Institute of Clinical Neuroscience, Göteborg University, Göteborg, Sweden
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ABSTRACT |
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Endothelin-1 (ET-1) is a potent vasoconstrictor peptide that is also known to induce a wide spectrum of biological responses in nonvascular tissue. In this study, we found that ET-1 (100 nM) inhibited the glutamate uptake in cultured astrocytes expressing the glutamate/aspartate transporter (GLAST); astrocytes did not express the glutamate transporter-1 (GLT-1). The Vmax and the Km of the glutamate uptake were reduced by 57% and 47%, respectively. Application of the ETA and ETB receptor antagonists BQ-123 and BQ-788 partly inhibited the ET-1-evoked decrease in the glutamate uptake, whereas the nonspecific ET receptor antagonist bosentan completely inhibited this decrease. Incubation of the cultures with pertussis toxin abolished the effect of ET-1 on the uptake. The ET-1-induced decrease in the glutamate uptake was independent of extracellular free Ca2+ concentration, whereas the intracellular Ca2+ antagonists thapsigargin and 3,4,5-trimethoxybenzoic acid 8-(diethylamino)octyl ester abolished the effect of ET-1 on the glutamate uptake. Incubation with the protein kinase C (PKC) antagonist staurosporine, but not with the fatty acid-binding protein bovine serum albumin, prevented the ET-1-induced decrease in the glutamate uptake. These results suggest that ET-1 impairs the high-affinity glutamate uptake in cultured astrocytes through a G protein-coupled mechanism, involving PKC and changes in intracellular Ca2+.
glutamate/aspartate transporter; glutamate transport
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INTRODUCTION |
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ENDOTHELIN (ET) is a potent vasoactive 21-amino acid peptide, which was first isolated from porcine aortic endothelial cells (70). Subsequently, three highly homogeneous isoforms of ET were identified through cloning: ET-1, ET-2, and ET-3 (34). The biochemical characterization of these compounds has attracted intensive research over the past few years, and it has been shown that, apart from being the most powerful vasoconstrictor known, ET exerts a wide spectrum of effects on various nonvascular tissues, including those of the central nervous system (CNS) (for a review, see Ref. 44). By now, the variety of described actions of ET on the CNS has led to the recognition of ET as a neuropeptide (38).
In the mammalian CNS, ET-1 and ET-3 are produced in neurons in the brain and spinal cord (19, 39), in the endothelium of cerebral microvessels (71), and in glial cells. However, ET expression in the glia appears to be limited to the early developmental stage and to gliosis or pathological conditions such as ischemia and Alzheimer's disease (43, 48, 72). ET has a stimulatory effect on DNA synthesis (57), which is implicated in reactive gliosis (25) and has been observed to initiate astrocytic growth after spinal cord injury (66). ET has also been demonstrated to act as a strong inhibitor of gap junctional communication in primary astrocyte cultures (7, 20).
Two types of mammalian ET receptors, ETA and ETB, have been characterized. ET-1, ET-2, and ET-3 activate ETB receptors with equal potency, whereas only ET-1 and ET-2 activate the ETA receptors. Glial cells in rat brain in vivo express predominantly the ETB receptor (29). Cultured astrocytes have also been shown to exhibit ET-binding sites (30, 32).
The production of ET in endothelium is increased in hypoxia and anoxia (9). During hypoxia, ET-1 has been demonstrated to be released from astrocytes (50). In addition, an increased level of ET-like immunoreactivity has been observed in glial cells in experimental models of ischemia (16, 48, 49). These pathological conditions are characterized by a decreased high-affinity uptake of the excitatory amino acid glutamate into the astrocytes (60), while the extracellular glutamate concentration is known to rise significantly during ischemia and anoxia (24). It has been established that the removal of the released glutamate from the extracellular space is dependent on the activity of neuronal and glial glutamate transporters (for recent reviews, see Refs. 14, 17, 18, and 52). Furthermore, the major role of the glial glutamate transporter-1 (GLT-1) and glutamate/aspartate transporter (GLAST) in preventing excitotoxicity has been demonstrated with the antisense oligodeoxynucleotide technique (53). It is proposed that the elevated extracellular glutamate concentration in ischemia is partly caused by the reversed operation of the glutamate transporters as a result of cell membrane depolarization after an increase in the extracellular K+ concentration ([K+]e) (62). In this respect, it is noteworthy that ET-1 has been demonstrated to evoke the efflux of glutamate in cultured astrocytes (54).
