Shifting the metallocentric molybdoenzyme paradigm: the importance of pyranopterin coordination.

J Biol Inorg Chem (2015) 20:349–372 DOI 10.1007/s00775-014-1194-6

MINIREVIEW

Shifting the metallocentric molybdoenzyme paradigm: the importance of pyranopterin coordination Richard A. Rothery • Joel H. Weiner

Received: 30 June 2014 / Accepted: 15 September 2014 / Published online: 30 September 2014 Ó SBIC 2014

Abstract In this review, we test the hypothesis that pyranopterin coordination plays a critical role in defining substrate reactivities in the four families of mononuclear molybdenum and tungsten enzymes (Mo/W-enzymes). Enzyme families containing a single pyranopterin dithiolene chelate have been demonstrated to have reactivity towards two (sulfite oxidase, SUOX-fold) and five (xanthine dehydrogenase, XDH-fold) types of substrate, whereas the major family of enzymes containing a bispyranopterin dithiolene chelate (dimethylsulfoxide reductase, DMSOR-fold) is reactive towards eight types of substrate. A second bis-pyranopterin enzyme (aldehyde oxidoreductase, AOR-fold) family catalyzes a single type of reaction. The diversity of reactions catalyzed by each family correlates with active site variability, and also with the number of pyranopterins and their coordination by the protein. In the case of the AOR-fold enzymes, inflexibility of pyranopterin coordination correlates with their limited substrate specificity (oxidation of aldehydes). In examples of the SUOX-fold and DMSOR-fold enzymes, we observe

three types of histidine-containing charge-transfer relays that can: (1) connect the piperazine ring of the pyranopterin to the substrate-binding site (SUOX-fold enzymes); (2) provide inter-pyranopterin communication (DMSOR-fold enzymes); and (3) connect a pyran ring oxygen to deeply buried water molecules (the DMSOR-fold NarGHI-type nitrate reductases). Finally, sequence data mining reveals a number of bacterial species whose predicted proteomes contain large numbers (up to 64) of Mo/W-enzymes, with the DMSOR-fold enzymes being dominant. These analyses also reveal an inverse correlation between Mo/W-enzyme content and pathogenicity. Keywords Cofactor Electrochemistry Electron transfer Metallocenter assembly Abbreviations AOR AOR-fold DMSOR DMSOR-fold LUA Mo-bisPGD Mo-PCD

Responsible Editors: Jose´ Moura and Paul Bernhardt. R. A. Rothery J. H. Weiner (&) Department of Biochemistry, University of Alberta, Edmonton, AB T6G 2H7, Canada e-mail: [email protected]

Mo-PPT Mo/ W-enzymes NIA SUOX SUOX-fold W-bisPPT XDH XDH-fold

Aldehyde oxidoreductase Aldehyde oxidoreductase protein fold Dimethylsulfide reductase DMSO reductase protein fold Last universal ancestor Molybdo-bis(pyranopterin guanine dinucleotide) Molybdo-pyranopterin cytosine dinucleotide Molybdo-pyranopterin Mononuclear molybdenum or tungsten enzymes Plant-type nitrate reductase Sulfite oxidase Sulfite oxidase protein fold Tungsto-bispyranopterin Xanthine dehydrogenase Xanthine dehydrogenase protein fold

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Introduction Scope of the review Advances in structural biology and biophysics are providing remarkable insights into the reactions catalyzed by the mononuclear Mo- and W-enzymes (Mo/W-enzymes) [1– 6]. At the heart of these enzymes lies the pyranopterin dithiolene chelate that helps define a catalytically competent metal coordination sphere. Depending on the type of enzyme, the metal environment can also be defined by oxogroups, sulfido-groups, and/or coordination by amino acid side chains such as those of Ser, Cys, and Asp [5]. Overall, these factors enable redox cycling through the ?4, ?5, and ?6 metal oxidation states and the catalysis of two electron redox chemistry. Across the range of Mo/W-enzymes, the active site can catalyze redox transformations spanning a reduction potential range in excess of 1 V.1 Unsurprisingly, the critical role of the metal ion in catalysis has resulted in a metallocentric view of enzyme catalysis, with the extended pyranopterin until recently receiving comparatively little attention [4, 5, 13–16]. Recent studies of model compounds able to mimic Mo/W-enzyme active sites suggest that the pyranopterin dithiolene may have a noninnocent role in defining active site redox chemistry [17– 19]. These observations were supported by analyses of pyranopterin conformations within high-resolution Mo/Wenzyme structures, suggesting that the pyranopterin can exist in both tetrahydro- and 10,10a-dihydro-redox states [20]. In this review, we explore the relationship between pyranopterin–protein interactions and the observed diversity of substrate specificity. Because of the large number of excellent reviews on the subject, we do not consider the environment of the Mo/W and the specifics of bondbreaking and making catalysis [4, 5, 13, 14, 21]. For the sake of brevity, we also do not consider the modularity and multimeric nature of many of the enzymes, which have also been extensively reviewed [1–3, 22–25]. A cofactor of scalable complexity The Mo/W-enzymes contain a Mo or W atom coordinated by one or two pyranopterin dithiolene chelates (Fig. 1). The pyranopterin is a tricyclic heterocycle comprising pyrimidine, piperazine, and pyran rings (labeled a, b, and c in Fig. 1a). Protein crystallography has permitted their assignment to four families based on the four protein folds 1

Reduction potentials of some common substrates are as follows: methanofuran/formylmethanofuran, -500 mV; SO42-/SO32-, -442 mV; uric acid/xanthine, -440 mV; CO2/HCO2-, -432 mV; Me2SO/Me2S, ?160 mV; NO3-/NO2-, ?420 mV; ClO4-/ClO3-, ?806 mV [1, 7–12].

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J Biol Inorg Chem (2015) 20:349–372

observed to coordinate catalytically active Mo- or W-pyranopterin [22]. In increasing complexity of cofactor, these are as follows. (1) The sulfite oxidase (SUOX-fold) family, which includes bacterial, plant, and mammalian sulfite oxidases, plant assimilatory nitrate reductase, and two types of prokaryotic enzymes of currently unknown function (YedY and YuiH) [2]. The SUOX-fold enzymes contain the simplest manifestation of the molybdo-pyranopterin (Mo-PPT) illustrated in Fig. 1a. (2) The xanthine dehydrogenase (XDH-fold) family, which participates in a range of reactions including oxidation of purines, isoquinoline, nicotinic acid (vitamin B3) and carbon monoxide. While eukaryotic and some prokaryotic XDH-fold enzymes contain Mo-PPT, most prokaryotic examples contain the more complex molybdo-pyranopterin cytosine dinucleotide (Mo-PCD) illustrated in Fig. 1b [26]. The aerobic carbon monoxide dehydrogenases have additional modifications, including the presence of a dinuclear heterometal active site (CuSMo(=O)OH) (inset, Fig. 1b) or incorporation of a selenium atom at the active site (as a Mo–S–Se moiety) [23, 27, 28]. (3) The aldehyde oxidoreductase (AOR-fold) family, of which the archetype is the archaeal aldehyde:ferredoxin oxidoreductase from the hyperthermophile Pyrococcus furiosus [29–31]. These contain the tungsto-bispyranopterin (W-bisPPT) illustrated in Fig. 1c, and catalyze oxidations of aldehydes to their corresponding carboxylic acids. (4) The dimethylsulfoxide reductase (DMSOR-fold) family, of which the eponymous member is the Rhodobacter dimethylsulfoxide reductase [32, 33]. These typically contain the molybdo-bis(pyranopterin guanine dinucleotide) (Mo-bisPGD) illustrated in Fig. 1d. Almost all DMSOR-fold enzymes contain an iron– sulfur cluster in close juxtaposition to one of the pyranopterins, which is thus referred to as the proximal pyranopterin, whereas the other is referred to as the distal pyranopterin [3, 20]. A final cofactor variant is observed in two structurally characterized DMSOR-fold enzymes, Aromatoleum aromaticum ethylbenzene dehydrogenase (EbdABC) and Escherichia coli respiratory nitrate reductase (NarGHI), wherein the distal pyranopterin has a bicyclic rather than tricyclic structure [34, 35]. The scalable complexity of the mononuclear Mo/Wcofactors illustrated in Fig. 1 comes at great metabolic cost and requires a biosynthetic machinery comprised of up to ten proteins [36, 37]. This raises the following question: why did evolution lead to such diversity of cofactor and protein fold design? This diversity has led to three examples of convergent evolution. (1) Nitrate reductases exist in the SUOX-fold family (with the simpler Mo-PPT cofactor) and in the DMSOR-fold family (incorporating the more complex Mo-bisPGD cofactor). (2) Two unrelated carbon monoxide dehydrogenases exist, the aerobic enzymes with the XDH-fold discussed herein (containing the Mo-PCD


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Fig. 1 Variations on the pyranopterin dithiolene theme. a The molybdo-pyranopterin cofactor (Mo-PPT) found in eukaryotic molybdoenzymes, some bacterial xanthine dehydrogenases (XDH) and all sulfite oxidases (SUOX). The tricyclic pyranopterin comprises a pyrimidine, b piperazine, and c pyran rings. b The molybdo(pyranopterin cytosine dinucleotide) cofactor (MoPCD) found in most bacterial XDH-fold enzymes [23]. Inset the dinuclear heterometal (CuSMo(=O)OH) found in CO dehydrogenase from Oligotropha carboxidovorans [27], but not in the CO dehydrogenase from H. pseudoflava [76]. c The tungstobis(pyranopterin) cofactor (WbisPPT) found in the archaeal aldehyde oxidoreductases [30, 31]. d The molybdobis(pyranopterin guanine dinucleotide) cofactor (MobisPGD) found in the majority of bacterial molybdoenzymes that can also exist in a W-bisPGD form [1, 3]. In two examples, the E. coli respiratory nitrate reductase (NarGHI) and the ethylbenzene dehydrogenase (EbdABC) from A. aromaticum, one of the pyranopterins has a bicyclic structure [34, 35]

cofactor), and an anaerobic enzyme that contains a completely unrelated nickel–iron–sulfur active site [38]. (3) Aldehyde oxidoreductases exist in the AOR-fold family,

and in the XDH-fold family (e.g., the aldehyde dehydrogenase from Desulfovibrio gigas [39]). Herein, we review evidence that diversity of cofactor design enables

