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Keywords:

  • cystine/glutamate antiporter;
  • excitatory amino acid transporter;
  • glutathione;
  • neuroprotection;
  • oxidative glutamate toxicity

Summary

  1. Top of page
  2. Summary
  3. Experimental procedures
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

Oxidative glutamate toxicity in the neuronal cell line HT22 is a model for cell death by oxidative stress. In this paradigm, an excess of extracellular glutamate blocks the glutamate/cystine-antiporter system inline image, depleting the cell of cysteine, a building block of the antioxidant glutathione. Loss of glutathione leads to the accumulation of reactive oxygen species and eventually cell death. We selected cells resistant to oxidative stress, which exhibit reduced glutamate-induced glutathione depletion mediated by an increase in the antiporter subunit xCT and system inline image activity. Cystine uptake was less sensitive to inhibition by glutamate and we hypothesized that glutamate import via excitatory amino acid transporters and immediate re-export via system inline image underlies this phenomenon. Inhibition of glutamate transporters by l-trans-pyrrolidine-2,4-dicarboxylic acid (PDC) and DL-threo-β-benzyloxyaspartic acid (TBOA) exacerbated glutamate-induced cell death. PDC decreased intracellular glutamate accumulation and exacerbated glutathione depletion in the presence of glutamate. Transient overexpression of xCT and the glutamate transporter EAAT3 cooperatively protected against glutamate. We conclude that EAATs support system inline image to prevent glutathione depletion caused by high extracellular glutamate. This knowledge could be of use for the development of novel therapeutics aimed at diseases associated with depletion of glutathione like Parkinson's disease.

Abbreviations used
ARE

antioxidant-response element

DMEM

Dulbecco's modified minimal essential medium

EAAT

excitatory amino acid transporter

FCS

fetal calf serum

GSH

glutathione

GPx

GSH peroxidase

HBSS

Hank's buffered salt solution

MAP2

microtubule-associated protein 2

MCB

monochlorobimane

MTT

methylthiazoltetrazolium

PBS

phosphate-buffered saline

PDC

l-trans-pyrrolidine-2,4-dicarboxylic acid

ROS

reactive oxygen species

TBOA

DL-threo-β-benzyloxyaspartic acid

Neuronal cell death in diverse neurological disorders including ischemic stroke and neurodegenerative diseases can in part be attributed to excitotoxicity, where excessive glutamate release overstimulates ionotropic glutamate receptors resulting in massive calcium influx and cell death (Choi 1988). In addition to this rapid process, increased extracellular glutamate also leads to a more prolonged cell death by oxidative stress called oxidative glutamate toxicity (Tan et al. 2001). Here, the increased extracellular glutamate depletes cells of cystine by blocking the gradient-driven glutamate/cystine antiporter system inline image (Murphy et al. 1989; Murphy et al. 1990). This transport system consists of two subunits, the specific subunit xCT and the 4F2 heavy chain (Sato et al. 1999). Cystine is required for the synthesis of glutathione (GSH), an important antioxidant in the brain (Dringen 2000; Schulz et al. 2000). GSH depletion renders the cells incapable of removing reactive oxygen species (ROS), which are constantly produced in the mitochondria as well as during certain enzymatic reactions, and ultimately leads to cell death by oxidative stress. Oxidative stress is involved in the pathophysiology of Alzheimer's and Parkinson's disease, as well as ischemic stroke (Coyle and Puttfarcken 1993; Beal 1996; Pratico 2002) and seems to involve, at least in Parkinson's disease, a decrease in GSH levels in the substantia nigra that precedes neuronal degeneration (Schulz et al. 2000).

Oxidative glutamate toxicity has been described in neuronal cell lines (Miyamoto et al. 1989; Murphy et al. 1989; Davis and Maher 1994; Maher and Davis 1996), immature primary neurons (Murphy et al. 1990; Davis and Maher 1994; Ratan et al. 1994), oligodendroglia (Oka et al. 1993), and astrocytes (Chen et al. 2000). It was also shown recently that part of the cell death after excitotoxic stimuli can be attributed to oxidative glutamate toxicity (Schubert and Piasecki 2001). The immortalized hippocampal cell line HT22 is an excellent model to study oxidative glutamate toxicity, as it lacks ionotropic glutamate receptors (Maher and Davis 1996) and responds to oxidative glutamate toxicity with programmed cell death in a very reproducible manner (Tan et al. 2001). The sequence of events that leads to glutamate-induced cell death of HT22 cells, after depletion of intracellular GSH (Davis and Maher 1994), involves the activation of 12-lipoxygenase (Li et al. 1997a), the accumulation of intracellular ROS (Tan et al. 1998), and the activation of a cyclic GMP-dependent calcium channel close to the end of the death cascade (Li et al. 1997b).

In this study, we generated HT22 cells resistant to oxidative stress and investigated the mechanisms underlying this resistance. By looking at mRNA levels of candidate genes, which could be involved in the resistance, we identified a prominent up-regulation of the system inline image subunit xCT reflected functionally by a similar increase in cystine uptake. In addition, the inhibitory properties of extracellular glutamate on cystine uptake were diminished in resistant HT22 cells. We present data that suggest that excitatory amino acid transporters (EAAT) are also involved in the resistant phenotype and that the two transporter systems work cooperatively to protect cells from GSH depletion and subsequent cell death by oxidative stress.

