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NEUROPHARMACOLOGY
Istituto di Ricerche Farmacologiche "Mario Negri," Milano, Italy (S.C., E.F., T.M., M.G.); Department of Medicinal Chemistry, Faculty of Pharmaceutical Sciences, University of Copenhagen, Copenhagen, Denmark (A.A.J.); Dipartimento di Farmacologia Preclinica e Clinica, Università di Firenze, Firenze, Italy (E.L., D.E.P.-G.); and Istituto di Chimica Farmaceutica e Tossicologica "Pietro Pratesi," Università di Milano, Milan, Italy (P.C., A.P., M.D.A., C.D.M.)
Received for publication
December 13, 2007
Accepted
April 30, 2008.
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Abstract |
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 Top Abstract Materials and MethodsResultsDiscussionReferences |
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). This reuptake is essential for maintaining of the balance of Glu receptor signaling in the central nervous system. In contrast to their neuroprotective role under normal conditions, EAATs also contribute to the severe neuronal damage caused by high levels of extracellular Glu (excitotoxicity). Especially under conditions where energy levels fall and the transmembrane gradients of Na+ collapse, EAATs mediate Glu release into the synaptic cleft, through the so-called "reverse transport" (McMahon and Nicholls, 1991; Levi and Raiteri, 1993). In fact, most of the Glu release in early ischemic states has been attributed to this reverse transport (Rossi et al., 2000).
To date, five EAAT subtypes have been identified. The human transporter subtypes EAAT1, EAAT2, and EAAT3 are expressed throughout the central nervous system (Arriza et al., 1994) and correspond to the rat homolog GLAST (Storck et al., 1992), GLT1 (Pines et al., 1992), and EAAC1 (Kanai et al., 1995). The EAAT4 and EAAT5 subtypes are predominantly expressed in cerebellar Purkinje cells (Fairman et al., 1995; Nagao et al., 1997) and in the retina (Arriza et al., 1997), respectively. Except in the cerebellum, where EAAT1/GLAST is the predominant EAAT subtype (Lehre and Danbolt, 1998), the GLT1/EAAT2 subtype quantitatively dominates in the forebrain because anti-GLT1 antibodies immunoprecipitate more than 90% of the reconstitutable glutamate transport (Haugeto et al., 1996). In agreement with this dominant role, a GLT1-KO mouse had lethal spontaneous seizures and showed increased susceptibility to acute forebrain injury (Watase et al., 1998), whereas mice knockouts of the other EAAT subtypes had less dramatic phenotypes (Peghini et al., 1997; Watase et al., 1998).
Very few pharmacological tools are currently available to elucidate the EAAT's function and to probe perspectives in the transporters as therapeutic targets. Our studies aimed at identifying new EAAT inhibitors gave very interesting results with (±)-3-hydroxy-4,5,6,6a-tetrahydro-3aH-pyrrolo-[3,4-d]isoxazole-4-carboxylic acid (HIP-A) and 3-hydroxy-4,5,6,6a-tetrahydro-3aH-pyrrolo[3,4-d]isoxazole-6-carboxylic acid (HIP-B), two conformationally constrained analogs of D-aspartate (Asp) and Glu, respectively, with low or no affinity for ionotropic and metabotropic glutamate receptors (Conti et al., 1999a,b). More interestingly, in rat brain cortex synaptosomal preparations, (±)-HIP-A and (±)-HIP-B inhibit Glu-induced [3H]Asp release, a model of EAAT-mediated reverse transport, with inhibitory potencies 10 times those for inhibition of [3H]Glu uptake (Funicello et al., 2004). The remarkable ability of these compounds to differentiate Glu uptake and Glu-induced release mediated by the EAATs makes them very interesting tools for clarifying the mechanisms of these processes. Moreover, the possibility of inhibiting the EAAT-mediated efflux of Glu more potently than EAAT-mediated Glu uptake might offer therapeutic potential, e.g., to target ischemia-induced Glu release (neurotoxic mechanism) with marginal effect on Glu uptake (neuroprotective mechanism). To explore this opportunity, the present work compares the effects of (–)-HIP-A, identified here as the most active isomer, and DL-threo-β-benzyloxyaspartic acid (TBOA) in mixed mouse cortical cultures and organotypic rat hippocampal slices exposed to oxygen-glucose deprivation (OGD), two established in vitro models of cerebral ischemia (Pellegrini-Giampietro et al., 1999). We also did in vitro studies in synaptosomal preparations and hEAAT1–3-expressing HEK293 cells to gain further information on the molecular mechanisms of action of (–)-HIP-A.
