We reported previously that inhibition of Na+-K+-Cl− cotransporter isoform 1 (NKCC1) by bumetanide abolishes high extracellular K+concentration ([K+]o)-induced swelling and intracellular Cl− accumulation in rat cortical astrocytes. In this report, we extended our study by using cortical astrocytes from NKCC1-deficient (NKCC1−/−) mice. NKCC1 protein and activity were absent in NKCC1−/− astrocytes. [K+]o of 75 mM increased NKCC1 activity approximately fourfold in NKCC1+/+ cells (P< 0.05) but had no effect in NKCC1−/− astrocytes. Intracellular Cl− was increased by 70% in NKCC1+/+ astrocytes under 75 mM [K+]o (P < 0.05) but remained unchanged in NKCC1−/− astrocytes. Baseline intracellular Na+ concentration ([Na+]i) in NKCC1+/+ astrocytes was 19.0 ± 0.5 mM, compared with 16.9 ± 0.3 mM [Na+]i in NKCC1−/− astrocytes (P < 0.05). Relative cell volume of NKCC1+/+ astrocytes increased by 13 ± 2% in 75 mM [K+]o, compared with a value of 1.0 ± 0.5% in NKCC1−/− astrocytes (P < 0.05). Regulatory volume increase after hypertonic shrinkage was completely impaired in NKCC1−/− astrocytes. High-[K+]o-induced 14C-labeledd-aspartate release was reduced by ∼30% in NKCC1−/− astrocytes. Our study suggests that stimulation of NKCC1 is required for high-[K+]o-induced swelling, which contributes to glutamate release from astrocytes under high [K+]o.
- cell swelling
- high potassium ion concentration
- cultured astrocytes
- glutamate release
- intracellular chloride
- excitatory amino acid
na+ -k+-cl−cotransporter (NKCC) isoform 1 (NKCC1) has been shown to play a role in cell volume regulation and K+ uptake in astrocytes (10, 12, 33). NKCC1 activity in cultured rat cortical astrocytes is significantly stimulated by high extracellular K+ concentration ([K+]o) (32). Inhibition of NKCC1 activity by the potent cotransporter inhibitor bumetanide decreases the high-[K+]o-mediated uptake of Cl− and blocks astrocyte swelling (Ref. 33; this issue). Moreover, administration of bumetanide in brain cortex significantly reduces edema and infarct volume after focal ischemia (41). Thus NKCC1 could contribute to an overload of intracellular Na+, K+, and Cl− and cell swelling in conditions such as cerebral ischemia, in which [K+]o is elevated.
Astrocytes release organic osmolytes such as excitatory amino acid (EAA) under high-[K+]o conditions to regulate cell volume (13, 18). One possible mechanism for the EAA release under high-[K+]o conditions is via volume-sensitive organic anion channels (VSOACs; Refs. 1,27). High-[K+]o-induced3H-labeled d-aspartate (Asp) release from cultured astrocytes is inhibited by the anion channel inhibitors L-644711 and dideoxyforskolin (27). We have reported (Ref.33; this issue) that blocking of NKCC1 activity by bumetanide significantly reduces high-[K+]o-induced 14C-labeledd-Asp release. These results imply that NKCC1-mediated swelling may contribute to the high-[K+]o-triggered glutamate release.
The role of NKCC1 in high-[K+]o-triggered swelling and glutamate release described above has been studied by pharmacological techniques. Although bumetanide is a potent inhibitor of the NKCCs, it also inhibits other ion transport proteins and GABAA channels (26, 34). In this study, we used astrocytes isolated from the NKCC1-deficient mice established by Flagella et al.(6). We report here that high-[K+]o-induced swelling was absent in NKCC1−/− astrocytes. A significant decrease in intracellular Cl− and Na+ and release of glutamate was observed in NKCC1−/− astrocytes. The results suggest that NKCC1 is important in regulation of astrocyte volume and intracellular Cl− and Na+.
MATERIALS AND METHODS
Bumetanide, digitonin, Triton X-100, monensin, and gramicidin were purchased from Sigma (St. Louis, MO). Eagle's modified essential medium (EMEM) and Hanks' balanced salt solution (HBSS) were from Mediatech Cellgro (Herndon, VA). Fetal bovine serum was obtained from Hyclone Laboratories (Logan, UT). Collagen type I was from Collaborative Biomedical Products (Bedford, MA). 86RbCl was purchased from NEN Life Science Products (Boston, MA).d-[14C]Asp was from American Radiolabeled Chemicals (St. Louis, MO). Chloride-36 was purchased from Amersham Pharmacia Biotech (Piscataway, NJ). Sodium-binding benzofuran isophthalate (SBFI)-AM was purchased from Molecular Probes (Eugene, OR). Pluronic acid was purchased from BASF (Ludwigshafen, Germany).
Animals and genotype analysis.
NKCC1 homozygous mutant and wild-type mice were obtained by breeding gene-targeted NKCC1 heterozygous mutant mice, and genotypes were determined by a polymerase chain reaction (PCR) of DNA from tail biopsies as described previously (6).
Primary culture of mouse cortical astrocytes.
Dissociated cortical astrocyte cultures were established based on a method used in our previous study of rat cortical astrocyte cultures (32). Cerebral cortices were removed from 1-day-old NKCC1+/+ or NKCC1−/− mice. The cortices were incubated in a solution of 0.25 mg trypsin /ml HBSS for 25 min at 37°C. The tissue was then mechanically triturated and filtered through nylon meshes. The dissociated cells were rinsed and resuspended in EMEM containing 10% fetal bovine serum. Viable cells (1 × 104 cells /well) were plated in 24-well plates coated with collagen type 1. Cultures were maintained in a 5% CO2atmosphere at 37°C. The cultures were subsequently refed every 3 days throughout the study. To obtain morphologically differentiated astrocytes, confluent cultures (days 12–15 in culture) were treated with EMEM containing 0.25 mM dibutyryl cAMP (DBcAMP) for 7 days to induce differentiation. DBcAMP has been widely used to mimic neuronal influences on astrocyte differentiation (9, 35). Experiments were routinely performed (see Figs. 4-10) on cultures treated with DBcAMP for 7 days.
