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Originally published In Press as doi:10.1074/jbc.M410771200 on December 3, 2004

J. Biol. Chem., Vol. 280, Issue 7, 5875-5883, February 18, 2005
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Reversible Oxidation of ERK-directed Protein Phosphatases Drives Oxidative Toxicity in Neurons*

David J. Levinthal{ddagger} and Donald B. DeFranco{ddagger}§

From the {ddagger}Center for Neuroscience and §Department of Pharmacology, University of Pittsburgh School of Medicine, Pittsburgh, Pennsylvania 15261

Received for publication, September 20, 2004 , and in revised form, November 18, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Oxidative stress links diverse neuropathological conditions that include stroke, Parkinson's disease, and Alzheimer's disease and has been modeled in vitro with various paradigms that lead to neuronal cell death following the increased accumulation of reactive oxygen species. For example, immortalized neurons and immature primary cortical neurons undergo cell death in response to depletion of the antioxidant glutathione, which can be elicited by administration of glutamate at high concentrations. We have demonstrated previously that this glutamate-induced oxidative toxicity requires activation of the mitogen-activated protein kinase member ERK1/2, but the mechanisms by which this activation takes place in oxidatively stressed neurons are still not fully known. In this study, we demonstrate that during oxidative stress, ERK-directed phosphatases of both the serine/threonine- and tyrosine-directed classes are selectively and reversibly inhibited via a mechanism that is dependent upon the oxidation of cysteine thiols. Furthermore, the impact of ERK-directed phosphatases on ERK1/2 activation and oxidative toxicity in neurons was tested in a neuronal cell line and in primary cortical cultures. Overexpression of the highly ERK-specific phosphatase MKP3 and its catalytic mutant, MKP3 C293S, were neuroprotective in transiently transfected HT22 cells and primary neurons. The neuroprotective effect of the MKP3 C293S mutant, which enhances ERK1/2 phosphorylation but blocks its nuclear translocation, demonstrates the necessity for active ERK1/2 nuclear localization for oxidative toxicity in neurons. Together, these data implicate the inhibition of endogenous ERK-directed phosphatases as a mechanism that leads to aberrant ERK1/2 activation and nuclear accumulation during oxidative toxicity in neurons.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Oxidative stress is a common feature of a diverse range of neuropathological conditions, including stroke, Parkinson's disease, Alzheimer's disease, and amyotrophic lateral sclerosis (1, 2). Glutamate-induced oxidative toxicity provides an excellent model for studying the effects of oxidative stress in immortalized neurons and in primary neuronal cultures (3–5). In this model, inhibition of a glutamate/cystine antiporter, known as system xc-, leads to decreased accumulation of intracellular free cysteine, a necessary precursor of glutathione, and to eventual glutathione depletion. As a result, reactive oxygen species (ROS)1 accumulate and activate cellular signaling events that contribute to neuronal cell death. We have shown previously that oxidative toxicity causes a delayed, sustained activation of extracellular signal-regulated kinase (ERK) 1/2 that is necessary for neuronal cell death (6–8), but the mechanisms by which oxidative stress drives ERK1/2 activation have not been fully elucidated.

The regulation of ERK1/2 phosphorylation and activation reflects a subtle balance between ERK-directed kinase and phosphatase activity. A diverse range of phosphatases directed against phospho-serine/threonine, phospho-tyrosine, or both have been identified as negative regulators of ERK1/2. Protein phosphatase 2A (PP2A) (9, 10) and strial-enriched phosphatase (11), among numerous others, can function as ERK-directed phosphatases in neurons. Dynamic changes in ERK-directed phosphatase activity can result from several mechanisms, including ERK-dependent up-regulation of phosphatase expression (12), phosphorylation-dependent increases in phosphatase stability (13), and protein-protein interaction-dependent activation of phosphatase activity (14). Collectively, these events have been shown to function in a negative feedback loop that terminates ERK signaling (12).

Recently, the role of oxidative stress in the regulation of phosphatase activity has received much attention (15–17). Several phosphatases have been shown to be redox-sensitive and can be either reversibly or irreversibly inhibited, depending upon the degree and mechanism of oxidation (15, 18). Oxidative phosphatase inhibition can impact various cellular signaling pathways and accounts for a mechanism now referred to as oxidative signaling. The extent to which oxidative inhibition of phosphatases plays a role in driving signaling events during neurotoxicity remains relatively unexplored.

We sought to characterize the effect of glutamate-induced oxidative toxicity on ERK-directed phosphatases in primary neuronal cultures. In this study, we show that endogenous ERK-directed phosphatase activity is specifically and reversibly inhibited during oxidative stress, consistent with a mechanism involving the oxidation of cysteine thiols. This inhibition of ERK-directed phosphatases correlates with an increase in phosphorylated ERK levels. We show that this aggregate phosphatase activity is likely composed of PP2A and a vanadate-sensitive component. The impact of ERK-directed phosphatase activity on oxidative toxicity was revealed by the neuroprotective effects of overexpressed mitogen-activated protein kinase phosphatase 3 (MKP3) and its catalytic mutant, MKP3 C293S, in both HT22 cells and primary cortical neurons. Collectively, these data implicate oxidative inhibition of ERK-directed phosphatases in neuronal oxidative toxicity induced by glutathione depletion.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Plasmids—The expression plasmids for MKP3 WT and MKP3 C293S, as well as the parent plasmid (pSG5), were kind gifts from Drs. Steven Keyse and Anne Brunet (19–21). The mitochondria-targeted eYFP expression plasmid was a gift from Dr. Ian Reynolds, and the expression plasmid for the Elk-1/GAL4 fusion protein, the luciferase reporter, and the constitutive Renilla plasmid (all from Stratagene, La Jolla, CA) were purchased or donated by Dr. Elias Aizenman.

Primary Cortical Cultures—Cortices from embryonic day 17 Spague-Dawley rat fetuses (Hilltop Lab Animals, Scottdale, PA) were dissected and manually dissociated by repeated trituration using fire-polished glass pipettes in Hanks' balanced salt solution (5.4 mM KCl, 0.3 mM Na2HPO4, 0.4 mM KH2PO4, 4.2 mM NaHCO3, 137 mM NaCl, and 5.6 mM D-glucose, pH 7.4) without Ca2+ or Mg2+ (Invitrogen) followed by passage through a 40-µm cell strainer (BD Biosciences) to remove clumped cells. Cells were counted and plated on 50 µg/ml poly-D-lysine-treated culture plates at a density of ~2.1 x 104 cells/cm2. Cell viability was routinely greater than 80% as assessed by uptake of trypan blue dye upon plating. Cultures were maintained for 3–4 days in media (Dulbecco's modified Eagle's medium; Invitrogen), 10% fetal calf serum (Hyclone, Logan, UT), 10% Ham's F12 nutrient supplement (Invitrogen), 1.9 mM glutamine, 24 mM Hepes buffer, and 4.5 mg/ml glucose at 37 °C and 5% CO2. At this time, these mixed cortical cultures are predominately neuronal, with ~20% glial fibrillary-associated protein staining cells (3). Unless otherwise stated, all chemicals and reagents used were purchased from Sigma.

