A Cytotoxic,Co-operative Interaction Between Energy Deprivation and Glutamate Release From System x c − Mediates Aglycemic Neuronal Cell Death

Abstract

The astrocyte cystine/glutamate antiporter (system x_c⁻) contributes substantially to the excitotoxic neuronal cell death facilitated by glucose deprivation. The purpose of this study was to determine the mechanism by which this occurred. Using pure astrocyte cultures, as well as, mixed cortical cell cultures containing both neurons and astrocytes, we found that neither an enhancement in system x_c⁻ expression nor activity underlies the excitotoxic effects of aglycemia. In addition, using three separate bioassays, we demonstrate no change in the ability of glucose-deprived astrocytes—either cultured alone or with neurons—to remove glutamate from the extracellular space. Instead, we demonstrate that glucose-deprived cultures are 2 to 3 times more sensitive to the killing effects of glutamate or N-methyl-D-aspartate when compared with their glucose-containing controls. Hence, our results are consistent with the weak excitotoxic hypothesis such that a bioenergetic deficiency, which is measureable in our mixed but not astrocyte cultures, allows normally innocuous concentrations of glutamate to become excitotoxic. Adding to the burgeoning literature detailing the contribution of astrocytes to neuronal injury, we conclude that under our experimental paradigm, a cytotoxic, co-operative interaction between energy deprivation and glutamate release from astrocyte system x_c⁻ mediates aglycemic neuronal cell death.

Keywords

aglycemia cystine glutamate neuronal injury glucose deprivation cell culture

Introduction

Hypoglycemia is a common and serious medical emergency that arises as a complication in patients attempting to tightly regulate glucose levels with insulin and is the limiting factor in the development of strategies that aim to maintain normoglycemia in diabetics (Lincoln et al., 1996; Lacherade et al., 2009). Hypoglycemia is also a complication in neonates and in patients with insulin-producing tumors (Anderson et al., 1967; de Herder, 2004) and occurs as a consequence of brain ischemia. Idiopathic, diabetic, and reactive hypoglycemia, (i.e., low blood glucose concentrations of <3.9 mM) can produce a variety of cognitive symptoms resulting from an inadequate supply of glucose to the brain. Owing to the fact that there is a linear relationship between blood and brain glucose levels (Choi et al., 2001), a large drop in blood glucose concentration (<2.0 mM) renders the brain aglycemic, which can result in a severe impairment of cognition, seizures, unconsciousness, and coma. If glucose levels are not restored within 30 min time, death of vulnerable cortical and hippocampal neurons also ensue (Suh et al., 2003; Auer, 2004).

Despite being initiated by glucose deprivation (GD), both in vitro and in vivo investigations showed that hypoglycemic neuronal cell death is not a direct result of energy failure but instead is mediated by glutamate excitotoxicity (Engelsen et al., 1986; Linden et al., 1987; Monyer et al., 1989; Papagapiou and Auer, 1990; Jackman et al., 2012). Adding to this understanding, we recently described system x_c⁻—a transporter that exports L-glutamate in exchange for L-cystine (Bannai, 1984b)—in astrocytes as a source of glutamate required for the initiation of non-cell autonomous neuronal injury following GD in vitro (Jackman et al., 2012). The precise mechanism by which injury is facilitated by astrocyte system x_c⁻ was not ascertained therein and thus is the focus of this study.

Maintenance of extracellular glutamate concentrations within a narrow physiological range involves control of its release as well as its uptake, both of which are efficiently managed by glutamate transporters expressed predominantly by astrocytes. Pertinently, numerous studies demonstrate that system x_c⁻ is an important contributor to the ambient extracellular glutamate levels that bathe the central nervous system (CNS) in vivo (Jabaudon et al., 1999; Warr et al., 1999; Baker et al., 2002a, 2002b; Melendez et al., 2005; Augustin et al., 2007; Featherstone and Shippy, 2008; De Bundel et al., 2011; Massie et al., 2011) with maximal transporter activity estimated to theoretically increase extracellular glutamate by 0.6 µM/s (Warr et al., 1999; Cavelier et al., 2005). Conversely, nearly 90% of glutamate is removed by astrocytic Excitatory Amino Acid Transporters 1 (EAAT1) and 2 (EAAT2; Rothstein et al., 1996; Anderson and Swanson, 2000). Thus, excitotoxic processes subsequent to GD in our system could result via the enhancement of astrocytic glutamate release from system x_c⁻, by a reduction in astrocyte cellular reuptake or by a combination of both. Hence, the overall purpose of this study was to investigate whether changes in system x_c⁻ expression or alterations in glutamate handling following GD occur(s) and contributes to aglycemic neuronal cell death.

Materials and Methods

Animals

This study was conducted in accordance with the National Institutes of Health guidelines for the use of experimental animals and was approved by the Institutional Animal Care and Use Committee at both The University of Connecticut Health Center and Syracuse University. Time-pregnant CD1 mice were obtained from Charles River Laboratories (Wilmington, MA).

Cell Culture

Media stock (MS): L-glutamine-free modified Eagle’s medium (Earl’s salt; MediaTech, Herndon, VA) supplemented with L-glutamine, glucose, and sodium bicarbonate to a final concentration of 2.0, 25.7, and 28.2 mM, respectively; Glial plating media: MS containing 10% heat-inactivated fetal bovine serum (Hyclone, Logan, UT) and 10% heat-inactivated calf serum (CS; Hyclone, Logan, UT), 10 ng/ml epidermal growth factor (Invitrogen, Carlsbad, CA), and 50 IU penicillin/50 µg/ml streptomycin (Gibco®, Life Technologies, Grand Island, NY); Glial/mixed culture maintenance media: MS containing 10% CS and 50 IU penicillin/50 µg/ml streptomycin; Neuronal plating media: MS containing 5% CS and 5% heat-inactivated bovine growth serum (Hyclone, Logan, UT) and 50 IU penicillin/50 µg/ml streptomycin (Gibco®, Life Technologies, Grand Island, NY); Neuronal maintenance media: neurobasal medium (Gibco®, Life Technologies, Grand Island, NY) containing 2 mM L-glutamine, 1× B27 supplement (Gibco®, Life Technologies, Grand Island, NY), and 50 IU penicillin/50 µg/ml streptomycin (Gibco®, Life Technologies, Grand Island, NY); HEPES-controlled salt solution (HCSS): 120 mM NaCl, 5.4 mM KCl, 0.8 mM MgCl₂, 1.8 CaCl₂, 15 mM glucose, 20 mM HEPES, 10 mM NaOH, 10 µM glycine, and 0.001% phenol red; Glucose-free balanced salt solution (BSS₀): 116 mM NaCl, 5.4 mM KCl, 0.8 mM MgCl₂, 1 mM NaH₂PO₄, 26.2 mM NaHCO₃, 1.8 mM CaCl₂, 0.01 mM glycine, and 2 mM L-glutamine. Unless otherwise indicated, BSS contained 1 × MEM amino acids (Life Technologies, Grand Island, NY).

Primary astrocytes were cultured from cortices of postnatal Day 1 to 3 CD1 mouse pups as described previously (Trackey et al., 2001; Uliasz et al., 2012). Cortices were dissected, pooled, and cells dissociated by trypsin digestion (0.025%, 15 min, 37℃). Cells were plated 400 µl/well in glial plating medium (two hemispheres/10 ml/plate; Falcon Primaria 24-well plates; BD Biosciences, Lincoln Park, NJ). Once confluent, astrocyte monolayers were treated with 8 µM β-D-cytosine arabinofuranoside (AraC) once for 4 to 7 days to reduce the number of microglia. Cells were then placed in maintenance media (vide supra), which was replaced once per week until experimentation. Purified astrocyte cultures were generated by removing residual microglia by treating monolayers with 50 to 75 mM L-leucine methyl ester for 30 to 90 min, 1 day prior to experimentation (Hamby et al., 2006; Uliasz et al., 2012). Cultures were used for experimentation at ≤35 days in vitro.

