Nuclear Factor E2-Related Factor 2-Dependent Antioxidant Response Element Activation by tert-Butylhydroquinone and Sulforaphane Occurring Preferentially in Astrocytes Conditions Neurons against Oxidative Insult

Introduction

The intraneuronal accumulation of reactive oxygen species has been implicated in the pathogenesis of many neurodegenerative diseases such as Alzheimer's, Parkinson's, and amyotrophic lateral sclerosis (Coyle and Puttfarcken, 1993; Simonian and Coyle, 1996; Mattson, 2000). The ability of a cell to neutralize reactive intermediates is, in part, dependent on the activation of a cis-acting regulatory element termed the antioxidant response element (ARE). The ARE is located in the 5′-flanking region of many genes essential for both detoxification and maintenance of cellular reducing potential (Rushmore et al., 1990, 1991). Thus, a powerful cluster of protective genes can be coordinately upregulated through the ARE to reduce the damage caused by harmful toxicants.

The functional characterization of the ARE sequence identified a core, TGACnnnGC, necessary for this transcriptional response (Rushmore et al., 1991). This finding led to the discovery of similar sequences in the promoter region of a growing list of neuroprotective genes that bind the transcription factor nuclear factor E2-related factor 2 (Nrf2) (Moi et al., 1994).

ARE induction by small-molecule activators such as tert-butylhydroquinone (tBHQ) has been shown to protect neuronal cell lines against the oxidative insult initiated by dopamine, hydrogen peroxide (H₂O₂), and glutamate (Murphy et al., 1991; Duffy et al., 1998; Li et al., 2002). This activation is dependent on the translocation of Nrf2 to the nucleus through inducer interaction with Keap1, the cytoplasmic repressor of Nrf2 (Dinkova-Kostova et al., 2002). In the nucleus, Nrf2 heterodimerizes with small Maf proteins before binding the ARE (Marini et al., 1997; Nguyen et al., 2000).

Additional studies have shown that, in rodent primary neuronal cultures, both basal expression and activation of the ARE are seen predominately in the astrocyte, not neuronal, subpopulation (Ahlgren-Beckendorf et al., 1999; Eftekharpour et al., 2000; Murphy et al., 2001; Johnson et al., 2002). We speculate that this is attributable to selective Nrf2 expression in astrocytes over neurons (Shih et al., 2003). Our previous work has shown that Nrf2 overexpression in astrocytes can protect naive neurons (Shih et al., 2003), and that cortical neurons derived from Nrf2 knock-out (KO) mice have a heightened sensitivity to oxidative stressors (Lee et al., 2003b). These data suggest that activation of the Nrf2–ARE pathway in astrocytes confers protection to vulnerable neurons. The individual contributions of each cell type to this neuroprotection are yet to be defined.

To further elucidate the factors contributing to a protective response manifest by ARE activation, we used mouse primary cortical neuronal cultures to look at two paradigms of oxidative toxicity: H₂O₂ and glutamate. Cultures were made resistant to these toxicities through previous activation of the ARE by tBHQ and sulforaphane. The tBHQ-mediated protection was absent in Nrf2 KO cultures and inhibited by viral infection with dominant-negative (DN) Nrf2, as well as the phosphatidylinositol-3 (PI-3) kinase inhibitor LY294002 (LY). Lastly, the preferential adenoviral infection of glial cells in this mixed culture system was exploited, allowing us to use cell-sorting techniques to determine the cell type-specific, Nrf2-dependent genetic changes associated with tBHQ-mediated neuroprotection.

Materials and Methods

Animals. ARE–human placental alkaline phosphatase (hPAP) transgenic mice were created using 51 bp of the NAD(P)H:quinone oxidoreductase (NQO1) promoter upstream of a hPAP reporter construct (Johnson et al., 2002). Nrf2 KO mice were graciously provided by Drs. Kaiman Chan and Yuet Wai Kan (Howard Hughes Medical Institute, University of California, San Francisco, CA). The mice were created by replacement of the basic region-leucine zipper region with lacZ (Chan et al., 1996). Glial fibrillary acidic protein (GFAP)–green fluorescent protein (GFP) mice were a generous gift from Dr. Albee Messing (University of Wisconsin, Madison, WI). Green fluorescent protein (humanized GFP-S65T) was expressed under the control of the astrocyte-specific GFAP promoter (Zhuo et al., 1997). All of the ARE–hPAP mice were outbred onto C57B6/SJL mice (The Jackson Laboratory, Bar Harbor, ME) for experiments and thus were of mixed background.

Chemicals. Sulforaphane was purchased from LKT Labs (St. Paul, MN). All of the other chemicals were purchased from Fisher Scientific (Fair Lawn, NJ).

Mouse primary cortical cultures (modified from Hertz et al., 1985). On embryonic day 15 (E15), pregnant mothers were anesthetized with isoflurane and killed by cervical dislocation; pups were removed. The cortices were dissected and removed to HBSS. The tissue was dissociated with 0.05% trypsin in HBSS for 10 min at 37°C, and then filtered through a 70 μm cell strainer. Cells were plated on poly-d-lysine (PDL)-coated plates at ∼3.2 × 10⁵ cells/cm² in CEMEM [10% fetal bovine serum, 10% horse serum, 2 mm l-glutamine, and 1% penicillin/streptomycin] for 45 min in a 37°C humidified tri-gas incubator (5% CO₂ and 5% O₂). For day 2 in vitro (2DIV) studies, treatments were performed after the initial 45 min incubation in CEMEM. For 5DIV studies, the cells were incubated in CEMEM for 48 hr before a medium change to Neurobasal medium (Invitrogen, Carlsbad, CA) with a B27 supplement containing antioxidants (NB plus AO) to specifically enhance neuronal growth. On the following day, the treatments were performed. Unless otherwise specified, after incubation with the treatment for 48 hr, the medium was changed to Neurobasal medium containing B27 supplement without antioxidants (NB minus AO) before the toxins were applied. This was done to prevent the direct interaction of the toxin with either the treatment or antioxidants such as catalase present in the medium.