Gel electrophoresis and Western blotting.
Cortical astrocytes growing on culture dishes were washed with ice-cold phosphate-buffered saline (PBS; pH 7.4) that contained 2 mM EDTA and protease inhibitors. Cells were scraped from dishes and lysed by sonication at 4°C (32). To obtain cellular lysates, cellular debris was removed by centrifugation and protein content of the cellular lysate was determined (29). Samples and prestained molecular mass markers (Bio-Rad) were denatured in SDS reducing buffer (1:2 vol/vol; Bio-Rad) and heated at 37°C for 15 min before gel electrophoresis. The samples were then electrophoretically separated on 6% SDS gels, and the resolved proteins were electrophoretically transferred to a polyvinylidene difluoride membrane (32). The blots were incubated in 7.5% nonfat dry milk in Tris-buffered saline and then incubated with a primary antibody. After rinsing, the blots were incubated with horseradish peroxidase-conjugated secondary IgG. Bound antibody was visualized with an enhanced chemiluminescence assay (ECL; Amersham). Anti-NKCC monoclonal antibody against the human colonic T84 epithelial NKCC (20) was used for detection of the cotransporter protein, and the same blot was probed with anti-β actin antibody as a control. A linear curve with 5–30 μg of protein has been established for both anti-NKCC antibody and anti-β actin antibody (32). In addition, the linear range of the ECL exposure course on the film was within 15–120 s (32). Therefore, 30 μg of protein was loaded in all immunoblots of the present study. In addition, the ECL exposure time was within 60 s.
Cultured cells grown on collagen type I-coated coverslips were rinsed with PBS (pH 7.4) and fixed with 4% paraformaldehyde in PBS for 40 min at room temperature. After rinsing, cells were incubated with blocking solution (10% normal goat serum, 0.4% Triton X-100, and 1% bovine serum albumin in PBS) for 1 h. Cells were then incubated with anti-glial fibrillary acidic protein (GFAP) polyclonal (1:100) or anti-NKCC monoclonal antibodies (1:100) in blocking solution overnight at 4°C. Cells were rinsed with PBS and incubated with goat anti-mouse FITC-conjugated or goat anti-rabbit Texas red-conjugated antibodies (1:200) for 2 h. Cell images were captured by laser scanning confocal microscope (Bio-Rad MRC 1000) as described previously (32). Bio-Rad MRC-1024 Laser Sharp software (version 2.1T) was used to control the microscope and its settings. An identical setting was used to capture the negative control and experimental images.
Measurement of relative cell volume changes in single cell.
Relative cell volume changes were estimated with video-enhanced differential interference contrast (DIC) microscopy, as described in our previous study (Ref. 33, this issue). The same method has been reported by others (40). Astrocytes cultured on collagen-coated coverslips were placed in an open-bath imaging chamber (Warner Instruments, Hamden, CT; bath volume 40 μl) on the stage of a Nikon TE 300 inverted epifluorescence microscope. Astrocytes were equilibrated with isotonic HEPES-buffered minimal essential medium (MEM, 312 mosmol/kgH2O) for 15 min (33). Astrocytes were exposed sequentially to HEPES-MEM (5 min), 75 mM [K+]o HEPES-MEM (10 min), HEPES-MEM (10 min), HEPES-MEM + 10 μM bumetanide (20 min), 75 mM [K+]o HEPES-MEM + 10 μM bumetanide (10 min), and HEPES-MEM (10 min). In 75 mM [K+]o HEPES-MEM, 75 mM [K+]o was obtained by replacing NaCl in HEPES-MEM solutions with equimolar KCl. A single astrocyte was visualized with a Nikon ×60 Plan Apo oil-immersion objective lens, and cell images were recorded every minute as described previously (33). The mean cross-sectional area (CSA) of the cell body was calculated with MetaMorph image-processing software. The control CSA values were obtained when cells were exposed to HEPES-MEM only. Relative changes in CSA (CSAr) were calculated as experimental CSA divided by control CSA. After each experiment, relative cell volume changes in response to HEPES-MEM calibration buffers were measured. Salt concentrations in the buffers were held constant, and the osmolality (238, 277, and 312 mosmol/kgH2O) was adjusted by varying the buffer concentration of sucrose.
Assay for NKCC1 activity.
NKCC1 activity was measured as bumetanide-sensitive K+influx with 86Rb as a tracer for K+(32). Cultured astrocytes were equilibrated for 10–30 min at 37°C with isotonic HEPES-MEM (312 mosmol/kgH2O). Cells were preincubated for 10 min in HEPES-MEM containing either 0 or 10 μM bumetanide. For assay of cotransporter activity, cells were exposed to 1 μCi/ml of 86Rb in HEPES-MEM for 3 min in the presence or absence of 10 μM bumetanide. 86Rb influx was stopped by rinsing cells with ice-cold 0.1 M MgCl2. Radioactivity of the cellular extract in 1% SDS was analyzed by liquid scintillation counting (1900CA; Packard, Downers Grove, IL). K+ influx rate was calculated and expressed as nanomoles of K+ per milligram of protein per minute. It has been established that the slope of 86Rb uptake over 10 min is linear in astrocytes (32). Bumetanide-sensitive K+ influx was obtained by subtracting the K+influx rate in the presence of bumetanide from the total K+influx rate. Quadruplicate determinations were obtained in each experiment throughout the study, and protein content was measured in each sample with a method described previously (29). Statistical significance in the study was determined by ANOVA (Bonferroni-Dunn) at a confidence level of 95% (P < 0.05).
