J. Neurochem. (2011) 10.1111/j.1471-4159.2011.07290.x
Mitochondrial oxidative stress is a contributing factor in the etiology of numerous neuronal disorders. However, the precise mechanism(s) by which mitochondrial reactive oxygen species modify cellular targets to induce neurotoxicity remains unknown. In this study, we determined the role of mitochondrial aconitase (m-aconitase) in neurotoxicity by decreasing its expression. Incubation of the rat dopaminergic cell line, N27, with paraquat (PQ2+) resulted in aconitase inactivation, increased hydrogen peroxide (H2O2) and increased ferrous iron (Fe2+) at times preceding cell death. To confirm the role of m-aconitase in dopaminergic cell death, we knocked down m-aconitase expression via RNA interference. Incubation of m-aconitase knockdown N27 cells with PQ2+ resulted in decreased H2O2 production, Fe2+ accumulation, and cell death compared with cells expressing basal levels of m-aconitase. To determine the metabolic role of m-aconitase in mediating neuroprotection, we conducted a complete bioenergetic profile. m-Aconitase knockdown N27 cells showed a global decrease in metabolism (glycolysis and oxygen consumption rates) which blocked PQ2+-induced H+ leak and respiratory capacity deficiency. These findings suggest that dopaminergic cells are protected from death by decreasing release of H2O2 and Fe2+ in addition to decreased cellular metabolism.
fetal bovine serum
carbonyl cyanide p-trifluoromethoxyphenylhydrazone
Hank’s Buffered Saline Solution
manganese superoxide dismutase
oxygen consumption rates
reactive oxygen species
rhodamine B 4-[(1,10-phenanthrolin-5-yl)aminocarbonyl]benzyl ester
Roswell Park Memorial Institute
Oxidative stress has been implicated in the onset and progression of numerous chronic and acute neurodegenerative disorders such as Parkinson’s disease (PD), Huntington’s disease, Friedreich’s ataxia, amyotrophic lateral sclerosis and stroke (Schapira 1999, 2008; Lin and Beal 2006; Baron et al. 2007). Multiple factors such as metabolism, aging, environment and genetics can lead to increased steady-state levels of reactive oxygen species (ROS) (Ross and Eisenstein 2002) and in turn oxidative damage to cellular macromolecules (i.e. proteins, DNA, lipids and carbohydrates). Although the occurrence of oxidative stress and neurodegeneration has been established, the precise mechanism(s) by which ROS modify cellular targets to induce neuronal death remain incompletely understood.
Mitochondria are a major source and target of ROS (Murphy 2009). Estimates of superoxide (O2˙−) formation within mitochondria range widely between 0.4% and 1% under normal physiological conditions (Boveris and Chance 1973; Imlay and Fridovich 1991). While the majority of the O2˙− produced within this organelle is detoxified by manganese superoxide dismutase (MnSOD), the fate of mitochondrial O2˙− depends on its reactivity and abundance of other targets such as nitric oxide (NO.) and iron sulfur (Fe-S) containing proteins. Based on concentrations of Fe-S centers, NO. and MnSOD, labile Fe-S centers are major targets of O2˙− in E. coli and yeast (Liochev and Fridovich 1994; Longo et al. 1999). Because labile Fe-S containing proteins such as aconitase are abundant in the brain (Koen and Goodman 1969), they are an important target for O2˙−. In addition to aconitase, other tricarboxylic acid (TCA) cycle enzymes including α-ketoglutarate dehydrogenase and succinate dehydrogenase have been shown to be sensitive to inactivation by ROS (Tretter and Adam-Vizi 2000, 2005).
In mammals and other eukaryotes, there are two aconitases; one localized in the mitochondrial matrix and the other in the cytosol which also functions as iron regulatory protein-1. Of the known functions of mitochondrial aconitase (m-aconitase), two are most prominent. The first major function of m-aconitase is participation in the TCA cycle, where it catalyzes the reversible isomerization of citrate and isocitrate in a 2-step dehydration/hydration reaction via its intermediate form, cis-aconitate. The TCA cycle produces reducing equivalents whereby electrons are carried to the electron transport chain for oxidative phosphorylation which results in the production of ATP. A secondary role for m-aconitase is to act as a biosensor for ROS and iron. Mammalian aconitase and bacterial dehydratases contain a [4Fe-4S] prosthetic group in their catalytic centers which are susceptible to inactivation by ROS, particularly O2˙− (Gardner and Fridovich 1991a,b; Flint et al. 1993). Aconitase is uniquely sensitive to O2˙− mediated oxidative inactivation because of the presence of a single unligated iron atom, such that oxidation of the [4Fe-4S]2+ cluster renders it unstable and promotes removal of the labile iron atom, consequently forming hydrogen peroxide (H2O2) by the reduction of O2˙−. In the mammalian brain, approximately 85% of aconitase activity is localized to the mitochondria; and despite the fact that both aconitases contain a [4Fe-4S] prosthetic group in their catalytic sites, m-aconitase is more sensitive than cytosolic aconitase to oxidative inactivation perhaps in part because of cellular localization (Liang et al. 2000). Release of redox-active iron (Fe2+) from aconitase and other hydro-lyases has been reported in cell-free systems (Flint et al. 1993; Keyer and Imlay 1996). m-Aconitase has also been shown to be a source of ˙OH, presumably via Fenton chemistry initiated by the co-released Fe2+ and H2O2 in bovine heart purified m-aconitase (Vasquez-Vivar et al. 2000). Besides being a source of Fe2+, translation of m-aconitase can be regulated by fluctuations in iron via an iron regulatory element (Lemire et al. 2007) in its 5′ UTR (Kim et al. 1996) which functions similarly to that of the iron storage protein ferritin; whereby decreased iron levels allows an iron regulatory protein to bind the iron regulatory element of m-aconitase blocking its translation. This suggests that expression of m-aconitase can be controlled by changes in iron levels.