Primary neuronal cultures were derived from dissociated cortical cells of embryonic Day 15 CD1 mouse fetuses. Following dissection and dissociation by trypsin digestion (vide supra), cells were diluted to a concentration of 10⁶ cells/ml of neuronal plating medium and then plated into polyethyleneimine coated 24-well plates (Costar®, Corning, NY). Four hr later, the medium was exchanged to neuronal maintenance medium. Two days later, cultures were treated with 1 µM AraC once for 2 days to prevent glial cell growth. The medium was partially replenished (1/2 volume exchange) at DIV 4. Experiments were performed on purified neuronal cultures after 6 days in vitro.

Mixed cortical cell cultures containing an approximate 50:50 neuron-to-astrocyte ratio were prepared by culturing dissociated cells from embryonic Day 15 CD1 mouse fetuses on to a confluent layer of microglia-depleted astrocytes in neuronal plating media. The plating medium was partially replaced (2/3 exchange) with maintenance medium at Days 5 and 9 in vitro. At Day 7, cultures were treated with 8 µM of AraC once to prevent microglial cell growth. Two days prior to experimentation, mixed cortical cell cultures were placed into MS (vide supra). Experiments were performed at 14 days in vitro. All cultures were maintained at 37℃ in a humidified 6.0% CO₂, 21% O₂-containing incubator.

Glucose Deprivation

Mixed cortical cultures and astrocyte cultures were deprived of glucose by thorough washing with and into BSS₀. Glucose (final concentration = 10 mM) was immediately added to parallel cultures to serve as controls and added back to experimental conditions as indicated in each figure legend. In the MK-801 (R&D System, Rockford, IL) or LY367385 (R&D System, Rockford, IL) experiments, MK-801 was made as a 10 mM stock solution in H₂O, and LY367385 was made as a 50 mM stock solution in 0.1 NaOH. The drugs or their corresponding vehicle were either given at the initiation of GD or spiked into desire groups at the times indicated in each figure legend.

Measurement of Neuronal Cell Death

Cell death was quantitatively determined by the spectrophotometric measurement of lactate dehydrogenase (LDH) found in the cell culture medium as described in detail previously (Uliasz and Hewett, 2000). Data are expressed as a percentage of total neuronal LDH activity (defined as 100% cell death) determined by exposing parallel cultures to 200 µM of N-methyl-D-aspartate (NMDA) for 20 to 24 hr. As cultured astrocytes do not express NMDA receptors (Backus et al., 1989; Chan et al., 1990; B. Fogal and S.J. Hewett, unpublished data) and have previously been shown to survive up to 8 hr of GD (Jackman et al., 2012), LDH measurements can be used as a specific measure of neuron cell death.

Quantitative Real-Time Polymerase Chain Reaction

Total RNA was isolated and first-strand cDNA synthesized as previously described (Uliasz et al., 2012). Quantitative real-time polymerase chain reaction (qPCR) was performed using mouse-specific primer pairs (Taqman Gene Expression Assays, Applied Biosystems: xCT [Mm00442530_m1] and β-actin [Mm01205647_g1]) per manufacturer’s instructions. Reactions were run in the Eppendorf Real-Time PCR System and relative quantification performed using the comparative cycle threshold method (ΔΔC_T), where C_T values of xCT were normalized to β-actin C_T values from the same sample and then compared with a calibrator sample C_T value (control group, cultures kept in BSS₁₀) to determine the relative fold increase in mRNA. β-actin C_T values are unaffected by GD.

Radiolabeled L-Cystine and D-Aspartate Uptake

System x_c⁻-specific ¹⁴C-L-cystine (PerkinElmer; Waltham, MA) and system X_AG⁻-mediated ³H-D-aspartate (PerkinElmer; Waltham, MA) uptake was performed as previously described (Fogal et al., 2007). Cultures were washed into HCSS (3 × 750 µl) and allowed to equilibrate for 10 min (25℃). For cystine uptake, cells were incubated in HCSS containing 3 µM ¹⁴C-L-cystine (1 µCi/ml), 27 µM unlabeled cystine, 1 mM D-aspartate, and 0.5 mM acivicin (Enzo Life Sciences; Farmingdale, NY). D-aspartate and acivicin were included in the uptake buffer to block system X_AG⁻ and γ-glutamyltranspeptidase, respectively. Uptake was terminated after 30 min by washing in ice-cold PBS (3 × 750 µl). For D-aspartate uptake, cells were incubated in HCSS containing 0.1 µCi/ml ³H-D-aspartate and 50 µM unlabeled D-aspartate (25℃) for 5 min and uptake terminated by washing cells with an ice-cold Na⁺-free choline stop buffer containing in mM: 116 choline chloride, 0.8 MgSO₄, 1 KH₂PO₄, 10 HEPES, 5 KOH, 10 glucose, 0.9 CaCl₂, and 5 nonradioactive D-aspartate.

Cells were lysed with warm 0.5% SDS and accumulated radioactivity estimated using a liquid scintillation counter. Readings of counts per minute from experimental conditions were corrected back to the original volume of lysate, and the picomoles of cystine and aspartate transported per minute were calculated as described (Fogal et al., 2007).

Toxicity Bioassay

Purified astrocyte cultures were washed thoroughly into BSS containing or lacking 10 mM glucose. Sixty, 75, 90, or 105 min before the end of a 6-hr incubation period, glutamate—75 µM final well concentration—was spiked into the wells after which the supernatant was collected and transferred via 3/4 exchange to highly enriched murine neuronal cultures. Six hr later, neuronal cell death was quantified by measurement of the LDH released into the cell culture medium. Data are expressed as a percentage of total neuronal cell death facilitated by adding 200 µM glutamate to parallel neuronal cultures 1 day prior to experimentation. We demonstrated previously that the toxicity of the supernatant correlated with the glutamate-buffering capacity of astrocytes (Sen et al., 2011). Of note, glucose, to a final concentration of 10 mM, was added to all pooled media prior to its addition to neurons to prevent any confound associated with GD.

Measurement of Glutamate

Two hundred microliters of supernatant harvested from the toxicity bioassay experiment (vide supra) were analyzed for glutamate concentration in the media via phenylisothiocyanate derivatization, high-performance liquid chromatography (HPLC) separation using a Hypersil-ODS reverse-phase column, and ultraviolet detection at 254 nm as described in (Fogal et al., 2007).

Measurement of Adenosine Triphosphate

Adenosine triphosphate (ATP) levels were determined using the ATP Determination Kit (Molecular Probe™, Invitrogen; Eugene, OR) per manufacturer’s instruction. Culture wells were aspirated dry and the plate quick frozen on dry ice. After 5 min, plates were incubated at 37℃ for 10 min. One milliliter or 250 μl of a 1 × Cell Culture Lysis Reagent (Promega; Madison, WI) was added into each well of a 6-well or 24-well plate, respectively. Plates were gently shaken for 5 min. Cell lysates were collected, and intracellular ATP levels were measured and normalized to protein levels (BCA Assay; Pierce™, Pierce Biotechnology, Rockford, IL). When necessary, samples were diluted so that values fell within the linear range of the assay (0–1 µM).

