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. 2020 Apr 15;5(17):9830–9838. doi: 10.1021/acsomega.9b04489

Identification and Characterization of a DmoB Flavin Oxidoreductase from a Putative Two-Component DMS C-Monooxygenase

D Scott Hammers †, Caroline M Donaghy †, Sarah L Heiss †, Lydia M Harris †, Jackson M Gordon †, John W Stevens IV †, Lucian P Murray ‡, Alexander D Schwab †, Brooke C Hester ‡, Megen A Culpepper †,*
PMCID: PMC7203701  PMID: 32391470

Abstract

graphic file with name ao9b04489_0007.jpg

The compound dimethyl sulfide (DMS) links terrestrial and oceanic sulfur with the atmosphere because of its volatility. Atmospheric DMS is responsible for cloud formation and radiation backscattering and has been implicated in climate control mitigation. The enzyme DMS C-monooxygenase degrades DMS and has been classified as a two-component FMNH2-dependent monooxygenase. This enzyme requires a flavin reductase B subunit to supply electrons to the monooxygenase A subunit where DMS conversion occurs. One form of the enzyme from Hyphomicrobium sulfonivorans has been isolated and characterized. In this work, a putative DMS C-monooxygenase has been identified with bioinformatics in Arthrobacter globiformis. We report the expression, purification, and characterization of the DmoB flavin reductase subunit, termed DmoB, from A. globiformis. Data support DmoB preference and optimal activity for the cosubstrates flavin mononucleotide (FMN) and NADH. FMN binds at a 1:1 stoichiometry with high affinity (Kd = 1.11 μM). The reductase is able to generate product with the A subunit from H. sulfonivorans expressed in Escherichia coli, albeit at a lower turnover than the natively expressed enzyme. No static protein–protein interactions were observed under the conditions tested between the two subunits. These results provide new details in the classification of enzymes involved in the sulfur cycling pathway and emerging forms of the enzyme DMS monooxygenase.

Introduction

Greenhouse gas emissions continue to rise with increasing population, deforestation, and manufacturing, resulting in climate warming. As global temperatures continue to rise, methods must be developed to both diminish and remove CO2 and other potent greenhouse gases from our atmosphere. Geoengineering methods, such as solar radiation management (SRM) and carbon dioxide reduction (CDR), are large-scale efforts to reduce the warming trends caused by greenhouse gas emissions.1 One SRM approach currently utilizes stratospheric aerosol injections, specifically sulfate aerosols, to decrease global mean temperatures.

Aerosolized sulfur is generated by volatile organic sulfur compounds (VOSCs) by both anthropogenic and biological sources. Manmade VOSCs originate from waste-water treatment plants, paper and textile industries, and fossil fuel burning, whereas biogenic sources arise from oceans, fresh waters, plants, and terrestrial systems.2 The predominate contributor to atmospheric sulfur is the chemical dimethyl sulfide (DMS), CH3–S–CH3. Because of its high vapor pressure and low boiling point, DMS readily volatilizes into the atmosphere, where it undergoes oxidation to generate sulfate aerosols.2,3 These aerosols act as cloud condensation nuclei (CCN) and attract water molecules, resulting in an increase in cloud formation and albedo.4 The increase in cloud formation leads to a decrease in solar radiation absorbed by the Earth and an overall cooling effect.4,5 The linkage between DMS emission and the Earth’s climate is explained by the CLAW hypothesis.2−4 CLAW is an acronym taken from the author’s names Charlson, Lovelock, Andreae, and Warren who formulated this model.4

Although the direct impact DMS has on the global sulfur cycle as described above is without question, the exact pathways and mechanisms of DMS formation and degradation are poorly understood. It is important to investigate these pathways to assess their environmental impact and response to environmental change and stimuli in order to properly assess a sulfate aerosol-based method as a feasible SRM approach. DMS degradation occurs primarily by the enzyme dissimilatory DMS C-monooxygenase (EC 1.14.13.131), which catalyzes the conversion of DMS to methanethiol (MT) and formaldehyde (Scheme 1); the formaldehyde is then assimilated or converted to CO2.6 In 2019, DMS monooxygenase was split into two classes; an assimilatory form used by Pseudomonas spp. and Escherichia coli to acquire sulfur from DMS and a dissimilatory form used by methylotrophs and autotrophs to utilize DMS for energy.6 A single study on the isolation and purification of DMS C-monooxygenase from Hyphomicrobium sulfonivorans has been reported.7

Scheme 1. Proposed Reaction Scheme of DMS Monooxygenase Involving a DmoB Flavin Reductase Subunit and a DmoA Monooxygenase Subunit.

