Advances in BiologyVolume 2014 (2014), Article ID 808569, 21 pageshttp://dx.doi.org/10.1155/2014/808569

Review Article

The Biosynthesis of the Molybdenum Cofactor in Escherichia coli and Its Connection to FeS Cluster Assembly and the Thiolation of tRNA

Silke Leimkühler

Department of Molecular Enzymology, Institute of Biochemistry and Biology, University of Potsdam, Karl-Liebknecht-Straße 24-25, 14476 Potsdam, Germany

Received 16 January 2014; Accepted 28 March 2014; Published 29 April 2014

Academic Editor: Paul Rösch

Copyright © 2014 Silke Leimkühler. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Abstract.

The thiolation of biomolecules is a complex process that involves the activation of sulfur. The L-cysteine desulfurase IscS is the main sulfur mobilizing protein in Escherichia coli that provides the sulfur from L-cysteine to several important biomolecules in the cell such as iron sulfur (FeS) clusters, molybdopterin (MPT), thiamine, and thionucleosides of tRNA. Various proteins mediate the transfer of sulfur from IscS to various biomolecules using different interaction partners. A direct connection between the sulfur-containing molecules FeS clusters, thiolated tRNA, and the molybdenum cofactor (Moco) has been identified. The first step of Moco biosynthesis involves the conversion of 5′GTP to cyclic pyranopterin monophosphate (cPMP), a reaction catalyzed by a FeS cluster containing protein. Formed cPMP is further converted to MPT by insertion of two sulfur atoms. The sulfur for this reaction is provided by the L-cysteine desulfurase IscS in addition to the involvement of the TusA protein. TusA is also involved in the sulfur transfer for the thiolation of tRNA. This review will describe the biosynthesis of Moco in E. coli in detail and dissects the sulfur transfer pathways for Moco and tRNA and their connection to FeS cluster biosynthesis.

1. An Introduction to the Importance of Molybdenum and Molybdoenzymes in Bacteria

Molybdenum is the only second row transition metal essential for biological systems, which is biologically available as molybdate ion [1]. Molybdenum has a chemical versatility that is useful to biological systems: it is redox-active under physiological conditions (ranging between the oxidation states VI and IV); since the V oxidation state is also accessible, the metal can act as transducer between obligatory two-electron and one-electron oxidation-reduction systems and it can exist over a wide range of redox potentials [2, 3]. The metal forms the active site of molybdoenzymes, which execute key transformations in the metabolism of nitrogen, sulfur, and carbon compounds [3]. The catalyzed reactions are in most cases oxo-transfer reactions; for example, the hydroxylation of carbon centers and the physiological role are fundamental since the reactions include the catalysis of key steps in carbon, nitrogen, and sulfur metabolism.

There are two distinct types of molybdoenzymes: molybdenum nitrogenase has a unique molybdenum-iron-sulfur cluster, the [Fe₄S₃]-(bridging-S)₃-[MoFe₃S₃] center called FeMoco [4]. Nitrogenase catalyzes the reduction of atmospheric dinitrogen to ammonia. All other molybdoenzymes contain the molybdenum cofactor (Moco). In Moco the molybdenum atom is coordinated to a dithiolene group on the 6-alkyl side chain of a pterin called molybdopterin (MPT) [3, 5]. The chemical nature of Moco has been determined by Rajagopalan and coworkers in 1982 [5]. They postulated a structure of the cofactor consisting of a pterin derivative, with the pterin ring substituted at position 6 with a phosphorylated dihydroxybutyl side chain containing a cis-dithiolene bond (Figure 1). The sulfur atoms of the dithiolene group were proposed to coordinate the molybdenum atom, with a stoichiometry of one MPT per Mo. Moco is present in more than fifty enzymes in bacteria, including nitrate reductases, sulfite dehydrogenase, xanthine oxidoreductase, aldehyde oxidoreductase, DMSO reductase, formate dehydrogenase, CO dehydrogenase, and various other enzymes [3]. In the redox reactions catalyzed by molybdoenzymes electron transfer is mediated by additional redox centers present in the protein or on other protein domains, like FeS centers, cytochromes, or FAD/FMN cofactors [6]. During these redox reactions, the molybdenum atom can exist in various oxidation states and couple oxide or proton transfer with electron transfer [2].

Figure 1: The different structures of Moco in E. coli. The basic form of Moco is a 5,6,7,8-tetrahydropyranopterin with a unique dithiolene group coordinating the molybdenum atom, named Mo-MPT. Mo-MPT (shown in the tri-oxo structure [25]) can be further modified and in E. coli three different molybdenum-containing enzyme families exist classified according to their coordination at the molybdenum atom: the xanthine oxidase, sulfite oxidase, and DMSO reductase families. In E. coli, the xanthine oxidase family contains the sulfurated MCD cofactor. The sulfite oxidase family is characterized by a di-oxo Moco with an additional protein ligand, which usually is a cysteine. The DMSO reductase family contains two MGDs ligated to one molybdenum atom with additional ligands being an O/S and a sixth ligand X, which can be a serine, a cysteine, a selenocysteine, an aspartate or a hydroxide, and/or water molecule. The characterized molybdoenzymes in E. coli are shown in blue for each family.

The different enzymes contain different forms of Moco and were historically categorized into three families based on the ligands at the molybdenum atom, which are characteristic for each family (Figure 1): the xanthine oxidase (XO) family, the sulfite oxidase (SO) family, and the dimethyl sulfoxide (DMSO) reductase family [3, 7]. The XO family is characterized by an MPT- core in the oxidized state, with one MPT equivalent coordinated to the molybdenum atom, one oxo-group, one sulfido-group, and one hydroxo-group [3] (Figure 1). The sulfido-group is cyanide labile [8–10], and removal of the sulfido group results in formation of an inactive desulfo-enzyme, with an oxygen ligand replacing the sulfur at the Mo active site [10]. The enzymes of this family are involved in two-electron transfer hydroxylation and oxo-transfer reactions with water as the source of oxygen [7, 11]. Among the members of the XO family in E. coli are the xanthine dehydrogenase XdhABC, the periplasmic aldehyde oxidoreductase PaoABC, and the so far uncharacterized xanthine dehydrogenase homologue XdhD (Figure 1, Table 1) [6, 12].

Table 1: Overview on the E. coli molybdoenzymes and the involved chaperones required for maturation/cofactor insertion.

Mo-enzyme
Subunits
Chaperone
Moco

DMSO reductase family

Nitrate reductase A
NarGHI
NarJ
bis-MGD

Nitrate reductase Z
NarZYV
NarW
bis-MGD

Peripl. nitrate reductase
NapABCGH
NapD
bis-MGD

TMAO reductase A
TorAC
TorD
bis-MGD

TMAO reductase Z
TorZY
?
bis-MGD

DMSO reductase
DmsABC
DmsD
bis-MGD

Formate dehydrogenase N
FdnGHI
FdhD
bis-MGD

Formate dehydrogenase O
FdoGHI
FdhD
bis-MGD

Formate dehydrogenase H
FdhF
FdhD
bis-MGD

Biotin sulfoxide reductase
BisC
—
bis-MGD

Selenate reductase
YnfEFGH
DmsD
bis-MGD

?
YdhYVWXUT
?
?

?
YdeP
?
?

Xanthine oxidase family

Aldehyde oxidoreductase
PaoABC
PaoD
MCD

Xanthine dehydrogenase
XdhABC
YqeB
MCD

?
XdhD (YgfM)
YqeB
?

Sulfite oxidase family

Sulfite oxidase
YedYZ
—
Mo-MPT

?
YcbX
—
?

?
YiiM
—
?

—: no chaperone identified.

?: function unknown or not clear which chaperone is involved in maturation, form of Moco not characterized.

Enzymes of the SO family coordinate a single equivalent of the pterin cofactor with an core in its oxidized state and usually an additional cysteine ligand which is provided by the polypeptide [3]. Members of this family catalyze the transfer of an oxygen atom either to or from the substrate [13]. Among the members of this family in E. coli is the YedY protein (Figure 1 and Table 1), for which the physiological role is unknown so far [14].

The DMSO reductase family is diverse in both structure and function, but all members have two equivalents of the pterin cofactor bound to the metal [3]. The molybdenum coordination sphere is usually completed by a single Mo=O group with a sixth ligand in the core; however, a sulfido ligand has been recently described to replace the oxo-group in formate dehydrogenase-H from E. coli [15]. The sixth ligand, X, can be a serine, a cysteine, a selenocysteine, an aspartate or a hydroxide, and/or water molecule [16]. The reactions catalyzed by members of this family frequently involve oxygen-atom transfer, but dehydrogenation reactions also occur. Members of the DMSO reductase family are not present in eukaryotes and include, among other enzymes, the dissimilatory nitrate reductases, formate dehydrogenases, trimethylamine-N-oxide (TMAO) reductases, DMSO reductase, and biotin sulfoxide reductases of E. coli (Figure 1 and Table 1).

This review will give in the first part an overview on the recent discoveries in Moco biosynthesis in E. coli. Since MPT is a sulfur-containing cofactor, the biosynthesis of Moco is linked to the sulfur mobilization in the cell. Two sulfurs are present in MPT which coordinate the molybdenum atom. This sulfur is mobilized from L-cysteine in the cell, by a general pathway which links Moco biosynthesis to the synthesis of other sulfur-containing biomolecules like FeS cluster assembly and the thiolation of tRNA. Additionally, FeS cluster containing proteins are required for the biosynthesis of the first intermediate in Moco biosynthesis. Thus, we will dissect the sulfur transfer pathway for the biosynthesis of Moco and FeS clusters and their connection to the thiolation of tRNA in the second part of the review. The biosynthesis of the molybdenum cofactor in eukaryotes has been additionally studied and is also highly conserved. Since there are numerous reviews available in the literature describing Moco biosynthesis in humans and plants [17–20], we focus only on the biosynthesis in bacteria and the newly discovered connection to other biosynthetic pathways.

2. The Biosynthesis of the Molybdenum Cofactor in E. coli

Using a combination of biochemical, genetic, and structural approaches, Moco biosynthesis has been extensively studied for decades. In all bacteria Moco is synthesized by a conserved biosynthetic pathway that can be divided into four steps, according to the stable biosynthetic intermediates which can be isolated and were intensively studied in Escherichia coli (Figure 2) [21]: the synthesis of cyclic pyranopterin monophosphate (cPMP) [22], conversion of cPMP into MPT by introduction of two sulfur atoms [23], and insertion of molybdate to form Moco [24]. In bacteria, Moco is further modified by the attachment of GMP or CMP to the phosphate group of MPT, forming the dinucleotide variants of Moco, MPT-guanine dinucleotide (MGD) [25–27], and MPT-cytosine dinucleotide (MCD) [28]. After Moco biosynthesis, the different forms of Moco are inserted into the specific target enzymes by the help of molecular Moco-binding chaperones.

Figure 2: The biosynthesis of Moco in E. coli. Shown is a scheme of the biosynthetic pathway for Moco biosynthesis in E. coli and the proteins involved in this pathway. Mo-MPT is formed from 5′GTP with cPMP, MPT, and MPT-AMP as intermediates. Mo-MPT is directly inserted into enzymes of the sulfite oxidase family. For enzymes of the DMSO reductase family, Moco is further modified by formation of a bis-Mo-MPT intermediate and further addition of a GMP molecule to each MPT unit, forming the bis-MGD cofactor. Both reactions are catalyzed by the MobA protein. For enzymes of the xanthine oxidase family in E. coli, Mo-MPT is further modified by the addition of CMP to form the MCD form of the cofactor. Additionally, a terminal sulfur ligand is added to the molybdenum site, generating sulfurated MCD. An additional ligand at the Mo-center usually is a hydroxo-group. The names of the proteins involved in the reactions are colored in red, and additional molecules required for the reactions are shown in blue.

