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Am J Physiol Cell Physiol 292: C1113-C1122, 2007. First published October 11, 2006; doi:10.1152/ajpcell.00412.2006
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NERVOUS SYSTEM CELL BIOLOGY

Role of Na⁺-K⁺-Cl^– cotransport and Na⁺/Ca²⁺ exchange in mitochondrial dysfunction in astrocytes following in vitro ischemia

Departments of ¹Neurosurgery and ²Physiology, University of Wisconsin School of Medicine and Public Health, Madison, Wisconsin; and ³Department of Molecular Genetics, Biochemistry, and Microbiology, University of Cincinnati, Cincinnati, Ohio

Submitted 1 August 2006 ; accepted in final form 4 October 2006

	ABSTRACT

TOP ABSTRACT EXPERIMENTAL PROCEDURES RESULTS DISCUSSION GRANTS REFERENCES

 
Na⁺-K⁺-Cl^– cotransporter isoform 1 (NKCC1) and reverse mode operation of the Na⁺/Ca²⁺ exchanger (NCX) contribute to intracellular Na⁺ and Ca²⁺ overload in astrocytes following oxygen-glucose deprivation (OGD) and reoxygenation (REOX). Here, we further investigated whether NKCC1 and NCX play a role in mitochondrial Ca²⁺ (Ca_m²⁺) overload and dysfunction. OGD/REOX caused a doubling of mitochondrial-releasable Ca²⁺ (P < 0.05). When NKCC1 was inhibited with bumetanide, the mitochondrial-releasable Ca²⁺ was reduced by 42% (P < 0.05). Genetic ablation of NKCC1 also reduced Ca_m²⁺ accumulation. Moreover, OGD/REOX in NKCC1^+/+ astrocytes caused dissipation of the mitochondrial membrane potential (_m) to 42 ± 3% of controls. In contrast, when NKCC1 was inhibited with bumetanide, depolarization of _m was attenuated significantly (66 ± 10% of controls, P < 0.05). Cells were also subjected to severe in vitro hypoxia by superfusion with a hypoxic, acidic, ion-shifted Ringer buffer (HAIR). HAIR/REOX triggered a secondary, sustained rise in intracellular Ca²⁺ that was attenuated by reversal NCX inhibitor KB-R7943. The hypoxia-mediated increase in Ca_m²⁺ was accompanied by loss of _m and cytochrome c release in NKCC1^+/+ astrocytes. Bumetanide or genetic ablation of NKCC1 attenuated mitochondrial dysfunction and astrocyte death following ischemia. Our study suggests that NKCC1 acting in concert with NCX causes a perturbation of Ca_m²⁺ homeostasis and mitochondrial dysfunction and cell death following in vitro ischemia.

intracellular calcium ion; mitochondrial membrane potential; sodium ion influx; bumetanide; cytochrome c; glial cell death

In in vitro models of ischemia, 1 h of oxygen-glucose deprivation (OGD) does not lead to astrocyte death but does cause a long-lasting decrease in mitochondrial membrane potential (_m) and a loss of mitochondrial cytochrome c (Cyt C; see Ref. 33). Combinations of hypoxia and acidosis or hypoxia/acidosis and ion-shifted conditions in Ringer buffer (HAIR) quickly trigger astrocyte death in vitro (4, 9, 41). The influx of Ca²⁺ in cells is the most significant event in the pathogenesis of ischemic brain damage. Intracellular Ca²⁺ overload triggers mitochondrial dysfunction, leading to immediate or delayed cell death (2, 30). HAIR-induced astrocyte death depends on extracellular Ca²⁺ and is prevented by inhibition of the reverse mode of the Na⁺/Ca²⁺ exchanger (NCX; see Ref. 5). Thus this non-N-methyl-D-aspartate-mediated Ca²⁺ influx may play a significant role in astrocyte damage.

The magnitude and direction of Ca²⁺ flux by NCX depend in part on the Na⁺ electrochemical gradient. Elevated intracellular Na⁺, coupled with membrane depolarization, could reverse the forward direction of NCX and trigger a detrimental Ca²⁺-dependent signal transduction cascade (39). Na⁺/H⁺ exchange has been implicated in cytosolic Na⁺ loading and reversal of NCX in astrocytes following hypoxia and ischemia (5, 21, 22). We recently reported that Na⁺-K⁺-Cl^– cotransporter isoform 1 (NKCC1) contributes to an approximately fourfold increase in intracellular Na⁺ concentration ([Na⁺]_i) in astrocytes following OGD (23). Reversal of NCX as a result of this Na⁺ loading leads to a significant accumulation of Ca²⁺ in the endoplasmic reticulum (ER) and mitochondria (23). However, it is unknown whether this alteration in Ca²⁺ handling plays a role in mitochondrial dysfunction in astrocytes following ischemia.

In the present study, we report that NKCC1 and NCX were in part responsible for loss of mitochondrial Ca²⁺ (Ca_m²⁺) homeostasis, mitochondrial depolarization, and Cyt C release in astrocytes after hypoxia/ischemia.

	EXPERIMENTAL PROCEDURES

TOP ABSTRACT EXPERIMENTAL PROCEDURES RESULTS DISCUSSION GRANTS REFERENCES

 
Materials. Eagle's modified essential medium (EMEM) and Hanks’ balanced salt solution (HBSS) were from Mediatech Cellgro (Herndon, VA). FBS was obtained from Hyclone Laboratories (Logan, UT). Collagen type I was from Collaborative Biomedical Products (Bedford, MA). FCCP, oligomycin, gramicidin, monensin, bumetanide, cyclosporine A (CsA), bongkrekic acid, and propidium iodide (PI) were purchased from Sigma Chemical (St. Louis, MO). Sodium-binding benzofuran isophthalate-AM (SBFI-AM), fluorescein-methylene-iminodiacetate-AM, 5,5',6,6'-tetrachloro-1,1',3,3'-tetraethyl-benzimidazo-carbocyanine iodine (JC-1), fura 2-AM, MitoTracker Red CMXRos, MitoTracker Green FM, and rhod 2-AM were from Invitrogen (Eugene, OR). KB-R7943 was from Tocris (Ellisville, MO). Anti-Cyt C monoclonal antibody (clone 6H2.B4) was from PharMingen (San Jose, CA), and anti-NCX1 monoclonal antibody was from Swant (Bellinzona, Switzerland). ATP kit was from Perkin-Elmer Life Sciences (Wellesley, MA).

