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Cellular |
1 Ordway Research Institute, Albany, NY 12208, USA
2 Center for Neuropharmacology and Neuroscience, Albany Medical College, Albany, NY 12208, USA
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Abstract |
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(Received 17 December 2005;
accepted after revision 2 March 2006;
first published online 9 March 2006)
Corresponding author A. A. Mongin: Center for Neuropharmacology and Neuroscience, Albany Medical College, 47 New Scotland Ave. (MC-136), Albany, NY 12208, USA. Email: mongina{at}mail.amc.edu
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Introduction |
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One hypothetical route for swelling-activated release of organic osmolytes is the ubiquitously expressed volume-regulated anion channel(s) (VRAC), which is activated by cell swelling and is permeable to a variety of inorganic and small organic anions, including the amino acids taurine, glutamate and aspartate (Strange et al. 1996; Okada, 1997; Nilius & Droogmans, 2003). In electrophysiological studies VRAC currents are identified based on a combination of biophysical properties, such as moderate outward rectification, time-dependent inactivation at positive potentials, and Eisenman's type I anion selectivity sequence (SCN– > I–> Br– > Cl– > F– > gluconate) (Okada, 1997; Nilius et al. 1997). However, in spite of an extensive experimental search, the molecular nature of VRAC has not yet been identified.
Although the amino acid permeability of the VRAC has been demonstrated in a number of electrophysiological studies (Banderali & Roy, 1992; Jackson & Strange, 1993; Jackson et al. 1994; Roy, 1995; Boese et al. 1996), it is still uncertain whether the VRAC serves as the only, or even a major, pathway for cell swelling-activated amino acid release (Junankar & Kirk, 2000). Several groups have reported that, at least in some cell types, swelling-activated Cl– and organic osmolyte fluxes are mediated by separate transport mechanisms (Lambert & Hoffmann, 1994; Tomassen et al. 2004). Furthermore, other indirect evidence suggests that volume-sensitive organic osmolyte release may involve more than one permeability pathway (Ruhfus et al. 1996; Mongin et al. 1999).
Among Cl– channels that have been cloned to date, ClC-2, plasmalemmal VDAC (p-VDAC or VDACL), and ClC-3 exhibit volume sensitivity, but their other characteristics differ from VRAC properties (Grunder et al. 1992; Duan et al. 1997; Sabirov et al. 2001). At least two of these channels, VDAC and ClC-3, show a measurable permeability to excitatory amino acids (Duan et al. 1997; Sabirov et al. 2001). The objective of the present study was to identify Cl– channels that contribute entirely or partially to swelling-activated release of excitatory amino acids from swollen rat astrocytes. To achieve this aim we used a number of pharmacological agents, which discriminate between particular Cl– channels. Some of the data included in this manuscript have been presented in a preliminary form (Mongin et al. 2005).
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Methods |
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Confluent primary astrocyte cultures were prepared from the cerebral cortex of newborn Sprague-Dawley rats as previously described (Frangakis & Kimelberg, 1984), with minor modifications as listed below. All animal procedures were performed according to the NIH Guide for Animal Care and approved by the institutional animal care and use committee. Briefly, neonatal rats were killed by rapid decapitation, the cerebral cortices were removed and separated from meninges and basal ganglia, and tissue was dissociated with the neutral protease dispase. Dissociated cells were seeded on poly D-lysine-coated 18-mm glass coverslips (Carolina Biological Supply, Burlington, NC, USA) and grown for 3–4 weeks in minimal essential medium (MEM) supplemented with 10% heat-inactivated horse serum (HIHS), 50 U ml–1 penicillin, and 50 µg ml–1 streptomycin at 37°C in a humidified 5% CO2–95% air atmosphere. Culture medium was replaced twice a week. After 2 weeks of cultivation, penicillin and streptomycin were removed from the culture medium. Immunocytochemistry showed that 
98% of the cells stained positively for the astrocytic marker glial fibrillary acid protein (GFAP).
