Iron-Dependent Regulation of Molybdenum CofactorBiosynthesis Genes in Escherichia coli

Arkadiusz Zupok,a Michal Gorka,b Beata Siemiatkowska,b Aleksandra Skirycz,b Silke Leimkühlera

aInstitute of Biochemistry and Biology, Molecular Enzymology, University of Potsdam, Potsdam-Golm, GermanybMax Planck Institute of Molecular Plant Physiology, Potsdam-Golm, Germany

ABSTRACT Molybdenum cofactor (Moco) biosynthesis is a complex process that in-volves the coordinated function of several proteins. In recent years it has becomeobvious that the availability of iron plays an important role in the biosynthesis ofMoco. First, the MoaA protein binds two [4Fe-4S] clusters per monomer. Second, theexpression of the moaABCDE and moeAB operons is regulated by FNR, which sensesthe availability of oxygen via a functional [4Fe-4S] cluster. Finally, the conversion ofcyclic pyranopterin monophosphate to molybdopterin requires the availability of theL-cysteine desulfurase IscS, which is a shared protein with a main role in the assem-bly of Fe-S clusters. In this report, we investigated the transcriptional regulation ofthe moaABCDE operon by focusing on its dependence on cellular iron availability.While the abundance of selected molybdoenzymes is largely decreased under iron-limiting conditions, our data show that the regulation of the moaABCDE operon atthe level of transcription is only marginally influenced by the availability of iron.Nevertheless, intracellular levels of Moco were decreased under iron-limiting condi-tions, likely based on an inactive MoaA protein in addition to lower levels of theL-cysteine desulfurase IscS, which simultaneously reduces the sulfur availability forMoco production.

IMPORTANCE FNR is a very important transcriptional factor that represents the mas-ter switch for the expression of target genes in response to anaerobiosis. Among theFNR-regulated operons in Escherichia coli is the moaABCDE operon, involved in Mocobiosynthesis. Molybdoenzymes have essential roles in eukaryotic and prokaryotic or-ganisms. In bacteria, molybdoenzymes are crucial for anaerobic respiration using al-ternative electron acceptors. This work investigates the connection of iron availabil-ity to the biosynthesis of Moco and the production of active molybdoenzymes.

KEYWORDS Escherichia coli, FNR, iron regulation, iron-sulfur cluster, anaerobicrespiration, molybdenum cofactor

The molybdenum cofactor (Moco) is an important metallocofactor displaying versa-tile roles as the catalytic center of enzymes (1–6). It is an essential cofactor for the

activity of at least 15 enzymes in Escherichia coli. In Moco, the molybdenum atom iscoordinated to a dithiolene group on the 6-alkyl side chain of a pterin called molyb-dopterin (MPT) (7). In E. coli, Moco biosynthesis is divided into four general steps (7). Inthe first step, 5=GTP is converted to cyclic pyranopterin monophosphate (cPMP) by theproteins MoaA (GTP 3=,8-cyclase) and MoaC (cPMP synthase) (8–11). MoaA belongs tothe superfamily of radical S-adenosylmethionine (SAM)-dependent enzymes. Reductivecleavage of SAM by a conserved [4Fe-4S] cluster leads to the production of protein orsubstrate radicals by proteins of this enzyme family (12). MoaA contains two [4Fe-4S]clusters per monomer, which are oxygen sensitive and are essential for protein func-tionality (13). In the second step of Moco biosynthesis, MPT synthase catalyzes theinsertion of two sulfur atoms to the C1= and C2= positions of cPMP. MPT synthase is a

Citation Zupok A, Gorka M, Siemiatkowska B,Skirycz A, Leimkühler S. 2019. Iron-dependentregulation of molybdenum cofactorbiosynthesis genes in Escherichia coli. JBacteriol 201:e00382-19. https://doi.org/10.1128/JB.00382-19.

Editor Anke Becker, Philipps-UniversitätMarburg

Copyright © 2019 American Society forMicrobiology. All Rights Reserved.

Address correspondence to Silke Leimkühler,sleim@uni-potsdam.de.

Received 4 June 2019Accepted 15 June 2019

Accepted manuscript posted online 24June 2019Published

RESEARCH ARTICLE

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heterodimer in which each monomer consists of two MoaD subunits and two MoaEsubunits (14, 15). The sulfur is directly inserted from the C-terminal thiocarboxylategroup of each MoaD protein into cPMP (16). Initial mobilization of sulfur for thisreaction is provided by the L-cysteine desulfurase IscS, which transfers the sulfur in theform of a protein-bound persulfide to MoaD, with the additional involvement of theTusA protein (17, 18). After sulfur insertion into cPMP, the MoeB protein regeneratesthe sulfur at the C-terminal glycine of MoaD in an ATP-dependent reaction (19, 20).Further, the MogA and MoeA proteins (MPT adenylyltransferase and MPT molyb-dotransferase, respectively) catalyze the insertion of molybdenum into MPT, formingthe Mo-MPT cofactor (10, 21–23). This form of the cofactor can either be directlyinserted into molybdoenzymes or can undergo further modification to form theMPT-cytosine dinucleotide (MCD) and bis-MPT guanine dinucleotide (bis-MGD) deriv-atives of the molybdenum cofactors (24–26). Formation of MCD is catalyzed by theMocA (Moco cytidylyltransferase) protein (24), while bis-MGD formation is catalyzed bythe MobA (Moco guanylyltransferase) protein in a two-step process (25, 27, 28). Afterthe formation of MCD or bis-MGD, these cofactors can further be modified by theaddition of a sulfido ligand to the molybdenum atom (29, 30).

Overall, at least 13 genes are involved in the biosynthesis of Moco in E. coli (3, 31).They are combined in 6 different operons organized as moaABCDE, mobA, mocA,modABC, moeAB, and mogA (3). With the exception of the mod locus of genes for ahigh-affinity molybdate transport system (32), all of these genes are involved in thebiosynthesis of Moco.

Genetic analysis of the regulation of Moco biosynthesis revealed that mainly themoaABCDE and moeAB operons are the target for transcriptional and translationalregulation (33, 34). In E. coli, transcription of the moaABCDE operon is mainly regulatedby ModE and FNR (the fumarate and nitrate reduction regulatory protein) (33).Molybdate-bound ModE acts as a positive regulator and binds to the moa promoterregion, thereby enhancing transcription of the operon (33). FNR is a transcriptionalregulator that is essential for expressing anaerobic respiratory processes and has beenshown previously to activate the transcription of the moa locus in E. coli (33, 35). Ingeneral, FNR senses the oxygen concentration directly via the disassembly and reassemblyof its [4Fe-4S] clusters (36). ModE and FNR regulation ensure that the moaABCDE transcriptis created only under cellular conditions when needing (anaerobic growth conditionssensed by FNR) and enabling (sufficient levels of molybdate sensed by ModE) Mocoproduction (33). Two transcriptional start sites have been mapped in the moaA pro-moter region (33). The S2 start site downstream of the FNR binding site (�87) is usedunder anaerobic conditions, while the S1 site (�1) is used under both aerobic andanaerobic conditions. It has been suggested that ModE and Fnr mediate their regula-tory effects specifically and distinctly at the proximal and distal promoter sites, respec-tively (33).

Further, the moeAB operon was also shown to be regulated by FNR, ArcA (oxygen-responsive DNA-binding response regulator), and NarL (nitrate-responsive regulator)(34). Surprisingly, FNR represses the nitrate-dependent transcription of the moe operon,but it is expected that under anaerobic conditions, an intermediate level of transcrip-tion of the moeAB operon is ensured by the antagonistic effects of FNR, ArcA-P, andNarL (34).

For the moa locus an additional level of regulation was determined by the identi-fication of a Moco riboswitch (37). The Moco riboswitch upstream of moaA has beenrevealed to control gene expression in response to an excess of Moco production. Itwas shown that translation of the moa locus is prevented when the Moco RNA structureand Moco are present; however, a direct binding of Moco to this RNA motif has notbeen demonstrated to date, mainly due to the lability of isolated cofactor (37). Inaddition, a regulation of the moaABCDE operon by the carbon storage regulator CsrAhas been described (38). CsrA is a global regulator that represses stationary-phasemetabolism and activates central carbon metabolism (39). Two binding sites of CsrAwere identified within the moaA promoter region (38). For Moco biosynthesis, CrsA was

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shown to activate the transcription, and it has been suggested that CsrA enhancesMoco biosynthesis under conditions of high demand. Thus, in total the moaABCDEoperon is regulated by at least four different factors, FNR, ModE, CsrA, and Moco (viathe riboswitch) (33, 37, 38).