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Materials and Methods |
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TopAbstractMaterials and MethodsResultsDiscussionReferences |
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). L-Glutamic acid was obtained from Sigma-Aldrich (St. Louis, MO), and TBOA was obtained from Tocris Cookson Inc. (Ellisville, MO). [3H]Glu and [3H]Asp were purchased from GE Healthcare (Chalfont St. Giles, UK). The construction of the hEAAT1–3-HEK293 cell lines has been described elsewhere (Jensen and Bräuner-Osborne, 2004).
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Preparation of the Synaptosomal Fraction. Adult male CRL: CD(SD)BR rats (Charles River Italica, Calco, Italy) were used. Procedures involving animals and their care were conducted in conformity with the institutional guidelines that are in compliance with national (D.L. n.116 G.U., suppl. 40, 1992 Feb 18) and international laws and policies (Guide for Care and Use of Laboratory Animals, 1996).
Rats were decapitated, and their brains were quickly removed. The cortex was dissected out and homogenized on ice in 40 volumes of ice-cold phosphate-buffered sucrose (0.32 M), pH 7.4, in a glass-Teflon homogenizer. To eliminate the nuclear fraction and cellular debris, the homogenate was centrifuged at 1000g for 5 min at 4°C. The supernatant was then centrifuged at 12,000g for 20 min at 4°C, and the resulting P2 synaptosomal pellet was resuspended in a assay buffer with the following composition: 10 mM Tris acetate, 128 mM NaCl, 5 mM KCl, 1.5 mM NaH2PO4, 1 mM MgSO4, 1 mM CaCl2, and 10 mM D-glucose 10, pH 7.3 to 7.4 (Koch et al., 1999b).
[3H]Asp Uptake in Synaptosomal Preparations. [3H]Asp uptake was measured as described previously (Funicello et al., 2004), in conditions similar to those used in hEAAT-expressing cells (see below). Thus, the P2 pellets were diluted to a concentration of approximately 3 mg/ml initial tissue in assay buffer, and 0.5-ml aliquots were preincubated for 7 min at 35°C in a water bath with or without the compounds to be tested. Nonspecific uptake was determined in the presence of 300 µM L-glutamate. Uptake was started by adding 10 nM [3H]Asp (specific activity, 36 Ci/mmol) and was stopped 4 min later by adding 2 ml of ice-chilled assay buffer. Samples were immediately filtered through cellulose mixed ester filters (0.65-µm pore size; Millipore Corporation, Cork, MA) and washed with 2 ml of assay buffer. The radioactivity trapped on the filters was counted in 4 ml of Ultima Gold MV (PerkinElmer Life and Analytical Sciences, Waltham, MA) in a Wallac 1409 liquid scintillation counter (PerkinElmer Life and Analytical Sciences) with a counting efficiency of approximately 60%.
IC50 values were determined by fitting the concentration-response curves in the presence of different concentrations of inhibitors (in triplicate), using the "one-site competition" equation built into GraphPad Prism 4.0a (GraphPad Software Inc., San Diego, CA), which gives the IC50 with its 95% confidence interval (CI).
[3H]Glu Uptake in Synaptosomal Preparations. At variance with the classic uptake protocol used in our previous study (Funicello et al., 2004), in the present study, [3H]Glu uptake was measured with the superfusion apparatus, using exactly the same procedures as for release experiments (see below). Thus, P2 pellets were resuspended in approximately 300 volumes of prewarmed assay buffer, and 2.5-ml portions (approximately 3 mg/ml initial tissue) were distributed on cellulose mixed ester Millipore filters (0.65 µm) placed at the bottom of the superfusion chambers held thermostatically at 37°C. Superfusion was started (t = 0 min) with assay buffer at a rate of 0.5 ml/min. Ten micromolars of [3H]Glu (initial specific activity, 49 Ci/mmol; specific activity after isotopic dilution with unlabeled Glu, 0.049 Ci/mmol) was then added in the superfusion buffer from 37 to 41 min; inhibitors were present in the superfusion buffer according to different experimental protocols, detailed under Results. For saturation experiments, [3H]Glu concentrations were increased by isotopic dilution with unlabeled Glu.
The uptake reaction was terminated after 4 min by adding ice-cold assay buffer in the superfusion chambers, followed by rapid aspiration of the buffer from the bottom. Filters were then counted for radioactivity as described above.
The specific [3H]Glu uptake was measured by subtraction of nonspecific uptake, estimated in parallel chambers, containing 300 µM unlabeled Glu (Funicello et al., 2004). The effects of the inhibitors on specific [3H]Glu uptake were calculated as percentages of the uptake measured, in parallel, without inhibitors.
IC50 values were determined by fitting the concentration-response curves in the presence of different concentrations of inhibitors (in triplicate) using the one-site competition equation built into GraphPad Prism 4.0a, which gives the IC50 with its 95% CI.