Animals and genotype analysis. NKCC1 homozygous mutant and wild-type mice were obtained by breeding gene-targeted NKCC1 heterozygous mutant mice (13), and genotypes were determined by PCR analysis of DNA from tail biopsies as described previously (40).

Primary culture of mouse cortical astrocytes. Dissociated cortical astrocyte cultures were established as described before (40). Cerebral cortices were removed from 1-day-old NKCC1^+/+ or NKCC1^–/– mice. The cortices were incubated in a trypsin solution (0.25 mg/ml of HBSS) for 25 min at 37°C. The dissociated cells were rinsed and resuspended in EMEM containing 10% FBS. Viable cells (1 x 10⁴ cells/well) were plated in six-well plates coated with collagen type 1. Cultures were maintained in a 5% CO₂ atmosphere at 37°C. To obtain morphologically differentiated astrocytes, confluent cultures (7 days in culture) were treated with EMEM containing 0.25 mM dibutyryl-cAMP (DBcAMP) to induce differentiation. DBcAMP has been widely used to mimic neuronal influences on astrocyte differentiation (42). Experiments were routinely performed in day 10–21 cultures.

In vitro hypoxia/ischemia. NKCC1^+/+ or NKCC1^–/– astrocytes on cover slips were rinsed two times with an isotonic OGD solution (pH 7.4) containing (in mM, at 37°C): 0 glucose, 26 NaHCO₃, 120 NaCl, 5.36 KCl, 0.33 Na₂HPO₄, 0.44 KH₂PO4, 1.27 CaCl₂, and 0.81 MgSO₄. Cells were incubated in 1.0 ml of the OGD solution in a hypoxic incubator for 2 h (model 3130; Forma Scientific, Marietta, OH) with 94% N₂, 1% O₂, and 5% CO₂. An orbital shaker (model M48215; Thermolyne, Dubuque, IA) in the hypoxic chamber was used to facilitate equilibration of the hypoxic gases during the initial 30-min OGD. Normoxic control cells were incubated in 5% CO₂ and atmospheric air in an isotonic control solution for 2 h. For reoxygenation (REOX), OGD-treated cells were superfused with HCO₃^–-MEM buffer containing 5.5 mM glucose or placed in normoxic HEPES-MEM (38) for 60 min.

Astrocytes were also subjected to a more severe in vitro ischemic condition by exposing cells to a HAIR buffer, which conditions mimic the extracellular space of ischemic tissue (5). Astrocytes on cover slips were placed in an imaging chamber and continuously superfused (1 ml/min) with HCO₃^–-MEM at 37°C during a 5-min baseline perfusion. The cells were then superfused with HAIR buffer (bubbled with a gas containing 15% CO₂-85% N₂, pH 6.6) for 5–20 min. Oxygen content of the HAIR buffer was determined via a flow-through electrode (model 16–730; Microelectrodes, Bedford, NH) immediately before entering the chamber. Oxygen content dropped to 3.3 ± 1.6% within 1 min and averaged at 1.2 ± 0.4% after 5 min. HAIR buffer contained (in mM): 34 NaCl, 13 NaHCO₃, 3 sodium gluconate, 65 potassium gluconate, 38 N-methyl-D-glucamine chloride, 1 NaH₂PO₄, 0.13 CaCl₂, 1.5 MgCl₂, and 5.5 glucose (5). The ionic shifts of K⁺, Na⁺, and Ca²⁺ simulate in vivo ischemic conditions (5). For REOX, cells were superfused with HCO₃^–-MEM at 37°C for 30–45 min. For cell death studies, HAIR treatment was 20 min followed by 120 min of REOX.

Intracellular Ca²⁺ measurement. Astrocytes were incubated with 5 µM fura 2-AM for 30–120 min at 37°C (23). The cells were washed, and the cover slips were placed in the imaging chamber and superfused with HCO₃^–-MEM at 37°C. With the use of a Nikon TE 300 inverted epifluorescence microscope (x40 oil immersion objective lens), astrocytes were excited every 10–30 s at 340 and 380 nm, and the emission fluorescence at 510 nm was recorded. Images were collected and analyzed with the MetaFluor (Molecular Devices, Sunnyvale, CA) image-processing software. At the end of each experiment, the cells were exposed to 1 mM MnCl₂ in Ca²⁺-free HEPES-MEM. The Ca²⁺-insensitive fluorescence was subtracted from each wavelength before calculations (23). The MnCl₂-corrected 340- to 380-nm emission ratios were converted to concentration as described previously (23). To assess sequestration of Ca²⁺ by mitochondria, Ca²⁺ release from mitochondria was measured following application of FCCP (10 µM) alone or in combination with oligomycin (2.5 µg/ml). In the case of assessing ER Ca²⁺ stores, Ca²⁺ release was induced by 1 µM thapsigargin.

Ca_m²⁺ measurement. Astrocytes were incubated with 3 µM rhod 2-AM (reduced with a minimum of sodium borahydride) and 200 nM MitoTracker green FM for 90–120 min in buffers supplemented with 1 mM sodium succinate (27). The cells were placed in a perfusion chamber (Warner Instruments, Hamden, CT) on the stage of a Leica (Exton, PA) DMIRE2 confocal microscope and superfused with HCO₃^–-MEM at 37°C. Cells were visualized with a x40 oil-immersion objective. Cells (5–6 in the field) were scanned sequentially for MitoTracker green (excitation 488 nm argon laser line, emission 500–545 nm) and rhod 2 (excitation 543 HeNe laser, emission 544–677). The MitoTracker green signal was used to maintain focus before each sequential scan. Sequential scans were analyzed using Leica confocal software. Average grayscale values were collected from regions of interest drawn around perinuclear mitochondrial clusters exhibiting colocalization of MitoTracker green and rhod 2. Ca_m²⁺ values were expressed as relative change of rhod 2 signals from the baseline values.