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Table 1 Bait sequences used for sequence database mining Organism

PDB Codea

UNIPROT/SWISSPROT accession

Sulfite oxidase

Gallus gallus

1SOX

SUOX_CHICK

Mo-PPT

Sulfite oxidase

Arabidopsis thaliana

1OGP

SUOX_ARATH

Mo-PPT

Sulfite dehydrogenase

Starkeya novella

2BLF

Q9LA16_STANO

Mo-PPT

Assimilatory nitrate reductase

Pichia angusta

2BII

NIA_PICAN

Mo-PPT

Novel oxidoreductase

E. coli

1XDQ

YEDY_ECOLI

Mo-PPT

Xanthine dehydrogenase

Homo sapiens

2E1Q

XDH_HUMAN

Mo-PPT

Xanthine dehydrogenase

Bos taurus

3NRZ

XDH_BOVIN

Mo-PPT

Aldehyde dehydrogenase

Desulfovibrio gigas

3L4P

MOP_DESGI

Mo-PCD

Isoquinoline oxidoreductase

Pseudomonas putida

1T3Q

P72224_PSEPU

Mo-PCD

4-Hydroxybenzoyl-CoA reductase

Thauera aromatica

1RM6

HCRA_THAAR

Mo-PCD

Carbon monoxide dehydrogenase

Hydrogenophaga pseudoflava

1FFV

DCML_HYDPS

Mo-PCD

Carbon monoxide dehydrogenase

Oligotropha carboxidovorans

1N62

DCML_OLICO

Mo-PCD (CuSMo(=O)OH)

Aldehyde oxidoreductase

Pyrococcus furiosus

1AOR

AOR_PYRFU

W-bisPPT

Formaldehyde oxidoreductase

Pyrococcus furiosus

1B25

Q8U1K3_PYRFU

W-bisPPT

Arsenite oxidase

Alcaligenes faecalis

1G8 K

AIOA_ALCFA

Mo-bisPGD

Dimethylsulfoxide dehydrogenase

Rhodobacter capsulatus

1DMR

DSTOR_RHOCA

Mo-bisPGD

Formate dehydrogenase

Desulfovibrio gigas

1H0H

FDHA_DESGI

W-bisPGD

Formate dehydrogenase H

E. coli

2IV2

FDHF_ECOLI

Mo-bisPGD

Formate dehydrogenase N

E. coli

1KQF

FDNG_ECOLI

Mo-bisPGD

Periplasmic nitrate reductase

Desulfovibrio desulfuricans

2V3 V

NAPA_DESDA

Mo-bisPGD


E. coli

2NYA

NAPA_ECOLI

Mo-bisPGD


Rhodobacter sphaeroides

1OGY

NAPA_RHOS4

Mo-bisPGD


Cupriavidus necator

3ML1

NAPA_CUPNH

Mo-bisPGD

Respiratory nitrate reductase

E. coli

1Q16

NARG_ECOLI

Mo-bisPGD

Enzyme

Cofactor

SUOX-fold

XDH-fold

AOR-fold

DMSOR-fold

Pyrogallol hydroxytransferase

Pelobacter acidigallici

1VLF

PGTL_PELAC

Mo-bisPGD

Trimethylamine-N-oxide reductase

Shewanella massilia

1TMO

TORA_SHEMA

Mo-bisPGD

Ethylbenzene dehydrogenase

Aromatoleum aromaticum

2IVF

Q5P5I0_AROAE

Mo-bisPGD

Acetylene hydratase

Pelobacter acetylenicus

2E7Z

AHY_PELAE

W-bisPGD

Polysulfide reductase

Thermus thermophilus

2VPZ

Q72LA4_THET2

Mo-bisPGD

a

1SOX [50], 1OGP [119], 2BII [58], 2BLF [120], 2BII [58], 1XDQ [60], 2E1Q [71], 3NRZ [72], 3LP4 [39], 1T3Q [74], 1RM6 [75], 1FFV [76], 1N62 [27], 1AOR [31], 1B25 [30], 1G8K [111], 1DMR [103], 1H0H [110], 2IV2 [113], 1KQF [102], 2V3V [106], 2NYA [107], 1OGY [108], 3ML1 [109], 1Q16 [35], 1VLF [105], 1TMO [104], 2IVF [34], 2E7Z [112], 2VPZ [101]

additional facets of enzyme reactivity tuning that complements the role of the active site metal and its immediate environment. Taxonomic distribution of mononuclear Mo/W-enzyme folds As of January 2014, the UNIPROT sequence database (http://www.uniprot.org) contained approximately 49 million protein sequence entries, almost all of which are from taxonomically identified organisms. Its SWISSPROT

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subset contains curated entries for which a biological function is either known or can be predicted with reasonable confidence. Sequence database growth has been complemented by remarkable progress in obtaining highresolution protein structures. We used sequences of available mononuclear Mo/W-enzyme structures (Table 1) as bait to search the non-redundant subset of UNIPROT (cutoff at 90 % identity, UNIREF90) using the BLASTP [40] search algorithm, generating large sets of unique sequences similar to those of structurally characterized SUOX, XDH, AOR and DMSOR enzyme subunits that


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contain a mononuclear Mo/W cofactor. We further reduced redundancy in our data sets using the program CDHIT [41], eliminating sequences with greater than 70 % pairwise identity to any other in the data set. We confirmed assignment of our sequences to the fourfolds of Mo/Wenzymes by aligning them using the program MAFFT [42]. Functional assignments of protein sequences were obtained by performing BLASTP searches against the curated SWISSPROT database, and taxonomic information was obtained from the UNIPROT database. This strategy allowed assembly of non-redundant taxonomically assignable data sets comprising 2,773, 1,355, 2,079, and 120 sequences for the DMSOR-, SUOX-, XDH-, and AORfolds, respectively. Figure 2 shows the distribution of the four protein folds across the three domains of life, with emphasis on their occurrence in bacteria. The approximate evolutionary pathway from the last universal ancestor (LUA) is based on the 2013 version of the ‘‘All-Species Living Tree’’ project (http://www.arb-silva.de/projects/living-tree/) [43], modified to combine the Acidobacteria and Fibrobacteres as a single group (http://www.uniprot.org/taxonomy/). Only one bacterial phylum is devoid of Mo/W-enzymes: this is the Chlamydiae, comprised of obligate intracellular pathogens responsible for an important class of sexually transmitted diseases [44]. The genome of Chlamydia trachomatis, an ocular and genital pathogen, encodes approximately 900 predicted proteins (compared to *4,300 encoded by the E. coli K12 genome), suggesting exploitation of the host metabolome during the intracellular phase of the C. trachomatis lifecycle. This provides a plausible explanation for the lack of Mo/W-enzymes in the Chlamydiae.

the respective protein fold, and far fewer have multiple copies. In the case of the AOR-fold, our analyses are limited by the low-number AOR-fold sequences in the UNIPROT database (our data set contained representatives of 46 genera and a total of 120 sequences). Nevertheless, we identified a number of ‘‘heavy-hitting’’ genera, including Desulfobacterium (Proteobacteria; mesophilic sulfate reducer [45]), Pelotomaculum (Firmicutes, thermophilic [46]), Caldivirga (Archaea, thermophilic [47]), and Geobacter (Proteobacteria, metal reducer [48]). Although the well-characterized archetypes of the AORfold enzymes come from the hyperthermophilic Archaeon P. furiosus, it is notable that these enzymes are predicted to also appear in mesophilic species. To provide further insight into the role of the AOR-fold enzymes, we sequentially used each of them as bait to search the curated SWISSPROT database, enabling assignments of putative functions (Fig. 4). The overwhelming majority of AOR-fold enzymes catalyze the oxidation of aldehydes [31], with subsets predicted to specialize in formaldehyde oxidation [30] and glyceraldehyde-3-phosphate oxidation [49]. Finally, four sequences are assigned as members of the YdhV family of proteins (e.g., YDHV_ECOLI), for which no function has yet been demonstrated. As discussed below (see ‘‘Inflexibility of AOR family pyranopterin coordination’’), reactivity of AOR-fold proteins towards a single type of substrate (aldehydes) correlates with an inflexibility of coordination of the pyranopterin moieties of the W-bisPPT cofactor.