Adenovirus. Recombinant adenovirus (ad) for enhanced GFP (eGFP) (ad-GFP), Nrf2-GFP (ad-Nrf2), and DN-Nrf2-GFP (ad-DN-Nrf2) were prepared, in collaboration with Dr. Tim Murphy at the University of Vancouver (Vancouver, British Columbia, Canada), by the Adenovirus Core Facility (Canadian Stroke Network Core Facility, University of Ottawa, Ottawa, Ontario, Canada). The Nrf2-cDNA was excised from the pEF/Nrf2 (Alam et al., 1999) using NotI and HindIII. These replication-deficient constructs were created using the Cre-lox system (Hardy et al., 1997) and were titered on human embryonic kidney 293 cells (Shih et al., 2003).

Treatments. All of the chemical pretreatments [vehicle is 0.005% ethanol (EtOH)] were added directly to the medium and were washed out in a medium change before toxin exposure. Cultures were incubated with replication-deficient adenovirus in MEM for 45 min at 37°C, after which the conditioned medium was added back to the cells.

Cytotoxicity. Forty-eight hours after treatments, cultures were incubated in either hydrogen peroxide or glutamate in fresh NB minus AO. On 5DIV, cells were incubated with H₂O₂ for 3 hr at 37°C. Catalase was added to the cultures at 105 U/ml to inactivate H₂O₂. Cultures were allowed to recover for 5 hr before a 3.5 hr quantification of cell viability using the Promega (Madison, WI) 3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl) 2H tetrazolium, inner salt (MTS) assay. For 2DIV cultures, cells were incubated with glutamate for 24 hr at 37°C before quantification of cell viability using the MTS assay.

Immunofluorescence. Primary antibodies were used at the following dilutions: anti-β(III)-tubulin, 1:200 (Promega); anti-GFAP, 1:1000 (Dako, Glostrup, Denmark); and anti-hPAP (clone 8B6), 1:400 (Sigma, St. Louis, MO). All of the secondary antibodies conjugated to either fluorescein or Texas Red were used at a dilution of 1:200, and appropriate IgG controls were run simultaneously. Terminal deoxynucleotidyl transferase-mediated biotinylated UTP nick end labeling (TUNEL) staining was performed using the Cell Death Detection kit from Roche (Indianapolis, IN) according to the manufacturer's instructions. Incubation of slides in Hoechst 33258 (0.1 mm in PBS; Sigma) was used to stain nuclear DNA. All of the images were taken using a Zeiss (Oberkochen, Germany) photomicroscope and analyzed using Axiovision software.

Immunofluorescence counts. Image fields (200×) were used to count cells displaying given immunofluorescent markers. Counts were expressed as percentage of vehicle to normalize for differences in cell density, immunostaining, and basal survivability variations across cultures. Cells displaying highly condensed nuclei, visible by bright spotting of Hoechst 33258 staining, were no longer considered viable and thus were not counted. GFAP-expressing cell counts had higher error values attributable to fewer cell numbers, as well as clustering of the cells beneath neurons, making exact counts difficult. hPAP enzyme activity assay. Cells were harvested in TMNC lysis buffer (0.05 m Tris, 0.005 m MgCl₂, 0.1 m NaCl, 1% 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate [CHAPS]) and freeze-thawed at –80°C. Endogenous phosphatase activity was heat inactivated at 65°C for 30 min in 0.2 m diethanolamine buffer. hPAP activity was quantitated by addition of the Applied Biosystems (Bedford, MA) substrates disodium 3-(4-methoxyspiro {1,2-dioxetane-3,2′-(5′-chloro) tricyclo [3.3.1.1 3,7]decan}-4-yl) phenyl phosphate and Emerald. A Berthold Orion microplate luminometer was used to measure the samples at various times after substrate addition. Values were corrected to an ARE–hPAP-negative culture and normalized by cell number at initial plating.

Total reduced glutathione assay (modified from Tietze, 1969). Cells were harvested in TMNC lysis buffer (0.05 m Tris, 0.005 m MgCl₂, 0.1 m NaCl, 1% CHAPS) and frozen at –80°C. Protein was precipitated from the lysates by mixing with an equal volume of 6% perchloric acid (PCA). Samples were spun at 7400 × g for 10 min at 4°C. The supernatant was diluted with 9 vols of 0.1 m sodium phosphate to neutralize the PCA. Reduced glutathione (GSH) standards and samples were further diluted six times with assay buffer (0.1 mg/ml dithionitrobenzoic acid and 0.32 mg/ml NADPH, in 0.1 m potassium phosphate). The reaction was started using 0.08 U of glutathione disulfide reductase per well of a 96 well plate. The absorbance was kinetically read at 410 nm, and the resulting values were corrected for protein via the BCA method (Pierce, Rockford, IL).

Reverse transcription-PCR. Cells were treated with tBHQ for 24 hr and harvested in TRIzol reagent (Invitrogen). After reverse transcription with oligo-dT primers using the Promega reverse transcription kit, the cDNA was probed using the following primers (IDT, Coralville, IA): β-actin (393 bp), 5′-tgttaccaactgggacgaca-3′ and 5′-ctctcagctgtggtggtgaa-3′; hPAP (351 bp), 5′-tgggtagctgggactacagg-3′ and 5′-ttgccccaaatctcaacttc-3′; Nrf2 (430 bp), 5′-aggcatcttgtttgggaatg-3′ and 5′-agttctcgctgctcggacta-3′; NQO1 (500 bp), 5′-ttcaaccccatcatttccag-3′ and 5′-tccagacgtttcttccatcc-3′; heme oxygenase-1 (HO-1) (603 bp), 5′-tacacatccaagccgagaat-3′ and 5′-gttcctctgtcagcatcacc-3′; and glutathione S-transferase (GST) μ1 (GSTm1) (450 bp), 5′-ctcccgactttgacagaagc-3′ and 5′-caggaagtccctcaggtttg-3′.

Specific-primed reverse transcription-PCR. The labeled amplified RNA (aRNA) isolated (see below) was used in a reverse transcription reaction with the specific sense (L) primers listed below rather than oligo primers. This cDNA was then analyzed in a PCR reaction with both the sense and antisense (L and R) primers listed below: β-actin (L), 5′-tgttaccaactgggacgaca-3′, and (R) 5′-ctctcagctgtggtggtgaa-3′; GFAP (L), 5′-ggagagggacaactttgcac-3′, and (R), 5′-gctctagggactcgttcgtg-3′; and β(III)-tubulin (L), 5′-gcttccagctgacacactca-3′, and (R), 5′-ttcttggcatcgaacatctg-3′.

