We hypothesized that high extracellular K+ concentration ([K+]o)-mediated stimulation of Na+-K+-Cl− cotransporter isoform 1 (NKCC1) may result in a net gain of K+ and Cl−and thus lead to high-[K+]o-induced swelling and glutamate release. In the current study, relative cell volume changes were determined in astrocytes. Under 75 mM [K+]o, astrocytes swelled by 20.2 ± 4.9%. This high-[K+]o-mediated swelling was abolished by the NKCC1 inhibitor bumetanide (10 μM, 1.0 ± 3.1%; P < 0.05). Intracellular36Cl− accumulation was increased from a control value of 0.39 ± 0.06 to 0.68 ± 0.05 μmol/mg protein in response to 75 mM [K+]o. This increase was significantly reduced by bumetanide (P < 0.05). Basal intracellular Na+ concentration ([Na+]i) was reduced from 19.1 ± 0.8 to 16.8 ± 1.9 mM by bumetanide (P < 0.05). [Na+]i decreased to 8.4 ± 1.0 mM under 75 mM [K+]o and was further reduced to 5.2 ± 1.7 mM by bumetanide. In addition, the recovery rate of [Na+]i on return to 5.8 mM [K+]o was decreased by 40% in the presence of bumetanide (P < 0.05). Bumetanide inhibited high-[K+]o-induced 14C-labeledd-aspartate release by ∼50% (P < 0.05). These results suggest that NKCC1 contributes to high-[K+]o-induced astrocyte swelling and glutamate release.
- cell swelling
- high potassium ion concentration, cultured astrocytes
- glutamate release
- intracellular chloride
the na+ -k+-cl− cotransporters (NKCCs) are membrane proteins that mediate the coupled, electrically neutral movement of Na+, K+, and Cl− across the membrane of many cell types (25). NKCC isoform 1 (NKCC1) is important for accumulation of Cl− in neurons, astrocytes, and oligodendrocytes (17, 25, 34). High intracellular Cl− concentration ([Cl−]i) makes possible the depolarizing action of GABA and glycine that opens Cl− channels (1). The inhibition of spontaneous epileptiform activity in rat hippocampal slices by furosemide has been attributed to the blockade of K+ uptake mediated by NKCC1 in hippocampal glial cells (13, 14). In the recent study of Yan et al. (41), inhibition of NKCC1 by a more potent inhibitor, bumetanide, resulted in a significant reduction of edema and infarct volume in rat focal cerebral ischemia. In the current study, we investigated the role of NKCC1 in astrocyte swelling and glutamate release induced by elevated extracellular K+ concentration ([K+]o) to further understand the contribution of glial NKCC1 in ischemic cerebral damage. Both high [K+]o and glutamate release are associated with ischemic cerebral damage (28).
High-[K+]o-induced astrocyte swelling has been observed in both brain slices and cultured astrocytes. However, the cellular mechanisms underlying high-[K+]o-induced astrocyte swelling have not been completely defined. NKCC has been implicated in high-[K+]o-induced swelling in several cell types, and high-[K+]o-induced swelling was abolished either by the cotransporter inhibitor bumetanide or removal of extracellular Cl− or Na+ (39). Furosemide blocked 70% of the high-[K+]o-induced increase in intracellular K+ content observed in cultured mouse cortical astrocytes (36). In a recent study (32), we found that the activity of NKCC1 in cultured rat cortical astrocytes was significantly stimulated under 75 mM [K+]o. This led us to hypothesize that this high-[K+]o-induced stimulation of cotransporter activity may cause Na+, K+, and Cl− influx and result in swelling in astrocytes.
One consequence of high-[K+]o-induced swelling is the stimulation of excitatory amino acid (EAA) release from astrocytes (18). The release of EAA under high-[K+]o conditions could be mediated by volume-sensitive organic anion channels (VSOACs; Refs. 2,27). High-[K+]o-induced3H-labeled d-aspartate (Asp) release from cultured astrocytes is inhibited by the anion channel inhibitors L-644711 and dideoxyforskolin (26). We hypothesized that NKCC1 may play a role in a swelling-dependent release of EAA under high-[K+]o conditions.
We report here the effects of inhibition of NKCC1 activity on cell swelling, intracellular Cl− accumulation, changes of intracellular Na+ concentration ([Na+]i), and release of14C-labeled d-Asp in cultured cortical astrocytes under high [K+]o.
MATERIALS AND METHODS
Bumetanide, digitonin, Triton X-100, monensin, gramicidin, and 4,4′-diisothiocyanostilbene-2,2′-disulfonic acid (DIDS) 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).