Excitatory amino acid efflux measurements
Excitatory amino acid efflux measurements were performed as previously described (Mongin et al. 1999; Mongin & Kimelberg, 2002). Briefly, astrocytes grown on glass coverslips were loaded overnight with D-[3H]aspartate (4 µCi ml–1) in 2.5 ml of MEM containing 10% HIHS in an incubator set for 5% CO2–95% air at 37°C. Before the start of the efflux measurements, the cells were washed free of extracellular isotope and serum-containing medium in Hepes-buffered solution. The isoosmotic Hepes-buffered medium contained (mM) 135 NaCl, 3.8 KCl, 1.2 MgSO4, 1.3 CaCl2, 1.2 KH2PO4, 10 D-glucose and 10 Hepes, pH adjusted to 7.4 with NaOH. In those experiments where CdCl2 was tested, phosphate salts were removed from solutions. The coverslips were inserted into a Lucite perfusion chamber that had a depression precisely cut in the bottom to accommodate the coverslip and a Teflon screw top, leaving a space above the cells of 100–150 µm in height. The cells were superfused at a constant flow rate of 1.2 ml min–1 in an incubator set at 37°C with Hepes-buffered medium. In hypoosmotic medium NaCl concentration was reduced to 85 mM. The osmolarities of all buffers were checked with a freezing point osmometer (µOsmette, model 5004, Precision Systems, Natick, MA, USA) and were 287 ± 2 and 198 ± 2 for isoosmotic and hypoosmotic media, respectively. Superfusate fractions were collected at 1-min intervals. At the end of each experiment, the isotope remaining in the cells was extracted with a solution containing 1% sodium dodecylsulphate plus 4 mM EDTA. Four milliliters of Ecoscint scintillation cocktail (National Diagnostics, Atlanta, GA, USA) was added, and each fraction was counted for 3H in a Packard Tri-Carb 1900TR liquid scintillation analyser (PerkinElmer, Downers Grove, IL, USA). The percentage fractional isotope release for each time point was calculated by dividing radioactivity released in each 1-min interval by the radioactivity left in the cells (the sum of all the radioactive counts in the remaining fractions up to the beginning of the fraction being measured plus the radioactivity left in the cell digest) with a custom computer program.
Electrophysiological experiments
For electrophysiological measurements, primary astrocytes grown in confluent cultures were detached using recombinant protease TrypLE (Invitrogen, Carlsbad, CA, USA) and re-plated on poly D-lysine-treated glass coverslips at low density in MEM supplemented with 10% HIHS. The following day, serum-containing medium was replaced with serum-free Opti-MEM (Invitrogen) with addition of 300 µM dibutyryl-cyclicAMP (dbcAMP), and single cells were patch-clamped within 24–72 h. Whole-cell recordings were performed at room temperature as previously described (Kubo & Okada, 1992; Liu et al. 1998). Patch electrodes were fabricated from borosilicate glass capillaries using a micropipette puller (P-87, Sutter Instruments, Novato, CA, USA), and had a resistance of 3–3.5 M
when filled with pipette solution. Series resistance was 
15 M. Currents were recorded using an Axopatch 200B amplifier (Axon Instruments, Union City, CA, USA). pCLAMP software (version 9.2, Axon Instruments) was used for command pulse control, data acquisition and analysis. Current signals were filtered at 2 kHz using a four-pole Bessel filter and digitized at 4 kHz. The time course of current development was monitored by applying alternating 2-s step pulses every 15 s from a holding potential of 0 to ± 40 mV. After attaining significant activation of Cl– currents, we tested their biophysical properties by applying 2-s step pulses from 0 mV to test potentials of –100 to +100 mV in 20-mV increments. The isoosmotic external solution contained (mM): 110 CsCl, 2 CaCl2, 1 MgSO4, 5 glucose, 10 Hepes, and 60 mannitol (pH 7.4, 290 mosmol l–1). The hypoosmotic solution was made by omitting the mannitol from the isotonic solution and had an osmolarity of 230 mosmol. The pipette solution contained (mM): 110 CsCl, 1 MgSO4, 1 Na2-ATP, 0.3 Na2-GTP, 15 Na-Hepes, 10 Hepes, 1 EGTA and 10 mannitol (pH 7.3, 255 mosmol l–1). The osmolarity of the pipette solution was set lower than that of the isotonic bath solution in order to prevent spontaneous cell swelling after attaining the whole-cell mode (Worrell et al. 1989).