In E. coli five gene loci were identified which are directly involved in the biosynthesis of Moco, moa, mob, moc, moe, and mog, comprising 11 genes in total [21, 29] (Figure 3). The moa locus contains the genes moaABCDE, whose gene products are involved in cPMP formation from 5′GTP and in the formation of MPT [30]. The mob locus contains the genes mobAB, whose gene products are involved in the synthesis of bis-Mo-MPT and bis-MGD from Mo-MPT [31]. The moc locus contains the gene mocA, and the MocA protein is involved in MCD synthesis from Mo-MPT [28]. The moe locus contains the moeA and moeB genes [32]. While MoeA is involved in Mo-MPT formation from MPT-AMP [33], MoeB is involved in the activation of MoaD by adenylation [34]. The mog locus contains the mogA gene, whose gene product is involved in the formation of MPT-AMP [33, 35]. Additional essential reactions are the transport of molybdate [36] and the mobilization of sulfur [37]. The proteins and biochemical reactions involved in the four steps of Moco biosynthesis are described in detail below.

Figure 3: Organization of the genes involved in the biosynthesis of Moco in E. coli. In total, 11 genes are involved in the biosynthesis of Moco in E. coli. These are organized into 5 different gene loci termed moa, mob, moc, moe, and mog. Genes of known function are colored in white. Black dots indicate promotor regions. The genes are drawn approximately to scale. Additional operons with a role for in Moco biosynthesis, such as the isc operon or the mod operon for molybdate transport, are not shown.

2.1. Conversion of 5′GTP to cPMP

The biosynthesis of Moco starts from 5′-GTP, which results in the formation of cPMP, the first stable intermediate of Moco biosynthesis (Figure 4) [22]. The cPMP molecule is an oxygen-sensitive 6-alkyl pterin with a cyclic phosphate group at the C2′ and C4′ atoms [38, 39]. In E. coli, the moaA and moaC gene products are responsible for the complicated chemical reactions required to generate cPMP [32, 40–42]. An in vitro system for cPMP synthesis containing the MoaA and MoaC proteins purified from Staphylococcus aureus showed that 5′-GTP is the specific initial substrate for cPMP biosynthesis [43]. In this reaction, the C8 of GTP is inserted between the C2′ and C3′ carbons of the GTP ribose. MoaA belongs to the superfamily of S-adenosyl methionine- (SAM-) dependent radical enzymes [44]. Members of this family catalyze the formation of protein and/or substrate radicals by reductive cleavage of SAM by [4Fe4S] cluster [45]. MoaA is a protein containing two oxygen-sensitive FeS clusters, each of which is coordinated by only three cysteine residues. The N-terminal [4Fe4S] cluster, present in all radical SAM proteins, binds SAM and carries out the reductive cleavage of SAM to generate the 5′-deoxyadenosyl radical, which subsequently initiates the transformation of 5′-GTP bound through the C-terminal [4Fe4S] cluster [44, 46, 47]. Experiments with GTP isotopes showed that the ribose C3′ hydrogen atom is abstracted by the 5′-deoxyadenosyl radical of MoaA [48]. Further reactions involve the attack of the C8 in the guanine ring by the formed C3′ radical resulting in the formation of (8S)-3′, 8-cyclo-7,8-dihydroguanosine 5′triphosphate (3′,8-cH₂GTP) intermediate (Figure 4) [40, 41]. The additional reducing equivalents required for this step might be provided by the C-terminal [4Fe4S] cluster in MoaA. The intermediate serves as a substrate for MoaC which converts 3′,8-cH₂GTP to cPMP, including pyrophosphate cleavage and formation of the cyclic phosphate group by general acid/base catalysis [40].

Figure 4: Synthesis of cPMP from 5′GTP. All carbon atoms of the 5′GTP are found within cPMP. The C8 atom from the guanine ring is inserted between the C2′ and C3′ atoms of the ribose. This reaction is catalyzed by the MoaA protein, an S-adenosylmethionine- (SAM-) dependent enzyme. MoaA forms a dimer with two [4Fe4S] clusters bound to each monomer. The trimeric MoaC protein is suggested to cleave the pyrophosphate group of the cyclic intermediate. cPMP is shown in the tetrahydropyrano form with a keto group at the C1′ position, as suggested from the crystal structure [55].

2.2. Conversion of cPMP to MPT

The next step involves the conversion of cPMP to MPT in which two sulfur atoms are incorporated in the C1′ and C2′ positions of cPMP [23, 38, 49] (Figure 5). This reaction is catalyzed by MPT synthase, a protein consisting of two small (~10 kDa) and two large subunits (~21 kDa), encoded by moaD and moaE, respectively [50, 51]. It was shown that MPT synthase carries the sulfur in form of a thiocarboxylate at the C-terminal glycine of MoaD [52]. The central dimer is formed by two MoaE subunits containing one MoaD at each end, as revealed by the crystal structure [53]. It was shown that the two MoaD/MoaE dimers act independently. Thus, for the insertion of two sulfurs into cPMP, two MoaD proteins are required at each end of the MPT synthase tetramer [54]. The first sulfur is added by one MoaD molecule at the C2′ position of cPMP (Figure 5), a reaction which is coupled to the hydrolysis of the cPMP cyclic phosphate [55]. During the course of this reaction, a hemisulfurated intermediate is formed in which the MoaD C-terminus is covalently linked to the substrate via a thioester linkage, which subsequently is hydrolyzed by a water molecule. After the transfer of its thiocarboxylate sulfur to cPMP, the first MoaD subunit dissociates from the MPT synthase complex [54, 55]. During the reaction of the first sulfur transfer, the opening of the cyclic phosphate is proposed to shift the location of the intermediate within the protein so that the C1′ position now becomes more accessible to the attack by the second MoaD thiocarboxylate (Figure 4). This results in a second covalent intermediate that is converted to MPT via the elimination of a water molecule and hydrolysis of the thioester intermediate. During the reaction, cPMP and the hemisulfurated intermediate remain bound to one MoaE subunit [56].

Figure 5: The biosynthesis of MPT from cPMP. In the MPT synthase mechanism, cPMP is bound to the MoaE subunit. The initial attack and transfer of the first thiocarboxylated MoaD-SH sulfur atom occurs at the C2′ position of cPMP, coupled to the hydrolysis of the cPMP cyclic phosphate. An intermediate is formed in which the MoaD C-terminus is covalently linked to the substrate via a thioester linkage that is subsequently hydrolyzed by a water molecule to generate a hemisulfurated intermediate at C2′. Opening of the cyclic phosphate shifts the location of the intermediate within the complex to a position where C1′ becomes more accessible. A new MoaD-SH thiocarboxylate attacks the C1′ resulting in a second covalent intermediate which is converted to MPT via the elimination of a water molecule and hydrolysis of the thioester intermediate. During the reaction, cPMP remains bound to the MoaE molecule. The MPT synthase tetramer is built of two MoaE and two MoaD subunits. The MoaD-SH thiocarboxylate is formed on MoeB, where MoaD is first activated under ATP consumption to form an activated MoaD acyl-adenylate. Further, sulfur is transferred from a sulfur relay system by an L-cysteine desulfurase in conjunction with either TusA or YnjE to MoaD. The mechanism was adapted from the one proposed in [55].

The regeneration of sulfur at the C-terminal glycine of MoaD is catalyzed by MoeB [32, 57] and resembles the first step of the ubiquitin-dependent protein degradation system [58] (Figure 4). It was determined that, in E. coli, L-cysteine serves as the origin of the MPT dithiolene sulfurs and that the cysteine sulfur is transferred to the activated MoaD acyl-adenylate by the action of a persulfide-containing protein [37]. After the reaction, MoaD-SH dissociates from the complex and reassociates with MoaE to form active MPT synthase (Figure 5). The binding constants within the different complexes of MoaD were shown to follow the order (MoaD-SH-MoaE)₂ > (MoaD-MoeB)₂ > (MoaD-MoaE)₂ [56, 59]. This order is mechanistically logical given that, during the course of MPT biosynthesis, MoaD-SH first binds to MoaE to form the active MPT synthase complex where transfer of the MoaD-SH thiocarboxylate to cPMP occurs, yielding MPT and inactive MPT synthase. MoaD must then dissociate from this inactive complex to form a new complex with MoeB, a prerequisite for the regeneration of MoaD-SH. In addition, the (MoaD-MoeB)₂ complex is stabilized by ATP addition and the subsequent formation of the acyl-adenylate on MoaD [34]. In this form, the sulfur transfer to MoaD occurs, generating MoaD-SH. In the sulfur transfer reactions the proteins IscS and TusA are involved, forming a sulfur relay system [60]. However, under anaerobic conditions, TusA can be replaced by YnjE and SufS [61]. The exact mechanism of sulfur mobilization and transfer is further described below. After the formation of the (MoaD-SH-MoaE)₂ complex, introduction of the dithiolene moiety in MPT completes the formation of the chemical backbone necessary for binding and coordination of the molybdenum atom in Moco (Figure 2).

2.3. Insertion of Molybdate into MPT

In E. coli insertion of the metal into MPT is accomplished by the moeA and mogA gene products (Figure 2) [24, 32]. The structure showed that MogA is a trimer in solution with each monomer folded into a single compact domain, and MogA binds MPT with high affinity [62]. The crystal structures for E. coli MoeA showed a dimeric structure with an elongated monomer consisting of four distinct domains, one of which was structurally related to MogA [63, 64]. It was shown that the proteins have different functions in the molybdenum chelation reaction [33, 65]. MoeA appeared to mediate molybdenum ligation to newly synthesized MPT in vitro at low concentrations of . This reaction was strongly inhibited by MogA in the absence of ATP, but, in the presence of ATP, MogA doubled the rate of molybdenum ligation [65]. Later, the catalytic formation of an MPT-AMP intermediate during the reaction was shown by the crystal structure of the homologous protein Cnx1 from Arabidopsis thaliana with bound MPT-AMP [35, 66, 67]. The accumulation of a comparable MPT-AMP intermediate in E. coli moeA ⁻ extracts was verified later (Figure 2, unpublished data). After in vitro incubation of MoeA with Mo-MPT, EXAFS studies showed that Mo-MPT is bound in its tri-oxo structure to MoeA [25].

It was shown that under physiological molybdate concentrations (1–10 μM), MogA is required in E. coli to form an MPT-AMP intermediate that facilitates molybdate insertion on the dithiolene sulfurs. However, this reaction is not absolutely required under high molybdenum concentrations in vivo, since mogA cells were rescued for molybdoenzyme activities by the addition of high molybdate concentrations to the medium [68]. This suggested that, under high molybdate concentrations (>1 mM) in the cell, MPT-AMP formation by MogA is not required and molybdate can be directly inserted into MPT with the aid of the MoeA protein [69]. Additionally, it was shown that bivalent copper and cadmium ions as well as trivalent arsenite ions could all be inserted nonspecifically into MPT without the presence of either MoeA or MogA and that copper had a higher affinity for the dithiolene group of MPT than molybdate [69]. Thus, bivalent metal ions in high concentrations might inhibit Moco biosynthesis in E. coli. After its formation, the Mo-MPT cofactor can be directly inserted into enzymes from the sulfite oxidase family without further modification [68], like into the E. coli YedY protein (Figure 1) [70].