Primary culture of rat cortical astrocytes.
Dissociated cortical astrocyte cultures were established as described previously (32). Cerebral cortices were removed from 1-day-old rats (Sprague-Dawley). The cortices were incubated in a trypsin solution for 25 min at 37°C. The tissue was then mechanically triturated and filtrated through nylon meshes (70 μm). 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% CO2 atmosphere 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 then 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 (11, 35). Experiments were routinely performed on cultures treated with DBcAMP for 7 days. More than 95% of cells in culture yielded by this preparation were astrocytes (32).
Measurement of relative cell volume changes in single cell.
Relative cell volume changes were determined in cultured single astrocytes on coverslips with video-enhanced differential interference contrast (DIC) microscopy (7, 10, 39). Astrocytes were cultured on collagen-coated coverslips and placed in a home-made bath chamber mounted on the stage of a Nikon TE 300 inverted epifluorescence microscope. The bath chamber was perfused continuously at room temperature at 1.0 ml/min, and the dead space between the perfusion pump and the bath chamber was 1.15 ml. Astrocytes were equilibrated with an isotonic HEPES-buffered minimal essential medium (MEM; 312 mosmol/kgH2O) for 15 min. The concentrations of components in HEPES-MEM were (mM) 140 NaCl, 5.36 KCl, 0.81 MgSO4, 1.27 CaCl2, 0.44 KH2PO4, 0.33 Na2HPO4, 0.4 NaHCO3, 5.55 glucose, and 20 HEPES. Astrocytes were perfused sequentially with HEPES-MEM (10 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+]oHEPES-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 (1.4 NA, 0.21 WD). Cell images were recorded every minute as 16-bit TIF files with a Princeton Instruments MicroMax charge-coupled device (CCD) camera (model 1300 YHS; Roper Scientific, Trenton, NJ). For each image, the cell body was traced three separate times with a mouse and the mean cross-sectional area (CSA) of the cell body was calculated with MetaMorph image-processing software (Universal Imaging, Downingtown, PA). The control CSA values were obtained when cells were exposed to HEPES-MEM only. Relative volume changes were calculated as CSAr = experimental CSA ÷ control CSA. On the same image, a peripheral astrocytic process that was far distant from the cell body was selected and the mean cross-sectional distance of the process (CSD) and relative CSD (CSDr) were determined in a manner similar to the CSA measurement. After each experiment, a calibration curve was constructed by measuring relative cell volume changes in response to HEPES-MEM calibration buffers in which salt concentrations 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 0–30 min in HEPES-MEM containing 5.8 mM [K+]o and36Cl (0.4 μCi/ml). 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 1–13 min. Thus 145 mM Cl− in HEPES-MEM was maintained in 75 mM [K+]o and the specific activity of36Cl was constant in 5.8 mM [K+]oand 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).
[Na+]i was measured with the fluorescent dye SBFI-AM as described by Rose and Ransom (23). Cultured astrocytes grown on coverslips were loaded with 10 μM SBFI-AM at room temperature for 90 min in HEPES-MEM containing 0.1% pluronic acid. The coverslips were placed in an open-bath imaging chamber (volume = 40 μl; series 20, Warner Instruments, Hamden, CT) containing HEPES-MEM at ambient temperature. The chamber was mounted on the stage of a Nikon TE 300 inverted epifluorescence microscope, and the astrocytes were visualized with a ×40 Super Fluor oil-immersion objective lens (1.3 NA, 0.22 WD). The cells were excited every 10 or 60 s at 345 and 385 nm, and the emission florescences at 510 nm were recorded. In some experiments, the data were processed with a nonparametric digital filter (Peakfit; SPSS, Chicago, IL) to improve the signal-to-noise ratio. Images were collected as 16-bit TIF files with a Princeton Instruments MicroMax CCD camera and analyzed with MetaFluor image-processing software. Cytoplasmic regions with minimum punctuate fluorescence staining were chosen for measurement of fluorescent intensity changes of SBFI. 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 and correction of autofluorescence of bumetanide. To determine the percentage of SBFI dye in cytoplasm, astrocytes were clamped at an extracellular Na+ concentration ([Na+]o) of 20 mM with a calibration solution (see below). A decrease in SBFI fluorescence at 340 nm was recorded after plasma membrane was permeabilized by 20 μM digitonin. A further release of the dye from organelles was induced subsequently by 1% Triton X-100.
