Iron-sulfur clusters: Formation, perturbation, and physiological functions

Plant Physiol. Biochem., 1999, 37 (2), 81-97

Review Iron-sulfur functions

clusters:

Formation,

perturbation,

and

physiological

John Imsande Department of Agronomy and Department of Zoology and Genetics, (fax +l 515 294-2299; e-mail [email protected])

(Received June 15, 1998; accepted November

Iowa State University,

Ames, Iowa 50011, USA.

17, 1998)

Abstract - Iron-sulfur proteins occur in all life forms. Ferredoxins

and Rieske proteins each contain a (2Fe-2s) cluster whereas photosystem I (PSI) contains three (4Fe-4s) clusters. Essential enzymes such as sulfite reductase, nitrite reductase, nitrogenase, glutamate synthase, aconitase, succinate dehydrogenase, ferredoxin/thioredoxin reductase, as well as many other vital proteins, each contain a (4Fe-4s) cluster. Iron-sulfur clusters are formed enzymatically from cysteinyl-sulfur and ferritin-sequestered iron. Many iron-sulfur clusters are inactivated by O2 and/or reactive oxygen species (ROS) such as O;-. Perhaps 0.1 % of the electrons passing through either the mitochondrial electron transport chain or PSI result in the formation of O;-. Many plant stresses increase ROS formation, which subsequently may perturb iron-sulfur clusters. Plants have evolved three different superoxide dismutases (SODS) to control the internal concentrations of harmful ROS. Possible roles of functional and non-functional ironsulfur clusters in the coordination of metabolic activities of stressed and non-stressed plants are discussed. 0 Elsevier, Paris

Nitrogen metabolism / oxidative stress / photosynthesis / reactive oxygen species / senescence / sulfur metabolism CPl, P700-chlorophyll a-protein 1 / FNR, fumarate nitrate reduction / FTR, ferredoxin-thioredoxin reductase / GltS, glutamate synthase / IRE, iron response element / IRP, iron response protein / nif, nitrogen fixation / PSI, photosystem I / ROS, reactive oxygen species / SDH, succinate dehydrogenase / SOD, superoxide dismutase I SoxR, superoxide response protein / SoxRS, superoxide response regulon I TIC, translocon at the inner envelope membrane of chloroplasts / TOC, translocon at the outer envelope membrane of chloroplasts

1. INTRODUCTION Iron-sulfur clusters associated with spinach ferredoxin were first characterized in the mid- 1960’s [ 13, 341. Subsequently, (Fe-S) clusters have been shown to perform many diverse roles in various biological organisms. Because (Fe-S) clusters are frequently inactivated by 0, and/or other reactive oxygen species (ROS) [53, 571, it was hypothesized that (Fe-S) clusters may be a relic of early evolution when 0, levels were low and that as the levels of 0, rose, due to the abundance of green plants, the importance of (Fe-S) clusters receded [35]. Ironically, photosystem I, a ubiquitous photosynthetic unit, contains three (Fe-S) clusters that are essential for photosynthesis and the production of 0, by plants. Experimental evidence indicates that both the activity and stability of photoPlant Physiol. Biochem., 0981-9428/99/2/O Elsevier, Paris

system I, and many other diverse proteins as well, are dependent upon intact (Fe-S) clusters. The chemistry of (Fe-S) clusters has been examined in great detail and was recently reviewed [8, 421. This review summarizes current information concerning the formation, perturbation, and physiological functions of iron-sulfur clusters. 2. TYPES OF IRON-SULFUR CLUSTERS Six different types of iron-sulfur clusters have been defined: (a) rubredoxins; (b) rhombic (2Fe-2s) clusters; (c) cuboidal (3Fe-4s) clusters; (d) cubane-type (4Fe-4s) clusters; (e) complex P-type @Fe-8s) clusters; and (f) Rieske-type (2Fe-2s) clusters [42]. The rubredoxins are low molecular mass proteins that con-

88

J. Imsande

tain one Fe ligated by four cysteines. They lack an acid-labile sulfur and hence are, at most, a rudimentary (Fe-S) cluster. The ferredoxin-type (2Fe-2s) cluster is planar and each of the irons is ligated to each of the acid-labile sulfurs. In addition, each iron is ligated individually with two cysteinyl sulfurs. The (3Fe-4s) cluster is cuboidal and each iron is ligated to three of the four acid-labile sulfurs and to one individual cysteinyl sulfur. The (4Fe-4s) cluster is a cubane with each iron ligated to three of the four sulfurs and to one individual cysteinyl sulfur. The nitrogenase cluster has been considered a (8Fe-7s) cluster or two coordinated (4Fe-4s) clusters [43]. The Rieske-type (2Fe-2s) cluster can be described as a (2cys-Fe-2S-Fe-2his) unit in which one iron is ligated by two cysteinyl sulfurs and the two acid-labile sulfur atoms whereas the second iron is ligated by the two acid-labile sulfurs and two histidinyl nitrogens [42]. 3. IRON-SULFUR

CLUSTER

FORMATION

To unravel the mysteries of the N,-fixation process, including (Fe-S) cluster assembly, many bacterial nif mutants have been isolated. A functional nifs gene, required for nitrogen fixation, produces a cysteine desulfurase (NIFS) [20, 29, 911. A protein with NIFSlike activity participates directly in (Fe-S) cluster assembly by catalyzing a two-step reaction: cysteine + NIFS + NIFS-cysteinyl persulfide + alanine NIFS-cysteinyl persulfide + Fe’+ + apo-protein + (Fe-S)-protein + NIFS Thus, cysteine is the general source of the acidlabile sulfur present in (Fe-S) clusters. Ferritin-sequestered iron can be acquired and released as needed for cluster formation [44,64]. Cysteinyl groups, and occasionally histidinyl groups, present in the apoprotein help coordinate the integration of iron and acid-labile sulfur atoms into the cluster. 4. SUPEROXIDES AND PERTURBATION OF IRON-SULFUR CLUSTERS The reactive oxygen species Oi-, H,O,, and OH’ are produced by most, if not all, aerobic organisms. In animals, the mitochondrial respiratory chain is the most significant metabolic source of the superoxide Oi-. Indeed, 1 to 2 % of oxygens encountered by the terminal oxidase of the electron transport chain are conPlant Physiol. Biochem.

verted to 0;. In plant cells, the thylakoid membranebound primary electron acceptors of photosystem I and ferredoxin are additional major sources of 0; [321. Because of the abundance, ubiquity, and chemical reactivity of O;-, plants have evolved three distinct superoxide dismutases (SODS) for its disposal [12]. Superoxide dismutases inactivate or neutralize this chemically reactive and destructive superoxide by the following reaction: 2 O;- + 2H+ + O2 +H,O, [59,60]. Hydrogen peroxide, also produced by several peroxidases, can be converted to 0, and water by either ascorbate peroxidase or catalase [31]. Enzymatic formation of the highly reactive hydroxyl radical HO’ was first reported in 1970 [5]. It was concluded that H,O, and OT, produced by the aerobic xanthine oxidase reaction, were essential reactants in the production of HO’. Subsequently, it was shown that Fe3’, present as a contaminant of the buffer salts, was a necessary catalyst in this system [58]. After examining various reactive oxygen species, Liochev and Fridovich [57] concluded that production of HO’ proceeds as follows: Fe3++ O’2 TV Fe*++ 0 2

(1)

Fe*+ + H,O, @ Fe3+ +HO- + HO’

(2)