Excitotoxicity Assays

Mixed cortical cultures were deprived of glucose or incubated with glucose for the times indicated before being washed and exposed to either 25 µM NMDA or 100 µM glutamate in HCSS at room temperature. After 10 min, the exposure solution was washed away and replaced by MS supplemented with 10 µM glycine. The cells were transferred to a 37℃, normoxic (21%), 6% CO₂ containing incubator overnight. Neuronal cell death was determined by the LDH assay, and data were expressed as a percentage of total neuronal LDH activity obtained by exposing parallel cultures to 200 µM of NMDA for 20 to 24 hr (defined as 100% neuronal cell death).

Statistical Analysis

All statistical analyses were performed using GraphPad Prism (Version 6.0.1, GraphPad Software, Inc.) as described in each figure legend. As percentage data and normalized data are by nature nonnormally distributed, such data were first transformed via arcsin square root or −1 × log (Y), respectively, before analysis. Values of zero or less were set at 1 × 10⁻²⁰ before transformation. In the case of qPCR, statistics were performed on the geometric means of 2^−ΔΔCT values. In all experiments, data were expressed as mean + SEM. Significance was assessed at p < .05.

Results

In confirmation of prior literature from our own laboratory as well as that from others, we demonstrate that removal of glucose from the cell culture medium of mixed neuronal/astrocyte cortical cell cultures results in a time-dependent (Figure 1(a)) and NMDA receptor-mediated excitotoxic neuronal cell death (Figure 1(b); Monyer and Choi, 1988; Monyer et al., 1989; Jackman et al., 2012). That system x_c⁻ activity contributes to death is illustrated by the fact that removal of amino acids (Figure 2(a))—specifically cystine (Jackman et al., 2012)—from the glucose-free medium reduced aglycemic neuronal injury, whereas addition of cystine back to the medium lacking amino acids results in significant neuronal cell death (Figure 2(a)). Furthermore, addition of the dual mGluR1/system x_c⁻ antagonist LY367385 ameliorates aglycemic neuronal cell death, while the mGluR1-specific antagonist, YM 298198, shows no protection (Figure 2(b); Jackman et al., 2012). While neurons and astrocytes both express functional system x_c⁻ in our culture system (Jackman et al., 2010), we recently described astrocytic system x_c⁻ as the main source of glutamate required for the initiation of neuronal injury following GD in vitro (Jackman et al., 2012).

Figure 1.

Aglycemic neuronal cell death is time-dependent and is blocked by NMDA receptor antagonism. (a) Mixed cortical cultures were washed into BSS containing (0 hr) or lacking 10 mM glucose for the times indicated, after which neuronal cell death was determined via measurement of LDH release. An asterisk (*) represents values significantly different from control (0 hr) as assessed by one-way ANOVA followed by Dunnett’s post hoc test (n = 8–10 cultures from three separate experiments). (b) Mixed cortical cultures were washed into BSS lacking or containing 10 mM glucose (0 hr) ± MK-801 (10 µM). Neuronal cell death was determined via measurement of LDH release. An asterisk (*) denotes a significant within-group difference, whereas a pound sign (#) denotes a significant between-group difference as determined by two-way ANOVA followed by Bonferroni’s post hoc test for multiple comparisons (n = 6 cultures from three separate experiments).

Figure 2.

System x_c⁻ contributes to aglycemic neuronal cell death. (a) Mixed cortical cultures washed into BSS₀ containing (GD) or lacking (-AA) MEM amino acids save for supplementation with 100 µM L-cystine (+ Cyss). (b) Mixed cultures were deprived of glucose (GD) in the absence or presence of the dual system x_c⁻/mGluR1 inhibitor, LY367385 (LY; 50 µM) or the selective mGluR1 antagonist, YM298190 (YM; 10 µM). Neuronal cell death was determined 8 hr later via measurement of LDH release. An asterisk (*) equals values different from control (cultures in BSS₁₀), while a pound sign (#) represents a significant diminution from GD-induced injury as determined by one-way ANOVA followed by Bonferroni’s post hoc test for multiple comparisons (n = 10–11 from three independent experiments).

Cellular stress results in transcriptional adaptive increases in the substrate-specific light chain of system x_c⁻, xCT, and a subsequent increase in cystine uptake (Bannai, 1984a; Ishii et al., 1992; Miura et al., 1992; Ishii et al., 2000; Sasaki et al., 2002; Sato et al., 2004). Hence, whether xCT mRNA levels was altered following aglycemia in pure astrocyte cultures or in mixed cortical cultures containing both astrocytes and neurons was ascertained using quantitative PCR analysis. In both astrocyte and mixed cortical cell cultures, a significant increase in xCT mRNA occurs following 8 hr of GD but not earlier (Figure 3(a) and (b)). Because significant and nearly maximal cell death occurs by this time point (Figure 1), we find it unlikely that a change in system x_c⁻ expression underlies its neurotoxic effect. In keeping with this, actinomycin D (10 µg/ml), when given at the onset of GD, fails to prevent aglycemic neuronal cell death (Figure 4(a)), demonstrating no requirement for transcription. Death is similarly unaffected by cycloheximide (1 µg/ml), indicating no need for protein synthesis (Figure 4(b)).

Figure 3.

Aglycemia regulates xCT mRNA levels in a time-dependent manner. (a) Purified astrocyte cultures or (b) mixed cortical cultures containing both astrocytes and neurons were deprived of glucose for the times indicated and changes in xCT mRNA assessed via qPCR. An asterisks (*) represent values that are significantly different from 0 hr (i.e., glucose-containing control) as determined by one-way ANOVA followed by a Dunnett’s post hoc test for multiple comparison (n = 4 from four separate experiments each).

Figure 4.

Macromolecular synthesis inhibitors do not prevent aglycemic neuronal cell death. Mixed cortical cultures were rendered aglycemic in the presence or absence of (a) the transcriptional inhibitor actinomycin D (ACT D; 10 µg/ml; n = 6–10 from five separate experiments) or (b) the protein synthesis inhibitor cycloheximide (CHX; 1 µg/ml; n = 12 from three separate experiments). Six and 8 hr later, the amount of neuronal cell death was assessed. (a, b) Two-way ANOVA revealed no significant between-group differences in either paradigm.

To ascertain whether system x_c⁻ activity was enhanced by GD, we measured ¹⁴C-L-cystine uptake in the mixed cortical cell cultures. No change in activity is noted over the time-course of GD studied (Figure 5(a)), suggesting increased glutamate extrusion via system x_c⁻ does not underlie its neurotoxic effect. As maintenance of extracellular glutamate concentrations involves control of release as well as uptake, we next investigated whether aglycemic-mediated alterations in glutamate removal occurred. We find no significant difference in ³H-D-aspartate uptake—a measure of EAAT activity as verified by the ability of D,L-threo-β-benzyloxyaspartate to completely suppress the uptake—between control mixed cultures and those that were deprived of glucose (Figure 5(b)). Also, glucose-deprived astrocytes retain the ability to remove exogenously added glutamate just as effectively as control astrocytes (Figure 6(a)). Finally, glutamate-containing media removed from glucose-deprived astrocytes is equally as toxic to pure neuronal cultures as that from glucose-containing medium at every time point assessed (Figure 6(b)). Therefore, overstimulation of neuronal excitatory amino receptors via enhancement of the extracellular glutamate concentration cannot account for the excitotoxic insult mediated by system x_c⁻ under aglycemic conditions.

Figure 5.