Taxonomic distribution and functions of the AOR-fold enzymes

Enzymatic sulfite oxidation plays critical roles in human health and in the global sulfur cycle. In humans, SUOX functions in the breakdown of cysteine and methionine. Deficiency, either via enzyme-inactivating point mutations [50] or via mutations in the Mo-PPT biosynthesis pathway [51], typically results in death in early infancy. Lethality is at least partly due to intracellular sulfite build up, which causes sulfitolysis of disulfide bonds, resulting in protein instability and subsequent degradation [52]. Symptoms include neurodegeneration, intractable seizures, mental retardation, and ocular lens dislocation [53]. In the case of cofactor deficiency, a therapy has been developed that involves injection of the biosynthetic intermediate cyclic pyranopterin monophosphate [53, 54]. However, there remains no treatment for SUOX deficiency due to enzymeinactivating point mutations. While animal SUOX enzymes function in the mitochondrial inter-membrane space to pass electrons from sulfite to cytochrome c, plant SUOX is located in the peroxisome and transfers electrons from

Of the four Mo/W-enzyme protein folds, the AOR-fold has the sparsest taxonomic representation, appearing in the Archaea, Thermotogae, Firmicutes, Chloroflexi, Nitrospirae, Deinococcus-Thermus, and the Proteobacteria. The distribution of these groups on the putative evolutionary pathway, specifically the absence of AOR-fold enzymes between the Cyanobacteria and the Spirochetes, is convincing evidence for their dissemination via horizontal gene transfer. To gain further insights into the distribution of Mo/W-enzymes, we drilled down to the genus level, enabling identification of what we term ‘‘heavy-hitting’’ genera (viz. genera containing large numbers of genes for the respective Mo/W-enzyme folds). Figure 3 shows plots of the number of genera versus the number of Mo/W-fold enzyme sequences occurring in our data sets. In general, large numbers of genera have few copies of genes encoding

Taxonomic distribution and functions of the SUOX-fold enzymes

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Fig. 2 Evolutionary persistence of the four Mo/W-enzyme families. In some cases, a paucity of UNIPROT data limits our confidence in excluding enzyme classes from particular taxonomic groups. For example, protein sequences assigned to the phylum Caldiserica arise from the genome sequence of a single species, Caldisericum exile, which is predicted to contain all the 1,566 proteins assigned to this group in UNIPROT. Another example is the Thermodesulfobacteria, with all 3,778 of the 3,797 sequences in UNIPROT arising from just

two complete genomes, those of Thermodesulfatator indicus [115] and Thermodesulfobacterium geofontis [116]. In the case of the Chlamydiae (marked with an arrow), there are over 144,000 UNIPROT sequences, and 167 complete genome sequences, indicating that the absence of Mo/W-enzymes from this taxonomic group is significant. Phyla/domains in labeled in gray text are monoderms, whereas those labeled in blue text are diderms. LUA last universal ancestor

sulfite directly to oxygen, generating hydrogen peroxide [55–57]. Plant SUOX plays a role in adaptation to acid rain and resistance to sulfite toxicity. A third well-characterized SUOX-fold enzyme is the plant-type assimilatory nitrate reductase (NIA) which is also found in yeasts such as Pichia augustina [58]. A fourth type of SUOX-fold enzyme is represented by the YedY enzyme encoded by the E. coli genome [2, 59, 60]. It is periplasmically localized, has limited reactivity towards S- and N-oxides (i.e., kcat = *20 s-1 for trimethylamine-N-oxide [59, 60]), and its unusual electrochemical properties render it an excellent model system for studying the Mo(V) catalytic intermediate of the Mo-PPT cofactor [61–63]. Finally, a fifth group is identified as YuiH in SWISSPROT, and has similarities to YedY. The precise physiological roles of YedY and YuiH are unknown. We assembled a sequence data set comprising 1,355 SUOX-fold sequences. Our taxonomic analyses (Fig. 2) reveal that SUOX-fold enzymes are absent in the Spirochaetes, Fusobacteria, Chrysiogenetes, Chlorobi,

Chlamydiae, Lentisphaerae, Synergistetes, Thermodesulfobacteria, and Thermotogae. As will be seen below (see ‘‘Mo/W-enzyme multiplicity and pathogenicity’’, below), the Spirochaetes contain a number of important pathogens including those responsible for syphilis, leptospirosis, and Lyme disease. The Fusobacteria are normal constituents of oral flora, the Chlorobi are photolithotropic sulfur oxidizers [64]. The Chrysiogenetes are able to respire anaerobically using arsenate as terminal electron acceptor utilizing an enzyme belonging to the DMSORfold family [65]. The Lentisphaerae are closely related to the Chlamydiae [66], and appear to only have DMSORfold Mo/W-enzymes. The Synergistetes are found in a range of aqueous environments and as a constituent of the animal oral flora [67]. The Thermodesulfobacteria are thermophilic sulfate reducers [68]. Finally, the Thermotogae are thermophiles that can be found in proximity to ocean-floor thermal vents [69]. Our data set of 1,355 sequences represents a total of 581 genera (Fig. 3), with the following genera contributing

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Fig. 3 Genera with large numbers of predicted Mo/W-enzymes. For each enzyme family, plots are shown of the number of genera versus number of predicted enzymes. To limit scatter in the plot, the number of sequences per genera were rounded to the nearest log2 integer prior to plotting. In general, a large number of genera in each data set

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contain a small number of predicted enzymes belonging to each family, whereas a small number are predicted to contain a large number of predicted enzymes. Data points are labeled with the corresponding highly represented genera and the actual number of available Mo/W-enzyme sequences (unrounded)

Fig. 4 Distribution of predicted substrate specificity amongst the AOR-fold enzymes. Functional assignments of sequences are based on BLASTP searches against the curated SWISSPROT database. Sequences were assigned as aldehyde:ferredoxin oxidoreductases (AOR), formaldehyde:ferredoxin oxidoreductases (FOR), glyceraldehyde:ferredoxin oxidoreductases (GAPOR), and YDHV (unknown function), respectively

large numbers of sequences: Streptomyces (Actinobacteria, 41 sequences), Mycobacterium (Actinobacteria, 22 sequences), Methylobacterium (Proteobacteria, 16

sequences), Bacillus (Firmicutes, 17 sequences), and Rhodococcus (Actinobacteria, 13 sequences). It is instructive to compare the number of ‘‘heavy-hitting’’ SUOX-fold genera

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with the number of genera for which complete genome sequences are available (available at http://www.ncbi.nlm. nih.gov/genome/browse/). Figure 5 shows plots of the number genera versus the number of completed genome sequences for both prokaryotes and eukaryotes. 1,292 genera are represented in the prokaryotes, yielding a total of 21,236 genomes (as of February 2014). The genome ‘‘heavy-hitters’’ include the genera Staphylococcus (2,890 genomes, ranked 1st), Escherichia (1,795, ranked 2nd), Streptococcus (1,208, ranked 3rd), Salmonella (1,073, ranked 4th), Mycobacterium (1,048, ranked 5th), and Acinetobacter (790, ranked 6th). These genera all include important pathogenic species (Table 2). There are 145 Streptomyces (ranked 23rd), 26 Methylobacterium (ranked 76th), 416 Bacillus (ranked 11th), and 36 Rhodococcus (ranked 56th) genomes. In general, these observations confirm a bias in database submissions favoring either pathogenic genera (e.g., Salmonella, Mycobacterium), or those of biotechnological relevance (e.g., Streptomyces). However, genera amongst the SUOX-fold ‘‘heavy-hitters’’ do include Rhodococcus (36 genomes, ranked 56th), an example of which is strain RHA1, which has a proteome comprising over 9,000 predicted proteins (compared to, for example, the approximately 4,300 encoded by E. coli strain K12). Rhodococcus RHA1 is of biotechnological significance and is able to degrade a range of organic compounds, including halogenated aromatics, lignin, and the toxic industrial by-product N-nitrosodimethylamine [70]. Its genome encodes 19 predicted Mo/W-enzymes, but only three of these belong to the SUOX-fold family (Table 2). Thus, that the Rhodococcus genus appears to be a SUOXfold ‘‘heavy-hitter’’ is a reflection of the diversity and number of its SUOX sequences represented in UNIPROT (individual and genome submissions). A species that is a genuine ‘‘heavy-hitter’’ is Methylobacterium extorquens, with its predicted proteome containing 28 Mo/W-enzymes, of which 7 are members of the SUOX-fold family. Our data set of 1,355 sequences presented an excellent opportunity to evaluate the predicted substrate specificity of the SUOX-fold enzymes. Figure 6 shows the distribution of predicted functions within our data set. Remarkably, *50 % of SUOX-fold proteins are assigned functionally to the YedY or YuiH subgroups whose physiological substrates are still unknown. The taxonomic distribution of YedY is biased towards the Proteobacteria, whereas that of YuiH is biased towards the Actinobacteria, Chloroflexi, Firmicutes, and the Archaea (data not shown). The large number of YedY and YuiH sequences within UNIPROT indicates that these two enzymes must play a critical biological role. Within the SUOX-fold family the presence of a single Mo-PPT correlates with predicted reactivity towards a limited number of known substrates, currently corresponding to oxidation of sulfite or reduction of nitrate.

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Fig. 5 Representation of genera in completed genome sequences. Plots of genera versus number of completed genomes as of February 2014. To limit scatter in the plot, the numbers of genomes were rounded to the nearest log2 integer (as in Fig. 3). Plots for both prokaryotic and eukaryotic genomes are shown. In general, a large number of genera are represented by a small number of complete genome sequences, whereas a small number of genera are represented by a large number of complete genome sequences. Data points are labeled with their corresponding highly represented genera and the actual number of available genome sequences (unrounded)

Taxonomic distribution and functions of the XDH-fold enzymes Human XDH deficiency causes xanthinuria, where xanthine accumulates as tissue deposits, and as urinary tract calculi (kidney stones). In extreme cases this can lead to acute renal failure [53]. Treatments for xanthinuria include high fluid intake, dehydration prevention, and consumption of a low purine diet. As is the case with SUOX, deficiency arises either from point mutations causing expression of non-functional variants, or from generalized molybdenum cofactor deficiency. In general, XDH-fold enzymes play a critical role in purine metabolism, but are also reactive towards other types of substrates. Members of this family include human and bovine XDH [71, 72], Pseudomonas sp. strain CBB1 caffeine dehydrogenase [73], D. gigas aldehyde dehydrogenase [39], P. putida quinoline oxidoreductase [74], Thauera aromatica 4-hydroxybenzoyl-CoA reductase [75], the Hydrogenophaga pseudoflava and Oligotropha carboxidovorans carbon monoxide dehydrogenases [27, 76], and P. putida nicotinate (vitamin B3) dehydrogenase [77]. For the most part, the prokaryotic members contain Mo-PCD, whereas the eukaryotic members contain Mo-PPT. The CO-dehydrogenase from Oligotropha carboxidovorans [27] contains a novel dinuclear heterometal (CuSMo(=O)OH) active site, whereas that from H. pseudoflava contains a SeMo-active site [28].

