©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
Superoxide Radical and Iron Modulate Aconitase Activity in Mammalian Cells (*)

Paul R. Gardner (1)(§), Inés Raineri (2)(¶), Lois B. Epstein (2) (3), Carl W. White(**) (1)

From the (1) Department of Pediatrics, Division of Pulmonary Medicine, National Jewish Center for Immunology and Respiratory Medicine, Denver, Colorado 80206 and the (2) Cancer Research Institute and (3) Department of Pediatrics, School of Medicine, University of California, San Francisco, California 94143

ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES

ABSTRACT

Aconitase is a member of a family of iron-sulfur-containing (de)hydratases whose activities are modulated in bacteria by superoxide radical (O)-mediated inactivation and iron-dependent reactivation. The inactivation-reactivation of aconitase(s) in cultured mammalian cells was explored since these reactions may impact important and diverse aconitase functions in the cytoplasm and mitochondria. Conditions which increase O production including exposure to the redox-cycling agent phenazine methosulfate (PMS), inhibitors of mitochondrial ubiquinol-cytochrome c oxidoreductase, or hyperoxia inactivated aconitase in mammalian cells. Overproduction of mitochondrial Mn-superoxide dismutase protected aconitase from inactivation by PMS or inhibitors of ubiquinol-cytochrome c oxidoreductase, but not from normobaric hyperoxia. Aconitase activity was reactivated (t of 12 ± 3 min) upon removal of PMS. The iron chelator deferoxamine impaired reactivation and increased net inactivation of aconitase by O. The ability of ubiquinol-cytochrome c oxidoreductase-generated O to inactivate aconitase in several cell types correlated with the fraction of the aconitase activity localized in mitochondria. Extracellular O generated with xanthine oxidase did not affect aconitase activity nor did exogenous superoxide dismutase decrease aconitase inactivation by PMS. The results demonstrate a dynamic and cyclical O-mediated inactivation and iron-dependent reactivation of the mammalian [4Fe-4S] aconitases under normal and stress conditions and provide further evidence for the membrane compartmentalization of O.


INTRODUCTION

Several growth impairments caused by excess superoxide radical (O) production (1, 2, 3, 4, 5, 6) or SOD() deficiencies (5, 6, 7) in Escherichia coli can be attributed to the O-mediated inactivation of [4Fe-4S]-containing (de)hydratases (5, 6, 8, 9, 10, 11, 12, 13, 14) . Loss of sensitive enzymes including ,-dihydroxyacid dehydratase (8, 11, 12, 15) , 6-phosphogluconate dehydratase (5) , aconitase (6, 9, 10) , and fumarases A and B (13, 14) collectively impedes branched-chain amino acid synthesis, the Entner-Doudoroff pathway, and Krebs' citric acid cycle and may also cause pleiotropic effects (16, 17) . Knowledge of these and other sensitive targets (18) continues to illuminate mechanisms of toxicity and the adaptive defense against O.

A balanced mechanism of O-dependent inactivation and iron-dependent reactivation modulates aconitase activity and citric acid cycle activity in E. coli(9, 10) . Similar inactivation-reactivation mechanisms may control the activity of other [4Fe-4S]-dependent metabolic pathways (5, 8, 11, 12, 19) , and regulators of transcription (20) and translation (21-26). The conservation of in vitro O sensitivities (6, 9, 15, 27) , reactivatabilities (9, 10, 28) , and primary structures (29, 30, 31) between the mammalian mitochondrial and cytoplasmic aconitases and the prokaryotic E. coli aconitase led us to suppose that similar dynamic inactivation-reactivation of the aconitases may occur in mammalian cells and, moreover, that these reactions may be important for aconitase function in the citric acid cycle and in the control of cellular iron metabolism (21) under normal and stress conditions.

We now describe a O-dependent inactivation and iron-dependent reactivation of the mammalian aconitases and describe the use of aconitases as measures of O produced in the mitochondria and cytoplasm of cultured mammalian cells. The possible roles of a dynamic inactivation-reactivation in aconitase function are discussed.


MATERIALS AND METHODS

Cells and Reagents

The human epithelial lung cell carcinoma A549 (CCL 185), the murine fibrosarcoma L929 (CCL 1), and human A375 melanoma cells (CRL 1619) were obtained from the American Type Culture Collection (Rockville, MD). Rat 1A fibroblasts were provided by G. Johnson of this department. MnSOD overexpressing A375 clones, D5 and D11, were constructed by transfecting A375 cells with the expression vector pcDNAI/neo (Invitrogen, San Diego) containing the 1.0-kilobase EcoRI human MnSOD cDNA fragment from pSP65-HMS (32) ligated into the EcoRV site in the sense orientation; and the control clone, neo5, was created by transfecting cells with pcDNAI/neo.() Stable transfectants were selected in medium containing 450 µg/ml Geneticin, and independently-derived clones were screened for elevated MnSOD by measuring activity in cell lysates (34) . PMS, antimycin A, FCCP, barium (±)-fluorocitrate, fetal calf serum, bovine milk xanthine oxidase, horse heart cytochrome c, nitro blue tetrazolium, riboflavin, and hydrogen peroxide were from Sigma. Beef liver catalase (260,000 units/ml) and myxothiazole were obtained from Boehringer Mannheim.Bovine erythrocyte Cu,ZnSOD (3500 units/mg) was from DDI Pharmaceuticals, Inc., Mountain View, CA. Deferoxamine mesylate was from CIBA, Summit, NJ. F12K and RPMI 1640 growth media, Hank's balanced salt solution, trypsin-EDTA, gentamycin sulfate, Geneticin, and penicillin-streptomycin were obtained from Life Technologies, Inc. Acetylated cytochrome c was prepared as described (35) .