Intracellular Cl− content measurement.
Cells on 24-well plates were preincubated for 30 min in HEPES-MEM containing 5.8 mM [K+]o and 36Cl (0.4 μCi/ml). A steady-state level of intracellular 36Cl was established and maintained during the 30-min preincubation (33). The cells were then incubated in 75 mM [K+]o HEPES-MEM containing 36Cl (0.4 μCi/ml) in the presence or absence of 10 μM bumetanide for 4 or 13 min. Thus extracellular Cl− concentration ([Cl−]o) of 145 mM in HEPES-MEM was maintained in 75 mM [K+]o, and the specific activity of 36Cl was constant in 5.8 mM [K+]o and 75 mM [K+]o HEPES-MEM. Intracellular36Cl content measurement was terminated by three washes with 1 ml of ice-cold washing buffer (in mM: 118 NaCl, 26 NaHCO3, 1.8 CaCl2, pH 7.40). Radioactivity of the cellular extract in 1% SDS was analyzed by liquid scintillation counting (Packard 1900CA). In each experiment, specific activities (counts/μmol × min) of 36Cl were determined for each assay condition and used to calculate intracellular Cl− content (μmol/mg protein).
Intracellular Na+ measurement.
Intracellular Na+ concentration ([Na+]i) was measured with the fluorescent dye SBFI-AM as described previously (25). Cultured astrocytes grown on collagen-coated coverslips were loaded with 10 μM SBFI-AM at room temperature in HEPES-MEM containing 0.1% pluronic acid as described in our previous report (Ref. 33, this issue). The coverslips were placed in an open-bath imaging chamber containing HEPES-MEM at ambient temperature. With a Nikon TE 300 inverted epifluorescence microscope and a ×40 Super Fluor oil-immersion objective lens, astrocytes were excited every 10 s at 345 and 385 nm, and the emission fluorescences at 510 nm were recorded. Images were collected and analyzed with MetaFluor image-processing software (33). An area on the coverslip without cells was defined as the background region and used for subtraction of baseline fluorometric intensities at 345 and 385 nm. Approximately 65% of the SBFI fluorescence signals (340- to 380-nm ratios) measured in this study represented changes of Na+in the cytoplasm of astrocytes (33).
To monitor changes of [Na+]i, the SBFI-loaded cells were equilibrated with HEPES-MEM for 20 min. Ratios of 340- to 380-nm fluorescence were recorded, and the bath chamber buffer was changed with 75 mM [K+]o HEPES-MEM (10 min) followed by HEPES-MEM (10 min). Absolute [Na+]i was determined for each cell by calibrating the SBFI fluorescence ratio with solutions containing 0, 10, 20, 40 or 80 mM extracellular Na+ concentration ([Na+]o) and monensin (10 μM) and gramicidin (3 μM) to equilibrate [Na+]o and [Na+]i. The resulting ratios from each cell were fit with a three-parameter hyperbolic curve from which [Na+]i was calculated (4).
d-[14C]Asp release measurement.
Aspartate release was measured as described previously (Ref.33, this issue). Astrocytes grown on chamber slides were incubated overnight in 1 ml of complete EMEM containing 2 μCi/ml of d-[14C]Asp (specific activity of 55 mCi/mmol). The perfusing rate of the perfusion chamber was 1.5 ml/min. This chamber allows a complete change of the perfusing buffer within 2 min. The cells were perfused at a constant flow rate with HEPES-MEM containing 5.8 or 75 mM [K+]o in the presence or absence of 10 μM bumetanide. The buffers and perfusion chamber were kept at 37°C. The perfusate was collected in 1-min intervals. At the end of the experiment, the cells were digested in 1% SDS. The radioactivity of samples was measured by liquid scintillation counting (Packard 1900CA). Calculation of fractional release was based on the following formula: fractional release =Ct /[Sum(Ct :C end) + C remain], where Ct is the cpm value in the effluent at time t,C end is the cpm value in the effluent at the end of the experiment, Sum(Ct :C end) is the total cpm value in the effluent from time t to the end of the experiment (28), and C remainis the cpm value left in the cells at the end of the experiment.
Absence of NKCC1 protein expression in cultured cortical astrocytes isolated from NKCC1−/− mice.
Mating of heterozygous mice yielded live offspring of all three genotypes in a Mendelian ratio of 1:2:1 (23% NKCC1+/+, 54% NKCC1+/−, 23% NKCC1−/−). As shown in Fig. 1 A, a single DNA band (∼105 bp) was detected in a tail biopsy of the NKCC1+/+mouse. In contrast, a larger DNA band (∼156 bp) was found in the NKCC1−/− tail biopsy. In a heterozygous sample, both PCR products were present.