Intracellular Na⁺ measurement. [Na⁺]_i was measured in astrocytes with the fluorescent dye SBFI-AM as described previously (40). Astrocytes were loaded with 10 µM SBFI-AM at 37°C in HEPES-MEM containing 0.05% pluronic acid for 45–120 min. The cells were washed, and the cover slips were placed in the imaging chamber containing HEPES-MEM at ambient temperature. Astrocytes were excited every 10 s at 345 and 385 nm, and the emission fluorescence at 510 nm was recorded with a Nikon TE 300 inverted epifluorescence microscope (x40 oil immersion objective lens). Absolute [Na⁺]_i was determined for each cell as described previously (35, 40).

Imaging of mitochondrial _m. The fluorescent probe JC-1 was used to investigate mitochondrial _m in astrocytes (45). Astrocytes were loaded with 9 µM JC-1 for 30–120 min at 37°C and visualized using a Nikon TE 300 inverted epifluorescence microscope and a x60 oil immersion objective. Astrocytes were excited at 480 nm. Emission fluorescence images recorded at 535 nm represented the monomer and 640 nm for JC-1 aggregates. The ratio of the aggregate to monomer fluorescence was measured in perinuclear mitochondrial regions and reflected _m. Maximal _m dissipation was induced by FCCP (10 µM) alone or in combination with oligomycin (2.5 µg/ml) at the end of each experiment. _m in astrocytes following OGD/REOX was expressed as the percentage of the maximal FCCP-induced change in normoxic controls.

Cyt C immunofluorescence staining. Cyt C release was determined by a specific antibody against Cyt C. Briefly, cells on cover slips were fixed in 4% paraformaldehyde in PBS for 10 min. After being rinsed, cells were incubated with a blocking solution for 20 min followed by application of anti-Cyt C antibody (1:100 diluted in a blocking buffer) for 1 h at room temperature. After being rinsed in PBS, slices were incubated with goat anti-mouse FITC-conjugated IgG (1:100) for 1 h at 37°C. Fluorescence images were obtained using a Leica DMIRE2 inverted confocal laser-scanning microscope (x63) and Leica confocal software. Samples were excited at 488 nm (argon/krypton), and the emission fluorescence was recorded at 500–535 nm.

MitoTracker Dye staining. MitoTracker Red CMXRos was used to stain for mitochondria. Briefly, after removing the cell culture medium, cells were incubated with fresh culture medium containing 200 nM MitoTracker Red CMXRos at 37°C for 15 min. After incubation, the cells were washed with medium three times and further stained for Cyt C as described above. Fluorescence images were obtained using a Leica DMIRE2 inverted confocal laser-scanning microscope (x63) and Leica confocal software. MitoTracker signals were obtained by exciting samples at 543 nm (GreNe), and the emission fluorescence was recorded at 555–620 nm. In double-staining studies (Cyt C/MitoTracker), the samples were excited and emissions were collected sequentially.

Measurement of intracellular ATP content. Intracellular ATP content was measured using a luminescence ATP detection assay as described before (23). After 2 h OGD, 2 h OGD plus 1 h REOX, 5 min HAIR, or 5 min HAIR plus 3 min REOX, culture medium was removed, and 100 µl fresh OGD or normoxic control buffer was added to each well. Lysis solution (50 µl) was added, and plates were gently swirled for 5 min. Aliquots of cell lysates (diluted 1:10) were added to 50 µl substrate buffer. Luminescence was detected with an Lmax II Luminometer (Molecular Devices). In each experiment, ATP levels were determined using ATP standards run concurrently with samples. Protein content in cell lysates was determined in each sample using the bicinchoninic acid method (Pierce, Rockford, IL). Intracellular ATP content is expressed as nanomoles per milligram protein.

Cell death determination. Ca²⁺-insensitive fura 2 fluorescence was used as an indication of changes in intracellular Ca²⁺ concentration ([Ca²⁺]_i) and as a vital dye to monitor cell death. Cell death was evidenced by a loss of fura 2 from the cell and an abrupt decline in 340-nm excitable fluorescence (Ca²⁺ insensitive). Loss of dye has been shown to reflect loss of membrane integrity and the onset of cell death (5, 12). To further confirm cell death, in some studies, cells were stained with PI following 20 min HAIR and 120 min REOX.

Gel electrophoresis and Western blotting. Cultured cells were washed with ice-cold PBS (pH 7.4) that contained 2 mM EDTA and protease inhibitors. Cells were collected by scraping from the six-well plates. The protein content was measured with the bicinchoninic acid method. Samples and prestained molecular mass markers (Bio-Rad, Hercules, CA) were denatured in SDS reducing buffer (1:2 by volume; Bio-Rad) and heated at 37°C for 15 min before gel electrophoresis. The protein samples (30 µg lysate protein) were loaded and separated by SDS-PAGE (6%). After being transferred to a polyvinylidene difluoride membrane, the blots were incubated in 7.5% nonfat dry milk in TBS and then incubated with a primary antibody, anti-NCX1 monoclonal antibody (1:1,000). For a protein-loading control, the blot was restripped and reprobed with anti-glial fibrillary acidic protein (GFAP) monoclonal antibody (1:4,000). After being rinsed, the blots were incubated with horseradish peroxidase-conjugated secondary IgG. Bound antibody was visualized using the enhanced chemiluminescence assay (Amersham).

To elucidate any quantitative changes in NCX1, densitometry measurements of each protein band were performed. Protein bands on the blot were selected, and average pixel was recorded with Un-Scan-It Gel (Silk Scientific, Orem, UT). A ratio of NCX1 and GFAP was calculated in normoxic control or OGD/REOX samples.

Statistics. Routinely, parameters for live cell imaging were measured in 20 cells from a cover slip and averaged. Throughout the study, n values represented numbers of cultures used in each experiment. Mean ± SE values for each experimental group were reported as a gauge of the accuracy of the calculated mean. Statistical significance was determined by the Mann-Whitney nonparametric test at a confidence of 95% (P < 0.05).