Experimental procedures

  1. Top of page
  2. Summary
  3. Experimental procedures
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

Materials

Tissue culture dishes were from Greiner BIO-ONE (Frickenhausen, Germany); white polystyrene 96-well culture plates for GSH measurements from PerkinElmer Europe (Zaventen, Belgium); high glucose Dulbecco's modified Eagle medium (DMEM) from PAA Laboratories (Linz, Austria); fetal calf serum (FCS), l-glutamine, penicillin/streptomycin, and phenol red-free DMEM, TriZol reagent for RNA purification from Invitrogen (Karlsruhe, Germany); bovine serum albumin (BSA), l-cystine, l-glutamate, 6-diazo-5-oxo-l-norleucine (DON), l-glutamic acid, N-(2-hydroxyethyl)-piperazine-N′-(2-ethanesulfonic acid) (HEPES), hydrogen peroxide (H2O2), monochlorobimane (MCB), methylthiazoltetrazolium (MTT), paraformaldehyde, l-trans-pyrrolidine-2,4-dicarboxylic acid (PDC) were from Sigma-Aldrich (Schnelldorf, Germany). Poly-L-lysine was obtained from Biochrom (Berlin, Germany). Goat serum was from Seratec (Göttingen, Germany). DL-threo-β-benzyloxyaspartic acid (TBOA) was from Tocris (Northpoint, UK). L-[35S]-Cystine (specific activity 40–250 mCi/mmol), [α-32P]-CTP and Cy2- and Cy3-labeled secondary antibodies were obtained from Amersham (Freiburg, Germany). Oligonucleotides were synthesized by MWG Biotech AG (Ebersberg, Germany). All other chemicals and diluents were obtained from Merck Eurolabs (Darmstadt, Germany). MTT was reconstituted in PBS, MCB in ethanol, all other compounds in water. For viability assays, TBOA was dissolved in DMEM and pH re-adjusted by adding HCL monitored by phenyl red. For glutamate release assays, TBOA was dissolved in DMSO at a final concentration of 100 mm.

Cell culture and viability assays

HT22 cells were a gift of Wulf Paschen, Max-Planck-Institute for Neurological Research, Cologne, Germany. Cells were cultivated at 37°C in a 5% CO2 atmosphere and DMEM containing 5% FCS, 2 mm glutamine, 10 mm HEPES, pH 7.4, 100 IU/mL penicillin, and 100 µg/mL streptomycin. Cells were passaged by dissociation with 0.005% trypsin and 0.002% EDTA every other day. For viability assays, 5000 cells were seeded in 100 µL into 96-well plates. To induce cell death, glutamate or H2O2 were added 24 h later for 8 or 24 h, other experimental agents were added as indicated. PDC and TBOA were added immediately before the addition of glutamate. Cell survival was judged by phase contrast microscopy and assayed by the MTT method as described previously (Hansen et al. 1989; Lewerenz et al. 2003) in at least three independent experiments.

Selection of HT22 cells resistant to glutamate toxicity

For selection of glutamate resistant HT22 cells, 6 × 105 cells were seeded in 92 mm culture dishes and exposed 24 h later to 10 mm glutamate for 24 h. Surviving cells were expanded and again exposed to 10 mm glutamate, followed by two additional cycles of 10 mm glutamate with less cells (3 × 105) per dish. Finally, cells remained exposed to 20 mm glutamate for 48 h during the last of the four selections. These cells were further propagated in the presence of 10 mm glutamate and named resistant HT22 cells (HT22R).

Transient transfection

An xCT construct was a generous gift from Andy Shih (Vancouver, Canada); EAAT3 was obtained from clone IRAKp961F0847Q (RZPD, Berlin, Germany). Both open reading frames were subcloned into the expression vector pCI-neo (Promega, San Luis Obispo, USA) and high purity plasmid DNA prepared using Nucleobond AX 500 columns (Machery & Nagel, Düren, Germany) according to the manufacturer. For transfection, 15 × 106 confluent HT22 cells were resuspended in 500 µL PBS and incubated on ice for 10 min with 10 µg plasmid DNA. Electroporation was carried our with a Genepulser (Biorad, Munich, Germany) at 960 F and 250 mA with a time constant of 17–20 s. Immediately after transfection, cells were plated in three 92 mm culture dishes for 24 h before seeding 5000 cells into 96-well plates for toxicity or GSH assays.

Cystine uptake

2.5 × 104 HT22 or HT22R cells per well were seeded in 24-well plates. HT22R cells were grown in regular growth medium or in medium supplemented with 10 mm glutamate. After 24 h, cells were washed with a modified Hank's buffered salt solution (HBSS) as reported previously (Murphy et al. 1989) and incubated with 50 µm l-[35S]-Cystine (specific activity 40 mCi/mmol) for 20 min alone or in the presence of 1 mm unlabeled cystine to ensure specificity of cystine uptake. Cystine uptake was also measured in the presence of different concentrations of glutamate. After 20 min, cells were washed three times with 0.5 mL ice-cold HBSS and lysed with 150 µL 0.5 m NaOH. Cell lysates were neutralized with 150 µL 0.5 m HCl. Protein concentration was determined by the bicinchoninic acid-based method (Micro BCA Protein Assay; Pierce, Bonn, Germany) in 100 µL. Another 100 µL were dried overnight in XtalScint Ready Caps (Beckmann, Fullerton, USA) for solid-phase scintillation counting using a LS 6000IC counter (Beckmann, Fullerton, USA). Specific cystine uptake was calculated as counts per minute (cpm)/mg protein after subtraction of cpm/mg protein in the presence of 1 mm unlabeled cystine. Each experiment was done in triplicates and repeated three times.

