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Address correspondence and reprint requests to Karen L. O'Malley, PhD, Department of Anatomy and Neurobiology, Washington University School of Medicine, 660 S. Euclid, St Louis, MO 63110, USA. E-mail: email@example.com
Oxidative stress is a key player in a variety of neurodegenerative disorders including Parkinson's disease. Widely used as a parkinsonian mimetic, 6-hydroxydopamine (6-OHDA) generates reactive oxygen species (ROS) as well as coordinated changes in gene transcription associated with the unfolded protein response (UPR) and apoptosis. Whether 6-OHDA-induced UPR activation is dependent on ROS has not yet been determined. The present study used molecular indicators of oxidative stress to place 6-OHDA-generated ROS upstream of the appearance of UPR markers such as activating transcription factor 3 (ATF3) and phosphorylated stress-activated protein kinase (SAPK/JNK) signaling molecules. Antioxidants completely blocked 6-OHDA-mediated UPR activation and rescued cells from toxicity. Moreover, cytochrome c release from mitochondria was observed after the appearance of early UPR markers, suggesting that cellular stress pathways are responsible for its release. Mechanistically, the 6-OHDA-induced UPR was independent of intracellular calcium changes. Rather, evidence of protein oxidation was observed before the expression of UPR markers, suggesting that the rapid accumulation of damaged proteins triggered cell stress/UPR. Taken together, 6-OHDA-mediated cell death in dopaminergic cells proceeds via ROS-dependent UPR up-regulation which leads to an interaction with the intrinsic mitochondrial pathway and downstream caspase activation.
(phosphorylated) the α subunit of eukaryotic initiation factor 2
(phosphorylated) Stress-activated protein kinase
permeability transition pore
upregulated modulator of apoptosis
reactive oxygen species
sodium dodecyl sulfate
unfolded protein response
Oxidative stress, mitochondrial dysfunction and inflammation have long been implicated in Parkinson's disease (PD), the second most common neurological disorder (Eriksen et al. 2005). Aberrant protein degradation is also thought to be important based on the discovery of several genetic mutations linked to PD that share a common role in the ubiquitin–proteosome pathway (Hofer and Gasser 2004; Snyder and Wolozin 2004). The neurotransmitter dopamine itself has been implicated in PD pathology because it can be auto-oxidized, leading to an array of toxic reactive species including catecholquinones and their derivatives (Curtius et al. 1974; Fornstedt et al. 1986; Spencer et al. 1998; Stokes et al. 1999). Moreover, dopamine hydroxylation can generate 6-hydroxydopamine (6-OHDA), which is widely used to create animal models of PD (Blum et al. 2001). Although 6-OHDA itself forms superoxide, hydrogen peroxide and catecholquinones, the ensuing interactions of these toxic species with molecular or genetic pathways leading to dopaminergic cell death are not well defined.
Recent genomic studies and bioinformatic approaches have shown that 6-OHDA induces the up-regulation of genes involved in cell stress responses (Ryu et al. 2002, Ryu et al. 2005; Holtz and O'Malley 2003; Holtz et al. 2005). These data, combined with the discovery that mutations in the PD-related gene, Parkin, are also associated with endoplasmic reticulum (ER)-mediated unfolded protein response (UPR; Takahashi and Imai 2003), suggest that stress-activated signaling pathways constitute a common mechanism by which both genetic and environmental factors might interact in a disorder such as PD. As one manifestation of ER stress, the UPR regulates both protein translation and gene transcription in response to perturbations affecting protein folding. Thus UPR induction is always protective to begin with. If, however, adjusting these two processes fails to remedy the stress situation, apoptosis may be initiated (Schroder and Kaufman 2005).
Apoptosis is controlled by a conserved set of Bcl-2 family proteins, the balance of which can lead to cell death or survival. The ER is now being recognized as an organelle intimately involved in this cascade, although the signaling machinery associated with the process is still uncertain. Emerging evidence suggests that loss of calcium from the ER and concomitant uptake by mitochondria can trigger a chain of events leading to the collapse of the mitochondrial membrane potential and culminating in apoptosis (Nutt et al. 2002; Scorrano et al. 2003). The exact mechanism by which calcium is transferred from one organelle to another is not known but appears to require various pro-apoptotic proteins such as Bax, Bak and Bid as well as the inositol-1,4,5-trisphosphate (IP3) receptor (Nutt et al. 2002; Boehning et al. 2003; Darios et al. 2003; Oakes et al. 2005). ER stress may also lead to cell death via pathways involving the oxidation and reduction of disulfide bonds, which result in the accumulation of ROS (Tu and Weissman 2004; Harding et al. 2003; Haynes et al. 2004), reduced mitochondrial activity and ensuing cell death. Additionally, prolonged ER stress can lead to caspase activation via mitochondrial-dependent and -independent pathways. Thus in rodents, at least, ER stress induces caspase 12 cleavage (Nakagawa and Yuan 2000; Nakagawa et al. 2000). Activated caspase 12 can either bypass the mitochondria and directly cleave caspase 9 or it may potentially interact with other pro-apoptotic proteins such as Bap31, leading to mitochondrial fission and loss of cytochrome c. Which of these complex scenarios predominates following a particular insult probably depends upon both the cell type and the particular cell stressor involved.
Reactive oxygen species (ROS) induction has also been clearly linked to apoptosis via complex signaling pathways (Matsuzawa and Ichijo 2005) or direct alteration of key mitochondrial proteins (Le Bras et al. 2005). In particular, proteins associated with the mitochondrial permeability transition pore (PTP) such as the adenine nucleotide translocator, creatine kinase and voltage-dependent anion channel can be directly targeted by superoxide and hydrogen peroxide (Le Bras et al. 2005). Thus changes in the cellular redox state via ROS generation can trigger intricate signaling systems associated with both the ER as well as the mitochondria that might act in parallel and/or sequentially to orchestrate cell death.