2.4. Further Modification of Moco

2.4.1. bis-Mo-MPT and bis-MGD Formation for Enzymes of the DMSO Reductase Family

The proteins of the DMSO reductase family in E. coli contain a dinucleotide derivative of Moco, the MPT-guanine dinucleotide (MGD) cofactor [3]. Additionally, the molybdenum atom is ligated by two dithiolene groups of two MGD moieties, forming the bis-MGD cofactor [27]. The synthesis of the bis-MGD cofactor occurs in a stepwise reaction which only requires Mo-MPT (which was previously formed by the MogA/MoeA reaction), MobA, and Mg-GTP [25]. In the first reaction, the bis-Mo-MPT intermediate is formed on MobA (Figure 2). In the second reaction, two GMP moieties from GTP are added to the C4′ phosphate of each MPT via a pyrophosphate bond leading to release of the β- and γ-phosphates of GTP as pyrophosphate [71]. However, the molecular mechanism of bis-Mo-MPT formation as well as its binding mode to MobA is not clear so far. The crystal structure of MobA showed that the protein is a monomer with an overall α β architecture, in which the N-terminal domain of the molecule adopts a Rossmann fold, and a possible MPT binding site that is localized to the C-terminal half of the protein [72, 73]. Since for bis-Mo-MPT formation two MPT moieties have to be bound to monomeric MobA, this might occur by using the MPT and predicted GTP-binding sites (Figure 6). During this reaction, one molecule of molybdate has to be released when two Mo-MPT molecules are combined; however, the underlying chemistry remains elusive and the release of molybdate has not been proven so far. It also remains possible that MobA binds one Mo-MPT molecule and one MPT molecule from which the bis-Mo-MPT could be formed. After the attachment of two GMP molecules to the bis-Mo-MPT intermediate, bis-MGD is formed [25]. Facilitated release of bis-MGD from MobA in the presence of GTP was observed, which suggested that GTP might compete with the same binding site occupied by the MPT units, thus, resulting in the release of formed bis-MGD. We propose that two MPT moieties bind to both the predicted GTP-binding sites in addition to the predicted MPT binding site on MobA, thus, enabling bis-Mo-MPT and bisMGD synthesis by monomeric MobA (Figure 5). The favourable release only of the final product may then be induced by a different binding mode of bis-MGD compared to bis-Mo-MPT to MobA. Both binding modes can be modeled on MobA, which is shown in Figure 6.

Figure 6: Model of the formation of bis-Mo-MPT and bis-MGD by MobA. It is proposed that two Mo-MPT moieties bind to the proposed MPT and GTP-binding sites on the MobA monomer, thus, enabling first bis-Mo-MPT and then bisMGD synthesis by the addition of two GMP molecules to bis-Mo-MPT. bis-Mo-MPT and bis-MGD are believed to have different binding modes on MobA, resulting in the favourable release of bis-MGD as final product. The structures of MobA were modeled using the coordinates from the Protein Data Bank (1FRW) and the figure was adapted from the one shown in [25].

While the role of MobA in MGD formation of Moco was already discovered in 1991 [74], the role of MobB, the second protein encoded by the mob locus [31], remains uncertain [75]. Based on its crystal structure, it was postulated that MobB could be an adapter protein, acting in concert with MobA to achieve the efficient biosynthesis and utilization of MGD [75]. Another possible role might be that MobB protects formed bis-MGD or prevents its release from MobA until an acceptor protein is present. A docking model of MobA and MobB suggested that GTP is bound to a shared binding site at the interface between both proteins [75, 76]. However, conditions under which MobB is essential for bis-MGD synthesis were not reported so far. However, in some bacteria like Rhodobacter capsulatus, MobB is not present and, thus, is not essential for bis-MGD formation [77]. After bis-MGD biosynthesis, the cofactor is released from MobA and either is bound to Moco-binding chaperones or is inserted into the target molybdoenzymes of the DMSO reductase family [71, 78].

2.4.2. Molecular Chaperones for bis-MGD Insertion into Target Enzymes

The last step of Moco modification including the formation of bis-MGD prepares the cofactor for insertion into the specific apo-enzymes. Until now it is not completely understood how Moco is inserted into the folded protein. The crystal structures of several molybdoenzymes revealed that Moco is deeply buried inside the proteins, at the end of a funnel-shaped passage giving access only to the substrate [79]. The insertion step is catalyzed by Moco-binding molecular chaperones, which bind the respective Moco variant and insert it into the target molybdoenzyme [78]. With a few exceptions, most of the molybdoenzymes of the DMSO reductase family in E. coli have a specific chaperone for Moco insertion (Table 1) [6]: NarJ is the chaperone for nitrate reductase A NarGHI [80], NarW is the chaperone for nitrate reductase Z NarZYV [81], DmsD is the chaperone for DmsABC [82] and YnfE/F [83], and FdhD is the chaperone for FdhF [84]. One well-studied example is the TorD/TorA system for TMAO reductase in E. coli. TorD was shown to be the specific chaperone for TorA [85] and plays a direct role in the insertion of Moco into apoTorA [86]. During this reaction, TorD interacts with both MobA and apoTorA and further stabilizes apoTorA for Moco insertion to avoid a proteolytic attack of the latter. This is consistent with its role as “facilitator” of the bis-MGD insertion and maturation of the apo-enzyme [6, 78, 87]. For the chaperone TorD it was described that it is able to bind to the signal peptide of apo-TMAO reductase until the bis-MGD is inserted and TMAO reductase is correctly folded [87]. Pre-TorA is then translocated to the periplasm where the active TorA enzyme is finally generated after cleavage of the signal peptide [6].

Recently, it was shown that these chaperones not only facilitate the insertion of bis-MGD into the target enzyme, but also directly bind bis-MGD [88]. This was shown by the FdsC-FdsA system for maturation of R. capsulatus formate dehydrogenase (FDH). R. capsulatus FDH consists of (α βγ)₂ heterotrimer in which the large α-subunit FdsA (105 kDa) harbors the bis-MGD cofactor and a set of four [Fe₄S₄] clusters and one [Fe₂S₂] cluster (Figure 7). FdsA is linked to the β-subunit FdsB (52 kDa) that binds one additional [Fe₄S₄] cluster and the FMN cofactor [89]. The γ-subunit FdsG (15 kDa) binds [Fe₂S₂] cluster. The terminal electron acceptor was shown to be NAD⁺. For R. capsulatus FDH so far two proteins, named FdsC and FdsD, were identified to be essential for the production of an active FDH, but are not subunits of the mature enzyme [89]. While FdsD has only counterparts in some oxygen-tolerant FDHs, FdsC shares high amino acid sequence homologies to E. coli FdhD, the chaperone for the membrane-bound FDH (FdhF) [84]. FdsC was directly copurified with bound bis-MGD [88]. The definitive proof that bis-MGD was bound to FsdC and not only MGD was given by the reconstitution of E. coli TMAO reductase activity in a system solely consisting of FdsC and apo-TMAO reductase. It also was concluded that, in addition to bis-MGD binding, FdsC might have a similar role in the maturation of FDH like TorD for TMAO reductase. For TorD it was shown that it interacts with MobA and TorA [78, 87]. For FdsC an interaction with MobA, FdsD, and FdsA was shown. Thus, FdsC might also act as a platform connecting bis-MGD biosynthesis and its insertion into the target protein (Figure 7). FdsC binds directly bis-MGD and therefore might protect the bis-MGD cofactor from oxidation before its insertion into FdsA. Alternatively, it might be the factor that determines the specificity for bis-MGD insertion into FDH. The FdsD protein seems to be additionally essential in this reaction. Since FdsD was only identified in organisms which contain an oxygen-tolerant FDH, its role might also serve to protect the bis-MGD cofactor specifically in the presence of oxygen.

Figure 7: Model for the role of FdsC in bis-MGD insertion into R. capsulatus FDH. Shown is a scheme for bis-MGD biosynthesis from two Mo-MPT molecules via a bis-Mo-MPT intermediate. MobA binds Mo-MPT and catalyzes first the synthesis of bis-Mo-MPT and then the synthesis of bis-MGD by ligation of two GMP molecules to each MPT moiety in bis-Mo-MPT. Further, bis-MGD is transferred from MobA to FdsC. FdsC, FdsD, and MobA were shown to form a complex and are believed to protect the bis-MGD cofactor from oxidation before its insertion into the FdsA subunit of R. capsulatus FDH. Additionally, FdsC is believed to contribute to the specificity for bis-MGD insertion into FDH. The role of FdsD in the reaction is not completely resolved so far.

Additionally, it was reported that the bis-MGD cofactor can be further modified by sulfuration [15, 16, 90, 91]. Reinterpretation of the original crystal data of FdhF suggested that, at the molybdenum site, the apical ligand is rather a sulfur ligand instead of an oxygen ligand [15]. In the oxidized state, the enzyme contains the four pterin sulfur ligands at the Mo site, a selenocysteine ligand, and a −SH ligand. The chaperone involved in sulfuration of the Moco for FdhF was shown to be the FdhD protein [84]. For the E. coli FdhD protein it was reported that it acts as a sulfurtransferase between the L-cysteine desulfurase IscS and FdhF, a mechanism which is essential to yield active FdhF [84]. Conclusively, the additional role of bis-MGD binding chaperones might be the further modification of Moco by sulfuration. In addition to FDHs, the DMSO reductase family includes other members for which an additional sulfur ligand of the molybdenum atom has been reported at the catalytic site. The X-ray crystal structure of the periplasmic nitrate reductase (Nap) of Cupriavidus necator showed the presence of a terminal sulfur ligand at the molybdenum coordination sphere [90]. Similar data were obtained for the homologous NapA protein from Desulfovibrio desulfuricans ATCC 27774, for which the crystal structure showed a unique coordination sphere of six sulfur ligands bound to the molybdenum atom [91]. These observations might suggest that sulfuration of bis-MGD is more common of this group of enzymes than previously expected. In total, we believe that bis-MGD binding to chaperones for enzymes of the DMSO reductase family might be a common feature, which might serve as a platform for further modification of Moco or the specific interaction with target enzymes for bis-MGD insertion.

2.4.3. MCD Formation and Further Modification by MCD Sulfuration for Enzymes of the Xanthine Oxidase Family

More recently, three enzymes were identified in E. coli belonging to the xanthine oxidase family (XdhABC, XdhD, and PaoABC) which bind the MCD form of the cofactor (Figure 1) [92–94]. MCD formation is catalyzed by a protein which was named MocA, for molybdopterin cytosine dinucleotide synthesis (Figures 2 and 3) [28]. MocA was identified by amino acid sequence comparison to MobA, since they exhibit 22% amino acid sequence identity [28, 95]. The catalytic reaction of MocA is similar to the reaction of MobA, in that it acts as a CTP: molybdopterin cytidylyltransferase and covalently links MPT and CMP with the concomitant release of the β- and γ-phosphates of CTP as pyrophosphate [28]. However, bis-MCD is not the end product of the reaction; instead, only one MCD moiety is ligated to the molybdenum atom in this cofactor variant. Therefore, the end product of MocA must be a monopterin MCD cofactor. Instead, the MCD cofactor for enzymes of the xanthine oxidase family is further modified in E. coli and contains an equatorial sulfido ligand at its active site [12]. Thus, MCD sulfuration might prevent bis-MCD formation.

Comparison of the two MPT nucleotidyl transferases showed that MobA is highly specific for binding of the purine nucleotide GTP, while MocA is specific for binding of the pyrimidine nucleotide CTP [28]. The most significant sequence differences between the two proteins were observed in two conserved motifs at their N-terminal domain. The crystal structure of MobA with bound GTP showed that the guanine moiety is mainly bound by the ¹²LAGG¹⁵ and ⁷⁸GPLAG⁸² amino acid sequence segments [72, 96]. In MocA, these sequences are altered to ¹²TAAG¹⁵ and ⁷⁸GLLTS⁸² [95]. Site directed mutagenesis studies showed that the introduction of only 5 amino acid exchanges in the two N-terminal regions of either MobA or MocA was sufficient to cause loss of specificity for the pyrimidine or purine nucleotides so that both proteins were able to bind either CTP or GTP to almost the same extent. In addition, the C-terminal domains of MocA and MobA have been found to play an important role in determining the specificity of their interaction with the target molybdoenzymes [95].