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 under different experimental conditions. Absolute [Na+]i was determined for each cell by standardization of the SBFI fluorescence ratio with calibration solutions containing 0, 10, 20, or 30 mM [Na+]o plus monensin (10 μM) and gramicidin (3 μM) to equilibrate [Na+]i and [Na+]o.
d-[14C]Asp release measurement.
Aspartate release was measured as described by Rutledge and Kimelberg (27). Astrocytes grown on chamber slides (Fisher, Pittsburgh, PA) were incubated overnight in 1 ml of complete EMEM containing 2 μCi/ml of d-[14C]Asp (specific activity of 55 mCi/mmol). Radiolabeledd-[14C]Asp is used as a nonmetabolizable marker for the intracellular glutamate and aspartate pool (27). Both of these amino acids are transported by the same glutamate carrier proteins (5, 8). A perfusion chamber was formed by a special lid that contains influx and efflux tubings in the chamber. The perfusing rate 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 from time t to the end of the experiment (27), and C remain is the cpm value left in the cells at the end of the experiment.
d-[14C]Asp uptake assay.
The assay was performed according to a method described by Kimelberg et al. (18). Cells were refed with EMEM on the evening before the uptake assay. On the following day, cells were washed four times (1 ml each) with HEPES-MEM to remove growth medium. The cells were preincubated with HEPES-MEM containing 5.8 mM [K+]o in the presence or absence of 10 μM bumetanide for 20 min at 37°C. In the high-[K+]o study, the cells were preincubated with 75 mM [K+]o HEPES-MEM in the presence or absence of 10 μM bumetanide for 20 min at 37°C. The buffer was then rapidly removed, and 0.5 ml of the same medium containingd-[14C]Asp (0.2 μCi/0.5 ml in each well) + unlabeled d-Asp (100 μM) was added to each well. The cells were then incubated for 1, 2, or 3 min. The uptake assay was terminated by three washes with ice-cold 0.1 M MgCl2 (1 ml each). Radioactivity of cellular extract in 1% SDS was analyzed by liquid scintillation counting. Uptake rate was determined by analyzing the slope of the uptake over time.
Inhibition of NKCC1 abolishes high-[K+]o-induced astrocyte swelling.
To validate DIC microscopy for the determination of CSArand CSDr in single cells, we measured the changes in CSAr in cultured astrocytes perfused with isotonic and hypotonic HEPES-MEM buffers (Fig.1 A). CSArresponded linearly (r = 0.999) as the osmolality of the perfusion buffer decreased. CSAr returned to basal levels when cells were incubated in the isotonic buffer (data not shown). This suggests that changes of CSAr in astrocytes measured by DIC microscopy can be used as an estimate of cellular volume changes, as reported by others (7, 10, 39).
When astrocytes were exposed to 75 mM [K+]o, CSAr increased gradually and reached a maximum value of 19% after 8 min (Fig. 1 B, b). CSArreturned to basal levels within 10 min when the cell was perfused with 5.8 mM [K+]o HEPES-MEM (Fig. 1 B,a and b). To test whether NKCC1 plays a role in this high-[K+]o-induced volume increase, 10 μM bumetanide was used to block cotransporter activity. The cells were perfused for 20 min with normal HEPES-MEM buffer containing 10 μM bumetanide. There was no significant effect of 10 μM bumetanide on the basal level of CSAr (Fig. 1B, c). When the cell was subsequently exposed to 75 mM [K+]o buffer containing 10 μM bumetanide, the CSAr responses to high [K+] were absent (Fig. 1 B, d). We then further tested whether the lack of the swelling in the presence of 75 mM [K+]o + bumetanide could be due to a lack of a cellular response to the second 75 mM [K+]o stimulus. As shown in Fig.1 B (inset), a second exposure of cells to 75 mM [K+]o without bumetanide led to a degree of swelling similar to that from the first exposure. Figure 1 C,left, summarizes CSAr measurements on single astrocytes. Maximal CSAr was significantly increased during the high-[K+]o perfusion (P< 0.05). This response was abolished in the presence of 10 μM bumetanide (P < 0.05). This suggests that NKCC1 is involved in astrocyte swelling in response to high [K+]o.
NKCC1 is expressed in plasma membrane of the cell body and process of astrocytes (32, 40). To test whether NKCC1 also contributes to swelling in the cell process, CSDr was measured. Similar to CSAr, CSDr increased significantly during high-[K+] perfusion (a maximum value of 29 ± 4%, P < 0.05; Fig. 1 C,right). Exposure of cells to both 10 μM bumetanide and high [K+]o abolished the swelling response of the astrocyte to 75 mM [K+]o.