To analyze expression of the NKCC1 protein in cultured NKCC1−/− cortical astrocytes, cellular lysates were obtained from both cultured NKCC1+/+and NKCC1−/− astrocytes. Immunoblotting revealed that an ∼152 ± 5.7 kDa protein was recognized by an anti-NKCC antibody in undifferentiated (−DBcAMP) and differentiated (+DBcAMP) NKCC1+/+ astrocytes (Fig. 1 B). In contrast, no protein band was detected by the anti-NKCC antibody in either undifferentiated or differentiated NKCC1−/− astrocytes. The lack of NKCC1 bands was not due to an insufficient amount of protein having been loaded, because, as shown on the same blot, a similar amount of β-actin protein (∼58 ± 5.2 kDa) was observed in samples from both NKCC1+/+ and NKCC1−/− astrocytes (Fig. 1 B). These results indicate that NKCC1 is absent in NKCC1−/− astrocytes. Further support is provided by the results of the immunofluorescence study. As shown in Fig. 2, Aand D, both undifferentiated and differentiated NKCC1+/+ astrocytes were positively stained for the glial marker GFAP. Anti-NKCC antibody immunoreactive fluorescence signals were observed in both undifferentiated and differentiated NKCC1+/+ astrocytes (Fig. 2, B andE). Colocalization of GFAP and the cotransporter is shown in Fig. 2, C and F. This is consistent with our previous findings in cultured rat cortical astrocytes (32). In NKCC1−/− astrocytes, DBcAMP induced differentiation in a similar fashion as in NKCC1+/+astrocytes (Fig. 2, a and d). Expression of GFAP was shown in both undifferentiated and differentiated NKCC1−/− astrocytes (Fig. 2, a andd). However, no immunoreactive signals were detected in NKCC1−/− astrocytes with the anti-NKCC antibody (Fig. 2,b and e). Only GFAP signals appeared in a double-staining image (Fig. 2, c and f). Together, these results demonstrate that the NKCC1 protein is not expressed in undifferentiated and differentiated NKCC1−/−astrocytes.
Decrease in K+ influx in NKCC1−/− astrocytes under high [K+]o.
Previous studies suggested that NKCC is important for K+uptake in astrocytes and that cotransporter activity is significantly stimulated by high [K+]o (32,37). In this study, we first examined whether the K+influx rate was different in NKCC1+/+and NKCC1−/− astrocytes (undifferentiated) under 5.8 mM [K+]o. As shown in Fig.3 A, at 5.8 mM [K+]o the total K+ influx rate was 111.8 ± 6.8 nmol/mg protein × min in NKCC1+/+ astrocytes; however, it was decreased to 86.1 ± 4.8 nmol/mg protein × min in NKCC1−/− astrocytes (P > 0.05). 10 μM bumetanide did not significantly affect the K+ influx rate in either NKCC1+/+ or NKCC1−/− astrocytes under 5.8 mM [K+]o. When NKCC1+/+ astrocytes were exposed to 75 mM [K+]o, the total K+ influx rate was increased to 348.7 ± 25.8 nmol/mg protein × min (P < 0.05). Inhibition of cotransporter activity by 10 μM bumetanide decreased the total K+ influx rate by 20% (P < 0.05). Interestingly, genetic ablation of NKCC1 caused a similar reduction in the total K+ influx rate under 75 mM [K+]o. Moreover, no additional reduction of total K+ influx was observed when NKCC1−/−astrocytes were treated with 10 μM bumetanide (Fig. 3 A). An identical pattern of changes in the total K+ influx was found in differentiated NKCC1+/+ and NKCC1−/−astrocytes (data not shown). These data suggest that the cotransporter is important in K+ influx in astrocytes, particularly under high-[K+]o conditions.
The inhibitory effect of bumetanide on the total K+ influx rate implies that cotransporter activity was increased in NKCC1+/+ astrocytes under high [K+]o. We next examined the bumetanide-sensitive K+ influx rate in NKCC1+/+and NKCC1−/− astrocytes. Figure3 B shows that high [K+]ostimulated the bumetanide-sensitive K+ influx in both undifferentiated and differentiated NKCC1+/+ astrocytes. In contrast, bumetanide-sensitive K+ influx in NKCC1−/− astrocytes was within experimental variation under 5.8 mM [K+]o. It was not changed under high-[K+]o conditions. This is in agreement with the finding that bumetanide has no effect on the total K+ influx rate in NKCC1−/− astrocytes.
Decrease in 36Cl− accumulation in NKCC1−/− astrocytes under high [K+]o.
Pharmacological inhibition of NKCC suggested that the cotransporter contributes to an accumulation of intracellular Cl− in astrocytes (15, 33). We investigated this further with NKCC1-deficient astrocytes. The basal levels of intracellular36Cl− were 0.48 ± 0.04 μmol/mg protein when NKCC1+/+ astrocytes were exposed to 5.8 mM [K+]o for 4 min (Fig.4 A). A similar level of intracellular 36Cl was maintained after the cells were incubated in 5.8 mM [K+]o for 13 min. The intracellular 36Cl level was increased to 0.81 ± 0.06 μmol/mg protein when NKCC1+/+ astrocytes were exposed to 75 mM [K+]o for 4 min (P < 0.05) and maintained at 0.79 ± 0.05 μmol/mg protein after a 13-min incubation. Blocking of cotransporter activity by 10 μM bumetanide significantly decreased the high-[K+]o-induced 36Cl increase in NKCC1+/+ astrocytes (P < 0.05; Fig.4 A). As shown in Fig. 4 B, the basal36Cl levels in NKCC1−/− astrocytes were not significantly different from those of NKCC1+/+ astrocytes under 5.8 mM [K+]o (P > 0.05). However, when NKCC1−/− astrocytes were exposed to 75 mM [K+]o for 4 min, no significant increase in 36Cl accumulation was observed (P > 0.05; Fig. 4 B). After a 13-min incubation with 75 mM [K+]o, the intracellular 36Cl level in the NKCC1−/−astrocytes was slightly increased but still lower than in control NKCC1+/+ astrocytes and insensitive to bumetanide. Moreover, 10 μM bumetanide did not significantly affect the intracellular 36Cl levels in NKCC1−/−astrocytes under either 5.8 or 75 mM [K+]o(P > 0.05). Collectively, the results show that the absence of NKCC1 in cortical astrocytes abolished the high-[K+]o-mediated accumulation of intracellular 36Cl.
Decrease in intracellular Na+ in NKCC1−/− astrocytes.
