Non-redox IronâSulfur Proteins Sensitive to Reactive Species

Article pubs.acs.org/accounts

Cite This: Acc. Chem. Res. XXXX, XXX, XXX−XXX

Aconitases: Non-redox Iron−Sulfur Proteins Sensitive to Reactive Species Laura Castro,*,†,‡ Veroń ica Toŕ tora,†,‡,§ Santiago Mansilla,†,‡ and Rafael Radi*,†,‡

Downloaded via UNIV OF SOUTHERN INDIANA on July 23, 2019 at 13:36:38 (UTC). See https://pubs.acs.org/sharingguidelines for options on how to legitimately share published articles.

†

Departamento de Bioquímica, Facultad de Medicina, Universidad de la República, Av. General Flores 2125, 11800 Montevideo, Uruguay ‡ Centro de Investigaciones Biomédicas (CEINBIO), Facultad de Medicina, Universidad de la República, Av. General Flores 2125, 11800 Montevideo, Uruguay § Departamento de Educación Médica, Facultad de Medicina, Universidad de la República, Av. General Flores 2125, 11800, Montevideo, Uruguay CONSPECTUS: Mammalian aconitases (mitochondrial and cytosolic isoenzymes) are unique iron−sulfur cluster-containing proteins in which the metallic center participates in the catalysis of a non-redox reaction. Within the cubane iron−sulfur cluster of aconitases only three of the four iron ions have cysteine thiolate ligands; the fourth iron ion (Feα) is solvent exposed within the active-site pocket and bound to oxygen atoms from either water or substrates to be dehydrated. The catalyzed reaction is the reversible isomerization of citrate to isocitrate with an intermediate metabolite, cis-aconitate. The cytosolic isoform of aconitase is a moonlighting enzyme; when intracellular iron is scarce, the complete disassembly of the iron−sulfur cluster occurs and apo-aconitase acquires the function of an iron responsive protein and regulates the translation of proteins involved in iron metabolism. In the late 1980s and during the 1990s, cumulative experimental evidence pointed out that aconitases are main targets of reactive oxygen and nitrogen species such as superoxide radical (O2•−), hydrogen peroxide (H2O2), nitric oxide (•NO), and peroxynitrite (ONOO−). These intermediates are capable of oxidizing the cluster, which leads to iron release and consequent loss of the catalytic activity of aconitase. As the reaction of the Fe−S cluster with O2•− is fast (∼107 M−1 s−1), quite specific, and reversible in vivo, quantification of active aconitase has been used to evaluate O2•− formation in cells. While •NO is modestly reactive with aconitase, its reaction with O2•− yields ONOO−, a strong oxidant that readily leads to the disruption of the Fe−S cluster. In the case of cytosolic aconitase, it has been seen that H2O2 and •NO promote activation of iron responsive protein activity in cells. Proteomic advances in the 2000s confirmed that aconitases are main targets of reactive species in cellular models and in vivo, and other post-translational oxidative modifications such as protein nitration and carbonylation have been detected. Herein, we (1) outline the particular structural features of aconitase that make these proteins specific targets of reactive species, (2) characterize the reactions of O2•−, H2O2, •NO, and ONOO− and related species with aconitases, (3) discuss how different oxidative post-translational modifications of aconitase impact the different functions of aconitases, and (4) argue how these proteins might function as redox sensors within different cellular compartments, regulating citrate concentration and efflux from mitochondria, iron availability in the cytosol, and cellular oxidant production.

■

tumor target cells.3 In the same decade, the nature of the arginine-derived metabolite was identified as nitric oxide (•NO) produced by enzyme catalyzed reactions by the family of nitric oxide synthases.4 Meanwhile, the citric acid cycle enzyme aconitase was identified as a main mitochondrial target of •NO or •NO-derived metabolites in target cells.5 Superoxide radical (O2•−), which is formed by both enzymatic and nonenzymatic reactions in all aerobic organisms, has long been proposed to be an important species in oxygen toxicity, but mechanistic data of direct O2•− targets were scarce until mitochondrial aconitase (m-aconitase) and

INTRODUCTION Mammalian aconitases (mitochondrial and cytosolic isoenzymes) are iron−sulfur proteins that catalyze the reversible isomerization of citrate to isocitrate via cis-aconitate and participate in iron homeostasis. These proteins contain a [4Fe−4S]2+ prosthetic group in which one of the irons is not ligated to a protein residue and thus can bind to oxygen atoms of the substrates or water. Oxidation of the cluster promotes iron release with formation of a catalytically inactive [3Fe− 4S]+ cluster; on the other hand, in the case of cytosolic aconitase (c-aconitase), complete cluster removal activates its function as an iron responsive protein.1,2 Seminal works from Hibbs and co-workers in the 1980s showed that an arginine-derived metabolite from activated macrophages induced mitochondrial dysfunction in cocultured © XXXX American Chemical Society

Received: March 24, 2019

A

DOI: 10.1021/acs.accounts.9b00150 Acc. Chem. Res. XXXX, XXX, XXX−XXX

Article

Accounts of Chemical Research

Figure 1. Mitochondrial aconitase structure. The left side shows the tridimensional structure of Sus scrofa (pig) heart m-aconitase, colored by domain (1 in yellow, 2 in purple, 3 in blue, 4 in cyan, hinge linker in pink). The Fe−S cluster is indicated with a circle. The right side shows the [4Fe−4S]2+ cluster of aconitase within its active site, ligated to protein cysteines by Fe−S bonds.

acids, just after this activity was disengaged from the oxidative decarboxylation of isocitrate to α-ketoglutarate in an experiment performed by Martius in 1937. In the same year, the term “aconitase” was coined by Breusch. The purified m-aconitase was very unstable in aqueous solutions, as its activity tended to decay rapidly, but when other cellular components were retained enzyme activity was preserved. In 1951 stabilization of the pure enzyme was achieved by addition of ferrous ion in combination with a reducing agent. In 1970, Eanes and Kun, were able to separate the m- and c-aconitases in pig tissue as they differed on its isoelectric point while catalyzing the same reaction. They also noticed a difference in stability, being the isolated c-aconitase less labile to inactivation (reviewed in refs 1 and 20). In 1983, Kennedy defined the best conditions for purified aconitase activation consisting in the anaerobic incubation of the enzyme with a source of ferrous ion and a reducing agent like dithiothreitol. Under aerobic conditions, enzyme activity can be measured as aconitase is protected from inactivation by binding of its substrates to the Fe−S cluster.1,21 m-Aconitase (EC 4.2.1.3) is an 83 kDa protein encoded by the nuclear gene ACO2 (chromosomal location 22q13.2 for human). The enzyme is expressed with a 27 amino acid long peptide on its N-terminus responsible for its transport into the mitochondrial matrix. This peptide serves as a mitochondrial targeting sequence, being an amphipathic helix with an overall positive charge, which can also be recognized as a substrate for the mitochondrial processing peptidase. It has been suggested, at least in yeast, that the C-terminal domain might have a “chaperone-like function” toward the N-terminal domain, modulated by the cytosolic Hsp70, important for efficient aconitase import into mitochondria.22 The most characterized m-aconitase is the Sus scrofa (pig) heart aconitase. The crystallographic structure of pig maconitase containing its cubane [4Fe−4S]2+ cluster was first solved by Robbins and Stout in 1988 using X-ray crystallography (PDB 6ACN) with a resolution value of 2.5 Å. Before that, they had solved the inactive enzyme ([3Fe− 4S]+ cluster) with a 2.1 Å resolution (PDB 5ACN), thus providing direct evidence for an Fe−S cluster interconversion,1 confirming previous analytical, biochemical, and spectroscopic evidence, which support that m-aconitase Fe−S cluster can be present in different states (reviewed in ref 20). m-Aconitase is a monomeric globular heart-shaped protein, folded in four domains (Figure 1). Domain one contains residues 1−200, domain 2 contains residues 201−319, domain 3 contains residues 320−512, and domain 4 contains residues 537−754. The segment containing residues 513−536 is termed hinge