	RESULTS

TOP ABSTRACT EXPERIMENTAL PROCEDURES RESULTS DISCUSSION GRANTS REFERENCES

 
NKCC1-mediated Na⁺ influx is increased in astrocytes following mild hypoxia. Under normoxic conditions, [Na⁺]_i was stable and did not differ significantly between NKCC1^+/+ and NKCC1^–/– astrocytes. Furthermore, 2 h OGD did not alter [Na⁺]_i in either NKCC1^+/+ or NKCC1^–/– astrocytes (Fig. 1A). Upon REOX following 2 h OGD, [Na⁺]_i in NKCC1^+/+ astrocytes increased at a maximum rate of 5.0 ± 2.2 mM/min and reached a value of 78 ± 11 mM at 60 min (Fig. 1, A-C). However, in the presence of the NKCC1 inhibitor bumetanide, the maximum rise in [Na⁺]_i was reduced to 0.62 ± 0.28 mM/min in NKCC1^+/+ astrocytes, and the maximal value of [Na⁺]_i was 27 ± 4 mM. Genetic ablation of NKCC1 also attenuated the maximum Na⁺ influx rate (1.23 ± 0.78 mM/min) in response to OGD/REOX, and [Na⁺]_i was only 42 ± 2 mM at 60 min REOX (Fig. 1, A-C). These findings indicate that NKCC1 activity is increased following OGD/REOX and contributes to Na⁺ accumulation in astrocytes. The remaining Na⁺ rise mediated by OGD/REOX in NKCC1^–/– astrocytes may be via activation of Na⁺/H⁺ exchange (see data in Fig. 4).

View larger version (19K): [in this window] [in a new window]   Fig. 1. Changes of intracellular Na+ concentration ([Na+]i) in astrocytes following oxygen-glucose deprivation (OGD)/reoxygenation (REOX). A: [Na+]i was determined every 5 min during REOX in Na+-K+-Cl– cotransporter isoform 1 (NKCC1)+/+, NKCC1–/–, or NKCC1+/+ astrocytes treated with bumetanide (Bum) following 2 h OGD. Controls (Con) were sister NKCC1+/+ astrocytes subjected to 3 h normoxic incubation. Bum (5 µM) was present throughout the time course in the drug treatment experiments. B: summary of maximal values of [Na+]i during 60 min REOX. C: summary of maximum rate of [Na+]i increase during 60 min REOX. Data are means ± SE; n = 4–5 experiments. P < 0.05 vs. Con (*), vs. NKCC1+/+ REOX (#), and vs. NKCC1–/– REOX ($). D: changes in [Na+]i were determined in the presence of ouabain (1 mM) during normoxia or during 60 min REOX following 2 h OGD in NKCC1+/+ astrocytes. Arrow: addition of 1 mM ouabain at 1 min of REOX. Data are means ± SE; n = 3–5 experiments.

View larger version (28K): [in this window] [in a new window]   Fig. 2. Changes in mitochondrial Ca2+ (Cam2+) in astrocytes following OGD/REOX. A: representative tracings of intracellular Ca2+ concentration ([Ca2+]i) in single NKCC1+/+, NKCC1–/– following 2 h OGD/1 h REOX. Arrow, FCCP/oligomycin triggered release of Cam2+. Con were done in sister NKCC1+/+ astrocytes subjected to 3 h normoxic incubation. B: summary data of the integrated area under the intracellular Ca2+ transient (IACa2+) of A. Data are means ± SE; n = 4–5 experiments. P < 0.05 vs. Con (*) and vs. NKCC1+/+ OGD/REOX (#). Bum (5 µM) was present throughout hypoxic, acidic, ion-shifted Ringer buffer (HAIR)/REOX in the drug-treated group. C: MitoTracker (green in a) and rhod 2 (red in b) images were shown in NKCC1+/+ astrocytes at the end of 2 h OGD and at 60 min REOX (c and d). Scale bar = 12.3 µm. D: average tracings of relative change in rhod 2 fluorescence during 60 min REOX following 2 h OGD in NKCC1+/+ and NKCC1–/– astrocytes. Arrow: response of rhod 2 fluorescence to FCCP. E: summary data. Data are means ± SE; n = 4–5 experiments. P < 0.05 vs. control (*) and vs. NKCC1+/+ OGD/REOX (#).

View larger version (41K): [in this window] [in a new window]   Fig. 3. Changes in membrane potential (m) in astrocytes following OGD/REOX. A: 5,5',6,6'-tetrachloro-1,1',3,3'-tetraethyl-benzimidazo-carbocyanine iodine (JC-1) aggregate (a) and monomer (b) fluorescence in NKCC1+/+ cells under normoxic control conditions. Insets in A: changes of JC-1 fluorescence following 3 min of FCCP (10 µM). Scale bar = 12.3 µm. B: representative tracing of changes of the JC-1 ratios. C: summary data of m. Maximal m dissipation was induced by FCCP in the presence of oligomycin in each experiment. In normoxic controls, the mean baseline ratio was considered to be 100%, and the mean ratio obtained after FCCP treatment was 0%. The JC-1 ratios were normalized to the initial baseline ratios. In OGD studies, JC-1 ratios were obtained at the start of REOX. Therefore, the FCCP-induced decrease in the JC-1 would be less when the mitochondria were partially depolarized following OGD. m in astrocytes following OGD/REOX was normalized to the normoxic controls and expressed as %normoxic controls. Data are means ± SE; n = 4–5 experiments. #P < 0.05 vs. NKCC1+/+ OGD/REOX. D: average change in m during 60 min REOX following 2 h OGD in NKCC1+/+ astrocytes and NKCC1+/+ astrocytes treated with 5 µM Bum.

View larger version (31K): [in this window] [in a new window]   Fig. 4. Changes in [Na+]i in astrocytes following HAIR/REOX. A: [Na+]i was monitored in single NKCC1+/+, NKCC1–/–, or NKCC1+/+ astrocytes treated with Bum during 5 min HAIR and followed by 30 min HEPES-MEM superfusion. B: summary of data. Cells were treated with 5 µM Bum 30 min before hypoxia and during HAIR/REOX. Data are means ± SE; n = 5–7 experiments. C: representative tracings of changes in intracellular Na+. HOE-642 (1 µM) or Bum (5 µM) was used. D: summary data of peak value in [Na+]i at the beginning of REOX and at the end of 30 min REOX. Data are means ± SE; n = 5–6 experiments. E: [Na+]i was monitored in single NKCC1+/+ astrocytes following HAIR/REOX with or without 1 mM ouabain. *P < 0.05 vs. baseline. #P < 0.01 vs. NKCC1+/+ HAIR/REOX.