Like other oxidants, 6-OHDA has also been shown to cause apoptotic cell death as defined by caspase activation in cellular models of dopaminergic cell death (Choi et al. 1999; Lotharius et al. 1999; von Coelln et al. 2001). In contrast to canonical mitochondrial-mediated apoptosis, however, overexpression of the anti-apoptotic protein Bcl-2 is not protective against 6-OHDA, nor is knocking out the pro-apoptotic protein Bax (Oh et al. 1995, 1998; O'Malley et al. 2003). Finally, the ‘extrinsic’ death receptor pathway does not seem to be involved either (Holtz and O'Malley 2003; Choi et al. 2004). Thus many questions remain as to how signaling pathways coupled to intrinsic cell death processes and ER stress interact in models of dopaminergic cell death.
Here we show that 6-OHDA-generated ROS are evident several hours before the appearance of UPR markers, and that antioxidants robustly block this process. ROS formation is rapidly followed by the appearance of carbonylated proteins suggesting that protein oxidation is the trigger of ER stress, as opposed to a disruption of calcium homeostasis. Release of cytochrome c from the mitochondria is observed late in relation to UPR-associated events, strengthening the observation that death induced by parkinsonian mimetics in dopaminergic cell types involves cross-talk between ER stress mechanisms and the mitochondria.
Materials and methods
Cells from the murine mesencephalic cell line, MN9D, were plated on dishes coated with 0.5 mg/mL poly-d-lysine (Sigma, St Louis, MO, USA) for 1 h at 37°C and then rinsed with sterile H2O. Culture medium contained Iscove's Dulbecco's modified Eagle's medium (DMEM) with 10% fetal calf serum. MN9D cells were maintained in a 37°C incubator with 10% CO2. Before addition of experimental reagents, the medium was switched to serum-free Iscove's DMEM/F-12 supplemented with 1 × B27 (Invitrogen, Carlsbad, CA, USA).
For primary cultures, the ventral mesencephalon was removed from embryonic day 14 CF1 murine embryos (Charles River Laboratories, Wilmington, MA, USA) as described previously (Lotharius et al. 1999). Animals were treated in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals. All procedures were approved by the Washington University School of Medicine animal experimentation committee. Cells were maintained in serum-free Neurobasal medium (Invitrogen) supplemented with 1 × B27 supplement (Invitrogen), 0.5 mm l-glutamine (Sigma), and 0.01 µg/mL streptomycin plus 100 U penicillin. Half of the culture medium was replaced with fresh Neurobasal medium on the fifth day, and cells were treated after 6 or 7 days in vitro.
Measurement of ROS
2′,7′-Dichlorodihydrofluorescein (DCF) and dihydroethidium (DHE) were purchased from Molecular Probes (Eugene, OR, USA). For DCF, MN9D cells were plated at a density of 400 000 cells/35-mm culture dish 2 days before experiments. Cells were washed twice with serum-free Iscove's DMEM/F-12 medium, and incubated with 10 µm DCF/0.001 Pluronic F-127 (Molecular Probes) in serum-free Iscove's DMEM/F-21 without B27 supplement for 45 min at room temperature (25°C). Cells were washed twice with Hank's balanced salt solution (HBBS) medium without phenol red indicator and then imaged in real time using a laser scanning confocal microscope at 60-s intervals. A baseline level was established for 5 min, and then either 1–10 mm H2O2 or 100–500 µm 6-OHDA was added to the culture dish and measurements were taken up to 20 min. The average relative change in fluorescence (ΔF/F0) was obtained using a computerized image analysis program (Metamorph, Molecular Devices, Sunnyvale, CA, USA).
Superoxide formation in MN9D cells was detected via the oxidation of DHE. Cells were plated as described for DCF experiments. Cells were washed twice with serum-free Iscove's DMEM/F-12 medium, and incubated with 10 µg/mL DHE (dissolved in dimethylsulfoxide) in serum-free Iscove's DMEM/F-21 without B27 supplement for 15 min at 37°C. 6-OHDA (100 µm) was added and the cells were incubated for an additional 0.5, 1, 2 and 3 h. DHE was removed by washing twice with HBBS without phenol red, and DHE fluorescence was recorded by confocal microscopy. The average change in DHE fluorescence was calculated using Metamorph image analysis software.