Many enzymes which contain (4Fe-4s) clusters are inactivated by 0; [30, 421. Loss of an iron from the (4Fe-4s) cluster lowers the specific activity of aconitase more than lOO-fold [9]. For native clusters containing two Fe*+ and two Fe3+ atoms, the oxidation and inactivation by O;- is explained as follows: (2Fe2+2Fe3+-4s) + O;- + 2H+ -+ (Fe2+3Fe3+-4s) + I-W? &_ The solitary Fe*+ atom is subsequently lost from the cluster because it is less tightly bound to the sulfide ligands than are the Fe3+. Thus, oxidation of the cluster by O;- inactivates the cluster and increases the supply of free Fe*+ in the cell. Consequently, O;- facilitates HO’ production from H,O, by making Fe*+ available for the Fenton reaction (i.e. reaction 2 above) [57]. The HO’ radical is highlymutagenic. 5. FUNCTIONS

OF IRON-SULFUR

CLUSTERS

Iron-sulfur clusters play many diverse roles in the aerobic growth of cells [8, 30, 421. These roles include: (a) single electron-transfer reactions, essential in photosynthesis, respiration, and dinitrogen fixation; (b) binding and activation of substrates; (c) protein dimerization and stabilization; (d) chloroplast mem-

Iron-sulfur clusters

brane assembly and function; (e) regulation of enzyme activity; (f) regulation of gene expression; (g) posttranslational regulatory switches and cross-talk; (h) sensors of iron, oxygen, and superoxide radicals; (i) participation in oxidative stress and senescence; (j) radical production [52, 54, 791; and (k) proton transport. A partial listing of enzymes and other proteins containing (Fe-S) clusters, and the specific reactions facilitated by these proteins, are presented in tableI. 5.1. (Fe-S) clusters and genetic regulation In Escherichiu coli, two well-characterized genetic regulatory proteins contain (Fe-S) clusters, the superoxide response (SoxR) protein and the regulator for fumarate and nitrate reduction (FNR protein). Transcription of the soxRS regulon, which consists of more than ten genes, is controlled by the SoxR protein [33]. The homodimeric SoxR protein contains two ferredoxin-like (2Fe-2s) clusters that are autooxidizable, the oxidation state of which is controlled by the cellular redox status, which includes glutathione [28]. In turn, the redox state of these (2Fe-2s) clusters determines the functionality of the SoxR protein in controlling the transcription rate of the soxRS regulon, which includes glucose-6-phosphate dehydrogenase [33]. The FNR protein activates and represses a family of genes, perhaps more than 100, required for anaerobic and aerobic metabolism [39, 481. The FNR protein assists in coordinating the activities of the citric acid cycle and nitrate reduction with 0, availability [78]. Under anaerobic conditions, each subunit of the dimeric FNR protein contains a (4Fe-4S)2+ cluster. In the presence of O,, the anaerobic (4Fe-4S)2+ clusters are rapidly converted to (2Fe-2S)2+ clusters. This form of the FNR protein lacks the ability to bind DNA and is transcriptionally inactive [48]. Consequently, the FNR regulatory protein is described as an aerobicanaerobic transcriptional switch. Thus, the involvement of (Fe-S) clusters in controlling some of the global responses of an organism to environmental changes is well documented. Iron-sulfur clusters in multicelluar plants may fulfill similar regulatory functions [l, 621, because higher plants also respond to stress by increasing ROS production [4], and because many (Fe-S) clusters are sensitive to oxygen radicals [56,57]. 5.2. Coordination of general metabolic activities Three enzymes of the citric acid cycle, aconitase, succinate dehydrogenase and fumarase, may contain

89

(Fe-S) clusters, some of which may coordinate or regulate metabolic activities. Most organisms contain both a mitochondrial and a cytosolic aconitase, each of which contains a (4Fe-4s) cluster [67]. As a first step in the citric acid cycle, mitochondrial aconitase catalyzes the conversion of citrate to isocitrate via aconitate. In animals, cytosolic aconitase seemingly helps coordinate citrate metabolism with the uptake of iron via iron response elements (IREs) and iron response proteins (IRPs) [49,70]. In its native state, the cytosolic enzyme contains a (4Fe-4S)2+ cluster and has aconitase activity [9]. When one iron is lost from the cluster, enzymatic activity is reduced over lOO-fold and a reasonably stable (3Fe-4s) cluster is formed. The apoprotein lacks aconitase activity. The physiological function of the cytosolic aconitase is not well defined and may differ in plants and animals [ 14,21,40,62]. Iron, an essential nutrient and yet a potential toxin, poses an interesting regulatory problem for all living organisms [41]. In mammals, interorgan transport, uptake and storage of iron are mediated by transfer& the transferrin receptor and ferritin, respectively. In addition, two cytoplasmic RNA-binding iron-response proteins, IRP-1 and IRP-2, have been identified. The IRPs bind to specific stem-loop motifs (i.e. ironresponsive elements) present in the 5’- or 3’-untranslated region of ferritin and transferrin receptor mRNAs, respectively [70]. Iron regulates the RNAbinding function of IRP-1 and IRP-2 through fundamentally different mechanisms. For IRP-1, which is a bifunctional protein, iron inhibits RNA-binding activity by promoting the assembly of a (4Fe-4s) cluster in the binding protein, thereby converting it to cytosolic aconitase [9]. A 30 % amino acid identity over the entire length of the protein was found between pig heart mitochondrial aconitase and the human IRP-1 protein [67]. Interactions between citrate and iron are well known because citrate frequently is used as an iron chelator. It is reported now, however, that synthesis of human mitochondrial aconitase is iron-regulated and that citrate may play a role in iron metabolism [70]. In some plants, cytosolic aconitase seems to participate in the glyoxylate cycle. Aconitase has been purified from potato mitochondria [86], from pumpkin cotyledons [25], and from melon [62]. Brouquisse et al. [15] found no differences between the cytosolic and the mitochondrial aconitases of higher plants; however, in both plants and animals, the two aconitases appear to be encoded by two different genes [40]. In plants, glyoxylate and acetyl-CoA, derived from fatty acid oxidation, are vol. 37 (2) 1999

90

J. Imsande

Table I. Enzymes and other proteins reported to contain (Fe-S) clusters. EC#

Name

Reaction

la. 1.1.1.204 lb. 1.1.3.22 lc. 1.2.3.1 2. 1.1.3.17 3. 1.2.3.10 4. 1.3.1.31 5a. 1.3.5.1 5b. 1.3.99.1 6a. 1.4.1.13 6b. 1.4.7.1 7. 1.5.5.1 8. 1.5.99.7 9a. 1.6.5.3 9b. 1.6.99.3 10. 1.7.7.1 11. 1.7.7.2 12a. 1.8.1.2 12b. 1.8.7.1 12~. 1.8.99.1 13. 1.8.99.3 14. 1.10.2.2 15. 1.14.12.3 16. 1.14.12.7

Xanthine dehydrogenase Xanthine ox&se Aldehyde oxidase Choline oxidase Carbon-monoxide oxidase Enoate reductase Succinate dehydrogenase Succinate dehydrogenase Glutamate synthase Glutamate synthase Flavoprotein dehydrogenase Trimethylamine dehydrogenase NADH dehydrogenase NADH dehydrogenase Nitrite reductase Nitrate reductase Sulfite reductase Sulfite reductase Sulfite reductase Trithionate oxidoreductase Ubiquinol-cytochrome-c reductase Benzene 1,2-dioxygenase Phthalate 4,5-dioxygenase

17. 1.14.12.10 18. 1.14.99.24 19. 1.17.4.1 20. 1.18.6.1 21. 1.18.99.1 22. 2.3.1.54 23. 2.4.2.14 24. 4.2.1.2 25.4.2.1.3 26. 4.2.1.9 27. 4.2.1.12 28. 4.2.1.13 29. 4.2.1.16 30. 4.2.1.31 31. 4.2.1.33 32. 4.2.1.34 33. 4.2.1.35 34. 4.2.1.85 35. 4.2.99.18 36. 4.99.1.1 37. 5.4.3.2 38. Unassigned 39. Unassigned 40. Unassigned 41. Unassigned 42. 43. 44. 46. 47. 48. 49.