Glutamate release assays

2 × 105 HT22R cells per well were seeded in six-well plates and grown for 24 h in medium with 10 mm glutamate. Cells were washed twice with 1 mL HBSS and then layered with 200 µL HBSS, HBSS containing 200 µm cystine ± 20 µm TBOA. Each condition was performed in triplicates. Cells were incubated for 20 min at 37°C. Then HBSS was aspirated, mixed with an equal volume ice-cold 8% 5-sulfosalicylic acid in H2O and stored on ice for one hour. After centrifugation with 16.000 × g for 15 min at 4°C, supernatant was removed and stored at − 20°C until analysis. For amino acid analysis, samples were first adjusted to 10% 5-sulfosalicylic acid, then neutralized by adding HCl at a final concentration of 50 mm. 300 mg/L norvaline were added as internal standard. Amino acids were separated using a Biochrom 20 Amino Acid Analyser (Amersham Pharmacia Biotech, Freiburg, Germany), an Ultrapac 8 column (Amersham Pharmacia Biotech), 0.2–0.9 m lithium citrate, pH 2.8–3.5, temperature 35°C− 65°C, followed by fluorometrical detection at 450 nm/400 nm after postcolumn derivatisation with O-phthalaldehyde.

Intracellular glutamate measurement

105 HT22R cells per well were seeded in six-well plates and grown for 24 h. Then, glutamate at a final concentration of 40 mm ± 900 µm PDC were added. Each condition was done in six replicates with triplicates of separate six-well plates. After 15 h, half of the plates were stored on ice, medium removed and cells washed twice with ice-cold PBS. PBS was completely removed and 200 µm ice-cold 4% 5-sulfosalicylic acid in H2O added. Cells in plates were shock-frozen by placing then on dry ice and thawed three times. The cell extract was aspirated and stored on ice for one additional hour. After centrifugation with 16 000 × g for 15 min at 15°C, supernatant was removed and stored at − 20°C until analysis. Glutamate concentration was quantified as described above. Control plates were handled equally, but after 15 h of glutamate exposure, viability was measured by MTT. Glutamate concentrations were normalized to the relative MTT optical densities of control cells.

Detection of intracellular GSH

5000 HT22 and HT22R cells were plated in white 96-well plates. After 24 h, cells were incubated with or without glutamate as indicated. Cells were carefully washed twice with phenol red-free DMEM and incubated with 20 µm MCB for 5 min at 37° protected from light. MCB fluorescence was measured by a SpectraMax Gemini (Molecular Devices, Ismaning, Germany) using SoftmaxPro 3.1.1 software (Softmax, San Diego, USA). Excitation wavelength was 393 nm and emission wavelength 485 nm. After measurement, one volume normal medium and MTT were added and cell viability was assessed two hours later. Results therefore indicate MCB fluorescence normalized to cell viability.

Northern blotting

5 × 105 HT22 and HT22R cells were plated in 92 mm culture dishes and cultivated for 24 h with or without glutamate, respectively. Additionally, sensitive HT22 cells were harvested after 6 h of 10 mm glutamate. RNA was prepared using the TriZol reagent procedure according to the manufacturer. 5 µg total RNA was separated by denaturing 1.2% agarose gel electrophoresis in the presence of 2.2 m formaldehyde and transferred to a nylon membrane. Specific probes were generated by linear PCR in the presence of 50 µCi [α-32P]-dCTP using reverse primers and 25 ng template generated by PCR with specific primers for catalase (forward 5′-AGAAGCCTAAGAACGCAATTC-3′ and reverse 5′-ATGTGAAATCACTGCGTATTAGC-3′), GSH peroxidase 1 (GPx, forward 5′-GGCACCACGATCCGGGACTA-3′ and reverse 5′-TTAGGTGGAAAGGCATCGGGAAT-3′), and xCT (forward 5′-TGTTCGCTGTCTCCAGGTTATTC-3′ and reverse 5′-GCTTGCCTCACTGTATGACTTGC-3′). Specific activity was > 105 cpm/µg DNA. Blots were hybridized overnight in ExpressHyb (Clontech, Palo Alto, USA) solution at 68°C and washed under high stringency conditions. Autoradiography was performed by exposure to Biomax MS films (Kodak, Stuttgart, Germany) and a phospho-imaging system (Fujix, Japan) for quantitative analysis. Blots were stripped and reprobed for the housekeeping gene glyceraldehyde-3-phosphate dehydrogenase (GPDH) using a template generated by PCR with the forward primer 5′-ACC ACA GTC CAT GCC ATC AC-3′ and the reverse primer 5′-TCCACCACCCTGTTGCTGTA-3′ as described above. Cycle parameters for all PCR reactions were 95°C 30 s, 60°C 10 s and 72°C 5 min for 30 cycles. Mean ± SEM of regulation over GPDH was calculated by comparing the band hybridization intensities obtained by phosphoimaging and analysis using TINA software (Raytest, Straubenhardt, Germany) from two individual Northern blots.