Table 1 demonstrates that the concentration of intracellular ATP was reduced by 20% at the end of 2 h OGD and 20% following 5 min HAIR treatment. However, at 3 min REOX following HAIR or 1 h REOX following 2 h OGD, intracellular ATP recovered nearly to the control level. This further implies that intracellular ATP is not a factor that would limit the functions of either primary active ion transport by the Na⁺-K⁺-ATPase, which consumes metabolic energy derived from the hydrolysis of ATP, or secondary active ion transport systems such as NKCC1 and Na⁺/H⁺ exchanger isoform 1 (NHE1), which are driven by energy stored in the ion gradients.

The accumulation of Ca_m²⁺ during OGD/REOX was then directly confirmed by using the fluorescence of the low-affinity Ca²⁺ dye rhod 2. Figure 2C shows that a MitoTracker green dye (green in Fig. 2Ca) was colocalized with rhod 2 at 2 h OGD (red in Fig. 2Cb). Rhod 2 signals were lost upon FCCP treatment (Fig. 2C, inset). After 60 min of REOX, the rhod 2 dye fluorescence increased (Fig. 2Cd), indicating loading of mitochondria with Ca²⁺. Rhod 2 fluorescence increased by approximately sevenfold in NKCC1^+/+ astrocytes at 60 min REOX (Fig. 2, D and E). In contrast, NKCC1^–/– astrocytes only exhibited 30% increase in Ca_m²⁺ (Fig. 2, D and E).

Disruption of _m during OGD/REOX. Ca_m²⁺ overload can trigger mitochondrial dysfunction (6). Therefore, we investigated whether _m was affected following in vitro ischemia. Changes of _m were monitored by the mitochondrial dye JC-1. Figure 3A illustrates the distribution of JC-1 punctate aggregate (red in Fig. 3Aa) and monomer (green in Fig. 3Ab) fluorescence in NKCC1^+/+ astrocytes under normoxic control conditions. As a positive control, NKCC1^+/+ astrocytes were exposed to the mitochondrial uncoupler FCCP for 1 min. JC-1 green fluorescence was increased and the red aggregate signal was largely eliminated (Fig. 3A, insets), reflecting depolarization of _m.

Figure 3, B and C, shows changes of the JC-1 ratio under different experimental conditions. A sharp reduction of _m in NKCC1^+/+ astrocytes under normoxic control conditions was recorded when mitochondrial function was inhibited by FCCP and oligomycin (Fig. 3, B and C). On the other hand, following OGD/REOX, the FCCP-induced decrease in _m in NKCC1^+/+ astrocytes was reduced to 42% of the controls (Fig. 3, B and C). The reduction in response to FCCP implies a partial depolarization of the mitochondria membrane. Inhibition of NKCC1 activity with 5 µM bumetanide did not affect _m under normoxic conditions but significantly attenuated the OGD/REOX-induced depolarization of _m, which remained at 66% of control levels (Fig. 3, B and C). Likewise, _m was depolarized only to 75% of control values in NKCC1^–/– astrocytes. The change in FCCP-sensitive _m was also monitored during 60 min REOX following 2 h OGD (Fig. 3D). In either NKCC1^+/+ astrocytes or in NKCC1^+/+ astrocytes treated with bumetanide, _m was depolarized to 40% of control values at the end of 2 h OGD. No recovery of _m was found in NKCC1^+/+ astrocytes during 60 min REOX. However, in NKCC1^+/+ astrocytes treated with bumetanide, _m increased to 88% of control within 15 min REOX. It was 50% more polarized at 60 min REOX compared with untreated NKCC1^+/+ astrocytes. Taken together, these findings imply that NKCC1 activity in part contributes to mitochondrial depolarization following hypoxia.

Sustained increase in [Na⁺]_i during HAIR/REOX in part depends on NKCC1 activity. To further determine the role of NKCC1 in deregulation of astrocyte Na⁺ homeostasis, we investigated changes of Na⁺ in astrocytes under HAIR conditions. As shown in Fig. 4A, the basal level of [Na⁺]_i in NKCC1^+/+ astrocytes was 13.5 ± 4.7 mM. Exposing NKCC1^+/+ astrocytes to HAIR for 5 min caused a decrease in [Na⁺]_i (6.6 ± 2.3 mM), which may result from the reduced Na⁺ concentration in the HAIR buffer. Interestingly, [Na⁺]_i in NKCC1^–/– astrocytes decreased further to a level of 1.8 ± 0.7 mM, which was significantly lower than NKCC1^+/+ astrocytes (P < 0.01). This suggests that NKCC1 activity plays a role in maintaining [Na⁺]_i under HAIR conditions (Fig. 4, A and B).

Upon REOX following 5 min HAIR, [Na⁺]_i in NKCC1^+/+ astrocytes increased at a rate of 42 mM/min and reached an average peak value of 75 ± 33 mM (Fig. 4, A and B). [Na⁺]_i then declined at 5 min and plateaued (56 ± 27 mM) during the 30 min REOX. This new steady-state level of [Na⁺]_i was higher than baseline values (P < 0.05). However, in NKCC1^–/– astrocytes, the rate of [Na⁺]_i rise during REOX was reduced by 50% (22 mM/min) and peaked at a value (38 ± 4 mM) that was significantly lower than in NKCC1^+/+ astrocytes (Fig. 4, A and B; P < 0.05). [Na⁺]_i in NKCC1^–/– astrocytes also reached a significantly lower steady state following REOX (18 ± 2 mM; P < 0.005). Inhibition of NKCC1 activity by bumetanide significantly reduced both peak and steady-state [Na⁺]_i values in NKCC1^+/+ astrocytes (30.0 ± 3.3 and 22.5 ± 3.4 mM, respectively).