Total RNA isolation and RT-PCR
Total RNA was isolated from cultured astrocytes grown to confluency in 60-mm Petri dishes. TRIzol reagent (Invitrogen) was used according to the manufacturer's instructions as an improvement to the single-step RNA isolation method developed by Chomczynski & Sacchi (1986). RNA samples were incubated for 15 min at 29°C with DNase I mix in the presence of RNase inhibitor to digest any contaminating genomic DNA. The RNA samples were then transferred onto ice and 25 mM EDTA (pH 8.0) was added to each tube. After 5-min incubation at 75°C, the PCR tubes were immediately placed on ice again. A concentrated reagent mix, containing 10x PCR buffer, 10 mM dNTPs, 0.1 M DTT, RNAse inhibitor and 5 ng µl–1 random hexamers, was added to the reaction tubes. The reaction mixture was then heated for 3 min at 42°C. One-hundred units of Moloney murine leukaemia virus reverse transcriptase (Invitrogen) was then added, and the incubation continued for another 60 min. The enzyme was then inactivated by heating the reaction mixture for 10 min at 65°C. The RT reaction products were stored at –20°C until used in PCR.
We used the previously published sequences of oligonucleotide primers for ClC-1, ClC-2, ClC-3, ClC-4, ClC-5, CFTR, MDR-1 and VDAC-1 (see Table 1 for primer sequences and references). Ten-microlitre aliquots containing 10 pmol of each 5'- and 3'-primer were added to 10 µl of the RT reaction mixture and overlaid with 30 µl of mineral oil. The reaction tubes were placed in a thermocycler block (iCycler, Bio-Rad, Hercules, CA, USA), and heated for 2 min at 94°C. Five microlitres of the mixture containing dNTPs and 1.5 U of Taq polymerase (Invitrogen) was then added. The final concentrations of all components were as follows: 1 x PCR buffer without Mg2+ (Roche), 200 µM of each dNTP (Invitrogen), 2.0 mM MgCl2, 1 µM of each primer, and 30 mU µl–1 of Taq polymerase. Reactions were set for 38 cycles. Denaturation temperature was set at 94°C, elongation temperature at 72°C, and annealing temperature at 60°C.
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Materials
D-[3H]Aspartate (specific activity 18 Ci mM–1) was obtained from PerkinElmer Life Sciences (Boston, MA, USA) or Amersham Biosciences (Piscataway, NJ, USA). Dispase (neutral protease dispase grade II) was purchased from Roche (Baton Rouge, LA, USA). All cell culture reagents were from Invitrogen (Carlsbad, CA, USA). 4-[(Butyl-6,7-dichloro-2-cyclopentyl-2,3-dihydro-1-oxo-1H-inden-5-yl)oxy]butanoic acid (DCPIB) and verapamil were obtained from Tocris (Ellisville, MO, USA). Phorbol 12,13-dibutyrate (PDBu) was from Calbiochem (La Jolla, CA, USA). Phloretin and all other reagents and salts were purchased from Sigma (St Louis, MO, USA) and were of highest grade available.
Data analysis
Amino acid release during the hypoosmotic medium exposure was analysed using repeated measures ANOVA to test for effects of drug and the effect of time and their interaction (P-values for the effects of drug are included in figure legends). Analysis also included planned comparison of mean peak release values in the presence or absence of the tested drug, using Fisher's LSD test (these P-values are presented in the text of the Results section). Peak release values were observed between the 2nd and 3rd minutes of exposure to hypoosmotic medium and represent the maximal activity of the amino acid permeability pathway. In electrophysiological experiments, effects of inhibitors were normalized to control Cl– currents in the same experiment. In these experiments significance of drug effects was tested using Student's t test comparing normalized currents in the presence of drug to 100% (P-values are presented in the text of the Results section). Statistical analysis was performed using Statistica 6.1 (StatSoft, Tulsa, OK, USA) or Origin 7.5 (OriginLab, Northampton, MA, USA).