other Fe−S cluster-containing dehydratases were detected as direct O2•− targets6−8 in the early 1990s. Augmented mitochondrial O2•− formation through the electron transport chain or matrix dehydrogenases9 will shift the reactivity of •NO toward O2•−, because the reaction becomes kinetically favored, leading to peroxynitrite (ONOO−) formation.10 Peroxynitrite is a strong oxidant formed by the diffusion-controlled reaction between the signaling radical •NO and O2•−.11 Actually, in the middle 1990s it was shown that even though •NO can cause a transient inhibition of electron transport components such as complexes I and IV, the inhibition pattern of mitochondrial respiration observed in the presence of authentic ONOO− is the one that closely resembles that found secondary to •NO interactions with intact cells and strongly pointed to ONOO− as the ultimate reactive intermediate accounting for •NOdependent inactivation of electron transport components and ATPase in living cells and tissues.12,13 Indeed, ONOO−, but not •NO, rapidly reacts with the Fe−S cluster of m-aconitase yielding an inactive [3Fe−4S]+-protein14,15 being later further confirmed.16,17 In mammals, a cytosolic form of aconitase is expressed, the product of a different gene from its mitochondrial counterpart and having an important role in iron homeostasis. Cytosolic aconitase is an iron responsive protein (IRP-1), a translational regulator for iron status in the cell, acting as a trans-element to the cis-acting sequences termed iron-responsive elements (IREs) present in the untranslated regions (UTRs) of transferrin receptor mRNA and of ferritin H−L chains, among others. Low intracellular levels of iron shift cytosolic aconitase to IRP-1 activation. In addition, there is still another group of factors that contribute to this switch of activity; among these factors, •NO and hydrogen peroxide (H2O2) were postulated to inhibit c-aconitase activity and increase IRE binding activity in vivo.18,19 Herein, we will outline structural aspects of aconitases that make these proteins selective targets of reactive species, disclose the proximal intermediates leading to aconitase Fe−S disassembly under biologically relevant conditions, and envision the biological significance of these oxidative modifications in the context of cell metabolism.

■

STRUCTURE OF ACONITASES: Fe−S CLUSTER-CONTAINING DEHYDRATASES m-Aconitase was first described as an “enzyme system” that catalyzed the interconversion of citric, cis-aconitic, and isocitric B


Article


Figure 2. Graphical representation of the reaction catalyzed by aconitase. Aconitase binds citrate stereospecifically and removes a hydroxyl group and a hydrogen (*) on a dehydration reaction where His101 acts as the acid and Ser642 as the base. Thus, a water molecule is released, and cisaconitate is formed binding to the Fe−S cluster in a “citrate mode” as the dehydrated substrate was citrate. The cis-aconitate is released, and then a new molecule of cis-aconitate binds the cluster in an “isocitrate mode” and is hydrated on a complementary hydration reaction leading to isocitrate formation.1,30 Only one of the structures depicted corresponds to an existing crystal structure, whereas the others were modeled based on mechanistic evidence. Adapted with permission from ref 23. Copyright 1999 Wiley and Sons

Figure 3. Aconitase−IRP-1 interconversion regulates cell iron homeostasis. Under abundant cellular iron conditions, c-aconitase contains a [4Fe− 4S]2+ cluster and catalyzes the isomerization of citrate to isocitrate as does the mitochondrial counterpart (left side). When cellular iron scarce, caconitase loses its cluster, and the apoprotein acquires the function of the iron responsive protein-1 (IRP-1). IRP-1 controls the expression of several proteins regulating its translation by mRNA stabilization or mRNA repression (right side). Proteins involved in IRP-1-dependent regulation and its function are depicted in the scheme.

directed mutagenesis and isotopic labeling of the substrate and solvent in Mössbauer, EPR, and ENDOR experiments1,20,23 (Figure 2). As an alternative function of aconitase unrelated to citrate dehydration, it was seen at least in yeast that m-aconitase is able to stabilize mtDNA by direct binding, protecting it from damage through control of replication, recombination, or repair.24 In a human lung adenocarcinoma cell line, overexpression of m-aconitase was found to protect mtDNA from oxidative damage, although no evidence of binding to any mammalian mtDNA has been found yet.25 The cytosolic isoform of the aconitase (ACO1, c-aconitase) is encoded by a different nuclear gene than the m-isoform (chromosomal location 9p21.1 in humans). This isozyme contains a [4Fe−4S]2+ cluster when active, but in the apo

linker, and the possibility has been suggested of a hinge motion around this region. The third domain contains three cysteine residues that bind the [4Fe−4S]2+ cluster.1 While 3 irons of the cluster coordinate to cysteines and inorganic sulfurs, one iron coordinates only with inorganic sulfurs. This iron, termed alpha (Feα), is exposed to solvent and acts as the binding site for the hydroxyl leaving group of a substrate to be dehydrated (Figure 1). The absence of a fourth cysteine ligand and an overall positive charge (2+) of the cluster makes it accessible and vulnerable to anionic one electron oxidants (see later).23 The multistep reaction mechanism of aconitase is wellknown (Figure 2), as the enzyme has been cocrystallized with substrates and competitive inhibitors and different reaction models have been tested against kinetic data.1 The assays performed to elucidate the mechanism include also siteC


Article

Accounts of Chemical Research form, the enzyme functions as IRP-1, binding to IREs, which are highly conserved sequences found in the 5′ or 3′ UTRs of hairpin loops of mRNA molecules related to iron metabolism, like ferritin and the transferrin receptor. Under iron depleted conditions, IRPs bind to IREs. When binding in the 5′-UTR, the IRP prevents translation of mRNA, while binding to the 3′UTR protects the mRNA from degradation.2 The binding of IRP-1 to the IREs occurs during cellular iron starvation, whereas the opposite scenario (c-aconitase active−IRP-1 inactive) develops when iron in the intracellular transit pool is plentiful18 (Figure 3). In humans, c-aconitase is 135 amino acids larger than the mitochondrial one, weighing 98 kDa, and they share a 43% sequence similarity (Q99798 and P21399 in UniProt database). In spite of this, the active site and the fold of the protein are highly conserved. Both isozymes fold in four globular domains, with a linker joining domains 3 and 4 with the Fe−S cluster close to the center of the enzyme and ligated to protein cysteines of domain 3. The extra amino acids of c-aconitase introduce differences at the protein surface. c-Aconitase has extra α-helices and β-sheets distributed particularly on the Nterminal region (domain 1), putatively involved in IRE binding.26

Figure 4. Aconitase cluster cycle of oxidative disruption and reductive reactivation. Anionic oxidants such as superoxide radical, peroxynitrite, and carbonate radical react fast with the [4Fe−4S]2+ cluster of aconitase with second order rate constant ranging from 105 to 108 M−1 s−1 yielding an inactive [3Fe−4S]-containing enzyme. During this process an iron ion (Feα) is released as Fe2+ and secondary oxidants are produced. Reactivation of the enzyme can be achieved in vitro or in vivo by addition of Fe2+ and a reductant that provides an additional electron (e−) such as glutathione. The formation of H2O2 and HCO3̅ requires H+ from the media, and the reaction of peroxynitrite yield H2O in addition to •NO2.