Aglycemia does not significantly alter cystine or aspartate uptake. (a) Mixed cortical cultures were deprived of glucose for the times indicated, then incubated with buffer containing ¹⁴C-L-cystine for 30 min to measure system x_c⁻ activity. Data are expressed as mean + SEM ¹⁴C-L-cystine uptake in pmol/min/mg protein. Aglycemia had no significant effect on radiolabeled cystine uptake as determined by one-way ANOVA followed by a Dunnett’s post hoc test (n = 6 from two separate experiments). (b) Mixed cortical cultures were deprived of glucose for the times indicated, then ³H-D-aspartate was added for 5 min to measure EAAT activity. D,L-threo-β-benzyloxyaspartate (500 µM) was used to confirm uptake via EAATs (n = 6 from five separate experiments). Data are expressed as mean + SEM ³H-D-aspartate uptake in pmol/min/mg protein. (a, b) ANOVA revealed no significant within-group differences (compared with 0 hr) in each paradigm.

Figure 6.

Aglycemic astrocytes are still capable of glutamate uptake. Purified astrocyte cultures were glucose deprived (− Glucose) or washed into BSS containing glucose (+Glucose; 10 mM) for 6 hr before glutamate (final concentration = 75 µM) was added to cultures. At the times indicated after addition, bathing medium was removed and like groups pooled. (a) HPLC analysis was used to measure remaining glutamate in an aliquot of the pooled media. Data are expressed as percentage of initial glutamate concentration (75 µM; n = 3–6 from three separate experiments). (b) The pooled medium was then added to pure neuronal cultures (2/3 exchange). Six hr later, neuronal cell death was assessed. (n = 10 from five separate experiments). (a, b) No significant between-group differences were found using two-way ANOVA.

Consequently, we set out to test whether the weak excitotoxic hypothesis (Albin and Greenamyre, 1992)—which posits that bioenergetic deficiencies enhance the toxic effect of glutamate—might explain the results described earlier. Following GD in vitro, rapid ATP depletion has been reported to occur in neurons with astrocytes being more resistant (Choi et al., 2008). Likewise, we find that aglycemic astrocytes maintain cellular ATP levels (Figure 7(a)), whereas the concentration of ATP in mixed cortical cultures is significantly and severely depressed when glucose was removed from the media (Figure 7(b)). These changes are not simply due to neuronal loss from the mixed cultures because the addition of MK-801 and LY367385, which protects the neurons from aglycemia-induced death (Figures 1(b) and 2(b)), does not rescue the culture ATP levels (Figure 7(c)).

Figure 7.

Aglycemia decreases ATP levels in cortical neurons but not astrocytes. (a) Purified cortical astrocyte cultures (n = 7–8 from two independent experiments) or (b) mixed cortical cultures (n = 3–4 from one experiment) were placed in medium containing glucose for 6 hr (0) or deprived of glucose for the times indicated, after which ATP levels were measured as described in methods. (c) Mixed cortical cultures were washed into BSS lacking (− Glucose, black bars) or containing 10 mM glucose (+Glucose, white bars) ± MK-801 (MK; 10 µM) or LY367385 (LY; 50 µM). Six hr later, ATP levels were measured (n = 6 from two independent experiments). Data are expressed as mean + SEM. Individual values are normalized to each well’s protein levels. (a, b) An asterisk (*) indicates a significant difference from 0 hr as determined by one-way ANOVA followed by a Dunnett’s post hoc test for multiple comparison. (c) An asterisk indicates a significant between-group difference as determined by two-way ANOVA followed by Bonferroni’s post hoc test for multiple comparisons.

To investigate whether aglycemic cortical neurons would show an enhanced susceptibility to excitotoxic insult, we deprived mixed cultures of glucose for increasing periods of time (2.5 to 4.5 hr) after which glutamate (100 µM final well concentration) was added for 10 min. Compared with cells maintained in glucose-containing medium, results show that glutamate kills significantly more neurons when the cultures are glucose deprived at every time point tested (Figure 8(a)). Likewise, neurons in glucose-deprived cultures are more susceptible to death than control cultures when exposed to equimolar concentrations of NMDA (Figure 8(b)).

Figure 8.

Glucose-deprived mixed cultures are more susceptible to the toxic effects of glutamate and NMDA. Mixed cortical cultures (a) were deprived of glucose for the times indicated followed by exposure to 100 µM glutamate (n = 6 from three separate experiments) or (b) were deprived of glucose for 2.25 hr followed by exposure to increasing concentrations of NMDA (n = 6 from three separate experiments). Ten min later, cultures were washed free of glutamate or NMDA and placed back into a glucose-containing medium. The percentage of neuronal cell death was assessed 20 to 24 hr later by measurement of LDH. An asterisk (*) indicates a significant between-group difference assessed by two-way ANOVA followed by a Bonferroni’s post hoc test for multiple comparison.

Finally, to study the optimal therapeutic time window for rescue, mixed cortical cultures were deprived of glucose for a total of 8 hr and either MK-801, to block NMDA receptors (Figure 9(a)), or LY367385 (Figure 9(b)), to mitigate glutamate release from system x_c⁻, was added to parallel cultures at 1- to 2-hr intervals postexperimental onset. The percentage of the dying neuronal population that could be rescued depended on the time interval between washout of glucose and addition of the pharmacological agents. When compared with 8 hr of GD alone, MK-801 is able to completely block aglycemic neuronal injury when given up to 3 hr after the start of GD, and it is still significantly effective when addition is delayed for 5 hr (Figure 9(a)). In contrast, maximal protection for LY367385 occurs at 1 hr, though significant protection is also demonstrated up to 5 hr (Figure 9(b)). No protection is observed with either compound if its administration is delayed by 7 hr (Figure 9(a) and (b)).

Figure 9.

Delayed pharmacological rescue of aglycemic neuronal injury in mixed cortical cultures. Mixed cortical cultures were deprived of glucose for a total 8 hr (GD). (a) MK-801 (10 µM) or (b) LY367385 (50 µM) were added at the times indicated after the initiation of GD. The percentage of neuronal cell death was assessed via the LDH assay. An asterisks (*) indicates a significant difference from GD, and a pound sign (#) indicates a significant difference from 1 hr determined by one-way ANOVA followed by Dunnett’s post hoc test for multiple comparison (n = 6 from two experiments).

Discussion

Symptomatic hypoglycemia, resulting in brain dysfunction ranging from mild behavioral impairments to coma (Wilson, 1983), is the principal problem associated with tight glucose regulation in patients undergoing insulin therapy (Amiel et al., 1987; Group, 1991; Cryer, 2001; Griesdale et al., 2009). The idea that the underlying cause of severe neurological complications that follow a dangerous drop in blood glucose levels is not directly due to substrate deprivation-induced energy failure has been recognized since the early 1970s (for review, see Auer, 1986). Instead, the neuronal cell death that ensues occurs via excitotoxicity (Wieloch, 1985; Monyer and Choi, 1988; Monyer et al., 1989; Papagapiou and Auer, 1990; Nellgard and Wieloch, 1992; Tasker et al., 1992; Ichord et al., 2001; Jackman et al., 2012), the process of which is most commonly ascribed to overexcitation of neuronal glutamate receptors (Bo et al., 1994). While multiple sources of glutamate could contribute to aglycemic neuron death (Danbolt, 2001), in vivo and in vitro studies indicate a contributory role for neuronal exocytotic release of glutamate (Wieloch et al., 1985; Monyer et al., 1989; Jackman et al., 2012) as well as an obligate requirement for astrocytic system x_c⁻ (Jackman et al., 2012). The mechanism by which astrocytic system x_c⁻ contributes to aglycemic neuronal injury has been explored in this study.