Cell sorting. Littermate embryos were separately cultured as above into PDL-coated six well plates at two wells per embryo. After 2 d, the medium was changed to NB plus AO, and treatments were performed the following day. After incubation with the treatments for 24 hr, the cells were gently lifted with Accutase (US Biotechnologies, Parker Ford, PA) at 37°C for 20 min, spun down at 300 × g for 2 min, and resuspended in diethylpyrocarbonate (DEPC)–PBS with 1% BSA. Samples were kept on ice and filtered through a 41 μm strainer before sorting to remove large aggregates. Cells were sorted using a FACSVantage SE equipped with FACS DiVa option into carrier solution (DEPC–PBS with BSA). To sort, cells were excited with a 488 nm argon laser at 100 mW, and GFP emission was collected through a 530/30 bandpass filter. The populations were sorted from viable cells [using forward scatter (FSC)/side scatter (SSC) size dimensions to discard debris] that did not display potential double characteristics (FSC-width windows). In addition, in an attempt to get solely GFP-expressing cells in one pool and completely GFP-negative cells in another, a third pool with low fluorescence was collected and discarded [fluorescence detector 1 (FL1)-GFP from ∼5 to 25] (see Fig. 7). The sorted populations were spun at 300 × g for 2 min, resuspended in TRIzol reagent (<0.25 ml per 50,000 cells), and stored at –80°C before RNA isolation. A sort check after the separation indicated >99.9% purity in the GFP-negative pool and >95% purity in the GFP-positive pool [using propidium iodide (PI), the outlying 5% in the GFP-positive pool revealed a significant percentage of dead cells that were swollen and/or had some autofluorescence; it should be noted that the RNA from these compromised cells would be quickly degraded and eliminated from additional isolation steps].

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Figure 7.

Cell sorting of adenovirus-infected cultures reveals two distinctly sized populations, correlating to an astrocyte-enriched versus a neuronal population. 3DIV cultures were infected with adenovirus encoding either GFP (ad-GFP) or DN-Nrf2-GFP (ad-DN-Nrf2) and treated with 10 μm tBHQ or vehicle for 24 hr. Cells were lifted with Accutase, filtered, and sorted according to their GFP content (indicating viral infection). A, The first sort gate was used to determine that ∼69% of the total population were deemed to be healthy cells free of debris according to size (F, R1). All of the subsequent gates were set on R1 so that only viable cells were analyzed and sorted. B, R2 shows the viable cell fluorescence for GFP (FL1, x-axis). The y-axis (FL2) is a measure of the orange fluorescence, because very strong GFP fluorescence can emit orange. These axes were used during postsort checks to compensate for dead cells that take up PI versus GFP fluorescence. R2 indicates that ∼21% of the viable cells were considered GFP-positive, and 79% were considered negative (see F). These, however, were not the final populations used to represent the GFP-positive and -negative populations. In C, the viable cells (R1) were gated for GFP fluorescence along the y-axis, but also for width along the x-axis (to remove doublets or clumps of cells from the sort that, potentially, could contain both GFP-positive and -negative cells). Thus, R3 represents the GFP-positive population isolated, and R4 represents the GFP negative [note that a large window was left between R3 and R4 in an attempt to get highly pure populations, eliminating cells with medium fluorescence (y-axis)]. Of the viable cells, the GFP-positive population was ∼14%, and the GFP-negative population was ∼52% (C; F, R3 and R4). D, The size of the cells isolated as the GFP-positive pool of cells is shown. E,The size of cells isolated as the GFP-negative population is indicated. Larger cells have higher FSC-height and/or higher SSC-height values (R3 cell size > R4 cell size). G, Mixed wild-type cultures were infected with 50 MOI ad-GFP on 3DIV and treated with tBHQ vehicle or 10 μm tBHQ 24 hr before cell sorting. Amplified RNA isolated from GFP-positive (+) and GFP-negative (–) sorted cell populations was then subjected to specific-primed reverse transcription with the sense primers for GFAP, β(III)-tubulin, and β-actin (internal control). PCR was then run on the samples using 3′ and 5′ primers. GFP-positive (+) populations (adenovirus-infected cells) from either tBHQ- or vehicle-treated cultures have transcripts highly enriched for the glial marker, GFAP, when compared with the neuronal marker, β(III)-tubulin. The GFP-negative (–) populations (uninfected) are nearly entirely made up of transcripts for the neuronal marker, β(III)-tubulin, with minimal to no expression of GFAP.

GeneChip analysis. E15 individual primary cultures were grown in CEMEM for 2DIV before changing the medium to NB plus AO. On the subsequent day, cultures were infected with either 50 multiplicity of infection (MOI) ad-GFP or ad-DN-Nrf2 for 45 min in MEM. After replacement of the conditioned medium, the cultures were treated with vehicle or 10 μm tBHQ. Twenty-four hours later, cells were lifted using Accutase and sorted in single-cell suspensions, resuspended in TRIzol, and frozen at –80°C. RNA was isolated from TRIzol following the Invitrogen protocol for RNA isolation. Double-stranded cDNA was synthesized from the RNA according to the Ambion (Austin, TX) protocol for use of their MessageAmp aRNA kit; purified using glycogen, ethanol, and ammonium acetate; and then in vitro transcribed to aRNA using the Ambion T7 ribonucleotides. The aRNA was purified, and the samples were reverse transcribed back to cDNA using second-round primers. A second round of in vitro transcriptional amplification to aRNA was performed, this time using the Enzo (Farmingdale, NY) biotin-labeled ribonucleotides at a final concentration of 7.5 mm. The end product was purified using Qiagen (Valencia, CA) RNeasy columns. The concentration and quality of the labeled aRNA were assessed, and 15 μg of fragmented aRNA was hybridized for 16 hr at 45°C to a murine U74Av2 array (Affymetrix, Santa Clara, CA). The arrays were automatically washed and stained with streptavidin–phycoerythrin using a fluidics station. Last, probe arrays were scanned at 3 μm resolution using the Genechip System confocal scanner made for Affymetrix by Agilent. Affymetrix Microarray Suite 5.0 was used to determine the relative intensities of each gene and the corresponding treatment comparisons. Intensity readings were scaled to a constant value (scale factor). To determine the rank of the changed genes, two intergroup comparisons (2 × 2 matrix) were performed based on the change called (0, no change; 1/–1, marginal increase/marginal decrease; 2/–2, increase/decrease). The sum of these four difference calls gave the final rank. The ranks varied from –8 to 8, and values were cut off at –4/4. Analysis parameters were set at γ1 = 0.01 and γ2 = 0.05; thus, ranks of –8/8 and –4/4 approximate p < 0.01 and p < 0.05, respectively. To increase the stringency of the analysis, genes with a coefficient of variation (CV) of >1.0 were eliminated^a (Li and Johnson, 2002). In addition, a value of 1.5 for average fold change (FC) was set as the cutoff for increased genes, and decreased genes below –3-fold are shown^a (note that some genes ranked 4/–4 may be omitted for reasons of space; see supplementary tables for a complete listing at http://www.pharmacy.wisc.edu/facstaff/sciences/JohnsonGroup/microdata.cfm).