After MCD formation, the cofactor is handed over to MCD binding chaperones, which are PaoD or YqeB in E. coli (Table 1). PaoD is the specific Moco-binding chaperone for the periplasmic aldehyde oxidoreductase PaoABC in E. coli [6]. PaoD belongs to the XdhC family of Moco-binding chaperones [12]. The best characterized chaperone from this family is the R. capsulatus XdhC protein which is essential for the maturation of R. capsulatus xanthine dehydrogenase (XDH) [12]. Investigation of R. capsulatus XdhC showed that it binds the Moco produced by MoeA/MogA and protects it from oxidation until the terminal molybdenum sulfur ligand is inserted [97, 98]. XdhC also interacts with the R. capsulatus L-cysteine desulfurase, NifS4, the protein that actually replaces the cofactor equatorial oxygen ligand with a sulfido ligand [99]. The sulfur for this reaction originates from L-cysteine, and NifS4 persulfide group is formed during the course of the reaction. After the sulfuration reaction, it is believed that XdhC with its bound sulfurated Moco dissociates from NifS4 and forms a new interaction with the XdhB subunits of the R. capsulatus (α β)₂ XDH heterotetramer [98, 100]. Thus, it appears from the R. capsulatus studies that XdhC-like proteins perform a number of functions including stabilization of the newly formed Moco and interaction with an L-cysteine desulfurase to ensure that Moco sulfuration occurs [99] as well as interaction with their specific target proteins for insertion of the sulfurated Moco [12]. Because Moco is deeply buried in the protein, it is also believed that the XdhC proteins may act as chaperones to facilitate the proper folding of the target proteins after Moco insertion [101]. This model implies that molybdoenzymes requiring the sulfurated form of Moco exist in a Moco competent “open” apo-molybdoenzyme conformation until the insertion of sulfurated Moco. After insertion, the protein adapts the final active “closed” conformation that can no longer accept Moco [101].

Since PaoD belongs to the XdhC family, it is expected to play a role similar to that of XdhC with the only difference being that PaoD binds the MCD cofactor rather than an MPT cofactor (Figure 8) [102]. The specific L-cysteine desulfurase involved in the sulfuration of PaoD-bound MCD has not been identified in E. coli to date, but it is expected that IscS performs this role in E. coli. The only other XdhC-like protein that is present in E. coli is the YqeB protein (Table 1). YqeB is expected to play a similar and shared role for both XdhABC and XdhD, since it is located in the vicinity of both gene regions in the E. coli genome [12]. YqeB is a little larger in size than other members of the XdhC family and it contains an NAD(P) binding Rossman fold. So far, no further data are available for YqeB.

Figure 8: Sulfur transfer to sulfur-containing biomolecules involving the IscS-bound persulfide. A protein-bound persulfide group is formed on the L-cysteine desulfurase IscS, which is further transferred to proteins like IscU, TusA, or ThiI. CyaY and IscX were also shown to bind to IscS. IscU is the primary scaffold for the assembly of FeS clusters. The IscS sulfur can also be further transferred for the thiolation of tRNA. In E. coli tRNA modification can be divided into an FeS cluster independent pathway and into a pathway which requires FeS clusters. The synthesis of s²C by TtcA and ms²i⁶A by MiaB require sulfur from the FeS clusters initially formed on IscU. ThiI can transfer sulfur either for the formation of thiamine or for the formation of s⁴U in tRNA. TusA can transfer sulfur for a sulfur relay system involving TusBCD, TusE, and MnmA for the formation of s²U in tRNA, or additionally TusA is involved in sulfur transfer for Moco biosynthesis. Persulfide-containing proteins are highlighted in red and names of the final sulfur-containing molecules are colored in blue.

3. The Distribution of Sulfur for Sulfur-Containing Biomolecules in Bacteria

L-Cysteine is the source for the biosynthesis of Moco and a variety of other biomolecules such as thiamin, iron-sulfur (FeS) clusters, and thionucleosides in transfer RNA (tRNA) (Figure 8) [103]. In E. coli the L-cysteine sulfur for these biomolecules is initially mobilized by the house-keeping L-cysteine desulfurase IscS [104]. The enzyme is a pyridoxal 5′-phosphate-containing homodimer that decomposes L-cysteine to L-alanine and sulfane sulfur via the formation of an enzyme-bound persulfide intermediate [104–106]. The persulfide sulfur from the L-cysteine desulfurase is further incorporated either directly or via sulfur relay systems into the biosynthetic pathways of several sulfur-containing biofactors, thus providing an elegant mechanism for making sulfur atoms available without releasing them in solution [107, 108]. IscS is encoded by a gene that is part of a larger operon, iscRSUA-hscBA-fdx-iscX, in which the other gene products are involved in the biosynthesis of FeS clusters in E. coli (Figure 9) [104]. For FeS cluster biosynthesis IscS interacts with IscU, which serves as a scaffold protein for FeS cluster assembly [109, 110]. However, IscU is not the only interaction partner of IscS, since IscS was shown to interact with a number of other proteins for delivery of sulfur to other sulfur-containing molecules, such as CyaY, IscX, TusA, and ThiI (Figure 8) [107, 110]. While CyaY and IscX are also involved in FeS cluster biosynthesis [110–112], ThiI and TusA are involved in tRNA thiolation (ThiI and TusA) [113, 114], thiamine biosynthesis (ThiI) [115], or Moco biosynthesis (TusA) [60]. Thus, the persulfide sulfur from IscS can be transferred in a relay flow to the conserved cysteine residues of acceptor proteins [116]. This chapter will describe how the sulfur transfer pathways for FeS clusters, tRNA, and Moco are connected and which factors determine the direction of the sulfur transfer mechanism to the various biomolecules.

Figure 9: General scheme for FeS cluster biosynthesis in E. coli. Shown is the isc operon containing the genes iscR-iscSUA-hscAB-fdx-iscX. IscR is the regulator for expression of the operon by sensing the FeS status of the cell. FeS clusters assemble on the scaffold protein IscU, which receives the sulfur from the L-cysteine desulfurase IscS. The iron donor in this reaction is not identified so far. CyaY is believed to stabilize a conformation of the IscS-IscU complex. The role of IscX is unclear. Ferredoxin is proposed to participate in the reductive coupling of two [2Fe2S] clusters to form a single [4Fe4S] cluster on IscU. Release of the clusters is catalyzed by the chaperones HscAB. The carrier protein IscA delivers the formed FeS clusters to the final target proteins.

3.1. The Biosynthesis of FeS Clusters

FeS clusters are among the earliest catalysts in the evolution of biomolecules and serve as electron carriers in redox reactions, regulatory sensors, stabilizers of protein structure, and chemical catalysts [103]. The most abundant types are [2Fe2S] clusters and [4Fe4S] clusters. Although FeS clusters can be formed spontaneously in vitro with inorganic iron and sulfide [117], the in vivo synthesis is a complex process which is highly regulated in the cell [118].

The isc operon encodes for IscS, the scaffold protein IscU, ferredoxin (Fdx), the scaffold protein IscA, the two molecular chaperones HscA and HscB, and IscX a protein of unknown function (Figure 9) [104]. The isc operon is controlled by IscR, FeS cluster binding transcriptional repressor [119]. The scaffold protein IscU is the central protein in FeS cluster biosynthesis and is responsible for the assembly of [2Fe2S] and [4Fe4S] clusters [109]. IscS provides the inorganic sulfur for FeS clusters [120]. In the presence of L-cysteine and Fe²⁺, IscS and IscU form a transient macromolecular complex and form sequentially [2Fe2S] clusters and then from two [2Fe2S] a [4Fe4S] cluster on IscU [121]. The sulfur is bound on IscS as a persulfide on Cys-328 and is subsequently transferred to the scaffold IscU. The crystal structure of the IscU-IscS complex showed that each IscU molecule interacts with one IscS molecule in a 1 : 1 stoichiometry [110]. In an intermediate structure, the Cys-328 residue of IscS is used as a ligand of the transient [2Fe2S] cluster together with the three cysteines on IscU [121]. Further, ferredoxin was proposed to participate in the reductive coupling of two [2Fe2S] clusters to form a single [4Fe4S] cluster on IscU [122]. The cluster is then subsequently transferred to a recipient apoprotein, which can be facilitated by the help of IscA [123]. The two chaperones HscA and HscB help in the formation and transfer of the [4Fe4S] cluster [124]. CyaY, the E. coli frataxin homologue, has also a role in FeS cluster biosynthesis and was shown to regulate the biosynthesis by competing about the binding site of IscS with other proteins like IscX [110, 112, 125]. Upon binding to IscS, CyaY was shown to inhibit FeS cluster biosynthesis. It has been suggested that CyaY binding may induce a conformational change in the IscS-IscU complex that does not significantly affect the IscS L-cysteine desulfurase activity, but abolishes FeS cluster synthesis [126]. This might occur by stabilizing a conformation on the flexible loop of IscS which disfavors FeS cluster assembly. The role of CyaY as an iron donor in this process is not completely resolved so far [127]. During stress conditions such as iron starvation and oxidative or heavy metal stress, the SufS protein is rather involved in biosynthesis of FeS clusters, replacing IscS in some functions [128, 129].

In total, FeS clusters are found to participate in diverse biological processes such as respiration, central metabolism DNA, or gene regulation. FeS clusters can act as catalysts or redox sensors and are predicted to be used by over 150 proteins in E. coli [118]. Among the FeS cluster containing molybdoenzymes are nitrate reductase, DMSO reductase and formate dehydrogenase, involved, for example, in anaerobic respiration [6].

3.2. The Thiolation of tRNA

TusA and ThiI are two proteins which are involved in the posttranscriptional modification of tRNA by producing thiolated nucleoside species at specific positions in the tRNA (Figure 8) [114, 130]. Thiolated nucleosides are found in several tRNAs. In E. coli these are s⁴U8, s²C32, ms²i(o)⁶A37, and mnm⁵s²U34, which, with the exception of s⁴U8, are located within the anticodon loop and are crucial for proper mRNA decoding (Figure 10) [107]. The base thiolations are mediated by several acceptor proteins, which are divided into an FeS cluster dependent and an independent pathway. In the iron-sulfur cluster independent pathway, direct transfer of sulfur from IscS to the acceptor ThiI leads to the s⁴U8 modification at position 8 in bacterial tRNAs (Figure 10) [113, 131]. The s⁴U serves as a near UV-photosensor since it undergoes photocrosslinking with a cytidine at position 13, which causes a growth delay since cross-linked tRNAs are inefficient aminoacylation substrates. However, ThiI has a dual role and also participates in thiamine biosynthesis by providing the sulfur of the thiazole species in thiamine pyrophosphate (Figure 12) [115].

Figure 10: Chemical structure of thio-containing tRNA modifications. Shown are the secondary structure and the positions of thiolated nucleosides in tRNA. Abbreviations: s⁴U, 4-thiouridine; s²U, 2-thiocytidine; mnm⁵s²U, 5-methylaminomethyl-2-thiouridine; ms²i⁶A, 2-methylthio-N6-isopentenyladenosine; ms²io⁶A, N6-(4-hydroxyisopentenyl)-2-methylthioadenosine. Each number represents the nucleoside position in the tRNA. On the left-hand side the sulfur transfer pathway for the formation of s²U is shown. The sulfur transfer involves the initial sulfur mobilization by IscS, sulfur transfer to TusA-C19, and further transfer via TusD in the TusBCD complex, TusE to MnmA, which binds and activates the tRNA by adenylation. After formation, s²U is further modified to form cmnm⁵s²U and mnm⁵s²U.