Proteobacteria Proteobacteria Proteobacteria Archaea

Escherichia coli (O127:H6, EPEC)

Geobacter uraniireducens (Rf4)

Desulfovibrio vulgaris (ATCC 29579)

Methanosarcina acetivorans (ATCC 35395)

Proteobacteria

Salmonella typhimurium (ATCC 68169)

Proteobacteria

Archaea

Proteobacteria

Rhizobium leguminosarum (WSM2304) Enterobacter cloacae (ATCC 13047)

Pyrobaculum aerophilum (ATCC 51768)

Proteobacteria

Escherichia coli (O139:H21, ETEC)

Proteobacteria

Gut commensal

Firmicutes

Pelotomaculum thermopropionicum (DSM 13744)

Shewanella frigidimarina (NCIMB 400)

Soil nitrogen fixer

Proteobacteria

Methane producer (implicated in end-Permian mass extinction)

Sulfate reducer

Bioremediation (e.g. heavy metals)

Enterohemorrhagic diarrrhea

Typhoid, food poisoning

Hyperthermophile

Psychrophile

Enterotoxigenic diarrhea

Thermophile, syntropic

Laboratory strain

Can produce bio-active steroids, acrylamide, etc.

Nosocomial, normal component of skin flora

Antibiotic production (e.g., neomycin, chloramphenicol)

Enterohemorrhagic diarrhea

Nosocomial (e.g. cystic fibrosis patients).

Actinobacteria

Proteobacteria

Burkholderia ambifaria (ATCC BAA-244)

Found in plants, facultative methylotroph

Escherichia coli (K12)

Proteobacteria

Methylobacterium extorquens (ATCC 14718)

Sulfate reducer

Rhodococcus (RHA1)

Proteobacteria

Desulfobacterium autotrophicum (ATCC 43914)

Found in the rumen of sheep

Proteobacteria

Actinobacteria

Slackia heliotrinireducens (ATCC 29202)

Soil nitrogen fixer

Pseudomonas aeruginosa (ATCC 15692)

Proteobacteria

Bradyrhizobium diazoefficiens (JCM 10833)

Gut commensal, link with Crohn’s disease

Proteobacteria

Actinobacteria

Eggerthella lenta (ATCC 25559)

Gut commensal, link with Crohn’s disease

Dehalogenates organic compounds

Actinobacteria

Actinobacteria

Gordonibacter pamelaeae (7-10-1-b)

Streptomyces coelicolor (ATCC BAA-471)

Firmicutes

Desulfitobacterium hafniense (Y51)

Applicable disease/commentb

Escherichia coli (O157:H7, EHEC)

Phyluma

Species (strain)

Table 2 Mo/W-enzyme content of representative predicted proteomes

4,468

3,517

4,255

4,568

4,539

2,590

3,990

5,411

6,403

4,915

2,884

4,307

9,075

5,563

8,038

6,486

6,600

6,233

4,846

2,750

8,253

3,054

2,027

5,014

Proteome sizec

11

11

12

14

14

14

15

16

17

18

18

18

19

20

21

24

27

28

29

34

38

44

49

64

Mo/Wenzymesd

5

8

6

11

13

7

11

15

9

13

5

13

8

9

9

19

11

9

16

27

12

34

49

59

DMSOR

1

0

0

1

1

1

1

1

6

1

0

1

3

2

4

1

3

7

0

5

7

8

0

0

SUOX

Protein folds

2

1

1

1

0

2

3

0

2

3

3

3

8

9

8

3

13

12

3

0

19

0

0

4

XDH

3

2

5

1

0

4

0

0

0

1

10

1

0

0

0

1

0

0

10

2

0

2

0

1

AOR

J Biol Inorg Chem (2015) 20:349–372 357

123

123 Proteobacteria Proteobacteria Proteobacteria Proteobacteria Firmicutes Firmicutes Firmicutes

Mycobacterium tuberculosis (ATCC 25177)

Brucella melitensis (ATCC 23457)

Bordetella pertussis (ATCC 9797)

Yersinia pestis

Campylobacter jejuni (RM122)

Enterococcus faecalis (62)

Clostridium botulinum (657)

Clostridium beijerinckii (ATCC 51743)

Acetone, ethanol, butanol production from starch Chronic gastritis/gastric ulcers

Firmicutes Firmicutes Firmicutes Firmicutes

Clostridium tetani (Massachusetts)

Staphylococcus aureus (MRSA252)

Lactobacillus plantarum (ATCC BAA-793)

Clostridium acetobutylicum (ATCC 824)

Diptheria Syphilis

Actinobacteria Spirochaetes

Corynebacterium diphtheriae (ATCC 700971) Treponema pallidum (SS14)

Pneumonia, meningitis

Proteobacteria Firmicutes

Helicobacter pylori (51)

Streptococcus pneumoniae (ATCC BAA-255)

Food industry (e.g. Sauerkraut, kimchi, sourdough)

Methicillin-resistant Staphylococcus aureus, infections of respiratory/urinary tract

Tetanus

Gangrene, food poisoning

Respiratory tract infections

Proteobacteria Firmicutes

Clostridium perfringens (13)

Saprophyte Atopic dermatitis

Anthrax

Cholera

Production of butanol, acetone, isopropanol

Botulism

Endocarditis, meningitis

Gastroenteritis

Bubonic plague (via fleas)

Pertussis (whooping cough)

Brucellosis

1,028

2,265

2,030

1,412

3,847

3,088

2,640

2,415

2,721

1,876

3,595 2,889

5,493

3,784

5,003

3,977

3,010

1,836

3,909

3,167

3,125

3,990

3,635

‘‘Iraqibacter’’ wound infectionse Tuberculosis

2,045

3,762

3,897

1,962

4,197

Proteome sizec

Hyperthermophile

Pseudomembranous colitis

Dysentery (shigellosis)

Hyperthermoacidophile

Gut commensal


Haemophilus influenzae (10810)

Spirochaetes Firmicutes

Actinobacteria

Acinetobacter baumannii

Leptospira biflexa (Patoc 1) Staphylococcus aureus (NCTC 8325)

Proteobacteria

Pyrococcus furiosus (ATCC 43587)

Proteobacteria

Archaea

Clostridium difficile (630)

Firmicutes

Firmicutes

Shigella dysenteriae (Sd197)

Bacillus anthracis

Proteobacteria

Caldivirga maquilingensis (ATCC 700844)

Vibrio cholerae (ATCC 39315)

Firmicutes Archaea

Bacillus subtilis (168)

Phyluma

Species (strain)

Table 2 continued

1

1

1

1

1

1

2

2

2

2

2 2

3

3

3

4

4

5

5

6

7

7

7

7

8

9

9

9

Mo/Wenzymesd

0

1

1

1

0

1

2

0

2

2

2 2

3

3

1

0

1

3

5

2

3

4

5

2

3

8

0

7

DMSOR

0

0

0

0

0

0

0

0

0

0

0 0

0

0

0

0

0

2

0

1

2

2

0

0

0

1

2

1

SUOX

Protein folds

1

0

0

0

0

0

0

2

0

0

0 0

0

0

2

3

2

0

0

3

2

1

2

0

5

0

0

1

XDH

0

0

0

0

1

0

0

0

0

0

0 0

0

0

0

1

1

0

0

0

0

0

0

5

0

0

7

0

AOR

358 J Biol Inorg Chem (2015) 20:349–372

1,599

629

3,135

2,307

917

1,289 3,654

3,183

1,963

1,887

2,424

2,136

Proteome sizec

0

0

0

0

0

0 0

0

0

0

0

0

Mo/Wenzymesd

0

0

0

0

0

0 0

0

0

0

0

0

DMSOR

0

0

0

0

0

0 0

0

0

0

0

0

SUOX

Protein folds

0

0

0

0

0

0 0

0

0

0

0

0

XDH

0

0

0

0

0

0 0

0

0

0

0

0

AOR

Predicted proteome sizes based on analyses of completed genome. Sequences were downloaded from http://www.uniprot.org in FASTA format

e

Known for nosocomial infections during Iraq and Afghanistan wars [121]

Each predicted protein sequence was used as bait to search the SWISSPROT curated protein sequence database (January 2014 version), generating a list of putative protein functions. Identification of proteins with DMSOR, SUOX, XDH, and AOR protein folds was confirmed by examination of bait sequences for appropriateness of length and electronic annotation in the UNIPROT databases

d

c

Pathogens are depicted in bold font, and are identified as being causative agents of well-characterized diseases. Note that some organisms cause opportunistic nosocomial infections (of immuno-compromised patients), and these are not treated as pathogens herein

b

Leprosy

Pneumonia, bronchitis

STDs: e.g. urethritis, proctitis, trachoma

For simplicity, members of the Archaeal domain are listed as ‘‘Archaea’’

Actinobacteria

a

Mycobacterium leprae (Br4923)