Western blot Analysis
MN9D cells were plated at a density of 200 000 cells/well in six-well plates. After 3 days, cells were treated with 75 µm 6-OHDA in the presence or absence of 5 mmN-acetylcysteine (NAC; Sigma) or 2.5 µm cyclosporin A. MN9D cells were also treated with 2 µg/mL tunicamycin in the presence or absence of NAC. Primary dopaminergic neurons were plated at a density of 1.5 × 106 cells/well in a 24-well plate, and treated on the seventh day in vitro with 30 µm 6-OHDA. Following the appropriate incubation time, cells were washed once with phosphate-buffered saline (PBS), harvested in ice-cold radioimmune precipitation assay buffer [150 mm NaCl, 1% Nonidet P-40, 0.5% sodium deoxycholate (NaDoc, 0.1% sodium dodecyl sulfate (SDS), 50 mm Tris, pH 8.0] together with protease inhibitors (Roche, Mannheim, Germany), and incubated on ice for 20 min. Lysates were centrifuged to remove insoluble cell debris, and the cell lysate protein concentration was determined with the Bio-Rad bicinchoninic protein (BCA) assay (Bio-Rad, Hercules, CA, USA). Equal amounts of protein were run on SDS–polyacrylamide (PAGE) gels, and then transferred to polyvinylidene difluoride membranes (Bio-Rad). Rabbit polyclonal antibodies against phosphorylated stress-activated protein kinase/c-Jun N-terminal kinase (APK/JNK), phospho-phosphorylated α subunit of eukaryotic initiation factor 2 (eIF2α), phospho-c-jun, cleaved caspase 3 (all 1 : 1000) and cleaved caspase 9 (1 : 500) were purchased from Cell Signaling Technologies (Beverly, MA, USA). Rabbit polyclonal anti-activating transcription factor 3 (ATF3) (1 : 200), goat polyclonal anti-heat-shock protein (Hsp)60 (1 : 500), and mouse monoclonal anti-C/EBP homologous protein (CHOP)/Gadd153 (1 : 100) and anti-ubiquitin (1 : 750) were purchased from Santa Cruz Biotechnologies (Santa Cruz, CA, USA). Rabbit polyclonal anti-heme oxygenase-1 (HO-1; 1 : 5000) was purchased from Stressgen (San Diego, CA, USA), mouse monoclonal anti-cytochrome c (1 : 250) was purchased from BD Pharmingen (San Jose, CA, USA), rabbit polyclonal anti-Bok (1 : 250) was from Abgent (San Diego, CA, USA), anti-Bak (1 : 1000) and anti-Bax (1 : 500) were from Upstate (Lake Placid, NY, USA), and anti-cyclo-oxygenase (COX) 1 (1 : 5000) was purchased from Molecular Probes. Following primary antibody incubation, membranes were incubated in appropriate horseradish peroxidase-conjugated secondary antibodies (anti-mouse 1 : 5000, Sigma; anti-goat 1 : 5000, Jackson Immunoresearch, West Grove, PA, USA; anti-rabbit 1 : 2000, Cell Signaling Technologies). Specific protein bands (or polyubiquitinated proteins) were detected by enhanced chemiluminescence substrate detection (ECL Plus; Amersham Biosciences, Piscataway, NJ, USA). Quantitative fluoroimaging analysis was performed to determine increases in polyubiquitinated proteins.
Primary culture cells were plated at a density of 100 000 cells/35-mm microwell plate (1.25 × 103 cells/mm2; MatTek Corp., Ashland, MA, USA). Cells were treated with 30 µm 6-OHDA in the presence or absence of 5 mm NAC or 50 µm Mn(III) tetrakis (4-benzoic acid) porphyrin (MnTBAP; A.G. Scientific, Inc., San Diego, CA, USA). Cultures were fixed with 4% paraformaldehyde in PBS after 8, 12 or 18 h. Cultures were stained with mouse monoclonal anti-CHOP (1 : 300; Santa Cruz), rabbit polyclonal anti-cleaved caspase 3 (1 : 300; Cell Signaling), or co-stained with rabbit polyclonal anti-ATF3 (1 : 200; Santa Cruz) together with mouse monoclonal anti-tyrosine hydroxylase (TH) (1 : 1000; Immunostar, Hudson, WI, USA). Secondary antibodies conjugated with Cy3 (anti-mouse and anti-rabbit, 1 : 300) and Alexa488 (anti-mouse 1 : 500, anti-rabbit 1 : 2000) were then used.
Determination of cell viability
MN9D cells were plated at a density of 40 000 cells/well in 24-well plates and treated after 3 days. Cultures were treated with a range of 6-OHDA concentrations from 10 to 100 µm in the presence or absence of 5 mm NAC. After 24 h, cells were washed with fresh medium and switched to medium without 6-OHDA or NAC. Cell survival was then assessed using the 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) reduction assay as described previously (Oh et al. 1995).
The subcellular fractionation protocol was adapted from Boehning et al. (2003). MN9D cells were plated at a density of 2.5 × 106 cells per P150 dish. After 3 days, cultures were treated with 75 µm 6-OHDA in the presence or absence of 5 mm NAC. Cells were washed twice with ice-cold PBS and harvested with a cell scraper followed by centrifugation. Cells from two P150 dishes were pooled together for each condition. Cellular pellets were resuspended in 10 volumes of a hypotonic buffer containing 10 mm NaCl, 1.5 mm MgCl2, 10 mm Tris-HCl (pH 7.5) and protease inhibitor mixture (Roche). Cells were incubated on ice for 10 min to swell, and then homogenized in a Wheaton glass homogenizer using 10 strokes with a ‘B’ pestle. Iso-osmotic buffer (2.5 × concentration containing 525 mm mannitol, 175 mm sucrose, 12.5 mm Tris-HCl, pH 7.5, 2.5 mm EDTA with protease inhibitor mixture) was then immediately added to achieve isotonic conditions. The crude homogenate was spun at 1000 g for 15 min at 4°C. The pellet (P1) containing whole cells and nuclei was discarded, and the supernatant (S1) was centrifuged at 10 000 g for 15 min at 4°C to pellet the heavy membrane fraction containing mitochondria (P2). P2 was washed once by resuspending in 1 × iso-osmotic buffer, repelleted, and resuspended again in 1 × iso-osmotic buffer. The supernatant (S2a) was centrifuged again at 15 000 g to remove any contaminating mitochondria. The resulting pellet (P0) was discarded, and the supernatant (S2b) was centrifuged at 100 000 g for 1 h to separate the cytosolic fraction in the supernatant (S3) and the light membrane fraction containing ER membranes in the pellet (P3). P3 was resuspended in 1 × iso-osmotic buffer. S3 was concentrated using a microsep centrifuge device with a 3-kDa cut-off (Pall Corporation, Ann Arbor, MI, USA). Protein concentrations were determined using the Bio-Rad BCA protein assay. Western blotting was performed as described above.
Cytochrome c ELISA
MBL Cytochrome C ELISA kit (MBL International Corp., Watertown, MA, USA) was used to determine the concentration of cytochrome c in nanograms in mitochondrial and cytosolic fractions. The ELISA was repeated in triplicate (n = 3 per replicate) to calculate the average amount of cytochrome c in mitochondrial and cytosolic fractions, expressed as nanograms per million cells.