We believe that bumetanide exerts its effect on high-[K+]-induced swelling by blocking of net influx of Na+, K+, and Cl− via NKCC1. To further support this view, we investigated whether impairment of cotransporter activity by removal of extracellular Na+could prevent the high-[K+]-mediated swelling. Figure2 illustrates that removal of extracellular Na+ resulted in ∼7% cell shrinkage under 5.8 mM [K+]o (P < 0.05). No cell swelling occurred when the cells were subsequently exposed to 75 mM [K+]o. The effect mediated by the Na+-free treatment was reversible. Astrocyte volume completely recovered when cells were returned to normal HEPES-MEM with 5.8 mM [K+]o. Thus inhibition of NKCC1 either by bumetanide or by removal of extracellular Na+ prevented high-[K+]-induced swelling.
Sustained elevation of cotransporter activity in presence of high-[K+]o-induced astrocyte swelling.
Cell swelling inhibits NKCC1 activity in many cell types (25). To investigate whether high-[K+]o-induced cell swelling under 75 mM [K+]o affects cotransporter activity, cotransporter activity was measured when astrocytes were exposed to either 5.8 or 75 mM [K+]o for 1–12 min. As shown in Fig. 3, under control conditions, cotransporter activity was 34.3 ± 5.1 nmol/mg protein × min (n = 5), and it did not change over the entire time course. In contrast, the activity of NKCC1 increased to the maximum level of 143.6 ± 38.1 nmol/mg protein × min (n = 5; P < 0.05) after 1 min of exposure to high [K+]o. It decreased gradually and reduced to 61.1 ± 9.0 nmol/mg protein × min (n = 5, P < 0.05) at 4 min. Cotransporter activity remained elevated during the rest of the exposure time (P < 0.05). The results imply that cotransporter activity remained stimulated under high [K+]o despite astrocyte swelling.
High-[K+]o-mediated increase in Cl− uptake is abolished by blocking of NKCC1.
To further examine whether NKCC1 contributes to high-[K+]o-induced cell swelling, we measured changes of intracellular Cl− content. Cells were preequilibrated in HEPES-MEM with 36Cl (0.4 μCi/ml) for 0–30 min. A steady-state level of intracellular36Cl− was obtained by only 4-min incubation and maintained during the 30 min-equilibration (data not shown). Thus, in the rest of the study, a 30-min preincubation was performed. After a 30-min equilibration with 36Cl (0.4 μCi/ml), the time course of intracellular Cl− content changes was measured under control or high-[K+]o conditions. Intracellular 36Cl− content was 0.49 ± 0.04 μmol/mg protein (n = 8) after exposure of cells to 5.8 mM [K+]o for 1 min (Fig.4 B). After 13 min of incubation in 5.8 mM [K+]o, the intracellular36Cl content was maintained at 0.39 ± 0.06 μmol/mg protein (P > 0.05). However, at 1 min of incubation of cells with 75 mM [K+]o,36Cl− content increased to 0.67 ± 0.05 μmol/mg protein (P < 0.05). It reached a peak value of 0.78 ± 0.07 μmol/mg protein (P < 0.05,n = 8) after 4 min of incubation with 75 mM [K+]o. A sustained elevation of intracellular36Cl− content was detected during the 13-min incubation period. In contrast, in the presence of 10 μM bumetanide, the high-[K+]o-induced intracellular36Cl rise was significantly inhibited. The values of intracellular 36Cl− content were significantly less than in non-bumetanide-treated cells at 1, 4, 6, 10, or 13 min (P < 0.05; n = 5). A small36Cl increase was observed at 10–13 min. The nature of the bumetanide-insensitive Cl− influx is unclear. It could reflect Cl− influx via Cl−channels or reversal of outward K-Cl cotransport (21). These results support a view that NKCC1 contributes to Cl−accumulation under high-[K+]o conditions and may lead to cell swelling.
We next investigated whether stimulation of NKCC1 activity under high [K+]o could affect [Na+]i. First, intracellular localization of SBFI dye in astrocytes was examined. Addition of 20 μM digitonin to permeabilize the plasma membrane resulted in a decrease of the 340-nm SBFI fluorescence signal (Na+-insensitive fluorescence) by 64.3 ± 6.9% (n = 1, 4 coverslips, 32 cells; Fig.5 A). This reflects a loss of SBFI from the cytoplasm. When the detergent Triton X-100 (1%) was subsequently added to cells, the 340-nm SBFI fluorescence signal decreased to near zero. This is presumably the result of release of the remaining SBFI dye from intracellular organelles. Typically, we selected a region of the cell cytoplasm that was as free of punctuate SBFI fluorescence as possible. Thus the SBFI fluorescence signals (340- to 380-nm ratios) measured in this study largely represent changes of Na+ in the cytoplasm of astrocytes. This pattern of intracellular SBFI dye localization in rat cortical astrocytes has been reported in rat hippocampal astrocytes (23).