Listeriosis

Firmicutes Firmicutes

Firmicutes

Streptococcus pyogenes (serotype M1) Listeria monocytogenes

Mycoplasma pneumoniae (ATCC 15531)

Strep throat, scarlet fever

Chlamydiae

Chlamydia trachomatis (ATCC VR571B)

Legionellosis (pneumonia) Lyme disease (via Ixodes tick) Leptospirosis

Proteobacteria Spirochaetes Spirochaetes

Borrelia burgdorferi (ATCC 35210) Leptospira interrogans (Fiocruz L1130)

Gonorrhea

Meningitis

Skin flora conmensal

Intestinal mucin degradation, gut commensal


Legionella pneumophila (2300/99)

Proteobacteria Proteobacteria

Firmicutes

Staphylococcus warneri (SG1)

Neisseria gonorrhoeae (ATCC 700825)

Verrucomicrobia

Akkermansia muciniphila (ATCC BAA-835)

Neisseria meningitidis (Z2491)

Phyluma

Species (strain)

Table 2 continued

J Biol Inorg Chem (2015) 20:349–372 359

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360

Six bacterial phyla lack XDH-fold enzymes, including the Chrysiogenetes, the Chlorobi, the Lentisphaerae, the Chlamydiae, the Nitrospirae, and the Thermodesulfobacteria (Fig. 2). Figure 7 shows the predicted distribution of substrate specificity within our data set of XDH-fold sequences towards five types of substrates. That the majority are predicted to catalyze oxidation of xanthine or caffeine is a reflection of the biological importance of purine metabolism. Quinoline oxidation is exclusively bacterial, whereas CO-oxidation occurs in both bacteria and archaea. Aldehyde oxidation occurs in all three domains. Likewise, nicotinate oxidation and 4-hydroxybenzoyl-CoA

Fig. 6 Distribution of predicted substrate specificity amongst the SUOX-fold enzymes. Assignments were carried out as described in the legend to Fig. 4. Sequences were assigned as sulfite oxidases (SUOX), nitrate reductases (NIA), YEDY (unknown function), and YUIH (unknown function)

Fig. 7 Distribution of predicted substrate specificity amongst the XDH-fold enzymes. Assignments were carried out as described in the legend to Fig. 4. SWISSPROT acronyms were assigned as follows: purine/caffeine dehydrogenases (XDH, XDH1, XDHA, XDHD, YAGR, and CDHA [caffeine dehydrogenase]); quinoline oxidoreductases (DHAQ [e.g. DHAQ_GLUPO [117]], IORB); nicotinate dehydrogenase or 4-hydroxybenzoyl-CoA reductase (NICB, NDLMS, KDHC, HCRA), aldehyde oxidoreductase (MOP, CUTA, ALDO, ADO); carbon monoxide dehydrogenase (DCML)

123


reduction are predicted to occur only in the bacteria and archaea. Overall, it is important to note that the XDH-fold enzymes coordinate cofactors containing a single pyranopterin dithiolene ligand, and that they are predicted to react with five overall types of substrate, with three of these being aromatic compounds. In the case of the CO dehydrogenases, it is clear that adaptations to the Moactive site are largely responsible for carbon monoxide oxidation. Our data set of 2,079 XDH-fold sequences represents 561 genera (Fig. 3), of which the following can be considered ‘‘heavy hitters’’: Clostridium (Firmicutes, 61 sequences), Streptomyces (Actinobacteria, 57 sequences), Pseudomonas (Proteobacteria, 46 sequences), Burkholderia (Proteobacteria, 40 sequences), Rhizobium (Proteobacteria, 32 sequences), and Bradyrhizobium (Proteobacteria, 31 sequences). Species of Clostridium include difficile (causative agent of pseudomembranous colitis, with five predicted XDH-fold enzymes, Table 2), botulinum (botulism, 3 enzymes), tetani (tetanus, 2 enzymes), perfringens (gangrene, 0 enzymes), beijerinckii (biofuel production, 2 enzymes), and acetobutylicum (biofuel production, 0 enzymes). Clearly, the genus Clostridium comprises both pathogens and species of biotechnological importance, resulting in a large number of complete genomes (488 genomes, ranked 10th) and this has contributed to its large number of sequence database entries. It is also notable that there is significant variability within the predicted Clostridium proteomes, with between 5 and 0 predicted instances of XDH-fold proteins. The predicted proteome of S. coelicolor contains eight XDH-fold enzymes, while that


of B. diazoefficiens contains 19 (there are 51 Bradyrhizobium genome sequences, ranked 47th). Taxonomic distribution and functions of the DMSORfold enzymes Enzymes containing a DMSOR-fold subunit and the Mo/ W-bisPGD cofactor are able to catalyze reactions spanning a reduction potential range from formylmethanofuran oxidation (to methanofuran; Em,7 & -500 mV [12]) to the reduction of perchlorate (to chlorate; Em,7 = ?806 mV), a reduction potential range of *1.3 V. The broad range of substrates and overall modularity of enzymes containing a DMSOR-fold subunit have been reviewed extensively [1, 3, 22, 78]. A fundamental question remains: how does the DMSOR-fold provide for such extreme redox and metabolic flexibility? DMSOR-fold enzymes occur in all the taxonomic groupings of Fig. 2, with the exception of the Caldiserica [79] and the Chlamydiae. In the former case, all 1,566 protein sequences present in UNIPROT arise from the completed genome sequence of the Caldisericum exile, which is an anaerobic thermophile [79]. 15 DMSOR-fold sequences were assigned as being fungal in origin, however, this compares with over 9 million sequences of eukaryotic proteins in UNIPROT. We therefore conclude, in agreement with previous studies, that DMSOR-fold

Fig. 8 Distribution of predicted substrate specificity amongst the DMSOR-fold enzymes. Assignments were carried out as described in the legend to Fig. 4. SWISSPROT acronyms were assigned as follows: formate dehydrogenase (FDHA, YJGC, FDHF, FDHL, FDOG, FDNG, FDXG), S- and N-oxidoreductases (DMSA, BISC, DSTOR, YNFE, YNFF, TORZ, DDHA, TORA), nitrate/selenate/ perchlorate reductases (NARB, NASA, NARG, NARZ, NAPA,

361

enzymes are essentially exclusively prokaryotic in origin [3, 22]. Our data set of 2,773 DMSOR-fold sequences represents 623 genera (Fig. 3). Compared to the other families of Mo/ W-enzymes, we noted a higher number of sequences per genera among the ‘‘heavy-hitters’’, including Desulfitobacterium (Firmicutes, 77 sequences), Desulfovibrio (Proteobacteria, 75 sequences), Clostridium (Firmicutes, 59 sequences), Slackia (Actinobacteria, 39 sequences), Bacillus (Firmicutes, 38 sequences), and Mycobacterium (Actinobacteria, 37 sequences). Of these, the Clostridium, Bacillus, and Mycobacterium genera are also among the genome heavy hitters (Fig. 5), and as noted above, this is due to the presence within them of important pathogenic species. However, Desulfitobacterium, Desulfovibrio, and Slackia have only 9 (ranked 157th), 43 (ranked 52nd), and 5 (ranked 242nd) sequenced genomes, respectively. We, therefore, examined the predicted proteomes of examples of these genera (Table 2): Desulfitobacterium hafniense, Desulfovibrio vulgaris, and Slackia heliotrinireducens are predicted to contain 59, 8, and 27 predicted DMSOR-fold proteins, respectively, confirming D. hafniense and S. heliotrinireducens as genuine ‘‘heavy hitters’’. Highlights of the predicted proteome of D. hafniense include 20 enzymes categorized as S- and N-oxide reductases and 6 categorized as nitrate reductases. Highlights of the predicted proteome of S. heliotrinireducens include 14 enzyme categorized as

NASC, PCRA, SERA), sulfur anion reductases (PHSA, PSRA, TTRA), acetylene hydratase (AHY), formylmethanofuran dehydrogenase (FWDB), arsenite oxidase (AIOA, 25 sequences, not labeled), Pyrogallol hydroxytransferase (PGTL, 18 sequences, not labeled). Sequences of unknown function returned the following SWISSPROT acronyms: YOAE, YYAE, Y2924, YDEP, and Y006

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362

S- and N-oxidoreductases and five categorized as nitrate reductases. The remarkable flexibility of the DMSOR-fold enzymes is reflected in the plethora of reactions they catalyze. Figure 8 summarizes the predicted reactivities of the enzymes assigned by performing BLASTP searches against SWISSPROT using our 2,773 sequence data set as bait. Reactivities are predicted towards formate, nitrate/selenate/ perchlorate, S- and N-oxides (dimethylsulfoxide, dimethylsulfide, biotin sulfoxide, and trimethylamine-N-oxide), sulfur anions (polysulfide, tetrathionate, and thiosulfate), acetylene, formylmethanofuran (43 sequences), arsenite (25 sequences), and pyrogallol (18 sequences). A further 580 sequences are assigned to the SWISSPROT entries for which functions are unknown, such as YOAE, YYAE, and YDEP. Based on this interpretation of our BLASTP results, the DMSOR family of enzymes has 8–9 generalized functions. However, if we were to treat the individual SWISSPROT acronyms as distinct functions, with individual sulfur anion reactions and oxyanion reactions treated as distinct reactivities, then there would be up to 35 subgroups, compared to 18 for the XDH-fold family, and 4 for the SUOX-fold family. Overall this supports the following hypothesis: the presence of the second pyranopterin in the DMSOR-fold enzymes presents opportunities for variability of cofactor coordination that facilitates the remarkable range of redox transitions catalyzed. Assigning functions to proteins using their sequences as bait in BLASTP searches against the SWISSPROT database generates results that are susceptible to erroneous annotation or omission of sequences corresponding to enzymes that have been biochemically characterized, but for which the details have not yet been added to the curated database. This is the reason why we present only generalized reactivity distributions in Figs. 4, 6, 7, and 8. A pertinent example of the omission issue is the ethylbenzene dehydrogenase (EbdABC) from A. aromaticum for which reactivity and high-resolution structural information are available [34, 80], but for which there is no SWISSPROT entry. Searching the electronic annotations within the UNIPROT database reveals the presence of EbdABC enzymes only in A. aromaticum, Azoarcus sp. ED1 (both closely related Proteobacteria) and Rhodococcus RHA1 (Actinobacteria). We interpret this to mean that ethylbenzene oxidation is a very rare metabolic functionality. Mo/W-enzyme multiplicity and pathogenicity Three observations led us to examine the relationship between Mo/W-enzyme multiplicity and prokaryotic pathogenicity: (1) our observation of significant numbers of pathogenic species in our analyses of ‘‘heavy-hitting’’ genera; (2) our observation that many pathogens can