As shown in Fig. 4, A and B, inhibition of NKCC1 only attenuated the Na⁺ entry by 60% following HAIR. This suggests that other Na⁺ entry pathways are also involved in this Na⁺ deregulation. Na⁺/H⁺ exchange activity in astrocytes is elevated during REOX following OGD or HAIR (9, 21). To investigate whether the Na⁺ entry that was insensitive to NKCC1 inhibition is via Na⁺/H⁺ exchange, we determined [Na⁺]_i in the presence of HOE-642, which is a potent inhibitor of NHE1. As shown in Fig. 4C, in the presence of HOE-642, REOX-induced rise in [Na⁺]_i was blocked by 40%. Inhibition of both NKCC1 and NHE1 resulted in 85% reduction of Na⁺ entry during REOX (Fig. 4, C and D). It is not clear why either inhibition of NHE1 or NKCC1 leads to a similar reduction in Na⁺ entry. One possible explanation is that they are localized in the same cellular microdomain complex and stimulated in response to the same stimulus.

We speculated that the initial decline in [Na⁺]_i following its peak rise during REOX may result from Na⁺-K⁺-ATPase-mediated Na⁺ extrusion. Inhibition of Na⁺-K⁺-ATPase with 1 mM ouabain during REOX triggered a sustained rise in Na⁺ (136 ± 13 mM) in NKCC1^+/+ astrocytes during 30 min REOX (Fig. 4E). The initial decline in Na⁺ (beginning at 5 min REOX) was absent in the presence of ouabain. This implies that Na⁺-K⁺-ATPase is functional following hypoxia but is only able to partially regulate intracellular Na⁺ during REOX. Therefore, the collapse of Na⁺ homeostasis results from an imbalance between NKCC1- and NHE1-mediated accelerated Na⁺ entry and Na⁺-K⁺-ATPase-mediated Na⁺ extrusion.

NKCC1- and NCX-dependent changes of intracellular Ca²⁺ following HAIR/REOX. Because REOX triggered such a significant increase in Na⁺ in astrocytes following HAIR, we anticipated that the latter change would affect Ca²⁺ homeostasis by reversing NCX. We monitored changes in intracellular Ca²⁺ during HAIR and REOX. Figure 5A shows an immediate transient rise in [Ca²⁺]_i in NKCC1^+/+ astrocytes at 1–2 min HAIR. The rise in [Ca²⁺]_i was sustained during the remaining 15 min of hypoxia (Fig. 5A). Similar changes were found in NKCC1^–/– astrocytes. REOX triggered a secondary rise in [Ca²⁺]_i during 45 min REOX in NKCC1^+/+ astrocytes (Fig. 5, A and B). In contrast, NKCC1^–/– astrocytes exhibited a significantly reduced rise in [Ca²⁺]_i (130 ± 40 vs. 262 ± 26 nM, P < 0.05, Fig. 5, A and B).

View larger version (37K): [in this window] [in a new window]   Fig. 5. Increase in [Ca2+]i following HAIR/REOX depends on NKCC1 and Na+/Ca2+ exchanger (NCX) activity. A: [Ca2+]i was monitored in single NKCC1+/+ or NKCC1–/– astrocytes during 15 min HAIR and 45 min REOX. In studies involving KB-R7943 (KB), 10 µM KB was present 30 min before HAIR and during HAIR/REOX. B: summary data of [Ca2+] i at 45 min REOX. Data are means ± SE; n = 4–5 experiments. P < 0.05 vs. control (*) and vs. NKCC1+/+ HAIR/REOX (#). C: representative Western blot and summarized densitometry data. Cells were exposed to 2 h OGD and 1 h REOX. Cell lysates were separated electrophoretically as described previously (21). NCX1 and glial fibrillary acidic protein (GFAP) proteins are shown on the same blot. Summarized data are means ± SE; n = 3 experiments.

NKCC1 activity contributes to mitochondrial dysfunction in astrocytes following HAIR/REOX. Figure 6A shows the changes in the ratio of the JC-1 fluorescence observed in NKCC1^+/+ and NKCC1^–/– astrocytes. HAIR treatment (15 min) did not trigger any changes in the JC-1 ratio. Upon REOX, the JC-1 fluorescence ratio decreased sharply and reached a plateau after 7 min (Fig. 6A), indicating a loss in _m. _m remained depressed over the remaining period of REOX (37 ± 4% of control; Fig. 6B). In contrast to NKCC1^+/+ astrocytes, _m was significantly less depolarized in NKCC1^–/– astrocytes (69 ± 2% of control; Fig. 6B). Interestingly, when NKCC1^+/+ astrocytes were treated with the permeability transition pore (PTP) inhibitor CsA (4 µM), there was no significant depolarization in _m during REOX (85 ± 1% of control values; Fig. 6B, P < 0.05). Because CsA also affects phosphatase 2B, another PTP inhibitor, bongrekic acid, was examined. Similar protective effects on _m (91% of control values) were observed with 4 µM bongrekic acid. These findings further indicate that NKCC1 activity affects mitochondrial function in astrocytes following hypoxia.

View larger version (13K): [in this window] [in a new window]   Fig. 6. NKCC1 activity affects dissipation of m, Cam2+, and endoplasmic reticulum (ER) Ca2+ in astrocytes following HAIR/REOX. A: representative changes in normalized JC-1 fluorescence ratio during 15 min HAIR/45 min REOX in single NKCC1+/+ or NKCC1–/– astrocytes. In studies with cyclosporine A (CsA), 4 µM CsA was present 30 min before HAIR and during HAIR/REOX. B: summary of m data. *P < 0.05 vs. NKCC1+/+ HAIR/REOX. C: representative tracings of relative rhod 2 fluorescence during 15 min HAIR/45 min REOX. D: summary of relative rhod 2 fluorescence. Data are means ± SE; n = 3–5 experiments. P < 0.05 vs. baseline (*) and vs. NKCC1+/+ HAIR/REOX (#). E: [Ca2+]i was monitored in NKCC1+/+ and NKCC1–/– astrocytes during 20 min HAIR and 15 min REOX when cells were exposed to 1 µM thapsigargin at the beginning of REOX (arrow). F: summary data of peak [Ca2+]i in response to thapsigargin. Data are means ± SE; n = 4–5 experiments. *P < 0.001 vs. control. #P < 0.05 vs. NKCC1+/+ HAIR/REOX.