Calibration of [Na+]i was performed by a four-point calibration at the end of each experiment (Fig.5 B). In these experiments, the Na+ ionophore gramicidin (5 μM) and the H+ ionophore monensin (10 μM) were used to equilibrate extra- and intracellular Na+ and H+ concentrations with calibration solutions containing 0, 10, 20, or 30 mM [Na+]o. Figure 5 Bshows that the SBFI fluorescence ratios changed in a linear fashion with [Na+]o over the range of 0–30 mM (r = 0.998; n = 3, 8 coverslips, 40 cells). Using the calibrations performed at the end of each experiment, we determined a baseline [Na+]i of 19.1 ± 0.8 mM (n = 4, 12 coverslips, 62 cells) in cultured rat cortical astrocytes. This value is similar to values reported for rat hippocampal astrocytes (14.6 mM; Ref. 23) and rat cortical astrocytes (15.3 mM; Ref. 19) but is significantly higher than the value reported for rat spinal cord astrocytes (8.3 mM; Ref.24).
Incubation of cells in 10 μM bumetanide caused a decrease in basal levels of [Na+]i, and a new steady-state [Na+]i was established after 10 min (Fig.6 A). The mean [Na+]i after 20 min of bumetanide treatment decreased from a control level of 19.1 ± 0.8 mM to 16.8 ± 1.9 mM (P < 0.05; n = 2, 6 coverslips, 26 cells; Fig. 6B). This indicates that NKCC1 plays a role in maintaining basal [Na+]i in rat cortical astrocytes.
NKCC1 has been proposed to provide Na+ for Na+-K+-ATPase function (36). NKCC1 and Na+-K+- ATPase may work synergistically to uptake K+ and maintain low [Na+]i. Under high [K+]o, a decrease in [Na+]i was observed (Fig. 6, A andB). This reduction in [Na+]i has been attributed to a stimulation of Na+-K+-ATPase (19). We then examined whether cotransporter activity affects the compensatory loss of intracellular Na+ under 75 mM [K+]o. In bumetanide-treated rat cortical astrocytes, [Na+]i decreased rapidly in response to 75 mM [K+]o and plateaued after 2–4 min (Fig. 6 A). A similar pattern was found in cells without bumetanide (data not shown). As shown in Fig. 6 B, in control astrocytes [Na+]i decreased from 19.1 ± 1.5 to 8.4 ± 1.0 mM (P < 0.05;n = 4, 12 coverslips, 62 cells; Fig. 6 B). In bumetanide-treated cells [Na+]i further dropped to 5.2 ± 1.7 mM (P < 0.05;n = 2, 6 coverslips, 27 cells; Fig. 6 B). This implies that Na+ influx via the cotransporter contributes to maintaining [Na+]i under both control 5.8 mM and 75 mM [K+]o conditions. The high-[K+]o-induced relative changes in [Na+]i were 11.6 mM in control cells vs. 10.7 mM in bumetanide-treated cells (P > 0.05). This suggests that the cotransporter function is not a rate-limiting factor for Na+ efflux via Na+-K+-ATPase. Furthermore, the decrease in [Na+]i in 75 mM [K+]o is reversible in both control (data not shown) and bumetanide-treated (Fig. 6B) astrocytes. The effect of bumetanide on changes of [Na+]i is also reversible. When bumetanide was removed from the perfusate, [Na+]i slowly returned toward basal levels (Fig. 6, A and B).
To further investigate the effect of NKCC1 on [Na+]i, the slopes of intracellular Na+ changes during and after 75 mM [K+]o treatment were determined (Fig.7, inset). When cells were exposed to 75 mM [K+]o, the slopes of [Na+]i reduction were similar in control and bumetanide-treated astrocytes (2.75 ± 1.42 vs. 2.28 ± 0.67 mM/min; Fig. 7). In contrast, when bumetanide-treated cells were returned to 5.8 mM [K+]o, the rate of [Na+]i recovery was significantly slower than in control cells (1.83 ± 0.97 vs. 2.96 ± 1.56 mM/min;P < 0.01; Fig. 7). This suggests that Na+influx via the cotransporter is important for reestablishment of basal [Na+]i after Na+-K+-ATPase activation under high [K+]o.