Taking these facts into consideration, we considered it relevant to investigate the possible effect of ET-1 on glutamate uptake. In the present study, we show that ET-1 decreases the high-affinity glutamate uptake in primary cultured astrocytes from rat cerebral cortex, and we address some aspects of the intracellular mechanisms of this decrease.
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MATERIALS AND METHODS |
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Astroglial cultures. Primary astrocyte cultures from 1- to 2-day-old Sprague-Dawley rats (Charles River, Uppsala, Sweden) were grown as previously described by Hansson and coworkers (27). In brief, the rats were decapitated, and the cerebral cortexes were dissected after carefully removing the meninges. The tissue was dissociated by mechanically passing it through an 80-µm nylon mesh and then placing it in 35-mm dishes (NUNC, Roskilde, Denmark). Cultures were maintained in Eagle's minimum essential medium (MEM; Life Technologies, Paisley, Scotland). This MEM was supplied with additional substances to make up the following final composition: a double concentration of amino acids (Life Technologies), quadruple concentrations of vitamins (Life Technologies), 2 mM L-glutamine (Life Technologies), 7.5 mM glucose (Sigma Chemical, St. Louis, MO), a double concentration of NaHCO3 (Merck, Darmstadt, Germany), 1% penicillin-streptomycin (Biological Industries), and 20% fetal calf serum (Harlan Sera-Lab, Sussex, UK). The medium was changed after 3 days of cultivation and subsequently three times a week. Cells were grown in 5% CO2-95% air at 37°C, pH 7.3, for 15 days, at which time they reached confluence.
Immunohistochemistry. Cells were rinsed free of medium with phosphate-buffered saline (PBS) containing 150 mM NaCl, 3.2 mM NaH2PO4, and 16.9 mM Na2HPO4. Thereafter, they were fixed using 4% buffered paraformaldehyde at pH 7.3 and 4°C for 10 min and then permeabilized using 0.05% saponin in PBS containing 1% bovine serum albumin (BSA; PBS-BSA) for 30 min. Immunostaining was performed by incubating the cells with primary antibodies diluted in PBS-BSA for 45 min at room temperature.
The astrocytes were stained with primary polyclonal rabbit antibodies against glial fibrillary acidic protein (Dako) at a 1:200 dilution. The glutamate transporter GLAST was visualized using a commercial guinea pig polyclonal antibody (Chemicon, Temecula, CA) against a peptide corresponding to an amino acid sequence (QLIAQDNEPEKVADSETKM) from the carboxy terminus of rat GLAST. The antibody dilution was 1:4,000. The guinea pig polyclonal antibody (Chemicon) against a carboxy-terminal amino acid sequence of the rat GLT-1 (AANGKSADCSVEEEPWKREK) was used in GLT-1 staining, diluted 1:5,000. The secondary antibodies used in the next step were sheep anti-rabbit rhodamine-conjugated IgG Fab'2 fragments (Boehringer Mannheim, Mannheim, Germany) and fluorescein-conjugated donkey anti-guinea pig IgG, both diluted 1:200. HOE-33258 (Aldrich, Steinheim, Germany), diluted 1:1,000, was used for staining the cell nuclei. Rinsing with PBS-BSA-saponin was performed between every incubation step. After the staining procedure, the coverslips were mounted on microscope slides using Dako fluorescent mounting medium and viewed under a Nikon Optiphot-2. The immunopictures were captured using a Hamamatsu C5810 color-intensified 3CCD camera.Western blot. Total protein from the phenol phase was extracted from cultured astrocytes or from rat brain homogenate using a modified version (6) of the technique described by Chomczynski (11). After the RNA-containing upper water phase was removed, 1:3 vol/vol 96% ethanol was added to the lower phenol phase. Precipitation of the protein pellet was performed with the addition of 1:1 vol/vol isopropanol and shaking. The samples were centrifuged at 3,150 g for 10 min. The supernatant was discarded, and the protein pellet was washed three times for 20 min and centrifuged, as before, in 0.3 M guanidinium hydrochloride, 96% ethanol, and, finally, 96% ethanol. The protein pellets were then dissolved in an urea-saturated buffer (9.0 M urea, 0.23 M sucrose, 97 mM dithiothreitol, 33 mM SDS, and 10 mM Tris, pH 8.0), and the total protein concentration was determined as described before (7).
