Vol. 282, Issue 5, C1136-C1146, May 2002
Contribution of Na+-K+-Cl
cotransporter to high-[K+]o- induced
swelling and EAA release in astrocytes
Gui
Su1,2,
Douglas B.
Kintner1, and
Dandan
Sun1,2
Departments of 1 Neurological Surgery and
2 Physiology, University of Wisconsin Medical
School, Madison, Wisconsin 53792
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ABSTRACT |
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). Intracellular
36Cl 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-labeled
D-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; bumetanide; intracellular chloride
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INTRODUCTION |
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-induced
3H-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 of
14C-labeled D-Asp in cultured cortical
astrocytes under high [K+]o.
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MATERIALS AND METHODS |
Materials.
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+]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 (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 and
36Cl (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 of
36Cl was constant in 5.8 mM [K+]o
and 75 mM [K+]o HEPES-MEM. Intracellular
36Cl 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.
[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). Radiolabeled
D-[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:Cend) + Cremain], where Ct is
the cpm value in the effluent at time t,
Cend is the cpm value in the effluent at the end
of the experiment,
Sum(Ct:Cend) is the total
cpm value from time t to the end of the experiment
(27), and Cremain 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 containing
D-[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.
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RESULTS |
Inhibition of NKCC1 abolishes
high-[K+]o-induced
astrocyte swelling.
To validate DIC microscopy for the determination of CSAr
and CSDr in single cells, we measured the changes in
CSAr in cultured astrocytes perfused with isotonic and
hypotonic HEPES-MEM buffers (Fig.
1A). CSAr
responded 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).
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Fig. 1.
Effect of bumetanide on high extracellular K+
concentration ([K+]o)-induced astrocyte
swelling in single cells. A: relative cross-sectional area
(CSAr) responds linearly to buffer osmolality.
CSAr of a single astrocyte was determined during isotonic
perfusion (312 mosmol/kgH2O, 5 min), followed by 277 mosmol/kgH2O perfusion (5 min), isotonic perfusion (10 min), 238 mosmol/kgH2O perfusion (10 min), and isotonic
perfusion (10 min). The maximum response in CSAr at 277 and
238 mosmol/kgH2O is plotted along with the average
CSAr during the initial isotonic perfusion. Data are
means ± SD and were fit with a linear regression
(r = 0.999); n = 2, 6 cells.
Inset: illustration of CSA. B: bumetanide (Bum)
inhibits the increase in CSAr during high
[K+]o exposure. Mean CSAr in a
single astrocyte was determined during 5 min of normal HEPES- minimal
essential medium (MEM) perfusion (a), followed by 10 min of
75 mM [K+]o perfusion (b), 10 min
of normal HEPES-MEM perfusion (a), 20 min of normal
HEPES-MEM + 10 µM bumetanide perfusion (c), 10 min 75 mM [K+]o MEM + 10 µM bumetanide
perfusion (d), and 10 min of normal HEPES-MEM perfusion
(a). Inset: swelling induced by 2 consecutive
high [K+]o exposures. C: the
maximum CSAr (left) or relative
cross-sectional distance (CSDr) (right) in
single astrocytes during 75 mM [K+]o
perfusion is plotted along with the average CSAr or
CSDr during the final 5 min of normal HEPES-MEM,
HEPES-MEM + 10 µM bumetanide, 75 mM
[K+]o MEM, or 75 mM
[K+]o MEM + 10 µM bumetanide. Data are
means ± SD; n = 2, 6 cells.
* P < 0.05 vs. 5.8 mM
[K+]o, #P < 0.05 vs. 75 mM
[K+]o (Mann-Whitney rank sum test).
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When astrocytes were exposed to 75 mM [K+]o,
CSAr increased gradually and reached a maximum value of
19% after 8 min (Fig. 1B, b). CSAr
returned to basal levels within 10 min when the cell was perfused with
5.8 mM [K+]o HEPES-MEM (Fig. 1B,
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. 1B, 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.
1B (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 1C,
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. 1C,
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. Figure
2 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.
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Fig. 2.
Effect of removal of extracellular Na+ on
astrocyte swelling. Mean CSAr in a single astrocyte was
determined during 5 min of normal HEPES-MEM perfusion followed by 5 min
of Na+-free HEPES-MEM, 5 min of Na+-free
HEPES-MEM containing 75 mM [K+]o, and 10 min
of normal HEPES-MEM perfusion. In Na+-free HEPES-MEM,
Na+ was replaced with equimolar
N-methyl- -glucamine. Data are means ± SE;
n = 2, 6 cells. * P < 0.05 vs. 5.8 mM [K+]o (Mann-Whitney rank sum test).