Inhibition of NKCC1 attenuates high-[K+]o-mediated preloaded d-[14C]Asp release.
To investigate whether cotransporter-mediated cell swelling is involved in glutamate release, we examined whether blocking cotransporter activity could inhibit aspartate release. As shown in Fig.8 A, under control conditions a trace level of release of preloadedd-[14C]Asp was detected. This is consistent with a previous report (27). After 6 min of exposure to high [K+]o, an increase ind-[14C]Asp release occurred (n = 4). The release developed progressively (Fig.8 A) and reached 1.5% fractional release ofd-[14C]Asp after 22 min (n = 4) in high [K+]o. On removal of high-[K+]o medium,d-[14C]Asp release returned to a resting level within 12 min (n = 4; Fig. 8 A). Figure8 B shows that exposing cells to 10 μM bumetanide did not significantly affect basal d-[14C]Asp release in 5.8 mM [K+]o. Moreover, in the presence of 10 μM bumetanide and 75 mM [K+]o, the peak value of d-[14C]Asp release was only about one-third of that in the absence of bumetanide (Fig. 8 B). This inhibition of aspartate release could be reversed by removal of 10 μM bumetanide (Fig. 8 B). Figure 8 C shows that the average peak value of the fractional release under high [K+]o was 1.52 ± 0.17% (n = 4). In contrast, the fractional release was 0.68 ± 0.13% (n = 4) in the presence of bumetanide (P < 0.05).
As a control experiment, d-[14C]Asp uptake rate was measured under both 5.8 and 75 mM [K+]o to rule out a role of the cotransporter in Na+-dependent glutamate uptake (Fig.9). d-[14C]Asp uptake was significantly inhibited under high [K+]o, and this is consistent with other reports (18). However, 10 μM bumetanide did not affect the d-[14C]Asp uptake rate under both normal and high-[K+]o conditions. Therefore, our observation of a bumetanide effect on aspartate release does not indirectly reflect the changes of aspartate reuptake.
Release of glutamate under high [K+]o is thought to occur via an activation of VSOACs (2,26). To further understand whether the high-[K+]o-triggeredd-[14C]Asp release is indeed associated with VSOACs, we examined whether DIDS, a broad-spectrum Cl− channel inhibitor, could block thed-[14C]Asp release. Figure10 shows that 100 μM DIDS did not affect the basal level of the d-[14C]Asp release. However, the high-[K+]o-triggeredd-[14C]Asp release was abolished by 100 μM DIDS. The data provide further support for the view that activation of VSOACs under high [K+]o causes efflux of EAA from astrocytes. As a control, we also examined whether DIDS has a nonspecific effect on NKCC1 activity. In the presence of 75 mM [K+]o and 100 μM DIDS, bumetanide-sensitive86Rb influx was measured. It was not statistically different from a control value at 75 mM [K+]o(P > 0.05; data not shown).
Role of NKCC1 in high-[K+]o-induced astrocyte swelling.
Astrocytes are thought to have a primary role in the clearance of K+ from the extracellular space in physiological and pathological conditions (38). Recently, Yan et al. (40) reported that NKCC1 protein is expressed in astrocytes in rat cortex, cerebellum, and hippocampus. An abundant level of NKCC1 protein is also detected in perivascular astrocytes (40). NKCC1 has been suggested to play a role in K+ uptake in cortical (36) or hippocampal (14) astrocytes. The cotransporter activity in astrocytes is significantly stimulated in response to high [K+]o in a Ca2+-dependent manner (32). Stimulation of NKCC1 under high [K+]o may result in cell swelling via a net increase of intracellular KCl and accompanying water. Cell swelling in rat hippocampal slices was detected with changes in intrinsic optical signals, and inhibition of NKCC1 increases extracellular space and blocks synchronized burst discharges (12).