RT-PCR

Total RNA from HT22R cells was digested with DNase I (Invitrogen) to eliminate residual genomic DNA and reverse transcribed with oligo(dT)-primers using the SUPERSRIPTTM. first strand cDNA synthesis system (Invitrogen). 50 ng of HT22R cDNA was incubated with 10 pg of primer (EAAT1: forward 5′-CTCTTCAGTCCCTAATCAGC-3′, reverse 5′-GACCATCCACCCAGCGTATG-3′; EAAT2: forward 5′-CCGCACACAACTCTGTCGTA-3′, reverse 5′-GCTGAGAATCGGGTCATTAT-3′; EAAT3: forward 5′-CAGCGGGTTCAGGGACAATAC-3′, reverse 5′-TAAGCCATCTTCGGGAATTA-3′; EAAT4: forward 5′-TGATTGTGCTCACATCCGTC-3′, reverse 5′-AATCGAATCCCCCAGTACAT-3′; EAAT5: forward 5′-CCGGCCAAGGGACTAGCAGTAC-3′, reverse 5′-TTTATCACATTATCGCAAACA T-3′), 25 ng of dNTPs, 10 mU of Taq-enzyme (New England Biolabs), 50 ng of Mg2+ and subjected to 35 cycles in a T-Gradient PCR-cycler (Biometra, Göttingen, Germany) with an annealing temperature of 60°C. Probes were separated on a 3% agarose gel containing 0.1‰ ethidium bromide. Experiments were repeated three times with similar results.

Western blotting

HT22 and HT22R cells were seeded at a density of 5 × 105 cells in 92 mm cell culture dishes and treated with or without 10 mm glutamate respectively. 24 h after plating, cells were washed with PBS and directly lysed in boiling protein lysate buffer (1% sodium dodecyl sulfate, 10% glycerol, 10%β-mercaptoethanol, 8% 0.5 m Tris pH 6.8, 0.01‰ bromophenol blue). Proteins were separated by 10% SDS-PAGE and transferred to a nitrocellulose membrane (Schleicher und Schüll, Dassel, Germany). Membranes were blocked with 5% skim milk in TBS-T (20 mm Tris buffer pH 7.5, 0.5 m NaCl, 0.1% Tween 20) and incubated with α-EAAT3 (1 : 100, generous gift from J. D. Rothstein, Johns Hopkins University). Subsequently, blots were washed in TBS-T and incubated for one hour with the secondary antibody (anti-rabbit, 1 : 7000 Promega, Madison, USA) conjugated to horseradish peroxidase. After a second wash, labeled proteins were detected using the ECL-reagent (Lumi-Phos WB, Pierce, Bonn, Germany). Experiments were done three times with similar results.

Immunofluorescence

2.5 × 104 HT22 and HT22R cells were plated onto 10 mm-cover slips coated with poly-L-lysine in 24 well-culture dishes and grown in either normal HT22 medium or medium supplemented 10 mm glutamate, respectively. After 24 h, medium was aspirated, and cells fixed in 4% paraformaldehyde in PBS for 20 min. After three washings with PBS, cells were permeabilized with ethanol and non-specific binding sites blocked by incubation with 10% goat serum and 0.2% BSA in PBS for 20 min. Primary antibodies were α-MAP2 (microtubule-associated protein 2, mouse monoclonal, Boehringer Mannheim, Germany, 1 : 1000), α-NFM-P+ (highly phosphorylated medium Mr neurofilament, mouse monoclonal, Zymed, San Francisco, USA, 1 : 4000) and α-NFH (high Mr neurofilament, rabbit polyclonal antibody, Serotec, Düsseldorf, Germany, 1 : 500). Cy3-coupled goat anti-rabbit-IgG and Cy2-coupled goat anti-mouse-IgG secondary antibodies were diluted 1 : 3000 and 1 : 2000, respectively.

Results

  1. Top of page
  2. Summary
  3. Experimental procedures
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

Reduced glutamate-induced GSH depletion in HT22 cells resistant to oxidative stress

We repeatedly exposed HT22 cells for 24 h to high concentrations of glutamate and propagated the few surviving cells in 10 mm glutamate to obtain cells resistant to oxidative stress. The new strain named HT22R withstood up to 40 mm glutamate for 24 h with only a ∼40% decrease in viability. In contrast, ∼96% of wild type HT22 cells died after exposure to only 5 mm glutamate. This resistant phenotype was not restricted to oxidative stress elicited by glutamate; HT22R cells were also more resistant to hydrogen peroxide (H2O2), which directly induces oxidative stress. This effect was, however, less prominent (Fig. 1a). Glutamate-induced GSH depletion was drastically reduced in HT22R cells (Fig. 1b); 2.5 mm glutamate for six hours, which led to a ∼60% reduction in wild type HT22 cells, was even accompanied by a non-significant trend to higher GSH levels in HT22R cells. Even so, HT22R cells had the same morphology as the parental cell line and both displayed the same immunoreactivity against the neuronal markers MAP2, NFM-P+, and NFH (Fig. 1c).

image

Figure 1.  Glutamate-resistant HT22 cells are partially cross-resistant to H2O2 and exhibit a reduction in glutamate-induced GSH depletion (a) 5000 HT22 (S, square) or HT22R (R, triangle) cells per well were seeded into 96-well plates and grown for 24 h before being exposed to the indicated concentrations of glutamate or H2O2 for 24 h. Viability was assessed by MTT assays. (b) Intracellular GSH content determined by MCB-fluorescence normalized to MTT optical density of HT22 and HT22R cells in response to six hours of the indicated concentrations of glutamate. Excitation wavelength for MCB was 393 nm and emission was 485 nm. Each data point in a and b shows the mean ± SEM of three pooled independent experiments with four replicates for survival assays and six for MCB-fluorescence. (c) Phase contrast microscopy and immunostaining of glutamate resistant and sensitive HT22 cells with antibodies against MAP2 (α-MAP2), NFH (α-NFH-p +), and NFM-P+ (α-NFM-P+).