Cell Line Culture—HT22 hippocampal cells, which are sensitive to glutamate-induced oxidative toxicity (5), were maintained in Dulbecco's modified Eagle's medium supplemented with 10% fetal calf serum (Atlanta Biologicals, Norcross, GA), 100 units of penicillin, and 100 µg/ml streptomycin at 37 °C and 5% CO2.

Transfections in Primary Cortical Neuronal Cultures—A number of transfection reagents and protocols that were tested proved toxic to our primary immature cortical cultures. However, limited toxicity was observed when the 25-kDa polyamine polymer polyethyleneimine was used to form DNA complexes and transfect primary neurons (22, 23). In brief, 2 µl of a 100 mM polyethyleneimine stock solution was added to 250 µl of 150 mM NaCl, whereas 6.5 µg of DNA was added to 250 µl of 150 mM NaCl. Two µg of a mitochondria-targeted eYFP (mt-eYFP) plasmid DNA and 4.5 µg of MKP3 plasmids were used per condition. After 5 min, the two solutions were mixed. Polyethyleneimine-DNA complexes were allowed to form for 10 min and then diluted into 10 ml of minimum Eagle's medium (Invitrogen). Conditioned medium was removed from the primary neuronal cultures and replaced with minimum Eagle's medium. One ml of transfection solution was added to each 35-mm well, and cultures were incubated at 37 °C for 1 h. The transfection media were then removed and replaced with conditioned media for the remainder of the experiment. The transfection efficiency with this method varied between 0.1% and 0.5%. Glutamate treatment was initiated 24 h after transfection.

Transfections in HT22 Cells—The cationic lipid reagent Lipofectamine 2000 (Invitrogen) was used to transiently transfect HT22 cells. For toxicity experiments, 2 µg of DNA was used per well of a 12-well tissue culture plate, in a DNA (µg)/Lipofectamine 2000 (µl) ratio of 1.5. Typically, 1 µg of mt-eYFP plasmid DNA and 1 µg of MKP3 plasmid DNA were used for these transfections. For the Elk-1-dependent luciferase expression experiments, ~8 µg of total plasmid DNA was used for each 60-mm plate (3 µg of pSG5, MKP3 WT, or MKP3 C293S; 4 µg of luciferase reporter plasmid pFR-Luc; 0.12 µg of Elk-1/GAL4 fusion expression plasmid pFC-Elk-1/GAL4 or the GAL4 DNA binding domain expression plasmid pFC-GAL4 DBD; and 1.2 µg of constitutive Renilla expression plasmid PRK-tk Renilla), with a DNA/Lipofectamine 2000 ratio of 1.5. DNA and Lipofectamine 2000 were diluted in Opti- MEM (Invitrogen) for 5 min and then combined to allow DNA complexes to form for 20 min. The transfection solution was added to HT22 cells in serum-free Dulbecco's modified Eagle's medium without antibiotics for 5 h, after which cells were returned to normal serum- and antibiotic-containing media (see "Transfections in Primary Cortical Neuronal Cultures"). Cells were harvested as described beginning 16 to 18 h after transfection, and luciferase/Renilla activity was measured (see "Elk-1/GAL4 Fusion Protein-dependent Luciferase Expression").

Elk-1/GAL4 Fusion Protein-dependent Luciferase Expression—Components of the PathDetect in vivo signal transduction pathway transreporting system (Stratagene) were used in order to monitor Elk-1-dependent gene expression in HT22 cells. Cells were transfected with an expression plasmid coding for an Elk-1/GAL4 fusion protein, a reporter plasmid containing the luciferase gene under a synthetic promoter containing five tandem GAL4 binding sites, and a MKP3 expression plasmid. In this system, luciferase activity is a measure of the extent of Elk-1 activation. HT22 cells were co-transfected with the Elk-1 fusion plasmid (pFA2-Elk-1) or its negative control containing only GAL4 (pFC2-DBD), the luciferase reporter plasmid (pFR-Luc), a constitutive expression plasmid coding for Renilla under control of the cytomegalovirus promoter (PRK-tk Renilla), and either the pSG5 empty vector, MKP3 WT, or MKP3 C293S plasmid. Sixteen hours after transfection, HT22 cells were left untreated or treated with glutamate for 7.5 h. Cells were then harvested and lysed in lysis buffer (50 mM Tris-Cl, pH 7.5, 2 mM EDTA, 100 mM NaCl, 1% Nonidet P-40, 100 µM Na3VO4, 100 µM NaF, and 2 mM DTT). Both luciferase activity and Renilla activity were measured with the Dual-Glo luciferase assay system (Promega, Madison, WI) using a luminometer (Wallac Victor3; PerkinElmer Life Sciences). All luciferase signals were normalized to Renilla within each sample.

Viability Measurements in Transfected Cells—Primary cortical neuronal cultures (DIV3) display cell body shrinkage, process retraction, and nuclear condensation at 18–20 h after treatment with 5 mM glutamate. After 24 h, cells begin to detach from the tissue culture plate. To assess cell viability in transfected neurons (i.e. mt-YFP-positive cells; see "Transfections in Primary Cortical Neuronal Cultures"), we treated cells with the DNA dye propidium iodide (PI), which is excluded from healthy, intact cells and only gains access into cells with a compromised plasma membrane. Eighteen hours after the initiation of all treatments, cultures were incubated for 10 min in media containing a final concentration of 6.25 µg/ml PI. Cells were observed under an inverted fluorescence microscope equipped with phase-contrast optics (x20 objective; Nikon Eclipse TE200), and mt-eYFP-positive, PI-labeled, and unlabeled cells were counted. The percentage of dual-labeled cells in each well was then calculated as a percentage of all transfected cells counted (~50 cells/well). Multiple wells were counted for each condition in at least three separate transfected cultures (total cell population of at least 250 per condition). Digital images were acquired by SimplePCI software (version 5.2.1; Compix Inc. Imaging Systems, Cranberry Township, PA) and formatted using Adobe Photoshop 7.0.1.