The current study directly establishes that NKCC1 is responsible for a high-[K+]o-mediated swelling in cultured rat cortical astrocytes. This conclusion is based on the following findings. 1) Astrocyte swelling (cell body and process) occurs within 3–4 min in response to high [K+]o and reaches a peak level by 8 min.2) Cotransporter activity is significantly stimulated when astrocytes are exposed to high [K+]o, and stimulation of the cotransporter precedes the swelling. 3) A high-[K+]o-mediated Cl− uptake correlates with the time course of the cotransporter stimulation.4) Inhibition of cotransporter activity by 10 μM bumetanide or removal of extracellular Na+ reversibly abolishes the high-[K+]o-induced astrocyte swelling. It could be argued that the lack of high-[K+]o-mediated swelling in 0 mM [Na+]o in our current study is due to astrocyte membrane depolarization followed by an inhibition of K+ uptake mediated by rectifying K+ channels (22). However, our cotransporter-mediated hypothesis is further supported by our recent study (Ref. 33; this issue) that shows that genetic ablation of NKCC1 abolished high-[K+]o-mediated swelling in mouse cortical astrocytes. Together, these data suggest that the NKCC1 in astrocytes may play an important role in astrocyte swelling when [K+]o reaches pathological levels under conditions such as ischemia and traumatic head injury.
Role of NKCC1 in high-[K+]o-mediated intracellular Cl− accumulation.
Astrocytes have a higher [Cl−]i level than predicted by passive distribution of the ion (9). NKCC1 is important for an accumulation of intracellular Cl− in neurons (34), so it is conceivable that NKCC1 may also contribute to intracellular Cl− accumulation in astrocytes. In the current study, inhibition of NKCC1 with 10 μM bumetanide did not significantly affect basal levels of intracellular36Cl. This implies that basal [Cl−]i can be maintained by other Cl− influx mechanisms. When astrocytes were exposed to high [K+]o intracellular 36Cl content was increased by ∼70%, and this increase in 36Cl level was significantly reduced by the cotransporter inhibitor bumetanide. An increase in Cl− influx, accompanied by K+ uptake, has been found in cultured mouse cortical astrocytes under high [K+]o (37,38). An increase of Cl− permeability by activation of voltage-dependent Cl− channels has been proposed as a major mechanism for the Cl− influx (37). Our current study suggests that the NKCC1-mediated Cl− influx also contributes to intracellular Cl− accumulation, and this may subsequently lead to astrocyte swelling under high [K+]o.
Role of NKCC1 in [Na+]i.
NKCC1 has been suggested to provide Na+ for Na+-K+-ATPase function in the so-called “transmembrane Na+ cycle” (36). Rose and Ransom (23) reported that application of 50 μM bumetanide to cultured hippocampal astrocytes caused a slow and reversible decrease in [Na+]i by <2 mM (14% of baseline). In our study, treatment of cortical astrocytes with 10 μM bumetanide caused a decrease in [Na+]iby 2.3 mM (12% of baseline), suggesting a role of the cotransporter in maintenance of a resting [Na+]i in cultured rat cortical astrocytes.
Consistent with reports by others (19, 38), we observed that [Na+]i in astrocytes significantly decreases when cells are exposed to high [K+]o. The decrease in [Na+]i can be partially attributed to the experimental decrease in [Na+]o (140 to 75 mM) that accompanies the high-[K+]otreatment. In addition, a stimulation of Na+-K+-ATPase by elevated external K+ may also play a role (36). In the current study, the decrease of [Na+]i was prevented by blocking Na+-K+-ATPase with 1 mM ouabain (data not shown). Moreover, inhibition of NKCC1 by bumetanide led to a further loss of [Na+]i by 3.2 mM under high [K+]o. However, the net decrease in [Na+]i was not changed in control vs. bumetanide-treated cells (Fig. 6 B), which suggests that the Na+ efflux via Na+-K+-ATPase was not affected by bumetanide treatment. This implies that it is not NKCC1, but other Na+ influx mechanisms such as Na+ channel, Na+/H+ exchanger, or Na+/HCO cotransporter, that may provide Na+ and maintain Na+-K+-ATPase function in 75 mM [K+]o in rat cortical astrocytes.
Interestingly, we found that inhibition of NKCC1 resulted in an ∼40% decrease in the slope of intracellular Na+ recovery when cells were returned to 5.8 mM [K+]o (Fig. 7). This effect is more profound than the effect of inhibition of NKCC1 on a steady-state level of [Na+]i under high [K+]o (a 12% decrease in the latter case). The differential effects imply that when the Na+ efflux mediated by Na+-K+-ATPase is reduced, the role of NKCC1 in intracellular Na+ accumulation is unmasked.
Role of NKCC1 in high-[K+]o-inducedd-[14C]Asp release.