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Functional up-regulation of the xCT subunit of the glutamate/cystine antiporter inline image

The observed lack of glutamate-induced GSH depletion and the concomitant increase in viability could be explained by an increase in the general competence of HT22R cells to detoxify ROS, by a specific increase in the capacity to generate GSH or by a combination of the two. Quantification of RNA coding for enzymes that metabolize ROS like catalase and GSH peroxidase (GPx) on one hand, and xCT, the specific subunit of the inline image antiporter, on the other hand revealed a prominent and specific seven-fold up-regulation of xCT in HT22R cells. Catalase mRNA was much less increased by 1.9-fold (Fig. 2a). GPx expression did not differ between the two cell lines. The observed up-regulation of xCT mRNA was reflected in a similar increase in radioactive cystine uptake of 8.2-fold (Fig. 2b) indicating that the up-regulated xCT mRNA is translated into functional protein. We already noted this almost direct relationship between xCT mRNA and cystine uptake in a previous study, where we showed that treating HT22 cells with cholera toxin protects from oxidative glutamate toxicity by inducing xCT mRNA (Lewerenz et al. 2003). Sensitive HT22 cells only expressed very little xCT RNA, which was increased approximately three-fold in response to GSH depletion induced by 6 h of 10 mm glutamate (Fig. 2a, right panel). In addition to the prominent up-regulation of system Xc in HT22R cells, glutamate-mediated inhibition of cystine uptake was shifted to significantly higher concentrations in HT22R cells. In wild type HT22 cells the EC50 for glutamate inhibition of cystine uptake was ∼66.6 µm (95% confidence interval (CI): 20.5–216.6 µm) as compared to 365 µm (95% CI: 269.5–494.4 µm) in HT22R cells (Fig. 2c). Omission of glutamate from HT22R cells for 24 h did not reduce cystine uptake in the absence of glutamate nor did it induce significant differences in the glutamate-mediated inhibition of cystine uptake (not shown). Interestingly, low concentrations of glutamate seemed to stimulate rather than inhibit cystine transport in HT22R cells. We conclude that an absolute increase in system inline image activity and a reduction in glutamate-mediated inhibition of cystine uptake contribute to the preserved GSH homeostasis in HT22R cells.

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Figure 2.  Functional up-regulation of the cystine/glutamate antiporter system inline image subunit xCT and EAATs in HT22R cells (a) Quantitative analysis of two Northern blots (representative blot probed with xCT and gapdh to the right) loaded with 5 µg total RNA from HT22R (R) and HT22 (S ± 6 h 10 mm Glu) cells probed with xCT, catalase (Cat), and GSH peroxidase-1 (GPx). Shown is mean fold regulation ± SEM R over S normalized to the housekeeping gene gapdh. (b) Corresponding uptake of radioactively labeled cystine normalized to cell protein in HT22 cells or HT22R cells. (c) Cystine uptake in the presence of the indicated concentrations of glutamate. Graphs show mean ± SEM of pooled data from four experiments each performed in triplicate and normalized to cystine uptake in the absence of glutamate. Statistical analysis in b and c was made using two-way anova and Bonferroni's multiple comparisons test (*p < 0.001 S vs. R). (d) Functional EAAT transporters are expressed in HT22R cells. 2 × 105 HT22R cells per well were grown for 24 h and exposed to HBSS containing 200 µm cystine ± 20 µm TBOA for 20 min. Glutamate release was quantified by chromatography and normalized to control conditions without cystine. Graph represents the mean ± SEM of two experiments performed in triplicates (*p < 0.05, unpaired two-tailed student's t-test). Right panel: Expression of EAAT1-5 mRNA in HT22R cells as shown by PCR with 35 cycles. PCR specificity was verified by sequencing. Western blot of extracts from HT22 and HT22R cells probed with antiserum against EAAT3. Equal amounts of protein were applied per lane. Molecular mass standards are indicated.

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Excitatory amino acid transporters are functionally expressed in HT22 cells

We hypothesized that excitatory amino acid transporter (EAATs) might underlie the phenomenon of decreased glutamate sensitivity of cystine import in HT22R cells. These transporters import glutamate with high affinity (Dunlop et al. 1993), which reduces the glutamate concentration at the extracellular surface and might support the glutamate/cystine antiporter by increasing the intracellular glutamate concentration. To demonstrate that EAATs are functionally expressed in HT22R cells and closely cooperate with system inline image antiporters, we investigated whether glutamate exported by system inline image is re-imported by EAATs. Indeed, in HT22R cells, cystine-stimulated glutamate release is increased by the specific non-transportable EAAT inhibitor TBOA (Fig. 2d). Next, we determined which of the five EAATs is expressed in HT22R cells. RT-PCR revealed the presence of EAAT1-3 but not EAAT 4 or 5 in HT22R cells (Fig. 2d right panel). As shown by immunoblotting, EAAT3 protein is also expressed in HT22 cells and appeared increased in resistant cells.