However, apart from the dependence on functional [4Fe-4S] clusters being presentin the FNR protein, a regulation of Moco biosynthesis by cellular iron levels has notbeen investigated in the past. This is surprising, since it has become evident that Mocobiosynthesis depends on Fe-S clusters or components of the Fe-S cluster assemblymachinery at several levels (40). In the first step of Moco biosynthesis, the MoaA proteinrequires two [4Fe-4S] clusters for activity (12, 41). Further, most molybdoenzymes in E.coli harbor numerous Fe-S clusters that are involved in intramolecular electron transferreactions, which are essential for the activity of the enzymes (4, 40). In E. coli two majorsystems for the assembly of Fe-S clusters have been identified, namely, the iron sulfurcluster (ISC) system and the sulfur mobilization (SUF) system (42, 43). The centralcomponents of these systems are the two L-cysteine desulfurases, IscS and SufS, thatmobilize sulfur from L-cysteine to form L-alanine and a protein-bound persulfideintermediate (44). IscS was shown to act as housekeeping L-cysteine desulfurase foriron-sulfur (Fe-S) cluster assembly (45–47). Further, the IscS protein has been revealedto be the primary sulfur donor for numerous other sulfur-containing molecules withimportant biological functions, among which is Moco (48). In contrast, the SUF pathwayencoded by the sufABCDSE operon was identified as an alternative Fe-S cluster assem-bly route under iron-limiting conditions (49). The expression of the ISC and SUFmachineries is regulated by IscR, among other factors (50–52). IscR is a [2Fe-2S]transcriptional regulator encoded by the first gene of the iscRSUA-hscBA-fdx-iscXoperon. IscR, in its [2Fe-2S] cluster-bound holoform, represses its own expression aswell as that of the isc operon. Under iron limitation and oxidative stress, IscR is presentin its apo form and activates expression of the suf operon (50, 51). In this report, weinvestigated the transcriptional regulation of the moaABCDE operon depending oncellular iron availability.

RESULTSThe iron-dependent expression of moaA::lacZ fusions. To investigate the influ-

ence of iron on the expression of the moaABCDE operon, three moaA::lacZ fusionswere constructed and transformed into different E. coli mutant strains (Fig. 1A). ThemoaA-L::lacZ fusion includes a 477-bp fragment upstream of the moaA ATG start codoncontaining the reported FNR, ModE, CrsA, and Moco binding sites (Fig. 1A and B) inaddition to a 255-bp-long fragment within the moaA coding region. The moaA-S(�FNR)::lacZ fusion contains the same 477-bp region upstream of the ATG start codonbut lacks the coding region within moaA and directly fuses the lacZ gene to the ATGstart codon of moaA. The moaA-S(�FNR)::lacZ fusion encompasses a 220-bp regionupstream of the moaA ATG start codon that truncates the FNR binding site and directlyfuses the moaA ATG start codon to the lacZ coding region.

To investigate differences in �-galactosidase activities of the three lacZ fusions, theplasmids were transformed into E. coli strain BW25113 (referred to as the wild-type [WT]strain) and grown under anaerobic conditions in LB medium supplemented with KNO3

or with KNO3 and trimethylamine N-oxide (TMAO) (Fig. 1C and D) as alternative electronacceptors for comparison. KNO3 was added to the medium, since the moeAB genes forMoco biosynthesis in addition to the operons encoding several molybdoenzymes (e.g.,fdhF, narGHJI, napFDAGHBC, ynfEFGH, ydhYVWXUTdmsD, fdnGHI, and dmsABC) wereshown to be regulated by NarL and NarP depending on nitrate availability (34, 53–55).To ensure optimal growth of the cultures, TMAO was also added to the medium as aniron-independent electron acceptor for respiration, since nitrate reductase requiresseveral Fe-S clusters for activity (56, 57), while TMAO reductase is expressed indepen-dent of FNR and does not contain any Fe-S clusters (58, 59).

The results in Fig. 1 show that the activity of the moaA-S(�FNR)::lacZ fusion was 2/3decreased compared to that of the lacZ fusions moaA-S(�FNR) and moaA-L. No

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difference in �-galactosidase activity was obtained in LB medium containing KNO3 (Fig.1C) or containing KNO3 and TMAO (Fig. 1D). moaA-S(�FNR) lacks the FNR binding site,confirming that FNR acts as a positive regulator of the moaA operon (33). To identifytargets for putative translational regulators (ncRNAs) within the moaABCDE codonregion, the activities of moaA-S(�FNR) and moaA-L were compared. No differences in�-galactosidase activities of both lacZ fusions were obtained (Fig. 1C and D), showingthat there are no regulatory ncRNA binding sites within the moaA coding region.

Further, we analyzed the influence of iron on the activities of the three lacZ fusions.BW25113 cells with the three moaA::lacZ fusions were grown under anaerobic condi-

FIG 1 Construction of moaA::lacZ fusions and influence of the availability of iron on moaA::lacZ �-galactosidaseactivities. (A) The moaA promoter region and construction of moaA::lacZ fusions. moaA-L contains the FNR bindingsite and a fragment of the moaA coding region; moaA-S(�FNR) contains the FNR binding site, and moaA-S(�FNR)does not contain the FNR binding site. Both fusions are directly fused to the moaA ATG start codon. Boxes withinthe promoter region show schematically the binding sites of regulators: black, reported FNR binding site; gray,Moco riboswitch region; blue, ModE binding site; green, CsrA binding site; red, CueR binding site. Binding sites arenot drawn to the scale. S2 and S1, transcription start sites. (B) DNA sequence of the moaA promoter region. Bindingsites of regulatory elements within the moaABCDE promoter region are boxed. Boldface, Moco riboswitch region;black box, FNR binding site; blue, ModE binding site; green, CsrA binding site; red, CueR binding site; red bases Gand A, S2 and S1 transcription start sites, respectively; green ATG, ATG start codon of moaA. (C and D)�-Galactosidase activities in Miller units (M.U.) of the three moaA::lacZ fusions in BW25113 grown under anaerobicconditions with KNO3 or KNO3 and TMAO. (E to G) �-Galactosidase activities in Miller units of the three moaA::lacZfusions in BW25113 (BW) and the Δfnr and ΔiscS mutant lines grown under anaerobic conditions with KNO3. Whitebars, 0 �M 2,2-DIP; gray bars, 150 �M 2,2-DIP. (H to J) �-Galactosidase activities of the three moaA::lacZ fusions inBW25113 and the Δfnr and ΔiscS mutant lines grown under anaerobic conditions with KNO3 and TMAO. White bars,0 �M 2,2-DIP; gray bars, 150 �M 2,2-DIP. All of the values in graphs E to J were normalized to the level of BW25113grown without 2,2-DIP and are shown as percentages of the WT level. Standard deviations were calculated fromthree biological replicates.

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tions with KNO3 (Fig. 1E to G) or KNO3 and TMAO (Fig. 1H to J) with and withoutdipyridyl (2,2-DIP), used as an iron chelator, until cultures reached mid-log phase. Toassay for strain-dependent effects on expression, we additionally expressed the threelacZ fusions in the E. coli MG1655 wild-type strain background (see Fig. S1 in thesupplemental material). The results show that no differences in �-galactosidase activ-ities were obtained in BW25113 and MG1655 strains. Toxicity of 2,2-DIP was examinedby the addition of 500 �M FeCl3 to the medium together with the iron chelator. Theactivity of the lacZ fusions in BW25113 grown in medium containing 150 �M 2,2-DIPtogether with 500 �M FeCl3 was overall comparable to that of cells grown in mediumdevoid of 2,2-DIP (data not shown). To confirm that addition of 150 �M 2,2-DIP leadsto cellular iron limitation, we investigated the abundance of the proteins of the ISC andSUF machineries under these conditions. Analysis of proteins encoded by the iscRSUA-hscBA-fdx-iscX (Fig. 2A) and sufABCDSE (Fig. 2B) operons confirmed the low iron contentof the LB medium after the addition of 150 �M 2,2-DIP. As expected, the accumulationof the transcriptional regulator IscR was significantly increased under iron-limitingconditions, which resulted in a decreased abundance of the IscS, IscU, and IscA proteinsin the presence of 2,2-DIP. Under iron limitation, an ryhB ncRNA binds downstream ofiscR and cleaves the mRNA of the isc operon (60). In this case only IscR is expressed (60).Under low-iron conditions, apo-IscR binds to the promoter of sufABCDSE and induces itsexpression. Moreover, transcription of the SUF machinery is negatively regulated byFur::Fe2� and under iron-limiting conditions; thus, an increased abundance of theproteins SufA, SufB, SufC, SufD, SufE, and SufS was expected.

In addition, the three above-mentioned lacZ fusions were measured in Δfnr andΔiscS mutant strains to analyze the dependence of moaA expression on the availabilityof Fe-S clusters. The analysis of moaA-S(�FNR) and moaA-L lacZ fusions confirmed theFNR-dependent regulation of the moaABCDE operon, since the activities of moaA-S(�FNR) and moaA-L were decreased in Δfnr and ΔiscS mutant lines (Fig. 1E, F, H, and I).Interestingly, a higher activity of moaA-S(�FNR) was observed in Δfnr and ΔiscSmutants (Fig. 1G and J). This shows that either a second FNR binding site might bepresent within the moaA promoter region or that other FNR-dependent regulatorscould be involved in regulation of the operon. However, it also remains possible thatthe level of transcript initiating at the S1 promoter is elevated in the absence of FNR.Due to an overlap of the promoter sites, the S1 site might become accessible in theabsence of FNR (33). This requires further investigations in future studies.