We previously observed that pharmacological inhibition of NKCC1 in rat cortical astrocytes resulted in a decrease in basal levels of [Na+]i by ∼2 mM (33). In the current study, the basal levels of [Na+]i were 19.0 ± 0.5 mM in NKCC1+/+ astrocytes. In contrast, [Na+]i was 16.9 ± 0.3 mM in NKCC1−/− astrocytes (P < 0.001; Fig.5, A and B). In response to high [K+]o, a compensatory decrease in [Na+]i occurred in both NKCC1+/+ and NKCC1−/− astrocytes (Fig. 5,A and B), which has been suggested to be attributable in part to activation of Na+-K+-ATPase (19). [Na+]i in NKCC1+/+astrocytes recovered to basal levels after cells were returned to 5.8 mM [K+]o. However, in NKCC1−/−astrocytes, [Na+]i remained significantly lower than in NKCC1+/+ astrocytes. Thus the results suggest that NKCC is important in maintaining intracellular Na+under control conditions.
Lack of high-[K+]o-induced swelling in NKCC1−/− astrocytes.
A significant decrease in intracellular Cl− and K+ uptake in NKCC1−/− astrocytes under high [K+]o implies that ablation of NKCC1 may prevent astrocyte swelling. Thus in the following experiments, relative volume changes were examined in both NKCC1+/+ and NKCC1−/− astrocytes. As shown in Fig.6 A, when NKCC1+/+astrocytes were exposed to 75 mM [K+]o, CSAr reached a maximum value of 1.13 ± 0.02. Inhibition of NKCC1 activity by 10 μM bumetanide abolished the high-[K+]o-induced swelling. Ten micromolar bumetanide had no significant effect on the basal level of CSAr (Fig. 6 A). In contrast, no cell swelling was observed when NKCC1−/− astrocytes were exposed to high [K+]o (Fig. 6 B). Ten micromolar bumetanide did not affect CSAr in NKCC1−/− astrocytes under either control or high-[K+]o conditions (Fig. 6 B).
Pharmacological inhibition and genetic ablation of NKCC1 have similar effects on astrocyte swelling. This indicates that NKCC1 is essential in astrocyte volume regulation. Therefore, we further investigated how NKCC1−/− astrocytes respond to osmotic stress. First, we evaluated regulatory volume increase (RVI) in both NKCC1+/+and NKCC1−/− astrocytes. As shown in Fig.7 A, when NKCC1+/+astrocytes were exposed to hypertonic HEPES-MEM (367 mosmol/kgH2O), CSAr decreased quickly and was 0.92 ± 0.01 at maximum cell shrinkage. CSAr in NKCC1+/+ recovered by 63.0 ± 4.4% at 20 min of hypertonic exposure (Fig. 7, A and B,inset). As NKCC1+/+ astrocytes were returned to an isotonic HEPES-MEM (310 mosmol/kgH2O), CSArwas further increased, followed by a secondary regulatory volume decrease (RVD; Fig. 7 A). As shown in Fig. 7 B, CSAr in NKCC1−/− astrocytes decreased in response to the hypertonic challenge. At maximum cell shrinkage, CSAr was 0.93 ± 0.01 (Fig. 7 B). Moreover, RVI was impaired in NKCC1−/− astrocytes (Fig.7 B). Overall, there was only 4.3 ± 1.7% RVI in NKCC1−/− astrocytes at 20 min of hypertonic exposure (Fig. 7 B, inset). In addition, after returning to isotonic conditions, NKCC1−/− astrocytes remained shrunken at a CSAr of 0.96 ± 0.01, which was significantly lower than basal levels (P < 0.01). These results indicate that the NKCC1 is essential for RVI in mouse cortical astrocytes.
We next examined RVD in NKCC1+/+ and NKCC1−/−astrocytes in response to a hypotonic challenge. When NKCC1+/+ astrocytes were exposed to hypotonic HEPES-MEM (247 mosmol/kgH2O), CSAr increased and reached a maximum level of 1.11 ± 0.02 (Fig.8 A). Cell volume subsequently decreased, and a RVD of 48.5 ± 5.0% developed after 20 min of hypotonic exposure (Fig. 8, A and B,inset). In NKCC1−/− astrocytes, cell swelling was not significantly different from that of NKCC1+/+astrocytes (CSAr = 1.08 ± 0.01 in NKCC1−/− vs. 1.11 ± 0.02 in NKCC1+/+;P > 0.05). Interestingly, the rate of RVD in NKCC1−/− astrocytes was faster (CSAr = 0.002%/min in NKCC1+/+ vs. 0.009%/min in NKCC1−/− astrocytes; P < 0.05). After 20 min of hypotonic exposure, an RVD of 86.6 ± 8.3% was obtained, which was significantly higher than the 48.5% in NKCC1+/+astrocytes (Fig. 8 B, inset). This suggests that other ion extrusion systems may be upregulated in NKCC1−/− astrocytes.
Decrease in high-[K+]o-induced release of d-[14C]Asp in NKCC1−/−astrocytes.
We previously observed that NKCC1 activity in rat astrocytes contributes to high-[K+]o-induced swelling and glutamate release (33). If the high-[K+]o-induced glutamate release is due in part to swelling, we anticipated that preventing cell swelling by inhibition of NKCC1 activity would reduce glutamate release. As shown in Fig. 9 A, in NKCC1+/+ astrocytes there was a trace amount ofd-[14C]Asp release under 5.8 mM [K+]o in the presence or absence of bumetanide (data not shown in the latter case). A small increase in the release of d-[14C]Asp occurred when cells were exposed to 75 mM [K+]o (phase I). The release was further developed with time and reached a peak value after 20 min (phase II). On removal of the high-[K+]o medium, the release returned to the resting level within 10 min. In the presence of 10 μM bumetanide, the peak value of the d-[14C]Asp release under 75 mM [K+]o was 0.65 ± 0.12% (fractional release per minute; Fig. 9 A). However, in the absence of bumetanide, high [K+]oinduced substantially more d-[14C]Asp release (P < 0.05; Fig. 9 A). In summary, 10 μM bumetanide has no effect on phase I ofd-[14C]Asp release; however, it blocked ∼30% of the phase II release ofd-[14C]Asp under high [K+]o (n = 12,P < 0.05; Fig. 9 A and inset). This is consistent with our finding in rat cortical astrocytes (Ref.33, this issue).