Inhibition of glutamate import increases glutamate sensitivity

Next, we tested the impact of EAAT inhibition on glutamate sensitivity of HT22R cells. TBOA up to concentrations of 40 mm did not exacerbate the limited toxicity induced by 40 mm glutamate, whereas higher concentrations showed an unspecific toxic effect (Fig. 3 A left). On the other hand, PDC dose-dependently induced cell death in the presence of glutamate but did not affect cell survival in the absence of glutamate (Fig. 3a, right). PDC is a transportable EAAT inhibitor inducing hetero-exchange of glutamate, thereby reversing glutamate transport (Waagepetersen et al. 2001). Thus, PDC might be much more effective to abolish the glutamate-concentrating action of EAATs. Furthermore, the high glutamate concentrations needed to induce cell death in HT22R cells demand high concentrations of EAAT inhibitors. To demonstrate that TBOA can also increase glutamate-induced cell death, we investigated the effect of TBOA and PDC in sensitive HT22 cells, in which 40-fold lower glutamate concentrations are partially toxic. TBOA proved to be effective to induce cell death in concentrations higher than glutamate (Fig. 3b, left). As predicted, much lower concentrations of PDC were effective to exacerbate cell death induced by 1 mm glutamate in HT22 cells compared to HT22R cells (Fig. 3b, right). Thus, inhibition of EAATs by PDC and TBOA exacerbates glutamate-induced cell death. On the other hand, inhibition of intracellular glutamate synthesis by the phosphate-activated glutaminase inhibitor, DON, had no effect on cell death glutamate-induced in HT22 or HT22R cells (not shown).

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Figure 3.  Inhibition of glutamate transport enhances glutamate-induced cell death in HT22R cells (a) 5000 HT22R (a) or HT22 (b) cells per well were seeded into 96-well plates and grown for 24 h before being exposed to 40 mm (a), respectively, 1 mm (b) glutamate (triangle) or vehicle (square) in the presence of the indicated concentrations of TBOA or PDC. Glutamate and experimental substances were removed in b by medium change after eight hours. Viability was assessed 24 h later by MTT assays. Each data point shows the mean ± SEM of three pooled independent experiments with four replicates. Statistical analysis was made by two-way anova and Bonferroni's multiple comparisons test (*p < 0.05, **p < 0.01, ***p < 0.001 Glu vs. no Glu).

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PDC exacerbates glutamate-induced cell death by decreasing intracellular glutamate accumulation and increasing GSH depletion

If PDC increases glutamate toxicity by inhibiting and partially reversing EAAT function, PDC should decrease intracellular glutamate accumulation and increase glutamate-induced GSH depletion in HT22R cells. We first determined the time course of PDC toxicity. In the presence of 40 mm glutamate, PDC starts to be toxic at 18 h (Fig. 4a). At 15 h no cell death was noticeable but PDC significantly decreased glutamate-induced glutamate accumulation in HT22R cells (Fig. 4b). Simultaneously, intracellular GSH content dropped by 25% shifting the residual GSH to ∼31% of cells not treated with glutamate or PDC (Fig. 4b right panel), thereby becoming considerably closer to the 20% shown previously to be sufficient to induce the exponential increase in ROS, which initiates cell death (Tan et al. 2001). Thus, inhibition of EAATs by PDC exacerbates glutamate-induced cell death by decreasing intracellular glutamate accumulation and increasing GSH depletion.

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Figure 4.  PDC exacerbates glutamate-induced cell death by decreasing intracellular glutamate and GSH (a) Time course of PDC-induced cell death. 5000 HT22R cells were grown as in 2a and treated with 40 mm glutamate ± 900 µm PDC for the indicated periods followed by MTT assay. Data are normalized to control cells without glutamate and PDC. (b) Effect on intracellular glutamate and GSH before the onset of cell death. 105 HT22R cells per well were seeded in six-well plates and grown for 24 h before exposure to 40 mm glutamate ± 900 µm PDC for 15 h. Intracellular glutamate was quantified by chromatography and GSH by MCB fluorescence. Both were normalized to MTT optical density of control cultures and to control cells ± PDC without glutamate set as 100%. Each graph represents data of three independent experiments performed in six replicates. Statistical analysis was made separately for each time point. Unpaired two-tailed student's t-test (a, *p < 0.0001; b, *p < 0.05; c, *p < 0.001 PDC vs. control).

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Overexpression of the neuronal glutamate transporter EAAT3 and xCT cooperatively protects against oxidative glutamate toxicity by increasing intracellular GSH

The glutamate-resistant phenotype of HT22R cells might therefore be at least in part a consequence of increased residual GSH synthesis driven by improved glutamate turnover through cooperation of system inline image providing cystine for GSH synthesis and EAATs providing glutamate for inline image exchange. To provide further evidence for this hypothesis, we transiently transfected sensitive HT22 cells with the neuronal EAAT, EAAT3, the specific system inline image subunit, xCT, or both. Transfection of both constructs alone or together was highly protective against eight hours of glutamate at various concentrations (Fig. 5a). Transfected cells survived 5 mm glutamate for eight hours from ∼10% in mock- to ∼44% in EAAT3- and ∼59% in xCT-transfected cells. Approximately 80% of double-transfected cells survived suggesting that the transporters work in concert to enhance survival. The degree of protection was paralleled by an increase in residual GSH after six hours of 5 mm glutamate (Fig. 5b), suggesting that both transporters protect by increasing GSH. Interestingly, higher concentrations of glutamate (20 mm, Fig. 5a) or longer exposure (24 h of 5 mm, Fig. 5c) revealed a marginal or absent protective effect of EAAT3 over-expression, whereas additional EAAT3 over-expression in xCT over-expressing cells had a prominent effect on cell survival. This again supports the assumption of a synergistic action of EAAT3 and xCT.