HT22 cells become PI-positive beginning 6–7 h after treatment with 5mM glutamate and begin to detach from the culture plate after 9 h. We thus assessed toxicity in transfected, mt-eYFP marker-positive cells at 7 h using the PI staining method as described above. Cells were observed under an inverted fluorescence microscope equipped with phase-contrast optics (x20 objective; Nikon Eclipse TE200), and mt-eYFP-positive, PI-labeled, and unlabeled cells were counted. The percentage of dual-labeled cells in each field was then calculated as a percentage of all transfected cells counted (~60 cells/field at x100), and multiple fields were counted for each condition in at least three separate transfected cultures (total cell population of at least 500 per condition). Digital images were acquired by SimplePCI software (version 5.2.1; Compix Inc. Imaging Systems) and formatted using Adobe Photoshop 7.0.1.

Indirect Immunofluorescence—HT22 cells were plated on 12-well tissue culture plates in normal media (see "Cell Line Culture") and then transfected with either the pSG5 parent plasmid, MKP3 WT, or MKP3 C293S expression plasmids for 16–18 h. Cells were then washed with phosphate-buffered saline (PBS) (137 mM NaCl, 2.7 mM KCl, 4.3 mM Na2HPO4, and 1.4 mM KH2PO4, pH 7.4), fixed with -20 °C methanol for 5 min, washed with PBS, permeabilized with PBS containing 0.1% Triton X-100 for 10 min, and then blocked for 1 h with PBS supplemented with 3% bovine serum albumin at room temperature. The fixed cells were then incubated with an anti-c-myc antibody (1:500; Cell Signaling, Beverly, MA) or an anti-GAL4 DBD antibody (1:500; Sigma) in PBS with 3% bovine serum albumin for 1 h at room temperature and washed three times with PBS before incubation with a fluorescencelabeled secondary antibody (1:1000; anti-mouse IgG conjugated to Alexa Fluor 488; Molecular Probes, Eugene, OR) and the nuclear stain Hoescht 33258 (1 µg/ml). After washing stained cells twice with PBS, images were viewed with a Nikon Eclipse TE200 inverted fluorescence microscope. Digital images were acquired by SimplePCI software (version 5.2.1; Compix Inc. Imaging Systems) and formatted using Adobe Photoshop 7.0.1.

ERK-directed Phosphatase Activity Assay—We modified a nonradioactive method for determining ERK-directed phosphatase activity in whole cell lysates (24). This method relies on detecting dephosphorylation of a purified, dual-phosphorylated, His6-tagged ERK upon incubation with whole cell lysate. Thus, alterations in ERK-directed phosphatase activity within the lysate can be monitored by measuring changes in the phosphorylation state of the isolated substrate, as assessed by Western blotting with a phospho-specific ERK1/2 antibody. In brief, 150 µg of whole cell lysate (see "Western Blot Analysis") lacking DTT, Na3VO4, or NaF was diluted into a total volume of 250 µl in phosphatase assay buffer (10 mM MgCl2, 10 mM Hepes, pH 7.5, and 10 µM of the MEK inhibitor U0126). Recombinant, phosphorylated His6-ERK2 (Biomol, Plymouth Meeting, PA) was added to each sample (30 ng/sample), and the reactions were maintained at 37 °C for 40 min. As a positive control for phosphatase activity, 1200 units of the dual specificity phosphatase {lambda} protein phosphatase ({lambda}-PPase; New England BioLabs, Beverly, MA) was included in a separate reaction. Treatment with 50 mM DTT or phosphatase inhibitors was accomplished during a 30-min incubation on ice, prior to the addition of purified ERK. When DTT and inhibitors were used, the samples were pre-incubated for 30 min on ice with DTT, followed by a 15-min incubation with inhibitors. After 40 min at 37 °C, the reactions were stopped by the addition of 250 µl of wash buffer (8 M urea, pH 8.6, containing 10 mM imidazole), and 30 µl of Ni2+-conjugated, magnetic beads (Qiagen, Valencia, CA) was added to each reaction. After 90 min of rocking at 4 °C, the samples were washed twice with wash buffer and once in 300 mM NaCl, 25 mM Tris, pH 7.5. The beads were then suspended in NaCl/Tris and Laemmli buffer, boiled for 5 min, loaded onto a 10% polyacrylamide gel, transferred to polyvinylidine fluoride membranes (Millipore, Bedford, MA), and subjected to Western blotting to detect phosphorylated ERK and total ERK.

JNK-directed Phosphatase Activity Assay—This method was performed exactly as described above, except that each sample was incubated with 30 ng of purified, dual-phosphorylated His6-tagged JNK-1 protein (Upstate, Waltham, MA), and Western blots were probed with anti-phospho-JNK and total JNK antibodies.

Western Blot Analysis—Cells were treated as described, scraped and collected into PBS, pelleted at 3000 rpm for 5 min, and resuspended in lysis buffer supplemented with 5 µl of protease inhibitor mixture (Sigma) per milliliter of lysis buffer. Total extract protein concentrations were determined using the Bio-Rad reagent. Equivalent amounts of total protein (either 15 or 20 µg) were separated by SDS-PAGE on 10% polyacrylamide gels and then transferred to polyvinylidine membranes (Millipore). Membranes were blocked with 5% dry milk in PBS/0.1% (v/v) Tween 20 (PBST). Membranes were then incubated with primary antibodies (anti-phospho-ERK, anti-total ERK, anti-phospho-JNK, anti-total JNK, or anti-c-myc; all from Cell Signaling) overnight at 4 °C with 3% dry milk or 5% bovine serum albumin, subjected to three 10-min washes with PBST, and exposed to the appropriate horseradish peroxidase-conjugated secondary antibody for 1 h at room temperature. Membranes were again subjected to three 10-min washes with PBST, and immunoreactive bands were detected by enhanced chemiluminescence (ECL; Amersham Biosciences) using standard x-ray film (Eastman Kodak Co., Rochester, NY). Several different exposure times were used for each blot to ensure linearity of band intensities. Densitometry was performed using a Personal Densitometer SI (Amersham Biosciences) linked to the ImageQuant 5.2 software (Amersham Biosciences).

Statistics—Comparisons of multiple mean values were accomplished by analysis of variance with either Tukey's or Bonferroni's post hoc tests for significance. Comparisons of two means were performed using a paired t test. p values of <0.05 were taken to be significant, and all data were analyzed using GraphPad Prism version 3.00 for Windows (GraphPad Software, San Diego CA).