Proteomic analysis of proteins involved in Moco biosynthesis under iron-limiting conditions. To investigate the influence of iron on the abundance of proteinsinvolved in Moco biosynthesis and on different molybdoenzymes, a detailed proteomicanalysis was performed (Fig. 3A). Bacteria were grown under anaerobic conditions in LB

FIG 2 Iron-dependent protein abundance analysis of proteins of the ISC and SUF machineries in E. coli strainBW25113. Shown is the relative abundance of the indicated proteins in cells grown anaerobically for 4 h in LB with20 mM KNO3 and 20 mM TMAO without the addition of 2,2-DIP (white bars) or in the presence of 150 �M 2,2-DIP(black bars). Standard deviations were calculated from 3 or 4 biological replicates. Asterisks indicate significantlychanged values, with P values of �0.05 (t test). P values are the following: IscR, 0.04; IscS, 0.01; IscU, 0.01; IscA, 0.03;SufA, 0.003; SufB, 0.002; SufC, 0.003; SufD, 0.001; SufE, 0.16; SufS, 0.003.

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medium with KNO3 and TMAO as electron acceptors. Iron was removed from themedium by the addition of 150 �M 2,2-DIP.

The results of the proteomic analysis in Fig. 3A show that iron availability does notinfluence the abundance of proteins involved in Moco biosynthesis, since similarabundances of the corresponding peptides were obtained in the presence or absenceof 2,2-DIP in the medium. The results were additionally confirmed by immunodetectionof MoaC, MoaE, and MogA using antisera raised against the purified proteins forcomparison (Fig. 3B). For immunodetection, bacteria were grown under anaerobicconditions in LB medium with KNO3 and TMAO in the presence or absence of 2,2-DIPbefore 50 �g of total protein lysates was separated by 15% SDS-PAGE and blotted ontoa polyvinylidene difluoride (PVDF) membrane. No changes in protein levels wereobtained in the presence of 2,2-DIP during growth.

Due to different migration properties of peptides during mass spectrometry (MS),the abundance of proteins cannot be compared quantitatively. To quantify the proteinconcentrations, we labeled peptides of the proteins involved in Moco biosynthesis withisobaric tags, which allows for the quantitative comparison of different proteins underthe analyzed conditions. The BW25113 strain was grown with and without the additionof 150 �M 2,2-DIP in the presence of KNO3 or TMAO (Fig. 3C) as an electron acceptorunder anaerobic conditions for comparison.

FIG 3 Iron-dependent protein abundance analysis of proteins involved in Moco biosynthesis in E. coli strainBW25113. (A) Relative abundance of the indicated proteins in cells grown anaerobically for 4 h in LB with 20 mMKNO3 and 20 mM TMAO without the addition of 2,2-DIP (white bars) or in the presence of 150 �M 2,2-DIP (blackbars). (B) Quantification of protein levels of MoaC, MoaE, and MogA by immunodetection. The strains were grownanaerobically in the presence of 20 mM KNO3 and 20 mM TMAO in the presence or absence of 150 �M 2,2-DIP. Cellswere lysed, and 50 �g of the total protein fraction was separated by 15% SDS-PAGE. Protein bands were transferredonto a membrane and incubated with MoaC (1:1,000)-, MoaE (1:5,000)-, or MogA (1:1,000)-specific antiserum.Target proteins were visualized by enhanced chemiluminescence. GroEL served as a loading control. Proteins fromrespective deletion strains were loaded in the middle pocket of the gel. (C) Absolute quantification of the indicatedproteins in cells grown anaerobically for 4 h in LB with 20 mM KNO3 or 20 mM TMAO without the addition of2,2-DIP (white bars) or in the presence of 150 �M 2,2-DIP (black bars). Twenty-five-microgram samples of eachpurified peptide extracted from BW25113 cells were labeled with the TMT10plex isobaric label reagent set andquantified by mass spectrometry. Standard deviations were calculated from 3 biological replicates. The asteriskindicates a significantly changed MoaB value with a P value of 0.005 (t test).

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We were able to label peptides of 6/10 enzymes involved in Moco biosynthesis:MoaB, MoaC, MoaD, MoaE, MoeA, and MoeB. The proteins MoaA and MobA were notlabeled, since their abundance was below the detection limit, as already obvious fromthe global proteomic analysis shown in Fig. 3A. No major differences in peptideabundances were observed when nitrate or TMAO was present during growth. Further,only slight differences in protein abundance were observed when 150 �M 2,2-DIP waspresent in the medium containing nitrate, while in medium containing TMAO the ironconcentration had no influence on protein abundance. Surprisingly, the abundance ofthe MoaB peptides was 4 or 16 times higher than that of the peptides of the MoaC orMoeB proteins, respectively. This result is particularly puzzling, since so far no role ofthe MoaB protein could be assigned in E. coli. Further, the MoaA, MoaB, MoaC, MoaD,and MoaE proteins are encoded by one operon, so that posttranslational factors mustexist that lead to the different abundances of these proteins under the studied growthconditions.

Proteomic analysis of molybdoenzymes and transcription factors involved inmolybdoenzyme biosynthesis under iron-limiting conditions. We analyzed theabundance of three molybdoenzymes under iron-limiting conditions, namely, formatedehydrogenase (FdnG, FdnH, and FdnI), nitrate reductase (NarG, NarH, NarI, and thechaperone NarJ), and dimethyl sulfoxide (DMSO) reductase (DmsA, DmsB, and DmsC)(Fig. 4). All three molybdoenzymes contain Fe-S clusters, and it has been reported thatthe transcription of the respective operons encoding the three enzymes is FNR depen-dent (56, 61–63). The results in Fig. 4 show that under iron-limiting conditions, theprotein abundance was largely reduced for all proteins analyzed. This likely reflects thefact that under the iron-limiting conditions with 150 �M 2,2-DIP, FNR is inactive anddoes not induce the transcription of the respective operons encoding the proteins.

Furthermore, the effect of iron on the protein abundance of different transcriptionfactors, which play a role in the expression of molybdoenzymes and enzymes involvedin Moco biosynthesis, was investigated (Fig. 5). While the abundance of ModE remainedconstant under iron-limiting conditions, the abundance of detected FNR peptides wasslightly increased under these conditions. Enhanced accumulation of FNR peptides wasexpected, since FNR inhibits its own expression (51). Further, the abundances of CsrA,ArcA, NarL, and NarP peptides were in general about 20% decreased in the presence of2,2-DIP. The arcA and narL genes were reported to be regulated by FNR (64, 65). Inaddition, the regulation of NarP by the availability of iron that was observed might bebased either on iron directly or on the presence of Fe-S clusters that influence itsexpression.

Overall Moco content under iron-limiting conditions. We analyzed the overallcontent of Moco (Fig. 6, top) and cPMP (Fig. 6, bottom) in BW25113 and ΔmoaD cellsgrown anaerobically with nitrate in the presence or absence of 150 �M 2,2-DIP. Therelative Moco and cPMP contents can be quantified from cell extracts after conversionof overall Moco (combining MPT, Mo-MPT, MCD, and bis-MGD) and cPMP to their

FIG 4 Iron-dependent protein abundance analysis of selected FNR-regulated molybdoenzymes in E. coli strainBW25113. Shown is the relative abundance of the indicated proteins in cells grown anaerobically for 4 h in LB with20 mM KNO3 and 20 mM TMAO without the addition of 2,2-DIP (white bars) or in the presence of 150 �M 2,2-DIP(black bars). Standard deviations were calculated from 3 to 4 biological replicates. Asterisks indicate significantlychanged values, with P values of �0.05 (t test). P values are the following: FdnG, 0.001; FdnH, 0.04; FdnI, 0.12; NarG,0.0004; NarH, 0.0004; NarI, 0.0000009; NarJ, 0.00001; DmsA, 0.001; DmsB, 0.003; DmsC, 0.04.

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fluorescent oxidation products, FormA and compound Z, respectively, which are gen-erated after acidic I2/KI oxidation at 95°C for 30 min. While the peptide abundance ofproteins involved in Moco biosynthesis was not changed by the iron content in themedium, surprisingly the absence of iron resulted in a lower Moco and no cPMP

FIG 5 Iron-dependent protein abundance analysis of selected transcription factors involved in theregulation of Moco biosynthesis and molybdoenzymes in E. coli strain BW25113. Shown is the relativeabundance of the indicated proteins in cells grown anaerobically for 4 h in LB with 20 mM KNO3 and20 mM TMAO without the addition of 2,2-DIP (white bars) or in the presence of 150 �M 2,2-DIP (blackbars). Standard deviations were calculated from 3 or 4 biological replicates. Asterisks indicate significantlychanged values, with P values of �0.05 (t test). P values are the following: FNR, 0.001; CsrA, 0.340; ArcA,0.01; NarL, 0.007; NarP, 0.05; ModE, 0.76.

FIG 6 Quantification of relative amounts of Moco and cPMP depending on iron availability duringgrowth. (Top) Moco content in BW25113 and ΔmoaD strain. (Bottom) cPMP content in BW25113 andΔmoaD strain. The strains were grown anaerobically in the presence of 20 mM KNO3 for 4 h in LB mediumin the absence or presence of 150 �M 2,2-DIP. Total Moco in crude extracts was oxidized overnight withacidic iodine into its fluorescence derivative, FormA. FormA was isolated and its fluorescence monitoredby excitation at 383 nm and emission at 450 nm via HPLC. Total cPMP was oxidized overnight into itsfluorescent derivative, compound Z, with acidic iodine. Compound Z was isolated, and its fluorescencewas monitored by excitation at 383 nm and emission at 450 nm via HPLC. ND, no FormA or compoundZ was detected. Standard deviations were calculated from 3 biological replicates.