image

Figure 5.  Cooperative action of the excitatory amino acid transporter EAAT3 and xCT on cell death (a) 5000 HT22 cells were electroporated with empty vector (square), EAAT3 plus empty vector (upside-down triangle), xCT plus empty vector (triangle), or xCT and EAAT3 (diamond). Equal amounts of DNA were transfected. 24 h later cells were seeded into 96-well plates and grown for 24 h before being exposed to the indicated amounts of glutamate for eight hours. Viability was assessed 24 h after the addition of glutamate by MTT assays. (b) Intracellular GSH content determined by MCB-fluorometry normalized to MTT optical density in transfected HT22 cells ± 5 mm glutamate for six hours. Statistical analysis was made by two-way anova and Bonferroni's multiple comparisons test (*p < 0.01 vs. empty vector and 5 mm glutamate). (c). Same experimental setup as in (a) with 5 mm glutamate for 24 h. Statistical analysis as above (n.s. p > 0.05: EAAT3 vs. empty vector, *p < 0.001: EAAT3 + xCT vs. xCT alone). (d) HT22 cells treated as in (a) but exposed to the indicated concentrations of H2O2 for eight hours. Graphs show mean ± SEM of pooled data from three experiments each performed in four replicates for survival assays normalized to viability in the absence of glutamate for each transfection condition separately, or six replicates for GSH-measurements normalized to mock-transfected HT22 cells in the absence of glutamate.

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If EAATs protect against glutamate by allowing system inline image to import cystine despite high extracellular glutamate, EAAT-overexpression should be less effective in paradigms of oxidative stress not dependent on system inline image inhibition. Indeed, in contrast to xCT, EAAT3 alone did not significantly decrease cell death induced by H2O2, a direct oxidant, which rapidly oxidizes GSH (Dringen et al. 2005) (Fig. 5d).

Discussion

  1. Top of page
  2. Summary
  3. Experimental procedures
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

Selection of neuronal cell lines resistant to various insults has been used previously to identify novel mechanisms of protection against oxidative stress (Sagara et al. 1996, 1998; Dargusch and Schubert 2002). Sagara et al. (1998) described three HT22 cell clones adapted to high concentrations of glutamate with the ability to rapidly replenish GSH levels after depletion. The resistant cells described here, in contrast, exhibited a massive basal increase in cystine uptake most probably mediated by system inline image as suggested by the increase in mRNA of the specific transporter subunit xCT and the fact that cystine uptake could be inhibited by glutamate. These differences might be due to the polyclonal nature of our cells and/or the selection procedures used.

xCT belongs to a family of phase II response genes involved in defense against oxidative stress (Sasaki et al. 2002), which contain an antioxidant-response element (ARE) in their promoter regions (Ishii et al. 2000). The nuclear factor Nrf2 binds to this element and activates transcription (Nguyen et al. 2003). Other antioxidative enzymes like catalase and GPx (Thimmulappa et al. 2002; Shih et al. 2003) are also induced by Nrf2, but the observed up-regulation of xCT far exceeds the transcriptional changes of these other ARE-dependent genes suggesting additional factors like activating transcription factor 4 (ATF4), which can induce xCT transcription via an amino acid response element (Sato et al. 2004). This could explain the up-regulation of xCT in wild type HT22 cells caused by six hours of 10 mm glutamate (Fig. 2a), which induces a significant decrease in intracellular cystine and subsequent cell death 6 h later. However, HT22R cells do not suffer from cystine deprivation in the presence of 10 mm glutamate as judged by the lack of GSH depletion and nevertheless exhibit the prominent induction of xCT mRNA, which speaks against a strong participation of this pathway. The exact mechanism of xCT induction in these cells therefore remains to be investigated.

We conclude that the up-regulation of xCT is at least in part responsible for the resistant phenotype as transient overexpression induced a robust protection against glutamate toxicity. Apparently, the other transporter subunit, the 4F2 heavy chain, must be expressed in excess in HT22 cells as described for HEK293 cells (Shih and Murphy 2001). Overexpression of xCT leads to decreased glutamate-induced GSH depletion suggesting that even the residual system inline image activity in the presence of high extracellular glutamate is sufficient to support GSH homeostasis. We also observed a reduced glutamate inhibition of cystine uptake, which should decrease glutamate-induced GSH depletion even further. EAATs probably contribute to this phenomenon, because blocking EAATs with PDC and TBOA exacerbates oxidative glutamate toxicity. PDC reduces glutamate-induced intracellular glutamate and GSH probably not by direct inhibition of system inline image (Patel et al. 2004). We think that the marked difference in effective concentrations of the two inhibitors is most probably a consequence of different transport characteristics: Although TBOA is slightly more potent than PDC in inhibiting [3H]d-aspartate uptake in neurons and astrocytes, only PDC induced an [3H]d-aspartate release from preloaded cells (Waagepetersen et al. 2001). Therefore, PDC is a transportable inhibitor leading to heteroexchange of glutamate. In contrast, TBOA is a non-transportable inhibitor. In our system, substances that not only interfere with glutamate up-take but reverse that function like PDC should be much more effective, which is exactly what we found. Therefore, we propose that EAATs are involved in GSH homeostasis in HT22 cells. Indeed, overexpression of the neuronal glutamate transporter EAAT3 protects HT22 cells from oxidative glutamate toxicity with an over-additive effect to xCT alone at high glutamate concentrations and long exposure.