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
ERK-directed Phosphatase Activity Is Specifically Inhibited during Glutamate-induced Oxidative Toxicity in Primary Immature Cortical Cultures—We have reported previously that in primary immature cortical cultures, ERK1/2 phosphorylation is elevated during oxidative stress generated by glutamate-induced glutathione depletion (6, 8). Given the sensitivity of some protein phosphatases to cellular redox state (15), we set out to examine whether ERK-directed phosphatase activity was altered during glutamate-induced oxidative toxicity. To this end, we adapted a novel in vitro phosphatase assay that uses whole cell lysates to dephosphorylate purified, dual-phosphorylated, His6-tagged ERK2 (Fig. 1A, lane 1; see "Experimental Procedures"). This assay revealed the robust ERK phosphatase activity of exogenously applied {lambda} protein phosphatase, a nonspecific dual specificity phosphatase that we employed as a positive control (Fig. 1A, lane 2). Furthermore, the method proved to be quite sensitive in detecting changes in phosphatase activity because the application of well-characterized phosphatase inhibitors was effective in inhibiting phosphatase activity in preparations of whole cell lysate (Fig. 2, A and B). Although not as active as purified {lambda} phosphatase, whole cell lysates prepared from primary cortical neurons possess robust ERK-directed phosphatase activity (Fig. 1A, lane 3). When extracts were prepared from cultures treated with glutamate for 14 h, a decrease in ERK-directed phosphatase activity was revealed (Fig. 1A, lane 6). Quantification of several independent experiments revealed a significant decrease in ERK phosphatase activity in cultures treated for 14 h with glutamate, as reflected in a 5-fold increase (p < 0.01) in the normalized intensity of the phosphorylated ERK2 band when compared with untreated cultures (Fig. 1B). Shorter treatments with glutamate (i.e. 1 and 6 h) did not significantly affect ERK-directed phosphatase activity in the primary neuron culture extracts (Fig. 1A, lanes 4 and 5). Thus, the observed inhibition of ERK-directed phosphatase activity in these extracts coincided with times of maximal oxidative stress, in which glutathione levels are depleted, and ERK phosphorylation is increased (5, 8). Interestingly, inhibition of ERK-directed phosphatase activity could be completely reversed by treatment of the extracts with 50 mM DTT (Fig. 1A, lane 7), implying that phosphatase function is impaired in oxidatively stressed cells due to cysteine oxidation. The restoration of ERK phosphatase activity in lysates from oxidatively stressed cells with DTT consistently yielded a level of activity greater than that found in untreated cells (e.g. compare lanes 3 and 7, Fig. 1A). Indeed, DTT treatment of lysates from primary cortical neuronal cultures not exposed to glutamate led to increased ERK-directed phosphatase activity (data not shown). Thus, a small pool of ERK phosphatases may be normally under tonic, reversible oxidative inhibition in primary neurons, despite intracellular conditions that are generally reducing.



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  		FIG. 1.
ERK-directed phosphatase activity is specifically inhibited during glutamate-induced oxidative toxicity in primary immature cortical cultures. A, primary immature cortical cultures were left untreated (lane 3) or treated with glutamate for 1, 6, or 14 h (lanes 4–6). Whole cell lysates from these samples were utilized in an in vitro ERK-directed phosphatase activity assay (see "Experimental Procedures"). ERK-directed phosphatase activity is significantly reduced during oxidative stress (i.e. 14 h after glutamate treatment; compare lanes 3 and 6), and this inhibition is completely reversible with 50 mM DTT (lane 7). B, the results of between four and eight independent experiments were quantified, revealing a significant increase in ERK-directed phosphatase inhibition during oxidative stress and a significant restoration of this activity with DTT. C, the effects of oxidative toxicity on MAPK-directed phosphatases are specific to ERK because JNK-directed phosphatase activity is not altered by glutamate treatment (n = 3). D, furthermore, endogenous JNK phosphorylation is not altered by glutamate treatment (n = 3). I, phosphatase assay input; {lambda}, {lambda} phage protein phosphatase (positive control); C, lysates from control cultures.

 



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  		FIG. 2.
ERK-directed phosphatase activity is inhibited by okadaic acid and orthovanadate. The nature and identity of the components of the basal ERK-directed phosphatase activity were probed using the phosphatase inhibitors okadaic acid, an inhibitor of PP2A, and sodium orthovanadate, a general inhibitor of tyrosine-directed and dual specificity phosphatases. Whole cell lysates from untreated primary immature cortical neurons were incubated with vehicle (A, lane 3), 0.1 µM (A, lane 4) or 1 µM okadaic acid (A, lane 5), or 1 mM orthovanadate (B, lane 4) for 30 min before use in the in vitro ERK-directed phosphatase activity assay. Both doses of okadaic acid were sufficient to inhibit activity (A), and the inhibition observed with orthovanadate (B) was quantified and found to be statistically significant (C). Because orthovanadate led to less inhibition of ERK-directed phosphatase activity than okadaic acid, it is likely that both tyrosinedirected and dual specificity phosphatases contribute to this activity, whereas PP2A is the major component. All experiments were conducted at least four times. *, p < 0.05; I, phosphatase assay input; {lambda}, {lambda} phage protein phosphatase (positive control); C, lysates from control cultures.

 
To determine the substrate specificity of the phosphatase activity affected by oxidative stress, we assayed the activity of JNK-directed phosphatases using purified His6-tagged dual-phosphorylated JNK1 protein in the same primary neuron lysates used to monitor ERK dephosphorylation. As shown in Fig. 1C, JNK1-directed phosphatase activity is not significantly altered during glutamate-induced oxidative toxicity. Furthermore, the addition of 50 mM DTT did not enhance JNK phosphatase activity in the lysates (Fig. 1C, compare lanes 3 and 7), unlike the restoration of ERK-directed phosphatase activity by DTT in the same lysates. However, these extracts do contain some active JNK phosphatases because phosphorylated JNK levels were reduced upon incubation with extracts as compared with input (Fig. 1C, compare lanes 1 and 3). To examine the phosphorylation state of endogenous JNK1 (p46) and JNK2/3 (p54) in the primary neurons, we performed Western blot analysis using a phospho-specific JNK antibody. As can be seen in Fig. 1D, JNK phosphorylation does not change during glutamate-induced oxidative toxicity, even during times of increased oxidative stress (i.e. 12–14 h after the addition of glutamate). This finding is consistent with the lack of regulation of JNK phosphatase activity by oxidative stress. Thus, oxidative stress in immature cortical neuron cultures specifically inhibits the activity of a subset of phosphatases that act upon specific mitogen-activated protein kinase (MAPK) members. Furthermore, the selectivity of oxidative inhibition of MAPK-directed phosphatases revealed in the in vitro assay correlates with the specificity of MAPK activation in oxidatively stressed neurons.

ERK-directed Phosphatase Activity Can Be Inhibited by Okadaic Acid and Orthovanadate—In order to ascertain the contribution of tyrosine/dual specificity versus serine/threonine-directed phosphatases to ERK dephosphorylation in primary neuron cultures, we utilized two inhibitors: okadaic acid (OA) and sodium orthovanadate. OA is a well-characterized serine/threonine-directed protein phosphatase inhibitor with a high degree of selectivity at low concentrations for PP2A. Furthermore, a major component of ERK phosphatase activity in neurons is contributed by PP2A (25). As shown in Fig. 2A, treatment with either 0.1 or 1.0 µM OA led to near complete inhibition of ERK-directed phosphatase activity in lysates from untreated control cultures (compare lanes 4 and 5 with lane 3). Thus, our in vitro assay confirms the expected contribution of PP2A to ERK dephosphorylation in neurons.