of natural mitochondrial iron chelators that may increase upon aconitase inactivation (i.e. citrate, glutamylcitrate, ADP), iron protein chaperones (e.g., frataxin), and H2O2-metabolizing systems (e.g., GSH peroxidase and peroxiredoxin 3) would largely limit the formation of •OH under most physiologically relevant conditions, except under situations of severe mitochondrial dysfunction and oxidative damage. Repair of the oxidatively disassembled Fe−S aconitase cluster occurs through univalent reduction and iron reinsertion both in vitro and in vivo29 (Figure 4). For E. coli and mammalian cell aconitases, reactivation of the Fe−S cluster was achieved with pseudo-first-order rates of 0.004 s−1 (t1/2 = 3 min) and 0.0014 s−1 (t1/2 = 12 min) respectively.8 The reactivation rate constant reflects a process that involves three species in which iron and glutathione are assumed to be constant. As both the inactivation and reactivation constant rates were determined, a measure of the steady-state concentration of O2•− within E. coli was estimated to be approximately 20−40 pM in aerobic log phase and increased to 300 pM in an E. coli mutant strain lacking SOD.8 This approach was also used to measure O2•− steady-state concentration in mammalian cell cytosol and mitochondria.29 The selective targeting for O2•− has led to aconitases being used as sensitive endogenous indicators for O2•− formation within different cell compartments. For instance, determination of aconitase activity has become an assay for measuring the variation in O2•− levels inside cells and mitochondria in a wide variety of cell types and conditions.30,31 Using this approach and as an example, we have determined that bovine endothelial cells exposed to hyperglycemic conditions exhibited an 8.8-fold increase in the mitochondrial O2•− steady-state concentration with respect to normoglycemic conditions. Furthermore, under hyperglycemic conditions, a mitochondrial O2•− steady-state concentration of 250 pM was estimated.31 Hydrogen peroxide, produced directly or by enzymatic dismutation of O2•−, also reacts with the [4Fe−4S]2+ cluster of aconitase with smaller rate constants than O2•−, around 102 M−1 s−16 (Figure 4). In intact cardiac mitochondria, H2O2 induced formation of the [3Fe−4S]+ form of aconitase as determined by EPR as well as isoelectric focusing native gel electrophoresis.32 In addition, aconitase amino acid carbonylation and increase in protein degradation was observed

■

LABILITY OF THE Fe−S CLUSTER AND REACTION WITH SUPEROXIDE RADICAL In E. coli, exposure to high oxygen concentrations or superoxide dismutase (SOD) deficiencies produced profound growth impairment due to O2•−-dependent inactivation of the [4Fe−4S]2+ cluster-containing dehydratases7,8 including aconitases. In vitro experiments using E. coli dihydroxy-acid dehydratase, fumarases A and B, and beef heart aconitase confirmed that cubane Fe−S clusters containing a Feα are rapidly inactivated by O2•−.6 Superoxide-mediated inactivation of [4Fe−4S] 2+ dehydratases results in iron release from the enzyme and promotes the formation of a paramagnetic [3Fe− 4S]+ cluster as detected by low temperature EPR.6 Enzyme substrates afford protection against O2•−-mediated inactivation. In addition, reactivation of the enzymes is achieved by addition of Fe2+ and a reductant, conditions that lead to cluster reconstitution.6−8 Overall, the experimental evidence suggested that the oxidative disassembly of the iron−sulfur cluster by O2•− was similar to that described for the aconitase [4Fe−4S]2+ center by O2 and ferricyanide21,27 (Figure 4). Second-order reaction rate constants for the reaction of mammalian m-aconitases with O2•− were determined to be in the range of 106 to 107 M−1 s−1.6,14 In aconitases and other 4Fe−4S-containing dehydratases, O2•− leads to the formation of an inactive [3Fe−4S]+ cluster (Figure 4). Feα (as Fe2+) is released, and H2O2 is likely to be formed (Figure 4), which together facilitate the formation of hydroxyl radical (•OH) by the Fenton reaction.6 Indeed, in vitro experiments with purified bovine heart mitochondria exposed to xanthine and xanthine oxidase (an enzymatic source of O2•− and H2O2) confirmed the formation of a paramagnetic [3Fe−4S]+ cluster detected at 10−15 K and revealed the formation of •OH by EPR spin trapping experiments with 5-diethoxyphosphoryl-5-methyl-1pyrroline N-oxide and α-phenyl-N-tert-butylnitrone.28 A scenario in which •OH may be continuously generated in the mitochondria was postulated,28 although the relevance of this source in oxidant-mediated damage is presumably modest and remains to be decisively established; indeed, the presence D


Article


withdrawing capacity of the Feα, thus diminishing NO+ formation and reactivity. Indeed, using pure preparations of recombinant porcine aconitase, we reconfirmed that a reversible [4Fe−4S]−•NO complex is formed by short exposures at low concentrations of •NO. However, sustained • NO production slowly promotes an irreversible disassembly of the Fe−S cluster due to NO+ reaction with the nucleophilic sulfur of the [4Fe−4S]2+ center, leading to irreversible enzyme inactivation.17 An overall second order rate constant of 0.65 M−1 s−1 for the •NO-dependent inactivation m-aconitase was determined; this rather slow inactivation probably reflects a more complex process that involves more than one constant for different reaction steps.17 The •NO-dependent irreversible inactivation was also confirmed by others39 for mammalian and E. coli aconitase. Taken altogether, the accumulated results from different groups support that the exposure of bovine maconitase to high concentrations of •NO for extended periods of time leads to the formation of protein-bound dinitrosyl− iron−dithiol complex along with other transient EPR signals, in both the presence and absence of substrates, ultimately leading to irreversible inactivation of the enzyme.40 In spite of the slow and multistep reactivity of •NO toward m-aconitase, ONOO−, the product of the diffusion-controlled reaction of •NO with O2•−, inactivates m-aconitase much faster (k = 1.4 × 105 M−1 s−1).14,15,17 Mitochondria are the main intracellular sites of O2•− production in complex I and III of the mitochondrial respiratory chain and matrix dehydrogenases.9 Unlike the highly diffusible •NO in biological milieu, O2•− diffuses very short distances; therefore ONOO− is typically formed close to the O 2 •− production sites, mitochondrial matrix being a preferential site of ONOO− generation.41 The direct reaction of the [4Fe−4S]2+ metal center of aconitase with ONOO− leads the enzyme cluster to a [3Fe−4S]+ form that can be reactivated both in vitro and in vivo14,17,40,42 in the presence of Fe2+ and a reducing agent17 (Figure 4). The role of glutathione in providing protection against ONOO−-mediated aconitase inactivation and promoting reactivation was evidenced in cultured fibroblasts in which GSH depletion or cellular clones with lower GSH content exhibited significantly enhanced sensitivity for aconitase inactivation due to ONOO−.37 Peroxynitrite-mediated aconitase inactivation becomes also noticeable in the presence of CO2, a reagent that readily reacts with ONOO− to yield carbonate radical (CO3•−) and •NO2.11 Indeed, CO3•− reacts rapidly with the Fe−S cluster of aconitase yielding the [3Fe−4S]+ form (k = 3 × 108 M−1 s−1)17 (Figure 4). In addition to inactivating aconitase due to cluster disruption, ONOO− can also cause oxidative post-translational modifications in the protein moiety; among these, ONOO− promotes aconitase tyrosine nitration secondary to the formation of radical species such as CO3•− and •NO2,17,42 which leads to the formation of protein 3-nitrotyrosine.11 Actually, nitrated aconitase has been identified as an oxidatively modified mitochondrial protein in proteomic analysis of animal models of sepsis, diabetes, amyotrophic lateral sclerosis, and aging (reviewed in ref 41). Moreover, the preferentially nitrated tyrosines were mapped in isolated pig heart m-aconitase as Tyr151 and -472; both tyrosines are adjacent to the active site, possibly reflecting a metal-catalyzed site-specific formation of •NO2.11 Nevertheless, nitration of maconitase does not affect the enzyme activity, as saturating substrate concentration completely protects against ONOO−-