The effect of bumetanide implies that NKCC1 may contribute tod-[14C]Asp release under high-[K+]o conditions. To investigate this further, we examined d-[14C]Asp release in NKCC1−/− astrocytes. As shown in Fig. 9 B, similar to the results in NKCC1+/+ astrocytes, high [K+]o triggered a small increase ind-[14C]Asp release in NKCC1−/−astrocytes (phase I). The peak release ofd-[14C]Asp in NKCC1−/−astrocytes developed slowly and was smaller than in NKCC1+/+ astrocytes (Fig. 9 B). Compared with the value in NKCC1+/+ astrocytes, the peak release was ∼30% less in NKCC1−/− astrocytes (P < 0.05; Fig. 9 B and inset). This is in agreement with the bumetanide-mediated effect in NKCC1+/+ astrocytes. Together, the results suggest that NKCC1 contributes to the high-[K+]o-induced glutamate release.
We hypothesized that high-[K+]o-inducedd-[14C]Asp release (phase II) is mediated by volume-sensitive Cl− channels because it has a characteristic delayed onset rate and is partially blocked by inhibition of NKCC1 activity. If this hypothesis is correct, blocking of Cl− channels should prevent high-[K+]o-inducedd-[14C]Asp release. Therefore, we tested whether blocking of Cl− channels with 100 μM 4,4′-diisothiocyanostilbene-2,2′-disulfonic acid (DIDS), a broad-spectrum Cl− channel blocker, could reduced-[14C]Asp release. In NKCC1+/+astrocytes, as shown in Fig.10 A, a trace level ofd-[14C]Asp release was found under control conditions (0.16 ± 0.02%) and DIDS did not affect the basal levels of release. In the absence of DIDS, the two-phase release ofd-[14C]Asp was the same as that described above. In the presence of DIDS, there was no change in phase I release; however, the peak release of phase II was blocked by DIDS. In the DIDS-treated group, the peak release ofd-[14C]Asp during phase II was not significantly different from basal levels (0.25 ± 0.04 vs. 0.16 ± 0.02% fractional release per minute, n = 6–12; Fig. 10 A and inset). In NKCC1−/− astrocytes, DIDS did not significantly affect the phase I release of d-[14C]Asp (0.13 ± 0.01 vs. 0.17 ± 0.02% fractional release per minute; P < 0.05). However, the remaining release during phase II was abolished in the presence of DIDS (Fig.10 B and inset). Our data imply that phase II of d-[14C]Asp release is largely mediated by Cl− channels.
We have demonstrated here that pharmacological inhibition of NKCC1 function or genetic ablation of the NKCC1 protein abolished high-[K+]o-induced swelling, but it only blocked 30% of d-[14C]Asp release. This suggests that, in addition to swelling, other signals under high-[K+]o conditions may also stimulate Cl− channels. Exposing cells to high [K+]o leads to an increase in intracellular Ca2+ in rat and mouse astrocytes (21, 28, 32,42). Preventing Ca2+ influx by exposing cells to Ca2+-free HEPES-MEM inhibited the Ca2+ rise in rat astrocytes (28, 32). However, astrocytes in situ may use the mechanisms other than voltage-dependent Ca2+channels to regulate intracellular Ca2+ signals (Ref.2; see discussion). To test whether the Cl− channels could be activated via a Ca2+-mediated pathway and subsequently lead to the release of d-[14C]Asp, we evaluated whether removal of extracellular Ca2+ could block high-[K+]o-inducedd-[14C]Asp release. As shown in Table1, 75 mM [K+]otriggered a sevenfold increase in the peak release ofd-[14C]Asp during phase II release in NKCC1+/+ astrocytes. In NKCC1−/−astrocytes, only a fourfold increase was observed (P < 0.05). Interestingly, in Ca2+-free HEPES-MEM, 75 mM [K+]o failed to induced-[14C]Asp release in either NKCC1+/+ or NKCC1−/− astrocytes (Table 1). The results further support our hypothesis that Cl−channels could be stimulated in a Ca2+-dependent fashion under high [K+]o and result in the glutamate release. Cell swelling-mediated stimulation of the channels only partially contributes to the high-[K+]o-mediated glutamate release.
Role for NKCC1 in regulation of intracellular Na+, K+, and Cl− in astrocytes.
Previous pharmacological studies suggested that NKCC1 plays a role in maintenance of intracellular Na+ and Cl−(25, 26). Inhibition of NKCC1 by bumetanide in rat hippocampal (25) or cortical (33) astrocytes reduces the basal levels of intracellular Na+ by ∼2 mM. Consistent with these studies, we observed a reduction of 2.1 mM in basal [Na+]i in NKCC1-deficient cortical astrocytes. The collective results further support the notion that the steady state of Na+ influx via NKCC1 is important in maintenance of resting [Na+]i in astrocytes. We have also examined whether deletion of NKCC1 activity would affect basal levels of K+ and Cl− influx in NKCC1−/− astrocytes. Unlike [Na+]i, our data indicate that the basal levels of K+ influx and accumulation of intracellular36Cl were not significantly different in NKCC1−/− and NKCC1+/+ cortical astrocytes. This implies that a role for NKCC1 in maintenance of resting intracellular levels of K+ and Cl− in astrocytes is negligible. It is possible that the intracellular K+ concentration is maintained by mechanisms such as Na+-K+-ATPase and K+ channels, whereas intracellular Cl− could be regulated by other ion transport pathways, including Cl−/HCO exchangers or Cl− channels.