Cell Growth, Harvest, and Extract Preparation

Cell cultures were grown at 37 °C under a humidified atmosphere containing 5% CO in 10-cm Falcon dishes and cells were routinely counted with a hemacytometer. A549, Rat 1A, and L929 cell cultures were grown in F12K medium containing 10% fetal calf serum, with 100 units/ml penicillin and 100 µg/ml streptomycin. Exposures to normobaric hyperoxia (pO of 600 mm Hg) were as described previously (36). A375 cell clones, neo5, D5, and D11 were grown in RPMI 1640 medium containing 10% fetal calf serum, 10 µg/ml gentamycin sulfate, and 225 µg/ml Geneticin, and A375 parental cells were grown in the absence of Geneticin. Cells were harvested and extracts were prepared as described previously (36) .

Enzyme and Protein Assays

Aconitase activity was measured as described previously (36) using a Beckman DU-64 spectrophotometer equipped with a thermostatted and oscillating multicuvette holder. SOD activity was assayed in a 0.5-ml assay mixture containing 10 µM acetylated-cytochrome c, 50 µM xanthine, 1 mM EDTA, and 200 units/ml catalase in 50 mM potassium phosphate buffer, pH 7.8, at 25 °C. Xanthine oxidase was added to produce an initial increase in absorbance at 550 nm of 0.0095 ± 0.03/min. SOD units were calculated from a standard curve prepared with bovine erythrocyte Cu,ZnSOD using McCord-Fridovich units (37) . SOD activities were calculated from the inhibition of cytochrome c reduction between 30 and 60%, a range corresponding to 0.02-0.1 units of SOD activity. Cell extracts were dialyzed against four changes of 100 volumes of 50 mM potassium phosphate, pH 7.8, and 0.1 mM EDTA at 4 °C. MnSOD activity was measured in dialyzed extracts in which Cu,ZnSOD was inactivated by prior incubation with 50 mM diethyl dithiocarbamate for 60 min at 30 °C (38) . Protein concentration was measured by the method of Bradford (39) using Coomassie Brilliant Blue staining reagent (Bio-Rad), and bovine serum albumin, fraction V (Calbiochem, La Jolla, CA), as the standard. Lactate dehydrogenase and glutamate dehydrogenase activities were assayed at 25 °C by following the change of absorbance at 340 nm in a 1-ml reaction mixture containing 50 mM potassium phosphate buffer, pH 7.4, plus 0.2 mM NADH with either 1 mM sodium pyruvate, or 2 mM -ketoglutarate and 50 mM NHCl, added as the respective substrates.

Respiration Measurements

Adherent cell cultures were washed with Dulbecco's phosphate-buffered saline (1.1 mM KHPO4, 8.1 mM NaHPO, 138 mM NaCl, 2.7 mM KCl, 0.5 mM MgCl, 0.9 mM CaCl) and gently trypsinized. Cells were washed with media and pelleted at 1000 g for 10 min. Oxygen consumption was measured using a Clark-type oxygen electrode. Assays were performed at 37 °C in a 2.0-ml volume of the original growth medium containing 0.6-1.4 10 cells with respiratory effectors as indicated. Rates of cyanide-resistant respiration (40) were measured in the presence of 0.25 mM sodium cyanide. An O saturation of 167 µM at 37 °C (41) corrected for Denver atmospheric pressure of 635 mm Hg was used for calculations of respiration rates.

Subcellular Fractionation

Six to ten 10-cm dishes of near confluent cultures (0.5-1.0 10 cells/dish) were washed with 5 ml of ice-cold phosphate-buffered saline, scraped in 5 ml of phosphate-buffered saline, and centrifuged in 15-ml conical tubes at 1500 g for 60 s. Cells were pooled and washed in 1.0 ml of ice-cold fractionation buffer containing 0.25 M sucrose, 10 mM Tris-HCl, pH 7.4, 0.1 mM EDTA, 2 mM sodium citrate, and 1 mM sodium succinate and were centrifuged at 1500 g for 60 s. Cells were resuspended in 0.5 ml of fractionation buffer and homogenized on ice in a Teflon-glass Duall tissue homogenizer (Kontes Glass Co., Vineland, NJ) by making 6-7 passes at a maximum speed of 500 rpm at 4 °C. Cell lysates were centrifuged for 5 min at 1000 g to pellet nuclei, plasma membranes, unruptured cells, and other debris away from the supernatant containing cytoplasm and mitochondria. Mitochondria were then separated from the cytoplasmic components by centrifugation at 10,000 g for 10 min. The mitochondrial pellet was gently overlaid with 0.1 ml of ice-cold fractionation buffer and centrifuged at 10,000 g for 2 min to remove residual cytoplasmic protein, and the wash was pooled with the cytoplasmic fraction. The mitochondria were resuspended in 0.1 ml of ice-cold fractionation buffer, and both fractions were sonicated for 10 s and clarified by centrifugation at 14,000 g for 30 s. Aconitase activity was assayed immediately since delay or freezing of extracts under these conditions resulted in the loss of activity. Extracts were stored at -70 °C, and the respective cytoplasmic and mitochondrial marker enzymes, lactate dehydrogenase and glutamate dehydrogenase, were assayed to evaluate fractionations. The amount of lactate dehydrogenase in the mitochondrial fraction, and glutamate dehydrogenase in the cytoplasmic fraction, was 16% of the total in each cell type.