123


contain few or no predicted Mo/W-enzymes; and (3) our identification of species that contain many copies of genes for Mo/W-enzymes, particularly arising from our analyses of the DMSOR-fold taxonomic distribution. We, therefore, analyzed the predicted proteomes of species selected on the basis of their representation in the data points of Figs. 3 and 5. As a control group, we selected a number of wellcharacterized pathogenic species, as well as non-pathogens of general interest (e.g., Methanosarcina acetivorans [81]). Predicted proteome sequence data sets were obtained from UNIPROT (at http://www.uniprot.org/taxonomy/completeproteomes), and each protein had a function assigned to it by using it as bait to search the curated SWISSPROT database. This enabled us to estimate the number of predicted Mo/W-enzymes belonging to the AOR, SUOX, XDH, and DMSOR families in each species. Table 2 summarizes the predicted number of the various Mo/Wenzymes for 64 species belonging to the bacteria and archaea. We analyzed 29 non-pathogenic species and 35 pathogenic species. Inspection of the data clearly indicates that enzymes belonging to the DMSOR family are largely responsible for the high number of predicted Mo/Wenzymes in ‘‘heavy-hitting’’ species. The species predicted to contain the most Mo/Wenzymes is the Firmicute D. hafniense, a strict anaerobe originally isolated from environments contaminated by halogenated organic compounds, including the potent toxin pentachlorophenol [82, 83]. Its genome encodes a possible total of 59 DMSOR enzymes, of which 20 are predicted to act on S- and N-oxide substrates, even though such enzymes do not appear to be directly involved in dehalogenation reactions. Six of the DMSOR enzymes are predicted to catalyze sulfur anion reduction, and 11 are predicted to be acetylene hydratases. Ranked second in Table 2 is the Actinobacterium Gordonibacter pamelaeae, whose genome encodes 49 predicted Mo/W-enzymes, all of which are members of the DMSOR family. Remarkably 26 of these are predicted to be acetylene hydratases, and a further eight are predicted to act on S- and N-oxide substrates. G. pamelaeae was found in the intestinal flora of a patient with acute Crohn’s disease [84]. The third ranked species in Table 2 is Eggerthella lenta, an Actinobacterium which has also been found in the intestine of a Crohn’s disease patient [85], which is predicted to have 34 DMSOR-fold enzymes, 8 SUOX-fold enzymes and 2 AOR-fold enzymes. Of the 34 DMSOR-fold enzymes, 11 are predicted to be S- and Noxidoreductases, and eight are predicted to catalyze sulfur anion reduction. A final organism of note in Table 2 is S. heliotrinireducens, which was isolated from the ruminal flora of a sheep [86]. In this case, there are 27 predicted DMSOR enzymes, of which 14 are predicted to reduce Sand N-oxide substrates. Although two of these organisms


363

Fig. 9 Inverse correlation between pathogenicity and the presence of Mo/W-enzymes. Complete proteomes were selected based on the occurrence of ‘‘heavy-hitting’’ genera in our analyses of the taxonomic distribution of DMSOR-, SUOX-, XDH-, and AOR-fold enzymes. These were complemented by those selected on the basis of ‘‘heavy-hitting’’ genera amongst available genome sequences (e.g., from the data used to generate Fig. 5). Genomes of a range of wellcharacterized pathogens were also selected. Species for which proteomes were analyzed are listed in Table 2. a A chart showing that the occurrence of predicted Mo/W-enzymes is biased against strain pathogenicity. Numbers in parentheses indicate either the number of pathogenic or non-pathogenic strains, or the number of

strains with C1 predicted Mo/W-enzyme. Of the 64 predicted proteomes analyzed, 2 and 10 non-pathogens and pathogens lacked Mo/W-enzymes, respectively. b Plot of the number of predicted Mo/ W-enzymes per proteome versus species ranked by Mo/W-enzyme content. Data bars for non-pathogens are colored blue, and those for pathogens are colored red. Proteomes lacking predicted Mo/Wenzymes are indicated by the presence of an appropriately colored bar below the abscissa. c Comparison of the mean values for Mo/Wenzyme content amongst non-pathogens (18.6 ± 15.8, n = 29) and pathogens (4.3 ± 5.6, n = 35). A t test revealed a P value of 0.000054 between the non-pathogen and pathogenic data sets

were found in Crohn’s disease patients, they are also known as common gut commensals, so are unlikely to be the causative agent. However, the presence of a large number of acetylene hydratases and S- and N-oxidoreductases is not inconsistent with Crohn’s patients having an unusual intestinal chemical composition, perhaps including high concentrations of potential substrates for DMSOR enzymes. Clearly, the DMSOR-fold enzymes dominate the Mo/W-enzyme proteomes of ‘‘heavy-hitting’’ organisms. One exception is Bradyrhizobium diazoefficiens [87], a soil nitrogen fixer, whose predicted proteome encodes 19 XDH-fold proteins, six of which are predicted to be involved in purine metabolism. In the case of the SUOXfold enzymes, no organism is predicted to have [8 (the DMSOR-fold ‘‘heavy-hitter: E. lenta), and this is probably a consequence of the limited substrate specificity of this family of enzymes [2]. In the case of the AOR-fold

enzymes, two organisms have ten predicted enzymes, the marine sulfate-reducing Proteobacterium Desulfobacterium autotrophicum [45] and the syntropic thermophile Firmicute Pelotomaculum thermopropionicum [88]. We wanted to ask a critical question: what is the relationship between Mo/W-enzyme composition and pathogenicity? We addressed this question in three ways using the data of Table 2. First, we generated a chart based simply on whether or not pathogens contain predicted Mo/ W-enzymes in their proteomes (Fig. 9a). Of the species listed in Table 2, 35 are pathogens and 29 are non-pathogens. Of these, the proteomes of 25 pathogens and 27 nonpathogens contain predicted Mo/W-enzymes, respectively. Or, to put it another way, of the 64 predicted proteomes analyzed, 2 and 10 non-pathogens and pathogens lacked Mo/W-enzymes, respectively. These observations are consistent with a bias against Mo/W-enzymes within the proteomes of pathogenic bacteria. When organisms are

123

364


laboratory strain is predicted to contain a total of 18 Mo/Wenzymes, and this compares to the pathogenic strains O157:H7, O139:H21, and O126:H6, which are predicted to contain 24, 18, and 14 such enzymes, respectively. Thus, there is not a convincing difference between non-pathogenic and pathogenic E. coli based on predicted Mo/Wenzyme composition. A similar situation exists within the species of Clostridium analyzed in Table 2, the non-pathogenic species C. acetobutylicum and C. beijerinckii contain 1 and 3 predicted enzymes, respectively, whereas the pathogenic species C. tetani, C. botulinum, C. perfringens, and C. difficile contain, 2, 4, 2, and 8 such enzymes, respectively. However, when non-pathogenic and pathogenic Bacillus species are compared (subtilis and anthracis), the data are consistent with the conclusion drawn from Fig. 9, with the non-pathogenic B. subtilis and the pathogenic B. anthracis species predicted to contain 9 and 3 Mo/ W-enzymes, respectively. We also analyzed non-pathogenic and pathogenic species of Leptospira, with L. interrogans (causative agent of leptospirosis) and L. biflexa (a saprophyte) containing zero and two predicted Mo/Wenzymes, respectively. Overall, comparisons of pathogenic and non-pathogenic strains/species within the same species/genera are not as convincing as when a broader data set is compared. This is likely due to the close relationship between the pathogenic and non-pathogenic species. One final observation is that intestinal pathogens tend to have higher numbers of predicted Mo/W-enzymes than nonintestinal ones. Fig. 10 Pyranopterin coordination in the AOR-fold enzymes. a Rendering using the Pymol molecular graphics system [118] of the moieties connecting the two pyranopterins of formaldehyde:ferredoxin oxidoreductase from P. furiosus (PDB code [30]). b Summary of hydrogen-bonding interactions generated using the MarvinSketch software package (http://www.chemaxon.com). In both panels, some predicted hydrogens have been added for clarity

ranked for the predicted number of enzymes (Fig. 9b), this bias becomes more convincing, with non-pathogens dominating organisms with large numbers of predicted Mo/Wenzymes, and pathogens dominating organisms with few or zero Mo/W-enzymes. When the average number of enzymes is compared between pathogens and non-pathogens (Fig. 9c), we find a statistically significant difference between the two groups, with most pathogens having few or no Mo/W-enzymes (P = 0.000054). Thus, within our data set there is convincing evidence that pathogenicity correlates with genomes encoding few or no Mo/Wenzymes. Given the very large number of bacterial genome sequences available, we had the opportunity to compare pathogenic and non-pathogenic strains within the same genus or species. The non-pathogenic E. coli K12