We speculated that there was more Ca²⁺ sequestered in the ER of NKCC1^+/+ than NKCC1^–/– astrocytes following HAIR/REOX. To test this, astrocytes were subjected to 1 µM thapsigargin at the beginning of REOX (Fig. 6, E and F). Thapsigargin caused a transient rise in [Ca²⁺]_i to a peak value of 597 ± 29 nM in NKCC1^+/+ astrocytes, which was significantly higher than in normoxic controls (366 ± 19 nM). In contrast, the thapsigargin-induced Ca²⁺ transient in NKCC1^–/– astrocytes was reduced to 325 ± 18 nM (P < 0.05), which was not different from controls. Taken together, these findings demonstrate that NKCC1 activity is in part responsible for mitochondrial and ER Ca²⁺ overload following HAIR/REOX.

Inhibition of NKCC1 activity reduces Cyt C release and cell death following HAIR/REOX. The increase in Ca_m²⁺ and decrease in _m may be accompanied by altered mitochondrial membrane integrity and cell death. Therefore, we first investigated the release of Cyt C from mitochondria. In Fig. 7, a punctate perinuclear pattern for Cyt C was seen in mitochondria of normoxic NKCC1^+/+ astrocytes, which colocalized with the mitochondrial dye MitoTracker Red CMXRos (Fig. 7, B and C). As a positive control, NKCC1^+/+ astrocytes were exposed to 10 µM FCCP for 1 min. In this experiment, Cyt C was released in the cytosol, and the immunoreactivity signals became diffuse (Fig. 7D). Some Cyt C was released into the nuclei (Fig. 7D). Such a phenomenon has also been seen in apoptotic muscle cells (19). As shown in Fig. 7E, 5 min HAIR/3 min REOX caused a redistribution of Cyt C in NKCC1^+/+ astrocytes that was similar to that observed in FCCP-treated cells. The loss of _m at 3 min REOX in these studies was confirmed by monitoring changes of the JC-1 ratio (data not shown). Cyt C staining was diffuse, and the intensity of immunoreactive signal in the cytosol was greater. However, when NKCC1 was either inhibited with 5 µM bumetanide or genetically ablated, Cyt C release from mitochondria was not elicited by HAIR/REOX (Fig. 7, F and G). Taken together, these data clearly demonstrate that mitochondrial membrane integrity was damaged following HAIR/REOX and that loss or inhibition of NKCC1 reduced this damage.

View larger version (98K): [in this window] [in a new window]   Fig. 7. Inhibition of NKCC1 activity reduces HAIR/REOX-mediated cytochrome c (Cyt C) release and cell death. The release of Cyt C in astrocytes was visualized by staining with an anti-Cyt C antibody. Colocalization of immunoreactive signals of Cyt C and MitoTracker Red CMXRos was observed in normoxic NKCC1+/+ control astrocytes incubated for 8 min in HEPES-MEM buffers (A-C). A: Cyt C staining. B: MitoTracker red CMXRos staining. C: overlay images of A and B. D: as a positive control, Cyt C release in NKCC1+/+ astrocytes was triggered with 10 µM FCCP for 1 min. E: 5 min HAIR/3 min REOX induced Cyt C release in NKCC1+/+ astrocytes. F: to study the effects of NKCC1 inhibition, NKCC1+/+ astrocytes were incubated in the presence of 5 µM Bum following 5 min HAIR/3 min REOX. G: less Cyt C was released in NKCC1–/– astrocytes following 5 min HAIR/3 min REOX. Arrow: Cyt C release from mitochondria. *Nuclei. Scale bar = 20 µm. Images shown are representative of 4 experiments. H: fluorescent staining of live (fura 2, 380 nm, green) and dead [propidium iodide (PI), red] NKCC1+/+ and NKCC1–/– astrocytes following 20 min HAIR and 120 min REOX. Scale bar = 25 µm. I: summary of cell death with a loss of fura 2 signals. Data are means ± SE; n = 4–6 experiments. *P < 0.001 vs. NKCC1+/+ HAIR/REOX.

	DISCUSSION

TOP ABSTRACT EXPERIMENTAL PROCEDURES RESULTS DISCUSSION GRANTS REFERENCES

 
Sustained loss of Na⁺ homeostasis in astrocytes following in vitro hypoxia/ischemia. In the present study, two hypoxic insults were applied, OGD and HAIR. The onset of REOX under both conditions triggered rapid intracellular Na⁺ overload in astrocytes. The rise in [Na⁺]_i was sustained during 30–60 min REOX. Loss of NKCC1 activity either by treatment with the inhibitor bumetanide or by genetic ablation blocked 64% of the rise in Na⁺. NKCC1 mediates (17) chemical gradients of the three transported ions (37). We have previously shown that NKCC1 activity in astrocytes was increased during REOX via protein phosphorylation (23), which is a major regulatory mechanism for stimulation of NKCC1 activity (14). The current study further demonstrates that stimulation of NKCC1 activity evokes excessive Na⁺ entry and disrupts astrocyte Na⁺ homeostasis following hypoxia/ischemia. However, it remains to be studied whether the OGD-mediated stimulation of NKCC1 results from activation of regulatory kinases of NKCC (10, 17), such as Ste20p-related proline alanine-rich kinase, oxidative response kinase 1, WNK1 and WNK4 (with no lysine K kinase), or altered activities of phosphatases.

Interestingly, following HAIR treatment, REOX triggered a rise in intracellular Na⁺ that had two phases (a peak increase and a sustained elevation), although the first phase was absent in OGD-treated cells. Although inhibition of NKCC1-mediated Na⁺ entry significantly reduced the amplitude of the sustained accumulation of intracellular Na⁺, it did not eliminate the first phase of the increased Na⁺. This implies that Na⁺ entry in the initial phase was not mediated by NKCC1. We recently reported that NHE1 activity in mouse cortical astrocytes was increased by 75% upon REOX following OGD (21, 22). In rat cortical astrocytes, REOX following 22 min HAIR treatment triggered a rapid alkaline shift in intracellular pH, which was mediated by NHE1 activity (5). These findings led us to speculate that the peak rise in [Na⁺]_i during onset of REOX may result from NHE1-mediated Na⁺ entry. Blocking of NHE1 activity with the potent inhibitor HOE-642 abolished the peak rise in Na⁺. In addition, inhibition of both NHE1 and NKCC1 eliminated 90% of the intracellular Na⁺ accumulation during REOX, but they are not additive. These data suggest that both pathways independently contribute to Na⁺ entry during REOX. However, it is unknown whether NKCC1 and NHE1 are thermodynamically linked, and future studies are needed to address this issue.