Sodium orthovanadate is a potent inhibitor of tyrosine and dual specificity phosphatases that does not affect serine/threonine-directed phosphatases. Currently, there are few compounds available that inhibit specific tyrosine or dual specificity phosphatases, yet orthovanadate remains a useful tool to delineate the contribution of these classes of phosphatases to the regulation of protein phosphorylation. To examine the role of tyrosine and dual specificity phosphatases in the observed phosphatase activity in untreated neuronal cultures, we treated lysates from these untreated cells with 1 mM Na3VO4. As shown in Fig. 2, B and C, orthovanadate significantly inhibited ERK-directed phosphatase activity in lysates from untreated cultures. Importantly, the degree of inhibition observed with orthovanadate was less than that observed with OA, further confirming the predominant role of PP2A in ERK-directed phosphatase activity in neurons. Nonetheless, tyrosine and/or dual specificity phosphatases appear to operate in primary neurons as ERK-directed phosphatases.

The Reversibly Oxidized Pool of ERK-directed Phosphatase Activity Can Be Inhibited by Okadaic Acid and Orthovanadate—Having established parameters for pharmacologic inhibition of ERK-directed phosphatases in vitro using lysates from control cultures, we next sought to reveal the nature of the phosphatase activity recovered by DTT treatment of extracts from oxidatively stressed cells. As shown in Fig. 3, 0.1 µM okadaic acid and 1 mM Na3VO4 were both capable of blocking ERK phosphatase activity restored by DTT in lysates from oxidatively stressed cells (compare lanes 6–8 with lane 5). Thus, the reversibly inhibited pool of ERK-directed phosphatases that we have identified in oxidatively stressed neuronal cultures is likely composed of PP2A and vanadate-sensitive activities.



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  		FIG. 3.
The reversibly oxidized pool of ERK-directed phosphatase activity is inhibited by okadaic acid and orthovanadate. Whole cell lysates were prepared from primary immature cortical neurons that had been left untreated (lane 3) or treated with glutamate for 14 h (lanes 4–8). The lysates from glutamate-treated cells were incubated with DTT alone (45 min; lane 5) or DTT (30 min) followed by a 15-min incubation with 0.1 (lane 6)or1 µM (lane 7) okadaic acid or 1 mM sodium orthovanadate (lane 8) before use in the in vitro ERK-directed phosphatase activity assay. The reversible pool of oxidized ERK-directed phosphatases is inhibited by okadaic acid and orthovanadate. Results shown are representative of four independent experiments. I, phosphatase assay input; {lambda}, {lambda} phage protein phosphatase (positive control); C, lysates from control cultures; 14, lysates from cultures treated with glutamate for 14 h.

 
Overexpression of MKP3 WT and MKP3 C293S Significantly Alters ERK Phosphorylation and Translocation in HT22 Cells—The extent of reversible inhibition of ERK-directed phosphatases generated by glutamate treatment of cells (Fig. 1A) is much less than that observed upon pharmacological inhibition of phosphatase activity in vitro (Fig. 2A). Thus, it is conceivable that sufficient overexpression of a specific phosphatase or associated factor could overcome the effects of oxidative stress on endogenous phosphatase activity and reduce ERK phosphorylation. The robust inhibitory effect of OA on ERK-directed phosphatase activity in primary neuron extracts highlights PP2A as a logical candidate for overexpression analysis. Although PP2A exists as a multimeric complex in vivo, expression of the catalytic subunit alone (i.e. C subunit) can be sufficient for activity (26). However, manipulation of PP2A activity in vivo upon catalytic subunit overexpression is complicated by the auto-inactivation of PP2A activity that results from C subunit overexpression (26, 27). Furthermore, PP2A has a wide range of substrates other than ERK1/2, making it difficult to ascribe biological effects of PP2A activity manipulation to effects on a particular substrate (i.e. ERK1/2).

We therefore decided to assess the effects on glutamate toxicity resulting from the overexpression of a single subunit ERK-directed phosphatase. MKP3 is a dual specificity phosphatase with a high degree of selectivity toward ERK1/2 that is predominantly localized in the cytoplasm (28). Thus, if sufficient overexpression of MKP3 can be obtained in transfected cells, some fraction may escape the damaging effects of glutamate-induced oxidative stress and act to dephosphorylate ERK1/2. Furthermore, a catalytically inactive mutant of MKP3 (i.e. C293S) is available that maintains its ability to bind to ERK1/2 (21) and therefore functions as a dominant negative to limit the access of endogenous ERK1/2 to endogenous phosphatases. Furthermore, the C293S mutant of MKP3 can be used to assess the impact of nuclear localization of active ERK1/2 because its overexpression leads to the trapping of active, phosphorylated ERK1/2 in the cytoplasm (21).

As expected from previous analyses of MKP3 compartmentalization, MKP3 WT and MKP3 C293S are clearly localized in the cytoplasm of transfected HT22 cells (Fig. 4, B-D). In HT22 cells that overexpressed transfected MKP3 WT, phospho-ERK levels were dramatically reduced (Fig. 4A, lane 2). The high degree of transfection efficiency that we obtain with the HT22 cells (i.e. ~80% or higher) allowed for such significant reductions of ERK phosphorylation to be detected in extracts prepared from the total pool of cells. Overexpression of the MKP3 C293S mutant in HT22 cells actually led to an enhancement of ERK phosphorylation that may be explained by the dominant negative effect of this mutant on basal MKP3 activity (Fig. 4A, lane 3). Given the ability of MKP3 overexpression to affect ERK phosphorylation, we examined their effects on glutamate-induced oxidative toxicity, which we have shown previously to be ERK-dependent (6, 8). In agreement with earlier studies from our group, glutamate treatment increased phospho-ERK levels in HT22 cells transfected with an empty plasmid (Fig. 4A, lanes 1 and 4). As observed in untreated HT22 cells, overexpression of MKP3 dramatically reduced ERK phosphorylation levels in oxidatively stressed cultures (Fig. 4A, lanes 2 and 5). Furthermore, MKP3 C293S overexpression was as effective as in untreated cultures in increasing phospho-ERK to levels even higher than those attained with glutamate treatment alone (Fig. 4A, lanes 3 and 6). Thus, MKP3 overexpression can be used to manipulate ERK phosphorylation in HT22 cells under either basal or oxidative stress conditions.