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accumulation in the cell (Fig. 6, top and bottom). The undetectable cPMP content underiron-limiting conditions in ΔmoaD cells, which accumulate cPMP under normal ironconditions, is likely based on an inactive MoaA protein, the activity of which dependson two functional [4Fe-4S] clusters. However, since low levels of Moco are neverthelessproduced in BW25113 cells, small amounts of active MoaA must be present, but allproduced cPMP is immediately converted to Moco so that no cPMP accumulation isobserved. Further, in the absence of the apo-molybdoenzyme backbones under iron-limiting conditions, we assume that synthesized bis-MGD in its free form is rapidlydegraded in the cell. This would explain the lower levels of overall Moco that weredetected as FormA.

DISCUSSION

Moco biosynthesis is a complex process that is regulated by different transcriptionfactors in response to the availability of oxygen by FNR and molybdenum by ModE anddepending on the central carbon metabolism by CsrA (33, 38). It has been reported inrecent years that iron also represents an important factor for Moco biosynthesis, bothat the level of transcription and the level of activity of the proteins involved in itssynthesis (12, 33, 41). It is known that the production of Moco is directly dependent ona functional Fe-S cluster assembly by the activity of the MoaA protein, which binds two[4Fe-4S] clusters per monomer (40). Further, the transcription of the moaABCDE andmoeAB operons depends on FNR as a transcriptional factor that binds a functional[4Fe-4S] cluster only under anaerobic conditions (33, 66). The aim of our study was toinvestigate in detail the dependence of iron availability on the regulation of genesinvolved in Moco biosynthesis, with a particular focus on the moaABCDE operon.

In the past, the expression of the moaABCDE operon was mainly studied underconditions depending on the molybdate concentration in the cell (33). These studiesidentified the ModE binding site within the moaA promoter region (33). Additionalstudies were performed in LB medium with glucose under anaerobic conditions, whichensures fermentation of the cells (33, 66). The expression of the moa locus underconditions of nitrate respiration has not been studied in the past. Since the expressionof most molybdoenzymes is regulated by the two-component regulatory systems NarLand NarP, depending on the availability of nitrate (53, 56, 67, 68), we reinvestigated theiron-dependent regulation of the moaABCDE operon under conditions of nitrate res-piration. NarL and NarP belong to two-component systems, which are involved in theregulation of numerous genes under anaerobic conditions (69–71). It has been shownthat NarL represses the transcription of torCAD (53), dmsABC (72), napA (73), ynfEFGH(74), and ydhV (67) operons when nitrate is present. In contrast, the narGHJI (56), fdnGHI(68), and fdhF (68) operons are activated by NarL under conditions of nitrate respiration.Some of the molybdoenzymes are also regulated by NarP. NarP negatively regulatesthe expression of napA (73), fdnGHI (68), and ydhV (67) and was shown to be involvedin the activation of transcription of napA (73) and fdhF (68).

For comparison, we analyzed the transcription of different moaA::lacZ fusions underanaerobic conditions, in the presence of either nitrate or nitrate and TMAO. As a result,we could exclude the regulation of the moaA gene by ncRNAs. Surprisingly, weobserved a higher activity of the moaA-S(�FNR)::lacZ fusion in the Δfnr strain. Thishigher expression might be based on (i) a second FNR binding site within the promoterregion that represses the transcription of moaABCDE, (ii) regulation by an unidentifiedFNR-dependent protein, or (iii) an elevated initiation of transcripts from the S1 pro-moter site that becomes better accessible in the absence of FNR. Further work isrequired to investigate the FNR-dependent expression from each of the two promotersand their dependence on cellular iron availability.

In particular, the proteomic analyses revealed constant protein levels of proteinsinvolved in Moco biosynthesis in the presence or absence of iron. The proteins MoaAand MobA, catalyzing the first and last steps in bis-MGD biosynthesis (2), were presentonly in small amounts and could not be quantified by the proteomics approach. Arather puzzling observation was the enhanced accumulation of MoaB peptides com-

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pared to that of other proteins involved in Moco biosynthesis, like MoaC, MoaD, MoaE,MoeA, and MoeB. The results revealed that MoaB is the most abundant protein of allproteins encoded by the moaABCDE operon. So far, a role of MoaB in Moco biosynthesishas not been identified (75). However, the protein is present in high abundance in thecell, which suggests that this protein indeed does have a role in E. coli that needs to beidentified in future studies. A slightly higher abundance of proteins under iron-limitingconditions was observed when the cells were grown in the presence of nitratecompared to that of the TMAO-grown cells. This suggests that iron is more importantunder conditions of nitrate respiration, since the three E. coli nitrate reductases are Fe-Scluster-containing proteins, while TMAO reductase does not bind an Fe-S cluster (3).This iron-dependent difference in protein abundance might be based on the antago-nistic effects of the FNR-binding site within the moaA promoter region and theFNR-dependent regulation of the moeAB operon (34). However, the exact regulation ofboth operons under different conditions needs to be investigated in future studiesto clarify in detail how the expression of both moaABCDE and moeAB is fine-tunedin the cell.

In contrast, the regulation of selected molybdoenzymes by cellular iron availabilitywas, as expected, very pronounced. Peptides for nitrate reductase NarGHI, formatedehydrogenase N FdnGHI, or DMSO reductase DmsAB were almost undetectable underiron-limiting conditions, underlining the strong dependence of the expression of therespective operons on FNR. Further, we showed that under iron-limiting conditions thepresence of the cellular Moco content is largely decreased. However, we cannotexclude that in the absence of synthesized Moco the above-mentioned apo-enzymesare degraded in the cell. While the protein abundance of molybdoenzymes wasrevealed to be highly iron dependent, the protein abundance of the transcriptionalfactors involved in the expression of Moco biosynthesis or molybdoenzymes wasrevealed to only marginally depend on the cellular iron availability. Here, we investi-gated the protein abundance of FNR, CsrA, ArcA, NarL, NarP, and ModE. While ModEprotein levels remained almost constant under iron-limiting conditions and the levelsof FNR were slightly increased, the protein levels of CsrA, ArcA, NarL, and NarP wereabout 20% reduced in 2,2-DIP-treated cells.

Additionally, we also analyzed the iron-dependent protein abundance of proteinsinvolved in Fe-S cluster assembly. This quantification also served as a control to testwhether our treatment with 150 �M 2,2-DIP indeed caused iron-limiting conditions inthe cell. As expected, proteins of the ISC machinery were downregulated underiron-limiting conditions, while proteins of the SUF machinery were increased under ironlimitation. The [2Fe-2S] cluster-responsive regulator of both operons was increasedunder iron-limiting conditions. For IscR regulation, it has been shown that underiron-limiting conditions, Fe-S clusters are not inserted into IscR and apo-IscR binds tothe suf promoter region, leading to the activation of its transcription (50, 76).

Overall, our studies reveal that under iron limitation the abundance of proteinsinvolved in Moco biosynthesis was not significantly changed, while in contrast, theproduction of Moco was significantly decreased. This decrease in the cellular Mococoncentrations might be the result of two different levels of regulation. Moco biosyn-thesis was revealed to be feedback regulated by the Moco riboswitch, allowing theproduction of Moco only under conditions where it is needed (37). Under iron-limitingconditions, the production of molybdoenzymes was largely decreased. However, whenMoco is nevertheless produced under these conditions, Moco in its free protein-unbound form will be rapidly degraded due to its instability. Further, the iron-dependent regulation of IscS might additionally by an important factor for Mocobiosynthesis. IscS was shown to be involved in MPT production by providing the sulfurfor the assembly of the dithiolene group. Under iron-limiting conditions the proteinabundance of IscS was decreased to half its concentration compared to “normal” ironconcentrations in LB medium. Therefore, IscS might present the limiting factor for MPTproduction. Further, cPMP did not accumulate under iron-limiting conditions, likelybased on the inactivity of MoaA under these conditions, the activity of which depends

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on the assembly of functional [4Fe-4S] clusters and the assembly of which also dependson the presence of the IscS protein (47).

In sum, Moco biosynthesis and the assembly of active molybdoenzymes is a tightlyregulated process that additionally depends on cellular iron availability. In contrast,under iron-limiting conditions, most molybdoenzymes are not present in E. coli and thecellular content of Moco was decreased, likely as a result of an inactive MoaA proteinlacking Fe-S clusters and the additional downregulation of the abundance of theL-cysteine desulfurase IscS.

MATERIALS AND METHODSBacterial strains, plasmids, media, and growth conditions. BW25113 (referred to as the wild-type

strain) and the �iscS, �fnr, �moaC, �moaD, �moaE, and �mogA isogenic mutant strains (Keio collection)were obtained from the National BioResource Project (National Institute of Genomics, Japan) (77, 78).Bacteria were grown anaerobically in LB medium at 37°C in the presence of 20 mM KNO3 and/or 20 mMTMAO. When required, 150 �g/ml ampicillin or 25 �g/ml kanamycin was added to the medium duringgrowth. As indicated, the iron chelator dipyridyl (2,2-DIP) was added to the medium at a concentrationof 150 �M. Bacteria were grown to mid-log phase (optical density at 600 nm [OD600] of 0.5 to 1.5),collected, frozen in liquid nitrogen, and stored at �80°C for further experiments.