To achieve a list of potential neuronal versus astrocyte markers, only those genes with a rank of 8 from a 2 × 2 comparison of the GFP vehicle-positive and -negative pools were considered. These genes with FC greater than threefold and CV of <1.0 were then analyzed against a single comparison (n = 1) of populations from a set of sorted cultures derived from GFAP-driven GFP transgenic mice. Only those genes ranking 8 from the 2 × 2 comparison that were also appropriately called increased or decreased from the GFAP–GFP cultures were chosen to demonstrate potential cell type-specific markers in this culture system.

Supplementary material. Additional data for this article are available online at http://www.pharmacy.wisc.edu/facstaff/sciences/JohnsonGroup/microdata.cfm.

Results

tBHQ and sulforaphane activate the ARE in astrocytes

Cells derived from animals expressing the ARE–hPAP transgene (Johnson et al., 2002) were evaluated for hPAP expression 48 hr after treatment with either tBHQ or sulforaphane. Both tBHQ (Fig. 1A) and sulforaphane (B) induced a dramatic, dose-dependent increase in hPAP activity. The mRNA levels for ARE-driven hPAP, Nrf2-dependent GSTm1 (Chanas et al., 2002), and other ARE-dependent genes (Fig. 1C), as well as total GSH levels (D), were correspondingly increased in the cortical neuronal cultures.

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Figure 1.

ARE activity in mixed (neurons and glia) cortical cultures after treatment with tBHQ or sulforaphane. A, B, Fold change indicating the relative conversion of an hPAP substrate in cortical cultures derived from transgenic animals containing an hPAP reporter construct under the control of 41 bp of the NQ01 promoter containing the ARE sequence. Cultures were treated with tBHQ (A) or sulforaphane (B) for 48 hr before an enzyme assay for hPAP activity. Cultures from animals negative for the ARE–hPAP transgene were used to subtract background luminescence. Values indicate the fold increase in ARE-driven hPAP substrate conversion compared with vehicle (n = 4; mean ± SE; *p < 0.05). C, RT-PCR of RNA isolated from ARE–hPAP cultures treated with 10 μm tBHQ for 24 hr. Treated cultures were assessed for the presence of increased mRNA of known ARE-dependent genes [GSTm1, HO-1, NQO1, hPAP (reporter gene), β-actin (internal control)]. D, Total intracellular glutathione was determined for individual, mixed primary cultures treated with either 10μm tBHQ or vehicle (0.005% EtOH) for 48 hr (n = 3; mean ± SE; *p < 0.05). GSH, Total intracellular glutathione (reduced form). E, Immunofluorescence staining of ARE–hPAP cultures 48 hr after treatment with 2.5 μm sulforaphane (bottom panels) versus vehicle (top panels). Anti-hPAP antibody is shown as punctate red staining (Texas Red) and anti-GFAP is green (FITC). The majority of cells displaying hPAP protein have GFAP-positive processes or GFAP staining overlaid with hPAP fluorescence (arrows). Small cells stained with Hoechst 33258 (blue) that do not display GFAP staining are likely neurons.

tBHQ has been shown previously to activate the ARE in the astrocyte subpopulation rather than neurons using analyses of either ARE-responsive genes such as NQO1 and GSTp1 (GST π1) (Ahlgren-Beckendorf et al., 1999) or an hPAP reporter construct (Murphy et al., 2001; Johnson et al., 2002) (data not shown). Immunofluorescence staining using an antibody to the hPAP protein (Texas Red) in conjunction with an antibody to the astrocyte marker GFAP (FITC) demonstrated a large increase in the presence of hPAP protein in GFAP-positive cells displaying astrocytic morphology (flattened with large cell bodies) after treatment with tBHQ (data not shown) or sulforaphane (Fig. 1E, arrows). Very few cells displaying neuronal morphology exhibited increased hPAP protein. Although all of the astrocytes visible in the sulforaphane-treated fields are not expressing the hPAP reporter protein, the majority of them are likely ARE activated. This reflects the fact that mixed embryos (+/– and –/– for the transgene because of breeding +/–males to –/–females) were used to generate these cultures; therefore, not all of the cells contain the hPAP transgene. In addition, chemicals structurally related to tBHQ such as butylated hydroxyanisole (–OCH₃ in 4′ position instead of –OH) and di-tert-butylhydroquinone (additional tert-butyl group in the 5′ position) did not increase activation of the ARE as determined by hPAP enzymatic activity (data not shown).

tBHQ and sulforaphane pretreatment protects against oxidative stress-induced death in neurons