NKCC1 has been suggested to play a role in K+ uptake in astrocytes under elevated [K+]o (10,11, 37). Inhibition of NKCC1 in astrocytes by furosemide or low [Cl−]o significantly reduced K+uptake (39) and spontaneous epileptiform activity in rat hippocampal slices (12). We found previously (32) that the bumetanide-sensitive K+ influx rate was significantly increased in rat cortical astrocytes at 75 mM [K+]o. In the current report, we demonstrate that 1) the total K+ influx rate is increased in NKCC1+/+ cortical astrocytes in the presence of 75 mM [K+]o; 2) inhibition of NKCC1 activity by bumetanide decreases the total K+ influx rate by 20%; and 3) ablation of NKCC1 in cortical astrocytes causes a similar reduction in total K+ influx rate. Moreover, the bumetanide-mediated effect on the total K+influx rate is absent in NKCC1−/− cortical astrocytes. Together, the results of our pharmacological and knockout studies clearly suggest that NKCC1 is important in K+ uptake in astrocytes under high-[K+]o conditions. However, other K+ uptake pathways also play a major role because 80% of the total K+ uptake rate in astrocytes was not sensitive to bumetanide or genetic ablation of NKCC1. In addition, we found that either inhibition of NKCC1 with bumetanide or genetic ablation of NKCC1 blocks 33% of the high-[K+]o-mediated Cl−accumulation in cortical astrocytes. NKCC1 appears to have a greater effect on Cl− uptake than on K+ uptake in astrocytes, consistent with the 2:1 stoichiometry for Cl−and K+. These results support a role for NKCC1 in mediating the net gain of both K+ and Cl− in astrocytes as [K+]o is elevated.
Role for NKCC1 in regulation of astrocyte volume.
NKCC1 is important in cell volume regulation in a variety of mammalian cell types (24, 26). Inhibition of NKCC1 by bumetanide leads to cell shrinkage under isotonic conditions in cells such as retinal pigment epithelia, ventricular myocytes, and vascular endothelial cells (26). However, bumetanide has no effect on the basal volume of C6 glioma and vascular smooth muscle cells (26). In our study, neither inhibition of NKCC1 activity by bumetanide nor genetic ablation of NKCC1 affected the baseline volume in astrocytes. One possible explanation for the lack of an effect is that NKCC1 is not essential for maintenance of cell volume in mouse cortical astrocytes. Other mechanisms, such as the coupled activities of Na+/H+ and Cl−/HCO exchangers, which are expressed in astrocytes (3), may be able to maintain cell volume under control conditions.
NKCC1 is one of the principal volume regulatory transporters in RVI (24, 26). However, contradictory results about RVI have been observed in rat astrocytes (24). No RVI was observed when cultured rat cortical astrocytes were exposed to a hypertonic medium made with mannitol or NaCl (17, 23). However, rat astrocytes shrank and exhibited RVI in hypertonic buffer made with sucrose (5). The basis of the lack of RVI in rat astrocytes in these studies is unclear, although it could be attributable to intracellular ion concentrations such as Cl− [see review by O'Neill (24)]. To our knowledge, RVI in mouse cortical astrocytes has not been extensively examined before. In the present study, NKCC1+/+ mouse cortical astrocytes underwent RVI within 10 min after a hypertonic shrinkage. Cell volume was completely recovered as NKCC1+/+cortical astrocytes were returned to isotonic conditions. In contrast, RVI was absent in NKCC1−/− cortical astrocytes. In addition, on returning cells to the isotonic solutions, cell volume of NKCC1−/− cortical astrocytes failed to recover to basal levels. This is the first study to firmly establish that hypertonic RVI occurs in mouse cortical astrocytes and that the absence of NKCC1 completely impairs the RVI function in mouse cortical astrocytes.
In addition to RVI, NKCC1 is also involved in a secondary RVI when cells are returned to isotonic conditions after hypotonic cell swelling (post-RVD RVI; Ref. 8). NKCC1+/+cortical astrocytes exhibited RVD and post-RVD RVI. Interestingly, in NKCC1−/− cortical astrocytes, post-RVD RVI occurred but at significantly slower rates (Fig. 8 B). This implies that other cell volume regulatory ion transport mechanisms can compensate for the loss of NKCC1 during this process. However, it is unclear why these ion transport mechanisms are functional after isosmotic shrinkage but not in hypertonic shrinkage. It could be that a loss of osmotically active molecules, such as KCl, under isosmotic shrinkage removes a factor(s) that otherwise inhibits these ion transport mechanisms (24). Moreover, the RVD rate in NKCC1−/−cortical astrocytes is significantly faster than that in NKCC1+/+ cortical astrocytes. We speculate that some ion efflux pathways, such as K+ and Cl− channels or K-Cl cotransporters, could be upregulated in NKCC1−/−cortical astrocytes. Further investigation is needed to verify this speculation.
Role for NKCC1 in high-[K+]o-induced astrocyte swelling and glutamate release.