Gel Electrophoresis and SOD Activity Staining

Samples containing 10 µg of protein were loaded in a well of a Universal 8 (Ciba-Corning Diagnostics, Inc.) thin film agarose gel pre-equilibrated with running buffer containing 20 mM Tris glycine, pH 8.2, and 1 mM EDTA. Gels were electrophoresed at 100 V and 2-3 mA for 60 min and were stained for SOD activity using the method of Beauchamp and Fridovich (42) .

Anoxic Reactivation of Aconitase

Cells were harvested by gentle trypsinization, washed with medium, and resuspended at 4 10 cells/ml in the original growth medium with cycloheximide added at 100 µg/ml. Septum sealed vacutainer tubes (Becton-Dickinson, 47 10 mm) were filled with 2.6 ml of cells, and the excess volume was allowed to escape through a syringe needle as the rubber septum was inserted. Cells in vacutainer tubes were incubated for 60 min in a 37 °C water bath and were manually agitated to maintain a uniform cell suspension and to achieve anaerobic conditions. Aerobic incubations were performed in parallel by adding cells to 10-cm dishes containing 15 ml of the original medium at 37 °C with 100 µg/ml cycloheximide. Vacutainer tubes were centrifuged at 7000 g in a microcentrifuge for 2 min, media was aspirated, and the cell pellet was lysed by sonicating for 10 s in 200 µl of cold lysis buffer containing 50 mM Tris-Cl, pH 7.4, 0.6 mM MnCl, and 20 µM fluorocitrate. Cell lysates were immediately frozen in dry-ice ethanol and stored at -70 °C.

Exposure of Cells to Xanthine Oxidase and HO

A549 cells were grown to a density of 4-5 10 cells per 10-cm dish and were washed twice with Hank's balanced salt solution at 37 °C. Washed cell monolayers were incubated at 37 °C for 15 min in 5.0 ml of Hank's balanced salt solution containing 0.5 mM xanthine and 500 units/ml catalase with or without 25 milliunits of xanthine oxidase, or washed cells were treated with the indicated concentrations of HO in Hank's balanced salt solution. Cycloheximide was present at 100 µg/ml throughout the exposures. Rates of O production by xanthine oxidase were estimated by following the initial rate of cytochrome c reduction at 550 nm under mock exposure conditions with 10 µM cytochrome c. HO concentrations were measured spectrophotometrically (43) .

Data Analysis

The Tukey-Kramer (HSD) statistical analysis method in the program JMP (SAS Institutes Inc.) was used for the analysis of significance (p < 0.05) between multiple pairs.


RESULTS

Inactivation of Aconitase by PMS

A variety of natural and synthetic quinones, viologens, anthracyclines, and phenazine pigments increase cellular O and HO production by diaphorase-catalyzed redox-cycling mechanisms (40) . We surveyed several redox-cycling agents for their ability to increase O production in cultured mammalian cells as measured by their effects on cyanide-resistant respiration (CRR) and on aconitase activity. Both are reliable indices of O production in E. coli(9, 40) . PMS was effective in rapidly increasing CRR and inactivating aconitase and was therefore selected for further studies. As shown in Fig. 1A, PMS exposure elicited a concentration-dependent increase in the CRR of cultured human epithelial A549 lung cells. Increases in CRR occurred within a few seconds of PMS addition, and rates were linear for at least 10 min. PMS increased CRR rates at concentrations in human cells similar to those required in E. coli(40) . Measurements of aconitase activity in A549 cells after a 120-min exposure to PMS revealed proportional decreases in aconitase activity with increases in CRR rates (Fig. 1B), and >90% of the total cellular aconitase activity was sensitive to PMS-mediated inactivation. Unlike the rapid increases in CRR rates, inactivation of aconitase was progressive during the initial 60 min of exposure to 1.0 µM PMS and then subsequently plateaued (Fig. 1C), suggesting a relatively slow response of aconitase activity to elevated O levels upon PMS exposure. Similar concentration and time-dependent effects of PMS on CRR rates and aconitase activity were observed in human A375 (neo5) melanoma cells.


Figure 1: Effect of PMS on cyanide-resistant respiration and aconitase activity in A549 cells. Panel A, A549 cells grown to a density of 4-5 10 cells per 10-cm dish were measured for CRR in the presence of 0.25 mM sodium cyanide and the indicated concentrations of PMS. The respiration rate without cyanide was 35.7 ± 4.7 nmol/min/10 cells. Panel B, A549 cells grown to a density of 4-5 10 cells/dish were treated with the indicated concentrations of PMS for 120 min in the presence of 100 µg/ml cycloheximide; or Panel C, cells were exposed to 1.0 µM PMS and harvested at intervals, and were assayed for aconitase activity. MeSO, the solvent control for PMS, was added to a concentration of 0.1% (v/v). Data represent the average ± S.D. of three independent exposures.