Construction of lacZ fusions. Promoter fragments for lacZ fusions were amplified by PCR accordingto standard protocols. Oligonucleotides contained EcoRI and/or BamHI restriction sites. Amplified PCRproducts were inserted into pJET1.2 vector (Thermo Fischer Scientific, USA) followed by digestion withappropriate restriction enzymes. Obtained fragments were purified and cloned into BamHI and/or EcoRIsites of pGE593 vector containing the lacZ gene downstream of the multiple-cloning site to createmoaA-S(�FNR), moaA-S(�FNR), and moaA-L fusions. Correct fragment insertion was confirmed by DNAsequencing.

Quantification of �-galactosidase activities. Cells were grown anaerobically in the presence of20 mM KNO3 or 20 mM KNO3 and 20 mM TMAO at 37°C until mid-log phase, and �-galactosidaseactivities were measured by using the SDS-chloroform method. Bacterial cells in 500 �l of Z buffer(60 mM Na2HPO4, 40 mM NaH2PO4, 10 mM KCl, 1 mM MgSO4, 0.05 mM �-mercaptoethanol; pH 8.0) werepermeabilized with 25 �l of 0.1% SDS and 50 �l of chloroform. Samples were incubated at 28°C for 5 min.Reactions were started by the addition of 100 �l of o-nitrophenyl-�-D-galactoside (ONPG) at a concen-tration of 4 mg/ml. Reactions were stopped by the addition of 0.25 ml of 1 M Na2CO3. The amount offormed ortho-nitrophenol was measured at 420 nm, corrected for light scattering at 550 nm, andnormalized to the volume of cells, their optical density at 600 nm, and the reaction time (Miller units). Foreach assay, a respective blank reaction containing BW25113 cells transformed with the vector controlwas subtracted.

Proteomic analysis. Bacteria were grown anaerobically in LB medium until the OD600 reachedmid-log phase. After a few washing steps in 50 mM Tris-HCl, pH 8.0, the bacterial pellet was resuspendedin 50 mM Tris-HCl, 150 mM NaCl, and 0.5 NP-40, pH 8.0, and sonicated. One hundred micrograms ofprotein lysate was mixed with 8 M urea in 10 mM Tris-HCl, pH 8.0, and loaded on a filter column(Microcon 30-kDa centrifugal filter unit with Ultracel-30 membrane). Columns were washed with 8 M ureain 10 mM Tris-HCl, pH 8.0, and reduced using 10 mM dithiothreitol in 8 M urea and alkylated using 27 mMiodoacetamide in 10 mM Tris-HCl, pH 8.0. Afterwards columns were mixed at 600 rpm in a thermomixerfor 1 min and incubated without mixing for a further 5 min. Eight molar urea in 10 mM Tris-HCl, pH 8.0,was added to each column and centrifuged. After this step, 14 h of digestion with trypsin was performed.The reaction was stopped by addition of 10% trifluoroacetic acid (TFA). Peptides were purified on C18

SepPack columns (Teknokroma) and eluted with 800 �l 60% acetonitrile (ACN), 0.1% TFA, dried in aspeed vacuum concentrator, and stored at �80°C prior to mass spectrometry analysis. Measurementswere performed on a Q Exactive HF mass spectrometer coupled with an nLC1000 nano-high-performance liquid chromatograph (nano-HPLC) (both from Thermo Scientific). Eight-microliter aliquotsof the samples were loaded onto an Acclaim PepMap RSLC reverse-phase column (75-�m inner diameter,25-cm length, 2-�m bead size; Thermo Scientific) at a flow rate of 0.4 �l min�1 in a buffer consisting of3% (vol/vol) acetonitrile, 0.5% (vol/vol) acetic acid. Peptide elution was facilitated by increasing theacetonitrile gradient from 0% to 30% (vol/vol) over 85 min, from 30% to 40% for the next 15 min, andfrom 40% to 60% for the last 8 min. The column was then washed with 80% (vol/vol) acetonitrile for6 min at a flow rate of 0.5 �l/min. Peptide ions were detected in a full scan from a mass/charge ratio of150 to 1,600 at a resolution of 60,000. Tandem MS (MS/MS) scans were performed for the ten peptideswith the highest MS signal (ddMS2 resolution of 15,000, automatic gain control [AGC] target of 2e5,isolation width mass/charge ratio of 1.6, and relative collision energy of 30). Peptides for which MS/MSspectra had been recorded were excluded from further MS/MS scans for 20 s. Quantitative analysis ofMS/MS measurements was performed with a Progenesis LC-MS (Nonlinear Dynamics). The reference runwas selected automatically. Further, an alignment and peak picking was also performed automatically.The spectra for each MS1 signal peak were exported to Mascot. Mascot search parameters were set asfollows. The Escherichia coli UniProt annotation database was used, trypsin and lysine were selected asdigesting enzymes, two missed cleavages were allowed, a fixed modification was set to carbamidom-ethylation (cysteine), and variable modifications were oxidation (methionine), peptide mass toleranceof �10 ppm, MS/MS tolerance of �0.8 Da, and allowed peptide charges of �2 and �3. Spectra were alsosearched against a decoy database of the Escherichia coli proteome, and results were filtered to obtain

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a false discovery rate below 1% at the protein level. Additionally, peptide identifications with a Mascotscore below 25 were excluded. Mascot results were imported into Progenesis QI, quantitative peak areainformation extracted, and the results exported for further analysis.

Peptide labeling and absolute quantification. Purified peptides (see “Proteomic analysis,” above)were labeled with the TMT10plex isobaric label reagent set (ThermoFisher Scientific, USA) according tothe manufacturer’s protocol, with some changes. Twenty-five micrograms of peptide mixture extractedfrom each sample was labeled with different isobaric tags. For labeling, 41 �l of different TMT isobariclabel reagents was added to each 90 �l of sample with purified peptides. Reaction mixtures wereincubated for an hour, followed by addition of 9 �l of 1 M Tris-HCl, pH 8.0, to quench the labelingreaction and incubated for 15 min at room temperature (RT). Thirty microliters of 10% TFA was added tothe samples to ensure an acidic environment for labeled peptides. Each reaction mixture was combinedin one Eppendorf tube, and labeled peptides were purified on C18 SepPack columns (Teknokroma) andeluted with 800 �l 60% ACN, 0.1% TFA, dried in a speed vacuum concentrator, and stored at �80°C priorto MS analysis. MS measurements were performed on a Q Exactive HF coupled to the Acquity UPLCM-class system (Waters). Five-microliter aliquots of the samples were loaded onto an Acquity UPLCM-class peptide CSH column (75-�m inner diameter, 25-cm length, 1.7-�m bead size; Waters) at a flowrate of 0.4 �l min�1 in a buffer consisting of 0.1% (vol/vol) formic acid. Peptide elution was facilitated byincreasing the acetonitrile gradient from 0% to 12% (vol/vol) over 20 min, from 12% to 24% for the next70 min, from 24% to 36% for 30 min, and from 36% to 85% for the last minute. The column was thenwashed with 85% (vol/vol) acetonitrile for 5 min at a flow rate of 0.3 �l/min. Peptide ions were detectedin a full scan from a mass/charge ratio of 300 to 1,600 at a resolution of 120,000. MS/MS scans wereperformed for the ten peptides with the highest MS signals (ddMS2 resolution of 15,000, AGC target of1e5, isolation width mass/charge ratio of 1.2, relative collision energy of 27). Peptides for which MS/MSspectra had been recorded were excluded from further MS/MS scans for 30 s. Quantitative analysis ofMS/MS measurements was performed with MaxQuant software (79). Escherichia coli protein sequenceswere used by the search engine Andromeda for identification of peptides. The settings used for a searchwere a 10-ppm peptide mass tolerance, 0.8-Da MS/MS tolerance, maximum of two missed cleavagesallowed, threshold for validation of peptides set to 0.01 using a decoy database, carbamidomethylationof cysteine set as a fixed modification, and oxidation of methionine set as a variable modification. Theminimum peptide length of six amino acids was used. The quantification was performed for proteins witha minimum of one unique and one razor peptide. Known contaminants, such as keratins, were removedfrom further analysis.