A number of neurodegenerative diseases in which oxidative stress has been implicated, as well as in vivo and in vitro models of these disorders, collectively demonstrate decreased aconitase activity presumably via oxidative inactivation by ROS (Patel et al. 1996; Rotig et al. 1997; Melov et al. 1999; Schapira 1999; Tabrizi et al. 1999; Park et al. 2001; Liang and Patel 2004b; Vielhaber et al. 2008). Because of the labile iron atom found within the [4Fe-4S]2+ cluster of m-aconitase, and its proximity to mitochondrially generated ROS, it is an ideal candidate for oxidative inactivation. Although aconitase has been well established as a sensitive target of ROS, the consequences of oxidative inactivation of this important mitochondrial enzyme are still being explored. The role of aconitase in mediating O2˙− toxicity has been shown in both bacteria (Gardner and Fridovich 1991b) and yeast (Longo et al. 1999). These observations are based on the principle that oxidation of the [4Fe-4S]2+ cluster of aconitase by O2˙−, in the presence of protons, results in the formation of Fe2+ and H2O2 and in turn produce ˙OH via the Fenton reaction (Liochev and Fridovich 1994). Further understanding the role of m-aconitase in mediating neurotoxicity by acting as a source of ROS may provide a mechanism by which oxidative inactivation of ROS-sensitive targets can lead to neurodegeneration. The role of iron and mitochondrial oxidative stress as major contributors to neurodegenerative diseases such as PD has been shown (Beal 1998; Jenner 2003; Kaur and Andersen 2004; Berg and Youdim 2006). Work in our laboratory has demonstrated that m-aconitase can become oxidatively inactivated in mice treated with the parkinsonian toxin MPTP, and that this inactivation correlates with increased chelatable mitochondrial iron in the ventral midbrain region (Liang and Patel 2004b). Over-expression of m-aconitase in primary midbrain neuronal-glial cultures, which occurred predominantly in astrocytes, resulted in increased Fe2+ and H2O2-dependent neuronal death (Cantu et al. 2009). However, whether oxidative inactivation of neuronal m-aconitase can release Fe2+ and H2O2 and contribute to neurotoxicity remains unknown. To verify the role of m-aconitase in Fe2+ and H2O2-dependent neurotoxicity, it is important to specifically decrease m-aconitase expression and thereby decrease a major contributor to the production of Fenton reactants. The goal of this study was to determine the role of m-aconitase in the death of dopaminergic cells. To accomplish this, we decreased the expression of m-aconitase via RNA interference and increased steady-state levels of ROS in immortalized rat dopaminergic (N27) cells by using paraquat (PQ2+), a bipyridyl herbicide that redox cycles with molecular oxygen and serves as a continuous source of O2˙−. N27 cells have been extensively characterized as having key features of dopaminergic cells, such as expressing tyrosine hydroxylase and the dopamine transporter (Clarkson et al. 1998). Here, we demonstrate that knockdown of m-aconitase in dopaminergic cells increases their resistance to PQ2+-induced cell death, predominantly via a mechanism involving decreased Fe2+ and H2O2 release.
Materials and methods
Cell culture media and reagents were obtained from Invitrogen (Carlsbad, CA, USA). All other materials were obtained from Sigma (St. Louis, MO, USA). Immortalized rat dopaminergic N27 cells were a kind gift from Drs. Curt Freed and Kedar Prasad at University of Colorado, Anschutz Medical Campus.
N27 cell culture
N27 cells were grown in RPMI 1640 medium supplemented with 10% fetal bovine serum (FBS), penicillin (100 U/mL) and streptomycin (100 U/mL). Cells were plated for experimentation in RPMI 1640 medium supplemented with 1% FBS, penicillin (10 U/mL) and streptomycin (10 U/mL) and maintained at 37°C in a 5% CO2 humidified atmosphere.