To determine whether these small-molecule inducers of the ARE can protect cells against oxidative stress, cultures were exposed to hydrogen peroxide (Fig. 2) and glutamate (Fig. 3). H₂O₂-induced cellular toxicity is characterized by a depletion of intracellular glutathione (GSH) and ATP, increased Ca²⁺ levels, and lipid peroxidation (Fernandez-Checa et al., 1997; Li et al., 1997). Oxidative glutamate toxicity results in high levels of extracellular glutamate that competitively inhibits the uptake of cystine by immature neural cells via the system x_c^– [cystine transporter (xCT)] (Murphy et al., 1989, 1990). The subsequent decrease in intracellular concentrations of cysteine results in lowered GSH levels and NMDA receptor-independent cell death, presumably because of the accumulation of free radicals. It is very important to note that all of the cytotoxicity experiments were performed in fresh medium (NB minus AO), and that both tBHQ and sulforaphane were not present in the medium with H₂O₂ or glutamate. This was done in an effort to examine the changes in gene expression rather than any direct antioxidant properties the treatments may possess. As observed by immunofluorescent staining, tBHQ pretreatment for 48 hr showed a remarkable attenuation of cytotoxicity induced by H₂O₂ (Fig. 2A–C). Sulforaphane was just as effective as tBHQ at blocking this cytotoxicity (Fig. 2D). Actual cell counts of the fluorescent-stained cultures support the visual observation that tBHQ or sulforaphane can protect these cells from H₂O₂-induced cytotoxicity. By setting vehicle-treated images at 100% (n = 8), it was determined that 50 μm H₂O₂ significantly decreased the total cells (Hoechst; n = 6) present to 20.7 ± 3.90% of vehicle. Within the cell subpopulations, this correlated to a significant decrease in neuronal [β(III)-tubulin; n = 6] viability to 4.30 ± 1.40% of vehicle and astrocytes (GFAP; n = 6) to 59.9 ± 20.7% of vehicle. Pretreatment with 10 μm tBHQ (n = 8) significantly attenuated the total cell death (81.2 ± 6.30%), the neuron-specific death (53.0 ± 6.80%), and the astrocytic death (86.7 ± 9.00%) caused by treatment with H₂O₂. Pretreatment with 2.5 μm sulforaphane (n = 4) resulted in similar protection against H₂O₂-induced cytotoxicity [Hoechst, 115 ± 11.6%; β(III)-tubulin, 76.5 ± 10.8%; and GFAP, 111 ± 13.7% of vehicle]. Interestingly, cell death induced by hydrogen peroxide was more selective for neurons than astrocytes at the doses used. Pretreatment of mixed cultures with tBHQ or sulforaphane exhibited a pattern of increased GFAP-positive and β(III)-tubulin-positive cells, indicating an enhanced survival of astrocytes and neurons, respectively. In addition, hydrogen peroxide leads to an increase in TUNEL-positive cells that was also attenuated by tBHQ pretreatment (data not shown). Furthermore, the overall cell viability, as measured by the MTS assay, of cultures exposed to increasing doses of H₂O₂ was enhanced when pretreated with 10 μm tBHQ for 48 hr (Fig. 2E). Previous studies have implicated PI-3 kinase in ARE activation by tBHQ (Lee et al., 2001; Johnson et al., 2002; Kang et al., 2002). Hence, we pretreated cultures with the PI-3 kinase inhibitor LY for 30 min before tBHQ treatment. Indeed, LY pretreatment blocked both the activation of the ARE promoter, similar to that seen previously (Johnson et al., 2002) (data not shown), and inhibited the tBHQ-induced protection against hydrogen peroxide (Fig. 2F).

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Figure 2.

tBHQ-mediated attenuation of H₂O₂-induced cell death and loss of protection by inhibition of PI-3 kinase. A–D, Cells were pretreated with vehicle (A and B), 10 μm tBHQ(C), or 2.5 μm sulforaphane (D) for 48 hr before 50μm H₂O₂ treatment (B–D). A, Hydrogen peroxide vehicle (PBS) treatment is represented. Neurons are indicated by red staining for β(III)-tubulin and astrocytes are shown in green (GFAP). The nuclear counterstain (Hoechst 33258; blue) shows condensed (bright) and fragmented DNA in apoptotic cells (magnification, 200×). E, 5DIV cultures pretreated for 48 hr with 10μm tBHQ were exposed to a 3 hr incubation with hydrogen peroxide. Cell viability was assessed using the soluble MTS assay (n = 4; mean ± SE; *p < 0.05, compared with vehicle). Error bars indicate mean ± SEM for four experiments. F, ARE–hPAP cultures were pretreated for 30 min with the PI-3 kinase inhibitor LY (10 μm) before a 10 μm tBHQ (T) treatment. H₂O₂ cytotoxicity at 75 μm was used 48 hr after tBHQ treatment to observe the effect of LY pretreatment on tBHQ-mediated neuroprotection (LY, 10 μm LY294002; LY Veh, LY vehicle, 0.01% DMSO; T, 10 μm tBHQ; V, tBHQ vehicle, 0.005% EtOH; n = 3; *p < 0.05, compared with LY Veh/H₂O₂ vehicle; **p < 0.05, compared with LY Veh/H₂O₂ vehicle and compared with LY vehicle/tBHQ with 75 μm H₂O₂). Error bars indicate mean ± SEM for three experiments.

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Figure 3.

tBHQ-mediated attenuation of glutamate-induced cell death. A, 2DIV cultures were incubated with a high extracellular concentration of glutamate for 24 hr after a 48 hr pretreatment with either 10 μm tBHQ or ethanol vehicle (*p < 0.05; n = 4; mean ± SE). Cell viability was assessed using the soluble MTS assay. B–D, Oxidative glutamate (Glu) (1 mm) toxicity after a 48 hr treatment with vehicle (C) or 10μm tBHQ (D). Error bars indicate mean ± SEM for four experiments. B, Cells after treatment with glutamate vehicle (double distilled H₂O) are shown. β(III)-tubulin (red) indicates the remaining neurons and GFAP (green), the scattered astrocytes (magnification, 400×). E–H, TUNEL stain of cells displaying fragmented DNA (bright green; arrows) overlaid with Hoechst 33258 (blue) and anti-hPAP antibody (red) immunofluorescence 24 hr after treatment with glutamate vehicle (E, F) or 1 mm glutamate (G, H) pretreated for 48 hr with tBHQ vehicle (E, G) or 10μm tBHQ (F, H) (magnification, 400×). Note: B–D represent a separate culture from E–H.

Similar results to that described above were observed with respect to glutamate-induced cytotoxicity. Cell viability (Fig. 3A), immunofluorescent staining (B–D), and cell counts of cultures pretreated with 10 μm tBHQ (or 2.5 μm sulforaphane) (data not shown) before glutamate exposure, demonstrated similar survival characteristics. In addition, TUNEL-positive cells were dramatically decreased in cultures pretreated with tBHQ before glutamate exposure (Fig. 3E–H). Immunofluorescent counts reflected the visual observations in that only 28.6 ± 0.80% of the Hoechst-stained cells and 7.70 ± 1.20% of the β(III)-tubulin-positive cells remained after 1 mm glutamate treatment (n = 4). Pretreatment with 10 μm tBHQ (n = 4) resulted in a significantly enhanced viability of both total cells and neurons to 75.9 ± 3.90% and 67.8 ± 3.90% of vehicle controls, respectively, when a 1 mm glutamate insult was applied. In addition, pretreatment with tBHQ resulted in marked attenuation of TUNEL-positive cell counts (percentages expressed as TUNEL-positive cells divided by total, Hoechst-stained nuclei). Whereas vehicle pretreatment resulted in 58.4 ± 4.90% of the cells displaying fragmented DNA after 1 mm glutamate insult, tBHQ-pretreated cultures exhibited only 13.8 ± 4.20% (glutamate vehicle treatment resulted in 4.80 ± 1.80% TUNEL positive; n = 3).

