sensitive K+ channel. To verify these observations, the apoptosis phenomenon was also assessed on the basis of morphological criteria. Wild type and task2-/- 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. In sharp contrast, after STS exposure (1 μm, 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 the plasma membrane integrity. The condensation and fragmentation of DNA clearly show that STS induced apoptosis. Less than 8.9 ± 1.2% of the total cells exhibited propidium iodide-labeled nuclei. These morphological characteristics could also be observed independently with orcein staining: control cells (without STS) did not exhibit chromatin condensation. By contrast, a dense and thin crown of nuclear coloration, typical of chromatin condensation, could be observed after STS exposure. Hoescht-33258 and orcein staining (data not shown) on STS-treated task2-/- cells revealed a significant decrease of condensed and fragmented chromatin figures as compared with wild type cells. On the basis of these morphological criteria, the number of apoptotic cells was determined in both cell lines. Fig. 1B shows the percentage of apoptotic cells determined after 4, 6, or 8 h in the presence STS (1 μm). In both cell lines, the percentage of apoptotic cells increased significantly with the time of incubation. At each time (4, 6, and 8 h), the percentage of apoptotic cells was higher in wild type than in task2-/- cell lines. At 8 h, the percentage of apoptotic cells reached 40.2 ± 5.2% in wild type cell lines but only 19.9 ± 4.4% in task2-/- cell lines. At the same time (8 h), the percentage of necrotic cells reached 33.4 ± 4.2% in the task2-/- cell but only 10.1 ± 4.1% in wild type cells (Fig. 1C). STS-induced AVD in Proximal Cell Lines—To check whether the STS-induced apoptotic process was related to an AVD phenomenon, the time course of relative cell volume variation during STS treatment was measured in proximal cell lines from wild type and task2-/- mice. In both cell lines, cell shrinkage started as early as 1 h after STS exposure (1 μm, Fig. 2A). Six h after application of STS, the relative mean cell volume decreased by 34.2 ± 2.6% in wild type cell lines but only by 20.2 ± 4.5% in task2-/- cell lines. The STS-mediated AVD was then studied in the presence of clofilium, ChTX, high [K+]e, or IbTX. As illustrated in Fig. 2B, in wild type cell lines, the STS-induced AVD was strongly inhibited by clofilium, high [K+]e but not by ChTX or IbTX. Moreover, AVD was completely blocked by the acidification of the external solution (Fig. 2B). In task2-/- cell lines, the moderate STS-induced AVD was inhibited by ChTX, IbTX, or high [K+]e and was insensitive to clofilium or external acidic pH (Fig. 2C). STS Activates Two Different Types of K+ Currents in Wild Type and task2-/- Proximal Cell Lines—The above experiments suggested the involvement of the TASK2 channel in the AVD process in wild type proximal cells. Whole cell experiments were then performed to further analyze the nature of the K+ conductance triggered by STS exposure. Fig. 3A illustrates the K+ currents recorded in wild type proximal cells before the addition of STS, the voltage step protocol elicited small outwardly rectifying currents. The corresponding I/V curve (Fig. 3B) indicated a reversal potential of -71.5 ± 5 mV and a maximal slope conductance (calculated between +80 and +120 mV) of 3.3 ± 0.7 nS (n = 15). STS exposure induced a large increase of the outward currents, with a maximal slope conductance of 20.3 ± 1.4 nS (n = 15) without significant modification of the reversal potential (-74.3 ± 3 mV). To determine the possible role of cytosolic Ca2+ in the development of STS-induced currents, experiments were performed using a pipette solution containing the Ca2+ chelating agent EGTA (30 mm) to greatly reduce the intracellular free Ca2+ concentration. In 100% of the cells tested, the STS-induced conductance was not modified. These STS-activated K+ currents were inhibited by clofilium (10 μm, maximal slope conductance 3.5 ± 0.8 nS, n = 15). Fig. 3C showed the mean currents recorded at +100 mV under different experimental conditions; the STS-induced currents exhibited no sensitivity to TEA, IbTX, or ChTX but were strongly reduced in the presence of clofilium or when perfusing an acidic external bath solution (pH 6). These characteristics tightly correspond to TASK2 K+ conductance. Whole cell experiments were also performed in the task2-/- cell line to better characterize the IbTX-sensitive K+ permeability involved in the residual AVD described in Fig. 2, A-C. Fig. 3D illustrates the K+ currents recorded in task2-/- proximal cells before and after STS exposure. Without STS, the voltage step protocol elicited small outwardly rectifying K+ currents (maximal slope conductance = 2.6 ± 1.3 nS, Erev = -69.2 ± 2.5 mV, n = 16, Fig. 3E). STS exposure significantly increased the outward currents, which reached a maximal conductance of 15.6 ± 2.3 nS (Erev =-70.2 ± 2.5 mV, n = 16) without significant change of the reversal potential. These STS-induced K+ conductances were inhibited by the addition of ChTX or IbTX to the external solution. Experiments were also performed using pipette solution containing Ca2+ chelating agent EGTA (30 mm) to greatly reduce the intracellular free Ca2+ concentration. In 100% of the cells tested, the STS-induced conductance was completely blocked (n = 5). As shown in Fig. 3F, the mean STS-induced K+ currents (measured at +100 mV) exhibited a different pharmacological profile as compared with wild type cells; they were inhibited by TEA, IbTX, and ChTX but remained insensitive to clofilium or perfusion with an acidic external bath solution (pH 6). These pharmacological properties of K+ channel were analyzed in both cell types using the highest currents recorded in the presence of STS. Thus, the mean currents recorded at +100 mV were smaller in task2-/- cells (986 ± 23 pA, n = 16) than in wild type cells (1315 ± 36 pA, n = 15). Fig. 3G further illustrates this difference by showing the distribution of the STS-induced current recorded in wild type and task2-/- cell lines. The individual records were arranged in 5 arbitrary groups according to the maximal current recorded at +100 mV in the presence of STS. In wild type cells, the STS-induced K+ currents measured at +100 mV were 1103.8 ± 55 pA (n = 27) and only 658.3 ± 65 pA in task2-/- cells (n = 28). The difference was statistically significant (p ≤ 0.0001 by t test). All together, in the wild type cell line, 93% of the STS-induced currents are between 800 and 1800 pA. In sharp contrast, in the task2-/- cell line only 57% of the records are in this range of currents. Moreover 12 records are moderately stimulated by STS in the task2-/- cell li
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