Benzoate 1,2dioxygenase Steroid 9a-monooxygenase Ribonucleotide reductase @) Nitrogenase Hydrogenase Pyruvate-formate lyase Amidophosphoribosyltransferase Fumarase (A&B of E.&i) Aconitate reductase (aconitase) Dihydroxy-acid dehydratase Phosphogluconate dehydratase Serine dehydratase Threonine dehydratase Maleate hydratase 3-Isopropylmalate dehydratase Mesaconate hydratase Citraconate hydratase Dimethylmaleate hydratase Endonuclease III Ferrochelatase Lysine 2,3-aminomutase Ferredoxin:thioredoxin reductase 4-OHbutyryl-CoA dehydratase Biotin synthase Quinolinate synthetase (subunit B) Fumarate nitrate reduction protein Superoxide response protein (soxR) Chloroplast Rieske protein (lumen side) Ferredoxin (fdx) Photosystem I (PSI) Fx Photosystem I (PSI) FA Photosystem I (PSI) F,

xanthine + NAD+ + H,O + mate + NADH xanthine + Hz0 + 0, + urate + H,O, aldehyde + H,O +O, + acid + O;choline + 0, + betaine aldehyde + HzO, CO + Hz0 + 0, + CO1 + H,O, 2-butenoate + NADH + butanoate + NAD+ succinate + ubiquinone --t fumarate + ubiquinol succinate + acceptor + fumarate + r-acceptor glutamine + 2-oxoglutarate + NADPH -+ 2 glutamate + NADP+ glutamine + 2-oxoglutarate +2 r-fdx + 2 glutamate + 2 o-fdx r-flavoprotein + ubiquinone --f o-flavoprotein + uhiquinol trimethylamine + Hz0 + o-acceptor + dimethylamine + formaldehyde + r-acceptor NADH + ubiquinone + NAD+ + ubiquinol NADH + o-cytochrome c + NAD+ + r-cytochrome c nitrite + 3-r-ferredoxin + NH, + 3 o-fdx + 2 H,O nitrate + 2 r-ferredoxin + nitrite +2 o-fdx + Hz0 sulfite + 3 NADPH + H,S + 3 NADP+ + 3 Ha0 sulfite + 3 r-ferredoxin + H,S + 3 o-fdx + 3 Ha0 sulfite + r-acceptor + H,S + o-acceptor + 3HaO 3 HSO; + r-acceptor + (OsS-S-SO,)*- + o-acceptor + 2H,O + OHQHz + 2 ferricytochrome c + Q + 2 ferrocytochrome c benzene + NADH + 0, -_;)cyclohexa-3,5-diene-1,2-diol + NAD+ phthalate + NADH + 0, -+ 4,5-diOHcyclohexa1(6X2-diene-1,2_dicarboxylate + NAD+ + H,O benzoate + NADH + 0, + catechol + COZ + NAD+ pregna-49 (1 I)-diene-3, 20-dione + AH, + 0, + epoxy-steroid +A + H,O NDP+ r-thioredoxin + dNDP + o-thioredoxin + H,O N, + nATP +6H+ + 3 r-ferredoxin + 2NHs +3 o-fdx+ nADP + nPi H, + 2 o-ferredoxin -+ 2 H+ + 2 r-ferredoxin pyruvate + CoA + acetyl-CoA + formate glutamine + PRPP + H,O + 5-phosphoribosylamine + glu + PP, fumarate + H,O --f malate citrate -_) aconitate + H,O 2,3,diOH-3-methylbutanoate + 3-methyl-2-oxobutanoate + Hz0 6-P-gluconate + 2-dehydro-3-deoxy-6-P-gluconate + H,O serine + H,O -+ pyruvate + NH, + H,O threonine + Hz0 + 2-oxobutanoate + NH, + H,O malate 3 maleate + H,O 3-isopropylmalate --f 2-isopropylmaleate + Hz0 2-methylmalate --f 2-methylfumarate + H,O 2-methylmalate --f 2-methylmaleate + H,O 2,3_dimethylmalate -+ dimethylmaleate + H,O DNA repair protophorphyrin + Fe*+ + protoheme + 2 H+ lysine -_) 3,6_diaminohexanoate r-ferredoxin + o-thioredoxin -+ o-ferredoxin + r-thioredoxin 4-hydroxybutyryl-CoA + crotonyl-CoA + H,O dethiobiotin + biotin (reaction unknown) aspartate + H,O -_) iminoaspartate + H,O, DNA binding protein (E. cd) DNA binding protein (/I?. coli)

Plant Physiol. Biochem.

Iron-sulfur clusters condensed via the glyoxylate cycle to form succinate. The five enzymes of the glyoxylate cycle (aconitase, isocitrate lyase, malate synthase, malate dehydrogenase and citrate synthase) were thought to be confined to glyoxysomes [6]; however, little or no aconitase activity is found in the glyoxysome [21]. It now appears likely that cytosolic aconitase, whose gene has been cloned from pumpkin cotyledons, participates in the glyoxylate cycle [40]. Because H,O, is a normal constituent of glyoxysomes and because aconitase is known to be inhibited by H,O, [86], it seems clear that an aconitase bearing a (4Fe-4s) cluster could not function in the glyoxysome. Oxidative stress signals (i.e. H,O, and OF) can perturb (Fe-S) clusters and thus convert cytosolic aconitase to IRP-1 191. Indeed, for humans and insects, it has been reported that mitochondrial aconitase is a prime target of oxidative damage that occurs during aging and/or hyperoxia [90]. Less is known about these processes in plants; however, a full-length cDNA clone of the unstable aconitase has been isolated from a library derived from immature Arabidopsis pods [62] and from pumpkin cotyledons [40]. When compared to seven other aconitase sequences, the Arabidopsis gene showed 86 % identity with melon, 60 % with mouse and rabbit, 58 % with human, 54 % with E. coli, and 30 % with pig and yeast [62]. E. coli contain two aconitase genes, acnA and acnB. acnA, expressed later in the growth cycle than acnB, is

specifically subject to SoxRS-mediated activation and is induced by iron and redox stress as the growth rate slows. Further, acnA can be repressed by FNR [24]. Thus, both soxRC and FNR are active genetic regulators. Succinate dehydrogenase (SDH) is an essential enzyme that links the citric acid cycle to the mitochondrial electron transport chain. The human enzyme is part of complex II that resides on the inner mitochondrial membrane. Complex II is made up of four peptides, all encoded by nuclear genes. The 27-kDa ironsulfur protein (Ip) and the 70-kDa flavoprotein associate to form SDH, which is a component of complex II. The mRNA encoding the Ip subunit in Drosophila contains an iron-response element in the S-untranslated region that, like aconitase, responds to the IRP [38]. The gene encoding the Ip protein has been cloned [2]. The Ip subunit contains a (2Fe-2S), a (4Fe4s) and a (3Fe-4s) cluster [2]. The sequence of the human Ip gene has been compared to that of Saccharomyces cerevisiae, Ustilago maydis, Haemonchus contortus, and Drosophila melanogaster [2]. The