In contrast, the TusA protein was identified to function as a sulfur mediator for the synthesis of 2-thiouridine of the modified wobble base 5-methylaminomethyl (mnm)⁵s²U in tRNA (Figure 10) [114]. It interacts with IscS and stimulates its L-cysteine desulfurase activity 3-fold [114]. TusA transfers the sulfur from Cys19 to Cys78 of TusD in the TusBCD complex in a TusE-dependent manner. TusE is likely to accept the sulfur from TusD to form a persulfide of Cys108. Finally, TusE transfers the sulfur to MnmA, which directly incorporates sulfur into the tRNA wobble position. MnmA has a P-loop in its active site that recognizes tRNA and activates the C2 position of the uracil ring at position 34 by forming an adenylate intermediate [132–134]. In total, little is known about the function of the modification of 2-thiouridine. It is proposed that in E. coli thio-modified confers efficient ribosome binding and 2-thio-modified is required for specific recognition by glutaminyl-tRNA synthetase. It also assists proper codon-anticodon base pairing at the ribosome A site and prevents frameshifting during translation. Thus, the 2-thio-modification of uridine 34 plays a critical role in the decoding mechanism [135].

The modification of tRNA also seems to affect lambda phage infection in E. coli [136, 137]. The process of phage development is dependent on tRNA thiolation. It was shown that a tusA mutant is more resistant to lambda phage infection because of a reduced tRNA modification efficiency and as a consequence has a defect in an associated frame-shift reprogramming control. In contrast, E. coli hscA, hscB, or iscU mutants were shown to be hypersensitive to lambda phage infection [137]. Inactivation of the FeS assembly machinery increases sulfur flux to TusA, resulting in an increased tRNA thiolation and as a consequence a “phage hypersensitivity” phenotype. Thus, tRNA modification and FeS cluster biosynthesis seem to be linked by the control of the sulfur flux to each pathway, which influence each other.

3.3. Sulfur Transfer for Moco Biosynthesis

The first step in Moco biosynthesis, the conversion of 5′GTP to cPMP, directly depends on the availability of FeS clusters [43]. The MoaA protein belongs to the class of radical SAM enzymes and contains two [4Fe4S] clusters which are essential for its activity (Figure 4) [45]. Thus, when no FeS clusters are present in the cell, cPMP is not formed. Further, for the formation of MPT from cPMP, two sulfur molecules are inserted [53]. The primary sulfur for the dithiolene group of the MPT backbone of Moco was shown to be the IscS protein [37]. Further studies suggested that TusA is additionally involved in sulfur transfer for the synthesis of MPT, by a balanced regulation of the availability of IscS to various biomolecules in E. coli (Figure 5) [60]. It was shown that deletion of tusA in E. coli affected the activity of molybdoenzymes under aerobic and anaerobic conditions. Characterization of the tusA strain under aerobic conditions showed an overall low MPT content and an accumulation of cPMP. Under anaerobic conditions the activity of nitrate reductase was only 50% reduced, showing that TusA is not essential for Moco biosynthesis and can be replaced by other proteins. One sulfur-carrier is the rhodanese-like protein YnjE, which was shown to be preferentially sulfurated by IscS [61]. The expression of ynjE is increased under anaerobic conditions and in the absence of TusA, thus making it more available for sulfur transfer under these conditions. Additionally, SufS can replace IscS in its role in providing the sulfur under certain conditions [60].

However, overexpression of IscU reduced the level of active molybdoenzymes in E. coli (Figure 11). When IscU is present in high amounts, it forms a complex with IscS, making it unavailable for the interaction with TusA, thus resulting in a lack of sulfur transfer for the conversion of cPMP to MPT. A similar situation was obtained when TusA was overexpressed in E. coli (Figure 11). Here, the levels of FeS clusters are decreased which further result in an inactive MoaA protein and thus in a decreased activity of molybdoenzymes.

Figure 11: Model for the effect of elevated concentrations of TusA or IscU on Moco biosynthesis. IscS interacts with IscU, TusA, and ThiI for sulfur transfer to FeS clusters, tRNA, Moco biosynthesis, and thiamine/tRNA, respectively. FeS clusters are inserted into target proteins. Among these proteins is MoaA, involved in Moco biosynthesis (conversion of 5′GTP to cPMP). TusA is involved in the sulfur transfer for the conversion of cPMP to MPT. Increased levels of IscU reduce the level of active molybdoenzymes in E. coli. When IscU is present in high amounts, it forms a complex with IscS, making it unavailable for the interaction with TusA, thus, resulting in a lack of sulfur transfer for the conversion of cPMP to MPT. When the amount of TusA is increased, the levels of FeS clusters are decreased which further result in an inactive MoaA protein and thus in a decreased activity of molybdoenzymes. Detailed descriptions are given in the text.

Figure 12: Model for the connection of Moco biosynthesis, FeS cluster assembly, thiamine biosynthesis, and tRNA thiolation in E. coli. Shown is a general scheme for the sulfur mobilization by IscS, the interaction partners of IscS and the pathways for the synthesis of s⁴U and thiamine involving ThiI, FeS clusters assembly involving IscU, Moco biosynthesis involving TusA and YnjE, and s²U formation involving TusA. FeS clusters are also required for Moco biosynthesis, connecting both pathways. The acceptor proteins for Moco are shown in addition to their localization in the cell. Detailed descriptions are given in the text.

Conclusively, the pleiotropic effects of a tusA deletion might be caused by changes in the FeS cluster concentration, leading to differences in the sulfur transfer to tRNA and for Moco biosynthesis. It was proposed that TusA is involved in regulating the IscS pool and shifting it away from IscU, thereby making IscS available for sulfur transfer for the biosynthesis of MPT. Additionally, major changes in the gene regulation were observed.

3.4. The Complex Network of Sulfur Transfer to Various Biomolecules

It seems that a lot of pathways are regulated by the availability of FeS clusters in the cell, since either they require directly FeS clusters or their synthesis is indirectly regulated by transcriptional regulators that depend on FeS clusters in the cell [138, 139]. Under these conditions when FeS clusters are required, the isc operon is induced and IscS then forms a complex with IscU for FeS cluster biogenesis, making it unavailable for the synthesis of other sulfur-containing biomolecules [138]. However, the other interaction partners of IscS like TusA and ThiI also direct the sulfur for important cellular processes like thiamine biosynthesis, tRNA thiolation, and Moco biosynthesis (Figure 12) [110]. It thus seems likely that, within the cell, the relative affinities of the interaction partners of IscS in addition to their concentration will determine which is the preferred partner protein that interacts with IscS. It was shown that IscU and TusA are not able to bind simultaneously to IscS, due to steric clashes of both proteins upon binding [110]. Since IscU binds with higher affinity to IscS than TusA, this would support the synthesis of FeS clusters under sulfur limitation in the cell [125]. Since the binding sites of ThiI and TusA also overlap, this would suggest that tRNA modification for s⁴U and s²U formation is a competitive process [110].

Conclusively, changes in the concentration of the interaction partners of IscS in the cell globally affect pathways requiring sulfur. Microarray analysis of a tusA strain showed that the expression of genes regulated by FNR (hypABCDE, narGHJI, moaABCDE, soxS, cyoABCDE, sdhAB, tdcABCDEFG, and feoB) was increased [60]. The same effect was observed after overexpression of FNR in E. coli under aerobiosis, which led to the induction of the isc operon due to a higher FeS cluster demand in the cell [138]. A higher level of FeS clusters in the cell stabilizes holoFNR, thus stimulating the transcription of FNR regulated genes (like the genes for NR) [138]. Conclusively, the absence of TusA changes the level of IscS which is available for FeS cluster biosynthesis. However, a higher level of FeS clusters in the cell reduces the sulfur transfer of IscS to other biosynthetic pathways like Moco, thiamine, or thiolated tRNA. This was shown by an overexpression of the isc operon, which resulted in a decreased activity of molybdoenzymes [60]. The same effect was obtained after overexpression of IscU alone or the absence of TusA in the cell, which both decreased the amount of active nitrate reductase or TMAO reductase. Thus, the availability and amount of IscS have an effect on various pathways in the cell.

Since FeS clusters are very important for cellular processes, it is conclusively suggested that one part of IscS in the cell is mostly in complex with IscU for FeS cluster biosynthesis and the other part might be available for the other interaction partners like TusA or ThiI (Figure 12). Thus, during FeS cluster formation, a portion of IscS would not be available for acceptor proteins like TusA or ThiI. When the concentration of one of the interaction partners is changed (as tested for IscU and TusA), the IscS pool is shifted to one or the other direction, with a drastic effect on gene regulation. Conclusively, our studies show that the pleiotropic effect of a tusA deletion might be caused by changes in the FeS cluster concentration in the cell leading to major differences in gene regulation [138]. It is proposed that TusA is involved in regulating the IscS pool and shifting it away from IscU, thereby making IscS available for sulfur transfer for the biosynthesis of MPT, thiamine, and thiolated tRNA.

However, TusA might have additional roles in the cell, since Ishii et al. [140] showed that the FtsZ-ring formation appeared to be severely impaired in tusA-deficient E. coli cells resulting in formation of a nondivided filamentous cell. These effects need to be clarified in future studies.

Conflict of Interests

The author declares that there is no conflict of interests regarding the publication of this paper.

Acknowledgments

The author thanks all current and former members of the research group in addition to collaboration partners who were involved in this work over the past years and decades. Special thanks go to K. V. Rajagopalan, the founder of the field of Moco biosynthesis, for his support over the years and the helpful discussions. The work was mainly supported by continuous grants of the Deutsche Forschungsgemeinschaft.

References

M. P. Coughlan, “The role of molybdenum in human biology,” Journal of Inherited Metabolic Disease, vol. 6, supplement 1, pp. 70–77, 1983.

R. Hille, “Molybdenum and tungsten in biology,” Trends in Biochemical Sciences, vol. 27, no. 7, pp. 360–367, 2002.

R. Hille, “The mononuclear molybdenum enzymes,” Chemical Reviews, vol. 96, no. 7, pp. 2757–2816, 1996.

G. Schwarz, R. R. Mendel, and M. W. Ribbe, “Molybdenum cofactors, enzymes and pathways,” Nature, vol. 460, no. 7257, pp. 839–847, 2009.

K. V. Rajagopalan, J. L. Johnson, and B. E. Hainline, “The pterin of the molybdenum cofactor,” Federation Proceedings, vol. 41, no. 9, pp. 2608–2612, 1982.

C. Iobbi-Nivol and S. Leimkuhler, “Molybdenum enzymes, their maturation and molybdenum cofactor biosynthesis in Escherichia coli,” Biochimica et Biophysica Acta, vol. 1827, pp. 1086–1101, 2013.

R. Hille, T. Nishino, and F. Bittner, “Molybdenum enzymes in higher organisms,” Coordination Chemistry Reviews, vol. 255, no. 9-10, pp. 1179–1205, 2011.

S. Gutteridge, S. J. Tanner, and R. C. Bray, “Comparison of the molybdenum centres of native and desulpho xanthine oxidase. The nature of the cyanide-labile Sulphur atom and the nature of the proton-accepting group,” Biochemical Journal, vol. 175, no. 3, pp. 887–897, 1978.

M. P. Coughlan, J. L. Johnson, and K. V. Rajagopalan, “Mechanisms of inactivation of molybdoenzymes by cyanide,” The Journal of Biological Chemistry, vol. 255, no. 7, pp. 2694–2699, 1980.

R. C. Wahl and K. V. Rajagopalan, “Evidence for the inorganic nature of the cyanolyzable Sulfur of molybdenum hydroxylases,” The Journal of Biological Chemistry, vol. 257, no. 3, pp. 1354–1359, 1982.

R. Hille and R. F. Anderson, “Coupled electron/proton transfer in complex flavoproteins: solvent kinetic isotope effect studies of electron transfer in xanthine oxidase and trimethylamine dehydrogenase,” The Journal of Biological Chemistry, vol. 276, no. 33, pp. 31193–31201, 2001.

S. Leimkühler and M. Neumann, “The role of system-specific molecular chaperones in the maturation of molybdoenzymes in bacteria,” Biochemistry Research International, vol. 2011, Article ID 850924, 13 pages, 2011.

M. S. Brody and R. Hille, “The kinetic behavior of chicken liver sulfite oxidase,” Biochemistry, vol. 38, no. 20, pp. 6668–6677, 1999.