Using a mixed cortical cell culture system in vitro, we demonstrated previously that enhancement of astrocyte system x_c⁻ (cystine/glutamate antiporter) expression and activity facilitated by interleukin-1β contributed to the development and progression of hypoxic neuronal injury—a model of the ischemic penumbra (Fogal et al., 2007; Jackman et al., 2010). While the enhanced extrusion of glutamate from astrocytic system x_c⁻ activity was not deleterious in and of itself, we found that under hypoxic conditions, astrocyte glutamate uptake was also compromised, thus leading to glutamate buildup (Fogal et al., 2007). Thus, we speculated that similar mechanisms might be in play in our hypoglycemia model described herein.

With respect to upregulation of system x_c⁻, we did find that xCT mRNA expression in astrocytes and mixed cultures was increased in a time-dependent manner following GD. However, in agreement with our previous study (Jackman et al., 2012), a statistically significant increase did not occur in either culture system until 8 hr following removal of glucose from the medium (Figure 3). Given that much, if not all, of the death has already occurred by this time point, it seems unlikely that a change in system x_c⁻ expression is necessary to mediate the deleterious effects of system x_c⁻ under aglycemic conditions. Indeed, actinomycin D or cycloheximide—used at concentrations shown previously by us to effectively decrease the transcriptional (10 µg/ml) and translational (1 µg/ml) increases in astrocyte xCT facilitated by interleukin-1β, respectively (Jackman et al., 2010)—failed to ameliorate aglycemic neuronal injury (Figure 4). However, because system x_c⁻ transport can be enhanced via phosphorylation of putative protein kinase A sites in xCT (Baker et al., 2002b) and changes in protein trafficking could lead to changes in xCT protein insertion in the membrane (Chase et al., 2013; Cramer and Chase, 2013; Ladd et al., 2014), we also investigated the activity of system x_c⁻ activity using a radiolabeled cystine uptake assay. Our results directly demonstrate that no change in system x_c⁻ activity occurs following GD (Figure 5(a)).

Following release, the concentration of glutamate in the extracellular space is carefully maintained at subtoxic levels by its rapid reuptake. We initially surmised that glucose-deprived astrocytes might show a diminution in their ability to remove glutamate from the extracellular space, thus leading to a toxic buildup. However, using three different assays—³H-D-asparte uptake (Figure 5(b)), direct measurement of residual glutamate levels in astrocyte media via HPLC following exogenous addition of glutamate (Figure 6(a)), and a toxicity bioassay (Figure 6(b)) developed by our laboratory and validated for its ability to accurately reflect glutamate uptake into astrocytes (Sen et al., 2011)—we demonstrate no change in the ability of glucose-deprived astrocytes alone or astrocytes contained in mixed culture to remove glutamate from the extracellular environment. While this might initially seem surprising, others have shown that there were no changes in astrocytic glutamate uptake in astrocytes for up to 2 hr of GD (Bakken et al., 1998) and only a 20% loss after 24 hr (Swanson and Benington, 1996). Indeed, unlike neurons (Auer et al., 1984; Monyer and Choi, 1988; Monyer et al., 1989; Goldberg and Choi, 1993), astrocytes have been demonstrated to be more resistant (Monyer and Choi, 1988; Monyer et al., 1989; Goldberg and Choi, 1993; Lyons and Kettenmann, 1998) to the ATP-depleting effects of glucose starvation by virtue of their ability to use glycogen stores (Cataldo and Broadwell, 1986; Swanson et al., 1990) that can be metabolized to meet their metabolic needs (Swanson et al., 1990; Erecinska and Silver, 1994; Dienel and Cruz, 2006; Walls et al., 2009). In addition, astrocytes can metabolize glutamate via the TCA cycle to provide energy (Yu et al., 1982; Yu et al., 1984; McKenna et al., 1996; McKenna, 2007), especially when glucose levels are low (Bakken et al., 1998). Indeed, we found that astrocytes are able to fully maintain their ATP levels under the conditions of GD used in this study (Figure 7(a)). Thus, the loss in mixed culture is then likely due to changes in neuronal ATP production (Figure 7(b)). These results are in good agreement with what has been reported previously (Choi et al., 2008).

It is important to point out that neither enhanced release of glutamate nor its diminished uptake may be needed to facilitate excitotoxic neuronal cell death under aglycemic conditions, as Novelli et al. demonstrated nearly 20 years ago that glutamate concentrations needed to kill energy-deprived cerebellar neurons are far less than those required to kill healthy neurons (Novelli et al., 1988; Henneberry et al., 1989). These observations, among others, led to the development of the weak excitotoxin hypothesis (Albin and Greenamyre, 1992), which postulates that a reduction in cellular ATP leads to an impairment in Na⁺/K⁺ ATPase activity resulting in a slight membrane depolarization (Calabresi et al., 1997) that ultimately allows for voltage-dependent NMDA receptors to be more easily activated (Henneberry et al., 1989; Beal, 1992). Indeed, the reduction in neuronal ATP correlated with the toxicity of glutamate (Henneberry et al., 1989) and poisoning of the Na⁺ pump with ouabain replicated what was found when glucose was removed from the medium (Novelli et al., 1988). In keeping with this hypothesis, we found that glucose-deprived cultures showed a two- to threefold enhancement in neuronal cell death when exposed to 100µM glutamate at nearly every time point of GD analyzed when compared with their glucose-containing controls (Figure 8(a)). Similar results were seen when NMDA was used (Figure 8(b)). As one might expect, a longer therapeutic window of opportunity, at least when it comes to maximal or essentially full protection, was achieved by blocking NMDA receptors when compared with blocking system x_c⁻ itself, although both drugs used showed significant protection up to a remarkable 5 hr following removal of glucose (Figure 9).

Finally, the fact that system x_c⁻ contributes to injury might seem paradoxical given its importance in the biosynthesis of the antioxidant molecule glutathione (Watanabe and Bannai, 1987; Bannai et al., 1989; Miura et al., 1992; Sato et al., 1995; Bridges et al., 2001; Dun et al., 2006; Lewerenz et al., 2009). Ironically, while this pathway allows the CNS to rapidly upregulate glutathione production in response to cellular stress, we and others demonstrate its potential to exacerbate CNS pathology both in vitro and in vivo (for reviews, see Bridges et al., 2012; Lewerenz et al., 2013; Massie et al., 2015). As such, it is becoming increasingly clear that the consequences of system x_c⁻ activity are complex and context-dependent. Hence, system x_c⁻ activity can either be beneficial (Tanaka et al., 1999; Shih et al., 2003; Jakel et al., 2007; He et al., 2015) or can contribute to pathophysiology (Piani and Fontana, 1994; Ye et al., 1999; Barger and Basile, 2001; Qin et al., 2006; Fogal et al., 2007; Savaskan et al., 2008; Sontheimer, 2008; Jackman et al., 2010; Massie et al., 2011; Jackman et al., 2012), the ultimate effect being dependent on the environmental and cellular milieu.

In sum, we find that aglycemic neuronal cell death does not result from changes in astrocyte system x_c⁻ expression or activity or from impairment of glutamate removal. Instead, our results are consistent with the weak excitotoxic hypothesis (Albin and Greenamyre, 1992), suggesting that the bioenergetic deficiencies, which are present in our mixed cultures, enhance the toxic effect of physiological levels of glutamate released from astrocyte system x_c⁻. Whether system x_c⁻ plays a similar role in hypoglycemic neurodegeneration in vivo requires further experimentation.

Footnotes

Declaration of Conflicting Interests

The authors declared no potential conflicts of interest with respect to the research, authorship, and/or publication of this article.

Funding

The authors disclosed receipt of the following financial support for the research, authorship, and/or publication of this article: This work was supported by grants awarded to SJH and JAH by the National Institutes of Health (NINDS: NS051445). During her training, NAJ was supported by the National Institutes of Health in the form of a T32 training grant (NINDS: NS041224), an F31 NRSA (NINDS: NS066745), and an R36 dissertation award (NIA: AG035036).