Inactivation of Aconitase by Inhibitors of Ubiquinol-cytochrome c Oxidoreductase

Respiratory inhibitors offered a specific tool for eliciting O production in the mitochondria and for investigating the interactions of O and aconitase in mitochondria of intact cultured mammalian cells. Elevated O production can be elicited by treating respiring submitochondrial particles with the high affinity ubiquinol-cytochrome c oxidoreductase inhibitor antimycin A. Protonophores, including FCCP, stimulate O production by antimycin A-supplemented mitochondria by >13-fold (44, 45, 46, 47) .

Thus, we tested the effects of various combinations of these ubiquinol-cytochrome c oxidoreductase inhibitors on aconitase activity in several mammalian cell types. As shown by Fig. 2, blocking mitochondrial respiration with antimycin A caused modest but significant (p < 0.05) decreases in aconitase activity in all cell types tested. The amount of inactivation ranged from 14% for A549 cells to 38% for L929 cells. Uncoupling mitochondria with FCCP stimulated respiration by 2.6-, 1.3-, and 3.3-fold in A549, Rat 1A, and L929 cells, respectively. However, FCCP alone did not significantly decrease aconitase activity in any cell type. The results are consistent with the observation that FCCP, and increased respiration, does not increase O production in isolated mitochondria (44, 46) . The combination of antimycin A and FCCP decreased aconitase activity dramatically to 73.6 ± 1.6, 50.5 ± 0.4, and 32.1 ± 2.3% (average ± S.D.; n = 3) of its control activity in A549, Rat 1A, and L929 cells, respectively. These decreases were significant (p < 0.05) relative to measurements with antimycin A alone as well as those of untreated controls. Myxothiazole is a potent inhibitor which blocks electron transfer at a site upstream of the antimycin A site in the ubiquinol-cytochrome c oxidoreductase (48) and which diminishes O production by antimycin A-supplemented mitochondrial membranes (49) . As anticipated, myxothiazole mitigated aconitase inactivation by antimycin A and FCCP exposure in all cell types (Fig. 2). By itself, myxothiazole caused a modest loss of aconitase activity in all cells possibly by affecting the release of O from the NADH dehydrogenase flavoprotein or from other reduced upstream sources (44, 50) .


Figure 2: Effect of ubiquinol-cytochrome c oxidoreductase inhibitors on aconitase activity in mammalian cells. A549, Rat 1A, and L929 cells were grown to 90% confluence in F12K medium and were exposed to 4 µM FCCP, 4 µM myxothiazole (Myx), 0.5 µM antimycin A (Anti A) individually or in various combinations in the presence of 100 µg/ml cycloheximide. After a 60-min exposure, cells were harvested, extracts were prepared and assayed for aconitase activity. The efficacy of inhibitors was verified by measurements of cell respiration rates as described under ``Materials and Methods.'' Ethanol was present as the solvent control at a concentration 0.06% (v/v) in all exposures. 100% aconitase activity equaled 6.4 ± 0.0, 6.0 ± 0.1, and 18.1 ± 0.3 in A549, Rat 1A, and L929 cells, respectively. Data were normalized to the control values and represent the average ± S.D. of three independent exposures. Results are representative of two or more experiments. The asterisks indicate p < 0.05 relative to the control value.



These results demonstrate the diminution of aconitase activity under conditions in which O production is elevated. Since approximately 5.5-fold increases in O production have been measured in intact isolated mitochondria treated with antimycin A (44, 47) , it seems probable that the depression of total aconitase activity by 14-38%, observed upon exposure of cultured cells to antimycin A (Fig. 2), was caused by increases in mitochondrial O production of a similar magnitude.

Elevated Mitochondrial MnSOD Protects Aconitase against Inactivation by PMS or Antimycin A, but Not by Hyperoxia

MnSOD activity was measured in A375 cell clones stably transfected with expression vector alone, or expression vector containing the human MnSOD cDNA. Control clone neo5 contained 0.8 ± 0.1 units of MnSOD/mg of protein and the independently-derived MnSOD overproducing clones D5 and D11 contained 12.8 ± 1.0 units of MnSOD/mg and 13.9 ± 1.1 units of MnSOD/mg of protein (average ± S.D.; n = 3), respectively. Both clones D5 and D11 produced MnSOD activity approximately 15-17-fold over control levels and expressed most of this MnSOD activity in their mitochondria as demonstrated by subcellular fractionation and SOD gel analysis (Fig. 3A). Clones neo5, D5, and D11 were used to investigate further the role of O in aconitase inactivation by PMS, and antimycin A plus FCCP. As shown in Fig. 3B, antimycin A plus FCCP or PMS exposure decreased aconitase activity in the control neo5 clone by 54.6 ± 2.8 and 52.2 ± 5.5% (average ± S.D.; n = 3), respectively. In comparison, the A375 clones D5 and D11 which overproduce mitochondrial MnSOD were significantly more resistant to aconitase inactivation by antimycin A plus FCCP or PMS exposure in all cases (Fig. 3B). These results directly implicate O in the inactivation of mitochondrial aconitase by PMS or antimycin A, and they suggest that mitochondrial MnSOD plays an important role in preventing this inactivation under normal conditions.