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Results |
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In cultured rat astrocytes, 30% reduction in medium osmolarity induced transient efflux of D-[3H]aspartate with the maximal release rate of 1.0–1.5%, which was 10–20 times higher that that seen in non-stimulated cells under isoosmotic conditions (Fig. 1A). The broad spectrum Cl– channel blocker NPPB, at 100 µM, blocked the peak release by 67 ± 7% (significantly different from control release, P= 0.003, Fig. 1A). In parallel experiments, we measured whole-cell Cl– currents in response to 20% reduction in medium osmolarity. The extremely flat morphology of cultured astrocytes made it difficult to patch cells and maintain the seal during exposure to hypoosmotic medium. Therefore, in electrophysiological experiments we pretreated cells with dibutyryl-cAMP to make the cell bodies rounder. Such treatment improved experimental success rate, while not causing any substantial changes in either current densities or the biophysical properties of swelling-activated Cl– currents (data not shown). As seen in Fig. 1B, hypotonic stress activated large whole-cell Cl– currents. Average steady-state current density increased from 3.1 ± 0.5 pA pF–1 under isoosmotic conditions to 19.7 ± 3.3 pA pF–1 at +40 mV (n= 5, data not shown). These currents exhibited moderate outward rectification and time-dependent inactivation at large positive potentials (Fig. 1B, inset). As shown in Fig. 1B, 100 µM NPPB strongly and reversibly inhibited swelling-activated Cl– currents, reducing them by 95.2 ± 3.4% (n= 5, P < 0.001) of control. Because of the slower activation of Cl– currents in electrophysiological experiments (Fig. 1B), compared to activation of D-[3H]aspartate release in efflux experiments (Fig. 1A), we additionally performed simultaneous measurements of D-[3H]aspartate and 36Cl– fluxes in hypoosmotically swollen cells. In intact cells, activation of D-[3H]aspartate and 36Cl– releases occurred at the same time (online Supplemental material, Supplemental Fig. 1).
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To determine which candidate Cl– channels are expressed in our preparation of cultured astrocytes, we assayed the mRNA in these cells using an RT-PCR method with previously published sequence-specific primers (Huber et al. 1998; Enz et al. 1999; Bres et al. 2000; Kulka et al. 2002; Albermann et al. 2005; for primer sequences see Table 1). As seen in Fig. 2, all the candidate Cl– channels were present in our preparation at the mRNA level. The resulting PCR products corresponded to their predicted molecular weights, and their molecular identity was confirmed by sequencing. The protein expression for Cl– channels of interest in cultured astrocytes, with the exception of ClC-4, as well as similar mRNA expression has been shown previously by Parkerson & Sontheimer (2004). We used two negative controls for the RT-PCR reactions. To check for genomic DNA contamination we omitted reverse transcriptase from the reaction mixtures. This resulted in disappearance of the PCR signal. Additionally, we used the sequence-specific primers for the ClC-1 gene product, which is not expressed in the brain or in cultured astrocytes. As expected, there was no signal corresponding to ClC-1 (Fig. 2).
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Maxi chloride channels (p-VDAC, or VDACL) were initially cloned from the bovine brain (Dermietzel et al. 1994) and are functionally expressed in cultured rat cortical astrocytes (Jalonen, 1993; Parkerson & Sontheimer, 2004). p-VDACs are volume sensitive and have a large enough pore to conduct excitatory amino acids (Sabirov et al. 2001; Sabirov & Okada, 2004). To test for p-VDAC contribution to excitatory amino acid release, we used 30 µM Gd3+, which at this concentration discriminates between p-VDAC and VRAC (Sabirov et al. 2001). As seen in Fig. 3, Gd3+ had a trend to slightly stimulate both swelling-activated D-[3H]aspartate release (P= 0.219) and Cl– currents (P= 0.050).
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We have used 300 µM CdCl2 to probe for the potential involvement of the ClC-2 channel. ClC-2 is a broadly expressed Cl– channel, which is also regulated by cell swelling (Grunder et al. 1992), and is expressed in cultured astrocytes (Ferroni et al. 1997; Parkerson & Sontheimer, 2004). Cd2+ (300 µM) inhibits ClC-2 currents by 70% (Clark et al. 1998). However, in our experiments Cd2+ failed to inhibit either swelling-activated D-[3H]aspartate release (Fig. 4A, P= 0.134) or Cl– currents (Fig. 4B, P= 0.689).