Western blot was performed using polyacrylamide gels (with 10% separation gel, pH 8.8, and 4% stacking gel, pH 6.8, in 0.1% SDS) run for 2-3 h at 20 mA using a Protean Cell apparatus (Bio-Rad). Protein was transferred to a polyvinylidene difluoride membrane (Immobilon-P; Millipore, Bedford, MA) at 80 mA overnight. The membranes were washed and probed with a guinea pig GLT-1 antibody at 1:4,000 or a guinea pig GLAST antibody at 1:2,500 (Chemicon) in 5-10 ml of blocking solution [1% polyvinylpyrrolidone 40 in 0.05% Tween 20 in 0.5 M Tris-buffered saline (TBS), pH 7.5]. After three TBS washes, horseradish peroxidase-conjugated goat anti-guinea pig antibody (Jackson Immunoresearch Laboratories, West Grove, PA) was applied to the membranes at 1:5,000 in blocking solution for 30 min. After five washes with 0.1% Tween 20 in TBS, the membranes were analyzed with a chemiluminescence kit (BM chemiluminescence blotting substrate, Boehringer-Mannheim). Autoradiography was done using Kodak XAR-5 film (Eastman Kodak, Rochester, NY).Glutamate uptake assay. Transport assays were conducted according to Hansson and coworkers (26) with some modifications. Twenty minutes before the experiments, individual cultures were washed three times with prewarmed (37°C) HEPES-buffered Hanks' balanced salt solution (HHBSS) containing 137 mM NaCl, 5.4 mM KCl, 0.4 mM MgSO4, 0.4 mM MgCl2, 1.26 mM CaCl2, 0.64 mM KH2PO4, 3 mM NaHCO3, 5.5 mM glucose, and 20 mM HEPES, pH 7.4. In the experiments requiring Ca2+-depleted HHBSS, CaCl2 was replaced by MgCl2, and 1 mM of EGTA (Sigma) was added. To determine the uptake kinetics, 1 µCi/ml of L-[2.3-3H]glutamate (NEN, Boston, MA) was added to unlabeled L-glutamate (Sigma) to give the final concentrations of 10, 50, 100, 200, 400, 1,000, or 1,500 µM. In all other uptake experiments, the glutamate concentration of 1,000 µM was used. The cultures were incubated with the isotope on a heating plate at 37°C for 4 min. Over this time period, uptake of glutamate was linear. The uptake process was terminated by a rapid removal of the radioactive medium and three washes with ice-cold uptake buffer. Cells were harvested into 400 ml of 1 M NaOH solution. Separate aliquots were taken for protein determination according to Lowry and coworkers (42) and for measurements of radioactivity by liquid scintillation counting. The samples for measurements of radioactivity were transferred to vials containing 2 ml of aqueous scintillation mixture and counted using a Wallac 1215 RackBeta liquid scintillation counter. Uptake rates were calculated from the measured radioactivity of the cells, the cultures' protein content, and the specific activity of the medium, and were expressed in nanomoles per milligram of protein per minute.
To isolate Na+-dependent transport, the cultures were exposed to tritiated substrate in the presence or absence of Na+. In the Na+-free buffer, NaCl was replaced with choline chloride. The radioactivity that accumulated in the cultures in the absence of Na+ was treated as blank and was subtracted from that accumulated in the cultures in the presence of Na+. ET-1 (100 nM), 5-hydroxytryptamine (5-HT; 100 µM), isoproterenol (1 µM),Statistical analyses. The kinetic parameters Km and Vmax were obtained by a least-squares, nonlinear regression analysis. Statistical analyses of the data were performed by Student's t-test for simple comparisons between two groups and by one-way ANOVA followed by post hoc Tukey's analysis for multiple comparisons. Independent data are expressed as means ± SE. P < 0.05 was considered statistically significant.
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RESULTS |
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Expression of glutamate transporter subtypes in astrocyte primary
cultures.
The astrocytes exhibited immunoreactivity for the glial glutamate
transporter GLAST. Individual cells were stained to varying degrees, and occasionally, labeling was absent (Fig.
1A). By contrast, there was no
staining for GLT-1. Similarly, Western blot of homogenates from
enriched astrocyte cultures detected immunoreactive bands at ~70 kDa
when anti-GLAST antibodies were used. In analogy with the
immunohistochemical assay, no bands of expected molecular weight were
observed in Western blot when using GLT-1 antibodies. When using a rat
brain homogenate as a positive control for the GLT-1 antibody, Western
blot showed a band at ~75 kDa, which corresponds to the molecular
mass of the GLT-1 (40) (Fig. 1B). The present data are consistent with previously published reports, according to
which only GLAST is expressed in undifferentiated astrocyte cultures,
whereas coculture with neurons or treatment with dibutyryl cyclic
adenosine 3',5'-cyclic monophosphate induces expression of GLT-1 in
astrocytes (61).