S. J. Brokx, R. A. Rothery, G. Zhang, D. P. Ng, and J. H. Weiner, “Characterization of an Escherichia coli sulfite oxidase homologue reveals the role of a conserved active site cysteine in assembly and function,” Biochemistry, vol. 44, no. 30, pp. 10339–10348, 2005.

H. C. A. Raaijmakers and M. J. Romão, “Formate-reduced E. coli formate dehydrogenase H: the reinterpretation of the crystal structure suggests a new reaction mechanism,” Journal of Biological Inorganic Chemistry, vol. 11, no. 7, pp. 849–854, 2006.

S. Grimaldi, B. Schoepp-Cothenet, P. Ceccaldi, B. Guigliarelli, and A. Magalon, “The prokaryotic Mo/W-bisPGD enzymes family: a catalytic workhorse in bioenergetic,” Biochimica et Biophysica Acta, vol. 1827, pp. 1048–1085, 2013.

F. Bittner, “Molybdenum metabolism in plants and crosstalk to Iron,” Frontiers in Plant Science, vol. 5, article 28, 2014.

R. R. Mendel and T. Kruse, “Cell biology of molybdenum in plants and humans,” Biochimica et Biophysica Acta—Molecular Cell Research, vol. 1823, no. 9, pp. 1568–1579, 2012.

G. Schwarz, R. R. Mendel, and M. W. Ribbe, “Molybdenum cofactors, enzymes and pathways,” Nature, vol. 460, no. 7257, pp. 839–847, 2009.

R. R. Mendel, “The molybdenum cofactor,” The Journal of Biological Chemistry, vol. 288, pp. 13165–13172, 2013.

K. V. Rajagopalan, “Biosynthesis of the molybdenum cofactor,” in Escherichia Coli and Salmonella Cellular and Molecular Biology, F. C. Neidhardt, Ed., pp. 674–679, ASM Press, Washington, DC, USA, 1996.

M. M. Wuebbens and K. V. Rajagopalan, “Structural characterization of a molybdopterin precursor,” The Journal of Biological Chemistry, vol. 268, no. 18, pp. 13493–13498, 1993.

D. M. Pitterle, J. L. Johnson, and K. V. Rajagopalan, “In vitro synthesis of molybdopterin from precursor Z using purified converting factor. Role of protein-bound Sulfur in formation of the dithiolene,” The Journal of Biological Chemistry, vol. 268, no. 18, pp. 13506–13509, 1993.

M. S. Joshi, J. L. Johnson, and K. V. Rajagopalan, “Molybdenum cofactor biosynthesis in Escherichia coli mod and mog mutants,” Journal of Bacteriology, vol. 178, no. 14, pp. 4310–4312, 1996.

S. Reschke, K. G. Sigfridsson, P. Kaufmann, et al., “Identification of a Bis-molybdopterin intermediate in molybdenum cofactor biosynthesis in Escherichia coli,” The Journal of Biological Chemistry, vol. 288, pp. 29736–29745, 2013.

T. Palmer, A. Vasishta, P. W. Whitty, and D. H. Boxer, “Isolation of protein FA, a product of the mob locus required for molybdenum cofactor biosynthesis in Escherichia coli,” European Journal of Biochemistry, vol. 222, no. 2, pp. 687–692, 1994.

J. C. Hilton and K. V. Rajagopalan, “Identification of the molybdenum cofactor of dimethyl sulfoxide reductase from Rhodobacter sphaeroides f. sp. denitrificans as bis(molybdopterin guanine dinucleotide)molybdenum,” Archives of Biochemistry and Biophysics, vol. 325, no. 1, pp. 139–143, 1996.

M. Neumann, G. Mittelstädt, F. Seduk, C. Iobbi-Nivol, and S. Leimkühler, “MocA is a specific cytidylyltransferase involved in molybdopterin cytosine dinucleotide biosynthesis in Escherichia coli,” The Journal of Biological Chemistry, vol. 284, no. 33, pp. 21891–21898, 2009.

K. T. Shanmugam, V. Stewart, R. P. Gunsalus et al., “Proposed nomenclature for the genes involved in molybdenum metabolism in Escherichia coli and Salmonella typhimurium,” Molecular Microbiology, vol. 6, no. 22, pp. 3452–3454, 1992.

S. L. Rivers, E. McNairn, F. Blasco, G. Giordano, and D. H. Boxer, “Molecular genetic analysis of the moa operon of Escherichia coli K-12 required for molybdenum cofactor biosynthesis,” Molecular Microbiology, vol. 8, no. 6, pp. 1071–1081, 1993.

C. Lobbi-Nivol, T. Palmer, P. W. Whitty, E. McNairn, and D. H. Boxer, “The mob locus of Escherichia coli K12 required for molybdenum cofactor biosynthesis is expressed at very low levels,” Microbiology, vol. 141, no. 7, pp. 1663–1671, 1995.

M. E. Johnson and K. V. Rajagopalan, “Involvement of chlA, E, M, and N loci in Escherichia coli molybdopterin biosynthesis,” Journal of Bacteriology, vol. 169, no. 1, pp. 117–125, 1987.

J. Nichols and K. V. Rajagopalan, “Escherichia coli MoeA and MogA: function in metal incorporation step of molybdenum cofactor biosynthesis,” The Journal of Biological Chemistry, vol. 277, no. 28, pp. 24995–25000, 2002.

S. Leimkühler, M. M. Wuebbens, and K. V. Rajagopalan, “Characterization of Escherichia coli MoeB and its involvement in the activation of molybdopterin synthase for the biosynthesis of the molybdenum cofactor,” The Journal of Biological Chemistry, vol. 276, no. 37, pp. 34695–34701, 2001.

J. Kuper, A. Llamas, H.-J. Hecht, R. R. Mendel, and G. Schwarz, “Structure of the molybdopterin-bound Cnx1G domain links molybdenum and copper metabolism,” Nature, vol. 430, no. 7001, pp. 803–806, 2004.

J. A. Maupin-Furlow, J. K. Rosentel, J. H. Lee, U. Deppenmeier, R. P. Gunsalus, and K. T. Shanmugam, “Genetic analysis of the modABCD (molybdate transport) operon of Escherichia coli,” Journal of Bacteriology, vol. 177, no. 17, pp. 4851–4856, 1995.

S. Leimkühler and K. V. Rajagopalan, “A sulfurtransferase is required in the transfer of cysteine Sulfur in the in vitro synthesis of molybdopterin from precursor Z in Escherichia coli,” The Journal of Biological Chemistry, vol. 276, no. 25, pp. 22024–22031, 2001.

S. Leimkühler, M. M. Wuebbens, and K. V. Rajagopalan, “The history of the discovery of the molybdenum cofactor and novel aspects of its biosynthesis in bacteria,” Coordination Chemistry Reviews, vol. 255, no. 9-10, pp. 1129–1144, 2011.

J. A. Santamaria-Araujo, B. Fischer, T. Otte et al., “The tetrahydropyranopterin structure of the Sulfur- and metal-free molybdenum cofactor precursor,” The Journal of Biological Chemistry, vol. 279, no. 16, pp. 15994–15999, 2004.

B. M. Hover, A. Loksztejn, A. A. Ribeiro, and K. Yokoyama, “Identification of a cyclic nucleotide as a cryptic intermediate in molybdenum cofactor biosynthesis,” Journal of the American Chemical Society, vol. 135, pp. 7019–7032, 2013.

A. P. Mehta, S. H. Abdelwahed, and T. P. Begley, “Molybdopterin biosynthesis: trapping an unusual purine ribose adduct in the MoaA-catalyzed reaction,” Journal of the American Chemical Society, vol. 135, pp. 10883–10885, 2013.

M. M. Wuebbens, M. T. Liu, K. V. Rajagopalan, and H. Schindelin, “Insight into molybdenum cofactor deficiency provided by the crystal structure of the molybdenum cofactor biosynthesis protein MoaC,” Structure, vol. 8, no. 7, pp. 709–718, 2000.

P. Hänzelmann and H. Schindelin, “Crystal structure of the S-adenosylmethionine-dependent enzyme MoaA and its implications for molybdenum cofactor deficiency in humans,” Proceedings of the National Academy of Sciences of the United States of America, vol. 101, no. 35, pp. 12870–12875, 2004.

P. Hänzelmann, H. L. Hernández, C. Menzel et al., “Characterization of MOCS1A, an Oxygen-sensitive Iron-Sulfur protein involved in human molybdenum cofactor biosynthesis,” The Journal of Biological Chemistry, vol. 279, no. 33, pp. 34721–34732, 2004.

H. J. Sofia, G. Chen, B. G. Hetzler, J. F. Reyes-Spindola, and N. E. Miller, “Radical SAM, a novel protein superfamily linking unresolved steps in familiar biosynthetic pathways with radical mechanisms: functional characterization using new analysis and information visualization methods,” Nucleic Acids Research, vol. 29, no. 5, pp. 1097–1106, 2001.

N. S. Lees, P. Hänzelmann, H. L. Hernandez et al., “ENDOR spectroscopy shows that guanine N1 binds to [4Fe-4S] cluster II of the S-adenosylmethionine-dependent enzyme MoaA: mechanistic implications,” Journal of the American Chemical Society, vol. 131, no. 26, pp. 9184–9185, 2009.

P. Hänzelmann and H. Schindelin, “Binding of 5′-GTP to the C-terminal FeS cluster of the radical S-adenosylmethionine enzyme MoaA provides insights into its mechanism,” Proceedings of the National Academy of Sciences of the United States of America, vol. 103, no. 18, pp. 6829–6834, 2006.

A. P. Mehta, J. W. Hanes, S. H. Abdelwahed, D. G. Hilmey, P. Hänzelmann, and T. P. Begley, “Catalysis of a new ribose Carbon-insertion reaction by the molybdenum cofactor biosynthetic enzyme MoaA,” Biochemistry, vol. 52, pp. 1134–1136, 2013.

D. M. Pitterle and K. V. Rajagopalan, “The biosynthesis of molybdopterin in Escherichia coli. Purification and characterization of the converting factor,” The Journal of Biological Chemistry, vol. 268, no. 18, pp. 13499–13505, 1993.

D. M. Pitterle, J. L. Johnson, and K. V. Rajagopalan, “Molybdopterin formation by converting factor of E. coli chlA1,” The FASEB Journal, vol. 4, p. A1957, 1990.

D. M. Pitterle and K. V. Rajagopalan, “Two proteins encoded at the chlA locus constitute the converting factor of Escherichia coli chlA1,” Journal of Bacteriology, vol. 171, no. 6, pp. 3373–3378, 1989.

G. Gutzke, B. Fischer, R. R. Mendel, and G. Schwarz, “Thiocarboxylation of molybdopterin synthase provides evidence for the mechanism of dithiolene formation in metal-binding pterins,” The Journal of Biological Chemistry, vol. 276, no. 39, pp. 36268–36274, 2001.

M. J. Rudolph, M. M. Wuebbens, K. V. Rajagopalan, and H. Schindelin, “Crystal structure of molybdopterin synthase and its evolutionary relationship to ubiquitin activation,” Nature Structural Biology, vol. 8, no. 1, pp. 42–46, 2001.

M. M. Wuebbens and K. V. Rajagopalan, “Mechanistic and mutational studies of Escherichia coli molybdopterin synthase clarify the final step of molybdopterin biosynthesis,” The Journal of Biological Chemistry, vol. 278, no. 16, pp. 14523–14532, 2003.

J. N. Daniels, M. M. Wuebbens, K. V. Rajagopalan, and H. Schindelin, “Crystal structure of a molybdopterin synthase-precursor Z complex: insight into its Sulfur transfer mechanism and its role in molybdenum cofactor deficiency,” Biochemistry, vol. 47, no. 2, pp. 615–626, 2008, Erratum in: Biochemistry, vol. 47, no. 10, pp. 3315, 2008

J. Schmitz, M. M. Wuebbens, K. V. Rajagopalan, and S. Leimkühler, “Role of the C-terminal Gly-Gly motif of Escherichia coli MoaD, a molybdenum cofactor biosynthesis protein with a ubiquitin fold,” Biochemistry, vol. 46, no. 3, pp. 909–916, 2007.