References

Albin

R. L.

Greenamyre

J. T.

(1992) Alternative excitotoxic hypotheses. Neurology 42: 733–738.

Amiel

S. A.

Tamborlane

W. V.

Simonson

D. C.

Sherwin

R. S.

(1987) Defective glucose counterregulation after strict glycemic control of insulin-dependent diabetes mellitus. The New England Journal of Medicine 316: 1376–1383.

Anderson

C. M.

Swanson

R. A.

(2000) Astrocyte glutamate transport: Review of properties, regulation, and physiological functions. Glia 32: 1–14.

Anderson

J. M.

Milner

R. D.

Strich

S. J.

(1967) Effects of neonatal hypoglycaemia on the nervous system: A pathological study. Journal of Neurology, Neurosurgery & Psychiatry 30: 295–310.

Auer

R. N.

(1986) Progress review: Hypoglycemic brain damage. Stroke; A Journal of Cerebral Circulation 17: 699–708.

Auer

R. N.

(2004) Hypoglycemic brain damage. Metabolic Brain Disease 19: 169–175.

Auer

R. N.

Wieloch

Olsson

Siesjo

B. K.

(1984) The distribution of hypoglycemic brain damage. Acta Neuropathologica 64: 177–191.

Augustin

Grosjean

Chen

Featherstone

D. E.

(2007) Nonvesicular release of glutamate by glial xCT transporters suppresses glutamate receptor clustering in vivo. Journal of Neuroscience 27: 111–123.

Backus

K. H.

Kettenmann

Schachner

(1989) Pharmacological characterization of the glutamate receptor in cultured astrocytes. Journal of Neuroscience Research 22: 274–282.

10.

Baker

D. A.

Shen

Kalivas

P. W.

(2002a) Cystine/glutamate exchange serves as the source for extracellular glutamate: Modifications by repeated cocaine administration. Amino Acids 23: 161–162.

11.

Baker

D. A.

Z. X.

Shen

Swanson

C. J.

Kalivas

P. W.

(2002b) The origin and neuronal function of in vivo nonsynaptic glutamate. The Journal of Neuroscience: The Official Journal of the Society for Neuroscience 22: 9134–9141.

12.

Bakken

I. J.

White

L. R.

Unsgard

Aasly

Sonnewald

(1998) [U-13C]glutamate metabolism in astrocytes during hypoglycemia and hypoxia. Journal of Neuroscience Research 51: 636–645.

13.

Bannai

(1984a) Induction of cystine and glutamate transport activity in human fibroblasts by diethyl maleate and other electrophilic agents. The Journal of Biological Chemistry 259: 2435–2440.

14.

Bannai

(1984b) Transport of cystine and cysteine in mammalian cells. Biochimica et Biophysica Acta 779: 289–306.

15.

Bannai

Sato

Ishii

Sugita

(1989) Induction of cystine transport activity in human fibroblasts by oxygen. Journal of Biological Chemistry 264: 18480–18484.

16.

Barger

S. W.

Basile

A. S.

(2001) Activation of microglia by secreted amyloid precursor protein evokes release of glutamate by cystine exchange and attenuates synaptic function. Journal of Neurochemistry 76: 846–854.

17.

Beal

M. F.

(1992) Mechanisms of excitotoxicity in neurologic diseases. FASEB Journal 6: 3338–3344.

18.

Dawson

T. M.

Wesselingh

Mork

Choi

Kong

P. A.

Trapp

B. D.

(1994) Induction of nitric oxide synthase in demyelinating regions of multiple sclerosis brains. Annals of Neurology 36: 778–786.

19.

Bridges

Kekuda

Wang

Prasad

Mehta

Huang

Ganapathy

(2001) Structure, function, and regulation of human cystine/glutamate transporter in retinal pigment epithelial cells. Investigative Ophthalmology & Visual Science 42: 47–54.

20.

Bridges

Lutgen

Lobner

Baker

D. A.

(2012) Thinking outside the cleft to understand synaptic activity: Contribution of the cystine-glutamate antiporter (system x_c⁻) to normal and pathological glutamatergic signaling. Pharmacological Reviews 64: 780–802.

21.

Calabresi

Ascone

C. M.

Centonze

Pisani

Sancesario

D’Angelo

Bernardi

(1997) Opposite membrane potential changes induced by glucose deprivation in striatal spiny neurons and in large aspiny interneurons. The Journal of Neuroscience: The Official Journal of the Society for Neuroscience 17: 1940–1949.

22.

Cataldo

A. M.

Broadwell

R. D.

(1986) Cytochemical identification of cerebral glycogen and glucose-6-phosphatase activity under normal and experimental conditions. I. Neurons and glia. Journal of Electron Microscopy Technique 3: 413–437.

23.

Cavelier

Hamann

Rossi

Mobbs

Attwell

(2005) Tonic excitation and inhibition of neurons: Ambient transmitter sources and computational consequences. Progress in Biophysics & Molecular Biology 87: 3–16.

24.

Chan

P. H.

Chu

Chen

(1990) Effects of MK-801 on glutamate-induced swelling of astrocytes in primary cell culture. Journal of Neuroscience Research 25: 87–93.

25.

Chase

Smith

Natasha

(2013) Cell surface expression of xCT is regulated by the Akt signaling pathway. The FASEB Journal 27: 1017–1010.

26.

Choi

I. Y.

Lee

S. P.

Kim

S. G.

Gruetter

(2001) In vivo measurements of brain glucose transport using the reversible Michaelis-Menten model and simultaneous measurements of cerebral blood flow changes during hypoglycemia. Journal of Cerebral Bood Flow and Metabolism: Official Journal of the International Society of Cerebral Blood Flow and Metabolism 21: 653–663.

27.

Choi

J. W.

Shin

C. Y.

Choi

M. S.

Yoon

S. Y.

Ryu

J. H.

Lee

J. C.

K. H.

(2008) Uridine protects cortical neurons from glucose deprivation-induced death: Possible role of uridine phosphorylase. Journal of Neurotrauma 25: 695–707.

28.

Cramer

Chase

(2013) Real time trafficking of system x_c⁻ in COS-7 cells. The FASEB Journal 27: 1017–1018.

29.

Cryer

P. E.

(2001) Hypoglycemia risk reduction in type 1 diabetes. Experimental and Clinical Endocrinology & Diabetes 109(Suppl 2): S412–S423.

30.

Danbolt

N. C.

(2001) Glutamate uptake. Progress in Neurobiology 65: 1–105.

31.

De Bundel

Schallier

Loyens

Fernando

Miyashita

Van Liefferinge

Massie

(2011) Loss of system x_c⁻ does not induce oxidative stress but decreases extracellular glutamate in hippocampus and influences spatial working memory and limbic seizure susceptibility. Journal of Neuroscience 31: 5792–5803.

32.

de Herder

(2004) Insulinoma. Neuroendocrinology 80: 20–22.

33.

Dienel

G. A.

Cruz

N. F.

(2006) Astrocyte activation in working brain: Energy supplied by minor substrates. Neurochemistry International 48: 586–595.

34.

Dun

Mysona

Van Ells

Amarnath

Ola

M. S.

Ganapathy

Smith

S. B.

(2006) Expression of the cystine-glutamate exchanger (x_c⁻) in retinal ganglion cells and regulation by nitric oxide and oxidative stress. Cell and Tissue Research 324: 189–202.

35.

Engelsen

Westerberg

Fonnum

Wieloch

(1986) Effect of insulin-induced hypoglycemia on the concentrations of glutamate and related amino acids and energy metabolites in the intact and decorticated rat neostriatum. Journal of Neurochemistry 47: 1634–1641.