Knockdown of m-aconitase was achieved using RNA interference. Four pre-designed, individual gene specific siRNA duplexes were obtained from Dharmacon (Lafayette, CO, USA) and screened for transfection efficiency. Of the four, one duplex was selected based on its high knockdown efficiency (CCAGUGAGUACAUCCGAUA). N27 cells were plated in 6-well plates at 1 × 105 cells/well overnight in RPMI 1640 medium supplemented with 1% FBS, penicillin (10 U/mL) and streptomycin (10 U/mL). Cells were then washed once with RPMI 1640 (1% FBS, 10 U/mL penicillin, 10 U/mL streptomycin) followed by addition of 800 μL of same media. Cells were then transfected using Dharmacon transfection reagent DharmaFect 1 (T-2001–2003) and 100 nM siRNA (final concentration). For each well, 5 μL of 20 μM m-aconitase, non-targeting or lamin siRNA was combined with 45 μL 1× siRNA buffer and 50 μL serum-free RPMI 1640. A second mixture containing 1.5 μL DharmaFect 1 and 98.5 μL serum-free RPMI 1640 was made. After 5 min, 100 μL of the first mixture was combined with 100 μL of the second mixture and incubated at 25°C for 20 min. 200 μL of combined siRNA mixture (siRNA, 1× siRNA buffer, Dharmafect 1 reagent and serum-free RPMI 1640) was added to cells. After 48 h, 1 mL of media was added to each well. Mock transfection consisted only of DharmaFect reagent with 1× siRNA buffer without siRNA; control cells were not transfected or treated with any transfection reagents. All experiments were conducted 72 h post-transfection.
Verification of m-aconitase knockdown
RNA was isolated using the RNeasy kit® (Qiagen, Valencia, CA, USA) and quantified using RiboGreen® RNA Quantitation Kit (Molecular Probes, Eugene, OR, USA). An Applied Biosystems (Carlsbad, CA, USA) 7500 Fast Real-Time PCR System was used for real time PCR. RNA was reverse transcribed to cDNA using the High-Capacity cDNA Reverse Transcription Kit (Applied Biosystems). Thermal cycling conditions included 25°C for 10 min, 37°C for 120 min and 85°C for 5 s. PCR reaction conditions included one cycle at 50°C for 2 min, one cycle at 95°C for 10 min and 40 cycles of 95°C for 15 s and 60°C for 1 min. Primers and probes for rat m-aconitase were purchased from Integrated DNA Technologies (San Diego, CA, USA); forward primer: 5′-CCGCCTTCCTGTTCAGTTTG-3′, reverse primer: 5′-TGTAGAGGGAGTGCTGTCATCAA-3′, probe: 5′-TTTGTCTTTGAGCAACCCATGCAA-3′.
In addition, western blot analysis was used to confirm knockdown. Media was replaced with 150 μL ice-cold lysis buffer (0.1% Triton-X in phosphate-buffered saline, protease inhibitor cocktail), placed in 4°C for 30 min, and sonicated. 10–20 μg of protein was loaded onto a 12% gel and analyzed via sodium dodecyl sulfate–polyacrylamide gel electrophoresis. Proteins were detected with a rabbit polyclonal IgG against human m-aconitase capable of cross-reacting with rat (1 : 500, Atlas Antibodies, Stockholm, Sweden) and with rabbit affinity-purified monoclonal IgG against rat β-actin (1 : 1000, Sigma) to confirm equal loading in gels. Densitometry was analyzed using ImageJ (NIH, Bethesda, MD, USA) by normalizing m-aconitase protein bands to β-actin.
Detection of H2O2
Hydrogen peroxide was measured using Amplex Red (Invitrogen), a horse radish peroxidase-linked fluorometric assay. Naïve and transfected N27 cells were plated on 96-well plates at 1.4 × 105 cells/well and allowed to adhere overnight. Cell culture media was removed and replaced with 150 μL of Hank’s Buffered Saline Solution (HBSS) solution containing 1 mg/mL glucose, 0.1 U/mL horse radish peroxidase, 50 μM Amplex Red and PQ2+. Resorufin fluorescence was measured after 6 h by a Gemini fluorescence microplate reader equipped for excitation in the range of 530–560 nm and fluorescence emission detection at 590 nm (Molecular Devices, Sunnyvale, CA, USA).
Detection of Fe2+
Detection of Fe2+ was conducted using a fluorescent iron indicator, rhodamine B 4-[(1,10-phenanthrolin-5-yl)aminocarbonyl]benzyl ester (RPA) (Petrat et al. 2002), whose fluorescence is quenched by mitochondrial ferrous iron. Naïve and transfected N27 cells were incubated with PQ2+ in fresh cell culture media for 4 h. Cell culture media was then removed and replaced with 1 μM RPA dissolved in HBSS. Cells were kept at 37°C for 10 min, rinsed with HBSS, and placed at 37°C for an additional 10 min. Cultures were rinsed a final time with HBSS before five randomly selected images were captured on an Olympus (Center Valley, PA, USA) IX81 inverted motorized microscope. Images were quantified by measuring the mean pixel intensity using ImageJ (NIH).
Cell death analysis
Cell death analysis was performed by measuring release of lactate dehydrogenase (LDH) enzyme activity in addition to propidium iodide (PI) staining (Invitrogen, Eugene, OR, USA). Media samples from PQ2+ treated cells were collected and LDH activity was measured spectrophotometrically at 30°C as the amount of pyruvate consumed, by monitoring the decrease in absorbance because of the oxidation of NADH at 340 nm (Vassault 1983). For PI staining, cells were rinsed with 2× SSC (0.3 M NaCl, 0.03 M sodium citrate, pH 7.0), incubated in 2 μM PI for 5 min and washed once again with 2× SSC. Five randomly selected fluorescent images were captured using an Olympus IX81 inverted motorized microscope and merged with phase contrast images. PI+ cells from fluorescent and phase contrast overlay were counted to ensure accurate identification of PI+ cells.