following H2O2 treatment of intact mitochondria. Increase in citrate concentration protected aconitase and promoted its reactivation upon H2O2 treatment.32 While the reassembly (and reactivation) of the iron−sulfur cluster has been seen as a nonenzymatic process, in 2004 it was postulated that frataxin, one of the components involved in the maturation of cellular iron−sulfur proteins, was involved in repairing oxidatively inactivated [3Fe−4S]+ aconitase.33 This work reported that following H2O2 exposure to mitochondria from either Saccharomyces cerevisiae or rat heart in the presence of citrate, aconitase immunoprecipitation rendered frataxin as an associated protein. In addition, oligomeric frataxin was able to convert inactive [3Fe−4S]+ aconitase to the active [4Fe− 4S]2+ enzyme in vitro. However, co-immunoprecipitation of frataxin with m-aconitase could not be reproduced later,34 even when ISCU, NFS1, ISD11, and MPP (other protein components of the iron−sulfur machinery synthesis) were present, thus challenging the experiments described in ref 33. Yet, frataxin levels are directly related to aconitase activity in various cellular models including fibroblasts from Friedrich ataxia or in mice that exhibited low levels of frataxin by modulation of the transcription factor hypoxia inducible factor 2α35,36 due to cluster biosynthesis impairment and subsequently minor aconitase quantity. Overall, we still have limited understanding of the frataxin−aconitase interactions, which are probably more complex than other readily observed frataxin interactions with partner proteins and thus require further studies and characterization.

■

ACONITASE REACTION WITH NITRIC OXIDE AND NITRIC OXIDE-DERIVED SPECIES In the decade of the 1990s, a controversy was generated about the capacity of •NO to promote direct inhibition of maconitase, as a result of papers that published disparate and potentially contradictory results.14,15,37−40 In 1994, two sideby-side papers from Fridovich’s and our group were published in the same issue of the Journal of Biological Chemistry and concluded that m-aconitase was rapidly inactivated by ONOO− but not by its precursor, •NO.14,15 For instance, data from our group revealed that exposure of 30 μM commercial preparations of porcine m-aconitase to 100 or 200 μM authentic •NO for 25 min lead only to 20% or 40% inactivation of the enzyme, respectively. The inhibitory effect of •NO was reversed by its displacement with argon,14 suggesting reversible nitrosylation at the Feα. Nevertheless, afew years later another group reported that purified heart bovine aconitase40 was inhibited by •NO solution at a •NO/ enzyme ratio of 65:1, in the absence of substrate, 56% activity remained at 5 min and 25% at 60 min.40 Concomitantly, EPR signals with g = 2.02 corresponding to the [3Fe−4S]+-clusters and g ≈ 2.04 corresponding to a protein-bound dinitrosyl− iron−dithiol complex appear.40 Previously, it was shown that macrophage aconitase was also inhibited by •NO.38 To justify some of the differences observed, a criticism about the purity of the aconitase preparations used in 1994 (e.g., contaminating heme compounds) was elaborated.40 It is clear that •NO has high affinity for heme and non-heme iron. In aconitase, the Lewis acid character of the Feα is expected to withdraw electrons from •NO to produce an intermediate with NO+-like (nitrosonium-like) reactivity. Ligands for the Feα, such as substrates, can compete with • NO and may affect binding either via steric hindrance or the E


Article


Figure 5. Fates of m-aconitase upon oxidative post-translational modifications in cells. m-Aconitase can be inactivated by superoxide and other oxidants such as peroxynitrite and hydrogen peroxide (1). These oxidants promote the formation of an inactive [3Fe−4S]+ aconitase, but this oxidation can be reversed in intracellular conditions by iron and a reducing agent such as glutathione or cysteine (2). Depending on the state of aconitase, the citrate/isocitrate ratio can change and the flow of metabolites into the citric acid cycle would be controlled. Therefore, the oxygen consumption rate (JO2) as well as the production of reactive oxygen species (ROS) by mitochondria would be modulated. Aconitase amino acids are also targets of reactive species (3) leading to nitrated, glutathionylated, and carbonylated enzyme, among other possible post-translational modifications. Another possible fate of oxidatively modified aconitase is the formation of protein aggregates produced by formations of disulfide or dityrosine bonds (4). Oxidized aconitase can be recognized and degraded by Lon protease (5).

inactivated state of the enzyme. The S-nitrosothiol-mediated inactivation could not be recovered even in the presence of iron and sulfur in a reducing environment, supporting that GSNO can modify some of the amino acids of the enzyme, possibly some of the cysteines that coordinate the metal cluster.17 The role of GSNO mediated m-aconitase inactivation was also suggested by experiments using GSH-depleted fibroblasts which were protected from the inactivation induced by •NO-donors as compared to control cells.37

mediated inactivation while 3-nitrotyrosine is still observed; moreover, ONOO−-inactivated aconitase can recover 100% of its activity after incubation with Fe2+ and a reductant even when the enzyme remains nitrated.17 The fate of tyrosine nitrated proteins in vivo remains to be resolved, the Londependent degradation being a possibility for oxidatively modified m-aconitase in vivo (Figure 5) as it was seen to be preferably degraded by the proteasome in vitro.43 Taking in consideration both kinetic rate constants of ONOO− reactions with mitochondrial targets and the intramitochondrial target concentrations, we have recently estimated that most of the ONOO− generated in the matrix will be catalytically detoxified by peroxiredoxins 3 and 5.11 However, a fraction of ONOO− and, presumably, CO3•− will continuously react and inactivate aconitase. Whether a significant portion of reversibly inactivated m-aconitase accumulates will depend on the interplay of oxidant exposure levels and cluster reassembly. The feasibility of these processes is substantiated by the detection of inactive aconitase under a variety of oxidative stress conditions in vivo. Importantly, moderate levels of m-aconitase inactivation can have profound consequences on the control of mitochondrial metabolism (see below). Besides ONOO−, other •NO-derived species such as Snitrosothiols can inactivate m-aconitase. S-Nitrosothiols are able to transfer a NO+ group into proteins,44 and in the case of aconitase, the NO+ group could be transferred into the [4Fe− 4S]2+ cluster. In the case of S-nitrosoglutathione (GSNO), we have determined that it reacts with aconitase with a second order rate constant of 0.23 M−1 s−1, leading to an irreversibly

■

SIGNIFICANCE OF THE [4Fe−4S]2+ MITOCHONDRIAL ACONITASE CLUSTER AS A REDOX SENSOR The high susceptibility of m-aconitase to reversible inactivation by different reactive oxygen and nitrogen species poses it as a conceivable redox sensor of the citric acid cycle and controller of O2•− production by mitochondria. Hence, m-aconitase offers a hinge among different cellular processes such as iron homeostasis, aerobic respiration, fatty acid biosynthesis, or even cell death.45 In order to elucidate the metabolic impact of oxidantdependent aconitase inhibition over the citric acid cycle, the respiratory chain reactions, and reactive species formation, we reported a limited metabolic analysis using isolated mitochondria from different rat tissues with different metabolic profiles.46 The flux control coefficient elicited by aconitase over the citric acid cycle and the respiratory chain was determined by evaluating the impact of aconitase inhibition by fluorocitrate, a competitive aconitase inhibitor, on these processes. It was observed that aconitase inhibition thresholds F


Article


Figure 6. m-Aconitase Fe−S cluster state and its impact on metabolism. When m-aconitase is mostly in its [4Fe−4S]2+ cluster form, citrate in mitochondria will be consumed mainly by the citric acid cycle (A). Oxidant-dependent inactivation of m-aconitase leads to Fe−S cluster oxidation and its [3Fe−4S]+ form and enzyme inactivation (B). By this mechanism, citrate accumulation in mitochondria is facilitated and its efflux promoted. Therefore, the m-aconitase Fe−S state would shift the balance between carbohydrate and fat metabolism.