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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.
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Fig. 3.
Time course of the
high-[K+]o-mediated stimulation of
cotransporter activity. Astrocytes were preincubated in HEPES-MEM
containing 5.8 mM [K+]o in either 0 or 10 µM bumetanide. Cells were then exposed to 75 mM
[K+]o for 1, 2, 3, 4, 6, 8, 10, or 12 min. In
the last 3 min of exposure to high [K+]o,
cells were switched to the same HEPES-MEM containing 1 µCi
86Rb. In the case of 1- and 2-min exposures, astrocytes
were treated directly with 75 mM [K+]o
containing 1 µCi 86Rb. [K+]o of
75 mM was obtained by replacing NaCl in HEPES-MEM buffer with equimolar
KCl. Data are means ± SE. * P < 0.05 vs. 5.8 mM [K+]o (Bonferroni-Dunn).
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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 intracellular 36Cl 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.
4B). After 13 min of
incubation in 5.8 mM [K+]o, the intracellular
36Cl 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 intracellular
36Cl content was detected during the 13-min
incubation period. In contrast, in the presence of 10 µM bumetanide,
the high-[K+]o-induced intracellular
36Cl 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 small
36Cl 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.
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Fig. 4.
Time course of
high-[K+]o-induced Cl uptake
increase and an effect of bumetanide. Astrocytes were preincubated in
normal HEPES-MEM at 37°C for 10 min and then equilibrated in
HEPES-MEM with 36Cl (0.4 µCi/ml) for 30 min. After 30-min
equilibration, astrocytes were treated with 75 mM
[K+]o containing 36Cl (0.4 µCi/ml) in the presence or absence of 10 µM bumetanide (Bum) for 1, 2, 3, 4, 6, 8, 10, or 13 min. In the control experiments, cells were
exposed to 5.8 mM [K+]o HEPES-MEM + 36Cl (0.4 µCi/ml) in the presence or absence of 10 µM
bumetanide for 1, 3, 5, or 13 min. Data are means ± SE.
* P < 0.05 vs. 75 mM
[K+]o + bumetanide (Bonferroni-Dunn).
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[Na+]i measurement.
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.
5A). 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).
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Fig. 5.
Estimation of cytosolic dye and calibration of
intracellular Na+ concentration
([Na+]i) in rat cortical astrocytes.
A: the fluorescence intensity of sodium-binding benzofuran
isophthalate (SBFI) was excited at 340 nm and monitored at 510 nm in
cortical astrocytes. [Na+]i was clamped at 20 mM Na+ with monensin (10 µM) and gramicidin (3 µM).
Plasma membrane was permeabilized by addition of 20 µM digitonin to
release the dye from the cytosol. Remaining dye was released by
addition of 1% Triton X-100. This is a representative experiment from
4 studies. B: calibration of
[Na+]i. Astrocytes were incubated in
calibration solutions containing 0, 10, 20, or 30 mM
[Na+]o and monensin (10 µM) and gramicidin
(3 µM). Data are means ± SE; n = 3, 8 coverslips, 40 cells. Data were fit with a linear regression.
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Calibration of [Na+]i was performed by a
four-point calibration at the end of each experiment (Fig.
5B). 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 5B
shows 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.
6A). 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.
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Fig. 6.
Effect of bumetanide on [Na+]i
under control and high-[K+]o conditions.
A: representative trace of [Na+]i
in a single cell during perfusion of 5.8 mM
[K+]o (5 min), 5.8 mM
[K+]o + 10 µM bumetanide (20 min), 75 mM [K+]o + 10 µM bumetanide (10 min),
5.8 mM [K+]o + 10 µM bumetanide (10 min), or 5.8 mM [K+]o (10 min). B:
summary data showing effects of 10 µM bumetanide on
[Na+]i under different conditions. Data are
means ± SD; control: n = 4, 12 coverslips, 62 cells; experimental: n = 2, 6 coverslips, 27 cells.
* P < 0.05 vs. 5.8 mM
[K+]o, ** P < 0.01 vs. 75 mM [K+]o, *** P < 0.01 vs. 5.8 mM [K+]o + bumetanide (after
high [K+]o).
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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 and
B). 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. 6A). A similar pattern was found in cells
without bumetanide (data not shown). As shown in Fig. 6B, 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. 6B). In
bumetanide-treated cells [Na+]i further
dropped to 5.2 ± 1.7 mM (P < 0.05;
n = 2, 6 coverslips, 27 cells; Fig. 6B).
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.
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Fig. 7.