Aconitase and fumarase activities
Aconitase and fumarase activities were measured spectrophotometrically as previously described (Patel et al. 1996).
Reactivation of aconitase
N27 cells were treated with 0, 0.5, and 1 mM PQ2+ for 4 h and lysed with 200 μL ice-cold aconitase lysis buffer (50 mM Tris–HCl, pH 7.4 containing 0.6 mM MnCl2, 1 mM l-cysteine, 1 mM citrate, and 0.5% Triton-X 100) for 30 min at 4°C. N27 cell lysates were collected and 90 μL from each treatment condition were either combined with reactivation buffer [0.5 M dithiothreitol (10 μL), 20 mM Na2S (1 μL), 20 mM ferrous ammonium sulfate (1 μL) in 50 mM Tris–HCl (pH 8.0)] or with vehicle for 30 min at 30°C.
Levels of m-aconitase apoprotein
m-Aconitase apoprotein levels were measured by western blot analysis as in Verification of m-aconitase knockdown section of Materials and Methods.
Measurement of oxygen consumption and extracellular acidification rate
Oxygen consumption and extracellular acidification rates were determined using a Seahorse XF24 analyzer. Naïve and transfected N27 cells were plated on XF24 microplates (Seahorse Bioscience, North Billerica, MA, USA) at 3.0 × 104 cells/well in RPMI 1640 supplemented with 1% FBS, penicillin (10 U/mL) and streptomycin (10 U/mL) and kept at 37°C in a 5% CO2 humidified atmosphere overnight. For experiments addressing the effects of PQ2+, incubation lasted 6 h. For measurement of oxygen consumption and glycolysis, growth media was replaced with incubation media which consisted of RPMI 1640 media (already containing 11 mM glucose and lacking sodium bicarbonate) supplemented with 1 mM sodium pyruvate and 2 mM Glutamax (Sigma). 10× stock solutions of oligomycin (oligo) (5 μM), carbonylcyanide p-trifluoromethoxyphenylhydrazone (FCCP) (30 μM) and antimycin A (AA) (3 μM) were diluted in incubation media and 75 μL of each 10× stock solution was loaded onto an XF24 cartridge. The XF24 microplate was loaded into the Seahorse XF24 analyzer following the manufacturer’s instructions. All experiments were carried out at 37°C. Respiratory parameters were quantified by subtracting respiration rates at times before and after addition of electron transport chain inhibitors according to Seahorse Biosciences; basal respiration: baseline respiration minus AA-dependent respiration; ATP turnover: baseline respiration minus oligo-dependent respiration; H+ leak: oligo-dependent respiration minus AA-dependent respiration; respiratory capacity: AA-dependent respiration minus FCCP-dependent respiration. Values were calculated for each individual well and averaged for each condition. PQ2+-dependent respiratory capacity deficiency was calculated by subtracting the oxygen consumption rates (OCR) from 0.3 mM PQ2+ cells from OCR in 0 mM PQ2+ treated cells.
For comparison between two experimental groups, a Student’s t-test was used. For three or more experimental groups, a one-way anova with the Bonferroni’s post-hoc test was used. A two-way anova was used for comparing two different treatment groups in the presence and absence of PQ2+. Values of p < 0.05 or more were considered statistically significant.
To determine the role of neuronal m-aconitase as a source of H2O2 and Fe2+, we asked whether oxidative inactivation of m-aconitase resulted in accumulation of H2O2, Fe2+ and cell death in N27 cells and whether decreasing m-aconitase expression would attenuate these effects. Furthermore, we determined the effects of decreased m-aconitase expression on cellular metabolism.
H2O2 production, mitochondrial Fe2+ accumulation, and cell death increase in N27 cells following oxidative inactivation of aconitase by PQ2+
To determine the effects of decreased m-aconitase expression on H2O2, Fe2+ and cell death, we first examined the effects of the O2˙− generating compound PQ2+ on aconitase activity. Consistent with our previous studies in primary midbrain and cortical cultures (Patel et al. 1996; Cantu et al. 2009), aconitase activity decreased in N27 cells after 4 h of 0.5 mM and 1 mM PQ2+ incubation while the activity of fumarase, a control mammalian enzyme which lacks an oxidation sensitive Fe-S center, remained unchanged (Fig. 1a). In order to verify that decreased aconitase activity by PQ2+ (and consequent production of O2˙−) results in a disruption of the [4Fe-4S] cluster of aconitase rather than degradation of the protein, we asked whether aconitase inactivation by PQ2+ was reversible. N27 cell lysates were collected after 4 h of 0, 0.5 and 1 mM PQ2+ incubation and treated with vehicle or with a combination of dithiothreitol, sodium sulfide (Na2S), and ferrous ammonium sulfate (reactivation reagents) for 30 min. Inactive aconitase resulting from PQ2+ incubation was reactivated after 30-min incubation with reactivation reagents (Fig. 1b). Furthermore, although aconitase enzymatic activity was significantly decreased with PQ2+, apoprotein levels of m-aconitase remained unchanged (Fig. 1c and d). Together, these results support that oxidative inactivation of aconitase occurred via disruption of the [4Fe-4S] cluster rather than degradation of the protein.