The saturation curves, i.e., in the presence of different concentrations of [3H]Glu, were fitted using the "one-site binding, hyperbola" equation (GraphPad Prism 4.0a). This estimates the maximal uptake velocity (Vmax) and Km, with their S.E.M.
[3H]D-Asp Release from Superfused Synaptosomes. P2 pellets were resuspended in approximately 40 volumes of assay buffer (see above), preincubated at 37°C for 10 min, diluted with an equal amount of a buffer containing [3H]Asp (final concentration, 60 nM; specific activity, 36 Ci/mmol) and incubated for 15 min. The suspension was then diluted 1:10 in prewarmed Koch buffer, and 2.5-ml portions (approximately 3 mg/ml initial tissue) were distributed on cellulose mixed ester Millipore filters (0.65-µm pore size) placed at the bottom of superfusion chambers held thermostatically at 37°C, and layered onto filters by aspiration from the bottom under moderate vacuum. Superfusion started (t = 0 min) with assay buffer at a rate of 0.5 ml/min. After 34 min of equilibration, fractions were collected every 2 min from 34 to 46 min. At the end, the filters were put into scintillation vials and counted for radioactivity, as the fractions, in 4 ml of Ultima Gold MV (PerkinElmer Life and Analytical Sciences).
[3H]Asp release was induced by either Glu (added in the superfusion buffer from t = 37 min at a concentration of 10 µM, if not otherwise indicated) or by inhibition of energy-dependent processes, i.e., another condition that favors reversal of transporter function. The latter was obtained by adding 5 µM rotenone and 2 mM 2-deoxyglucose in glucose-free medium resulting in inhibition of mitochondrial ATP synthesis, glycolysis, and glycogenolysis (McMahon and Nicholls, 1991; Santos et al., 1996; Koch et al., 1999a). Inhibitors were present in the superfusion buffer according to different experimental protocols, detailed under Results.
The fractional release rate (FRR) was calculated as 100 times the amount of radioactivity released into each 2-min fraction over total radioactivity present on the filter at the start of that fraction. The FRRs before the releasing stimulus (two fractions, 34–36 and 36–38 min), expressed as a percentage in 2 min, were considered as the basal outflow. Glu-induced release was calculated as the difference between the FRR in the presence of Glu (38–44 min) and the basal outflow.
The effects of the inhibitors on Glu-induced release were calculated as percentages of the release measured, in parallel, without inhibitors. IC50 values were determined by fitting the concentration-response curves in the presence of different concentrations of inhibitors (in triplicate), using the one-site competition equation built into GraphPad Prism 4.0a, which gives the IC50 with its 95% CI.
The saturation curves, i.e., in the presence of different concentrations of Glu, were fitted using the one-site binding, hyperbola equation (GraphPad Prism 4.0a). This estimates the maximal release rate (plateau of the releasing effect of Glu) and Km, with their S.E.M.
[3H]Asp Uptake in hEAAT1–3-Expressing Cells. The pharmacological properties of the compounds at hEAAT1-, hEAAT2-, and hEAAT3-HEK293 cell lines were determined as described previously (Jensen and Bräuner-Osborne, 2004). In brief, cells were split into poly-D-lysine-coated white 96-well plates (PerkinElmer Life and Analytical Sciences) in Dulbecco's modified Eagle's medium supplemented with penicillin (100 U/ml), streptomycin (100 µg/ml), 10% dialyzed fetal bovine serum, and 1 mg/ml G-418. Sixteen to 24 h later, the medium was aspirated, and cells were washed two times with room temperature 100 µl of Krebs buffer (140 mM NaCl, 4.7 mM KCl, 2.5 mM CaCl2, 1.2 mM MgCl2, 10 mM D-glucose, 11 mM HEPES, pH 7.4).
For the preincubation experiments depicted in Fig. 5B, the cells were incubated with 50 µl of Krebs buffer with or without 20 µM (–)-HIP-A for 30 min, after which the buffers were aspirated, and the cells were washed three times with buffer. Then, 50 µl of Krebs buffer supplemented with 3, 10, or 30 nM [3H]Asp or with 30 nM [3H]Asp and various concentrations of unlabeled Asp was added to the wells, and the plate was incubated at 37°C for 7 min.