Effect of bumetanide on slopes of
[Na+]i changes during and after 75 mM
[K+]o treatment. Summary data showing effects
of 10 µM bumetanide on the slopes of [Na+]i
changes. Data are means ± SD; n = 2, 6 coverslips, 26 cells. * P < 0.05 vs. 5.8 mM
[K+]o (after high
[K+]o). Inset: illustration of
measurement of slope of [Na+]i changes during
a transition between 5.8 and 75 mM [K+]o
(dotted line).
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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.
8A, under control conditions a
trace level of release of preloaded
D-[14C]Asp was detected. This is consistent
with a previous report (27). After 6 min of exposure to
high [K+]o, an increase in
D-[14C]Asp release occurred
(n = 4). The release developed progressively (Fig.
8A) and reached 1.5% fractional release of
D-[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. 8A). Figure
8B 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. 8B).
This inhibition of aspartate release could be reversed by removal of 10 µM bumetanide (Fig. 8B). Figure 8C 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).
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Fig. 8.
Effect of bumetanide on
high-[K+]o-mediated release of preloaded
D-[14C]aspartate (Asp) from astrocytes.
A: cells loaded with D-[14C]Asp
were perfused with 75 mM [K+]o (indicated by
bar) for 20 min. B: cells were treated with normal HEPES-MEM
containing 5.8 mM [K+]o in the presence of 10 µM bumetanide for 20 min, followed by 75 mM
[K+]o + 10 µM bumetanide for 20 min.
Bumetanide was then washed off for 25 min. Cells were reexposed to 75 mM [K+]o alone for 20 min. C:
summarized data are shown. Results are peak values of the percentage of
fractional release under 75 mM [K+]o in the
absence or presence of 10 µM bumetanide. Data are means ± SE;
n = 4. * P < 0.05 vs. absence of
bumetanide (Bonferroni-Dunn).
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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.
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Fig. 9.
Effect of bumetanide on
D-[14C]Asp uptake of astrocytes. Astrocytes
were preincubated in either HEPES-MEM (5.8 mM
[K+]o) alone or HEPES-MEM containing 10 µM
bumetanide for 10 min at 37 °C. In
high-[K+]o experiments, cells were
preincubated in HEPES-MEM (75 mM [K+]o) with
or without 10 µM bumetanide for 10 min at 37 °C.
D-[14C]Asp uptake (0.4 µCi/ml) in HEPES-MEM
containing either 5.8 or 75 mM [K+]o was
assayed for 1, 2, or 3 min at 37°C. Uptake rate was determined by
analyzing the slope of the uptake over time. Data are means ± SE;
n = 5. * P < 0.05 vs. 5.8 mM
[K+]o (Bonferroni-Dunn).
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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-triggered
D-[14C]Asp release is indeed associated
with VSOACs, we examined whether DIDS, a broad-spectrum
Cl channel inhibitor, could block the
D-[14C]Asp release. Figure
10 shows that 100 µM DIDS did not
affect the basal level of the D-[14C]Asp
release. However, the high-[K+]o-triggered
D-[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-sensitive
86Rb influx was measured. It was not statistically
different from a control value at 75 mM [K+]o
(P > 0.05; data not shown).
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Fig. 10.
Effect of 4,4'-diisothiocyanostilbene-2,2'-disulfonic
acid (DIDS) on the high-[K+]o-mediated
release of preloaded D-[14C]Asp. In the
control experiment, cells were perfused with 5.8 mM
[K+]o for 15 min, followed by 20 min of 75 mM
[K+]o and 20 min of 5.8 mM
[K+]o. In the experiment with DIDS treatment,
100 µM DIDS was present during the whole period. Inset:
summarized data. Data are means ± SE; n = 4-6. * P < 0.05 vs. 5.8 mM
[K+]o, #P < 0.05 vs. 75 mM
[K+]o (Bonferroni-Dunn).
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DISCUSSION |
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 intracellular
36Cl. 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+]i
by 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+]o
treatment. 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. 6B), 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-induced
D-[14C]Asp release.
In the current study, high-[K+]o-induced
D-[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 in
D-[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 
-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 induce
D-[14C]Asp release and the release progressed
steadily during 20 min of exposure of cells to high
[K+]o. DIDS (100 µM) abolished the
D-[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 preloaded
D-[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-mediated
D-[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).
| |
ACKNOWLEDGEMENTS |
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.
| |
FOOTNOTES |
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: sun{at}neurosurg.wisc.edu).
The costs of publication of this
article were defrayed in part by the
payment of page charges. The article
must therefore be hereby marked
"advertisement"
in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
10.1152/ajpcell.00478.2001
Received 9 October 2001; accepted in final form 7 December 2001.
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Am J Physiol Cell Physiol 282(5):C1136-C1146
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