Figure 3: Effect of elevated mitochondrial MnSOD on aconitase inactivation in A375 cells. Panel A, cell mitochondria and cytoplasm were fractionated from six 10-cm dishes of A375 clones neo5, D5, and D11 grown to a density of 8 10 cells per dish. Cytoplasmic (C) and mitochondrial (M) extracts (10 µg/lane) were separated by agarose gel electrophoresis and stained for SOD activity. A375 clones neo5, D5, and D11 were grown to 6-8 10 cells per 10-cm dish and were treated with 100 µg/ml cycloheximide and no additions (), 0.5 µM antimycin A plus 4 µM FCCP (), or 1.0 µM PMS () for 60 min (Panel B) and were assayed for aconitase activity. Percent aconitase activity was normalized to the appropriate control cell aconitase activity. 100% aconitase activity for controls equals to 2.5 ± 0.1, 3.3 ± 0.1, and 3.1 ± 0.1 milliunits/mg of protein in neo5, D5, and D11, respectively. Data are the average ± S.D. of three independent exposures. Asterisks indicate p < 0.05 when compared to corresponding neo5 values.



We recently reported the loss of >80% of the aconitase activity in A549 cells and a 73% loss of aconitase in rat lungs after a 24-h exposure to normobaric hyperoxia (36) , and we hypothesized that this was due to the effect of hyperoxia-induced mitochondrial O. Interestingly, mitochondrial MnSOD did not protect aconitase against inactivation during a 24-h exposure of the MnSOD overexpressing A375 clones to hyperoxia (pO = 600 mm Hg); aconitase was inactivated to 24.5 ± 0.6, 24.2 ± 2.0, and 29.9 ± 3.3% (average ± S.D., p > 0.05 for all pairs; n = 3 per condition) of the air control activity in neo5, D5, and D11 cells, respectively. The failure of MnSOD to protect suggests an O-independent mechanism for aconitase inactivation under hyperoxic conditions possibly involving direct attack by dioxygen.

Membrane Compartmentalization of Aconitase Activity and SOD Activity

We examined the subcellular distribution of aconitase activity since cell-type differences in the localization of aconitase activity to mitochondria may account for cell-type differences in aconitase inactivation by antimycin A plus FCCP ( Fig. 2 and Fig. 3B). As shown in Fig. 4A, total aconitase activity varied considerably from a low of 1.9 ± 0.1 milliunits/mg of protein in A375 (neo5) cells to a high of 16.5 ± 0.5 milliunits/mg of total cell extract protein in L929 cells. There was also significant variation in the mitochondrial and cytoplasmic distribution of aconitase activity. The mitochondrial aconitase specific activity was markedly higher than the cytoplasmic aconitase activity in L929 cells (Fig. 4A) in which 82.5 ± 0.7% (average ± S.D.; n = 2) of the total aconitase activity fractionated with the mitochondria (Fig. 4B). A549 cells did not show an enrichment of aconitase activity within mitochondria. Only 37.6 ± 4.7% (average ± S.D.; n = 3) of the total aconitase activity fractionated with the mitochondria in A549 cells. A375 (neo5) cells and Rat 1A cells more closely resembled L929 cells than A549 cells in that their aconitase activity was enriched in the mitochondrial fraction (Fig. 4, A and B). These results may explain the greater inactivation of total aconitase activity in L929, Rat 1A, and A375 (neo5) cells than in A549 cells since O produced inside the mitochondrial matrix may not have access to the 62% cytoplasmic fraction of aconitase activity in A549 cells.


Figure 4: Cytoplasmic and mitochondrial aconitase activities in A549, A375 (neo5), Rat 1A, and L929 cells. Aconitase activity and protein were measured in cytoplasmic, mitochondrial, and whole cell extracts (Panel A), and the percent mitochondrial aconitase activity was calculated from the total of mitochondrial plus cytoplasmic aconitase activities for each cell type (Panel B). The data in Panel A show the average ± S.D. and are representative data of two or three determinations for each cell line. The data in Panel B represent the average ± S.D. of two or three independent subcellular fractionations for each cell line. The asterisk indicates p < 0.05 relative to other cell lines.



We also measured the total SOD and mitochondrial MnSOD activity in cells since differences in aconitase inactivation may be, at least in part, due to differences in the amount of SOD. While total SOD activities were remarkably similar at 2.8 ± 0.2 units/mg (average ± S.D.), the MnSOD activites varied 1.6 ± 0.2 (A549), 0.8 ± 0.1 (A375 clone neo5), 0.4 ± 0.1 (Rat 1A), and 0.9 ± 0.1 (L929) units/mg (average ± S.D.; n = 3). It is interesting to note that while no gross differences in total SOD activity were detected in these different cell types, A549 cells contained 2-4-fold more MnSOD activity per total cell protein. Thus, higher mitochondrial MnSOD activity in A549 cells may contribute to the greater resistance of A549 cells to aconitase inactivation during exposure to antimycin A plus FCCP.