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ET-1 decreases high-affinity glutamate uptake in cultured
astrocytes.
The application of 100 nM ET-1 to the astrocyte culture medium 15 s before the glutamate incubation resulted in a prominent inhibition of
the glutamate uptake (Fig.
2A). Compared with the controls, the Vmax in ET-exposed cells was
reduced by 57% (from 24.3 ± 2.3 nmol/mg of protein per minute to
10.4 ± 1.2 nmol/mg of protein per minute, P < 0.001), while the Km was decreased by 47% (from
105.3 ± 11.6 µM in controls to 55.6 ± 10.6 µM in the
treated cells, P < 0.01). The ETA receptor
antagonist BQ-123 and the ETB receptor antagonist BQ-788
(both at 10 µM) partially inhibited the effect of ET-1 (100 nM) on
the glutamate uptake. Simultaneous application of BQ-123 and BQ-788
also resulted in a partial block of the ET-1-induced glutamate uptake
inhibition. The nonspecific ET receptor antagonist bosentan (10 µM)
completely abolished the effect of ET-1 on the glutamate uptake (Fig.
2B).
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Intracellular transduction mechanisms of the ET-1 effect on
glutamate uptake.
Incubation of the cultures with pertussis toxin (PTX; 100 ng/ml,
24 h) abolished the effect of ET-1 (100 nM) on the glutamate uptake, suggesting the involvement of Gi/Go
protein in the signal transduction cascade (Fig.
3A).
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Ligand specificity of the decrease in glutamate uptake.
Because astrocytes express a large number of receptors for various
neurotransmitters (51), we tested several agonists known to induce signal transduction responses in astrocytes to determine whether they have an effect similar to that of ET-1 on the glutamate uptake. Each ligand was applied to the cells 15 s before the
application of glutamate. 5-HT (100 µM), GABA (100 µM), ATP (100 µM), carbachol (1 mM), and ACPD (100 µM) had no effect on the
glutamate uptake, whereas isoproterenol (1 µM), a -adrenoceptor
agonist, reduced the uptake by ~50%. The results are summarized in
Fig. 4.
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Cell membrane depolarizing agents BaCl2 and a high
[K+]e reduce glutamate
uptake.
After a 10-min incubation with BaCl2 (1 mM), which
depolarizes astrocytes by blockage of K+ channels, the
glutamate uptake in the treated cells was reduced by 51% compared with
the controls. A high [K+]e (35 mM), when
applied to the cells 15 s before the glutamate, inhibited the
uptake of glutamate by 63% (Fig. 5).
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DISCUSSION |
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This work aimed to investigate the effect of ET-1 on the activity of glutamate transporters in rat cortical astrocytes in primary culture. Studies of the glutamate uptake kinetics in cultured astrocytes showed that ET-1 was able to inhibit the uptake in a dose-dependent manner by decreasing the Vmax and the Km by ~57% and 47%, respectively. The inhibitory effect of ET-1 could be blocked by the PKC antagonist staurosporine, by the inhibitors of intracellular Ca2+ release thapsigargin and TMB-8, and by PTX, but was independent of an extracellular Ca2+ influx into the astrocytes. ET-1 receptor antagonists BQ-123 and BQ-788 partially blocked the inhibitory effect of ET-1 on the glutamate uptake, whereas the nonspecific ET receptor antagonist bosentan completely blocked this effect.
It is well documented that ETs cause a sustained increase in intracellular free Ca2+ concentration ([Ca2+]i) in astrocytes (6, 21, 58). This increase contains two components: an instant Ca2+ spike seconds after the application of ET (due to Ca2+ release from intracellular stores), and then a sustained intracellular Ca2+ elevation that lasts for several minutes and depends on the influx of Ca2+ from the extracellular space. To investigate whether the effect of ET-1 was dependent on Ca2+ release from the intracellular stores, we used two agents: thapsigargin and TMB-8, known to act by different mechanisms. Thapsigargin blocks endoplasmic reticulum Ca2+-ATPases, preventing them from counterbalancing the passive leak of Ca2+ from the endoplasmic reticulum to the cytosol, and thus indirectly leading to the Ca2+ store depletion (65). By contrast, TMB-8 is described as an intracellular Ca2+ antagonist that inhibits evoked increases in [Ca2+]i (23). Both agents were able to abolish the ET-1-induced reduction in the uptake of glutamate. However, in Ca2+-depleted medium, the effect of ET-1 on the glutamate uptake was comparable to that in controls, suggesting that Ca2+ influx is not important with regard to this effect. These observations suggest that the effect of ET-1 on the glutamate uptake may be dependent on Ca2+ release from the intracellular stores but not on the influx of extracellular Ca2+ into the astrocytes.