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The ClC-3 channel has been cloned from rat brain, and is potently inhibited by activation of PKC (Kawasaki et al. 1994). The PKC activator PDBu strongly down-regulates swelling-activated Cl– currents in cardiomyocytes and pulmonary artery smooth muscle cells (Duan et al. 1999; Zhong et al. 2002). ClC-3 is expressed in cultured astrocytes (Parkerson & Sontheimer, 2004). In our hands, 500 nM PDBu had no effect on swelling activated Cl– currents (Fig. 5B, P= 0.727), and strongly up-regulated D-[3H]aspartate release (Fig. 5A, peak release 170.4 ± 11.6% of control, P < 0.001).
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Hypoosmotic media are known to elevate intracellular Ca2+ concentration in many cells (McCarty & O'Neil, 1992), which may trigger activation of calcium-sensitive Cl– channels. ATP and other Ca2+-mobilizing agents strongly potentiate swelling-induced release of organic osmolytes in astrocytes and several other cell types (Mongin & Kimelberg, 2002; Loveday et al. 2003; Franco et al. 2004), implying a potential contribution of calcium-sensitive Cl– channels. These channels are completely blocked by 100 µM niflumic acid (Large & Wang, 1996; Pedersen et al. 1998). In our experiments, 100 µM niflumic acid was ineffective against swelling-activated excitatory amino acid release (Fig. 6A, P= 0.585). When ATP was coapplied with hypoosmotic medium, it strongly potentiated D-[3H]aspartate release, which now was weakly sensitive to 100 µM niflumic acid (Fig. 6A, 22.5 ± 1.4% inhibition, P= 0.047). Swelling-activated Cl– currents were more sensitive to niflumic acid (Fig. 6B, 41.1 ± 2.5% inhibition, P < 0.001).
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The product of the multidrug resistance-1 gene, P-glycoprotein or MDR-1, was formerly proposed to be VRAC, but it is now thought to function only as a VRAC modulator (Valverde et al. 1992; Wine & Luckie, 1996). This protein is expressed in cultured astrocytes (Ronaldson et al. 2004). We used 100 µM verapamil, which at this concentration completely inhibits MDR-1 function (Luckie et al. 1994), to test for direct or indirect MDR-1 involvement in excitatory amino acid release. In our experiments, verapamil did not affect peak D-[3H]aspartate release (P= 0.402), but strongly inhibited the time-dependent inactivation of the release during exposure to hypoosmotic medium (Fig. 7A). In the corresponding electrophysiological study, 100 µM verapamil failed to affect swelling-activated Cl– currents (Fig. 7B, P= 0.323).
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As seen in Fig. 1B, application of hypotonic solution induced whole-cell Cl– currents with moderate outward rectification and inactivation at large positive potentials, which are typical for VRAC. To explore the contribution of VRAC activity to excitatory amino acid release, we tested a novel selective VRAC blocker, DCPIB. DCPIB inhibits VRAC currents in calf pulmonary artery endothelial cells (CPAE), guinea-pig atrial cardiomyocytes and Xenopus oocytes, but does not affect either endogenous or heterologously expressed ClC-1, ClC-2, ClC-4, ClC-5, CFTR, and calcium-activated Cl– channels (Decher et al. 2001). The IC50 for DCPIB inhibition of VRAC currents is 4.1 µM (Decher et al. 2001). In our experiments, 20 µM DCPIB blocked swelling-activated D-[3H]aspartate release by 76.0 ± 4.5% (Fig. 8A, P < 0.001). DCPIB was even more effective when hypoosmotic medium was coapplied with 20 µM ATP (Fig. 8A, 94.9 ± 1.7% inhibition, P < 0.001). Consistent with effects reported in the literature, 20 µM DCPIB nearly completely inhibited swelling-activated Cl– currents (Fig. 8B, 93.5 ± 1.7% inhibition, P < 0.001).
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80%, as originally shown by Fan et al. (2001) in T84, C127/CFTR and intestinal 407 cells.