acetyl-CoA), and subsequently acetyl-CoA levels increase.49 The increased entry of metabolites to the citric acid cycle potentially favors the generation of mitochondrial O2•−. Remarkably, O2•− inactivates aconitase through Fe−S cluster oxidation and consequently slows the citric acid cycle flux, down-modulating the increase in substrate supply to the electron transport chain. The previous analysis supports the concept that mitochondrial oxidants can modulate several sites of mitochondrial metabolism in a coordinated manner (Figure 6). In the state where acetyl-CoA is increased but aconitase is inhibited, citrate accumulation is promoted and might shift the balance between carbohydrate and fat metabolism (Figure 6). This metabolic shift would help to explain how an extremely O2•−-sensitive aconitase was kept during evolution.45 Supporting this statement, homozygous MnSOD knockout mice, which die within a week of birth and presented significantly decreased aconitase activity, had significant cardiomyopathy with fat accumulation in liver and muscle.50 Also, heterozygous MnSOD knockout mice are insulin-resistant,51 and mitochondria-targeted antioxidants decrease fat content and protect against insulin resistance,51 sustaining that controlled O2•− production may control the flux into the electron transport chain for oxidative phosphorylation or may channel citrate into the synthesis of fats through its effect on aconitase. A recent work from Rouault’s group showed that down regulation of primary Fe−S biogenesis scaffold protein iron−sulfur cluster assembly enzyme 2 (ISCU2) in human HEK293 cells produced a 12-fold increase in intracellular citrate content in Fe−S-deficient cells, a surge that was due to loss of aconitase activity.52 In ISCU2 deficient cells, fatty acid biosynthesis increased markedly relative to cellular proliferation rates and promoted cytosolic lipid droplet formation. Their findings highlight the remodeling of cellular metabolism that occurs during acute iron deficiency or defective Fe−S cluster assembly and support the idea that inhibition of aconitase will reprogram the metabolic flux though citrate accumulation (Figure 6). In addition, the role of m-aconitase as a regulator of metabolism and signaling was pointed out during erythropoiesis. In fact, it was seen that m-aconitase inhibition by iron

ranged from 63% to 95% of initial activity beyond which respiration decreased rapidly, depending on the tissue, brain and heart aconitase being the most sensitive (i.e., less than 5% inhibition of aconitase activity serves to control mitochondrial respiration rates in those tissues). In addition, in brain and heart, inhibition of aconitase led to a significant decrease in both mitochondrial membrane potential and H2O2 production.46 Therefore, m-aconitase could subtly modulate the flux of metabolites though the citric acid cycle and electron transport chain and diminish O2•− production by mitochondria with tissue or cell specificity (Figures 5 and 6). To operate as a redox sensor, m-aconitase has to be reversibly inactivated in vivo, as effectively occurs (Figures 4 and 5). For instance, during cardiac ischemia−reperfusion in rats, aconitase exhibited a loss and a regain of activity during cardiac reperfusion.47 Aconitase inactivation under a short reperfusion period not only would limit O2•− (and H2O2 via Mn-SOD-catalyzed dismutation) production by the electron transport chain but also prevents Ca2+ overload by decreasing the proton gradient across inner mitochondrial membrane; by these mechanisms, m-aconitase could function as a mitochondrial “circuit breaker“ for protective purposes. Also, it was shown that high-intensity sprint training in human volunteers promoted aconitase inactivation in skeletal muscle.48 Muscle aconitase inactivation promoted inhibition of muscle mitochondrial respiration as well as diminished H2O2 release from mitochondria and augmented citrate accumulation.48 Aconitase can operate as part of a coordinated redox mitochondrial regulation by modulating the flux of the citric acid cycle and mitochondrial oxidant production. In this regard, mitochondrial thiol proteins, which reacted with low (physiologically relevant) levels of H2O2 were identified by a proteomic approach and postulated to afford some kind of redox regulation in mitochondrial metabolism. One of them, pyruvate dehydrogenase kinase 2, a major regulator of pyruvate dehydrogenase complex, was revealed to be very sensitive to H2O2 by oxidation of Cys residues 45 and 392, which result in its inactivation; this process, in turn, decreases the phosphorylation of the pyruvate dehydrogenase complex (which becomes activated to catalyze the conversion of pyruvate to G


Article


capacity in intact cells when applied extracellularly or even produced from intracellular sources.37,63 H2O2-dependent activation of IRP-1 proceeds through a multistep pathway that involves an energy dependent phosphorylation process.64 Interestingly, removal of the inductive external signal of H2O2 after approximately 15 min of treatment (called induction phase) permits complete IRP-1 activation within 60 min (execution phase), which is sustained for several hours in murine fibroblasts. This contrast with the IRP-1 activation pathway by •NO and iron starvation, in which •NO-releasing drugs or iron chelators need to be present during the entire activation phase.19 While •NO and H2O2 activate IRP-1 (with concomitant caconitase inactivation) in vivo or in cell systems, strong nitrooxidative stress could promote both c-aconitase and IRP-1 inactivation due to critical thiol oxidation (Cys437, -503, and -506, which bind to the Fe−S clusters or IREs) and tyrosine nitration.65−67 In addition, the impact of oxidant-mediated reactions with components of the cytosolic iron−sulfur cluster assembly machinery would also impact c-aconitase/IRP-1 activities. In this regard, it has been proposed that an outer mitochondrial membrane (OMM) protein, mitoNEET, is responsible for reverting H2O2-dependent IRP-1 activation.68 MitoNEET, the first identified mammalian iron−sulfur protein anchored to the OMM consists of a dimer that accommodates one [2Fe−2S]+ cluster per monomer. MitoNEET [2Fe−2S]+ clusters are very stable but can be reversibly oxidized by H2O2. In vitro and cell culture experiments support that mitoNEET is able to transfer its oxidized Fe−S cluster to an apoprotein receptor such as IRP-1 (reviewed in ref 69). During acute inflammation, H2O2 and •NO-mediated IRP-1 activation in cells such as macrophages may reduce the availability of essential iron to invading bacteria, which in turn is probably beneficial for the host, reflecting a role of these mechanisms as part of innate immunity. However, in chronic inflammation, the diversion of iron traffic reduces iron availability for erythropoiesis, eventually contributing to the anemia of chronic diseases.70 Interestingly, structural motifs in the 5′-untranslated regions (UTRs) of IREs are also present in the mRNA encoding maconitase;18 therefore activation of IRP-1 by H2O2 and •NO would also impact the quantity of m-aconitase. Hence, in a scenario of augmented cellular formation of reactive species, m-aconitase would be affected by both the direct (oxidantdependent) inhibition of its activity and, eventually, decrease in its quantity (via translational regulation and degradation). Thus, the Fe−S cluster disassembly of m- and c-aconitase and the metabolic impact on carbohydrate, lipid, and iron metabolism may be intertwined and rapidly respond to changes of cellular redox homeostasis.

deprivation disturbed erythropoietin signaling pathway in erythroid progenitor cells and that isocitrate can enhance the effectiveness of erythopoietin during iron deficiency in vitro and in mice with hypoplastic anemia and in rats with anemia of chronic inflammation. These observations support that maconitase is an iron-sensing regulator of mitochondrial metabolism.53 Oxidative damage to proteins is a hallmark of the aging process and oxidized protein degradation is an important mechanism to prevent dysfunctional protein accumulation. In the housefly, the age-related accumulation of oxidized aconitase (carbonylated enzyme) has been found to be associated with a corresponding loss of activity in the flight muscles and a shortening of the lifespan of flies.54,55 Inactive aconitase seems to be a preferential target of Lon proteases, which degrade oxidized proteins in the mitochondrial matrix. Data regarding whether [4Fe−4S]2+ cluster oxidation, tyrosine nitration, or other amino acid oxidation contribute to the increase in protein degradation by Lon protease is still insufficient. As Lon protease activity declines with aging, oxidized mammalian aconitase accumulates and may contribute to age-related mitochondrial dysfunction56 (Figure 5).