36.

Erecinska

Silver

I. A.

(1994) Ions and energy in mammalian brain. Progress in Neurobiology 43: 37–71.

37.

Featherstone

D. E.

Shippy

S. A.

(2008) Regulation of synaptic transmission by ambient extracellular glutamate. The Neuroscientist: A Review Journal Bringing Neurobiology, Neurology and Psychiatry 14: 171–181.

38.

Fogal

Lobner

McCullough

L. D.

Hewett

S. J.

(2007) System x(c)- activity and astrocytes are necessary for interleukin-1β-mediated hypoxic neuronal injury. The Journal of Neuroscience: The Official Journal of the Society for Neuroscience 27: 10094–10105.

39.

Goldberg

M. P.

Choi

D. W.

(1993) Combined oxygen and glucose deprivation in cortical cell culture: Calcium-dependent and calcium-independent mechanisms of neuronal injury. The Journal of Neuroscience: The Official Journal of the Society for Neuroscience 13: 3510–3524.

40.

Griesdale

D. E.

de Souza

R. J.

van Dam

R. M.

Heyland

D. K.

Cook

D. J.

Malhotra

Talmor

(2009) Intensive insulin therapy and mortality among critically ill patients: A meta-analysis including NICE-SUGAR study data. CMAJ: Canadian Medical Association Journal=Journal de l’Association Medicale Canadienne 180: 821–827.

41.

Group

T. D. R.

(1991) Epidemiology of severe hypoglycemia in the diabetes control and complications trial. The DCCT Research Group. The American Journal of Medicine 90: 450–459.

42.

Hamby

M. E.

Uliasz

T. F.

Hewett

S. J.

Hewett

J. A.

(2006) Characterization of an improved procedure for the removal of microglia from confluent monolayers of primary astrocytes. Journal of Neuroscience Methods 150: 128–137.

43.

Jackman

N. A.

Thorn

T. L.

Vought

V. E.

Hewett

S. J.

(2015) Interleukin-1beta protects astrocytes against oxidant-induced injury via an NF-kappaB-dependent upregulation of glutathione synthesis. Glia 63: 1568–1580.

44.

Henneberry

R. C.

Novelli

Cox

J. A.

Lysko

P. G.

(1989) Neurotoxicity at the N-methyl-D-aspartate receptor in energy-compromised neurons. An hypothesis for cell death in aging and disease. Annals of the New York Academy of Sciences 568: 225–233.

45.

Ichord

R. N.

Johnston

M. V.

Traystman

R. J.

(2001) MK801 decreases glutamate release and oxidative metabolism during hypoglycemic coma in piglets. Brain Research. Developmental Brain Research 128: 139–148.

46.

Ishii

Itoh

Takahashi

Sato

Yanagawa

Katoh

Yamamoto

(2000) Transcription factor Nrf2 coordinately regulates a group of oxidative stress-inducible genes in macrophages. The Journal of Biological Chemistry 275: 16023–16029.

47.

Ishii

Sato

Miura

Sagara

Bannai

(1992) Induction of cystine transport activity by stress. Annals of the New York Academy of Sciences 663: 497–498.

48.

Jabaudon

Shimamoto

Yasuda-Kamatani

Scanziani

Gahwiler

B. H.

Gerber

(1999) Inhibition of uptake unmasks rapid extracellular turnover of glutamate of nonvesicular origin. Proceedings of the National Academy of Sciences U S A 96: 8733–8738.

49.

Jackman

N. A.

Melchior

S. E.

Hewett

J. A.

Hewett

S. J.

(2012) Non-cell autonomous influence of the astrocyte system x_c⁻ on hypoglycaemic neuronal cell death. ASN Neuro 4: pii:e00074. doi:00010.01042/AN20110030.

50.

Jackman

N. A.

Uliasz

T. F.

Hewett

J. A.

Hewett

S. J.

(2010) Regulation of system x(c)(-)activity and expression in astrocytes by interleukin-1beta: Implications for hypoxic neuronal injury. Glia 58: 1806–1815.

51.

Jakel

R. J.

Townsend

J. A.

Kraft

A. D.

Johnson

J. A.

(2007) Nrf2-mediated protection against 6-hydroxydopamine. Brain Research 1144: 192–201.

52.

Lacherade

J. C.

Jacqueminet

Preiser

J. C.

(2009) An overview of hypoglycemia in the critically ill. Journal of Diabetes Science and Technology 3: 1242–1249.

53.

Ladd

Unterbrink

Georges

Lang

Chase

(2014) Identification of trafficking motifs in the C-terminus of the cystine/gluatamate exchanger, system x_c⁻ (997.3). The FASEB Journal 28(Suppl. 1).

54.

Lewerenz

Albrecht

Tien

M. L.

Henke

Karumbayaram

Kornblum

H. I.

Methner

(2009) Induction of Nrf2 and xCT are involved in the action of the neuroprotective antibiotic ceftriaxone in vitro. Journal of Neurochemistry 111: 332–343.

55.

Lewerenz

Hewett

S. J.

Huang

Lambros

Gout

P. W.

Kalivas

P. W.

Maher

(2013) The cystine/glutamate antiporter system x(c)(-) in health and disease: From molecular mechanisms to novel therapeutic opportunities. Antioxidants & Redox Signaling 18: 522–555.

56.

Lincoln

N. B.

Faleiro

R. M.

Kelly

Kirk

B. A.

Jeffcoate

W. J.

(1996) Effect of long-term glycemic control on cognitive function. Diabetes Care 19: 656–658.

57.

Linden

Kalimo

Wieloch

(1987) Protective effect of lesion to the glutamatergic cortico-striatal projections on the hypoglycemic nerve cell injury in rat striatum. Acta Neuropathologica 74: 335–344.

58.

Lyons

S. A.

Kettenmann

(1998) Oligodendrocytes and microglia are selectively vulnerable to combined hypoxia and hypoglycemia injury in vitro. Journal of Cerebral Blood Flow and Metabolism: Official Journal of the International Society of Cerebral Blood Flow and Metabolism 18: 521–530.

59.

Massie

Boillee

Hewett

Knackstedt

Lewerenz

(2015) Main path and byways: Non-vesicular glutamate release by system x_c⁻ as an important modifier of glutamatergic neurotransmission. Journal of Neurochemistry. doi: 10.1111/jnc.13348.

60.

Massie

Schallier

Kim

S. W.

Fernando

Kobayashi

Beck

Michotte

(2011) Dopaminergic neurons of system x(c)(-)-deficient mice are highly protected against 6-hydroxydopamine-induced toxicity. FASEB Journal 25: 1359–1369.

61.

McKenna

M. C.

(2007) The glutamate-glutamine cycle is not stoichiometric: Fates of glutamate in brain. Journal of Neuroscience Research 85: 3347–3358.

62.

McKenna

M. C.

Sonnewald

Huang

Stevenson

Zielke

H. R.

(1996) Exogenous glutamate concentration regulates the metabolic fate of glutamate in astrocytes. Journal of Neurochemistry 66: 386–393.

63.

Melendez

R. I.

Vuthiganon

Kalivas

P. W.

(2005) Regulation of extracellular glutamate in the prefrontal cortex: Focus on the cystine glutamate exchanger and group I metabotropic glutamate receptors. Journal of Pharmacology and Experimental Therapeutics 314: 139–147.

64.

Miura

Ishii

Sugita

Bannai

(1992) Cystine uptake and glutathione level in endothelial cells exposed to oxidative stress. American Journal of Physiology 262: C50–C58.

65.

Monyer

Choi

D. W.

(1988) Morphinans attenuate cortical neuronal injury induced by glucose deprivation in vitro. Brain Research 446: 144–148.