ET receptors are known to be coupled to a diversity of intracellular transduction mechanisms (for a review, see Refs. 33, 55, and 56). A number of these pathways have been reported to interact with glutamate transporters and could be involved in the observed decrease in the glutamate uptake: 1) ET-1 evokes the release of arachidonic acid (AA) from astrocytes in primary culture (64); this fatty acid is, in turn, a potent inhibitor of the glutamate uptake (3, 68); 2) ET-1 activates PKC (37), and PKC-mediated phosphorylation of GLAST has been shown to inhibit the activity of this transporter (12); and 3) ET-1 depolarizes astrocytic cell membranes (31, 32), and the glutamate uptake is sensitive to changes in membrane potential because it is an electrogenic process characterized by an inward movement of one positive charge per glutamate molecule (8).
Several observations indicated that AA was not involved in the observed glutamate uptake decrease. First, the ET-evoked release of AA has been shown to require extracellular Ca2+ (64, 69), and, in the present study, the ET-1-induced decrease in the glutamate uptake was independent of the extracellular Ca2+. Second, AA inhibits the uptake of glutamate mainly by reducing the maximum uptake rate, with only a minor effect on the affinity for external glutamate (3, 68). In our experiments, both the Vmax and the Km of the glutamate uptake were reduced by ET-1. Third, the free fatty acid-binding protein BSA did not abolish the inhibitory effect of ET-1 on the activity of the glutamate carriers. Together, these observations suggest that the ET-1-induced release of AA was probably not essential for the reduced activity of the glutamate transporters.
Inhibition of the activity of the glutamate transporter GLAST by PKC has been demonstrated by several groups. Phosphorylation of GLAST by PKC has not been shown; however, the functional effects of PKC activation on this transporter are consistent among existing reports (12, 22; for a review, see Ref. 2). It has been suggested that the reduction in GLAST activity in astrocytes resulting from PKC activation might be due to rapid intracellular sequestration of GLAST transporters (13). PKC activation could, at least in part, be involved in the ET-1-induced inhibition. This hypothesis is supported by the fact that the preincubation of the astrocytes with the protein kinase inhibitor staurosporine abolished the inhibitory effect of ET-1 on the glutamate uptake. The role of active PKC involved in the studied mechanism is further supported by the absence of an ET-1-induced inhibition when intracellular Ca2+ is blocked by TMB-8 or thapsigargin. Several isoforms of PKC require Ca2+ for a full activation of the enzyme, and the lack of Ca2+ release from intracellular stores may impair the PKC function. However, the PKC-mediated decrease in glutamate carrier efficacy is characterized by a decrease in the Vmax, but not in the Km (12), as opposed to the present findings of an almost constant Vmax:Km ratio.
As shown here and by others (41, 63), the depolarizing agents BaCl2 and extracellular K+ at high concentrations inhibit the glutamate uptake, and this inhibition is characterized by a similar pattern of parallel decreases in both the Vmax and the Km (41). ET-1 has been reported to evoke cell membrane depolarization in astrocytes (31, 32). If the membrane depolarization is strong enough, glutamate transporters may start operating in a reversed manner, releasing glutamate from the astrocytes instead of taking it up (62). The amplitude of ET-1-induced astrocyte membrane depolarization has been shown to vary between individual cells, reaching as high as 23 mV (32). A 20-mV depolarization induced by BaCl2 has been demonstrated to be sufficient to reverse the operation of the glutamate transporters, leading to excitatory amino acid release (41). Moreover, recently, Sasaki and coworkers (54) reported that ET-1 evoked the efflux of glutamate from rat cortical astrocytes and concluded that this was probably due to a reversal of glutamate carriers. Together, these findings support the hypothesis that an impairment of the glutamate carriers, or even carrier reversal, resulting from the ET-1-induced membrane depolarization, may contribute to the observed decrease in the glutamate uptake.
The fact that the effect of ET-1 on the glutamate uptake could be blocked by incubation with staurosporine is compatible with the hypothesis above, because it has been shown that activation of PKC leads to a significant depolarization of the astrocyte membrane (1). A PKC-dependent depolarization of astrocyte membranes in response to membrane receptor stimulation has been reported by Köller and coworkers (36). However, in the case of ET-1, the exact mechanism of the astrocyte membrane depolarization is still not clarified and will require further examination.