predicted amino acid sequences of the Ip subunits are

91

highly conserved. Thus, one might speculate that the Ip subunit of plant SDH may be similar to that of other organisms. Indeed, Chauveau and Roussaux [19] reported that a complex II preparation from potato mitochondria contains three (Fe-S) clusters, one of which is a component of the SDH protein. E. coli produce 3 different fumarases (FumA, FumB, and FumC). Both FumA, an aerobic enzyme, and FumB, an anaerobic enzyme, contain (4Fe-4s) clusters and are oxygen-sensitive [24]. FumC, an oxygen-stable fumarase, closely resembles mitochondrial fumarases and is fully functional during aerobic growth. Fumarases have been divided into two classes: (a) homodimeric iron-sulfur proteins (FumA and FumB) and (b) homotetrameric proteins (FumC). Fumarase genes from potato [61] and Arabidopsis [7] have been cloned and are of the homotetrameric class. Fumarase from pea mitochondrial has been purified and partially characterized [7]. To date, there is no evidence that plant fumarase contains a (Fe-S) cluster. The first energy-coupling site of the mitochondrial respiratory chain, complex I, has NADH:ubiquinone oxidoreductase activity. This complex, recently isolated from potato tuber mitochondria, is composed of at least 32 distinct protein subunits and contains at least 1 (2Fe-2s) cluster and 3 (4Fe-4s) clusters [55]. Perturbation of a (Fe-S) cluster in aconitase, SDH, or the NADH:ubiquinone oxidoreductase complex could readily alter basic carbohydrate metabolism and many aspects of plant growth. 5.3. Coordination of sulfur and nitrogen assimilation Microbial sulfite reductases (SiRs) and related nitrite reductases (NiRs) couple a metalloporphyrin to a (4Fe-4s) cluster and carry out the six electron reduction of sulfite to sulfide or nitrite to ammonia, respectively [22, 231. These sulfite reductases (EC 1.8.1.2) contain 8 a-subunits (66 kDa each) and 4 P-subunits (64 kDa each) and rely on NADPH for reducing power. Each P-subunit contains a (4Fe-4s) cluster and a siroheme. Green plants carry out similar reactions; however, they are ferredoxin-dependent (EC 1.8.7.1) rather than NADPH-dependent. Sulfite reductase has been purified from turnip and several other sources [81]. The turnip enzyme contains a siroheme cofactor but the presence or absence of an iron-sulfur cluster was not addressed. Subunits of the enzyme have a molecular mass of approximately 64.5 kDa. Using the PCR technique, a putative P-subunit has been cloned from Arabidopsis [ 161. The transcript would produce a protein of 642 amino acids, of which 66 amino acids vol. 37 (2) 1999

92

J. Imsande

would be a transit peptide. Thus, the mature protein would be composed of 576 amino acids with a molecular mass of 65.28 kDa. Sequence analysis revealed conservation of the cysteinyl groups required for the (4Fe4s) cluster [ 161. The effect of ROS on the reduction of sulfite or nitrite in higher plants has not been established. It is noteworthy, however, that yeast mutants lacking cytosolic superoxide dismutase (SODl) and E. coli SOD- mutants are auxotrophic for organic sulfur [IO, 821. Enhanced NADPH production, however, is reported to alleviate the reduced sulfur requirement in yeast [75]. Therefore, it was speculated that the SODmutation allowed ROS to accumulate, which in turn, inactivated the respective sulfite reductases. Genes encoding nitrite reductase have been cloned from several higher plants, including spinach, maize, bean and tobacco, and from microbes. The derived amino acid sequences of the higher plant nitrite reductases show an 80 % identity and reveal a molecular mass of approximately 63 kDa [3,68]. The 4 cysteines involved in binding the (4Fe-4s) cluster are conserved in all higher-plant nitrite reductases examined. In addition, all enzymes contain a siroheme cofactor. Genetic regulation of nitrite reductase in E.coli has been examined in great detail and several proteins, including FNR, help coordinate gene expression [89]. Thus, in E. coli, both the synthesis and the activity of nitrite reductase rely on iron-sulfur clusters. The glutamate synthases (GltS) of several bacteria are dimers composed of an a-subunit (162 kDa) and a P-subunit (53.2 kDa) [85]. The P-subunit of the bacterial enzyme seemingly contains the NADPH binding site and hence, is not required in photosynthethic tissue. The mature ferredoxin-dependent glutamate synthase present in green tissue of higher plants is a monomer of 165 kDa [80]. The NAD(P)H-dependent GltS, found primarily in non-photosynthetic shoots, roots and root nodules, is a monomer of approximately 120 kDa. Both bacterial subunits and the Fd-GltS of green plants are reported to contain a (3Fe-4S)+%”cluster [50]. Little is known about the effect of ROS on the lability of these iron-sulfur clusters. It is possible, however, that the oxidation state of plant cells could have a profound effect on these iron-sulfur clusters and consequently upon the assimilation of nitrogen and sulfur.

the amide group of glutamine with an activated phosphoribosyl group to yield phosphotibosylamine. The enzyme from Bacillus subtilus is well studied and is said to be a model for all higher eukaryotes [201. This enzyme contains a (4Fe-4s) cluster and, although the cluster does not have a role in catalysis, the intact cluster is necessary for optimal propeptide processing and the stability of the mature protein. Perturbation of the cluster by molecular oxygen results in-a loss of native structure, leading to proteolysis and enzyme turnover [36]. Xanthine dehydrogenase has several roles in purine catabolism. A homodimer that contains two (2Fe-2s) clusters, xanthine dehydrogenase can exist in two states, a reduced form containing 2 RSH groups and an oxidized form containing a RS-SR group [66]. The reduced form, which contains two specific sulfhydryl groups, catalyzes the following reaction: xanthine + NAD+ + H,O + urate + NADH + H+. In the presence of oxidized glutathione and a transhydrogenase, the two sulfhydryls of xanthine dehydrogenase are oxidized and glutathione is reduced. Xanthine oxidase, the oxidized form of xanthine dehydrogenase, catalyzes the following reaction: xanthine + H,O + 0, + urate + H,O, (table I>. Upon storage at -20 “C, xanthine dehydrogenase may be spontaneously converted to xanthine oxidase. Xanthine dehydrogenase-oxidase participates in purine catabolism and in ureide formation, which is an essential step in N, fixation by the ureide transporters [ 11, 841. In vitro, nitrogenase must be handled anaerobically because oxygen inactivates its complex (Fe-S) clusters. The primary function of leghemoglobin in the legume root nodule is to provide sufficient oxygen for microbial respiration without the oxygen destroying the activity and/or inducibility of nitrogenase. However, if oxygen, or the redox state of the nodule, were to significantly inhibit xanthine dehydrogenase-xanthine oxidase activities, purines would accumulate in the nodules and might repress symbiotic nitrogen fixation.

5.4. Puke metabolism, senescence and indole acetic acid biosynthesis

Senescence, a genetically regulated oxidative process, involves the degradation of cellular structures and enzymes and the remobilization of these metabolites to other plant parts. Reactive oxygen species (ROS) are the primary mediators in this process and chloroplasts are among the earliest sites of catabolism [26]. Increases in the activities of peroxisomal and glyoxysomal xanthine oxidase and urate oxidase during this process suggest a role for these organelles and enzymes in the senescence cycle.