Immunodetection. Bacterial cultures were grown anaerobically at 37°C in LB containing 20 mMKNO3 and 20 mM TMAO until mid-log phase. Cells were collected by centrifugation, diluted in 50 mMTris-HCl, 150 mM NaCl, 0.5% NP-40 (pH 8.0), and sonicated. Protein concentrations were quantified by theBradford method. Fifty micrograms of total protein was loaded on a 15% SDS-polyacrylamide gel and,after separation, transferred onto a PVDF membrane (GE Healthcare, Buckinghamshire, UK). Membraneswere blocked with 5% bovine serum albumin in TBS-T buffer (50 mM Tris-HCl, 150 mM NaCl, 0.1% Tween20, pH 7.4) and washed three times for 5 min each time with TBS-T buffer. Afterwards, membranes wereincubated overnight at 4°C with appropriate primary antibodies (anti-MoaC at a dilution of 1:1,000,anti-MoaE at a dilution of 1:5,000, and anti-MogA at a dilution of 1:1,000), followed by 1 h of incubationwith secondary anti-rabbit antibody (anti-mouse IgG–peroxidase antibody at a dilution of 1:10,000;Sigma). Protein bands were visualized by chemiluminescence of 1:1 solution A (100 �l 250 mM luminol,44 �l 90 mM p-cumaric acid, 8.85 ml H2O, and 1 ml of 1 M Tris-HCl, pH 8.5) and solution B (6 �l of 30 H2O2,9 ml of H2O, and 1 ml of 1 M Tris-HCl, pH 8.5) mixture. Visualization was performed on a Fusion FX7(Vilber).

Detection of Moco and cPMP in cell extracts. Bacteria were grown anaerobically at 37°C with20 mM KNO3 and/or 20 mM TMAO until the cultures reached mid-log phase, followed by centrifugation,and sonicated in 100 mM Tris-HCl, pH 7.2. Samples were divided for Moco and cPMP quantification. Mocowas quantified by adding 50 �l of solution A (1,063 �l of KI solution and 100 �l of 37 HCl; KI solutionwas prepared by dissolving 1 g of I2 and 2 g of KI in 91.4 ml of Millipore water) and 150 �l of KI solutionto 400 �l of bacterial lysate. Samples were incubated at 95°C for 30 min and kept in darkness at RTovernight. Supernatant was recovered to the new Eppendorf tube, followed by addition of 100 �l of 1%ascorbic acid and 100 �l of 1 M Tris-HCl. After addition of 40 mM MgCl2 and 1 U of fast alkalinephosphatase, FormA was obtained. For purification of FormA, the samples were loaded onto a 500-�lQAE ion exchange resin (Sigma) equilibrated with water. The column was washed with 10 columnvolumes of water and with 1,300 �l 10 mM acetic acid. FormA was eluted six times with 500 �l 10 mMacetic acid. The fractions were separated on a C18 reverse-phase HPLC column (4.6- by 250-mm ODSHypersil; particle size, 5 �m) equilibrated with 5 mM ammonium acetate, 15% (vol/vol) methanol at aflow rate of 1 ml/min. Elution of FormA was monitored by an Agilent 1100 series fluorescence detectorwith excitation at 383 nm and emission at 450 nm. The total FormA content was normalized to the cellculture density (OD600). Compound Z was obtained from cPMP after the addition of 50 �l of solution A(1,063 �l of KI solution and 86 �l of 37% HCl) and 150 �l of KI solution, followed by overnight incubationat RT in the dark. Prior to elution from the QAE column, the column was washed with 10 column volumesof water and 10 mM acetic acid before compound Z was eluted five times 1,000 �l 100 mM HCl. Thefractions were separated on a C18 reverse-phase HPLC column (4.6- by 250-mm ODS Hypersil; particlesize, 5 �m) equilibrated with 10 mM potassium dihydrogen phosphate (pH 3.0), 1% (vol/vol) methanol ata flow rate of 1 ml/min. Compound Z was monitored by its fluorescence with excitation at 383 nm andemission at 450 nm. The total compound Z content was normalized to the cell culture density (OD600).

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SUPPLEMENTAL MATERIALSupplemental material for this article may be found at https://doi.org/10.1128/JB

.00382-19.SUPPLEMENTAL FILE 1, PDF file, 0.2 MB.

ACKNOWLEDGMENTSThis work was funded by the Deutsche Forschungsgemeinschaft (DFG) priority

program SPP1927 grant LE1171/15-1.We thank Jan Dahl for cloning of the moaA-S(�FNR)::lacZ fusion plasmid. We thank

Ute Armbruster (Max Planck Institute of Molecular Plant Physiology) and Alexander Graf(Max Planck Institute of Molecular Plant Physiology) for sharing their laboratory andequipment while performing MS analyses.

REFERENCES1. Leimkühler S, Iobbi-Nivol C. 2016. Bacterial molybdoenzymes: old en-

zymes for new purposes. FEMS Microbiol Rev 40:1–18. https://doi.org/10.1093/femsre/fuv043.

2. Mendel RR, Leimkühler S. 2015. The biosynthesis of the molybdenumcofactors. J Biol Inorg Chem 20:337–347. https://doi.org/10.1007/s00775-014-1173-y.

3. Iobbi-Nivol C, Leimkühler S. 2013. Molybdenum enzymes, their matura-tion and molybdenum cofactor biosynthesis in Escherichia coli. BiochimBiophys Acta 1827:1086 –1101. https://doi.org/10.1016/j.bbabio.2012.11.007.

4. Hille R, Hall J, Basu P. 2014. The mononuclear molybdenum enzymes.Chem Rev 114:3963– 4038. https://doi.org/10.1021/cr400443z.

5. Mendel RR. 2013. The molybdenum cofactor. J Biol Chem 288:13165–13172.https://doi.org/10.1074/jbc.R113.455311.

6. Rajagopalan KV. 1996. Biosynthesis of the molybdenum cofactor, p674 – 679. In Neidhardt FC, Curtiss R, III, Ingraham JL, Lin ECC, Low KB,Magasanik B, Reznikoff WS, Riley M, Schaechter M, Umbarger HE (ed),Escherichia coli and Salmonella cellular and molecular biology, vol1.ASM Press, Washington, DC.

7. Rajagopalan KV, Johnson JL. 1992. The pterin molybdenum cofactors. JBiol Chem 267:10199 –10202.

8. Hänzelmann P, Schindelin H. 2004. Crystal structure of theS-adenosylmethionine-dependent enzyme MoaA and its implications formolybdenum cofactor deficiency in humans. Proc Natl Acad Sci U S A101:12870 –12875. https://doi.org/10.1073/pnas.0404624101.

9. Hover BM, Loksztejn A, Ribeiro AA, Yokoyama K. 2013. Identification ofa cyclic nucleotide as a cryptic intermediate in molybdenum cofactorbiosynthesis. J Am Chem Soc 135:7019 –7032. https://doi.org/10.1021/ja401781t.

10. Wuebbens MM, Liu MT, Rajagopalan K, Schindelin H. 2000. Insights intomolybdenum cofactor deficiency provided by the crystal structure of themolybdenum cofactor biosynthesis protein MoaC. Structure 8:709 –718.https://doi.org/10.1016/S0969-2126(00)00157-X.

11. Hover BM, Tonthat NK, Schumacher MA, Yokoyama K. 2015. Mechanismof pyranopterin ring formation in molybdenum cofactor biosynthesis.Proc Natl Acad Sci U S A 112:6347– 6352. https://doi.org/10.1073/pnas.1500697112.

12. Sofia HJ, Chen G, Hetzler BG, Reyes-Spindola JF, Miller NE. 2001. RadicalSAM, a novel protein superfamily linking unresolved steps in familiarbiosynthetic pathways with radical mechanisms: functional character-ization using new analysis and information visualization methods. Nu-cleic Acids Res 29:1097–1106. https://doi.org/10.1093/nar/29.5.1097.

13. Hänzelmann P, Schindelin H. 2006. Binding of 5’-GTP to the C-terminalFeS cluster of the radical S-adenosylmethionine enzyme MoaA providesinsights into its mechanism. Proc Natl Acad Sci U S A 103:6829 – 6834.https://doi.org/10.1073/pnas.0510711103.

14. Pitterle DM, Rajagopalan KV. 1993. The biosynthesis of molybdopterin inEscherichia coli. Purification and characterization of the converting fac-tor. J Biol Chem 268:13499 –13505.

15. Pitterle DM, Johnson JL, Rajagopalan KV. 1993. In vitro synthesis ofmolybdopterin from precursor Z using purified converting factor. Role ofprotein-bound sulfur in formation of the dithiolene. J Biol Chem 268:13506 –13509.

16. Daniels JN, Wuebbens MM, Rajagopalan KV, Schindelin H. 2008. Crystal

structure of a molybdopterin synthase-precursor Z complex: insight into itssulfur transfer mechanism and its role in molybdenum cofactor deficiency.Biochemistry 47:615–626. https://doi.org/10.1021/bi701734g.

17. Zhang W, Urban A, Mihara H, Leimkühler S, Kurihara T, Esaki N. 2010. IscSfunctions as a primary sulfur-donating enzyme by interacting specificallywith MoeB and MoaD in the biosynthesis of molybdopterin in Esche-richia coli. J Biol Chem 285:2302–2308. https://doi.org/10.1074/jbc.M109.082172.

18. Dahl J-U, Radon C, Bühning M, Nimtz M, Leichert LI, Denis Y, Jourlin-Castelli C, Iobbi-Nivol C, Méjean V, Leimkühler S. 2013. The sulfur carrierprotein TusA has a pleiotropic role in Escherichia coli that also affectsmolybdenum cofactor biosynthesis. J Biol Chem 288:5426 –5442. https://doi.org/10.1074/jbc.M112.431569.