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  		FIG. 4.
Overexpression of MKP3 WT and MKP3 C293S significantly alters ERK phosphorylation and translocation in HT22 cells. HT22 cells transfected with either empty plasmid (pSG5), MKP3 WT, or MKP3 C293S were left untreated or treated with 5 mM glutamate for 7.5 h, and whole cell lysates were prepared. Western blots from these cells demonstrate glutamate-dependent ERK activation in empty vector-transfected cells (A, lanes 1 and 4), whereas MKP3 WT abrogated ERK activation independently of glutamate treatment (A, lanes 2 and 5), and MKP3 C293S markedly accentuated ERK activation independently of glutamate treatment (A, lanes 3 and 6). Immunocytochemistry against the myc epitope of the transfected myc-tagged MKP3 WT and MKP3 C293S demonstrated a lack of staining in empty plasmid-transfected HT22 cells (B) and a clearly cytoplasmic pattern of staining in cells transfected with the MKP3 WT (C) and MKP3 C293S (D) expression plasmids. HT22 cells were transfected with an Elk-1/GAL4 fusion expression plasmid, a luciferase reporter plasmid, a Renilla expression plasmid, and either empty plasmid or MKP3 WT or MKP3 C293S expression plasmid. Glutamate-treated, empty plasmid-transfected cells demonstrated a significant increase in luciferase expression, whereas both MKP3 WT and MKP3 C293S overexpression, as well as U0126 treatment, abrogated this increase (E). *, p < 0.001 compared with pSG5 control; #, p < 0.05 compared with pSG5 control.

 
MKP3 C293S not only acts as a dominant negative to sequester ERK away from endogenous MKP3 but also restricts ERK to the cytoplasm via its selective, high-affinity interaction with ERK via its NH2-terminal domain (21, 29). Because we have previously proposed a role for nucleus-localized ERK in glutamate-induced oxidative toxicity (7), we sought to establish a sensitive, functional test for detecting active ERK in the nuclei of MKP3-transfected cells. Therefore, HT22 cells were transfected with MKP3 expression plasmids along with an expression vector for an Elk-1/GAL4 DBD fusion protein and a luciferase reporter gene under the control of a promoter containing five tandem GAL4 DNA binding sites. Given that Elk-1 is an established nuclear target of ERK, enhanced transactivation activity resulting from its phosphorylation by nuclear localized ERK is easily monitored through the activity of the luciferase reporter in this system.

As confirmation of the sensitivity of this system to the presence of activated ERK in the nucleus, luciferase activity was induced nearly 2-fold upon glutamate treatment of HT22 cells transfected with an empty vector (Fig. 4E). Treatment with the MEK inhibitor U0126 blocked the glutamate-induced increase in luciferase activity. As shown in Fig. 4E, MKP3 WT overexpression reduced luciferase expression below baseline in both untreated and glutamate-treated cells, as would be expected from the dramatic reduction in ERK phosphorylation (Fig. 4A, lanes 2 and 5). Interestingly, MKP3 C293S overexpression, which led to a significant increase in ERK phosphorylation (Fig. 4A, lanes 3 and 6), led to reduction in luciferase activity in both untreated and glutamate-treated cells (Fig. 4E). This result again confirms the sensitivity of the system to detect the activity of phosphorylated ERK within the nucleus and establishes the ability of MKP3 C293S to restrict nuclear translocation of active ERK in a neuronal cell line, consistent with previous work in fibroblast cell lines (21). Collectively, these data show that glutamate-induced oxidative toxicity in HT22 cells is indeed accompanied by a functional increase in ERK signaling in the nucleus.

Overexpression of MKP3 WT and MKP3 C293S Protects Both HT22 Cells and Primary Immature Cortical Neurons from Glutamate-induced Oxidative Toxicity—Given that MKP3 WT and MKP3 C293S overexpression was effective in manipulating both ERK activation and translocation, we sought to determine the impact of MKP3 WT and MKP3 C293S overexpression on glutamate-induced oxidative toxicity in both HT22 and primary immature cortical cell cultures. A single-cell toxicity assay was devised that relied on the dual transfection of a mitochondria-targeted enhanced yellow fluorescent protein expression plasmid (mt-eYFP) along with either an empty plasmid (pSG5) or MKP3 WT or MKP3 C293S expression plasmid. After 6–7 h of glutamate treatment, HT22 cells with a compromised plasma membrane become permeable to PI, a DNA dye. Toxicity can be assessed in transfected HT22 cells by determining the percentage of dual-labeled, eYFP+/PI+ cells in untreated and glutamate-treated groups. As can be seen in Fig. 5A, after transfection with the pSG5 empty plasmid, a significant proportion of HT22 cells were dual-labeled upon glutamate treatment. Co-transfection with either MKP3 WT (Fig. 5B) or MKP3 C293S (Fig. 5C) significantly blocked oxidative toxicity, as reflected by a decrease in the percentage of duallabeled cells compared with cultures transfected with empty vector. These results are quantified and summarized in Fig. 5D.



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  		FIG. 5.
Overexpression of both MKP3 WT and MKP C293S protects HT22 cells from glutamate-induced oxidative toxicity. HT22 were co-transfected with a mitochondrial-targeted eYFP expression plasmid and either an empty plasmid (pSG5) or MKP3 WT or MKP3 C293S expression plasmid. Toxicity was measured as the percentage of PI-positive cells within the transfected population in untreated and glutamate (5 mM)-treated cells. Photomicrographs (x20 objective) of untreated and treated empty plasmid (A)-, MKP3 WT (B)-, and MKP3 C293S (C)-transfected HT22 cells are shown. A summary of counts from three random fields from at least three independent transfections is shown (D). *, p < 0.01 when compared with pSG5 control.

 
This finding was extended to primary immature cortical neuronal cultures. These cultures were transfected with both the mt-eYFP marker and either pSG5, MKP3 WT, or MKP3 C293S expression plasmid. We confirmed that the mt-eYFP transfection marker was localized in neurons by immunocytochemical staining of transfected cultures with an antibody to the neuronal marker NeuN (data not shown). Cultures were treated with glutamate, and PI uptake in transfected cells was determined 18–20 h later (a time at which cells become PI-labeled, but before a significant number of cells have detached from the culture dish). An example of a dual-labeled cell in glutamate-treated cultures transfected with empty vector is seen in Fig. 6A. However, in cultures transfected with MKP3 WT (Fig. 6B) or MKP3 C293S (Fig. 6C), transfected cells were significantly spared from glutamate-induced oxidative toxicity, as reflected by a reduced proportion of dual-labeled cells within the transfected population. These results are quantified and summarized in Fig. 6D.