■

IRP-1/CYTOSOLIC ACONITASE REACTIONS WITH REACTIVE SPECIES The Fe−S cluster of IRP-1/c-aconitase is more stable than that of m-aconitase.57 Early experiments employing immunologically stimulated murine primary peritoneal macrophages or macrophage cell lines showed that •NO biosynthesis correlates with activation of IRP1 (i.e., increase binding to IREs) and repression of ferritin mRNA translation.58,59 Further experiments using •NO-donors or stably transfected NOS2 cells confirmed •NO-dependent IRP-1 activation concomitant with changes in translation of IREs containing mRNAs (reviewed in ref 60). In vitro experiments using cytoplasmic extracts of macrophage cell lines or purified c-aconitase demonstrated that neither ONOO−, O2•−, nor H2O2 significantly induced IRP-1 activation, even though they inhibited c-aconitase activity.15,61 Interestingly SIN-1, an organic molecule that releases both • NO and O2•−, only promoted IRP-1 activation when SOD was present, supporting •NO-dependent IRP-1 activation. Furthermore, NONOates (•NO-donors) or GSNO activates IRE binding by IRP-1.61 Reaction of •NO with purified bovine c-aconitase was studied following EPR signals.40 In this study, the main species detected at 77 K was the g = 2.04 signal, typical for the dinitrosyl−iron−thiol complex, while at 10−20 K, varying intensities of the signal for the [3Fe−4S]+ cluster were also seen. In addition, a third signal was observed in some samples at both 77 and 10−20 K corresponding to the formation of a thiyl radical.40 Unfortunately, in this report only c-aconitase activity was followed without looking at the IRE binding capacity of the purified protein. Anyway, the overall data suggest that a general cluster disassembly is necessary for IRP-1 activation. Thioredoxin, a 12 kDa multifunctional protein that participates in redox reactions through dithiol− disulfide exchange reactions, seems to be necessary to enhance the RNA binding activity of •NO-treated IRP162 in cell lysates, supporting that •NO induces IRP-1 activation in a reductive medium (cytoplasmic extracts, GSH, or thioredoxin).61,62 H2O2, as already stated, does not activate IRP1 binding directly, but it induces a rapid activation of IRE binding

■

AUTHOR INFORMATION

Corresponding Authors

*L.C. E-mail addresses: [email protected]; [email protected]. *R.R. E-mail addresses: [email protected]; rafael.radi@gmail. com. ORCID

Santiago Mansilla: 0000-0002-3208-9981 Rafael Radi: 0000-0002-1114-1875 H


Article

Accounts of Chemical Research Notes

(9) Murphy, M. P. How mitochondria produce reactive oxygen species. Biochem. J. 2009, 417, 1−13. (10) Radi, R.; Cassina, A.; Hodara, R.; Quijano, C.; Castro, L. Peroxynitrite reactions and formation in mitochondria. Free Radical Biol. Med. 2002, 33, 1451−1464. (11) Ferrer-Sueta, G.; Campolo, N.; Trujillo, M.; Bartesaghi, S.; Carballal, S.; Romero, N.; Alvarez, B.; Radi, R. Biochemistry of Peroxynitrite and Protein Tyrosine Nitration. Chem. Rev. 2018, 118, 1338−1408. (12) Cassina, A.; Radi, R. Differential inhibitory action of nitric oxide and peroxynitrite on mitochondrial electron transport. Arch. Biochem. Biophys. 1996, 328, 309−316. (13) Radi, R.; Rodriguez, M.; Castro, L.; Telleri, R. Inhibition of mitochondrial electron transport by peroxynitrite. Arch. Biochem. Biophys. 1994, 308, 89−95. (14) Castro, L.; Rodriguez, M.; Radi, R. Aconitase is readily inactivated by peroxynitrite, but not by its precursor, nitric oxide. J. Biol. Chem. 1994, 269, 29409−29415. (15) Hausladen, A.; Fridovich, I. Superoxide and peroxynitrite inactivate aconitases, but nitric oxide does not. J. Biol. Chem. 1994, 269, 29405−29408. (16) Gardner, P. R. Aconitase: sensitive target and measure of superoxide. Methods Enzymol. 2002, 349, 9−23. (17) Tortora, V.; Quijano, C.; Freeman, B.; Radi, R.; Castro, L. Mitochondrial aconitase reaction with nitric oxide, S-nitrosoglutathione, and peroxynitrite: mechanisms and relative contributions to aconitase inactivation. Free Radical Biol. Med. 2007, 42, 1075−1088. (18) Fillebeen, C.; Pantopoulos, K. Redox control of iron regulatory proteins. Redox Rep. 2002, 7, 15−22. (19) Pantopoulos, K.; Weiss, G.; Hentze, M. W. Nitric oxide and oxidative stress (H2O2) control mammalian iron metabolism by different pathways. Mol. Cell. Biol. 1996, 16, 3781−3788. (20) Beinert, H.; Kennedy, M. C.; Stout, C. D. Aconitase as IronSulfur Protein, Enzyme, and Iron-Regulatory Protein. Chem. Rev. 1996, 96, 2335−2374. (21) Kennedy, M. C.; Emptage, M. H.; Dreyer, J. L.; Beinert, H. The role of iron in the activation-inactivation of aconitase. J. Biol. Chem. 1983, 258, 11098−11105. (22) Ben-Menachem, R.; Wang, K.; Marcu, O.; Yu, Z.; Lim, T. K.; Lin, Q.; Schueler-Furman, O.; Pines, O. Yeast aconitase mitochondrial import is modulated by interactions of its C and N terminal domains and Ssa1/2 (Hsp70). Sci. Rep. 2018, 8, 5903. (23) Lloyd, S. J.; Lauble, H.; Prasad, G. S.; Stout, C. D. The mechanism of aconitase: 1.8 A resolution crystal structure of the S642a:citrate complex. Protein Sci. 1999, 8, 2655−2662. (24) Chen, X. J.; Wang, X.; Butow, R. A. Yeast aconitase binds and provides metabolically coupled protection to mitochondrial DNA. Proc. Natl. Acad. Sci. U. S. A. 2007, 104, 13738−13743. (25) Kim, S. J.; Cheresh, P.; Williams, D.; Cheng, Y.; Ridge, K.; Schumacker, P. T.; Weitzman, S.; Bohr, V. A.; Kamp, D. W. Mitochondria-targeted Ogg1 and aconitase-2 prevent oxidant-induced mitochondrial DNA damage in alveolar epithelial cells. J. Biol. Chem. 2014, 289, 6165−6176. (26) Dupuy, J.; Volbeda, A.; Carpentier, P.; Darnault, C.; Moulis, J. M.; Fontecilla-Camps, J. C. Crystal structure of human iron regulatory protein 1 as cytosolic aconitase. Structure 2006, 14, 129−139. (27) Emptage, M. H.; Dreyers, J. L.; Kennedy, M. C.; Beinert, H. Optical and EPR characterization of different species of active and inactive aconitase. J. Biol. Chem. 1983, 258, 11106−11111. (28) Vasquez-Vivar, J.; Kalyanaraman, B.; Kennedy, M. C. Mitochondrial aconitase is a source of hydroxyl radical. An electron spin resonance investigation. J. Biol. Chem. 2000, 275, 14064−14069. (29) Gardner, P. R.; Raineri, I.; Epstein, L. B.; White, C. W. Superoxide radical and iron modulate aconitase activity in mammalian cells. J. Biol. Chem. 1995, 270, 13399−13405. (30) Estrada, D.; Specker, G.; Martinez, A.; Dias, P. P.; Hissa, B.; Andrade, L. O.; Radi, R.; Piacenza, L. Cardiomyocyte diffusible redox mediators control Trypanosoma cruzi infection: role of parasite