To determine the consequences of oxidative inactivation of aconitase, we established the time- and dose-dependent effects of H2O2 production, mitochondrial Fe2+ accumulation and cell death after short (4–6 h) and long (18 h) exposure to the O2˙− generating compound PQ2+. A concentration-dependent increase in H2O2 production was observed by measuring Amplex Red fluorescence in N27 cells after short exposure to 0.3 and 1 mM PQ2+ (Fig. 2a). PQ2+ concentrations in the mM range were necessary to significantly increase ROS production as with MPP+ (data not shown), although concentrations varied from other compounds used to increase ROS production such as rotenone (50 nM) and antimycin A (25 nM) (Sipos et al. 2003). Increased mitochondrial Fe2+ was detected in N27 cells after short exposure to PQ2+ via RPA fluorescence, whereby quenching indicates an increase in Fe2+ concentration. A significant decrease in RPA fluorescence intensity was observed at 1 mM PQ2+ indicating an increase in mitochondrial Fe2+ (Fig. 2b). Because of the sensitivity of RPA used to measure iron, we increased the concentration of PQ2+ from 0.3 mM to 0.5 mM. However, treatment of N27 cells with 0.5 mM PQ2+ was not statistically different from 0 mM or 1 mM PQ2+ (Fig. 2b).
Cell death was evaluated after short and long PQ2+ exposure via measurement of extracellular LDH release. Although no difference in LDH release was found after short PQ2+ exposure (Fig. 2c), a significant increase was observed after long exposure to 1 mM PQ2+ (Fig. 2d). Collectively, these studies indicate that oxidative inactivation of aconitase, increased H2O2 production and increased mitochondrial Fe2+ accumulation occur at times preceding cell death in N27 cells.
siRNA treatment of N27 cells decreases m-aconitase mRNA and protein expression
To confirm the role that oxidative inactivation of m-aconitase plays in mediating neuronal death, we knocked down its expression using RNA interference in N27 cells. m-Aconitase knockdown was successfully achieved by transfecting N27 cells with a siRNA duplex specific for m-aconitase. m-Aconitase mRNA expression was measured 72 h post-transfection using real-time PCR and compared to non-transfected cells (control), mock transfected cells (mock), lamin siRNA transfected cells (lamin) and non-targeting siRNA transfected cells (non-targeting). N27 cells transfected with m-aconitase siRNA demonstrated a significant decrease in m-aconitase mRNA expression of approximately ∼ 87% compared to control and ∼ 91% compared to non-targeting siRNA (Fig. 3a). To confirm that knockdown was successful at this time-point, western blot analysis was used to measure protein expression. Similarly, m-aconitase siRNA transfected cells showed decreased protein expression compared to control cells and non-targeting siRNA transfected cells (Fig. 3b). After quantifying m-aconitase protein expression, we observed a decrease of approximately ∼ 73% compared to control and ∼ 60% compared to non-targeting siRNA (Fig. 3c). As a positive control, we measured lamin protein expression in N27 cells transfected with lamin siRNA. We observed a significant decrease in lamin protein levels 72 h post-transfection by western blot analysis (data not shown).
m-Aconitase knockdown attenuates PQ2+-induced increase in H2O2 production, mitochondrial Fe2+ accumulation, and cell death
We next investigated whether decreased m-aconitase expression would attenuate release of Fenton reactants (H2O2 and Fe2+) and cell death after an oxidative insult. If m-aconitase was contributing to H2O2 and Fe2+ release, then decreasing its expression should attenuate this effect. Because no statistical difference in m-aconitase mRNA or protein levels was observed between control and non-targeting siRNA transfected cells, we proceeded to compare only non-targeting and m-aconitase siRNA transfected cells. This allowed us to account for any differences attributed to transfection alone. Cells expressing basal levels of m-aconitase (non-targeting) and cells with decreased m-aconitase expression (m-aconitase) were treated with PQ2+ (in order to increase O2˙− and inactivate aconitase). Although H2O2 production increased in both non-targeting and m-aconitase transfected N27 cells after short PQ2+ exposure, m-aconitase knockdown cells showed a significant decrease compared to cells expressing basal m-aconitase (Fig. 4a). We proceeded to conduct a similar experiment to determine if Fe2+ release was also decreased when m-aconitase deficient cells were treated with PQ2+. Indeed, cells transduced with non-targeting siRNA demonstrated a decrease in RPA fluorescence (indicating an increase in free mitochondrial Fe2+) after short PQ2+ exposure, while cells transfected with m-aconitase siRNA showed no change in RPA fluorescence (Fig. 4b). These data suggest that (i) H2O2 and Fe2+ are released from m-aconitase after the enzyme is oxidatively inactivated and (ii) when m-aconitase is knocked down, H2O2 and Fe2+ release is attenuated.