Dynamic Iron-dependent Reactivation of Aconitase in Vivo

To determine whether inactivated aconitase was reactivated upon removal of the O insult, PMS-treated A549 cell monolayers were washed and aconitase activity was measured at intervals in the presence of cycloheximide. Cells recovered 50% of the lost activity within 12 min, and 95% within 75 min (Fig. 5A, line 1). The half-time for reactivation of aconitase (12 ± 3 min) estimated from line 1 was slower than that determined for the E. coli aconitase (3-5 min) (9, 10) . Reactivation in E. coli was shown to be dependent upon iron. The importance of available iron for mammalian aconitase reactivation is demonstrated by the effects of the avid iron chelator deferoxamine on aconitase inactivation, reactivation rates, and total recovery. Simultaneous exposure of cells to PMS and deferoxamine for 30 min (Fig. 5A, line 2) resulted in increased aconitase inactivation, and subsequent incubation of cells with deferoxamine slowed the reactivation rate and decreased the maximum aconitase activity recovery. Preincubation of cells with deferoxamine (60 min), to allow its cellular uptake, caused an even more profound impairment of aconitase reactivation (Fig. 5A, line 3). Exposure to deferoxamine under steady-state conditions also produced a slow but progressive loss of cellular aconitase activity relative to that seen in cells exposed to cycloheximide alone (Fig. 5B, compare lines 1 and 2). Deferoxamine also enhanced O-mediated aconitase inactivation during a 2-h exposure of A375 clone neo5 to antimycin A (Fig. 6A), and A375 clones D5 and D11 producing elevated MnSOD were more resistant to deferoxamine-enhanced inactivation. Elevated MnSOD provided a significant 15-17% protection of aconitase activity during a prolonged 5.5-h incubation with deferoxamine (Fig. 6B). This experiment reveals the constant inactivation of aconitase by endogenously produced O under otherwise normal growth conditions. Iron-dependent reactivation thus appears to be a primary mechanism to maintain aconitase activity since even prolonged inhibition of protein synthesis with cycloheximide was comparatively without effect on aconitase activity in A549 and A375 cells (Fig. 5B and Fig. 6B).


Figure 5: Effect of deferoxamine on aconitase reactivation in A549 cells. Panel A, A549 cells (4-5 10 cells/dish) were exposed to 1.0 µM PMS for 30 min. Cells were washed twice with 10 ml of F12K medium (37 °C) and incubated in 10 ml of fresh medium at 37 °C, and the recovery of aconitase activity was measured at intervals in cells exposed and maintained either in the absence (line 1) or presence of 200 µM deferoxamine (line 2) or in cells treated with 200 µM deferoxamine for 60 min prior to PMS exposure and maintained with deferoxamine throughout the incubation (line 3). Cycloheximide (100 µg/ml) was present during all exposures and incubations. Panel B, A549 cells were exposed to cycloheximide alone (line 1) or with 200 µM deferoxamine (line 2), and aconitase activity was measured at intervals. Percent aconitase activity values are normalized to the control value at the time of PMS exposure. Data are the average ± S.D. of three independent exposures.




Figure 6: Effect of deferoxamine on O-mediated aconitase inactivation in A375 cells. A375 clonal cell lines neo5, D5, and D11 were grown in RPMI 1640 medium to 6-8 10 cells/dish and treated with 100 µg/ml cycloheximide. A, cells were incubated alone (), with 0.5 µM antimycin A (), or with antimycin A plus 200 µM deferoxamine () for 120 min. B, cells were incubated with () or without () 200 µM deferoxamine for 5.5 h and assayed for aconitase activity. Data represent the mean ± S.D. of three independent exposures. Aconitase activity is normalized to 100% and equals 2.8 ± 0.0, 3.5 ± 0.1, and 3.6 ± 0.1, and 2.5 ± 0.1, 2.7 ± 0.1, and 2.8 ± 0.1 milliunits/mg of protein (average ± S.D.; n = 3) for neo5, D5, and D11, in panels A and B, respectively. Asterisks indicate p < 0.05 relative to corresponding neo5 value.



A 10-15% inactive, O-sensitive fraction of aconitase was measured in E. coli during aerobic growth (9, 10) . This inactive, but activable, fraction of aconitase presumably reflects the steady-state balance of inactivation-reactivation. An inactive fraction of aconitase of 14% was also measured in A549 cells under normal aerobic incubation conditions as shown by the increase in the specific activity of aconitase from 5.40 ± 0.04 to 6.30 ± 0.09 milliunits/mg (average ± S.D., p < 0.05; n = 3) in cells incubated anaerobically in the presence of cycloheximide as described under ``Materials and Methods.''

Together the results demonstrate a significant contribution of O to aconitase inactivation under steady-state conditions and demonstrate the importance of the balanced iron-dependent reactivation of aconitase for the maintenance of aconitase activity in cultured mammalian cells.