Oligomerization of Bak or Bok was assessed by chemical crosslinking. Cross-linker bis-maleimidohexane (BMH; Pierce, Rockford, IL, USA) was added to mitochondrial fractions (50 µg) at a final concentration of 5 mm at room temperature for 30 min. Reactions were terminated by adding 5 × SDS–PAGE sample loading buffer containing 10% SDS and 10%β-mercaptoethanol. Mitochondria were then analyzed by SDS–PAGE and immunoblotting using Bak or Bok antibodies.
Fluorescence measurements of intracellular calcium
MN9D cells were plated at a density of 400 000 cells/35-mm culture dish 2 days before experiments. Cells were washed twice with serum-free Iscove's DMEM/F-12 medium, and incubated with 10 µm Oregon Green O,O′-Bis(2-aminophenyl)ethyleneglycol-N,N,N′,N′-tetraacetic acid, tetraacetoxymethyl ester (BAPTA)-1 AM/0.001 Pluronic F-127 (Molecular Probes) in serum-free Iscove's DMEM/F-21 without B27 supplement for 1 h at 37°C. Cells were washed twice with serum-free Iscove's DMEM/F-12 medium and then imaged in real time using an Olympus Fluoview laser scanning confocal microscope. After establishing a baseline level, 1 mm H2O2, 100 µm 6-OHDA, 100 nm thapsigargin or 1 mm calcium ionophore A23187 (all reagents from Sigma) were added into the culture dish and measurements were taken up to 20 min. The average relative change in calcium measured as change in fluorescence over baseline (ΔF/F0) was obtained using a computerized image analysis program (Metamorph).
Detection of oxidative protein modification
Detection of protein carbonyl groups generated by oxidative stress was performed by dinitrophenylhydrazine (DNPH) derivatization using the OxyBlotTM Protein Oxidation Detection Kit (Chemicon International, Temecula, CA, USA) following the manufacturer's standard protocol. Briefly, equal amounts of MN9D protein lysates (see western blot analysis) were incubated in 6% SDS with DNPH solution for 20 min, at which time the reaction was neutralized with the neutralization solution supplied. Samples were either blotted on to nitrocellulose filters or separated on SDS–PAGE gels and then transferred to polyvinylidene difluoride membranes. Membranes were incubated with the supplied rabbit anti-dinitrophenylhydrazone (DNP) (1 : 150) primary antibody followed by horseradish peroxidase-conjugated goat anti-rabbit secondary (1 : 300). Bands were visualized by enhanced chemiluminescence substrate detection (ECL Plus; Amersham Biosciences). Slot-blotted proteins were quantitated using Molecular Dynamics (Sunnyvale, CA, USA) software.
Proteasome activity assay
Proteasome activity was measured using a microtiter plate assay essentially as described by Snider et al. (2002). Protein concentrations were determined using BCA (Pierce). Separate assays were conducted for N-succinyl-Leu-Leu-Val-Tyr-7-amido-4-methyl coumarin (LLVY-AMC), which measures chymotrypsin-like activity, t-butoxycarbonyl-Leu-Arg-Arg-7-amido-4-methyl coumarin (LRR-AMC) (trypsin-like activity) and benzyloxycarbonyl-Leu-Leu-Glu-aminomethylcoumarin (LLE-AMC) (peptidyl-glutamyl peptidase activity). The specific inhibitor epoxomicin (1 mm) was used to ensure specificity. Standard curves were constructed using 7-amino-4-methylcoumarin. All samples were assayed in quadruplicate using a CytoFluor II plate reader (Thermo Electron Corporation, Waltham, MA, USA, excitation 380, emission 440).
Descriptive statistics (mean ± SEM) of cell survival or treatment effects were calculated with statistical software (GraphPad Prism Software, San Diego, CA, USA) on data collected from a minimum of three independent experiments done in triplicate. The significance of effects between control and drug treatments was determined by one-way anova and post hoc Student's t-tests. If significant differences were observed then post hoc pairwise comparisons were performed for individual drug concentrations or time points.
6-OHDA causes rapid generation of ROS in dopaminergic cells
In separate studies we have shown previously that 6-OHDA rapidly induces ROS (Lotharius et al. 1999) as well as triggering the ER stress response and UPR in primary dopaminergic neurons (Holtz and O'Malley 2003). Because identical responses were also observed in the CNS-derived dopaminergic cell line, MN9D, we utilized this unique resource to test the hypothesis that ROS generation is upstream of 6-OHDA-mediated UPRs. Specifically, we used a variety of reagents and techniques to determine the nature and timing of the creation of 6-OHDA-generated free radicals. Overall ROS production in situ was determined using the cell-permeable indicator DCF together with real-time confocal imaging of single cells following H2O2 or 6-OHDA addition (Fig. 1a). Application of either toxin led to rapid changes in DCF fluorescence intensity that plateaued in about 2–3 min. These effects could be blocked by the thiol antioxidant NAC as well as the superoxide dismutase mimetic MnTBAP (Fig. 1b). Neither of these reagents by themselves led to ROS production (Fig. 1b). Inasmuch as DCF detects various oxidizing species including superoxide anion, H2O2 and nitric oxide, any or all of these types of radicals might be present. The indicator DHE reacts specifically with superoxide anion to generate ethidium (Bindokas et al. 1996), which can be detected by its stable intercalation into DNA. Figure 1(c) shows robust DHE accumulation following 1 h of exposure to 75 µm 6-OHDA. DHE induction was completely blocked by NAC or MnTBAP (Fig. 1d). These results are consistent with those previously observed for 6-OHDA treatment of primary mesencephalic cultures (Lotharius et al. 1999). It should be noted that in a previous study using MN9D cells (Choi et al. 1999), a lower dose of 6-OHDA did not appear to induce changes in DCF intensity whereas DHE was also increased 3–4 fold, albeit with a slower time course. Taken together, these results demonstrate that in MN9D cells as well as primary cultures (Lotharius et al. 1999) 6-OHDA rapidly generates free radical species that include but are not limited to superoxide anions. This early increase in oxidative species precedes the appearance of markers of UPR and/or apoptosis that have been observed in the course of 6-OHDA-mediated cell death (Holtz and O'Malley 2003).