Functions of NKCC1 and NHE1 have been shown to require cellular ATP (8, 37). Our results indicated that intracellular ATP content recovered to the control level during REOX following either 2 h OGD or 5 min HAIR. Thus sufficient intracellular ATP levels are available to support accelerated activities of NKCC1 and NHE1 during REOX.

Na⁺-K⁺-ATPase activity removed 50% of the Na⁺ during REOX following either OGD or HAIR treatment. In the absence of Na⁺-K⁺-ATPase, the increase in [Na⁺]_i doubled during REOX. This further suggests that NKCC1 and NHE1 mediate excessive Na⁺ uptake in the astrocyte after hypoxia/ischemia and that Na⁺-K⁺-ATPase is unable to maintain Na⁺ homeostasis in astrocytes. However, additional work is needed to determine the relative kinetics of Na⁺ uptake compared with Na⁺ extrusion.

Intracellular Na⁺ overload triggers reversal of NCX in astrocytes. Following HAIR/REOX, the rise in astrocyte [Na⁺]_i was accompanied by increased entry of Ca²⁺. A significant rise in [Ca²⁺]_i was triggered in astrocytes following HAIR, which was insensitive to either inhibition of NKCC1 or NCX. In contrast, REOX induced a secondary increase in intracellular Ca²⁺ following either OGD or HAIR. We hypothesized that the REOX-induced secondary increase in intracellular Ca²⁺ results from Ca²⁺ entry through the reverse mode operation of NCX.

Approximately 70% of the REOX-triggered Ca²⁺ rise following 15 min HAIR was abolished with 10 µM KB-R7943. Interestingly, NKCC1^–/– astrocytes exhibited a similar low level of REOX-triggered Ca²⁺ rise. The data suggest that Ca²⁺ entry during REOX is largely via NKCC1-mediated reversal of NCX in astrocytes. This finding is consistent with our thermodynamic analysis that NCX would operate in the reverse mode and mediate Ca²⁺ influx when [Na⁺]_i in astrocytes is increased >25 mM (21). Maximum values of [Na⁺]_i reached 70–80 mM during REOX following either OGD or HAIR, a condition that favors the reverse operation of NCX. This view is supported by several recent reports. For example, inhibition of NCX with 100 nM KB-R7943 significantly attenuated the rise in intracellular Ca²⁺ in response to severe mechanical strain injuries in rat cortical astrocytes (15). The strain injury led to a rapid rise in [Na⁺]_i in astrocytes that was sustained for 50 min (15). Moreover, transient elevation of [Ca²⁺]_i following 25–30 min HAIR was blocked by either the NCX inhibitor KB-R7943 or the NHE1 inhibitor HOE-694 in rat astrocytes (5). Although [Na⁺]_i following HAIR was not measured in the latter study, the effects of these inhibitors implies that REOX evokes Na⁺ entry via NHE1 activation, which subsequently triggers the reversal of NCX. The elevation of [Na⁺]_i and the reversal of NCX have been suggested to play a role in spinal cord astrocyte ischemic damage during the reperfusion period (36). NCX inhibitors bepridil and KB-R7943 improve functional recovery of white matter tracts after anoxic and traumatic injury (24).

Ca_m²⁺ overload, mitochondrial dysfunction, and cell death. It has been proposed that Ca_m²⁺ overload leads to mitochondrial dysfunction via many pathways, including inhibition of oxidative phosphorylation, oxygen free radical formation, or formation of the mitochondrial PTP, which subsequently releases proapoptotic molecules such as Cyt C (6). In the current study, an approximately fivefold increase in the rhod 2 signal was found in astrocytes at 1 h REOX following 2 h OGD, indicating a significant increase in Ca_m²⁺. The elevation of Ca_m²⁺ was accompanied by 58% depolarization of _m. Interestingly, REOX-induced Ca_m²⁺ overload and loss of _m was significantly attenuated by inhibition of NKCC1 activity with bumetanide or by genetic ablation.

REOX following 15 min HAIR also caused dissipation of _m in NKCC1^+/+ astrocytes. In contrast, 50% of the _m was preserved in NKCC1^–/– astrocytes. Inhibition of NKCC1 activity not only attenuated REOX-induced depolarization of _m and Ca_m²⁺ overload, it also blocked Cyt C release from mitochondria. These findings imply that NKCC1-mediated perturbation of Na⁺ and Ca²⁺ homeostasis plays a role in Ca_m²⁺ overload and dysfunction [a positive correlation between an increase in intracellular Na⁺ and loss of _m (R² = 0.995) or Ca_m²⁺ overload and loss of _m (R² = 0.911) was found during REOX]. We subsequently demonstrated that, after 20 min HAIR treatment, REOX triggered 50% cell death, which was preceded by a delayed, sustained increase in Ca²⁺ (data not shown). These findings suggest that NKCC1-mediated dysregulation of cellular Na⁺ and Ca²⁺ homeostasis can seriously impair astrocyte function and survival.

In summary, we used two hypoxia/ischemia in vitro models to study the role of Na⁺ and Ca²⁺ dysregulation in astrocyte damage. We found that NKCC1 plays an important role in intracellular Na⁺ overload. Moreover, intracellular Na⁺ overload evokes reversal of NCX and subsequent Ca²⁺ dysregulation. We revealed here that inhibition of NKCC1 activity attenuated Na⁺ and Ca²⁺ overload, dissipation of the mitochondrial _m, release of Cyt C, and astrocyte death. These findings suggest that the concerted activities of multiple ion transport proteins are important in the perturbations of Na⁺ and Ca²⁺ homeostasis and astrocyte death in response to hypoxia/ischemia.

	GRANTS

TOP ABSTRACT EXPERIMENTAL PROCEDURES RESULTS DISCUSSION GRANTS REFERENCES

 
This work was supported in part by National Institutes of Health Grants R01NS-38118 (to D. Sun) and R01HL-61974 (to G. E. Shull) and American Heart Association Grant EIA 0540154 (to D. Sun).

	FOOTNOTES

 

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.

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