K. V. Rajagopalan, “Biosynthesis and processing of the molybdenum cofactors,” Biochemical Society Transactions, vol. 25, no. 3, pp. 757–761, 1997.

M. W. Lake, M. M. Wuebbens, K. V. Rajagopalan, and H. Schindelin, “Mechanism of ubiquitin activation revealed by the structure of a bacterial MoeB-MoaD complex,” Nature, vol. 414, no. 6861, pp. 325–329, 2001.

Y. Tong, M. M. Wuebbens, K. V. Rajagopalan, and M. C. Fitzgerald, “Thermodynamic analysis of subunit interactions in Escherichia coli molybdopterin synthase,” Biochemistry, vol. 44, no. 7, pp. 2595–2601, 2005.

J. U. Dahl, C. Radon, M. Bühning, et al., “The Sulfur carrier protein TusA has a pleiotropic role in Escherichia coli that also affects molybdenum cofactor biosynthesis,” The Journal of Biological Chemistry, vol. 288, pp. 5426–5442, 2013.

J.-U. Dahl, A. Urban, A. Bolte et al., “The identification of a novel protein involved in molybdenum cofactor biosynthesis in Escherichia coli,” The Journal of Biological Chemistry, vol. 286, no. 41, pp. 35801–35812, 2011.

M. T. W. Liu, M. M. Wuebbens, K. V. Rajagopalan, and H. Schindelin, “Crystal structure of the gephyrin-related molybdenum cofactor biosynthesis protein MogA from Escherichia coli,” The Journal of Biological Chemistry, vol. 275, no. 3, pp. 1814–1822, 2000.

S. Xiang, J. Nichols, K. V. Rajagopalan, and H. Schindelin, “The crystal structure of Escherichia coli MoeA and its relationship to the multifunctional protein gephyrin,” Structure, vol. 9, no. 4, pp. 299–310, 2001.

J. D. Schrag, W. Huang, J. Sivaraman et al., “The crystal structure of Escherichia coli MoeA, a protein from the molybdopterin synthesis pathway,” Journal of Molecular Biology, vol. 310, no. 2, pp. 419–431, 2001.

J. D. Nichols and K. V. Rajagopalan, “In vitro molybdenum ligation to molybdopterin using purified components,” The Journal of Biological Chemistry, vol. 280, no. 9, pp. 7817–7822, 2005.

A. Llamas, R. R. Mendel, and G. Schwarz, “Synthesis of adenylated molybdopterin: an essential step for molybdenum insertion,” The Journal of Biological Chemistry, vol. 279, no. 53, pp. 55241–55246, 2004.

A. Llamas, T. Otte, G. Multhaup, R. R. Mendel, and G. Schwarz, “The mechanism of nucleotide-assisted molybdenum insertion into molybdopterin: a novel route toward metal cofactor assembly,” The Journal of Biological Chemistry, vol. 281, no. 27, pp. 18343–18350, 2006.

S. Leimkühler and K. V. Rajagopalan, “In vitro incorporation of nascent molybdenum cofactor into human sulfite oxidase,” The Journal of Biological Chemistry, vol. 276, no. 3, pp. 1837–1844, 2001.

M. Neumann and S. Leimkühler, “Heavy metal ions inhibit molybdoenzyme activity by binding to the dithiolene moiety of molybdopterin in Escherichia coli,” The FEBS Journal, vol. 275, no. 22, pp. 5678–5689, 2008.

L. Loschi, S. J. Brokx, T. L. Hills et al., “Structural and biochemical identification of a novel bacterial oxidoreductase,” The Journal of Biological Chemistry, vol. 279, no. 48, pp. 50391–50400, 2004.

C. A. Temple and K. V. Rajagopalan, “Mechanism of assembly of the bis(molybdopterin guanine dinucleotide)molybdenum cofactor in Rhodobacter sphaeroides dimethyl sulfoxide reductase,” The Journal of Biological Chemistry, vol. 275, no. 51, pp. 40202–40210, 2000.

M. W. Lake, C. A. Temple, K. V. Rajagopalan, and H. Schindelin, “The crystal structure of the Escherichia coli MobA protein provides insight into molybdopterin guanine dinucleotide biosynthesis,” The Journal of Biological Chemistry, vol. 275, no. 51, pp. 40211–40217, 2000.

C. E. M. Stevenson, F. Sargent, G. Buchanan, T. Palmer, and D. M. Lawson, “Crystal structure of the molybdenum cofactor biosynthesis protein MobA from Escherichia coli at near-atomic resolution,” Structure, vol. 8, no. 11, pp. 1115–1125, 2000.

J. L. Johnson, L. W. Indermaur, and K. V. Rajagopalan, “Molybdenum cofactor biosynthesis in Escherichia coli: requirement of the chlB gene product for the formation of molybdopterin guanine dinucleotide,” The Journal of Biological Chemistry, vol. 266, no. 19, pp. 12140–12145, 1991.

K. McLuskey, J. A. Harrison, A. W. Schüttelkopf, D. H. Boxer, and W. N. Hunter, “Insight into the role of Escherichia coli MobB in molybdenum cofactor biosynthesis based on the high resolution crystal structure,” The Journal of Biological Chemistry, vol. 278, no. 26, pp. 23706–23713, 2003.

D. J. Eaves, T. Palmer, and D. H. Boxer, “The product of the molybdenum cofactor gene mobB of Escherichia coli is a GTP-binding protein,” European Journal of Biochemistry, vol. 246, no. 3, pp. 690–697, 1997.

S. Leimkühler and W. Klipp, “The molybdenum cofactor biosynthesis protein MobA from Rhodobacter capsulatus is required for the activity of molybdenum enzymes containing MGD, but not for xanthine dehydrogenase harboring the MPT cofactor,” FEMS Microbiology Letters, vol. 174, no. 2, pp. 239–246, 1999.

O. Genest, V. Méjean, and C. Iobbi-Nivol, “Multiple roles of TorD-like chaperones in the biogenesis of molybdoenzymes,” FEMS Microbiology Letters, vol. 297, no. 1, pp. 1–9, 2009.

C. Kisker, H. Schindelin, and D. C. Rees, “Molybdenum-cofactor-containing enzymes: structure and mechanism,” Annual Review of Biochemistry, vol. 66, pp. 233–267, 1997.

F. Blasco, J.-P. Dos Santos, A. Magalon et al., “NarJ is a specific chaperone required for molybdenum cofactor assembly in nitrate reductase A of Escherichia coli,” Molecular Microbiology, vol. 28, no. 3, pp. 435–447, 1998.

F. Blasco, J. Pommier, V. Augier, M. Chippaux, and G. Giordano, “Involvement of the narJ or narW gene product in the formation of active nitrate reductase in Escherichia coli,” Molecular Microbiology, vol. 6, no. 2, pp. 221–230, 1992.

N. Ray, J. Oates, R. J. Turner, and C. Robinson, “DmsD is required for the biogenesis of DMSO reductase in Escherichia coli but not for the interaction of the DmsA signal peptide with the Tat apparatus,” FEBS Letters, vol. 534, no. 1-3, pp. 156–160, 2003.

D. Guymer, J. Maillard, and F. Sargent, “A genetic analysis of in vivo selenate reduction by Salmonella enterica serovar Typhimurium LT2 and Escherichia coli K12,” Archives of Microbiology, vol. 191, no. 6, pp. 519–528, 2009.

R. Thomé, A. Gust, R. Toci et al., “A sulfurtransferase is essential for activity of formate dehydrogenases in Escherichia coli,” The Journal of Biological Chemistry, vol. 287, no. 7, pp. 4671–4678, 2012.

O. Genest, M. Ilbert, V. Méjean, and C. Iobbi-Nivol, “TorD, an essential chaperone for TorA molybdoenzyme maturation at high temperature,” The Journal of Biological Chemistry, vol. 280, no. 16, pp. 15644–15648, 2005.

O. Genest, F. Seduk, L. Théraulaz, V. Méjean, and C. Iobbi-Nivol, “Chaperone protection of immature molybdoenzyme during molybdenum cofactor limitation,” FEMS Microbiology Letters, vol. 265, no. 1, pp. 51–55, 2006.

O. Genest, M. Neumann, F. Seduk et al., “Dedicated metallochaperone connects apoenzyme and molybdenum cofactor biosynthesis components,” The Journal of Biological Chemistry, vol. 283, no. 31, pp. 21433–21440, 2008.

N. Böhmer, T. Hartmann, and S. Leimkühler, “The chaperone FdsC for Rhodobacter capsulatus formate dehydrogenase binds the bis-molybdopterin guanine dinucleotide cofactor,” FEBS Letters, vol. 588, no. 4, pp. 531–537, 2014.

T. Hartmann and S. Leimkuhler, “The Oxygen-tolerant and NAD(+) -dependent formate dehydrogenase from Rhodobacter capsulatus is able to catalyze the reduction of CO₂ to formate,” The FEBS Journal, vol. 280, pp. 6083–6096, 2013.

C. Coelho, P. J. González, J. G. Moura, I. Moura, J. Trincão, and M. João Romão, “The crystal structure of cupriavidus necator nitrate reductase in oxidized and partially reduced states,” Journal of Molecular Biology, vol. 408, no. 5, pp. 932–948, 2011.

S. Najmudin, P. J. González, J. Trincão et al., “Periplasmic nitrate reductase revisited: a Sulfur atom completes the sixth coordination of the catalytic molybdenum,” Journal of Biological Inorganic Chemistry, vol. 13, no. 5, pp. 737–753, 2008.

H. Xi, B. L. Schneider, and L. Reitzer, “Purine catabolism in Escherichia coli and function of xanthine dehydrogenase in purine salvage,” Journal of Bacteriology, vol. 182, no. 19, pp. 5332–5341, 2000.

S. G. Kozmin and R. M. Schaaper, “Molybdenum cofactor-dependent resistance to N-hydroxylated base analogs in Escherichia coli is independent of MobA function,” Mutation Research—Fundamental and Molecular Mechanisms of Mutagenesis, vol. 619, no. 1-2, pp. 9–15, 2007.

M. Neumann, G. Mittelstädt, C. Iobbi-Nivol et al., “A periplasmic aldehyde oxidoreductase represents the first molybdopterin cytosine dinucleotide cofactor containing molybdo-flavoenzyme from Escherichia coli,” The FEBS Journal, vol. 276, no. 10, pp. 2762–2774, 2009.

M. Neumann, F. Seduk, C. Iobbi-Nivol, and S. Leimkühler, “Molybdopterin dinucleotide biosynthesis in Escherichia coli: identification of amino acid residues of molybdopterin dinucleotide transferases that determine specificity for binding of guanine or cytosine nucleotides,” The Journal of Biological Chemistry, vol. 286, no. 2, pp. 1400–1408, 2011.

A. Guse, C. E. M. Stevenson, J. Kuper et al., “Biochemical and structural analysis of the molybdenum cofactor biosynthesis protein MobA,” The Journal of Biological Chemistry, vol. 278, no. 28, pp. 25302–25307, 2003.

S. Leimkühler, S. Angermüller, G. Schwarz, R. R. Mendel, and W. Klipp, “Activity of the molybdopterin-containing xanthine dehydrogenase of Rhodobacter capsulatus can be restored by high molybdenum concentrations in a moeA mutant defective in molybdenum cofactor biosynthesis,” Journal of Bacteriology, vol. 181, no. 19, pp. 5930–5939, 1999.

M. Neumann, W. Stöcklein, and S. Leimkühler, “Transfer of the molybdenum cofactor synthesized by Rhodobacter capsulatus MoeA to XdhC and MobA,” The Journal of Biological Chemistry, vol. 282, no. 39, pp. 28493–28500, 2007.