123

Variability of pyranopterin coordination correlates with diversity of substrate specificity Inflexibility of AOR family pyranopterin coordination Compared to the other families of Mo/W-enzymes, the number of available sequences and protein structures for the AOR enzymes is limited (Figs. 2, 3, 4). Figure 10a summarizes the coordination of the AOR pyranopterins in formaldehyde:ferredoxin oxidoreductase from P. furiosus [30], wherein the two piperazine rings are bridged at their N-5 positions by a combination of a Mg2? atom and two water molecules, with the Mg2? also being coordinated by two backbone amide oxygens and both of the pyranopterin phosphate moieties. A similar arrangement exists in the structure of the P. furiosus aldehyde:ferredoxin oxidoreductase [31]. In both cases, the N-10 atoms of the proximal and distal pyranopterins appear to be involved in hydrogen bonds with a [4Fe–4S] cluster coordinating Cys (to its SG atom) and to the carboxylate moiety of an Asp residue, respectively. The predicted hydrogen-bonding interactions are summarized in Fig. 10b. This inflexibility of


Fig. 11 a Charge-transfer relay connecting the N-5 atom of the pyranopterin to the molybdenum atom in sulfite oxidase subfamily of the SUOX-fold enzymes. Structures from the following sources were analyzed for conservation of pyranopterin contacts: Gallus gallus (PDB code 1SOX [50]), Arabidopsis thaliana (PDB code 1OGP [119]), and Starkeya novella (2BLF [120]). In each case, a conservation of a His and Tyr was observed connecting the N-5 hydrogen of the piperazine ring to the Mo-active site. The structure around the cofactor of the G. gallus enzyme is shown. b Summary of contact residues and putative mechanism of charge transfer-mediated tetrahydropyranopterin oxidation. Note that the metal-coordinating Cys is omitted for clarity and only one Mo=O group is shown. Pyranopterin contacts were analyzed in two additional SUOX-fold proteins: nitrate reductase (NIA) from P. angusta (PDB code 2BII [58]) and YedY from E. coli (PDB code 1XDQ [60]). In the case of NIA, the His is retained but the Tyr is missing. In the case of YedY, neither residue is present with no obvious hydrogen bonds being identified involving the piperazine N-5. Panel b was generated with the MarvinSketch software package (http://www.chemaxon.com)

pyranopterin coordination contrasts with the remarkable variability seen in the DMSOR family of enzymes, which also contain a bis-pyranopterin cofactor (see below).

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Fig. 12 Inflexible pyranopterin coordination in the XDH family of Mo-enzymes. a Pymol rendering of the pyranopterin coordination environment of H. pseudoflava CO-dehydrogenase [76]. Structures from the following sources were analyzed for conservation of pyranopterin contacts: carbon monoxide dehydrogenase from H. pseudoflava (PDB code 1FFV [76]), CO-dehydrogenase from Oligotropha carboxidovorans (PDB code 1N62, [27]), 4-hydroxybenzoylCoA reductase from Thauera aromatica (PDB code 1RM6 [75]), aldehyde dehydrogenase from D. gigas (PDB code 3L4P [39]), xanthine oxidase from Bos taurus (PDB code 3NRZ [72]), xanthine oxidase from Homo sapiens (PDB code 2E1Q [71]), and quinoline-2oxidoreductase from Pseudomonas putida (PDB code 1T3Q [74]). The structure around the cofactor of the CO-dehydrogenase from H. pseudoflava is shown. The C-8 amine acts as an H-bond donor to a conserved Gln residue that is part of the [2Fe–2S] cluster binding domain/subunit of the enzyme. A second Gln acts as an H-bond donor to the pyrimidine N-9 atom via its carboxamide nitrogen and acts as an acceptor from the piperazine N-5 hydrogen via its carboxamide oxygen. A backbone carbonyl oxygen is an H-bond acceptor from the N-7 nitrogen, while a backbone amide nitrogen acts as an H-bond donor to the pyrimidine C-6 carbonyl oxygen. b Illustration of pyranopterin–protein hydrogen-bonding interactions in the XDH family. All contacts are conserved, except in the case of the N-10 hydrogen, for which no contacts were observed in any of the structures

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A charge-transfer relay connecting the pyranopterin piperazine to the Mo-active site in the SUOX-fold enzymes When we examined the pyranopterin coordination environment in the SUOX-fold enzymes, we noticed that a majority have a conserved His residue in a position able to act as a hydrogen-bonding acceptor/donor (via its ND1 nitrogen) that is in close juxtaposition to the N-5 atom of the pyranopterin piperazine ring (Fig. 11a). A conserved Tyr residue completes a putative charge-transfer relay, connecting the piperazine ring to the substrate-binding site (which is occupied by a SO42- in the structure of chicken sulfite oxidase [50]). Figure 11b illustrates how this relay could participate in redox transitions of the pyranopterin between the tetrahydro- and 10,10a-dihydro forms. Although the Tyr residue has been demonstrated to play a critical role in catalysis [89–92], we are aware of no studies addressing the role of the conserved His residue. Although our approach is non-metallocentric in this review, we speculate that the role of the piperazine–His–Tyr–Mo charge-transfer relay may be to participate in the deprotonation of the Mo4?–(OH2/OH) step in the mechanism of sulfite oxidation [5, 15]. Also shown in Fig. 11b is a conserved Lys residue (Lys301). An Arg variant at this position in the human enzyme causes sulfite oxidase deficiency [93]. This residue links the pyranopterin pyrimidine ring (via its amide nitrogen and oxygen) to the phosphate group via its NZ atom, and may play a role in maintaining the overall pyranopterin conformation in the SUOX-fold enzymes. The His residue is conserved in the plant-type nitrate reductase from Pichia angusta [58], but is hydrogen bonded to the carboxylate of an Asp residue (Asp271) in the active site. In the case of YedY, the His and Tyr residues are absent and there is no obvious protein ligand to the piperazine N-5 atom [60]. The lack of the His–Tyr motif in YedY may partly explain its lack of reactivity towards sulfite. YedY also lacks a triad of Arg residues that stabilizes anion binding to the sulfite oxidases, and this also likely contributes to its lack of sulfite reactivity [2]. Inflexible pyranopterin coordination in the XDH family We analyzed the pyranopterin coordination environment of seven structurally characterized members of the XDH family (Table 1). Figure 12a shows the pyranopterin coordination environment of carbon monoxide dehydrogenase from H. pseudoflava [76], wherein there is no protein coordination of the N-5 piperazine nitrogen, and the N-10 nitrogen is predicted to be a H-bond donor to the carboxamide oxygen of Gln692, whose carboxamide nitrogen also acts as a H-bond donor to the N-9 atom of the

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pyranopterin pyrimidine ring. Figure 12b summarizes the pyranopterin contacts for the structurally characterized members of the XDH family, demonstrating a remarkable conservation and inflexibility of coordination, with the features observed in H. pseudoflava CO-dehydrogenase being conserved across the structurally characterized members of the family. Variability in DMSOR-fold pyranopterin coordination correlates with diversity of substrate reactivity We have previously reported the effects of variants of residues close to the Mo-bisPGD in two E. coli members of the DMSOR family of enzymes: NarGHI [94] and dimethylsulfoxide reductase (DmsABC) [95–97], and in both cases we demonstrated that a conserved Arg residue located between the [4Fe–4S] cluster (FS0) and the proximal pyranopterin plays a critical role in defining the electron transfer pathway to the substrate-reactive Mo atom. A Lys residue located in a similar position in the periplasmic nitrate reductase (NapA) of Ralstonia eutropha was demonstrated to have a similar function [98]. In the case of the NarG, mutation of the Arg residue to a Ser had little effect on the reduction potentials for the Mo(VI/V) and Mo(V/IV) transitions, but significantly lowered that of FS0 (from -55 to -170 mV) [94]. These observations are consistent with residues close to the pyranopterin playing a critical role in defining catalytic efficiency. The hypothesis that pyranopterin coordination plays a role in fine-tuning Mo/W reactivity is supported by the observation of bicyclic forms of the distal pyranopterins in two enzymes, E. coli NarGHI [35, 94] and A. aromaticum ethylbenzene dehydrogenase (EbdABC) [34]. Figure 13a shows the bicyclic form of the distal pyranopterin in NarG, with possible H-bonding interactions between the pyran oxygen and Ser719 and His1163 [99], which are both conserved in the NarGHI class of DMSOR-fold enzymes. Further inspection of available structures reveals the presence of a second conserved His residue (His1184), and three structurally conserved water molecules (e.g., in PDB codes 1Q16, 1Y4Z, 1R27 [35, 100]). Figure 13b shows how this charge-transfer relay could participate in stabilizing the alkoxide open ring form of the distal pyranopterin. In the case of ethylbenzene dehydrogenase, the open pyran ring is stabilized by an Arg residue (Arg612, with both NH1 and NH2 ˚ from the pyran oxyhydrogens being approximately 2 A gen), again favoring the anionic alkoxide form of the bicyclic distal pyranopterin. With the exception of NarGHI and EbdABC, all other structurally characterized DMSOR-fold enzymes contain the Mo/W-bisPGD form of the cofactor wherein both pyranopterins have a tricyclic structure. In all the