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Discussion |
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An important finding of this study is the nearly complete inhibition of the swelling-activated organic osmolyte release by the selective VRAC blocker DCPIB. In a recent study by Decher et al. (2001), DCPIB inhibited swelling-activated Cl– currents, i.e. VRACs, in several cell types, while not affecting Cl– currents carried by either endogenous or heterologously expressed ClC-1, ClC-2, ClC-4, ClC-5, CFTR and calcium-activated Cl– channels. Thus, VRAC is likely to be the predominant pathway responsible for the volume-dependent release of organic osmolytes in cultured astrocytes. Consistent with such a conclusion, 10 mM extracellular ATP, which at high concentrations acts as a VRAC open channel blocker (Tsumura et al. 1996), potently suppresses swelling-activated excitatory amino acid release in cultured astrocytes (Mongin & Kimelberg, 2002; Haskew-Layton et al. 2005). Furthermore, 100 µM phloretin, which distinguishes VRAC from CFTR and Ca2+-activated Cl– channels (Fan et al. 2001), also strongly inhibits astrocytic amino acid release (Mongin & Kimelberg, 2002; Haskew-Layton et al. 2005; and Fig. 10 in the present study).
Although the data of Decher and colleagues on the specific inhibition of VRAC by DCPIB are very convincing, the selectivity of this compound has not yet been verified by other laboratories. Therefore, the information obtained with other Cl– channel blockers is useful in order to corroborate the DCPIB data and test for the possible contribution of VRAC-independent permeability pathways. In our study Gd3+ and Cd2+ did not affect swelling-activated excitatory amino acid release. Since Gd3+ and Cd2+ potently block p-VDAC and ClC-2, respectively (Sabirov et al. 2001; Clark et al. 1998), these results rule out involvement of these two volume-sensitive pathways. Contribution of MDR-1 to organic osmolyte release was excluded based on the absence of inhibiton by verapamil, which potently blocks MDR-1 activity (Luckie et al. 1994). Interestingly, verapamil markedly inhibited time-dependent inactivation of D-aspartate release in hypoosmotically swollen cells (see Fig. 6A). Such inactivation is thought to be due to a regulatory volume decrease (RVD). Verapamil blocks RVD in several cell types and may act via inhibition of Ca2+ influx or K+ conductance (McCarty & O'Neil, 1990; De Smet et al. 1998).
Unlike all the candidate channels mentioned above, ClC-3 does not have a selective inhibitor. The sensitivity of ClC-3 to DCPIB has not been tested. However, a unique feature of this channel is a strong inhibition by PKC. Phorbol esters, such as phorbol 12-myristate 13-acetate (PMA) and PDBu, eliminate ClC-3-mediated Cl– currents in osmotically swollen cells (Duan et al. 1997; Zhong et al. 2002). On the contrary, PDBu and PMA stimulate VRAC-mediated Cl– currents and swelling-activated organic osmolyte release at least in some cell types (Jackson & Strange, 1993; Loveday et al. 2003; Gong et al. 2004). In our experiments, 500 nM PDBu strongly potentiated swelling-activated organic osmolyte fluxes, thus making ClC-3 involvement unlikely. PDBu did not stimulate Cl– currents in our electrophysiological experiments. One possible explanation is that PDBu acts via one of the Ca2+-dependent PKC isoforms, which is not activated in the absence of Ca2+ in our pipette solution.
We next tested for the involvement of Ca2+-activated Cl– channels (CaCCs). Although CaCCs are not volume sensitive, we included them in a list of candidates because cell swelling increases intracellular Ca2+ and therefore may indirectly activate Ca2+-sensitive transport pathways. It has been also found that ATP and other Ca2+ mobilizing agonists strongly potentiate volume-sensitive efflux of 125I– and organic osmolytes in swollen cells (Tilly et al. 1994; Loveday et al. 2003; Franco et al. 2004). In cultured astrocytes, ATP potentiates hypoosmotic medium-induced D-[3H]aspartate release by 2- to 3-fold and this effect is completely dependent on intracellular Ca2+ and calmodulin (Mongin & Kimelberg, 2005b). It is possible that ATP activates a separate Ca2+-sensitive osmolyte release route in swollen cells, additional to VRAC. Lee et al. (2004) have recently reported in a preliminary form that ATP, bradykinin and other Ca2+-releasing agonists induce astrocytic glutamate release via CaCCs. In their work, CaCCs were completely blocked by 100 µM niflumic acid, as in a number of other studies (Large & Wang, 1996; Pedersen et al. 1998; Lee et al. 2004). In our hands, 100 µM niflumic acid produced negligible inhibition of swelling-activated excitatory amino acid release both in the absence and in the presence of extracellular ATP. These results seem to rule out the involvement of CaCCs.