The first step in purine biosynthesis, catalyzed by phosphoribosylpyrophosphate amidotransferase, joins

Two genes encoding aldehyde oxidases in maize have been cloned and sequenced [73]. Both sequences

Plant Physiol. B&hem.

Iron-sulfur clusters show a significant similarity to animal xanthine dehydrogenase. Interestingly, indole-3-acetaldehyde is a good substrate for these aldehyde oxidases. Subsequently, genes encoding aldehyde oxidases in Arubido@ were cloned and found to contain consensus sequences for two (Fe-S) clusters [74]. Indole-3-acetaldehyde is also a substrate for these enzymes. Thus, enzymes containing (Fe-S) clusters are predicted to play key roles in hormone biosynthesis and plant development [73, 741. 5.5. Chloroplast assembly Chloroplast assembly requires proteins synthesized from two different genomes. These proteins, bearing signal-type peptides, are targeted into and across the thylakoid membrane. Transport of precursor proteins across the chloroplastic envelope membrane requires the interaction of protein translocons localized in both the outer and inner envelope membranes [ 181. Components of the outer envelope protein import apparatus are designated ‘Tot’ for translocon at the outer envelope membrane of chloroplasts. Correspondingly, components of the translocon of the inner envelope membrane of chloroplasts are named ‘Tic’ [72]. Tic55 which has a molecular mass of 52 kDa, contains a predicted Rieske-type iron-sulfur cluster and a mononuclear iron-binding site [18]. Because of its (2Fe-2s) cluster, it was predicted that Tic55 might serve as a regulating subunit of the inner envelope translocon [ 181. The lethal leaf spot 1 (Usl) locus of maize encodes a protein that serves as a suppressor of cell death [37]. This protein also contains a predicted Rieske-type iron-sulfur cluster and a mononuclear iron-binding site and exhibits a 30 % identity to Tic55. Because of its (2Fe-2s) cluster and sequence similar to a dioxygenase, it was speculated that Llsl might use dioxygenase activity to suppress cell death [37]. Spinach envelope membranes are reported to contain: (a) semiquinone and flavosemiquinone radicals, (b) a series of iron-containing electron-transfer centers, and (c) flavins (mostly FAD) loosely associated with the proteins [47]. Three iron-sulfur clusters were associated with envelope membranes, a (4Fe-4s) cluster, a (2Fe-2s) cluster, and an unusual (Fe-S) cluster termed ‘X’. It was hypothesized that the (4Fe-4s) and (2Fe-2s) clusters might be electron carriers associated with desaturase activities in the membranes or with mechanisms involved in the export of protons to the cytosol. The (4Fe-4s) and (2Fe-2s) clusters associated with chloroplast membranes were shown to be bio-

93

chemically active: however, their precise mechanisms of action remain to be identified. 5.6. Coordination of photosynthetic and plant metabolic activities The two large proteins of PSI, PSI-A and PSI-B (83.2 and 82.4 kDa, respectively), form a heterodimer known as P700 chlorophyll a-protein 1 (CPl). This complex binds most of the chlorophyll a and all of the p-carotene (approximately 90 and 15 molecules, respectively) associated with each PSI unit. In addition, the CPl reaction center complex contains electron acceptors A, and A, and the iron sulfur cluster Fx [71]. Subunit PSI-C, located on the stromal side of the PSI complex, contains two (4Fe-4s) clusters, FA and F,. The Fx cluster associated with the CPI complex, which is a component of the stroma lamella, passes electrons to the FA cluster associated with PSI-C [27]. In turn, the FA cluster passes electrons to the F, cluster. Chloroplasts are known to undergo photoinactivation. Initially, nonphotochemical thermal dissipation of excess energy and dynamic regulation of antenna size protect photosynthetic machinery. When these protective capacities are inadequate, destructive photoinhibition takes place, lowering net photosynthetic efficiency [63]. A combination of high light with other stress factors such as chilling or drought greatly increases photoinhibition. Two major types of photoinhibition have been described: photoinhibition of PSII, which requires only light [51] and that of PSI, which requires both light and 0, [69]. Originally, it was thought that PSI1 was the most sensitive site to photoinactivation; however, Inoue et al. [45] concluded that the main cause of photoinactivation in isolated spinach chloroplasts was inactivation of the iron sulfur clusters of PSI. Subsequent work with cucumber [77], spinach [76], and barley [83] showed that, with chilled plants exposed to weak illumination, hydroxyl radicals (ROS) produced at the reduced ironsulfur clusters in PSI trigger a conformational change in the CPl complex, allowing a set-me-type protease to hydrolyze PSI-A and possibly PSI-B. Chilling temperatures accompanied by weak illumination increase super-oxide formation in PSI of spinach [45], cucumber [77], and barley [83]. In each case, the increased concentrations of the superoxides cause progressive damage to PSI clusters FA, F,, and Fx. Damage to the iron sulfur clusters ultimately leads to inactivation of PSI and subsequent protein degradation [76, 831. Indeed, many forms of oxidative stress vol. 37 (2) 1999

94

J. Imsande

increase the formation of ROS and inflict plant damage [4]. It is clear from data presented above and below that the expression of many stress responses requires either a functional iron-sulfur cluster or the perturbation of iron-sulfur cluster. Glycine betaine is a prominent osmoprotectant among higher plants. Choline monooxygenase, which converts choline to betaine aldehyde, catalyzes the first step in the synthesis of glycine betaine. Choline monooxygenase, a stress-inducible enzyme, contains a Rieske-type (2Fe-2s) cluster that relies upon ferredoxin for its reducing power [65]. Thus, at least one response to salinity-stress relies upon the functional (Fe-S) clusters of ferredoxin and the Rieske-type (FeS) clusters of choline monooxygenase. The activities of five enzymes of the Calvin cycle are modulated by light intensity [17, 881. Some of the electrons generated by PSII and PSI are directed through the three (4Fe-4s) clusters of PSI and passed to ferredoxin. Subsequently, some of these electrons are transferred to thioredoxin via ferredoxin/thioredoxin reductase (FTR) [46]. The reducing power contained in one of the various forms of reduced thioredoxin can activate glyceraldehyde-3-phosphate dehydrogenase, phosphoribulokinase, fructose 1-6-bisphosphatase, and sedoheptulose 1-7-bisphosphatase by reduction of intramolecular sulfhydryl groups. Glucose 6-phosphate dehydrogenase (G-6-PDH), on the other hand, is inactivated by the FIR system. Thus, the light driven FTR system and the redox state of chloroplasts contribute strongly to the coordination and regulation of photosynthesis, plant metabolism and plant growth. Interestingly, yeast cells deficient in G-6-PDH are auxotrophic for reduced-sulfur or methionine [82]. The requirement for reduced-sulfur may result from inadequate NADPH, which is necessary for sulfite reduction in yeast and is normally provided by G-6PDH and phosphogluconate dehydrogenase of the pentose phosphate pathway [75]. The G-6-PDH deficiency is exacerbated by a SOD deficiency. In higher plants, the oxidative segment of the pentose phosphate pathway drives the Calvin cycle in the absence of photosynthesis.