To ensure that the oxidative glutamate toxicity was indeed oxidative stress induced and not a result of NMDA receptor-induced excitotoxicity, cultures were pretreated with 100 μm N-acetyl-l-cysteine (NAC). NAC provides cells with an alternative form of cysteine for GSH synthesis by circumventing the xCT. Pretreatment with NAC showed a complete block of glutamate-induced cellular death, whereas the synthetic NMDA receptor inhibitor MK-801 did not block cell death (data not shown).

ARE activation and protection against oxidative stress are dependent on Nrf2

The induction of genes in response to tBHQ has been shown to be dependent on binding of the transcription factor Nrf2 to the ARE core sequence (Lee et al., 2003a). To look at the dependence on Nrf2 in a mixed culture of astrocytes and neurons, we prepared cultures from ARE–hPAP × Nrf2 KO mice (Chan et al., 1996). Nrf2 knock-out mice have a heightened sensitivity to oxidative stress, including an increased susceptibility to carcinogenesis (Ramos-Gomez et al., 2001). In these cultures, tBHQ was unable to induce ARE activity (data not shown), exhibited no protection versus hydrogen peroxide (Fig. 4A) (note: KO cells are a bit more sensitive to H₂O₂) or glutamate (data not shown), and displayed no enhancement of β(III)-tubulin-positive cell survival after hydrogen peroxide treatment (Fig. 4B). Immunofluorescence cell counts revealed 20.8 ± 3.90% versus 27.2 ± 4.60% total (Hoechst-stained) cell survival after a 50 μm H₂O₂ insult subsequent to pretreatment with vehicle or 10 μm tBHQ, respectively. Neuronal survivability after insult was also unchanged (vehicle pretreatment, 3.2 ± 1.3%, and tBHQ, 2.6 ± 0.9% of H₂O₂ vehicle controls). It should be noted that tBHQ treatment alone resulted in no significant cell count changes from those seen with vehicle treatment (n = 4).

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Figure 4.

In Nrf2 KO cultures, in which the effect of tBHQ on ARE activity is lost, neuronal protection is abrogated. A, Pretreatment of Nrf2 KO cultures with 10μm tBHQ and vehicle for 48 hr was performed before H₂O₂ treatment. Error bars indicate mean ± SEM for four experiments. B, KO cultures were treated with 10 μm tBHQ or vehicle for 48 hr before a 50 μm insult of H₂O₂. Cultures were then stained for β(III)-tubulin (red) to qualify the presence of neurons and Hoechst 33258 (blue) to indicate additional cell types remaining, as well as apoptotic cells (displaying condensed or fragmented nuclei; magnification, 200×).

Overexpression of Nrf2 restores neuroprotection and ARE activity

In an effort to further establish that Nrf2 is necessary for protection, knock-out cultures were infected with replication-deficient adenovirus encoding Nrf2 (ad-Nrf2). An MOI (plaque-forming units per cell) of 50 resulted in a near-complete protection of the cultures against hydrogen peroxide (Fig. 5A) or glutamate (data not shown). Ad-Nrf2 infection also resulted in an increase in ARE reporter activation as seen by immunofluorescent staining as well as hPAP enzyme activity (Fig. 5B,D). This correlated with a significantly increased survivability of β(III)-tubulin-positive cells (Fig. 5C). Immunofluorescence counts indicate 3.1 ± 1.7% survival of β(III)-tubulin positive cells infected with ad-GFP after insult, whereas infection with 50 MOI of ad-Nrf2 completely protected the neurons from glutamate toxicity (survival, 94.1 ± 4.3%). Cytotoxicity appears to be selective for neurons, because staining for β(III)-tubulin is drastically decreased in unprotected cultures (Fig. 5C) (in 0 and 50 MOI GFP-infected cultures treated with H₂O₂). Lastly, overexpression of Nrf2 resulted in an increase in total intracellular GSH levels (Fig. 5E) similar to that seen with tBHQ treatment (Fig. 1D).

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Figure 5.

Overexpression of Nrf2 resulting in ARE activation and neuroprotection. A, Nrf2 KO cultures were infected with 50 MOI ad-Nrf2 48 hr before H₂O₂ insult. (n = 4; *p < 0.05). B, ad-GFP- and ad-Nrf2-GFP (50 MOI)-infected cultures were observed for ARE activation in primary cultures from ARE–hPAP transgenics using hPAP antibody staining [hPAP antibody (red); eGFP fluorescence (green); Hoechst 33258 (blue); magnification, 400×]. C, Staining for the neuronal marker, β(III)-tubulin (red), after a 50 μm insult with H₂O₂ [eGFP (green); Hoechst 3258 (blue); 0 MOI, no virus/MEM alone; magnification, 400×]. D, E, Forty-eight hours after treatment with 50 MOI ad-GFP or 50 MOI ad-Nrf2, ARE–hPAP cultures were harvested in lysis buffer and evaluated for hPAP enzyme activity (D) and total glutathione (E) (50 GFP, 50 MOI ad-GFP; 50 Nrf2, 50 MOI ad-Nrf2). GSH concentration (E) was corrected for protein using the BCA method (n = 4; mean ± SE; *p < 0.05). Error bars in A, D, and E indicate mean ± SEM for four experiments.

Introduction of DN-Nrf2 to wild-type cultures results in a decrease in tBHQ-mediated cellular protection and ARE activation

To determine the endogenous contributions of Nrf2 to tBHQ-mediated events, cortical cultures from ARE–hPAP mice were infected with ad-DN-Nrf2 before tBHQ treatment. This mutant lacks the N-terminal transcription activation domain, but still retains the C-terminal DNA binding and heterodimerization domain. It is thought to inhibit Nrf2 by competing with the transcription factor for binding partners and/or DNA binding sites (Alam et al., 1999). Interestingly, when primary cortical cultures were injected with low concentrations of adenovirus (<75 MOI), it appeared that the viral infection was selective for astrocytic cell types (Fig. 6A,B). Therefore, low concentrations of the adenovirus were used to selectively infect the astrocytes in this mixed-culture system. Infection of astrocytes with ad-DN-Nrf2 before tBHQ treatment resulted in a large attenuation of tBHQ-mediated activation of the ARE response (Fig. 6C). Amazingly, this also led to a reversal of the tBHQ-mediated neuroprotection against hydrogen peroxide-induced cytotoxicity (Fig. 6D). Knowing that ARE activation by tBHQ occurs predominately in the astrocyte subpopulation, ad-DN-Nrf2 preferentially infects astrocytes, and neurons are more susceptible than astrocytes to H₂O₂-induced toxicity, these data suggest that ARE activation in astrocytes confers protection to vulnerable neurons (compared with ad-GFP controls).