Several studies using pharmacological approaches suggested that NKCC1 plays a role in K+ uptake and astrocyte swelling in high [K+]o (37). In rat cortical astrocytes, we reported that inhibition of NKCC1 with bumetanide abolished the high-[K+]o-induced swelling (33). A similar observation was made in choroid plexus cells (40). In the current report, we conclude that activation of NKCC1 is responsible for high-[K+]o-induced swelling in mouse cortical astrocytes. Blocking of NKCC1 activity or ablation of NKCC1 abolishes high-[K+]o-induced swelling. We believe that NKCC1 leads to cell swelling via excessive influx of Na+, K+, and Cl−, with accompanying H2O under high [K+]o. This view is supported by the following findings. 1) NKCC1 activity is significantly stimulated in NKCC1+/+ astrocytes. 2) Either inhibition of NKCC1 with bumetanide or ablation of NKCC1 significantly reduces intracellular Cl− content under high [K+]o. Na+ entering the cell via NKCC1 is then subsequently pumped out of the cell via Na+-K+-ATPase. This speculation is based on our finding that inhibition of Na+-K+-ATPase with 1 mM ouabain prevents the reduction of intracellular Na+ in rat astrocytes under high [K+]o (data not shown). A voltage-dependent stimulation of Na+-K+-ATPase has been established (7).
Transport mechanisms underlying the reduction in [Na+]i under high-[K+]o conditions remain unknown. Longuemare et al. (19) recently suggested that an increase in Na+-K+-ATPase activity, which is stimulated by high [K+]o (16, 38), may contribute to reduced [Na+]i. However, because [Na+]i is normally a rate-limiting factor for Na+-K+-ATPase, it is likely that changes in Na+ entry mechanisms also contribute to the reduction in [Na+]i. For example, astrocytes express a plasma membrane Na+ conductance that has been suggested to provide an important leak pathway for the Na+-K+-ATPase (30) and a decrease in inwardly directed Na+ currents has been observed under conditions of high [K+]o (36). Thus the reduced set point for [Na+]i during high [K+]o could be the result of a balance between the decrease in the inwardly directed Na+ current and stimulation of intrinsic Na+-K+-ATPase activity.
High [K+]o causes glutamate release from astrocytes in response to high-[K+]o-induced swelling (1, 18). It is generally believed that astrocytes reduce cell volume by the efflux of chloride, glutamate, and other anions through VSOACs (1, 18, 27). However, the channels responsible for VSOAC currents in astrocytes remain to be identified (31). In addition to the swelling stimulus, Cl− channels are regulated by a variety of signal transduction pathways, such as those involving Ca2+/calmodulin, cAMP, and protein kinase C (1,14).
In the current study, high [K+]o triggers the release of d-[14C]Asp from both NKCC1+/+ and NKCC1−/− astrocytes. The stilbene derivative DIDS abolished the high-[K+]o-induced release ofd-[14C]Asp from both NKCC1+/+ and NKCC1−/− astrocytes. This implies that the release ofd-[14C]Asp is largely through VSOACs. The slow onset kinetics of the d-[14C]Asp release in our study (33) further supports this view. Interestingly, elimination of the high-[K+]o-mediated swelling by bumetanide or ablation of NKCC1 only blocks ∼30% of thed-[14C]Asp release. This suggests that under high-[K+]o conditions, the VSOACs can be activated by other mechanisms in addition to swelling. VSOAC activation in astrocytes is reportedly dependent on calmodulin and intracellular Ca2+ (22). The Ca2+ chelator 1,2-bis(2-aminophenoxy)ethane-N,N,N′,N′-tetraacetic acid (BAPTA)-AM and the calmodulin antagonist trifluoperazine significantly suppress [K+]o-induced taurine release in rat astrocytes (22). Intracellular Ca2+ rise has been observed in both rat and mouse astrocytes under high-[K+]o conditions (2, 21, 28, 32). In cultured astrocytes, the high-[K+]o-mediated Ca2+ influx was blocked by voltage-operated Ca2+ channel inhibitors (21, 32, 42). However, the depolarization-induced intracellular Ca2+ concentration ([Ca2+]i) increases in astrocytes in situ are attributed to metabotropic glutamate receptor-mediated Ca2+release from intracellular Ca2+ stores (2). In the current study, the release of d-[14C]Asp from astrocytes in the absence of cell swelling is abolished by removal of extracellular Ca2+. Therefore, it is possible that a rise in [Ca2+]i activates VSOACs and leads to glutamate release. However, Ca2+-free medium plus 50–100 μM EGTA only moderately inhibits the [K+]o-induced taurine release (22) and has no effect on glutamate release in rat astrocytes (28). This is different from our observations on the release of d-[14C]Asp in mouse astrocytes. The cause of this discrepancy is unclear. It could be due to species differences (rat vs. mouse) or different cell culture conditions (differentiated vs. undifferentiated astrocytes).
In summary, with the use of both pharmacological and transgenic knockout approaches, we report here that NKCC1 contributes to baseline [Na+]i in mouse cortical astrocytes. Ablation of NKCC1 leads to complete impairment of RVI in mouse cortical astrocytes. High-[K+]o-induced swelling and accumulation of intracellular Cl− are absent in NKCC1−/− astrocytes. In addition, the release ofd-[14C]Asp is inhibited by 30% in NKCC1−/− astrocytes. Together, these results demonstrate that NKCC1 is important in the regulation of cell volume and intracellular ion concentrations in astrocytes.
We thank Charanjeet Kaur of Dr. Albee Messing's laboratory for technical assistance in genotyping.
This work was supported in part by National Institutes of Health (NIH) Grant R01-NS-38118, NSF CAREER Award IBN9981826 to D. Sun, and NIH Grant R01-DK-50594 to G. E. Shull.
Address for reprint requests and other correspondence: D. Sun, Dept. of Neurological Surgery, Univ. of Wisconsin Medical School, H4/332 Clinical Sciences Center, 600 Highland Ave., Madison, WI 53792 (E-mail:)
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- Copyright © 2002 the American Physiological Society