One consequence of oxidative inactivation of aconitase is cell death, as reported in bacteria and yeast (Gardner and Fridovich 1991b; Longo et al. 1999), presumably via co-released H2O2 and Fe2+ undergoing Fenton chemistry and producing toxic ˙OH radicals. We therefore wanted to test whether decreasing m-aconitase expression would prevent cell death. Similar to the above experiments, N27 cells were transfected with non-targeting and m-aconitase siRNA and cell death was measured after long PQ2+ exposure. Because no cell death was observed after long exposure to 0.3 mM PQ2+ (Fig. 2d), we proceeded to increase the concentration of PQ2+ to 0.5 mM. PQ2+-mediated cell death was significantly decreased after long exposure to 1 mM PQ2+ in m-aconitase knockdown cells as measured by release of extracellular LDH (Fig. 4c). To verify these results, we used PI staining as an alternative for measuring cell death. Fluorescent images of PI stained cells were overlaid with phase contrast images to ensure accurate counting of PI+ cells. Similarly, m-aconitase knockdown cells showed a decrease in PQ2+-mediated cell death compared to cells expressing basal levels of m-aconitase (Fig. 4d). Together these results implicate oxidative inactivation of m-aconitase as a contributing factor to dopaminergic cell death.
Decreased m-aconitase expression changes cellular metabolism
As m-aconitase plays an essential metabolic role in the TCA cycle, we proceeded to determine the bioenergetic consequences of decreased m-aconitase expression and whether they conferred resistance to PQ2+ neurotoxicity. To address this question, we conducted a complete bioenergetic profile using a Seahorse XF24 Analyzer on cells expressing basal or decreased levels of m-aconitase in the presence or absence of PQ2+. A bioenergetic profile includes the addition of electron transport chain inhibitors in order to place the mitochondria in different states of respiration; they include the ATP synthase inhibitor oligomycin, the mitochondrial uncoupler FCCP, and the complex III inhibitor antimycin A. Cells were transfected with non-targeting and m-aconitase siRNA for 72 h and oxygen consumption and glycolytic rates were measured. Figure 5(a) shows decreased OCR in m-aconitase knockdown N27 cells compared to cells expressing basal m-aconitase (non-targeting). In addition, rates of glycolysis, as measured by changes in the extracellular acidification rate, were also decreased in m-aconitase knockdown cells (Fig. 5b). Respiration parameters can be extrapolated from the OCR data of the bioenergetic profile (as described by Seahorse Biosciences). This enables one to determine (i) basal respiration, (ii) ATP turnover (amount of O2 consumption devoted to ATP production), (iii) H+ leak (amount of O2 consumption devoted to maintaining the proton gradient) and (iv) respiratory capacity (the maximal respiratory rate under conditions of uncoupled respiration). m-Aconitase knockdown N27 cells showed significantly decreased basal respiration, ATP turnover, H+ leak and respiratory capacity (Fig. 5c–f). These data suggest that although m-aconitase knockdown cells were efficiently respiring, they were doing so at significantly lower rates.
PQ2+ treatment decreases respiratory capacity and glycolysis, and increases H+ leak
As m-aconitase knockdown uniformly decreases both OCR and glycolysis, it is possible that their resistance to PQ2+ neurotoxicity may be related to a metabolic effect in addition to Fe2+ and H2O2 released via oxidative inactivation. We therefore examined the effects of short PQ2+ exposure on OCR and glycolysis in naïve (non-transfected) N27 cells. After short PQ2+ exposure, a bioenergetic profile demonstrated that FCCP-dependent respiration (Fig. 6a) and glycolysis (Fig. 6b) were decreased in PQ2+ treated cells. Quantification of respiration parameters clearly indicated that basal respiration (Fig. 6c) and ATP turnover (Fig. 6d) remained unaltered after PQ2+ treatment while H+ leak (Fig. 6e) significantly increased and respiratory capacity (Fig. 6f) decreased. This suggests that PQ2+-dependent neurotoxicity may be a result of increased H+ leak in addition to decreased respiratory capacity.
PQ2+ exacerbates basal and stimulated OCR and glycolysis but attenuates H+ leak and respiratory capacity in m-aconitase knockdown N27 cells
We proceeded to determine the effect of PQ2+ on OCR and glycolysis in m-aconitase knockdown cells. N27 cells were transfected with non-targeting and m-aconitase siRNA for 72 h followed by short exposure to PQ2+. As observed in Fig. 5, knocking down m-aconitase decreased both OCR and glycolysis. A secondary insult by PQ2+ further decreased OCR and glycolysis (Fig. 7a and b). Likewise, PQ2+ incubation further decreased basal respiration and ATP turnover in m-aconitase knockdown cells (Fig. 7c and d). However, PQ2+-dependent increases in H+ leak were significantly attenuated in m-aconitase knockdown cells (Fig. 7e), suggesting that decreased H+ leak may be a potential mechanism by which m-aconitase knockdown cells are protected from PQ2+-dependent cell death. Respiratory capacity was unchanged between non-targeting and m-aconitase knockdown cells in the presence of PQ2+ (Fig. 7f). However, the decrease in respiratory capacity after PQ2+ exposure in m-aconitase knockdown cells (88 ± 13 pmolO2/min/30 K cells) was significantly lower than in cells expressing basal m-aconitase (non-targeting) (173 ± 28 pmolO2/min/30 K cells). Respiratory capacity deficiency has been described as a key index implicated in cell death (Yadava and Nicholls 2007). Therefore, neuroprotection resulting from decreased m-aconitase expression may be because of metabolic mechanisms via decreased H+ leak and decreased respiratory capacity deficiency.