In addition to glutamate, EAATs have been shown to transport cystine and cysteine (Knickelbein et al. 1997; McBean and Flynn 2001; Shanker et al. 2001; Allen et al. 2002; Chen and Swanson 2003; Himi et al. 2003). However, the affinities of system XAG- (the biochemical description of EAAT-mediated uptake (Kanai and Hediger 2004)) are very low for cystine (Km > 500 µm) and high for glutamate (7.5 µm) in rat alveolar type II cells (Knickelbein et al. 1997). Similar values were obtained in brain synaptosomes, which have a Km for sodium-dependent cystine uptake of 473 µm and an IC50 for transport of 300 µm cystine of 9.1 µm glutamate (McBean and Flynn 2001). Cysteine is not present in cell culture medium, but exported GSH has been shown to effectively reduce cystine to cysteine in the extracellular space (Wang and Cynader 2000). However, the affinities of system XAG- for cysteine are also considerably lower than those for glutamate (96.1 µm vs. 7.5 µm) (Knickelbein et al. 1997). In contrast, the affinity of system inline image is higher for cystine than for glutamate (7.5 µm vs. 35 µm) (Murphy et al. 1989). Thus, it is very unlikely that cystine or cysteine uptake via EAATs contributes to the diminished glutamate inhibition of cystine uptake observed in resistant HT22 cells. Given the concentration of cystine in the medium of 200 µm, uptake of both amino acids should also be negligible in the presence of millimolar concentrations of glutamate. We conclude that glutamate uptake via EAATs is responsible for the observed protective effects. This could be accomplished either by decreasing glutamate levels in proximity to the cell membrane, thereby reducing inhibition of cystine uptake by system inline image. Indeed, in HT22R cells, EAAT inhibition by TBOA increased in cystine-evoked release of glutamate. In another cell system, a robust increase in cystine-evoked system inline image-mediated glutamate release was found by inhibiting EAATs by switching to sodium-free conditions (Patel et al. 2004). Both findings indicate a tight functional cooperation of both transport systems with rapid re-uptake of glutamate released via system inline image by EAATs under physiological conditions. Thus, EAATs should facilitate cystine uptake via system inline image by extracellular glutamate accumulation close to the antiporter. On the other hand, Bannai (1986) showed that cystine uptake via system inline image is tightly regulated by intracellular glutamate concentrations. Thus, EAATs could also stimulate system inline image activity by increasing intracellular glutamate. We speculate that EAAT-mediated glutamate uptake supports system inline image and thereby protects against oxidative glutamate toxicity in both ways (Fig. 6). This view is further strengthened by the fact that the over-additive effect of EAAT3 expression on xCT-mediated protection is not evident after treatment with hydrogen peroxide, because glutamate inhibition of system inline image activity, which could be ameliorated by EAATs, does not play a significant role is this model of oxidative stress-induced cell death. Alternatively, H2O2 could rapidly inactivate EAATs by oxidation (Trotti et al. 1998).

image

Figure 6.  Diagram depicting the proposed cooperative action of EAAT3 and xCT. Glutamate import via EAATs can reduce glutamate concentrations in the extracellular space in proximity to the cell membrane, thus reducing competition between glutamate and cystine (CSSC) for binding to xCT, as well as fuel cystine import by providing intracellular glutamate for re-export.

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The cooperative action of EAAT and system inline image appears to be a more general mechanism not restricted to nerve cells as stimulation of GSH synthesis by EAAT-mediated glutamate uptake has also recently been demonstrated in human macrophages (Rimaniol et al. 2001) and retinal glial cells (Reichelt et al. 1997). It was also shown previously that CHO-K1 cells defective for glutamate transporters exhibit decreased system inline image activity (Igo and Ash 1998). In the rat striatum, the extracellular glutamate concentration seems to be determined mainly by the balance of glutamate release via system inline image and uptake via EAATs (Baker et al. 2002).

In summary, the results presented here show that up-regulation of system inline image is a prominent protective response in the adaptation against oxidative stress and support the idea that synergistic action of system inline image and EAATs is critical for GSH homeostasis in nerve cells and tissues where both are expressed like the brain. Whether this participates in death and survival of specific cell populations in neurodegenerative diseases like Alzheimer's or Parkinson's disease remains to be investigated.

Acknowledgements

  1. Top of page
  2. Summary
  3. Experimental procedures
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

We thank Dipl.-Ing. Maren Stehn for the amino acid analysis, Chica Schaller for continuous support and Pamela Maher for helpful discussions and critical reading of the manuscript. The Deutsche Forschungsgemeinschaft within the Graduiertenkolleg 255 and the Dr Kurt and Irmgard Meister-Stiftung funded this work.

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  5. Discussion
  6. Acknowledgements
  7. References
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