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Fluorometric Imaging Plate Reader Membrane Potential Assay in hEAAT1–3-Expressing Cells. The pharmacological properties of (–)-HIP-A on hEAAT-HEK293 cell lines in the fluorometric imaging plate reader membrane potential (FMP) assay (Molecular Devices, Crawley, UK) were determined essentially as described previously (Jensen and Braüner-Osborne, 2004). In brief, cells were split into poly-D-lysine-coated black clear-bottom 96-well plates (BD Biosciences, San Jose, CA) in Dulbecco's modified Eagle's medium supplemented with penicillin (100 U/ml), streptomycin (100 µg/ml), 5% dialyzed fetal bovine serum, and 1 mg/ml G-418; 16–24 h later, the medium was aspirated, and the cells were washed with 100 µlof Krebs buffer. Then, 50 µl of Krebs buffer, supplemented with FMP assay dye, with or without 30 µM (–)-HIP-A, was added to each well, and the plate was incubated at 37°C for 30 min. The plate was assayed at room temperature in a NOVOstar plate reader (BMG Labtech GmbH, Offenburg, Germany) measuring emission at 560 nm in response to excitation at 530 nm before and up to 1 min after addition of 33 µl of Glu solution (dissolved in Krebs buffer). The concentration-response curves were constructed based on the maximal responses for eight different concentrations of each compound. The experiments were run in duplicate three times.
OGD in Cortical Cell Cultures. Primary cultures of mixed cortical cells containing both neuronal and glial elements were prepared as described previously by (Pellegrini-Giampietro et al., 1999). In brief, cerebral cortices were dissected from fetal mice at 14 to 15 days of gestation, minced using medium stock (MS; composed of Eagle's minimal essential medium, with Earle's salts, glutamine- and NaHCO3-free, supplemented with 38 mM NaHCO3, 22 mM glucose, 100 U/ml penicillin, and 100 µg/ml streptomycin), and incubated for 10 min at 37°C in MS with 0.25% trypsin and 0.05% DNase. Enzymatic digestion was terminated by a second incubation (10 min at 37°C) in MS supplemented with 10% heat-inactivated horse serum and 10% fetal bovine serum, after which the cells were mechanically disrupted and counted. After brief centrifugation, cells were resuspended (approximately 4 x 105 cells/ml) and plated in 15-mm multiwell vessels on a layer of confluent astrocytes using a plating medium of MS supplemented with 10% heat-inactivated horse serum, 10% fetal bovine serum, and 2 mM glutamine. Cultures were kept in an incubator at 37°C, with 100% humidity and 95% air/5% CO2 atmosphere. After 4 to 5 days in vitro (DIV), non-neuronal cell division was halted by the application of 3 µM cytosine arabinoside for 24 h. Cultures were then shifted to a maintenance medium identical to the plating medium but lacking fetal bovine serum, which was then partially replaced twice a week. Experiments were done with mature cultures (14–15 DIV).
OGD was induced as described previously (Pellegrini-Giampietro et al., 1999). In brief, the culture medium was replaced by thorough exchange with a glucose-free balanced salt solution (composition: 116 mM NaCl, 5.4 mM KCl, 0.8 mM MgSO4, 1 mM NaH2PO4, 26 mM NaHCO3, 1.8 mM CaCl2, and 10 mg/l phenol red) previously saturated with 95% N2/5% CO2 and heated to 37°C. Multiwells were then placed in a gassed incubator equipped with an oxygen controller (BioSpherix, Redfield, NY) at 37°C for 60 min.
The chamber was then sealed and placed in the incubator at 37°C for 60 min. OGD was terminated by removing the cultures from the chamber, replacing the exposure solution with oxygenated MS, and returning the multiwells to the incubator under normoxic conditions. The extent of neuronal death was assessed 24 h later. To achieve maximal neuronal injury, the cultures were exposed for 24 h to 1 mM glutamate in MS at 37°C, with 100% humidity and 95% air/5% CO2 atmosphere.
Cell damage was evaluated by measuring the amount of lactate dehydrogenase (LDH) released from injured cells into the extracellular fluid 24 h after exposure to OGD or glutamate (Pellegrini-Giampietro et al., 1999). Background LDH release was determined in control cultures not exposed to OGD and was subtracted from all experimental values. The resulting value correlated linearly with the degree of cell loss estimated by observing the cultures under phase-contrast microscopy or under bright-field optics after 5 min of incubation with 0.4% trypan blue, which stains debris and nonviable cells.