6. CONCLUSION Most stressed ‘yellow’ plants are chlorophyll deficient. The normal green hue of the yellow plant ‘is sometimes regained following the addition of N, or sulfur, or iron [44]. Is there a common mechanism by Plant Physiol. Biochem.

which a deficiency of N, or S, or Fe might cause a normally green plant to develop a chlorophyll deficiency and the resulting yellow hue? For most higher plants grown on cultivated soils, nitrate is the most common source of N. Because chlorophyll contains N [87], one might expect that a N-deficiency would cause a plant to become chlorophyll deficient and yellow. Another reason, however, might be that nitrate reductase, nitrite reductase, and glutamate ‘synthase, three essential enzymes in the normal assimilation of N, all contain iron-sulfur clusters. Nitrogenase also contains (Fe-S) clusters. If N stress were to generate an increased concentration of ROS, and seemingly most if not all stresses do [4], then the increased ROS might inactivate one or more essential (Fe-S) cluster. Hence, there could be two reasons for the yellow chlorophyll-deficiency phenotype: (a) inadequate nitrogen in the rooting medium; or (b) an apparent deficiency of N, or other essential element, caused by defective (Fe-S) clusters. Thus, even if a small amount of N were made available, possibly by protein turnover, its assimilation would be impaired. Likewise, stress resulting from a sulfur deficiency would produce ROS which could destroy one or more essential (Fe-S) cluster, perhaps that of sulfite reductase. Consequently, both the reduction and assimilation of nitrate, nitrite, and sulfite could be impaired and a chlorophyll deficiency would occur. Similarly, an iron deficiency could impede N, S, and Fe assimilation in at least two ways: (a) iron deficiency-stress could produce ROS which could inactivate (Fe-S) clusters,, and (b) iron deficiency itself could hinder the formation of functional (Fe-S) clusters. Thus, there are known mechanisms by which N, S, or Fe deficiencies could produce the same yellow phenotype in a normal green plant. Functioning (Fe-S) clusters are central to plant metabolism and photosynthesis and deserve the careful attention of plant biochemists and physiologists. Acknowledgements. Journal paper No. J-17940 of the Iowa Agric. and Home Econ. Exp. Stn., Ames, Project No. 3412, and supported by Hatch Act and State of Iowa funds. The author apologizes to the many authors whose work could not be included because of the restriction on citations.

REFERENCES [l] Asada K., Production and action of active oxygen species in photosynthetic tissues, in: Foyer C.H., Mullineaux P.M. (Eds.), Causes of Photooxidative Stress and Amelioration of Defense Systems in Plants, CRC Press, Boca Raton, FL, 1994, pp. 77-104.

Iron-sulfur clusters

[2] Au H.C., Ream-Robinson D., Bellew L.A., Broomfield P.L.E., Saghbini M., Scheffler I.E., Structural organization of the gene encoding the human iron-sulfur subunit of succinate dehydrogenase, Gene 159 (1995) 249253. [3] Back E., Burkhart W., Moyer M., Privalle L., Rothstein S., Isolation of cDNA clones coding for spinach nitrite reductase: Complete sequence and nitrate induction, Mol. Gen. Genet. 212 (1988) 2&26. [4] Bartosz G., Oxidative stress in plants, Acta Physiol. Plant. 19 (1997) 47-64. [5] Beauchamp C.O., Fridovich I., A mechanism for the production of ethylene from methional: The generation of hydroxyl radical by xanthine oxidase, J. Biol. Chem. 245 (1970) 4641-4646. [6] Beevers H., Organelles from castor bean seedlings: Biochemical roles in gluconeogenesis and phospholipid synthesis, in: Gall&d T., Mercer E.I. (Ms.), Recent Advances in the Chemistry and Biochemistry of Plant Lipids, Academic Press, New York, 1975, pp. 287-299. [7] Behal R.H., Oliver D.J., Biochemical and molecular characterization of fumarase from plants: purification and characterization of the enzyme, cloning, sequencing, and expression of the gene, Arch. Biochem. Biophys. 348 (1997) 65-74. [8] Beinert H., Holm R.H., Munck E., Iron-sulfur clusters: Nature’s modular, multipurpose structures, Science 277 (1997) 653-659. [9] Beinert H., Kennedy M.C., Stout C.D., Aconitase as iron-sulfur protein, enzyme, and iron-regulatory protein, Chem. Rev. 96 (1996) 2335-2373. [lo] Benov L., Fridovich I., Superoxide imposes leakage of sulfite from Escherichia coli, Arch. Biochem. Biophys. 347 (1997) 271-274. [ll] Boland M.J., Blevins D.G., Randall D.P., Soybean nodule xantbine dehydrogenase: a kinetic study, Arch. Biochem. Biophys. 222 (1983) 435-441. [12] Bowler C., Van Camp W., Van Montagu M., Inze D., Superoxide dismutase in plants, CRC Crit. Rev. Plant Sci. 13 (1994) 199-218. [ 131 Brintzinger H., Palmer G., Sands R.H., On the ligand field of iron in ferredoxin from spinach chloroplasts and related nonheme iron enzymes, Proc. Nat1 Acad. Sci. USA 55 (1966) 397-404. [14] Brouquisse R., Gaillard J., Deuce R., Electron paramagnetic resonance characterization of membrane bound iron-sulfur clusters and aconitase in plant mitochondria, Plant Physiol. 81 (1986) 247-252. [15] Brouquisse R., Nishimura M., Gaillard J., Deuce R., Characterization of a cytosolic aconitase in higher plant cells, Plant Physiol. 84 (1987) 1402-1407. [16] Bruhl A., Haverkamp T., Gisselmann G., Schwenn J.D., A cDNA clone from Arabidopsis thaliana encoding plastidic ferredoxin:sulfite reductase, Biochim. Biophys. Acta 1295 (1996) 119-124. [17] Buchanan B.B., Regulation of CO, assimilation in oxygenic photosynthesis: the ferredoxinlthioredoxin system, Arch. Biochem. Biophys. 288 (1991) l-9.

95

[ 181 Caliebe A., Grimm R., Kaiser G., Lubeck J., Sol1 J., Heins L., The chloroplastic protein import machinery contains a Rieske-type iron-sulfur cluster and a mononuclear iron-binding protein, EMBO J. 16 (1997) 7342-7350. [19] Chauveau M., Roussaux J., EPR studies on 6-benzoylaminopurine and thenoyltrifluoroacetone inhibition sites of succinate dehydrogenase in plant mitochondria, Plant Cell Physiol. 37 (1996) 914-921. [20] Chen S., Zheng L., Dean D.R., Zalkin H., Role of nifS in maturation of glutamine phosphoribosylpyrophosphate amidotransferase, J. Bact. 179 (1997) 75877590. [21] Courtois-Vemiquet F., Deuce R., Lack of aconitase in glyoxysomes and peroxisomes, Biochem. J. 294 (1993) 103-107. [22] Crane B.R., Siegel L.M., Getzoff E.D., Structures of the siroheme- and Fe4-SCcontaining active center of sulfite reductase in different states of oxidation: Heme activation via reduction-gated exogenous ligand exchange, Biochemistry 36 (1997) 12101-12119. [23 ] Crane B.R., Siegel L.M., Getzoff E.D., Probing the catalytic mechanism of sulfite reductase by X-ray crystallography: Structures of the Escherichia coli hemoprotein in complex with substrates, inhibitors, intermediates, and products, Biochemistry 36 (1997) 12120-12137. [24] Cunningham L., Gruer M.J., Guest J.R., Transcriptional regulation of the aconitase genes (ucnA and ucnB) of Escherichiu coli, Microbiology 143 (1997) 3795-3805. [25] De Bellis L., Tsugeki R., Alpi A., Nishimura M., Purification and characterization of aconitase from etiolated pumpkin cotyledons, Physiol. Plant. 88 (1993) 485492. [26] de1 Rio L.A., Pastori G.M., Palma J.M., Sandalio L.M., Selvilla F., Corpas F.J., Jiminez A., Lopez-Huertas E., Hemandez J.A., The activated oxygen role of peroxisomes in senescence, Plant Physiol. 116 (1998) 11951200. [27] Diaz-Quintana A., Leibl W., Bottin H., Setif P., Electron transfer in photosystem I reaction centers follows a linear pathway in which iron-sulfur cluster Fa is the immediate donor to soluble ferredoxin, Biochemistry 37 (1998) 3429-3439. [28] Ding H., Demple B., Glutathione-mediated destabilization in vitro of (2Fe-2s) centers in the SoxR regulatory protein, Proc. Nat1 Acad. Sci. USA 93 (1996) 94499453. [29] Flint D.H., Escherichiu coli contains a protein that is homologous in function and N-terminal sequence to the protein encoded by the niJs gene of Azotobucter vineZundii and that can participate in the synthesis of the Fe-S cluster of dihydroxy-acid dehydratase, J. Biol. Chem. 271 (1996) 16068-16074. [30] Flint D.H., Allen R.M., Iron-sulfur proteins with nonredox functions, Chem. Rev. 96 (1996) 23 15-2334. [31] Foyer C.H., Lopez-Delgado H., Dat J.F., Scott I.M., Hydrogen peroxide- and glutathione-associated mechavol. 37 (2) 1999