19. Leimkühler S, Rajagopalan KV. 2001. A sulfurtransferase is required in thetransfer of cysteine sulfur in the in vitro synthesis of molybdopterin fromprecursor Z in Escherichia coli. J Biol Chem 276:22024 –22031. https://doi.org/10.1074/jbc.M102072200.

20. Leimkühler S, Wuebbens MM, Rajagopalan KV. 2001. Characterization ofEscherichia coli MoeB and its involvement in the activation of MPTsynthase for the biosynthesis of the molybdenum cofactor. J Biol Chem276:34695–34701. https://doi.org/10.1074/jbc.M102787200.

21. Xiang S, Nichols J, Rajagopalan KV, Schindelin H. 2001. The crystalstructure of Escherichia coli MoeA and its relationship to the multifunc-tional protein gephyrin. Structure 9:299 –310. https://doi.org/10.1016/S0969-2126(01)00588-3.

22. Nichols JD, Rajagopalan KV. 2005. In vitro molybdenum ligation tomolybdopterin using purified components. J Biol Chem 280:7817–7822.https://doi.org/10.1074/jbc.M413783200.

23. Nichols J, Rajagopalan KV. 2002. Escherichia coli MoeA and MogA.Function in metal incorporation step of molybdenum cofactor bio-synthesis. J Biol Chem 277:24995–25000. https://doi.org/10.1074/jbc.M203238200.

24. Neumann M, Mittelstadt G, Seduk F, Iobbi-Nivol C, Leimkühler S. 2009.MocA is a specific cytidylyltransferase involved in molybdopterin cyto-sine dinucleotide biosynthesis in Escherichia coli. J Biol Chem 284:21891–21898. https://doi.org/10.1074/jbc.M109.008565.

25. Lake MW, Temple CA, Rajagopalan KV, Schindelin H. 2000. The crystalstructure of the Escherichia coli MobA protein provides insight intomolybdopterin guanine dinucleotide biosynthesis. J Biol Chem 275:40211– 40217. https://doi.org/10.1074/jbc.M007406200.

26. Temple CA, Rajagopalan KV. 2000. Mechanism of assembly of the bis-(molybdopterin guanine dinucleotide)molybdenum cofactor in Rhodo-bacter sphaeroides dimethyl sulfoxide reductase. J Biol Chem 275:40202– 40210. https://doi.org/10.1074/jbc.M007407200.

27. Johnson JL, Indermaur LW, Rajagopalan KV. 1991. Molybdenum cofactorbiosynthesis in Escherichia coli. Requirement of the chlB gene productfor the formation of molybdopterin guanine dinucleotide. J Biol Chem266:12140 –12145.

28. Reschke S, Sigfridsson KG, Kaufmann P, Leidel N, Horn S, Gast K, SchulzkeC, Haumann M, Leimkühler S. 2013. Identification of a bis-molybdopterinintermediate in molybdenum cofactor biosynthesis in Escherichia coli. JBiol Chem 288:29736 –29745. https://doi.org/10.1074/jbc.M113.497453.

29. Thome R, Gust A, Toci R, Mendel R, Bittner F, Magalon A, Walburger A.2012. A sulfurtransferase is essential for activity of formate dehydroge-

Regulation of Moco Biosynthesis Genes by Iron Journal of Bacteriology

September 2019 Volume 201 Issue 17 e00382-19 jb.asm.org 13

on July 22, 2020 by guesthttp://jb.asm

.org/D

ownloaded from

nases in Escherichia coli. J Biol Chem 287:4671– 4678. https://doi.org/10.1074/jbc.M111.327122.

30. Kaufmann P, Duffus BR, Mitrova B, Iobbi-Nivol C, Teutloff C, Nimtz M, JanschL, Wollenberger U, Leimkühler S. 2018. Modulating the molybdenum coor-dination sphere of escherichia coli trimethylamine N-oxide reductase. Bio-chemistry 57:1130 –1143. https://doi.org/10.1021/acs.biochem.7b01108.

31. Shanmugam KT, Stewart V, Gunsalus RP, Boxer DH, Cole JA, Chippaux M,DeMoss JA, Giordano G, Lin ECC, Rajagopalan KV. 1992. Proposed no-menclature for the genes involved in molybdenum metabolism in Esch-erichia coli and Salmonella typhimurium. Mol Microbiol 6:3452–3454.https://doi.org/10.1111/j.1365-2958.1992.tb02215.x.

32. Rosentel JK, Healy F, Maupin-Furlow JA, Lee JH, Shanmugam KT. 1995.Molybdate and regulation of mod (molybdate transport), fdhF, and hyc(formate hydrogenlyase) operons in Escherichia coli. J Bacteriol 177:4857– 4864. https://doi.org/10.1128/jb.177.17.4857-4864.1995.

33. Anderson LA, McNairn E, Lubke T, Pau RN, Boxer DH, Leubke T. 2000.ModE-dependent molybdate regulation of the molybdenum cofactoroperon moa in Escherichia coli. J Bacteriol 182:7035–7043. https://doi.org/10.1128/jb.182.24.7035-7043.2000.

34. Hasona A, Self WT, Shanmugam KT. 2001. Transcriptional regulation ofthe moe (molybdate metabolism) operon of Escherichia coli. Arch Mi-crobiol 175:178 –188. https://doi.org/10.1007/s002030100252.

35. Unden G, Achebach S, Holighaus G, Tran HG, Wackwitz B, Zeuner Y.2002. Control of FNR function of Escherichia coli by O2 and reducingconditions. J Mol Microbiol Biotechnol 4:263–268.

36. Unden G, Schirawski J. 1997. The oxygen-responsive transcriptionalregulator FNR of Escherichia coli: the search for signals and reactions.Mol Microbiol 25:205–210. https://doi.org/10.1046/j.1365-2958.1997.4731841.x.

37. Regulski EE, Moy RH, Weinberg Z, Barrick JE, Yao Z, Ruzzo WL, BreakerRR. 2008. A widespread riboswitch candidate that controls bacterialgenes involved in molybdenum cofactor and tungsten cofactor metab-olism. Mol Microbiol 68:918 –932. https://doi.org/10.1111/j.1365-2958.2008.06208.x.

38. Patterson-Fortin LM, Vakulskas CA, Yakhnin H, Babitzke P, Romeo T. 2013.Dual posttranscriptional regulation via a cofactor-responsive mRNAleader. J Mol Biol 425:3662–3677. https://doi.org/10.1016/j.jmb.2012.12.010.

39. Sabnis NA, Yang H, Romeo T. 1995. Pleiotropic regulation of centralcarbohydrate metabolism in Escherichia coli via the gene csrA. J BiolChem 270:29096 –29104. https://doi.org/10.1074/jbc.270.49.29096.

40. Yokoyama K, Leimkühler S. 2015. The role of FeS clusters for molybde-num cofactor biosynthesis and molybdoenzymes in bacteria. BiochimBiophys Acta 1853:1335–1349. https://doi.org/10.1016/j.bbamcr.2014.09.021.

41. Hanzelmann P, Hernandez HL, Menzel C, Garcia-Serres R, Huynh BH,Johnson MK, Mendel RR, Schindelin H. 2004. Characterization ofMOCS1A, an oxygen-sensitive iron-sulfur protein involved in humanmolybdenum cofactor biosynthesis. J Biol Chem 279:34721–34732.https://doi.org/10.1074/jbc.M313398200.

42. Tanaka N, Kanazawa M, Tonosaki K, Yokoyama N, Kuzuyama T, TakahashiY. 2016. Novel features of the ISC machinery revealed by characteriza-tion of Escherichia coli mutants that survive without iron-sulfur clusters.Mol Microbiol 99:835– 848. https://doi.org/10.1111/mmi.13271.

43. Outten FW, Djaman O, Storz G. 2004. A suf operon requirement for Fe-Scluster assembly during iron starvation in Escherichia coli. Mol Microbiol52:861– 872. https://doi.org/10.1111/j.1365-2958.2004.04025.x.

44. Zheng L, Cash VL, Flint DH, Dean DR. 1998. Assembly of iron-sulfurclusters. Identification of an iscSUA-hscBA-fdx gene cluster from Azoto-bacter vinelandii. J Biol Chem 273:13264 –13272. https://doi.org/10.1074/jbc.273.21.13264.

45. Yuvaniyama P, Agar JN, Cash VL, Johnson MK, Dean DR. 2000. NifS-directed assembly of a transient [2Fe-2S] cluster within the NifU protein.Proc Natl Acad Sci U S A 97:599 – 604. https://doi.org/10.1073/pnas.97.2.599.

46. Nuth M, Cowan JA. 2009. Iron-sulfur cluster biosynthesis: characteriza-tion of IscU-IscS complex formation and a structural model for sulfidedelivery to the [2Fe-2S] assembly site. J Biol Inorg Chem 14:829 – 839.https://doi.org/10.1007/s00775-009-0495-7.

47. Schwartz CJ, Djaman O, Imlay JA, Kiley PJ. 2000. The cysteine desul-furase, IscS, has a major role in in vivo Fe-S cluster formation in Esche-richia coli. Proc Natl Acad Sci U S A 97:9009 –9014. https://doi.org/10.1073/pnas.160261497.