Oxygen-Glucose Deprivation in Organotypic Hippocampal Cultures. Organotypic hippocampal slice cultures were as described previously (Pellegrini-Giampietro et al., 1999). In brief, the hippocampi were removed from the brains of 7- to 8-day-old Wistar rats, and transverse slices (420 µm) were prepared using a McIlwain tissue chopper in a sterile environment. Isolated slices were first placed in ice-cold Hanks' balanced salt solution, supplemented with 5 mg/ml glucose and 1.5% amphotericin B (Fungizone; Invitrogen, Carlsbad, CA), then transferred to humidified semiporous membranes (30-mm Millicell-CM 0.4-mm tissue culture plate inserts; Millipore Corporation; four per membrane). These were placed in six-well tissue culture plates containing 1.2 ml of culture medium containing 50% Eagle's minimal essential medium, 25% heat-inactivated horse serum, 25% Hanks' balanced salt solution, 5 mg/ml glucose, 1 mM glutamine, and 1.5% amphotericin B. Slices were maintained at 37°C, with 100% humidity and 95% air/5% CO2 atmosphere, and the medium was changed every 3 days. Experiments were carried out after 14 DIV. OGD was induced as described previously (Pellegrini-Giampietro et al., 1999). In brief, the slices were exposed to a serum-free medium saturated with 95% N2/5% CO2 at 37°C in a gassed incubator equipped with an oxygen controller (BioSpherix). After 30 min, the cultures were transferred to oxygenated serum-free medium containing 5 mg/ml glucose and returned to the incubator under normoxic conditions. Neuronal injury was evaluated 24 h later. Maximal damage was achieved in this system by exposing the slices to 10 mM glutamate for 24 h.
Cell injury was assessed using the fluorescent dye propidium iodide (PI), a highly polar compound that is normally excluded from cells with an intact membrane. When the membrane is damaged, PI can enter the cells and upon binding to exposed DNA becomes highly fluorescent. PI (5 µg/ml) was added to the medium at the end of the 24-h post-OGD recovery period. Thirty minutes later, fluorescence was viewed using an inverted fluorescence microscope (Olympus IX-50; Solent Scientific, Segensworth, UK) equipped with a xenonarc lamp, a low-power objective (4x), and a rhodamine filter. Images were digitized using a video image obtained with a charge-coupled device camera (Diagnostic Instruments, Inc., Sterling Heights, MI) controlled by software (InCyt Im1TM; Intracellular Imaging Inc., Cincinnati, OH) and subsequently analyzed using the Image-Pro Plus morphometric analysis software (Media Cybernetics, Inc., Bethesda, MD). To quantify cell death, the CA1 hippocampal subfield was identified and encompassed in a frame using the drawing function in the image software (ImageJ; National Institutes of Health, Bethesda, MD), and the optical density of PI fluorescence was recorded. There was a linear correlation between CA1 PI fluorescence and the number of injured CA1 pyramidal cells, as indicated by morphological criteria.
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Results |
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TopAbstractMaterials and MethodsResultsDiscussionReferences |
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Synaptosomal Preparations. We compared the effects of (–)-HIP-A as an inhibitor of Glu uptake and Glu-induced reverse transport (release) in rat brain cortex synaptosomal preparations (P2). Uptake and release assays were done in identical experimental settings in the same superfusion apparatus, for a reliable comparison of the results (see Materials and Methods and legend to Table 2). When added 7 min before the substrate (classic protocol), the standard EAAT blocker TBOA affected the uptake of [3H]Glu (10 µM) and the Glu (10 µM)-induced [3H]Asp release with similar inhibitory potencies, giving IC50 of 6.4 and 7.3 µM, respectively. In contrast, the IC50 of (–)-HIP-A for inhibition of [3H]Glu uptake (2.2 µM) was 3 times its IC50 for inhibiting Glu-induced release (0.7 µM) (Table 2). The sizable difference between the IC50 in the two assays confirms the finding previously reported for racemic HIP-A (Funicello et al., 2004).
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Also consistent with these previous data (Funicello et al., 2004), we found that the inhibitory potency of (–)-HIP-A on Glu-induced release was highly dependent on the duration of preincubation because (–)-HIP-A had a higher IC50 (8.6 µM) when added concomitantly with Glu and significantly lower (0.7 and 0.4 µM) after 7- and 27-min preincubation. The IC50 of TBOA on Glu-induced release was not affected by the preincubation time, being 4.6, 4.1, and 4.6 µM when it was added concomitantly to Glu or after 7- and 27-min preincubation. The length of preincubation had much less effect on the inhibitory potency of (–)-HIP-A on Glu uptake (Table 2). Thus, with 27-min preincubation, (–)-HIP-A was 5 times more potent in inhibiting Glu-induced release than Glu uptake (Table 2).
It is interesting to note that although the 27-min preincubation with 1 µM (–)-HIP-A resulted in a marked impairment of Glu-induced [3H]Asp release (Fig. 2A), it did not significantly affect the [3H]Asp release induced by adding rotenone and 2-deoxy-glucose in glucose-free medium (Fig. 2B). In these conditions, in which the energy-dependent processes are inhibited and the ionic gradients are altered (McMahon and Nicholls, 1991), [3H]Asp is released through a reverse transport mediated by EAATs, as indicated by the almost complete inhibition observed in the presence of TBOA (Fig. 2B) (Funicello et al., 2004).