The authors declare no competing financial interest. Biographies Laura Castro obtained her M.D. and Ph.D. from Universidad de la República, Uruguay, in 1994 and 1999, respectively. She currently holds an appointment as Professor of Biochemistry at Facultad de Medicina, Universidad de la República. Her research interests include mitochondrial redox metabolism and oxidative post-translational modifications in proteins. Verónica Tórtora obtained her Ph.D. from Universidad de la República, Uruguay, in 2014 and now is an Associate Professor in the Departments of Medical Education and Biochemistry at Facultad de Medicina, Universidad de la República. Her work focuses on redox biochemistry of mitochondrial proteins. Santiago Mansilla is a Ph.D. student and Instructor of Biochemistry. His research project relates to postranslational modifications of human mitochondrial aconitase. Rafael Radi obtained his M.D. and Ph.D. from Universidad de la República, Uruguay, in 1989 and 1991, respectively. He is Professor and Chair of Biochemistry at Facultad de Medicina, Universidad de la República. He is a Foreign Associate of the US National Academy of Sciences. His research interests include free radical and redox biochemistry and its relation to human disease.

■

ACKNOWLEDGMENTS This work was supported by grants from Espacio Interdisciplinario (Centros 2015 to R.R.) and Comisión Sectorial de ́ Investigación Cientifica (CSIC I+D 2016 to L.C. and CSIC Grupos 2014 to R.R.). Additional support was obtained through Fundación Manuel Perez and from Programa de Desarrollo de Ciencias Básicas to all the authors. S.M. is partially supported by a fellowship from Universidad de la República.

■

REFERENCES

(1) Robbins, A. H.; Stout, C. D. The structure of aconitase. Proteins: Struct., Funct., Genet. 1989, 5, 289−312. (2) Rouault, T. A. The role of iron regulatory proteins in mammalian iron homeostasis and disease. Nat. Chem. Biol. 2006, 2, 406−414. (3) Granger, D. L.; Taintor, R. R.; Cook, J. L.; Hibbs, J. B., Jr. Injury of neoplastic cells by murine macrophages leads to inhibition of mitochondrial respiration. J. Clin. Invest. 1980, 65, 357−370. (4) Ignarro, L. J.; Buga, G. M.; Wood, K. S.; Byrns, R. E.; Chaudhuri, G. Endothelium-derived relaxing factor produced and released from artery and vein is nitric oxide. Proc. Natl. Acad. Sci. U. S. A. 1987, 84, 9265−9269. (5) Drapier, J. C.; Hibbs, J. B., Jr. Differentiation of murine macrophages to express nonspecific cytotoxicity for tumor cells results in L-arginine-dependent inhibition of mitochondrial iron-sulfur enzymes in the macrophage effector cells. J. Immunol 1988, 140, 2829−2838. (6) Flint, D. H.; Smyk-Randall, E.; Tuminello, J. F.; DraczynskaLusiak, B.; Brown, O. R. The inactivation of dihydroxy-acid dehydratase in Escherichia coli treated with hyperbaric oxygen occurs because of the destruction of its Fe-S cluster, but the enzyme remains in the cell in a form that can be reactivated. J. Biol. Chem. 1993, 268, 25547−25552. (7) Gardner, P. R.; Fridovich, I. Superoxide sensitivity of the Escherichia coli aconitase. J. Biol. Chem. 1991, 266, 19328−19333. (8) Gardner, P. R.; Fridovich, I. Inactivation-reactivation of aconitase in Escherichia coli. A sensitive measure of superoxide radical. J. Biol. Chem. 1992, 267, 8757−8763. I


Article

Accounts of Chemical Research mitochondrial iron superoxide dismutase. Biochem. J. 2018, 475, 1235−1251. (31) Quijano, C.; Castro, L.; Peluffo, G.; Valez, V.; Radi, R. Enhanced mitochondrial superoxide in hyperglycemic endothelial cells: direct measurements and formation of hydrogen peroxide and peroxynitrite. Am. J. Physiol Heart Circ Physiol 2007, 293, H3404− 3414. (32) Bulteau, A. L.; Ikeda-Saito, M.; Szweda, L. I. Redox-dependent modulation of aconitase activity in intact mitochondria. Biochemistry 2003, 42, 14846−14855. (33) Bulteau, A. L.; O’Neill, H. A.; Kennedy, M. C.; Ikeda-Saito, M.; Isaya, G.; Szweda, L. I. Frataxin acts as an iron chaperone protein to modulate mitochondrial aconitase activity. Science 2004, 305, 242− 245. (34) Schmucker, S.; Martelli, A.; Colin, F.; Page, A.; WattenhoferDonze, M.; Reutenauer, L.; Puccio, H. Mammalian frataxin: an essential function for cellular viability through an interaction with a preformed ISCU/NFS1/ISD11 iron-sulfur assembly complex. PLoS One 2011, 6, No. e16199. (35) Condo, I.; Ventura, N.; Malisan, F.; Rufini, A.; Tomassini, B.; Testi, R. In vivo maturation of human frataxin. Hum. Mol. Genet. 2007, 16, 1534−1540. (36) Oktay, Y.; Dioum, E.; Matsuzaki, S.; Ding, K.; Yan, L. J.; Haller, R. G.; Szweda, L. I.; Garcia, J. A. Hypoxia-inducible factor 2alpha regulates expression of the mitochondrial aconitase chaperone protein frataxin. J. Biol. Chem. 2007, 282, 11750−11756. (37) Castro, L. A.; Robalinho, R. L.; Cayota, A.; Meneghini, R.; Radi, R. Nitric oxide and peroxynitrite-dependent aconitase inactivation and iron-regulatory protein-1 activation in mammalian fibroblasts. Arch. Biochem. Biophys. 1998, 359, 215−224. (38) Drapier, J. C.; Pellat, C.; Henry, Y. Generation of EPRdetectable nitrosyl-iron complexes in tumor target cells cocultured with activated macrophages. J. Biol. Chem. 1991, 266, 10162−10167. (39) Gardner, P. R.; Costantino, G.; Szabo, C.; Salzman, A. L. Nitric oxide sensitivity of the aconitases. J. Biol. Chem. 1997, 272, 25071− 25076. (40) Kennedy, M. C.; Antholine, W. E.; Beinert, H. An EPR investigation of the products of the reaction of cytosolic and mitochondrial aconitases with nitric oxide. J. Biol. Chem. 1997, 272, 20340−20347. (41) Castro, L.; Demicheli, V.; Tortora, V.; Radi, R. Mitochondrial protein tyrosine nitration. Free Radical Res. 2011, 45, 37−52. (42) Han, D.; Canali, R.; Garcia, J.; Aguilera, R.; Gallaher, T. K.; Cadenas, E. Sites and mechanisms of aconitase inactivation by peroxynitrite: modulation by citrate and glutathione. Biochemistry 2005, 44, 11986−11996. (43) Grune, T.; Blasig, I. E.; Sitte, N.; Roloff, B.; Haseloff, R.; Davies, K. J. Peroxynitrite increases the degradation of aconitase and other cellular proteins by proteasome. J. Biol. Chem. 1998, 273, 10857−10862. (44) Stamler, J. S. S-nitrosothiols and the bioregulatory actions of nitrogen oxides through reactions with thiol groups. Curr. Top. Microbiol. Immunol. 1995, 196, 19−36. (45) Walden, W. E. From bacteria to mitochondria: aconitase yields surprises. Proc. Natl. Acad. Sci. U. S. A. 2002, 99, 4138−4140. (46) Scandroglio, F.; Tortora, V.; Radi, R.; Castro, L. Metabolic control analysis of mitochondrial aconitase: influence over respiration and mitochondrial superoxide and hydrogen peroxide production. Free Radical Res. 2014, 48, 684−693. (47) Bulteau, A. L.; Lundberg, K. C.; Ikeda-Saito, M.; Isaya, G.; Szweda, L. I. Reversible redox-dependent modulation of mitochondrial aconitase and proteolytic activity during in vivo cardiac ischemia/reperfusion. Proc. Natl. Acad. Sci. U. S. A. 2005, 102, 5987−5991. (48) Larsen, F. J.; Schiffer, T. A.; Ortenblad, N.; Zinner, C.; Morales-Alamo, D.; Willis, S. J.; Calbet, J. A.; Holmberg, H. C.; Boushel, R. High-intensity sprint training inhibits mitochondrial respiration through aconitase inactivation. FASEB J. 2016, 30, 417− 427.