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  		FIG. 6.
Overexpression of both MKP3 WT and MKP C293S protects primary immature cortical HT22 cells from glutamate-induced oxidative toxicity. Primary immature cortical cells were co-transfected with a mitochondrial-targeted eYFP expression plasmid and either empty plasmid (pSG5) or MKP3 WT or MKP3 C293S expression plasmid. Toxicity was measured as the percentage of PI-positive cells within the transfected population in untreated and glutamate (5 mM)-treated cells. Photomicrographs (x20 objective) of untreated and treated empty plasmid (A)-, MKP3 WT (B)-, and MKP3 C293S (C)-transfected HT22 cells are shown. A summary of counts from three random fields from at least three independent transfections is shown (D). *, p < 0.001 when compared with untreated pSG5.

 
Unlike the HT22 cells, which can be transfected with high efficiency, primary neurons are inefficiently transfected, and therefore it is conceivable that neuroprotective effects of MKP3 apply to a distinct subpopulation of neurons. However, the comparison of MKP3 (i.e. WT and C293S mutant) effects is always made against cells co-transfected with empty vector (pSG5) and the mt-eYFP expression plasmid. Thus, expression of mt-eYFP in transfected cells is not sufficient to confer protection against glutamate toxicity, and the population of neurons that are transfectable (i.e. mt-eYFP positive) is sensitive to glutamate toxicity (i.e. exhibit positive PI staining). The results of the transfection experiments in primary immature cortical neurons and HT22 cells confirm that ERK activation is necessary for cell death during glutamate-induced oxidative toxicity. Furthermore, the fact that MKP3 C293S significantly protected both HT22 cells and primary neurons from cell death implies that activation of ERK1/2 alone is not sufficient to induce toxicity in response to glutathione depletion. Active ERK1/2 must be capable of translocating to the nucleus in order to trigger cell death in oxidatively stressed neurons.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
Although thiol oxidation and inactivation of phosphatases have been shown in numerous cellular systems to be a feature of normal cellular signaling events (15) and occur during ROS accumulation in neuronal cell lines (30), the role of this mechanism in oxidative stress-induced neurotoxicity has not been assessed previously. In this report, ERK-directed phosphatase activity was found to be specifically inhibited during times of oxidative stress, whereas phosphatases regulating other MAPKs, such as JNK1, were not affected. This inhibition of ERK-directed phosphatase activity is coincident with the temporal pattern of ERK activation that occurs during glutamate-induced oxidative toxicity; only at times in which inhibition of ERK-directed phosphatase activity is detected do we observe a concurrent increase in phosphorylated ERK1/2. These findings of substrate specificity and temporal congruence strongly indicate that oxidative inhibition of ERK-directed phosphatase activity is a mechanism that drives ERK activation and propels neurons toward a cell death pathway. Based upon inhibitor studies, PP2A and vanadate-sensitive phosphatases appear to be the likely phosphatases that mediate this effect.

Many studies have demonstrated p38 and JNK activation during oxidative stress in numerous cell types, including primary cortical neuronal cultures (31, 32). We do not observe JNK or p38 activation in either HT22 cells or primary immature cortical neuronal cultures during glutamate-induced oxidative toxicity. A potential reason for this disparity could be attributed to differences in the complement of MAPK-directed phosphatases that are expressed in various cellular systems or to the specificity to which certain phosphatases, even within one cell type, may be inhibited by oxidative stress. We have shown that JNK-directed phosphatases are not inhibited by oxidative stress in primary cortical cultures and that this correlates with the lack of increased JNK phosphorylation during oxidative stress. Thus JNK-directed MAPK phosphatases, such as M3/6, may not be as sensitive to oxidation as ERK-directed phosphatases, such as MKP3 or strial-enriched phosphatase. Further work clearly needs to be done to determine the differential sensitivities of various phosphatases to oxidation because this has a direct implication for potential mechanisms of oxidative signaling.

A number of mechanisms could account for oxidation-dependent phosphatase inhibition. Protein-tyrosine phosphatases exhibit sensitivity toward oxidation of the catalytic cysteine (33), and inactivation can occur by both reversible and irreversible mechanisms. Because both tyrosine-directed and dual specificity phosphatases share a canonical HC(X)5R motif in their active sites, it is likely that both classes undergo similar mechanisms of inhibition. Conversion of the active-site cysteine to a metastable sulfenic acid (Cys-SOH) (15) and the involvement of this cysteine in the formation of an intramolecular disulfide bond (33) are both reversible with thiol-reducing agents. A novel mechanism of reversible oxidative inhibition of protein-tyrosine phosphatase 1B was recently identified that involves the formation of a sulfenyl-amide stable intermediate resistant to further oxidation (34, 35). However, the extent to which this mechanism operates in other phosphatases is currently unknown. Progression of these reversible states to sulfinic (Cys-SO2H) or sulfonic (Cys-SO3H) acid forms leads to irreversible phosphatase inhibition. Because we observe total reversibility of oxidative phosphatase inhibition with the thiolreductant DTT, oxidation of ERK-directed phosphatases to sulfinic and sulfonic forms is unlikely to occur in the oxidatively stressed neuronal cultures that we analyzed. Furthermore, the fact that activity is recoverable with DTT rules out the possibility that decreased ERK-directed phosphatase activity during oxidative stress reflects decreased phosphatase expression (36) or increased phosphatase degradation (37). The restoration of ERK-directed phosphatase activity with DTT is likely to represent a direct effect on the phosphatases themselves. However, it is formally possible that DTT is reversing the thiol oxidation of cofactors necessary for ERK-directed phosphatase function. The identification of both the sites of oxidation and the mechanism of inhibition of specific ERK-directed phosphatases during oxidative stress will be the subject of future studies.

Although redox modification of cysteine residues plays an important role in protein phosphatase inhibition, other mechanisms are possible, such as nitrosylation or glutathionylation of residues within the protein or oxidation of Fe-Zn metal clusters that are critical for the function of serine/threonine-directed phosphatases. In fact, oxidative-inhibition of serine/threonine-directed phosphatases is not well understood and may reflect effects on oxidant-sensitive regulatory subunits. Nonetheless, oxidative stress-induced inhibition of phosphatases is likely to represent a common mechanistic thread that drives cellular signaling events during neuronal cell death in a diverse range of conditions, such as stroke, Alzheimer's disease, or Parkinson's disease. For example, the activity of the serine/theonine phosphatase calcineurin was recently shown to be reduced in lymphocytes of amyotrophic lateral sclerosis patients due to an effect thought to be mediated by oxidation (38). Indeed, it has been known for some time that PP2A activity is reduced in the cortices of Alzheimer's patients as compared with control patients (39), and ERK has been found to be activated in these tissues (40). Tau is a well-described ERK1/2 target (41), and the hyperphosphorylation of tau protein and the development of neurofibrillary tangles in Alzheimer's disease pathology could reflect aberrant ERK1/2 activity (41). Interestingly, mice expressing a dominant negative form of PP2A in neurons display features of Alzheimer's pathology (42). Thus, it is intriguing to postulate that oxidative stressmediated PP2A inhibition in Alzheimer's disease may account for enhanced ERK1/2 activity and subsequent tau hyperphosphorylation and neurofibrillary tangle formation.