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Figure 6.

Selective adenovirus infection of astrocytes leads to ad-DN-Nrf2-mediated reduction in ARE activation by tBHQ and inhibition of tBHQ-mediated neuroproteion. Infection of primary cortical cultures with with low MOIs (25–75) of adenovirus show ad-infected cells colocalized with GFAP-positive astrocytes (B, arrows) (50 MOI; magnification, 200×) and, very infrequently, β(III)-tubulin-positive neurons (C, arrowheads) (75 MOI; magnification, 400×). On 3DIV, ARE–hPAP cultures were infected with ad-DN-Nrf2 (DN) virus or ad-GFP (G) virus (25 or 50 MOI) for 45 min, followed by treatment with 10 μm tBHQ or tBHQ vehicle. C, ARE activation by tBHQ was quantified by conversion of an hPAP substrate to a luminescent product (fold change of vehicle is shown above each bar; n = 5; mean ± SE; *p < 0.05, significant increase compared with vehicle; **p < 0.05, significant increase compared with vehicle and significant decrease compared with 25 MOI ad-GFP + 10 μm tBHQ). Veh, Vehicle; T, 10 μm tBHQ; V, vehicle. D, Cytotoxicity caused by 50μm H₂O₂ was calculated using the MTS viability assay. The percentage of cell death induced in the presence of 50 MOI ad-GFP with tBHQ vehicle or 10 μm tBHQ (50 G/Veh and 50 G/T, respectively), was compared with the cell death induced in the presence of 10, 25, and 50 MOI of the DN-Nrf2 adenovirus with 10 μm tBHQ (n=3; mean ± SE; *p < 0.05, significant attenuation of cell death compared with 50 MOI GFP: vehicle-treated cultures exposed to 50 μm H₂O₂). Error bars in C and D indicate mean ± SEM for three experiments.

Discussion

This study demonstrates the importance of ARE activation in astrocytes of a mixed primary culture system. Data presented herein are the first demonstrating that sulforaphane activation of the ARE prevents neuronal cell death. Sulforaphane, tBHQ, and Nrf2 overexpression via adenovirus infection all induce an ARE-mediated genetic response. This response is highly selective for astrocytes over neurons and conveys neuroprotection from oxidative insults initiated by H₂O₂ or nonexcitotoxic glutamate toxicity. These two stressors target neurons preferentially over astrocytes, whereas the genetic changes induced directly by tBHQ seem to be confined to the astrocytes. Thus, a tBHQ-induced, Nrf2-dependent, coordinated change in the gene expression of astrocytes (∼15% of total) is sufficient to confer protection to the majority of cortical neurons. Finally, the combined use of cell sorting and microarray technologies allowed us to determine the cell-specific changes in gene expression that facilitate ARE-mediated protection. These findings lend support to the hypothesis that treatment of cells with ARE inducers leads to modifications of glia–neuronal interactions.

It has been well established that Nrf2 is the key transcription factor mediating ARE-driven gene expression. In addition, Nrf2 knock-out cultures of primary astrocytes (Lee et al., 2003a) and neurons (Lee et al., 2003b) have been shown to be more sensitive to oxidative stressors. The complete loss of Nrf2 from mixed neural cultures, using Nrf2 knock-out mice, results in a total loss of both ARE inducibility by tBHQ and tBHQ-mediated protection from toxins (Fig. 4). Reexpression of Nrf2 using a replication-deficient adenovirus dramatically increases ARE activation and neuroprotection (Fig. 5). We also recently published that a very small percentage of ad-Nrf2-infected astrocytes (∼1%) are sufficient to confer protection against oxidative glutamate toxicity toward naive cortical neurons (Shih et al., 2003). In an attempt to understand the difference between Nrf2 overexpression and ARE activation by tBHQ in this mixed culture system, we exploited the selective adenoviral infection of astrocytes to test whether tBHQ-mediated protection could be reversed by overexpression of DN-Nrf2 selectively in astrocytes. The neuroprotective effects of tBHQ were nullified, suggesting that activation of the Nrf2–ARE pathway in astrocytes alone was sufficient to confer neuroprotection. The question still remained, however, as to which genetic changes specific to astrocytes and/or neurons precipitated this protective effect.

The combined use of cell sorting and gene chip technology has allowed us to begin to determine the cell-specific contributions to ARE-mediated protection by tBHQ. Not surprisingly, the majority of the genes increased by tBHQ treatment, particularly those previously shown to be ARE-driven, were seen in the astrocyte-enriched population. There were, however, many genes that changed in both the neuronal and astrocyte-enriched populations, indicating either stimulation of the ARE in both or induced changes in one population by the other. Genes increased in concert include GSTm1, malic enzyme, stearoyl-CoA desaturases (Scd), and glucose phosphate isomerase (GPI). More striking were the genes decreased in both of the sorted populations. These include Notch 1, cyclin D2, p21, MAP tau, and thymopoietin, most of which regulate cell cycle events. In addition, there were dramatic tBHQ-induced changes to protein modification, calcium homeostasis, and intracellular phosphorylation cascades [including TGFβ, insulin, and nuclear factor-κB (NF-κB) pathways]. These alterations may modify multiple facets of the complex signaling systems directing cellular events.

The most intriguing aspect of observing ARE activity in a mixed culture system is the presence of communication between astrocytes and neurons. Astrocyte–neuron interactions are proposed to be involved in synapse formation and conduction, neurotransmitter release, process outgrowth, chemotaxis, energy production, and response to injury (Pellerin and Magistretti, 1994; Dugan et al., 1995; Vernadakis, 1996; Pellerin et al., 1998; Goritz et al., 2002). The modified genetic profile of ARE-activated astrocytes alludes to an enhancement of environmental detoxification and increased trophic factor secretion.