(49) Hurd, T. R.; Collins, Y.; Abakumova, I.; Chouchani, E. T.; Baranowski, B.; Fearnley, I. M.; Prime, T. A.; Murphy, M. P.; James, A. M. Inactivation of pyruvate dehydrogenase kinase 2 by mitochondrial reactive oxygen species. J. Biol. Chem. 2012, 287, 35153−35160. (50) Li, Y.; Huang, T. T.; Carlson, E. J.; Melov, S.; Ursell, P. C.; Olson, J. L.; Noble, L. J.; Yoshimura, M. P.; Berger, C.; Chan, P. H.; Wallace, D. C.; Epstein, C. J. Dilated cardiomyopathy and neonatal lethality in mutant mice lacking manganese superoxide dismutase. Nat. Genet. 1995, 11, 376−381. (51) Rodriguez-Cuenca, S.; Cocheme, H. M.; Logan, A.; Abakumova, I.; Prime, T. A.; Rose, C.; Vidal-Puig, A.; Smith, A. C.; Rubinsztein, D. C.; Fearnley, I. M.; Jones, B. A.; Pope, S.; Heales, S. J.; Lam, B. Y.; Neogi, S. G.; McFarlane, I.; James, A. M.; Smith, R. A.; Murphy, M. P. Consequences of long-term oral administration of the mitochondria-targeted antioxidant MitoQ to wild-type mice. Free Radical Biol. Med. 2010, 48, 161−172. (52) Crooks, D. R.; Maio, N.; Lane, A. N.; Jarnik, M.; Higashi, R. M.; Haller, R. G.; Yang, Y.; Fan, T. W.; Linehan, W. M.; Rouault, T. A. Acute loss of iron-sulfur clusters results in metabolic reprogramming and generation of lipid droplets in mammalian cells. J. Biol. Chem. 2018, 293, 8297−8311. (53) Richardson, C. L.; Delehanty, L. L.; Bullock, G. C.; Rival, C. M.; Tung, K. S.; Kimpel, D. L.; Gardenghi, S.; Rivella, S.; Goldfarb, A. N. Isocitrate ameliorates anemia by suppressing the erythroid iron restriction response. J. Clin. Invest. 2013, 123, 3614−3623. (54) Das, N.; Levine, R. L.; Orr, W. C.; Sohal, R. S. Selectivity of protein oxidative damage during aging in Drosophila melanogaster. Biochem. J. 2001, 360, 209−216. (55) Yan, L. J.; Levine, R. L.; Sohal, R. S. Oxidative damage during aging targets mitochondrial aconitase. Proc. Natl. Acad. Sci. U. S. A. 1997, 94, 11168−11172. (56) Bota, D. A.; Van Remmen, H.; Davies, K. J. Modulation of Lon protease activity and aconitase turnover during aging and oxidative stress. FEBS Lett. 2002, 532, 103−106. (57) Kennedy, M. C.; Mende-Mueller, L.; Blondin, G. A.; Beinert, H. Purification and characterization of cytosolic aconitase from beef liver and its relationship to the iron-responsive element binding protein. Proc. Natl. Acad. Sci. U. S. A. 1992, 89, 11730−11734. (58) Drapier, J. C.; Hirling, H.; Wietzerbin, J.; Kaldy, P.; Kuhn, L. C. Biosynthesis of nitric oxide activates iron regulatory factor in macrophages. EMBO J. 1993, 12, 3643−3649. (59) Weiss, G.; Goossen, B.; Doppler, W.; Fuchs, D.; Pantopoulos, K.; Werner-Felmayer, G.; Wachter, H.; Hentze, M. W. Translational regulation via iron-responsive elements by the nitric oxide/NOsynthase pathway. EMBO J. 1993, 12, 3651−3657. (60) Kim, S.; Ponka, P. Nitric oxide-mediated modulation of iron regulatory proteins: implication for cellular iron homeostasis. Blood Cells, Mol., Dis. 2002, 29, 400−410. (61) Bouton, C.; Raveau, M.; Drapier, J. C. Modulation of iron regulatory protein functions. Further insights into the role of nitrogenand oxygen-derived reactive species. J. Biol. Chem. 1996, 271, 2300− 2306. (62) Oliveira, L.; Bouton, C.; Drapier, J. C. Thioredoxin activation of iron regulatory proteins. Redox regulation of RNA binding after exposure to nitric oxide. J. Biol. Chem. 1999, 274, 516−521. (63) Martins, E. A.; Robalinho, R. L.; Meneghini, R. Oxidative stress induces activation of a cytosolic protein responsible for control of iron uptake. Arch. Biochem. Biophys. 1995, 316, 128−134. (64) Pantopoulos, K.; Hentze, M. W. Activation of iron regulatory protein-1 by oxidative stress in vitro. Proc. Natl. Acad. Sci. U. S. A. 1998, 95, 10559−10563. (65) Bouton, C.; Hirling, H.; Drapier, J. C. Redox modulation of iron regulatory proteins by peroxynitrite. J. Biol. Chem. 1997, 272, 19969−19975. (66) Gonzalez, D.; Drapier, J. C.; Bouton, C. Endogenous nitration of iron regulatory protein-1 (IRP-1) in nitric oxide-producing murine macrophages: further insight into the mechanism of nitration in vivo J


Article

Accounts of Chemical Research and its impact on IRP-1 functions. J. Biol. Chem. 2004, 279, 43345− 43351. (67) Soum, E.; Brazzolotto, X.; Goussias, C.; Bouton, C.; Moulis, J. M.; Mattioli, T. A.; Drapier, J. C. Peroxynitrite and nitric oxide differently target the iron-sulfur cluster and amino acid residues of human iron regulatory protein 1. Biochemistry 2003, 42, 7648−7654. (68) Golinelli-Cohen, M. P.; Lescop, E.; Mons, C.; Goncalves, S.; Clemancey, M.; Santolini, J.; Guittet, E.; Blondin, G.; Latour, J. M.; Bouton, C. Redox Control of the Human Iron-Sulfur Repair Protein MitoNEET Activity via Its Iron-Sulfur Cluster. J. Biol. Chem. 2016, 291, 7583−7593. (69) Golinelli-Cohen, M. P.; Bouton, C. Fe-S Proteins Acting as Redox Switch: New Key Actors of Cellular Adaptive Responses. Current Chemical Biology 2017, 11, 70−88. (70) Mueller, S. Iron regulatory protein 1 as a sensor of reactive oxygen species. BioFactors 2005, 24, 171−181.

K


Non-redox IronâSulfur Proteins Sensitive to Reactive Species

Recommend Documents

Non-redox IronâSulfur Proteins Sensitive to Reactive Species