Because the endogenous pool of PP2A and vanadate-sensitive phosphatases is inactivated during oxidative toxicity in primary neurons, we sought to overexpress one of these phosphatases to attempt to overcome this inhibition and impact ERK activation. The choice of MKP3 as a tool to manipulate ERK activity in our system was made for several reasons. MKP3 is localized in the cytoplasm and physically interacts with ERK1/2 at a kinase-interacting motif domain in its NH2 terminus in a highly specific manner. Brunet et al. (21) elegantly showed that the overexpression of MKP3 WT could extinguish ERK1/2 signaling, whereas overexpression of the catalytic mutant MKP3 C293S could abolish ERK1/2 signaling specifically in the nucleus by acting to trap activated ERK1/2 in the cytoplasm via its physical interaction with the kinase-interacting motif domain of the MKP3 C293S protein. Interestingly, MKP3 C293S was not shown to impair ERK1/2 functioning in the cytoplasm (21).

Shuttling of phospho-ERK into and out of the nucleus is a dynamic process, and standard, static immunocytochemical fluorescence imaging techniques may not be sufficient to detect small shifts in the equilibrium of the subcellular distribution of phospho-ERK. However, these small changes in the distribution of phospho-ERK could have important biological consequences. We therefore employed a sensitive, quantifiable, and functional assay to assess nuclear residency of active ERK during glutamate-induced oxidative toxicity. In this assay, luciferase expression was controlled by the activation of an Elk-1-GAL4 fusion by active ERK in the nucleus. We found that Elk-1/Gal4-directed luciferase expression increases with oxidative stress in HT22 cells and that both MKP3 WT and MKP3 C293S block this increase. Thus, glutamate-induced oxidative stress leads to an increase in active ERK function within the nucleus. Although it is predominantly localized within the cytoplasm, overexpressed wild-type MKP3 can deplete the total cellular pool of active ERK, most likely by acting upon dynamically shuttling ERK during its transit through the cytoplasm. Overexpression of the MKP3 C293S mutant exerts a dominant negative effect to block endogenous MKP3 and elevate ERK phosphorylation. Elevated phospho-ERK in this case is not accessible to the Elk-Gal4 fusion in the nucleus due to its sequestration with the cytoplasm by the MKP3 C293S mutant (21).

Despite the likelihood that multiple phosphatases from a diversity of classes are responsible for the regulation of ERK phosphorylation, MKP3 overexpression was sufficient to manipulate ERK activity in oxidatively stressed neurons and revealed consequences of ERK activation and localization on toxicity. Thus, inhibition of ERK activation was brought about by MKP3 overexpression and led to significant protection from oxidative toxicity in both HT22 cells and immature cortical neuronal cultures. Obviously, the overexpression was sufficient to overcome the oxidation-induced inhibition of endogenous ERK-directed phosphatases in the primary neurons. Elevation of ERK activation brought about by MKP3 C293S overexpression also was protective in both contexts, presumably based upon the sequestering of active ERK in the cytoplasm. Thus, we hypothesize that ERK-directed toxicity in oxidatively stressed neurons requires some translocation of active ERK to the nucleus. We are currently in pursuit of specific targets of ERK that may be mobilized, even in response to the transient nuclear accumulation of active ERK, to trigger neuronal cell death in response to oxidative stress.

ERK activation occurs downstream of ROS production in primary immature cortical cultures undergoing glutamate-induced oxidative toxicity. We have previously monitored ROS in primary immature cortical cultures, and ERK inhibition via U0126 administration did not alter the rate or extent of ROS production in these cells during glutamate-induced oxidative toxicity (8). We observe that ERK-directed phosphatases are still inhibited during glutamate-induced oxidative toxicity in the presence of U0126 (data not shown), a treatment that protects these cultures from cell death. This indicates that ERK-directed phosphatase inhibition is not sufficient for toxicity, in and of itself, but must act through specific activation of ERK. Furthermore, it also implies that there is a requirement for MEK activity to provide a forward drive for ERK activation during oxidative stress in primary immature cortical cultures (8).

The results reported here represent the first demonstration of the critical toxic role of ERK1/2 activity in the nucleus using a molecular approach rather than relying upon inhibitor studies, which cannot resolve the distinction between ERK1/2 activation as a whole versus the nuclear translocation of active ERK1/2. Thus, the spatio-temporal pattern of ERK activation during glutamate-induced oxidative toxicity is critical for determining the cellular response to this insult, and nuclear targets of ERK must be mobilized to bring about oxidative toxicity in neurons. We also have implicated an intriguing mechanism by which oxidative stress may be coupled to ERK activation. Specific oxidation of ERK-directed phosphatases during glutamate-induced oxidative toxicity could drive ERK activation and nuclear accumulation, leading to neuronal cell death. We are now fully engaged in determining the specific phosphatases that are involved in mediating this effect and the mechanism by which they are inhibited by oxidative stress.


   FOOTNOTES
 
* This work was supported by National Institutes of Health Grants F30 NS43824 (to D. J. L.) and R01 NS38319 (to D. B. D.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

To whom correspondence should be addressed: Dept. of Pharmacology, University of Pittsburgh School of Medicine, E1352 BST, Pittsburgh, PA 15261. Tel.: 412-624-4259; Fax: 412-648-1945; E-mail: dod1{at}pitt.edu.

1 The abbreviations used are: ROS, reactive oxygen species; ERK, extracellular signal-regulated kinase; eYFP, enhanced yellow fluorescent protein; JNK, c-jun NH2-terminal kinase; MKP, mitogen-activated protein kinase phosphatase; mt-eYFP, mitrochondria-targeted eYFP; OA, okadaic acid; PP2A, protein phosphatase 2A; WT, wild type; DTT, dithiothreitol; PI, propidium iodide; PBS, phosphate-buffered saline; MEK, mitogen-activated protein kinase/extracellular signal-regulated kinase kinase; MAPK, mitogen-activated protein kinase. Back


   ACKNOWLEDGMENTS
 
We thank Dr. Ian Reynolds for the mt-eYFP expression plasmid; Drs. Steven Keyse and Anne Brunet for the expression plasmids pSG5, MKP3 WT, and MKP3 C293S; Dr. Elias Aizenman for PathDetect component plasmids; Dr. Floh Thiels for okadaic acid; Dr. Nevin Lambert for samples of polyethyleneimine and the protocol for neuronal transfections; and Julie Brick for assistance with the preparation of the manuscript.



   REFERENCES
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 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

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