ROS generation precedes UPR induction and is blocked by antioxidants
To confirm and extend the observation that 6-OHDA induces ROS formation in a time frame preceding the appearance of UPR markers, the antioxidants NAC and MnTBAP were used to order and block 6-OHDA toxicity. As a rapid, albeit indirect measure of cell viability, an MTT assay was used to assess the ability of these antioxidants to block 6-OHDA toxicity. Although MTT more accurately assesses mitochondrial function, the dose–response curve shown in Fig. 2(a) is very similar to that obtained in previous studies using propidium iodide to measure cell viability in MN9D cells (Jensen et al. 2003). Here, 5 mm NAC completely blocked death of MN9D cells induced by up to 100 µm 6-OHDA (Fig. 2a). Moreover, 6-OHDA-induced ROS was necessary for the 6-OHDA-induced UPR because pretreatment with NAC robustly blocked signs of stress including phosphorylation of eIF2α and increased ATF3 at 3 and 6 h (Fig. 2b). NAC also blocked phosphorylation of c-jun and SAPK/JNK, as well as increases in CHOP and HO-1 levels at 9 and 12 h (Fig. 2c). In addition, NAC blocked the downstream activation of caspase 3 and caspase 9 (Fig. 2c). To demonstrate that NAC blocks 6-OHDA-induced UPR markers via antioxidant properties and not by some alternative mechanism, ER stress was induced in MN9D cells with tunicamycin (Fig. 2d). This drug leads to ER stress through inhibition of proper protein glycosylation. Pretreatment with NAC was unable to block tunicamycin-induced up-regulation of CHOP, which occurs independently of ROS generation.
Although NAC has been used widely as an antioxidant, it can also inhibit many cellular pathways via its reducing activity (Zafarullah et al. 2003). To test whether another antioxidant could also block indices of UPR induction, we performed similar experiments using MnTBAP (Fig. 3). Like NAC, MnTBAP blocked 6-OHDA-generated ROS (Fig. 1), ATF3 activation, c-jun phosphorylation, CHOP induction, caspase 3 activation and cell death (Fig. 3). Taken together, these data demonstrate that 6-OHDA induces coordinated up-regulation of UPR genes through the generation of oxidative stress.
Earlier findings demonstrated that antioxidants such as fullerenes completely protected primary dopaminergic cells from cell death (Lotharius et al. 1999). Immunostaining similarly treated cultures revealed that NAC antioxidant treatment blocked 6-OHDA-mediated nuclear ATF3 and CHOP accumulation and caspase 3 activation, whereas by itself it was not toxic (Figs 4a and b, and not shown). MnTBAP has also been shown to be protective against 6-OHDA (Choi et al. 2004). Pretreatment of primary midbrain cultures with MnTBAP also prevented phosphorylation of c-jun, CHOP activation and caspase 3 up-regulation (Figs 4c–e). These results as well as our previous studies (Lotharius et al. 1999; Holtz and O'Malley 2003) not only demonstrate the similarity between MN9D cells and primary dopaminergic neurons, but also show that the prevention of 6-OHDA-induced UPRs via ROS attenuation is not unique to thiol antioxidants such as NAC.
Mitochondrial events are downstream of UPR up-regulation
Cell death induced by 6-OHDA leads to caspase activation although the pro-apoptotic protein Bax is not required in this system (O'Malley et al. 2003). Thus, an important question is whether the intrinsic mitochondrial apoptotic pathway plays a parallel or sequential role to UPR in 6-OHDA-mediated cell death and what protein and/or process triggers this response. Because cytochrome c release from the mitochondria precedes caspase activation and serves as a check point for mitochondrial involvement (Danial and Korsmeyer 2004), the timing of this release in relation to UPR activation was used as an aid to determining the role of the mitochondrial pathway. Subcellular fractionation of cells treated for 3, 6, 9 and 12 h with 6-OHDA along with vehicle-treated controls revealed that changes in cytochrome c distribution were still not apparent after 6 h (Fig. 5a), a time at which UPR markers were robustly up-regulated (Fig. 2; Holtz and O'Malley 2003). Loss of cytochrome c from the mitochondrial fraction did not become apparent until 9–12 h after drug treatment (Fig. 5a). Blotting membranes encompassing the appropriate molecular size range for the mitochondrially expressed COX 1 or cytoplasmic lactate dehydrogenase (LDH) confirmed the compartmental designations (Fig. 5a). These data are consistent with independent results of an experiment examining cytochrome c release using an ELISA assay (Fig. 5b). In either case, NAC pretreatment completely blocked the loss of cytochrome c (not shown).
In support of our earlier studies (O'Malley et al. 2003), Bax was not translocated from cytoplasmic to mitochondrial compartments over the time course of this experiment (Fig. 5a). We also examined changes in the distribution of the structurally related pro-apoptotic proteins, Bak or Bok/Mtd, to determine whether either of these proteins could substitute for Bax. In MN9D cells both Bak and Bok were primarily localized to the heavy membrane, mitochondrial fraction (Fig. 5a). In the presence of cross-linking agents, high molecular weight Bak oligomers were observed following 9, 12 and 24 h of 6-OHDA treatment which were not present in untreated or NAC-treated controls (Fig. 5c). No evidence of oligomerization was observed for Bok (Fig. 5c) or Bax (not shown). These data, together with the finding that activation of caspases 3 and 9 is a late event occurring 9–12 h after 6-OHDA addition (Figs 2–4; Choi et al. 2004), suggest that mitochondrial changes occur either in parallel or subsequent to UPR-associated events.