96

[32]

[33]

[34]

[35]

[36]

[37]

[38]

[39]

[40]

[41]

[42]

[43] [44]

[45]

[46]

[47]

J. Imsande

nisms of acclimatory stress tolerance and signaling, Physiol. Plant. 100 (1997) 241-2.54. Furbank R.T., Badger M.R., Oxygen exchange associated with electron transport and photophosphorylation in spinach chloroplasts, B&him. Biophys. Acta 723 (1983) 400-409. Gaudu P., Moon N., Weiss B., Regulation of the soxRS oxidative stress regulon, J. Biol. Chem. 272 (1997) 5082-5086. Gibson J.F., Hall D.O., Thomley J.H.M., Whatley F.R., The iron complex in spinach ferredoxin, Proc. Nat1 Acad. Sci. USA 56 (1966) 987-990. Grabowski R., Hofmeister A.E.M., Buckel W., Bacterial L-serine dehydratases: a new family of enzymes containing iron-sulfur clusters, Trends Biochem. Sci. 18 (1993) 297-300. Grandoni J.A., Switzer R.L., Makaroff C.A., Zalkin H., Evidence that the iron-sulfur cluster of Bacillus subtiZus glutamine phosphoribosylpyrophosphate amidotransferase determines stability to enzyme degradation in vivo, J. Biol. Chem. 264 (1989) 6058-6064. Gray J., Close P.S., Briggs S.P., Johal G.S., A novel suppressor of cell death in plants encoded by the Llsl gene of maize, Cell 89 (1997) 25-31. Gray N.K., Pantopoulos K., Dandekar T., Ackrell B.A.C., Hentze M.W., Translational regulation of mammalian and Drosophila citric acid cycle enzymes via iron-responsive elements, Proc. Nat1 Acad. Sci. USA 93 (1996) 4925-4930. Green J., Bennett B., Jordan P., Ralph E.T., Thompson A.J., Guest J.R., Reconstitution of the (4Fe-4s) cluster in FNR and demonstration of the aerobic-anaerobic transcription switch in vitro, B&hem. J. 316 (1996) 887-892. Hayashi M., de-Bellis L., Alpi A., Nishimura M., Cytosolic aconitase participates in the glyoxylate cycle in etiolated pumpkin cotyledons, Plant Cell. Physiol. 36 (1995) 669-680. Hentze M.W., Kuhn L.C., Molecular control of vertebrate iron metabolism: mRNA-based regulatory circuits operated by iron, nitric oxide, and oxidative stress, Proc. Nat1 Acad. Sci. USA 93 (1996) 8175-8182. Holm R.H., Kennepohl P., Solomon E.I., Structural and functional aspects of metal sites in biology, Chem. Rev. 96 (1996) 2239-23 14. Howard J.B., Rees D.C., Structural basis of biological nitrogen fixation, Chem. Rev. 96 (1996) 2965-2982. Imsande J., Iron, sulfur, and chlorophyll deficiencies: A need for an integrative approach to plant physiology, Physiol. Plant. 103 (1998) 139-144. Inoue K., Sakurai H., Hiyama T., Photoinactivation site of photosystem I in isolated chloroplasts, Plant Cell Physiol. 27 (1986) 961-968. Jacquot J.-P, Lancelin J.-M., Meyer Y., Thioredoxins: structure and function in plant cells, New Phytol. 136 (1997) 543-570. Jager-Voterro P., Domie A.-E., Jordanov J., Deuce R., Joyard J., Redox chains in chloroplast envelope membranes: Spectroscopic evidence for the presence of

Plant Physiol. Biochem.

electron carriers, including iron-sulfur centers, Proc. Nat1 Acad. Sci. USA 94 (1997) 1597-1602. [48] Jordan P.A., Thomson A.J., Ralph ET., Guest J.R., FNR is a direct oxygen sensor having a biphasic response curve, FEBS Lett. 416 (1997) 349-352. [49] Kennedy M.C., Emptage M.H., Dreyer J.-L., Beinert H., The role of iron in the activation-inactivation of aconitase, J. Biol. Chem. 258 (1983) 11098-l 1105. [50] Knaff D.B., Hirasawa M., Ameyibors E., Fu W., Johnson M.K., Spectroscopic evidence for a (3Fe-4s) cluster in spinach glutamate synthase, J. Biol. Chem. 266 (1991) 15080-15084. [51] Kok B., Gassner E.B., Rurainski H.J., Photoinhibition of chloroplast reactions, Photochem. Photobiol. 4 (1965) 215-227. [52] Kulzer R., Pils T., Kappl R., Huttermann J., Knappe J., Reconstruction and characterization of the polynuclear iron-sulfur cluster in pyruvate formate-lyase-activating enzyme, J. Biol. Chem. 273 (1998) 48974903. [53] Kuo C.F., Mashino T., Fridovich I., a,b-Dihydroxy isovalerate dehydrogenase: A superoxide-sensitive enzyme, J. Biol. Chem. 262 (1987) 4724-4727. [54] Lieder K.W., Booker S., Ruzicka F.J., Beinert H., Reed G.H., Frey PA., S-adenosylmethionine-dependent reduction of lysine 2,3-aminomutase and observation of the catalytically functional iron-sulfur centers by electron paramagnetic resonance, Biochemistry 37 (1998) 2578-2585. [55] Lin T.I., Sled V.D., Ohnishi T., Brennicke A., Grohmann L., Analysis of the iron-sulfur clusters within the complex I (NADH:ubiquinone oxidoreductase) isolated from potato tuber mitochondria, Eur. J. Biochem. 230 (1995) 1032-1036. [56] Liochev S.I., The role of iron-sulfur clusters in in vivo hydroxyl radical production, Free Rad. Res. 25 (1996) 369-384. [57] Liochev S.I., Fridovich I., The role of O;- in the production of HO’: In vitro and in vivo, Free Rad. Biol. Med. 16 (1994) 29-33. [58] McCord J.M., Day E.D., Superoxide-dependent production of hydroxyl radical catalyzed by iron-EDTA complex, FEBS L&t. 68 (1978) 139-142. [59] McCord J.M., Fridovich I., Superoxide dismutase: An enzymic function for erythrocuprein (hemocuprein), J. Biol. Chem. 244 (1969) 6049-6055. 1601 McCord J.M., Keele B.B., Fridovich I., An enzyme based theory of obligate anaerobiosis: the physiological function of superoxide dismutase, Proc. Natl Acad. Sci. USA 68 (1971) 1024-1027. [61] Nast G., Muller-Rober B., Molecular characterization of potato fumarate hydratase and functional expression in Escherichia coli, Plant Physiol. 112 (1996) 12191227. [62] Peyret P., Perez P., Alric M., Structure, genomic organization, and expression of the Arabidopsis thaliana aconitase gene, J. Biol. Chem. 270 (1995) 8131-8137. [63] Powles S.B., Photoinhibition of photosynthesis induced by visible light, Annu. Rev. Plant Physiol. 35 (1984) 15-44.