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We had shown previously that the inhibitory effects of (±)-HIP-A on synaptosomal [3H]Glu uptake cannot be overcome by increasing the concentration of the labeled substrate, suggesting a noncompetitive interaction (Funicello et al., 2004). In the present study, we found that (–)-HIP-A also had an insurmountable (noncompetitive-like) inhibitory effect on both [3H]Glu uptake and Glu-induced [3H]Asp release (Fig. 3, A and B), with a significant decrease in the maximal effect (see legend to Fig. 2). TBOA, a competitive inhibitor of EAAT-mediated uptake (Shimamoto et al., 1998; Funicello et al., 2004), acted as a competitive inhibitor of glutamate-induced [3H]Asp release as well (Fig. 3C). The data shown in Fig. 3 were obtained by adding the inhibitor 7 min before the substrate, but when (–)-HIP-A was added concomitantly with [3H]Glu, it acted mainly on the Km of Glu uptake, which increased from 11 to 52 µM, but not on Vmax, indicating a competitive interaction (Fig. 3D).
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) was very similar in cells expressing human EAAT1, EAAT2, and EAAT3 (7.4, 3.3, and 11 µM, respectively). These values were obtained by adding (–)-HIP-A 30 min before application of Glu. The compound did not elicit a fluorescence response per se, nor did the preincubation with (–)-HIP-A elevate the fluorescence baseline. Under this condition, the inhibition observed in hEAAT2 cells was noncompetitive (Fig. 5A).
Likewise, the preincubation of hEAAT2 cells for 30 min with 20 µM (–)-HIP-A resulted in a 50% decrease in the Vmax of [3H]Asp, indicating a noncompetitive inhibition, although Km was also increased (Fig. 5B). On the contrary, when (–)-HIP-A was added concomitantly with [3H]Asp, the inhibition is competitive with a significant 6-fold increase in the Km of the labeled substrate (Fig. 5C).
Mixed Cortical Cells and Organotypic Hippocampal Slices. Figure 6 shows the effects of TBOA and (–)-HIP-A on mixed mouse cortical cultures and organotypic rat hippocampal slices. Under basal conditions, 24-h exposure to TBOA caused a dose-dependent toxicity in cortical cultures, whereas in hippocampal slices, only the highest concentration of TBOA (100 µM) had a significant toxic effect. The lower inhibitory potency of TBOA in hippocampal slices may be because lower concentrations are reached at the active sites because the tissue is more compact, hence, less accessible than cortical cultures. In contrast to the excitotoxicity induced by TBOA, exposure to (–)-HIP-A, up to 100 µM, had no toxic effects in either of the two models. As described elsewhere (Pellegrini-Giampietro et al., 1999), exposure of mixed cortical cells to 60 min of OGD induced approximately 65% of the maximal neuronal damage caused by exposing the cultures to 1 mM glutamate for 24 h, whereas exposure of hippocampal slices to 30 min of OGD induced a selective CA1 injury that was approximately 60% of the maximal CA1 toxicity after 24-h exposure to 10 mM glutamate. OGD-induced toxicity was dose-dependently potentiated in both models when 30 to 100 µM TBOA was added to the medium during OGD and in the subsequent 24-h recovery period. In contrast, a significantly less OGD-induced neuronal damage was seen in both models in the presence of 10 and 30 µM (–)-HIP-A; this neuroprotective effect disappears at 100 µM.
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Discussion |
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TopAbstractMaterials and MethodsResultsDiscussionReferences |
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) provided explanations for the stereoselectivity.
Neither (–)-HIP-A nor (+)-HIP-B discriminate between hEAAT1–3 subtypes. The rank order of potency of (–)-HIP-A, (+)-HIP-B, and TBOA for inhibiting EAAT-mediated uptake in the synaptosomal fraction is different from the order for uptake mediated by hEAAT1–3 [(+)-HIP-B 
(–)-HIP-A > TBOA and TBOA > (–)-HIP-A > (+)-HIP-B, respectively]. These data question the hypothesis that synaptosomal EAATs correspond only to EAAT2/GLT1 transporters (Dunlop et al., 1999, 2005; Tan et al., 1999) and identify our novel compounds as interesting pharmacological tools for further investigating the nature of presynaptic transporters (Danbolt, 2001).