This study supports the hypothesis that oxidative inactivation of m-aconitase serves as a source of iron and ROS and as a contributor of neuronal death. First, N27 dopaminergic cells showed a PQ2+-dependent increase in H2O2, Fe2+ and cell death. Second, decreased m-aconitase expression in N27 cells by RNA interference significantly attenuated PQ2+-induced increases in H2O2, Fe2+ and cell death. Third, m-aconitase knockdown N27 cells showed lower basal and stimulated metabolic rates (OCR and glycolysis). Finally, PQ2+-dependent increase in H+ leak and respiratory capacity deficiency were attenuated in m-aconitase knockdown cells. Together these findings suggest that both oxidative and metabolic mechanisms may contribute to neuroprotection in m-aconitase knockdown cells by (i) decreasing ROS production and (ii) lowering metabolic function which in turn attenuates H+ leak and respiratory capacity deficiency under conditions of oxidative stress.
Aconitase and other [4Fe-4S] containing dehydratases have been extensively characterized as targets of ROS, specifically O2˙− and ONOO− (Gardner and Fridovich 1991a,b; Flint et al. 1993; Keyer and Imlay 1997). Aconitase inactivation has been reported in various human neurodegenerative disorders associated with mitochondrial oxidative stress including Huntington’s disease, Friedreich’s ataxia, progressive supranuclear palsy and temporal lobe epilepsy (Rotig et al. 1997; Schapira 1999; Tabrizi et al. 1999; Park et al. 2001; Vielhaber et al. 2008). Furthermore, aconitase inactivation has been shown in animal and cell models of neurodegeneration such as the Huntington R6/2 transgenic mouse (Tabrizi et al. 2000), MnSOD deficient mice (Melov et al. 1999; Liang and Patel 2004a), DJ1 knockout mice (Andres-Mateos et al. 2007), PINK1 knockout mice (Gautier et al. 2008), MPTP treated mice (Liang and Patel 2004b), cerebral ischemia (Mackensen et al. 2001), aging (Yan et al. 1997; Patel and Li 2003), oxygen glucose deprivation (Li et al. 2001), β-amyloid toxicity (Longo et al. 2000), excitotoxicity (Patel et al. 1996) and MPP+ neurotoxicity (Kalivendi et al. 2003; Shang et al. 2004). Aconitase inactivation is frequently used as a marker for oxidative stress (Gardner and Fridovich 1992). Because the mitochondrial isoform accounts for the majority of aconitase in the brain (∼ 85%) (Liang et al. 2000), it is suggested that m-aconitase is a sensitive target of ROS in dopaminergic cells.
In addition to being a target of ROS, there is evidence that m-aconitase can be a source of ROS and free mitochondrial iron (Liochev and Fridovich 1994). Aconitase inactivation by O2˙− was first suggested to be a source of ˙OH via the production of H2O2 and Fe2+ in cell free systems by Flint et al. (1993). Evidence of ˙OH production from bovine heart purified m-aconitase was later shown by Vasquez-Vivar et al. (2000). In addition, aconitase has been shown to be a source of iron and toxicity in bacteria and yeast (Gardner and Fridovich 1991b; Longo et al. 1999).
Work from our laboratory has been focused on determining a role for m-aconitase in mediating O2˙− neurotoxicity via oxidative inactivation. We have shown that m-aconitase may be an important source of iron in MPTP neurotoxicity (Liang and Patel 2004b). Importantly, we recently demonstrated that over-expression and inactivation of m-aconitase by PQ2+ increased free mitochondrial iron, H2O2 and neurotoxicity in primary midbrain cultures (Cantu et al. 2009). However, in the latter study, adenoviral m-aconitase transduction occurred predominantly in astrocytes and H2O2 release injured neighboring neurons. This prompted us to (i) investigate whether knocking down m-aconitase in dopaminergic cells would protect them by releasing less H2O2 and Fe2+ after inactivation of the enzyme and (ii) determine the relative roles of oxidative stress and metabolism in the mechanism. Consistent with our previous work, two major findings provide evidence that oxidative inactivation of m-aconitase releases redox active iron and H2O2 which contribute to cell death. First, PQ2+ incubation inactivates aconitase, increases iron and increases H2O2 at times preceding cell death (Figs 1 and 2). In N27 cells, PQ2+ leads to neuronal death at considerably earlier time-points of ∼ 18–24 h compared to MPP+ which predominately occurs at ∼ 48 h (Drechsel et al. 2007; Cantu et al. 2009). Therefore, it is highly unlikely that N27 cells are producing considerable amounts of H2O2 at 18 h, as cells are already dead or dying at this time-point and the production of ROS by PQ2+ depends on the inner mitochondrial transmembrane potential (Castello et al. 2007). Second, knocking down m-aconitase significantly attenuates PQ2+-induced increases in H2O2, Fe2+ and cell death (Fig. 4). A post-translational mechanism of inactivation is supported by the ability of iron and reactivation agents to reactivate PQ2+-induced inactivation of aconitase and unchanged apoprotein levels. Because our studies focused on the effects of mitochondrial aconitase inactivation via the production of O2˙− by PQ2+, we speculate that both necrotic and apoptotic mechanisms of cell death contributed to PQ2+-induced cell death (Patel et al. 1996; Li et al. 2001; Peng et al. 2004).