Iron-sulfur clusters

[64] Proudhon D., Wei J., Briat J.-F., Theil E.C., Ferritin gene organization: differences between plants and animals suggest possible kingdom-specific selective constraints, J. Mol. Evol. 42 (1996) 325-336. [65] Rathinasabapathi B., Burnet M., Russell B.L., Gage D.A., Liao P.-C., Nye G.J., Scott I!, Golbeck J.H., Hanson A.D., Choline monooxygenase, an unusual iron-sulfur enzyme catalyzing the first step of glycine betaine synthesis in plants: prosthetic group characterization and cDNA cloning, Proc. Nat1 Acad. Sci. USA 94 (1997) 3454-3458. [66] Romao M.J., Huber R., Structure and function of the xanthine-oxidase family of molybdenum enzymes, Structure Bonding 90 (1998) 69-95. [67] Rouault T.A., Stout C.D., Kaptain S., Harford J.B., Klausner R.D., Structural relationship between an ironregulated RNA-binding protein (IRE-BP) and aconitase: Functional implications, Cell 64 (1991) 881-883. [68] Sander L., Jensen P.E., Back L.F., Stummann B.M., Henningsen K.W., Structure and expression of a nitrite reductase gene from bean (Phaseolus vulgaris) and promoter analysis in transgenic tobacco, Plant Mol. Biol. 27 (1995) 165-177. [69] Satoh K., Mechanism of photoinhibition in photosynthetic systems II. The occurrence of two different types of photoinhibition, Plant Cell. Physiol. 11 (1970) 2938. [70] Schalinske K.L., Chen O.S., Eisenstein R.S., Iron differentially stimulates translation of mitochondrial aconitase and ferritin mRNAs in mammalian cells, J. Biol. Chem. 273 (1998) 3740-3746. [71] Scheller H.V., Naver H., Moller B.L., Molecular aspects of photosystem I, Physiol. Plant. 100 (1997) 842-85 1. [72] Schnell D.J., Blobel G., Keegstra K., Kessler F., Ko K., Sol1 J., A nomenclature for the protein-import components of the chloroplast envelope, Trends Cell Biol. 7 (1997) 303-304. [73] Sekimoto H., Seo M., Dohmae N., Takio K., Kamiya Y., Koshiba T., Cloning and molecular characterization of plant aldehyde oxidase, J. Biol. Chem. 272 (1997) 15280-15285. [74] Sekimoto H., Seo M., Kawakami N., Komano Y., Desloire S., Liotenberg S., Marionpoll A., Caboche M., Kamiya Y., Koshiba T., Molecular-cloning and characterisation of aldehyde oxidase in Arabidopsis thaliana, Plant Cell. Physiol. 39 (1998) 433-442. [75] Slekar K.H., Kosman D.J., Culotta V.C., Yeast copper/ zinc superoxide dismutase and the pentose phosphate pathway play overlapping roles in oxidative stress protection, J. Biol. Chem. 271 (1996) 28831-28836. ]76] Sonoike K., Kamo M., Hihara Y., Hiyama T., Enami I., The mechanism of the degradation of psaB gene product, one of the photosynthetic reaction center subunits of photosystem I, upon photoinhibition, Photosyn. Res. 53 (1997) 55-63. ]77] Sonoike K., Terashima I., Iwaki M., Itoh S., Destruction of photosystem I iron-sulfur centers in leaves of

97

Cucumis sativus L. by weak illumination at chilling temperatures, FEBS Lett. 362 (1995) 235-238. [78] Spiro S., Guest J.R., FNR and its role in oxygen-regulated gene expression in Escherichia coli, FEMS Microbial. Rev. 75 (1990) 399-428. [79] Staples C.R., Ameyibor E., Fu W., Gardet-Salvi L., Stritt-Etter A.-L., Schurmann P., Knaff D.B., Johnson M.K., The function and properties of the iron-sulfur center in spinach ferredoxin:thioredoxin reductase: A new biological role for iron-sulfur clusters, Biochemistry 35 (1996) 11425-l 1434. [80] Suzuki A., Rothstein S., Structure and regulation of ferredoxin-dependent glutamate synthase from Arabidopsis thaliana, Eur. J. Biochem. 243 (1997) 708-7 18. [81] Takahashi S., Yip W.-C., Tamura G., Purification and characterization of ferredoxin-sulfite reductase from turnip (Brassica rapa) leaves and comparison of properties with ferredoxin-sulfite reductase from turnip roots, Biosci. Biotech. Biochem. 61 (1997) 1486-1490. [82] Thomas D., Cherest H., Surdin-Kerjan Y., Identification of the structural gene for glucose-6-phosphate dehydrogenase in yeast. Inactivation leads to a nutritional requirement for organic sulfur, EMBO J. 10 (1991) 547-553. [83] Tjus S.E., Moller B.L., Scheller H.V., Photosystem I is an early target of photoinhibition in barley illuminated at chilling temperatures, Plant Physiol. 116 (1998) 755-764. 1841 Triplett E.W., Blevins D.G., Randall D.D., Purification and properties of soybean nodule xanthine dehydrogenase, Arch. Biochem. Biophys. 219 (1982) 39-46. [85] Vanoni M.A., Fischer F., Ravasio S., Verzotti E., Edmondson D.E., Hagen W.R., Zanetti G., Curti B., The recombinant a-subunit of glutamate synthase: spectroscopic and catalytic properties, Biochemistry 37 (1998) 1828-1838. [86] Verniquet F., Gaillard J., Neuburger M., Deuce R., Rapid inactivation of plant aconitase by hydrogen peroxide, Biochem. J. 276 (1991) 643-648. [87] Von Wettstein D., Gough S., Kannangara C.G., Chlorophyll biosynthesis, Plant Cell 7 (1995) 1039-1057. [88] Wedel N., Sol1 J., Paap B.K., CP12 provides a new mode of light regulation of Calvin cycle activity in higher plants, Proc. Nat1 Acad. Sci. USA 94 (1997) 10479-10484. [89] Wu H.-C., Tyson K.L., Cole J.A., Busby S.J.W., Regulation of transcription initiation at the Escherichia coli nir operon promoter: a new mechanism to account for co-dependence on two transcription factors, Mol. Microbial. 27 (1998) 493-505. ]90 Yan L.-J., Levine R., Sohal R.S., Oxidative damage during aging targets mitochondrial aconitase, Proc. Nat1 Acad. Sci. USA 94 (1997) I 1168-l 1172. ]91 Zheng L., White R.H., Cash V.L., Jack R.F., Dean D.R., Cysteine desulfurase activity indicates a role for NIFS in metallocluster biosynthesis, Proc. Nat1 Acad. Sci. USA 90 (1993) 2754-2758. vol. 37 (2) 1999