Because (–)-HIP-A was identified as the most potent EAAT inhibitor among the HIP-A and HIP-B isomers, this compound was selected for further studies. Our experiments brought to light marked differences between (–)-HIP-A and TBOA. In particular, 1) TBOA inhibits EAATs in a competitive and reversible manner (Shimamoto et al., 1998; present data), whereas (–)-HIP-A induces long-lasting inhibition, which, under certain experimental conditions, is apparently noncompetitive. 2) The inhibitory effect of (–)-HIP-A, but not TBOA, on Glu-induced reverse transport was potentiated (up to 20-fold) by preincubation at 37°C. Interestingly, preincubation had much less effect on the potency of (–)-HIP-A as Glu uptake inhibitor. 3) (–)-HIP-A, but not TBOA, has the fascinating property of inhibiting Glu-induced reverse transport at concentrations significantly lower than those required to inhibit Glu uptake, especially after preincubation. 4) TBOA completely inhibited the reverse transport induced by extracellular glutamate and rotenone/2-deoxy-glucose (mimicking "ischemic-like" conditions), whereas (–)-HIP-A inhibited only the former.
The binding mode of TBOA was recently clarified by a crystallographic study on the complex of EAAT from Pyrococcus horikoshii (GltPh), cocrystallized with TBOA itself (Boudker et al., 2007). The aspartate skeleton of TBOA fitted the same binding site as the endogenous substrate, in line with its competitive inhibitory effect, whereas the bulky benzyl group stabilized the apo state of the transporter, thus preventing the binding of sodium and halting further conformational changes along the transport cycle. We recently docked (–)-HIP-A in the apo-state model of Gltph, showing that our compound can fit the binding pocket of TBOA and interact with all but one of the amino acid residues involved in TBOA molecular recognition (Pinto et al., 2008). Our observations that: 1) TBOA prevents the lasting effect of (–)-HIP-A, and 2) inhibition is competitive when (–)-HIP-A is applied concomitantly with the labeled substrate, are consistent with the hypothesis that (–)-HIP-A, TBOA and the endogenous substrate may share the same binding site, although final proof will only be provided by X-ray analysis of the transporter cocrystallized with (–)-HIP-A.
If (–)-HIP-A does interact with the substrate site, it might be a substrate itself. (–)-HIP-A has some, although small, releasing effect at the concentrations inhibiting Glu-induced release, but even the highest concentrations had less releasing effect than Glu. If (–)-HIP-A is a substrate, it should be a "partial" one, such as 1,2,3,4-tetrahydro-9-aminoacridine and PDC (Dunlop, 2001). However, in the FMP assay, which is the most direct measurement of whether a compound is transported with concomitant transport of ions, 1,2,3,4-tetrahydro-9-aminoacridine and PDC had depolarizing effects (Jensen and Bräuner-Osborne, 2004), whereas (–)-HIP-A had no effects.
Our data seem to indicate that (–)-HIP-A cannot "freeze" the transporter in an inactive state, as TBOA does (rotenone/deoxyglucose experiments), but specifically interferes, in a complex manner, with Glu uptake and Glu-induced release, with a remarked preference for the latter. Therefore, lasting impairment induced by (–)-HIP-A might be due to persistent modifications of the transporter (e.g., phosphorylation, internalization, or other regulatory mechanisms). This would be consistent with the noncompetitive-like inhibition seen in the saturation experiments and might also explain why the duration of preincubation with (–)-HIP-A has a significantly greater effect on Glu-induced release than on Glu uptake. In monoamine transporters, the substrate-induced reverse transport, but not the substrate uptake, is a complex mechanism involving intracellular events, such as Ca2+-dependent activation of kinases (Kantor et al., 2001; Gnegy, 2003; Seidel et al., 2005) and transporter phosphorylation (Khoshbouei et al., 2004).
In the present study, we also evaluated the effects of (–)-HIP-A and TBOA in organotypic hippocampal slices and mixed brain cortical cultures, under basal and ischemic conditions. Significant neuronal damage was observed after 24-h exposure to TBOA, at concentrations compatible with selective inhibition of EAATs (10–100 and 100 µM in cortical cells and hippocampal slices, respectively). Bonde et al. (2003) had already shown that the cell death induced by TBOA in organotypic hippocampal slices was prevented by glutamate receptor antagonists. Microdialysis studies proved that administration of TBOA into the hippocampus raises the extracellular levels of glutamate and induces neuronal damage and a convulsive state (Montiel et al., 2005). Thus, the toxic effects of TBOA in our experimental models confirm that a reuptake system is normally active to keep Glu concentrations below a toxic level (Mitani and Tanaka, 2003) and that TBOA inhibits it. However, under the same conditions 10 to 100 µM (–)-HIP-A had no toxic effects, indicating that it does not inhibit Glu reuptake, at least not enough to reach neurotoxic levels of extracellular Glu. Thus, (–)-HIP-A appeared to be less potent than TBOA as an inhibitor of reuptake under basal conditions. This finding is at least partially consistent with in vitro data showing that (–)-HIP-A is approximately 4 times less potent than TBOA as an uptake inhibitor in hEAAT1/2-expressing cells (Table 1). However, (–)-HIP-A showed a similar or even greater potency than TBOA as uptake inhibitor in crude (or enriched; data not shown) rat brain synaptosomal preparations. Although species differences cannot be excluded, it could be speculated that, under basal conditions, glutamate reuptake in mixed cortical cultures and in hippocampal slices is mainly mediated by glial GLT1 (EAAT2), as previously suggested (for review, see Danbolt, 2001) and not by the neuronal (synaptosomal) transporters, more sensitive to (–)-HIP-A than to TBOA.