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Fig. 13 a Role of Ser719 and His1163 in maintaining the distal pyranopterin of NarG in a bicyclic state. The image was generated with the Pymol molecular graphics package [118] using the structure of NarGHI described by PDB code 1Q16 [35]. b Charge-transfer relay connecting the open pyran alkoxide to one of three conserved waters observed in the majority of NarG structures. The image was created using the MarvinSketch software package (http://www.chemaxon.com)

structures, the N-5 atoms of the two piperazine rings are bridged by either a His or an Arg residue, with the N-5 atom of the proximal pyranopterin having an additional H-bonding interaction with a second His, a Gln or a Ser, which we refer to as the ‘‘stabilizing’’ residue (which is absent in the Thermus thermophilus polysulfide reductase) [20, 101]. The function of the ‘‘stabilizing residue is proposed to be to maintain the N-5 atom of the proximal pyranopterin in an sp3-hybridized state, thus preventing oxidation from the tetrahydro- to the 10,10a-dihydro form. In both pyranopterins, the N-10 atom is H-bonded to a backbone amide oxygen. Figure 14a illustrates the pertinent area of the E. coli FdnG subunit [102], showing the bridging Arg residue (Arg896) linking the two N-5 atoms, a ‘‘stabilizing’’ His residue (His902), and the two N-10 atoms acting as H-bonding donors to backbone amide oxygens. With the constraints outlined above, very similar arrangements exist in the other structurally characterized DMSORfold enzymes. Figure 14b provides more detail of the H-bonding interactions. Examination of Fig. 14 in the context of all 15 DMSORfold structures reveals how the bridging and stabilizing residues could function to stabilize the N-5 atom of the proximal pyranopterin in a geometry consistent with an sp3 hybridized state (with a tetrahydro proximal pyranopterin). In the case of the distal pyranopterin, there is only one interaction with the bridging His/Arg, raising the possibility that the distal pyranopterin could exist in a 10,10adihydro form in the DMSOR-fold enzymes. (although in the case of NarG, it should be noted that the piperazine ring in bicyclic distal pyranopterin already resembles the 10,10a-dihydro tricyclic form in terms of it being partially oxidized [20]). In general, the proximal pyranopterins have conformations similar to that predicted for the tetrahydro form, whereas the distal pyranopterins have conformations

similar to that predicted for the 10,10a-dihydro form [20], and this is reflected in the structure of the distal pyranopterin depicted in Fig. 14b. Figure 15 shows the combinations of bridging and stabilizing residues observed thus far in the DMSOR-fold enzyme structures. Five enzymes have a His residue at both the bridging and stabilizing positions: R. sphaeroides DMSO reductase [32, 33, 103], Shewanella massilia TMAO reductase [104], E. coli NarG [35, 100], Pelobacter acidigallici pyrogallol hydroxytransferase [105], and EbdA [34]. The periplasmic nitrate reductases and formate dehydrogenase N [102, 106–109] have an Arg bridging residue and a His stabilizing residue. Tungsten-containing formate dehydrogenase from D. gigas [110], arsenite oxidase from Alcaligenes faecalis [111], and the tungstencontaining acetylene hydratase from Pelobacter acetylenicus [112] contain an Arg bridging residue and Gln stabilizing residue. One DMSOR-fold structure, the FdhF subunit of the formate hydrogen lyase [113], has an Arg bridging residue and a Ser stabilizing residue; and the polysulfide reductase from T. thermophilus has an Arg bridging residue, but no stabilizing residue [101]. Overall, these observations reveal a diversity of pyranopterin coordination in the DMSOR-fold enzymes that is likely to play a pivotal role in their contribution to bacterial metabolic diversity and their persistence from the last universal ancestor to extant species. The role of the bridging residue can also be considered in the context of both its charge and its potential role in inter-pyranopterin communication and pyranopterin protonation. The high pKa of the guanidinium side chain of Arg in aqueous solution (*12.5) allows us to predict that it is positively charged when functioning as a bridging residue in the DMSOR-fold enzymes. The imidazole side chain of His has a pKa much closer to neutrality in aqueous

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Fig. 15 Venn diagram analysis of bridging and stabilizing residues within the structurally characterized DMSOR-fold enzymes. Histidine can function as both a bridging and stabilizing residue, and can function as stabilizing residues with an Arg bridging residue. However, Ser or Gln can only function as stabilizing residue in combination with an Arg bridging residue. Analyses were based on 15 available DMSOR-fold structures. Acronyms used: DorA, Rhodobacter Me2SO reductase; TorA, Shewanella massilia Me3NO reductase; NarG, E. coli nitrate reductase; PgtL, P. acidigallici pyrogallol hydroxytransferase; EbdA, A. aromaticum ethylbenzene dehydrogenase; NapA, periplasmic nitrate reductase (various sources); FdnG, E. col respiratory formate dehydrogenase; WFdh, tungsten-containing D. gigas formate dehydrogenase; AioA A. faecalis arsenite oxidase; Ahy, tungsten-containing P. acetylenicus acetylene hydratase; FdhF, E. coli formate dehydrogenase (formate hydrogen lyase); PsrA, T. thermophilus polysulfide reductase

Fig. 14 a Residues controlling piperazine ring coordination in E. coli formate dehydrogenase (FdnG). The image was generated using the structure of formate dehydrogenase described by PDB code 1KQF [102]. An Arg functions to bridge the two pyranopterins, with the NE nitrogen functioning as a donor to the proximal pyranopterin N-5 atom, and the NH2 nitrogen functions as a donor to the distal pyranopterin N-5: the guanidinium group is thus rotated towards the distal pyranopterin, which is shown in a 10,10a-dihydro form. The NH1 nitrogen functions as a donor to the distal pyranopterin pyrimidine keto-oxygen. b Proposed H-bonding network around the piperazine rings of the two pyranopterins. The proximal pyranopterin is shown in its tetrahydro form, with this redox state stabilized by both the conserved bridging Arg and the conserved stabilizing His. Both N-10 atoms act as H-bond donors to backbone amide oxygens. Three interactions are shown for the distal pyranopterin, the conserved Arg acts as an H-bond donor to the piperazine N-5 and also to the pyrimidine keto-oxygen, and the N-10 acts as an H-bond donor to a backbone amide oxygen

solution (*6.0), and, depending on its immediate environment [114], it may be protonated or deprotonated. Thus, in those enzymes with a bridging His residue, it is likely that the imidazole side chain functions in inter-

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pyranopterin communication and/or charge transfer. A pertinent example of this is the putative protonation of the N-5 atom of the 10,10a-dihydro form of the distal pyranopterin [20], which could facilitate an internal redox reaction resulting in a monoanionic thiol–thione chelate. By formally decreasing the net charge of the chelate from -2 to -1, it is predicted to drastically shift the Mo/W reduction potentials to markedly more positive values, thus providing an additional mechanism whereby substrate reactivity could be modulated in the DMSOR enzymes2.

Conclusions and outlook We have deliberately used a non-metallocentric approach to explore the persistence and functions of the four families of Mo/W-enzymes. For each of these, we sought to investigate the potential roles of pyranopterin coordination 2

It should be noted that similar observations could be made regarding protonation of a putative 10,10a-dihydro form of the pyranopterin in the SUOX-fold enzymes containing the chargetransfer relay depicted in Fig. 11.


and overall cofactor complexity in facilitating substrate diversity. In general, the number of substrates increases with increasing cofactor complexity provided opportunities exist for variability of pyranopterin coordination. The SUOX-fold enzymes have thus far been demonstrated to act efficiently on the anions sulfite and nitrate. Remarkably, the evolutionary-persistent prokaryotic YedY and YuiH subunits comprise over half of the known SUOX-fold sequences, and yet their physiological functions remain unknown. In the case of the XDH-fold enzymes, reactivity is known towards a range of heterocyclic compounds, aldehydes and CO, with reactivity to the latter being facilitated by Se or Cu modifications adjacent to the active site Mo. Although the Mo-bisPPT cofactor of archaeal AOR family is second in complexity only to the DMSOR family Mo/W-bisPGD cofactor, its inflexible pyranopterin coordination is consistent with its limited reactivity towards aldehydes. Finally, the combination of cofactor complexity, the ability to coordinate either Mo or W, along with a diversity of pyranopterin coordination is entirely consistent with the remarkable diversity of interconversions catalyzed by the DMSOR-fold enzymes. By carefully analyzing available enzyme structures, we have revealed the existence of His-containing chargetransfer relays in the SUOX-fold and DMSOR-fold enzymes. In the former, these may provide communication between the pyranopterin piperazine ring and the Moactive site, and in the latter they may facilitate inter-pyranopterin communication and, in the case of NarGHI, may aid in maintaining the distal pyranopterin in a bicyclic state. The importance of these relays will need to be tested by generating and characterizing appropriate variants. Future work will also include theoretical studies of how variability of pyranopterin coordination may impact Mo/W redox chemistry. We also investigated the link between the presence of predicted Mo/W-enzymes and bacterial pathogenicity, revealing a tendency within our data set of pathogens for there to be few or no such enzymes encoded by the respective genomes. The flip side of this is the identification of bacterial species that are genuine ‘‘heavy-hitters’’ in terms of Mo/W-enzyme content, notably including D. hafniense, G. pamelaeae, E. lenta, and S. heliotrinireducens. The remarkable diversity of DMSOR-fold enzymes in these species clearly needs to be further investigated. In the case of G. pamalaeae and E. lenta, there is an intriguing link between the presence of large number of DMSOR-fold enzymes and the gut flora of Crohn’s disease patients. Acknowledgments The authors would like to thank Justin Fedor and Sheng Yi Wu for helpful discussion and for suggesting aspects of the analysis presented in Fig. 9. Sheng Yi Wu is also thanked for suggesting the analysis presented in Fig. 15. This work was funded by the Canadian Institutes of Health Research (Grant MOP106550 to J.

369 H. W.). We are indebted to the remarkable computing expertise and advice of Dean Schieve.

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