Effects of Extracellular O and HOon Intracellular Aconitase Activity

The O-sensitive aconitase provided, for the first time, a sensitive method for measuring how much O passes through the membrane of intact cells and for assessing the effect of extracellular [O] on intracellular [O]. O produced at a relatively high rate (2 nmol min ml for 15 min) with xanthine oxidase and xanthine did not significantly affect total aconitase activity in A549 cells. After exposure to xanthine oxidase-generated O, treated and control cells contained 4.84 ± 0.23 and 5.18 ± 0.12 milliunits/mg (n = 3; ± S.D., p > 0.05) aconitase activity, respectively. For comparison, this amount of xanthine oxidase-generated O would be roughly equivalent to that released by 5 10 activated neutrophils applying a rate of 2 nmol of O min per 10 neutrophils (51) . Moreover, Cu,ZnSOD (200 units/ml) added extracellularly did not protect aconitase activity during a 60-min exposure to 1.0 µM PMS. PMS inactivated aconitase to 31.3 ± 0.3% in the absence and to 29.4 ± 0.7% (average ± S.D., p > 0.05; n = 3) in the presence of added Cu,ZnSOD. The results suggest that the rate of passage of O into or out of the plasma membrane does not significantly affect intracellular or intramitochondrial [O] in intact cells.

In comparison, HO, which freely permeates biological membranes, did inactivate aconitase in A549 cells during a 15-min exposure. HO added in bolus concentrations of 10, 50, and 100 µM caused a concentration-dependent decrease of aconitase activity to 94.4 ± 1.4, 92.5 ± 1.1, and 42.5 ± 2.4% (average ± S.D., p < 0.05 for all pairs; n = 3) of the control activity, respectively. However, scavenging HO with exogenously added catalase (1000 units/ml) did not affect the inactivation of aconitase during exposure of the cells to PMS (1.0 µM; 60 min), thus HO appears to have a minimal role in PMS-mediated aconitase inactivation. Aconitase activity in catalase-treated and control cells was 28.5 ± 0.4 and 31.3 ± 0.3% (average ± S.D., p > 0.05; n = 3), respectively.


DISCUSSION

We have demonstrated a dynamic mechanism of O-dependent inactivation and iron-dependent reactivation of aconitase in mammalian cells. Aconitase was inactivated by conditions which increase the production of O (Fig. 1-3) and was protected from inactivation by MnSOD (Fig. 3B). Inactivated aconitase was rapidly reactivated, and reactivation was dependent upon available iron as evidenced by the inhibition of reactivation by the iron chelator deferoxamine (Fig. 5A and 6A). Moreover, overexpression of MnSOD partially protected against the loss of aconitase activity in the presence of deferoxamine (Fig. 6B), thus revealing the constant rate of O-mediated inactivation of aconitase occurring at physiological O levels. A 14% inactive fraction of aconitase activity was detected in growing A549 cells and presumably reflects the steady-state balance of inactivation-reactivation under normal growth conditions. Our results do not rule out the participation of other potentially important oxidants such as HO or O in aconitase inactivation. However, the requirement of comparatively high exposure levels in vivo and relatively poor reactivities in vitro(6, 15) suggest a lesser role of O and HO in the inactivation of aconitase under normal growth conditions.

The effects of O and iron on the cytoplasmic and mitochondrial aconitases cannot be precisely discriminated from measurements of total cell aconitase activity. Nevertheless, our results indicate that both cytoplasmic and mitochondrial aconitases are inactivated by O and subsequently reactivated in vivo. Thus, PMS inactivated >90% of the total aconitase activity in A549 cells. Since 62% of the total activity in A549 cells is located to the cytoplasm, we can conclude that the cytoplasmic aconitase was inactivated. On the other hand, the inactivation of aconitase by mitochondrion-specific generators of O in several cell types ( Fig. 2and Fig. 3B) and the protection against this inactivation by mitochondrial MnSOD (Fig. 3B) demonstrates an O dependent inactivation of the mitochondrial aconitase.

Our results are consistent with a limited diffusibility of O across membranes (52) and a compartmentalization of O. Thus, high levels of extracellular O or extracellular SOD failed to modulate aconitase activity in the cytoplasm or mitochondria of A549 cells. Furthermore, the fractional loss of total aconitase by antimycin A/FCCP-elicited mitochondrial O production ( Fig. 2and 3B) correlated with the enrichment of aconitase activity in mitochondria, suggesting O production and compartmentalization within the mitochondria. Correction for the fraction of aconitase localized to mitochondria (Fig. 4B) reveals similar losses of mitochondrial aconitase of 77 ± 8% (average ± S.D.) in the four cell lines examined.

An aconitase inactivation-reactivation cycle showing the opposing roles of O and iron in modulating aconitase activity is depicted in Fig. SI. Active mitochondrial aconitase, and presumably other homologous aconitases, contain [4Fe-4S] clusters which are attacked and oxidized by endogenous O with estimated second order rates of 8 10M s to 3 10M s (15, 27) to form inactive aconitases containing stable oxidized [3Fe-4S] clusters. Release of the solvent exposed iron atom in the ferrous state, cluster oxidation, and formation of HO are thought to occur during the inactivation process (15) . Regeneration of the active aconitase [4Fe-4S] cluster is efficiently achieved in vivo with a pseudo-first order rate of 0.0014 s (t = 12 min) (Fig. 5A) presumably by the univalent reduction of the oxidized [3Fe-4S] cluster to form a [3Fe-4S] cluster (28, 53, 54) and subsequent insertion of a ferrous ion from a deferoxamine-sensitive iron pool.