ET receptor antagonists BQ-123 (ETA receptor) and BQ-788 (ETB receptor) only partially blocked the inhibitory effect of ET-1 on the glutamate uptake. This was true even when a combination of BQ-123 and BQ-788 was applied to the cells. However, the nonspecific ET receptor antagonist bosentan completely abolished the ET-1-induced inhibition of the glutamate uptake. A possible explanation of this fact could be that the inhibitory effect is mediated, at least partially, by an atypical, yet not fully characterized, ET receptor. Expression in rat astrocytes of such an atypical ET receptor has been reported by Jensen and coworkers (35).
To further address the question of specificity of the ET-1-induced decrease of the glutamate uptake, we tested the effect of several other agents known to activate PKC and/or to increase the intracellular Ca2+ in astrocytes. Carbachol, a muscarinic receptor agonist, is known to trigger Ca2+ waves in cultured rat astrocytes dependent on phospholipase C activity and inositol phosphate production (67). A metabotropic glutamate receptor agonist ACPD has been shown to induce Ca2+ oscillations in cultured astrocytes in a PKC-dependent manner (45). Adenosine triphosphate induces the increase of intracellular Ca2+ levels in cultured astrocytes and activates PKC (10), and 5-HT and GABA evoke cytosolic Ca2+ mobilization in cultured astrocytes (46, 47). Carbachol, ACPD, ATP, and GABA had no effect on the function of the glutamate transporters, whereas 5-HT decreased the glutamate uptake into astrocytes.
Elevated extracellular levels of glutamate are currently considered to be the main cause of neuronal death in ischemia and have been implicated as a mechanism of neuronal death in several other acute and chronic neuropathological conditions (5, 15). On the other hand, increased levels of ET-1 have been demonstrated in the rat brain after transient ischemia (4) and in the cerebrospinal fluid of patients suffering from stroke or subarachnoidal hemorrhage (28, 59). The present work shows that ET-1 has an inhibitory effect on glutamate uptake into cultured astrocytes, and Sasaki and coworkers (54) found that ET-1 was able to induce glutamate release from cultured astrocytes. Furthermore, ET-1 is a potent blocker of gap junction communication in the astrocyte syncytium (7), which might have implications for the buffering of K+ and other low-molecular-weight neurotoxic substances from the damage site. These findings indicate that ET-1 may contribute to the neuronal damage in cerebrovascular disorders, not only by potent and long-lasting vasoconstrictor action (49) but also by affecting the astroglial cell functions. The inhibitory effect of ET-1 on the astroglial glutamate uptake may exacerbate glutamate-induced excitotoxicity. However, one should bear in mind that only one subtype of glutamate transporter, namely GLAST, was expressed in our cultures, whereas both GLAST and GLT-1 are expressed in astrocytes in vivo. Further experiments determining the effect of ET-1 on GLT-1 are required.
In conclusion, the present study demonstrates that the ET-1 induces a decrease in glutamate uptake in cultured astrocytes. The mechanism of this effect may involve PKC activation, an increase in [Ca2+]i, and cell membrane depolarization; moreover, interaction between several intracellular pathways may be necessary.
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ACKNOWLEDGEMENTS |
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The skillful technical assistance of Ulrika Björklund and Barbro Eriksson is greatly appreciated. The endothelin receptor antagonist bosentan was generously supplied by Dr. Martine Clozel (Hoffmann-LaRoche, Basel, Switzerland). We are grateful to Professor Torsten Olsson and Professor Sture Holm of Chalmers University of Technology (Göteborg, Sweden) for data analysis and statistical advice and to Professor Lou DeFelice (Vanderbilt University, Nashville, TN) for helpful discussion.
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FOOTNOTES |
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This work was supported by Swedish Medical Research Council Grant 33X-06812, Swedish Council for Working Life and Social Research, Edith Jacobsson's Foundation, Rune and Ulla Almlöv's Foundation for Neurological and Rheumatological Research, and John and Brit Wennerström's Foundation for Neurological Research.
Address for reprint requests and other correspondence: J. Leonova, Institute of Clinical Neuroscience, Göteborg Univ., Box 420, S-405 30 Göteborg, Sweden (E-mail: elisabeth.hansson{at}anatcell.gu.se).
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.
Received 1 December 2000; accepted in final form 2 July 2001.
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