Cytochrome c can also be released from mitochondria via the non-selective PTP, which may be opened by oxidative stress and blocked by cyclosporin A (Danial and Korsmeyer 2004; Le Bras et al. 2005). To test whether inhibition of the PTP prevents the up-regulation of the UPR marker CHOP, cells were pretreated with various concentrations of cyclosporin A with and without 6-OHDA. Cyclosporin A did not block CHOP or caspase 3 activation at any concentration tested in MN9D cells, whereas NAC did (Fig. 5d and not shown). Further, treatment with cyclosporin A did not prevent cytochrome c loss (not shown) or the activation of caspase 3 in primary cultures (Fig. 5e). Collectively, these data demonstrate that activation of the PTP is not the mechanism by which cytochrome c is released. Rather, induction of UPR precedes and possibly orchestrates cytochrome c release via a Bax and Bok-independent process.
6-OHDA does not cause rapid changes in intracellular calcium
Because maintenance of ER calcium is required for proper protein folding, one mechanism by which ROS are proposed to lead to ER stress is via disruption of cellular calcium homeostasis. To test whether 6-OHDA treatment leads to changes in intracellular calcium, MN9D cells were loaded with the calcium indicator dye Oregon Green whose spatio-temporal distribution was analyzed by confocal microscopy. Addition of the calcium ionophore A23187 caused an immediate increase in intracellular calcium (Fig. 6). Thapsigargin, a potent inhibitor of calcium re-uptake into the ER, and H2O2 resulted in a rapid increase in calcium (Fig. 6). As thapsigargin is commonly used to induce ER stress, these results are consistent with the proposed mechanism of ROS-induced calcium release. In contrast, addition of 6-OHDA at concentrations known to kill > 70% of the cells did not result in measurable increases in intracellular calcium (Fig. 6). Thus, these experiments demonstrate that, although 6-OHDA rapidly induces ROS formation, increased free radicals are not correlated with observable changes in intracellular calcium.
6-OHDA causes oxidative modification of proteins
Unchecked generation of ROS can result in oxidative damage to proteins which can impair proper protein degradation and lead to UPR up-regulation (Jenner 2003). A common indicator of oxidative damage is carbonylation (Nystrom 2005). After only 30 min of exposure to 6-OHDA, significant increases in protein carbonylation were detected in MN9D lysates, which could be blocked by NAC pretreatment (Fig. 7a). Size fractionation revealed at least six bands ranging from 42 to 123 kDa as specific ROS targets (Fig. 7b). Interestingly, most of these targets were more pronounced at all time points following 6-OHDA compared with H2O2 treatment. These results are consistent with a model in which 6-OHDA-generated ROS leads to protein oxidation; the ensuing accumulation of damaged proteins triggers a cell stress response.
6-OHDA induces a biphasic proteasome response and a late rise in polyubiquitinated proteins
Oxidized proteins may be polyubiquitinated and degraded via the proteasome system or may be directly sent to the proteosome (Shringarpure et al. 2003). To assess proteasome activity across the time course of UPR induction, cell lysates were prepared after 6-OHDA exposure and assayed for three known proteasome activities (trypsin-like, chymotrypsin-like and peptidyl-glutamyl peptidase activity). In keeping with the model postulating increased proteasome activity in the face of an increase in damaged proteins, proteasome activity increased over the first 6–9 h (Fig. 8a) before dropping to control levels as cells become committed to die (9 h; Holtz and O'Malley 2003).
Toxin-treated MN9D cells increased polyubiquitinated proteins in a time-dependent manner (Figs 8b and c). Quantification of band intensity across multiple experiments indicated a 1.5-fold increase over control in polyubiquitinated proteins at 9 and 12 h, which could be blocked by pretreatment with NAC. This time point coincided with the drop in proteasome activity (Fig. 8a) and the rise in caspase activity (Fig. 2c), suggesting that as cells begin to die the proteasome is no longer able to prevent the accumulation of damaged proteins that have been marked for disposal. Thus the 6-OHDA-induced ER stress response/UPR is not due to general impairment, but rather an overload, of the proteasome system.
The recent identification of genetic mutations linked to PD such as α-synuclein, parkin and ubiquitin C-terminal hydrolase L1 have highlighted the role of aberrant protein handling and degradation in this disorder (Snyder and Wolozin 2004; Eriksen et al. 2005). Previous results from this and other laboratories (e.g. Ryu et al. 2002, 2005) demonstrated that the parkinsonian mimetic 6-OHDA triggered ER stress resulting in UPR, thus providing a mechanistic link between toxicity and the identified PD mutations (Holtz and O'Malley 2003; Holtz et al. 2005). Although the toxicity of 6-OHDA is thought to stem from the production of ROS (Blum et al. 2001), whether ROS generation and coordinated ER stress induction are subsequent or parallel events has not been well established. Using molecular, biochemical and cellular techniques, the present study demonstrated that 6-OHDA-generated ROS precede the appearance of UPR and apoptotic markers all of which could be blocked by antioxidant exposure. Mechanistically, 6-OHDA-induced oxidative stress does not alter the spatio-temporal distribution of intracellular calcium. Instead, toxin exposure leads to the rapid accumulation of oxidized proteins preceding the appearance of UPR marker activation. Finally, UPR induction occurs upstream of mitochondrial events associated with control of apoptosis such as Bak oligomerization and cytochrome c release (Danial and Korsmeyer 2004). Thus, these studies demonstrate that 6-OHDA-generated ROS induces a UPR, which we hypothesize leads to apoptosis via downstream involvement of the mitochondrial pathway.