Figure SI: Scheme I



A balanced steady-state aconitase inactivation-reactivation cycle (Fig. SI) can be described by Equation 1 (see below) where the rate of aconitase inactivation equals the rate of aconitase reactivation and when the availability of iron and other reactants is assumed to remain constant. We can thus ideally solve for [O] from the inactive and active fraction of aconitase, and the rate constants for inactivation (k) and reactivation (K = [Fe] k) as shown by Equation 2. Applying the in vitro estimates of k(15, 27) and a value of 0.0014 s for K, we calculate [O] = 50-200 pM when aconitase is 50% active, and [O] = 8-30 pM under normal conditions when aconitase is 14% inactive and 86% active. These rough approximations of [O] are in the range of the [O] of 8 pM estimated from rates of O production and dismutation in mitochondria (55). We can also see from this relation that the aconitase reactivation rate must be 6 times greater than the rate of aconitase inactivation to maintain 86% active aconitase and, moreover, that the half-time for aconitase activity turnover due to oxidative inactivation at a normal balance of 86% active aconitase equals 6 12 or 72 min. Thus, the steady-state inactivation-reactivation cycle of aconitase turns over relatively rapidly (t = 72 min) when compared with aconitase turnover due to aconitase degradation and synthesis (Fig. 5B and 6B) (56) . Without absolute rate constants to apply in vivo, relative changes in steady-state [O] can be estimated from the change in the ratio of inactive and active aconitase under X and Y conditions as described by Equation 3. For example, by correcting for the fraction of aconitase inactivated in mitochondria of cells exposed to antimycin A/FCCP, and by applying an estimated 14% inactive fraction of mitochondrial aconitase during normal conditions, we calculate 20-40-fold increases in steady-state mitochondrial [O]. Furthermore, we can estimate that the 15-17-fold elevation of mitochondrial MnSOD in the MnSOD overexpressing A375 clones decreased mitochondrial aconitase inactivation by 60% (Fig. 3) and the corresponding mitochondrial [O] by 12-fold. It is expected that decreases in available [Fe] will alter the balance of inactivation-reactivation (Equation 1) by decreasing the rate of reactivation but not the rate of inactivation. Indeed, this effect can explain the greater loss of aconitase activity in cells treated with deferoxamine and the protection against this loss by elevated MnSOD (Fig. 6).

On-line formulae not verified for accuracy

On-line formulae not verified for accuracy

On-line formulae not verified for accuracy

The cyclical inactivation-reactivation of aconitase(s) may modulate aconitase function(s) under stress and normal conditions. The citric acid cycle flux capacity and homeostatic regulation of iron metabolism may be affected during oxidative stress (24, 36) , nutritional or immune-induced iron deficiencies, or SOD deficiencies. In addition, the control of the aconitase iron-sulfur state by intracellular [O] and [Fe] may also facilitate iron-sensing by the RNA-binding cytoplasmic aconitase during homeostasis (21, 22, 25, 26) . Interestingly, the opposing effects of iron and O on aconitase, and other [4Fe-4S]-containing (de)hydratases, can explain the previously puzzling and paradoxical benefit of MnSOD up-regulation during iron limitation in E. coli(9) . It may also provide a rationale for the increase in mitochondrial MnSOD synthesis (57) which accompanies increased ferritin synthesis (58) and hypoferremia (33) during the inflammatory cytokine-mediated immune response in mammals.

Further explorations of the effects of O on aconitase(s), the citric acid cycle, and cellular iron metabolism may shed light on the complex interactions of O and iron in normal and pathological states. In addition, the aconitase(s) can now be utilized as versatile compartmentalized sensors for the assay of O.


FOOTNOTES

*
This work was supported in part by a grant from the American Lung Association (to P. R. G.) and National Institutes of Health Grants HL46481 (to C. W. W.), CA27903, and CA44446 (to L. B. E.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§
Supported by National Institutes of Health Postdoctoral Training Grants T32 HL07670 and F32 HL08997 and an American Heart Association of Colorado Fellowship. To whom correspondence should be addressed: National Jewish Center for Immunology and Respiratory Medicine, D301, 1400 Jackson St., Denver, CO 80206. Tel.: 303-398-1402; Fax: 303-398-1851.

Supported by the Swiss National Foundation and Ciba Geigy Jubilaeumsstiftung.

**
Recipient of an Established Investigator Award sponsored by the Colorado Affiliate of the American Heart Association.

The abbreviations used are: SOD, superoxide dismutase; MnSOD, manganese-containing superoxide dismutase; Cu,ZnSOD, copper- and zinc-containing superoxide dismutase; CRR, cyanide-resistant respiration; PMS, phenazine methosulfate; FCCP, carbonyl cyanide p-trifluoromethoxyphenylhydrazone.

I. Raineri, T.-T. Huang, C. J. Epstein, and L. B. Epstein, manuscript submitted for publication.


ACKNOWLEDGEMENTS

We thank Dr. Ting-Ting Huang for assistance in the preparation of the MnSOD overexpressing A375 clones and Dr. Ye-Shih Ho for kindly providing human MnSOD cDNA. We thank Drs. Irwin Fridovich, Anne Gardner, and Kamuda Das for careful reading of the manuscript and for suggestions.


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