48. Leimkühler S, Buhning M, Beilschmidt L. 2017. Shared sulfur mobilization

routes for tRNA thiolation and molybdenum cofactor biosynthesis inprokaryotes and eukaryotes. Biomolecules 7:E5. https://doi.org/10.3390/biom7010005.

49. Bai Y, Chen T, Happe T, Lu Y, Sawyer A. 2018. Iron-sulphur clusterbiogenesis via the SUF pathway. Metallomics 10:1038 –1052. https://doi.org/10.1039/c8mt00150b.

50. Yeo WS, Lee JH, Lee KC, Roe JH. 2006. IscR acts as an activator inresponse to oxidative stress for the suf operon encoding Fe-S assemblyproteins. Mol Microbiol 61:206 –218. https://doi.org/10.1111/j.1365-2958.2006.05220.x.

51. Mettert EL, Kiley PJ. 2014. Coordinate regulation of the Suf and IscFe-S cluster biogenesis pathways by IscR is essential for viability ofEscherichia coli. J Bacteriol 196:4315– 4323. https://doi.org/10.1128/JB.01975-14.

52. Vinella D, Loiseau L, Ollagnier de Choudens S, Fontecave M, Barras F.2013. In vivo [Fe-S] cluster acquisition by IscR and NsrR, two stressregulators in Escherichia coli. Mol Microbiol 87:493–508. https://doi.org/10.1111/mmi.12135.

53. Iuchi S, Lin EC. 1987. The narL gene product activates the nitratereductase operon and represses the fumarate reductase and trimethyl-amine N-oxide reductase operons in Escherichia coli. Proc Natl Acad SciU S A 84:3901–3905. https://doi.org/10.1073/pnas.84.11.3901.

54. Bonnefoy V, Demoss JA. 1994. Nitrate reductases in Escherichia coli. AntonieVan Leeuwenhoek 66:47–56. https://doi.org/10.1007/BF00871632.

55. Blasco F, Iobbi C, Ratouchniak J, Bonnefoy V, Chippaux M. 1990. Nitratereductases of Escherichia coli: sequence of the second nitrate reductaseand comparison with that encoded by the narGHJI operon. Mol GenGenet 222:104 –111.

56. Stewart V. 1982. Requirement of Fnr and NarL functions for nitratereductase expression in Escherichia coli K-12. J Bacteriol 151:1320 –1325.

57. Pinske C, Sawers RG. 2012. A-type carrier protein ErpA is essential forformation of an active formate-nitrate respiratory pathway in Escherichiacoli K-12. J Bacteriol 194:346 –353. https://doi.org/10.1128/JB.06024-11.

58. Pascal MC, Burini JF, Chippaux M. 1984. Regulation of the trimethyl-amine N-oxide (TMAO) reductase in Escherichia coli: analysis of tor::Mud1 operon fusion. Mol Gen Genet 195:351–355. https://doi.org/10.1007/BF00332770.

59. Spiro S, Guest JR. 1990. FNR and its role in oxygen-regulated geneexpression in Escherichia coli. FEMS Microbiol Rev 6:399 – 428. https://doi.org/10.1111/j.1574-6968.1990.tb04109.x.

60. Desnoyers G, Morissette A, Prevost K, Masse E. 2009. Small RNA-induceddifferential degradation of the polycistronic mRNA iscRSUA. EMBO J28:1551–1561. https://doi.org/10.1038/emboj.2009.116.

61. McNicholas PM, Chiang RC, Gunsalus RP. 1998. Anaerobic regulation ofthe Escherichia coli dmsABC operon requires the molybdate-responsiveregulator ModE. Mol Microbiol 27:197–208. https://doi.org/10.1046/j.1365-2958.1998.00675.x.

62. Grainger DC, Aiba H, Hurd D, Browning DF, Busby SJ. 2007. Transcriptionfactor distribution in Escherichia coli: studies with FNR protein. NucleicAcids Res 35:269 –278. https://doi.org/10.1093/nar/gkl1023.

63. Myers KS, Yan H, Ong IM, Chung D, Liang K, Tran F, Keles S, Landick R,Kiley PJ. 2013. Genome-scale analysis of Escherichia coli FNR revealscomplex features of transcription factor binding. PLoS Genet9:e1003565. https://doi.org/10.1371/journal.pgen.1003565.

64. Compan I, Touati D. 1994. Anaerobic activation of arcA transcription inEscherichia coli: roles of Fnr and ArcA. Mol Microbiol 11:955–964. https://doi.org/10.1111/j.1365-2958.1994.tb00374.x.

65. Kumar R, Shimizu K. 2011. Transcriptional regulation of main metabolicpathways of cyoA, cydB, fnr, and fur gene knockout Escherichia coli inC-limited and N-limited aerobic continuous cultures. Microb Cell Fact10:3. https://doi.org/10.1186/1475-2859-10-3.

66. Hasona A, Ray RM, Shanmugam KT. 1998. Physiological and geneticanalyses leading to identification of a biochemical role for the moeA(molybdate metabolism) gene product in Escherichia coli. J Bacteriol180:1466 –1472.

67. Partridge JD, Browning DF, Xu M, Newnham LJ, Scott C, Roberts RE,Poole RK, Green J. 2008. Characterization of the Escherichia coli K-12ydhYVWXUT operon: regulation by FNR, NarL and NarP. Microbiology154:608 – 618. https://doi.org/10.1099/mic.0.2007/012146-0.

68. Wang H, Gunsalus RP. 2003. Coordinate regulation of the Escherichia coliformate dehydrogenase fdnGHI and fdhF genes in response to nitrate,nitrite, and formate: roles for NarL and NarP. J Bacteriol 185:5076 –5085.https://doi.org/10.1128/jb.185.17.5076-5085.2003.

69. Katsir G, Jarvis M, Phillips M, Ma Z, Gunsalus RP. 2015. The Escherichia

Zupok et al. Journal of Bacteriology

September 2019 Volume 201 Issue 17 e00382-19 jb.asm.org 14

on July 22, 2020 by guesthttp://jb.asm

.org/D

ownloaded from

coli NarL receiver domain regulates transcription through promoterspecific functions. BMC Microbiol 15:174. https://doi.org/10.1186/s12866-015-0502-9.

70. Stewart V. 1994. Dual interacting two-component regulatory systemsmediate nitrate- and nitrite-regulated gene expression in Escherichiacoli. Res Microbiol 145:450 – 454. https://doi.org/10.1016/0923-2508(94)90093-0.

71. Rabin RS, Stewart V. 1993. Dual response regulators (NarL and NarP)interact with dual sensors (NarX and NarQ) to control nitrate- andnitrite-regulated gene expression in Escherichia coli K-12. J Bacteriol175:3259 –3268. https://doi.org/10.1128/jb.175.11.3259-3268.1993.

72. Bearson SM, Albrecht JA, Gunsalus RP. 2002. Oxygen and nitrate-dependent regulation of dmsABC operon expression in Escherichia coli:sites for Fnr and NarL protein interactions. BMC Microbiol 2:13. https://doi.org/10.1186/1471-2180-2-13.

73. Stewart V, Bledsoe PJ. 2003. Synthetic lac operator substitutions forstudying the nitrate- and nitrite-responsive NarX-NarL and NarQ-NarPtwo-component regulatory systems of Escherichia coli K-12. J Bacteriol185:2104 –2111. https://doi.org/10.1128/jb.185.7.2104-2111.2003.

74. Xu M, Busby SJ, Browning DF. 2009. Activation and repression at the

Escherichia coli ynfEFGHI operon promoter. J Bacteriol 191:3172–3176.https://doi.org/10.1128/JB.00040-09.

75. Kozmin SG, Schaaper RM. 2013. Genetic characterization of moaB mu-tants of Escherichia coli. Res Microbiol 164:689 – 694. https://doi.org/10.1016/j.resmic.2013.05.001.

76. Ayala-Castro C, Saini A, Outten FW. 2008. Fe-S cluster assembly path-ways in bacteria. Microbiol Mol Biol Rev 72:110 –125. https://doi.org/10.1128/MMBR.00034-07.

77. Yamamoto N, Nakahigashi K, Nakamichi T, Yoshino M, Takai Y, Touda Y,Furubayashi A, Kinjyo S, Dose H, Hasegawa M, Datsenko KA, NakayashikiT, Tomita M, Wanner BL, Mori H. 2009. Update on the Keio collection ofEscherichia coli single-gene deletion mutants. Mol Syst Biol 5:335.https://doi.org/10.1038/msb.2009.92.

78. Baba T, Ara T, Hasegawa M, Takai Y, Okumura Y, Baba M, Datsenko KA,Tomita M, Wanner BL, Mori H. 2006. Construction of Escherichia coli K-12in-frame, single-gene knockout mutants: the Keio collection. Mol SystBiol 2:2006 0008. https://doi.org/10.1038/msb4100050.

79. Cox J, Mann M. 2008. MaxQuant enables high peptide identificationrates, individualized p.p.b.–range mass accuracies and proteome-wideprotein quantification. Nat Biotechnol 26:1367–1372. https://doi.org/10.1038/nbt.1511.

Regulation of Moco Biosynthesis Genes by Iron Journal of Bacteriology

September 2019 Volume 201 Issue 17 e00382-19 jb.asm.org 15

on July 22, 2020 by guesthttp://jb.asm

.org/D

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