Under ischemic conditions, there was a marked neuronal damage in both experimental models, and this excitotoxicity was antagonized by Glu receptor antagonists (Pellegrini-Giampietro et al., 1999), suggesting an OGD-induced Glu release. EAAT-mediated reverse transport is probably responsible for this effect because TBOA reduced the ischemia-induced Glu release in rat cortical superfusates (Phillis et al., 2000), and a subtoxic concentration (10 µM) of TBOA had a protective effect in organotypic hippocampal slices exposed to OGD (Bonde et al., 2003). However, in our assays, 10 µM TBOA failed to protect cortical cultures and hippocampal slices against OGD-induced neurodegeneration. Methodological differences in the study by Bonde et al. (2003), such as the use of serum-free medium during slice maturation and the presence of PI from the very beginning of the experiment, might explain the discrepancy. Previous data actually support the possibility that EAATs can still take up Glu even under ischemic conditions (Rao et al., 2001; Mitani and Tanaka, 2003; Selkirk et al., 2005). It has also been reported that the function of GLT1 may change from neuroprotective (uptake mode) to neurodegenerative (release mode) depending on ischemic conditions (Mitani and Tanaka, 2003).
However, unlike TBOA, 10 to 30 µM (–)-HIP-A significantly reduced the OGD-induced neurotoxicity in both cortical cultures and hippocampal slices. This neuroprotective effect cannot be ascribed to a direct antagonism at Glu receptors (Conti et al., 1999b), nor can it be due to inhibition of GABA reuptake (C. Thomsen, personal communication). Thus, we propose that it is the consequence of lower levels of extracellular Glu due to the inhibition of EAAT-mediated Glu release. However, the OGD-induced neurotoxicity was not affected by 100 µM (–)-HIP-A and was larger than with the lower concentrations. This suggests that the higher (–)-HIP-A concentration raises extracellular Glu levels by inhibiting EAAT-mediated Glu reuptake. This dual effect of (–)-HIP-A further supports the notion that, under ischemic conditions, EAATs mediate two concomitant and opposite processes: release through reverse transport and uptake. (–)-HIP-A distinguishes between the two because in the concentration window, 10 to 30 µM, the compound might have a greater effect on reverse transport, with a net neuroprotective effect. The two processes might be mediated by different EAATs, possibly located on different cell populations, and (–)-HIP-A might serve as a very promising tool to further investigate this.
Our studies in synaptosomal preparations (Funicello et al., 2004; present data) show that low concentrations of (±)-HIP-A and (–)-HIP-A efficiently and preferentially inhibit the reverse transport induced by Glu but not that induced by ischemic-like conditions. Although this is apparently at variance with the results described here in cells, it must be considered that the EAAT-mediated, OGD-induced release might be partly due to the collapse of ion gradients (in the early stage) and partly to the consequent rise of extracellular Glu (Glu-induced Glu release). Thus, our data might suggest that the latter mechanism is a relevant consequence of the ischemia-induced release, with a major, and potentially druggable, role in the subsequent neurodegeneration.
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Acknowledgements |
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Footnotes |
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Article, publication date, and citation information can be found at http://jpet.aspetjournals.org.
ABBREVIATIONS: EAAT, excitatory amino acid transporter; HIP-A, 3-hydroxy-4,5,6,6a-tetrahydro-3aH-pyrrolo[3,4-d]isoxazole-4-carboxylic acid; HIP-B, 3-hydroxy-4,5,6,6a-tetrahydro-3aH-pyrrolo[3,4-d]isoxazole-6-carboxylic acid; TBOA, DL-threo-β-benzyloxyaspartic acid; OGD, oxygen-glucose deprivation; CI, confidence interval; FRR, fractional release rate; FMP, fluorometric imaging plate reader membrane potential; MS, medium stock; DIV, day(s) in vitro; LDH, lactate dehydrogenase; PI, propidium iodide; ER, eudismic ratio.
Address correspondence to: Dr. Marco Gobbi, Istituto di Ricerche Farmacologiche "Mario Negri," Via La Masa 19, 20156 Milan, Italy. E-mail: Gobbi{at}marionegri.it
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