The difficulty in assessing cause and effect in post-mortem tissue from patients with PD makes assessing the role of a transient process such as UPR induction and/or apoptosis problematic. Recently, however, Silva et al. (2005) have shown that CHOP is up-regulated in a rodent model of dopaminergic cell death following intrastriatal injection of 6-OHDA. Moreover, CHOP knockout animals exhibited reduced apoptosis indicating that CHOP contributes to this process. Thus, these in vivo data (Silva et al. 2005) confirm and extend in vitro results such as the present findings as well as previous studies showing CHOP up-regulation following 6-OHDA (Ryu et al. 2002, 2005; Holtz and O'Malley 2003; Holtz et al. 2005).
6-OHDA-generated ROS precedes activation of the UPR
In a process that is dependent upon Bax, studies have shown that the quintessential oxidant, H2O2, releases ER Ca2+ which is then taken up by the mitochondria leading to cytochrome c release and apoptotic cell death (Hockenbery et al. 1993; Scorrano et al. 2003). Despite being rapidly oxidized into H2O2 among other products, 6-OHDA toxicity does not seem to be mediated by this component because it is independent of Bax (O'Malley et al. 2003) and does not release Ca2+ (Fig. 6). These results are consistent with a recent study showing that 6-OHDA-mediated cell death is due to quinone oxidation products rather than H2O2 (Izumi et al. 2005). Quinones are thought to react readily with nucleophiles such as cysteinyl residues in proteins (Graham et al. 1978). These in turn could inactivate critical proteins such as those involved in dopaminergic function (Hastings et al. 1996) and/or maintaining proper protein folding (Sitia and Molteni 2004).
Although ROS can damage DNA, lipids and proteins, their primary targets depend largely upon the cell type, proximity and severity of the stress (Gutteridge and Halliwell 2000). In dopaminergic systems, we and others have shown that an early consequence of 6-OHDA exposure is the induction of ER stress and UPR as exemplified by splicing of X box-binding protein 1 mRNA, increased levels of CHOP and the ER chaperone, GRP78/BiP (BiP), as well as phosphorylation of the ER-stress inducible kinase, PERK, the endoribonuclease inositol-requiring enzyme 1, IRE1 and eIF2α in 3–6 h (Ryu et al. 2002, 2005; Holtz and O'Malley 2003; Holtz et al. 2005). Temporally, these and even earlier markers of cell stress (ATF3, phosphorylated SAPK/JNK; Fig. 2) appeared well after ROS induction. Moreover, antioxidants such as NAC blocked toxin-induced cell death (Fig. 2a) and also inhibited ATF3 and CHOP up-regulation, phosphorylation of SAPK/JNK and c-jun, and downstream activation of caspases 3 and 9 (Figs 2b and c). Similarly, antioxidants rescued primary dopaminergic neurons from cell death (Lotharius et al. 1999) as well as the up-regulation of ATF3, CHOP and caspase 3 activation (Fig. 4). Together, these results link the generation of ROS to downstream UPR activation.
The mitochondrial intrinsic pathway is downstream of the UPR
Induction of ROS can trigger apoptosis via complex signaling pathways including direct effects on key mitochondrial proteins (Le Bras et al. 2005; Matsuzawa and Ichijo 2005). Hence 6-OHDA-generated ROS might induce the mitochondrial intrinsic pathway in parallel with ER stress via direct permeabilization of the PTP leading to cytochrome c release (Kowaltowski et al. 2001). This hypothesis seems unlikely, however, because cytochrome c release occurred long after both ROS generation and the up-regulation of UPR markers (Figs 2 and 4). Moreover, cyclosporin A, which is thought to block the PTP, did not prevent up-regulation of the UPR marker CHOP (Fig. 5d), cytochrome c release (not shown) or caspase 3 activation (Fig. 5e).
Many studies now indicate that ER stress can initiate apoptosis via mitochondrial-dependent and -independent processes (Breckenridge et al. 2003). For example, ER stress may induce changes in the ratios of Bcl-2 family members resulting in calcium release and/or the translocation of pro-apoptotic proteins to the mitochondria, leading to the release of cytochrome c (Momoi 2004). Alternatively, ROS alone may disrupt ER calcium homeostasis causing mitochondrial calcium overload, PTP opening and the mass exodus of cytochrome c (Nutt et al. 2002; Boehning et al. 2003; Oakes et al. 2005). Finally, the ER resident caspase 12 may directly cleave pro-caspase 9, bypassing the release of cytochrome c altogether (Rao et al. 2002). Of these possibilities it would appear that caspase 12-mediated apoptosis might be ruled out because neither caspase 12 transcripts nor proteins were detectable in cell extracts from MN9D or primary cells (not shown). Moreover, there was no evidence of calcium release from intracellular stores (Fig. 6) nor was cytochrome c re-distributed to the ER following toxin exposure (Fig. 5). Thus, although our observations are all consistent with the idea that cytochrome c release is dependent upon ER signals in this system, the exact mechanism by which this occurs is still unclear.