Perhaps the most perplexing result of our study is the relatively weak inhibition of swelling-activated amino acid release by tamoxifen. Tamoxifen is a potent VRAC blocker, which is commonly used in electrophysiological studies. However, the sensitivity of VRAC to this compound varies among cell types, from high sensitivity in HEK293 and I-407 cells to partial inhibition in human cancer KB3 and rat RBL-2H3 cells, to complete insensitivity in cultured cortical neurons (Nilius et al. 1994; Inoue et al. 2005). In our experiments, 10 µM tamoxifen completely blocked swelling-activated Cl– currents but, surprisingly, was only partially effective (40% inhibition) against the volume-dependent D-[3H]aspartate release. Moreover, its inhibitory effect on organic osmolyte release required preincubation. In the cells, where swelling-activated excitatory amino acid release was potentiated by application of ATP, tamoxifen suppressed the release by 80%, approaching the efficacy of the selective VRAC blocker, DCPIB. Such differential sensitivity to tamoxifen may be explained by the indirect, calmodulin-dependent effects of this compound on VRAC, as originally suggested by Kirk & Kirk (1994). Consistent with such an idea, other calmodulin antagonists more potently block swelling-activated amino acid release in cells stimulated by ATP, where increases in intracellular Ca2+ up-regulate calmodulin-dependent processes (Mongin & Kimelberg, 2005b).
As seen in the summary Fig. 10, in cultured rat astrocytes swelling-activated Cl– currents and excitatory amino acid release share a similar pharmacological profile. In particular, the recently characterized selective VRAC blocker DCPIB nearly completely inhibited volume-dependent organic osmolyte release, as well as VRAC currents. The data obtained with other Cl– channel inhibitors are consistent with the idea of DCPIB selectivity, and rule out contributions of ClC-2, ClC-3, p-VDAC, CaCC, CFTR and MDR-1 to organic osmolyte release from swollen cells. Therefore we now propose that VRAC is a principal candidate pathway for mediating organic osmolyte release. Furthermore, DCPIB is a valuable tool to probe for VRAC contribution to physiological and pathological release of organic osmolytes in vivo and in vitro.
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Supplemental material |
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Supplemental Figure 1. Simultaneous measurements of D-[3H]aspartate and 36Cl– release from cultured astrocytes exposed to hypoosmotic medium
Supplemental Figure 2. VRAC blocker phloretin strongly inhibits swelling-activated D-[3H]aspartate release and Cl–currents
This material can also be found as part of the full-text HTML version available from http://www.blackwell-synergy.com
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) or absence (
) of 100 µM NPPB. Data are means ±S.E.M. of 5 experiments. **P= 0.007, effect NPPB versus control, repeated measures ANOVA. B, representative of 5 electrophysiological recordings of Cl– currents in single cultured astrocytes exposed to hypoosmotic medium. Cells were held at 0 mV and alternative pulses to ± 40 mV were applied to measure activation of Cl– currents. NPPB (100 µM) was applied after Cl– currents approached steady-state level. Inset: Cl– current responses to step pulses from –100 to +100 mV in 20 mV increments at the time point indicated by asterisk.
, •). Data are means ±S.E.M. of 5 experiments. NS, not significant (P= 0.074), niflumic acid versus control; *P= 0.04, ATP versus ATP plus niflumic acid, repeated measures ANOVA. B, effect of 100 µM niflumic acid (NA) on swelling-activated Cl– currents. Representative of 6 electrophysiological recordings.