M. Neumann, W. Stöcklein, A. Walburger, A. Magalon, and S. Leimkühler, “Identification of a Rhodobacter capsulatus L-cysteine desulfurase that sulfurates the molybdenum cofactor when bound to XdhC and before its insertion into xanthine dehydrogenase,” Biochemistry, vol. 46, no. 33, pp. 9586–9595, 2007.

M. Neumann, M. Schulte, N. Jünemann, W. Stöcklein, and S. Leimkühler, “Rhodobacter capsulatus XdhC is involved in molybdenum cofactor binding and insertion into xanthine dehydrogenase,” The Journal of Biological Chemistry, vol. 281, no. 23, pp. 15701–15708, 2006.

S. Leimkühler and W. Klipp, “Role of XDHC in molybdenum cofactor insertion into xanthine dehydrogenase of Rhodobacter capsulatus,” Journal of Bacteriology, vol. 181, no. 9, pp. 2745–2751, 1999.

A. R. Otrelo-Cardoso, V. Schwuchow, D. Rodrigues, et al., “Biochemical, stabilization and crystallization studies on a molecular chaperone (PaoD) involved in the maturation of molybdoenzymes,” PLoS ONE, vol. 9, no. 1, Article ID e87295, 2014.

H. Beinert, “A tribute to Sulfur,” European Journal of Biochemistry, vol. 267, no. 18, pp. 5657–5664, 2000.

L. Zheng, V. L. Cash, D. H. Flint, and D. R. Dean, “Assembly of Iron-Sulfur clusters. Identification of an iscSUA-hscBA-fdx gene cluster from Azotobacter vinelandii,” The Journal of Biological Chemistry, vol. 273, no. 21, pp. 13264–13272, 1998.

L. Zheng, R. H. White, V. L. Cash, and D. R. Dean, “Mechanism for the desulfurization of L-cysteine catalyzed by the NIFs gene product,” Biochemistry, vol. 33, no. 15, pp. 4714–4720, 1994.

L. Zheng, R. H. White, V. L. Cash, R. F. Jack, and D. R. Dean, “Cysteine desulfurase activity indicates a role for NIFS in metallocluster biosynthesis,” Proceedings of the National Academy of Sciences of the United States of America, vol. 90, no. 7, pp. 2754–2758, 1993.

R. Hidese, H. Mihara, and N. Esaki, “Bacterial cysteine desulfurases: versatile key players in biosynthetic pathways of Sulfur-containing biofactors,” Applied Microbiology and Biotechnology, vol. 91, no. 1, pp. 47–61, 2011.

E. G. Mueller, “Trafficking in persulfides: delivering Sulfur in biosynthetic pathways,” Nature Chemical Biology, vol. 2, no. 4, pp. 185–194, 2006.

H. D. Urbina, J. J. Silberg, K. G. Hoff, and L. E. Vickery, “Transfer of Sulfur from IscS to IscU during Fe/S cluster assembly,” The Journal of Biological Chemistry, vol. 276, no. 48, pp. 44521–44526, 2001.

R. Shi, A. Proteau, M. Villarroya et al., “Structural basis for Fe-S cluster assembly and tRNA thiolation mediated by IscS protein-protein interactions,” PLoS Biology, vol. 8, no. 4, Article ID e1000354, 2010.

C. Iannuzzi, S. Adinolfi, B. D. Howes et al., “The role of cyay in Iron Sulfur cluster assembly on the E. coli iscu scaffold protein,” PLoS ONE, vol. 6, no. 7, Article ID e21992, 2011.

S. Adinolfi, C. Iannuzzi, F. Prischi et al., “Bacterial frataxin CyaY is the gatekeeper of Iron-Sulfur cluster formation catalyzed by IscS,” Nature Structural & Molecular Biology, vol. 16, no. 4, pp. 390–396, 2009.

E. G. Mueller, C. J. Buck, P. M. Palenchar, L. E. Barnhart, and J. L. Paulson, “Identification of a gene involved in the generation of 4-thiouridine in tRNA,” Nucleic Acids Research, vol. 26, no. 11, pp. 2606–2610, 1998.

Y. Ikeuchi, N. Shigi, J.-I. Kato, A. Nishimura, and T. Suzuki, “Mechanistic insights into Sulfur relay by multiple Sulfur mediators involved in thiouridine biosynthesis at tRNA wobble positions,” Molecular Cell, vol. 21, no. 1, pp. 97–108, 2006.

T. P. Begley, D. M. Downs, S. E. Ealick et al., “Thiamin biosynthesis in prokaryotes,” Archives of Microbiology, vol. 171, no. 5, pp. 293–300, 1999.

D. Bordo and P. Bork, “The rhodanese/Cdc25 phosphatase superfamily. Sequence-structure-function relations,” EMBO Reports, vol. 3, no. 8, pp. 741–746, 2002.

R. Malkin and J. C. Rabinowitz, “The reconstitution of clostridial ferredoxin,” Biochemical and Biophysical Research Communications, vol. 23, no. 6, pp. 822–827, 1966.

B. Roche, L. Aussel, B. Ezraty, P. Mandin, B. Py, and F. Barras, “Iron/Sulfur proteins biogenesis in prokaryotes: formation, regulation and diversity,” Biochimica et Biophysica Acta, vol. 1827, pp. 455–469, 2013.

C. J. Schwartz, J. L. Giel, T. Patschkowski et al., “IscR, an Fe-S cluster-containing transcription factor, represses expression of Escherichia coli genes encoding Fe-S cluster assembly proteins,” Proceedings of the National Academy of Sciences of the United States of America, vol. 98, no. 26, pp. 14895–14900, 2001.

C. J. Schwartz, O. Djaman, J. A. Imlay, and P. J. Kiley, “The cysteine desulfurase, IscS, has a major role in in vivo Fe-S cluster formation in Escherichia coli,” Proceedings of the National Academy of Sciences of the United States of America, vol. 97, no. 16, pp. 9009–9014, 2000.

E. N. Marinoni, J. S. de Oliveira, Y. Nicolet, et al., “(IscS-IscU)₂ complex structures provide insights into Fe₂S₂ biogenesis and transfer,” Angewandte Chemie, vol. 51, no. 22, pp. 5439–5442, 2012.

K. Chandramouli, M.-C. Unciuleac, S. Naik, D. R. Dean, H. H. Boi, and M. K. Johnson, “Formation and properties of [4Fe-4S] clusters on the IscU scaffold protein,” Biochemistry, vol. 46, no. 23, pp. 6804–6811, 2007.

S. Ollagnier-De-Choudens, T. Mattioli, Y. Takahashi, and M. Fontecave, “Iron-Sulfur cluster assembly. Characterization of IscA and evidence for a specific and functional complex with ferredoxin,” The Journal of Biological Chemistry, vol. 276, no. 25, pp. 22604–22607, 2001.

J. H. Kim, M. Tonelli, R. O. Frederick, D. C. Chow, and J. L. Markley, “Specialized Hsp70 chaperone (HscA) binds preferentially to the disordered form, whereas J-protein (HscB) binds preferentially to the structured form of the Iron-Sulfur cluster scaffold protein (IscU),” The Journal of Biological Chemistry, vol. 287, pp. 31406–31413, 2012.

F. Prischi, P. V. Konarev, C. Iannuzzi et al., “Structural bases for the interaction of frataxin with the central components of Iron-Sulphur cluster assembly,” Nature Communications, vol. 1, article 95, 2010.

J. Bridwell-Rabb, C. Iannuzzi, A. Pastore, and D. P. Barondeau, “Effector role reversal during evolution: the case of frataxin in Fe-S cluster biosynthesis,” Biochemistry, vol. 51, no. 12, pp. 2506–2514, 2012.

F. Bou-Abdallah, S. Adinolfi, A. Pastore, T. M. Laue, and N. Dennis Chasteen, “Iron binding and oxidation kinetics in frataxin CyaY of Escherichia coli,” Journal of Molecular Biology, vol. 341, no. 2, pp. 605–615, 2004.

L. Loiseau, S. Ollagnier-de-Choudens, L. Nachin, M. Fontecave, and F. Barras, “Biogenesis of Fe-S cluster by the bacterial suf system. SufS and SufE form a new type of cysteine desulfurase,” The Journal of Biological Chemistry, vol. 278, no. 40, pp. 38352–38359, 2003.

F. W. Outten, O. Djaman, and G. Storz, “A suf operon requirement for Fe-S cluster assembly during Iron starvation in Escherichia coli,” Molecular Microbiology, vol. 52, no. 3, pp. 861–872, 2004.

R. Kambampati and C. T. Lauhon, “Evidence for the transfer of sulfane Sulfur from IscS to ThiI during the in vitro biosynthesis of 4-thiouridine in Escherichia coli tRNA,” The Journal of Biological Chemistry, vol. 275, no. 15, pp. 10727–10730, 2000.

P. M. Palenchar, C. J. Buck, H. Cheng, T. J. Larson, and E. G. Mueller, “Evidence that ThiI, an enzyme shared between thiamin and 4-thiouridine biosynthesis, may be a sulfurtransferase that proceeds through a persulfide intermediate,” The Journal of Biological Chemistry, vol. 275, no. 12, pp. 8283–8286, 2000.

R. Kambampati and C. T. Lauhon, “Mnma and IscS are required for in vitro 2-thiouridine biosynthesis in Escherichia coli,” Biochemistry, vol. 42, no. 4, pp. 1109–1117, 2003.

T. Numata, Y. Ikeuchi, S. Fukai, T. Suzuki, and O. Nureki, “Snapshots of tRNA Sulphuration via an adenylated intermediate,” Nature, vol. 442, no. 7101, pp. 419–424, 2006.

T. Numata, Y. Ikeuchi, S. Fukai et al., “Crystallization and preliminary X-ray analysis of the tRNA thiolation enzyme MnmA from Escherichia coli complexed with tRNAGlu,” Acta Crystallographica F: Structural Biology and Crystallization Communications, vol. 62, no. 4, pp. 368–371, 2006.

I. Moukadiri, M. J. Garzon, G. R. Bjork, and M. E. Armengod, “The output of the tRNA modification pathways controlled by the Escherichia coli MnmEG and MnmC enzymes depends on the growth conditions and the tRNA species,” Nucleic Acids Research, vol. 42, no. 4, pp. 2602–2623, 2013.

N. D. Maynard, E. W. Birch, J. C. Sanghvi, L. Chen, M. V. Gutschow, and M. W. Covert, “A forward-genetic screen and dynamic analysis of lambda phage host-dependencies reveals an extensive interaction network and a new anti-viral strategy,” PLoS Genetics, vol. 6, no. 7, Article ID e1001017, 2010.

N. D. Maynard, D. N. MacKlin, K. Kirkegaard, and M. W. Covert, “Competing pathways control host resistance to virus via tRNA modification and programmed ribosomal frameshifting,” Molecular Systems Biology, vol. 8, article 567, 2012.

J. L. Giel, A. D. Nesbit, E. L. Mettert, A. S. Fleischhacker, B. T. Wanta, and P. J. Kiley, “Regulation of Iron-Sulphur cluster homeostasis through transcriptional control of the Isc pathway by [2Fe-2S]-IscR in Escherichia coli,” Molecular Microbiology, vol. 87, pp. 478–492, 2013.

K. S. Myers, H. Yan, I. M. Ong, et al., “Genome-scale analysis of Escherichia coli FNR reveals complex features of transcription factor binding,” PLoS Genetics, vol. 9, Article ID e1003565, 2013.

Y. Ishii, H. Yamada, T. Yamashino et al., “Deletion of the yhhP gene results in filamentous cell morphology in Escherichia coli,” Bioscience, Biotechnology and Biochemistry, vol. 64, no. 4, pp. 799–807, 2000.