Whatever the ER signal is, what is the process by which cytochrome c is released from the mitochondria? Although still an area of active investigation, one model suggests that pro-apoptotic proteins such as Bax or Bak oligomerize to form channels within the outer mitochondrial membrane that are capable of releasing cytochrome c (Danial and Korsmeyer 2004). Typically, Bax, at least, requires some sort of activation event such as induction of a Bcl-2 homology 3 (BH3)-only protein that leads to its translocation from the cytosol to the mitochondrial membrane. Bax does not appear to play an essential role in this system, however, because we have previously demonstrated that Bax deficiency does not protect dopamine neurons from 6-OHDA-induced cell death (O'Malley et al. 2003). Similarly, although Bax is highly expressed in MN9D cells, it remained localized to the cytosol, with no evidence of mitochondrial translocation (Fig. 5a). We also assessed whether Bak or Bok might be involved in this response. Both proteins were predominately expressed in the mitochondrial fraction throughout the time course of toxin treatment (Fig. 5a). As Bok is thought to serve a role similar to that of Bax, i.e. undergoes translocation and oligomerization (Yakovlev et al. 2004; Gao et al. 2005), these observations suggest that Bok was not involved. In contrast, Bak is primarily localized in the outer mitochondrial membrane where it is maintained in an inactive state via association with voltage-dependent anion channel (Cheng et al. 2003). Following an apoptotic stimulus, Bak either homo- or hetero-oligomerizes (Chandra et al. 2005). In agreement with this model, 6-OHDA treatment led to Bak but not Bok higher-order conjugates (Fig. 5c). These observations suggest that, in response to an as yet unidentified UPR-generated signal, Bak undergoes oligomerization leading to the release of cytochrome c.
What the Bak-oligomerizing signal might be is currently under investigation. Preliminary surveys of various BH3-only proteins rule out Bim, Bad or Noxa as possible candidates based on lack of induction and/or translocation to mitochondrial fractions (not shown). Another possibility is the BH3-only protein bbc3/p53 upregulated modulator of apoptosis (PUMA). The latter has recently been reported as an ER stress-induced protein that promotes the release of cytochrome c, leading to caspase activation (Reimertz et al. 2003). Two recent reports suggest that 6-OHDA-induced p53 leads to the up-regulation of PUMA which is required for 6-OHDA-mediated cell death (Biswas et al. 2005; Nair 2006). Thus PUMA may be the link between ER stress and subsequent mitochondrial dysfunction. Whether or not other pro-apoptotic Bcl-2 family proteins might be responsible for 6-OHDA-induced cytochrome c release and apoptotic cell death requires further study.
Oxidative modification of proteins: a link between ROS and UPR
ER stress is also induced by factors that impair disulfide bond formation (Tu and Weissman 2004). As this process requires complex redox-dependent modifications, excessive oxidation can inhibit proper folding leading to ER-associated degradation, protein aggregation and UPR (Rutkowski and Kaufman 2004). One stable marker of protein oxidation is carbonylation, a modification that affects many amino acids (Dalle-Donne et al. 2003). Carbonylation has become a common indicator of oxidative damage in various model systems as well as in aging and in age-associated disorders such as PD (Nystrom 2005). Previously thought to be a random, ubiquitous process, new data suggest that only subsets of proteins are prone to carbonylation, including chaperones, cytoskeletal components, disulfide isomerases and metabolic enzymes (Beal 2002; Choi et al. 2003; Dalle-Donne et al. 2003; Reverter-Branchat et al. 2004). Because many of these proteins are considered protective, their oxidation may trigger downstream sequelae such as the UPR and apoptosis. Although identification of the carbonylated proteins observed here (Fig. 7b) awaits further studies, these results confirm that protein oxidation is one of the first events following 6-OHDA-generated ROS.
Toxin-induced ER stress/UPR is independent of ubiquitin–proteasome impairment
Despite the rapid appearance of oxidatively modified proteins, accumulation of polyubiquitinated proteins did not occur until much later (Figs 8b and c). Interestingly, polyubiquitinated protein levels did not increase in a linear fashion over the course of 6-OHDA exposure, but instead remain steady for the first 6 h, and then rose significantly (Fig. 8d). The lag time may be accounted for by increased proteasome activity between 3 and 9 h (Fig. 8a). Apparently undamaged by the original oxidative stress, proteasome activity increased throughout the time period in which the ER was trying to mount a survival response via the UPR (Fig. 8a). Consistent with these results, Elkon et al. (2004) reported that 6-OHDA-induced protein carbonylation in the PC12 pheochromocytoma cell line concurrent with increased proteosome activity early in the cell death process (<10 h) whereas decreased activity was apparent after this time point. Qualitatively similar results were seen with dopamine treatment of PC12 cells (Keller et al. 2000). Collectively, these data support the notion that proteosome function is rapidly induced following an oxidative stress, presumably in response to increased levels of damaged proteins. The late drop in proteasome activity seen in this study as well as that of Elkon et al. (2004), together with the rise in polyubiquitinated proteins (Fig. 8), may represent the cell's inability to further cope with stress. Interestingly, the increase in polyubiquitinated proteins coincided with the appearance of apoptotic markers, including activated caspase 3 and 9 (Fig. 2c; Holtz and O'Malley 2003). Caspase 3 has been shown to cleave specific subunits of the proteasome, thereby inhibiting its action (Sun et al. 2004). This could be the explanation for the decrease in proteasome activity around 9 h as this decrease coincided with the time of appearance of activated caspase 3. Although PD-like ubiquitin-rich aggregates were not observed in this paradigm, they have been observed in in vivo and in vitro models of proteasome inhibition (McNaught et al. 2002a,b). Thus, the appearance of polyubiquitinated proteins here may represent the precursor material for aggregate formation if the cells remained viable for longer periods of time.
Timeline of 6-OHDA-induced UPR and apoptotic events
Taken together, the results of the present study can be used to construct a temporal model of events associated with 6-OHDA toxicity (Fig. 9). This linear model will be useful in determining the relationship of yet to be defined factors. Knowledge of the signaling pathways utilized by parkinsonian mimetics as well as their temporal induction may aid in designing better interventions in models of PD.
We thank Laura Dugan and Gene Johnson for valuable discussions. We also thank Rui Guan and Steve Harmon for technical assistance. This work was supported by National Institutes of Health grant NS39084, MH45330, and Department of Defense grant DAMD170110777.