66.

Monyer

Goldberg

M. P.

Choi

D. W.

(1989) Glucose deprivation neuronal injury in cortical culture. Brain Research 483: 347–354.

67.

Nellgard

Wieloch

(1992) Cerebral protection by AMPA- and NMDA-receptor antagonists administered after severe insulin-induced hypoglycemia. Experimental Brain Research 92: 259–266.

68.

Novelli

Reilly

J. A.

Lysko

P. G.

Henneberry

R. C.

(1988) Glutamate becomes neurotoxic via the N-methyl-D-aspartate receptor when intracellular energy levels are reduced. Brain Research 451: 205–212.

69.

Papagapiou

M. P.

Auer

R. N.

(1990) Regional neuroprotective effects of the NMDA receptor antagonist MK-801 (dizocilpine) in hypoglycemic brain damage. Journal of Cerebral Blood Fflow and Metabolism: Official Journal of the International Society of Cerebral Blood Flow and Metabolism 10: 270–276.

70.

Piani

Fontana

(1994) Involvement of the cystine transport system x_c⁻ in the macrophage-induced glutamate-dependent cytotoxicity to neurons. Journal of Immunology 152: 3578–3585.

71.

Qin

Colin

Hinners

Gervais

Cheret

Mallat

(2006) System Xc⁻ and apolipoprotein E expressed by microglia have opposite effects on the neurotoxicity of amyloid-beta peptide 1–40. Journal of Neuroscience 26: 3345–3356.

72.

Rothstein

J. D.

Dykes-Hoberg

Pardo

C. A.

Bristol

L. A.

Jin

Kuncl

R. W.

Schielke

J. P.

(1996) Knockout of glutamate transporters reveals a major role for astroglial transport in excitotoxicity and clearance of glutamate. Neuron 16: 675–686.

73.

Sasaki

Sato

Kuriyama-Matsumura

Sato

Maebara

Wang

Bannai

(2002) Electrophile response element-mediated induction of the cystine/glutamate exchange transporter gene expression. The Journal of Biological Chemistry 277: 44765–44771.

74.

Sato

Fujiwara

Sagara

Bannai

(1995) Induction of cystine transport activity in mouse peritoneal macrophages by bacterial lipopolysaccharide. Biochemical Journal 310(Pt 2): 547–551.

75.

Sato

Nomura

Maebara

Sato

Tamba

Bannai

(2004) Transcriptional control of cystine/glutamate transporter gene by amino acid deprivation. Biochemical and Biophysical Research Communications 325: 109–116.

76.

Savaskan

N. E.

Heckel

Hahnen

Engelhorn

Doerfler

Ganslandt

Eyupoglu

I. Y.

(2008) Small interfering RNA-mediated xCT silencing in gliomas inhibits neurodegeneration and alleviates brain edema. Nature Medicine 14: 629–632.

77.

Sen

Basu

Willing

L. B.

Uliasz

T. F.

Myrkalo

J. L.

Vannucci

S. J.

Levison

S. W.

(2011) Pre-conditioning induces the precocious differentiation of neonatal astrocytes to enhance their neuroprotective properties. ASN Neuro 3: e00062.

78.

Shih

A. Y.

Johnson

D. A.

Wong

Kraft

A. D.

Jiang

Erb

Murphy

T. H.

(2003) Coordinate regulation of glutathione biosynthesis and release by Nrf-2 expressing glia potently protects neurons from oxidative stress. Journal of Neuroscience 23: 3394–3406.

79.

Sontheimer

(2008) A role for glutamate in growth and invasion of primary brain tumors. Journal of Neurochemistry 105: 287–295.

80.

Suh

S. W.

Aoyama

Chen

Garnier

Matsumori

Gum

Swanson

R. A.

(2003) Hypoglycemic neuronal death and cognitive impairment are prevented by poly(ADP-ribose) polymerase inhibitors administered after hypoglycemia. The Journal of Neuroscience: The Official Journal of the Society for Neuroscience 23: 10681–10690.

81.

Swanson

R. A.

Benington

J. H.

(1996) Astrocyte glucose metabolism under normal and pathological conditions in vitro. Developmental Neuroscience 18: 515–521.

82.

Swanson

R. A.

A. C.

Chan

P. H.

Sharp

F. R.

(1990) Glutamate increases glycogen content and reduces glucose utilization in primary astrocyte culture. Journal of Neurochemistry 54: 490–496.

83.

Tanaka

Toku

Zhang

Ishihara

Sakanaka

Maeda

(1999) Astrocytes protect neuronal death induced by reactive oxygen and nitrogen species. Glia 28: 85–96.

84.

Tasker

R. C.

Coyle

J. T.

Vornov

J. J.

(1992) The regional vulnerability to hypoglycemia-induced neurotoxicity in organotypic hippocampal culture: Protection by early tetrodotoxin or delayed MK-801. Journal of Neuroscience 12: 4298–4308.

85.

Trackey

J. L.

Uliasz

T. F.

Hewett

S. J.

(2001) SIN-1-induced cytotoxicity in mixed cortical cell culture: Peroxynitrite-dependent and -independent induction of excitotoxic cell death. Journal of Neurochemistry 79: 445–455.

86.

Uliasz

T. F.

Hamby

M. E.

Jackman

N. A.

Hewett

J. A.

Hewett

S. J.

(2012) Generation of primary astrocyte cultures devoid of contaminating microglia. Methods in Molecular Biology 814: 61–79.

87.

Uliasz

T. F.

Hewett

S. J.

(2000) A microtiter trypan blue absorbance assay for the quantitative determination of excitotoxic neuronal injury in cell culture. Journal of Neuroscience Methods 100: 157–163.

88.

Walls

A. B.

Heimburger

C. M.

Bouman

S. D.

Schousboe

Waagepetersen

H. S.

(2009) Robust glycogen shunt activity in astrocytes: Effects of glutamatergic and adrenergic agents. Neuroscience 158: 284–292.

89.

Warr

Takahashi

Attwell

(1999) Modulation of extracellular glutamate concentration in rat brain slices by cystine-glutamate exchange. The Journal of Physiology 514(Pt 3): 783–793.

90.

Watanabe

Bannai

(1987) Induction of cystine transport activity in mouse peritoneal macrophages. Journal of Experimental Medicine 165: 628–640.

91.

Wieloch

(1985) Hypoglycemia-induced neuronal damage prevented by an N-methyl-D-aspartate antagonist. Science 230: 681–683.

92.

Wieloch

Engelsen

Westerberg

Auer

(1985) Lesions of the glutamatergic cortico-striatal projections in the rat ameliorate hypoglycemic brain damage in the striatum. Neuroscience Letters 58: 25–30.

93.

Wilson

D. E.

(1983) Excessive insulin therapy: Biochemical effects and clinical repercussions. Current concepts of counterregulation in type I diabetes. Annals of Internal Medicine 98: 219–227.

94.

Z. C.

Rothstein

J. D.

Sontheimer

(1999) Compromised glutamate transport in human glioma cells: Reduction-mislocalization of sodium-dependent glutamate transporters and enhanced activity of cystine-glutamate exchange. Journal of Neuroscience 19: 10767–10777.

95.

A. C.

Fisher

T. E.

Hertz

Tildon

J. T.

Schousboe

Hertz

(1984) Metabolic fate of [14C]-glutamine in mouse cerebral neurons in primary cultures. Journal of Neuroscience Research 11: 351–357.

96.

A. C.

Schousboe

Hertz

(1982) Metabolic fate of 14C-labeled glutamate in astrocytes in primary cultures. Journal of Neurochemistry 39: 954–960.