Most noticeable of the changes induced by tBHQ specifically in the astrocyte-enriched population is the large increase in cellular protective proteins. A battery of ARE-driven genes encoding detoxification enzymes were upregulated, as were enzymes associated with glutathione synthesis and conjugation reactions, although the latter were not significantly inhibited by DN-Nrf2 (supplementary Table 2, available at www.jneurosci.org). Interestingly, in addition to regulating the neural environment, astrocytes help to synthesize neuronal glutathione (Aschner, 2000). Thus, the astrocyte-dependent environmental detoxification that contributes to neuronal protection is, at least in part, regulated by this inducible system.

A striking number of genes associated with energy production were upregulated by tBHQ in the glia-enriched population, including the majority of the 10 genes catalyzing the glycolysis of glucose to pyruvate. Glucose metabolism is essential for calcium homeostasis in the brain via the synthesis of ATP (Nijjar and Belgrave, 1997). In addition, pyruvate can be further metabolized to lactate, the preferred energy substrate for neurons that helps to maintain synaptic transmission (Pellerin and Magistretti, 1994; Pellerin et al., 1998). The increases in astrocytic glycogen synthesis (glucan branching enzyme), gluconeogenesis (GPI), and neuronal creatine production [Gatm (glycine amidinotransferase) and creatine kinase] may also indicate an enhanced maintenance of neuronal activity by tBHQ. Interestingly, astrocytes are known to couple neuronal activity (glutamate release) to energy production and glutathione synthesis (Magistretti et al., 1999; Sonnewald et al., 2002). Additional metabolism of pyruvate to acetyl CoA leads to an increased synthesis of cholesterol (squalene epoxidase and lanosterol synthase) and fatty acids for phospholipids and energy storage (malic enzyme, fatty acid synthase, and Scd1 and -2). Increasingly, research reveals that neurons require glia-derived cholesterol to form efficient synaptic connections (Barres and Smith, 2001; Goritz et al., 2002; Pfrieger, 2003). Thus, stimulated astrocytes may, in addition to detoxification of the cellular environment, produce increasing levels of substrates essential for neuronal maintenance. An enhanced coupling of metabolism and energy utilization between astrocytes and neurons appears to be a key mechanism underlying tBHQ-inducible neuroprotection.

In addition to metabolic changes, cells responding to tBHQ treatment are modified in their extracellular interactions. An enhancement of physical interactions would help to facilitate much of the soluble and contact-dependent communication between neurons and glia. In the astrocyte-enriched subpopulation, tBHQ increased integrin signaling molecules [calpain, Nap1 (NCK-associated protein 1), and Thy1 (thymus cell antigen 1)], as well as cell adhesion proteins (fibronectin and laminin). These factors and integrin receptors are partially responsible for initiating neuron–glia interactions (Marchetti, 1997; Bialkowska et al., 2000; Leyton et al., 2001; Ding et al., 2002). In neurons, the elevated expression of neuropilin, calmyrin, and VCAM (vascular cell adhesion molecule) supports a tBHQ-induced enhancement of cellular contacts.

Astrocytes play a considerable role in neuronal synapse formation (Goritz et al., 2002; Pfrieger, 2003). In addition to glia-derived cholesterol and lactate, neurons are thought to require astrocytes during synapse maturation for soluble and contact-dependent factors. GABAergic signaling has been implicated in ARE activity (supplementary Table 3, available at www.jneurosci.org) (Lee et al., 2003b) and has been postulated to mediate neuron-to-glia signaling (Runquist and Alonso, 2003). Astrocytes respond to, and secrete, such neurochemicals (Danbolt, 2001; Fields and Stevens-Graham, 2002). Some genes increased in the astrocyte-enriched population are Rapsyn, neuroserpin, and syntaxin bp (binding protein) 3. Rapsyn clusters acetylcholine receptors; neuroserpin protects against plasminogen activators and helps form synapses; and syntaxin bp3 contributes to synaptic trafficking and glucose transport. In the neuronal subpopulation, transcripts for FGF14, a factor essential for synaptosomal function (Wang et al., 2002), synaptotagmins, which are essential for calcium-dependent synaptic vesicle docking, and galanin, which maintains neurotransmitter homeostasis, are all increased by tBHQ. Thus, it appears that tBHQ-induced astrocytes may be better primed to receive signals from neighboring neurons inducing motility, adhesion, or morphogenesis to facilitate energy and cholesterol production necessary for synapse formation and maintenance.

The ARE-inducible transmembrane and secreted proteins from both populations may be the direct mediators of astrocytic protection of neurons against insult. Basigin, a transmembrane glycoprotein, has been shown to induce the regrowth of damaged dopaminergic fibers after exposure to 1-methyl-4-phenylpyridinium (Mitsumoto et al., 2001). Gdf15 (growth/differentiation factor-15) is a secreted TGFβ family member that acts as a direct trophic factor for neurons (Strelau et al., 2000). CREG (cellular repressor of E1A genes) is a secreted glycoprotein that antagonizes E1A, an inhibitor of Nrf2 activity (Katoh et al., 2001). Endothelin-1 has been shown to increase astrocytic glial-derived neurotrophic factor secretion, induce extracellular and cytoskeletal reorganization, and protect against hypoxia and ischemia (Ho et al., 2001; Egnaczyk et al., 2003). Interestingly, the IK cytokine prevents immune response by inhibiting IFN-γ-induced [human leukocyte antigen (HLA) class II] gene expression (Krief et al., 1994). Transcripts induced by IFN were significantly decreased in both populations, whereas an HLA repressor, NFX1, was increased. This suggests a link between increased IK mRNA and resultant changes to gene expression. These factors, in conjunction with Thy1, FGF14, lactate, cholesterol, and glutathione, may comprise the soluble mediators of neuron–glia communication effecting neuroprotection.

The progressive neurodegeneration seen in many diseases of aging is due, in part, to the buildup of harmful oxidants. As a result of this toxin accumulation, these neurons display decreased synaptic efficiency or undergo cell death. Selective chemical activation of the ARE in astrocytes may provide a means of amplifying the existing metabolic, soluble, and physical coupling between astrocytes and neurons to boost synaptic function and provide protection against oxidative injury. Future work will confirm and analyze these changes within the intact architecture of the brain to pinpoint the exact protective mechanism(s) underlying this enhanced interaction in vivo.