Although other mechanisms exist for increased H2O2 and Fe2+ in N27 cells, neurons and parkinsonian substantia nigra (i.e. NADPH oxidase, uncoupling protein-2, prolyl hydroxylase, neuromelanin, ferritin, and lactotransferrin), the fact that decreased m-aconitase expression directly attenuates production of H2O2 and Fe2+ suggests a major role for m-aconitase as their source (Youdim and Riederer 1993; Gerlach et al. 1997; Double et al. 2000; Cristovao et al. 2009; Lee et al. 2009; Zhang et al. 2009). Together, these findings demonstrate that oxidative inactivation of m-aconitase mediates neurotoxicity.
The basis of the PQ2+-induced attenuation of H2O2, Fe2+ and cell death in m-aconitase knockdown N27 cells is most likely because of decreased availability of the target (m-aconitase) for oxidative inactivation and therefore eliminating an important source of ROS and iron. This is consistent with our previous observations that N,N′-bis (2-hydroxybenzyl) ethylenediamine-N,N′-diacetic acid and catalase inhibit PQ2+-induced neurotoxicity (Cantu et al. 2009). It is important to note that m-aconitase was not completely knocked out; approximately 27–40% m-aconitase protein was still detectable after 72 h of transfection with m-aconitase siRNA (Fig. 3). As m-aconitase is primarily involved in the isomerization of citrate via the TCA cycle, m-aconitase deficiency per se may have a deleterious effect because of interference with mitochondrial metabolism. Whereas a global decrease in total cellular respiration (i.e. basal and stimulated OCR and glycolysis) occurred in N27 cells deficient in m-aconitase (Fig. 5), they were resistant to PQ2+-induced oxidative stress and cell death (Fig. 4) rather than being more vulnerable. This is consistent with the observations of Tretter and Adam-Vizi (2005) that completely abolishing m-aconitase activity allowed the TCA cycle to remain functional via the transamination of glutamate to α-ketoglutarate. Furthermore, anapleurotic mechanisms have long been known to prevent TCA dysfunctions arising from individual enzyme deficiencies (Gibala et al. 2000; Owen et al. 2002). Based on these data, we do not speculate that inhibiting m-aconitase activity would have any significant effect on the production of NADPH. Therefore, cellular antioxidant systems which rely on NADPH for energy, such as the thioredoxin and glutathione systems, should remain unaffected. The observation that m-aconitase deficiency lowers overall rates of OCR and glycolysis is indicative of metabolic quiescence or ‘hunkering down,’ which has been proposed by Gardner (Gardner 1997) to explain the significance of O2˙− sensing and inactivation by aconitase. Lower basal OCR and glycolytic rates may offer m-aconitase deficient cells a metabolic advantage and be cytoprotective in itself. In fact, TCA cycle mutations resulting in deficiency as seen in the fumarate hydratase deficient (FH−/FH−) UOK262 cell line and in isocitrate dehydrogenase 1 and 2 mutations are known to have oncogenic phenotypes (Reitman and Yan 2010; Yang et al. 2010). In the case of fumarate hydratase deficiency, cells exhibit a quintessential Warburg effect, by using aerobic glycolysis for metabolism (Yang et al. 2010). In our model, glycolysis is decreased in m-aconitase knockdown cells (Fig. 5), suggesting that cells are not relying on aerobic glycolysis for respiration as seen in cancer cells. This study highlights a mechanistic role of m-aconitase in mitochondrial oxidative stress. It is unlikely that decreasing metabolism to achieve neuroprotection could be a viable therapeutic avenue for neurodegenerative diseases such as PD in which mitochondrial function is already impaired (Arthur et al. 2009) and improving mitochondrial bioenergetics is neuroprotective (Yang et al. 2009).
Decreased metabolism may also be playing a role in the resistance of m-aconitase deficient N27 cells to PQ2+ neurotoxicity. This is based on our observation that m-aconitase knockdown cells showed a global decrease in total cellular metabolism (i.e. basal and stimulated OCR and glycolysis) while H+ leak and respiratory capacity deficiency were significantly attenuated after PQ2+ exposure. Respiratory capacity has been identified as a critical deleterious metabolic signature (Yadava and Nicholls 2007). Therefore, the resistance of m-aconitase deficient cells to PQ2+ toxicity suggests that both oxidative and metabolic mechanisms may be playing a role in protection. However, Yadava and Nicholls (2007) concluded that spare respiratory capacity rather than oxidative stress regulates glutamate excitotoxicity after rotenone treatment (Yadava and Nicholls 2007). Interestingly, when naïve cells were treated with PQ2+, glycolysis and respiratory capacity decreased, basal respiration and ATP turnover remained unchanged, and H+ leak increased (Fig. 6). This suggests that in addition to respiratory capacity, increased H+ leak may also be a hallmark of neuronal death.
In conclusion, this study emphasizes the role of m-aconitase in mediating neurotoxicity via oxidative mechanisms involving deleterious release of Fe2+ and H2O2. Furthermore, by decreasing metabolism after m-aconitase knockdown, we were able to protect neurons from PQ2+-dependent cell death via attenuation of H+ leak and respiratory capacity deficiency.
This work was supported by National Institutes of Health Grants NS045748 (MP), NS039587 (MP), and Supplement NS039587-S1 (DC). We would also like to thank Kristen Ryan for helpful discussions.