In the current study, high-[K+]o-inducedd-[14C]Asp release was detected in cultured rat cortical astrocytes. High [K+]o could induce glutamate release from astrocytes via nonvesicular mechanisms, either a reversal of glutamate transporter or VSOACs (2,27). Under physiological conditions the largest factor influencing the extracellular/intracellular glutamate equilibrium potential and the direction of the glutamate transporter is the Na+ gradient (19). Thus, if there were no compensation of intracellular Na+, an increased [K+]o plus decreased [Na+]o would reduce the Na+gradient and extracellular/intracellular glutamate equilibrium potential. This would result in the release of glutamate via reversal of the transporter (4). However, in the current study, the transmembrane Na+ gradient increased from 8.7 ± 1.7 to 12.9 ± 5.2 in response to the decrease in [Na+]o during 75 mM [K+]o incubation. Therefore, as discussed by Longuemare et al. (19), it is unlikely that the reversal of the glutamate transporter is a primary cause of the increase ind-[14C]Asp release under 75 mM [K+]o conditions. However, a fast-developing small release of d-[14C]Asp, before the delayed peak release (shown in Fig. 8), has been suggested to be mediated via a reversal of the glutamate transporters (26,27). This small release was enhanced by an increase of intracellular Na+ with 1 mM ouabain and inhibited by the glutamate transporter inhibitor threo-hydroxy β-aspartic acid (26, 27). A low [Na+]o at 75 mM [K+]o and the low [Na+]o-induced depolarization in astrocytes (22) may facilitate the reversal of the glutamate transporters.
Glutamate release from astrocytes can also occur in response to cell swelling via VSOACs (2, 27). However, the channel(s) responsible for VSOAC function remain to be identified (30,31), although several members of the Cl− channel subfamilies (ClC-3, 4, and 5) are proposed candidates (2,15). Rutledge and Kimelberg (27) reported that the rate of [K+]o-evoked glutamate release through VSOACs appears steadily, peaking 20–30 min after exposure of cells to 100 mM [K+]o. The release was completely blocked by the general anion channel blocker 5-nitro-2-(3-phenylpropylamino)benzoic acid (NPPB) and reduced by >90% by the nonspecific anion transport inhibitor DIDS (100 μM; Ref. 26).
In the current study, it took ∼6 min for high [K+]o to induced-[14C]Asp release and the release progressed steadily during 20 min of exposure of cells to high [K+]o. DIDS (100 μM) abolished thed-[14C]Asp release. The release pattern observed here resembled the high-[K+]o-induced glutamate release mechanism described above (26). Our study suggests that the cotransporter may contribute to this high-[K+]o-induced swelling and glutamate release. This view is supported by the following findings.1) The high-[K+]o-induced swelling precedes the d-[14C]Asp release under 75 mM [K+]o. 2) Activation of cotransporter activity precedes the development of astrocyte swelling and is sustained under high [K+]o.3) Stimulation of NKCC1 leads to astrocyte swelling and intracellular Cl− accumulation under high [K+]o. 4) Inhibition of NKCC1 by bumetanide results in ∼50% reduction of the preloadedd-[14C]Asp release during high [K+]o.
Although the current study focused on high-[K+]o-mediated swelling and release of glutamate from astrocytes, several reports suggest that glutamate and glutamate-mediated increase of intracellular Ca2+ are involved in bidirectional communication between neurons and astrocytes (3, 6, 16, 20). High-[K+]o-induced release of glutamate from neurons triggers a release of Ca2+ from intracellular Ca2+ stores in astrocytes (6). Ca2+-dependent glutamate release from astrocytes has been reported, and it requires the presence of functional vesicle-associated proteins (3). These studies also suggest a release of glutamate from astrocytes through exocytosis.
In summary, the results of our study suggest that NKCC1 is important in maintenance of intracellular [Na+] under resting- and high-[K+]o conditions. Pathological levels of [K+]o stimulate NKCC1 activity that leads to accumulation of intracellular Cl− and astrocyte swelling. Moreover, inhibition of cotransporter activity significantly decreases high-[K+]o-mediatedd-[14C]Asp release. Our findings imply that NKCC1 may play a role in astrocyte swelling and high-[K+]o-induced glutamate release under pathophysiological conditions in the central nervous system. Such a role of NKCC1 could contribute to in vivo cerebral ischemic damage because pharmacological inhibition of NKCC1 is neuron protective (41).
The authors thank Dr. James Franklin for the use of the Nikon epifluorescence microscope in his laboratory. We also thank Dr. Robert Haworth for helpful discussion and comments.
This work was supported in part by a Scientist Development Grant from the National Center Affiliate of American Heart Association (no. 9630189N), National Institute of Neurological Disorders and Stroke Grant R01-NS-38118, and National Science Foundation CAREER Award IBN9981826 to D. Sun.
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