Scheme 1

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DMS C-monooxygenase belongs to a diverse family of enzymes known as the two-component FMNH2-dependent monooxygenases.8,9 There are six distinct flavoprotein monooxygenase subclasses, with DMS C-monooxygenase proposed to belong to class C.10 These enzymes catalyze a wide variety of reactions ranging from Baeyer–Villiger, desulfonation, hydroxylation, epoxidation, and sulfoxidation.8,11 It is proposed that DMS monooxygenase may represent a new class of two-component FMNH2-dependent enzymes distinct from other homologous members of class C flavoprotein monooxygenases, based on its substrate specificity, potential divergent cofactor requirements, and the molecular phylogeny of its predicted amino acid sequence.7

The DMS C-monooxygenase reaction specifically is tightly coupled to a flavin reductase (DmoB) and a flavin-dependent monooxygenase (DmoA).7 DmoB is proposed to catalyze the reduction of flavin mononucleotide (FMN) to FMNH2 in the presence of a nicotinamide adenine dinucleotide (NAD(P)H) cosubstrate. The DmoA subunit is the monooxygenase where DMS substrate conversion to MT occurs. A crystal structure of the DmoA subunit from H. sulfonivorans recombinantly expressed in E. coli has been solved, though there are no structures of the native, two-subunit enzyme.12 Though only a single form of the native DMS C-monooxygenase enzyme has been isolated and purified, DMS degradation has been detected in methylotrophic and autotrophic bacteria under aerobic conditions, and NADH-dependent oxygen uptake in the presence of DMS has been observed in cell extracts of Hyphomicrobium, Arthrobacter, Pseudarthrobacter, and Thiobacillus.7,13,14

Here, we report the expression, purification, and characterization of a flavin reductase from Arthrobacter globiformis, proposed to be the B subunit of a DMS C-monooxygenase. This work demonstrates that this flavin reductase from A. globiformis has the same affinity for FMN and NADH as the native enzyme and is able to provide the electrons necessary to drive MT conversion in the A subunit from H. sulfonivorans.

Results and Discussion

Gene Identification of the Oxidoreductase DmoB

The protein sequence of the DmoA subunit from H. sulfonivorans (accession ID E9JFX9) was used to find potential candidates for characterization of DMS C-monooxygenase from alternate bacteria.7,15 Only candidates containing a putative oxidoreductase on the same operon as the monooxygenase were selected and a phylogenetic tree was generated (Figure 1).19,20 The resulting protein candidates primarily belong to the Actinobacteria and Proteobacteria classes, and provide an excellent starting point to characterize the diversity among this protein family. Based on the phylogenetic tree analysis as well as reports that cell extracts of Arthrobacter species uptake oxygen in the presence of DMS in an NADH-dependent manner, we chose to study the genes corresponding to a putative DMS C-monooxygenase from A. globiformis NBRC 12137.13 From the Arthrobacter genomes that were screened, this gene is in close proximity to the DMS C-monooxygenase from H. sulfonivorans and is the only candidate annotated as an FMNH2-dependent monooxygenase (Figure S1).

Figure 1.

Figure 1

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Phylogenetic tree of the DmoA protein sequence from H. sulfonivorans (*represented by asterisk) and the A subunit of putative two-component FMNH2-monooxygenase proteins from alternate bacteria. The protein candidate hits arise from Proteobacteria and Actinobacteria classes. A blast search (BLASTp) of the DmoA sequence was run using the joint genomics institute database against permanent and draft genomes.18 A sequence alignment was run MAFFT19 followed by the construction of a maximum likelihood tree using MEGAX.20 The phylogeny tree was tested with 100 rounds of bootstrap.

A pairwise sequence alignment between the DmoA protein sequence of H. sulfonivorans and the putative FMNH2-dependent monooxygenase gene from A. globiformis results in a 47% sequence identity (Figure S2). Examining the sequence alignment, amino acids proposed to be involved in FMN binding (F10, Q79, N133, and F245) based off the crystal structure of the DmoA subunit from H. sulfonivorans are identical in the A. globiformis sequence.12 In addition, both operons contain open reading frames (orfs) which encode for a flavin oxidoreductase (Figure 2) in close proximity to the monooxygenase gene. Additional genes in this cluster encode for a monoamine oxidase, an aldehyde dehydrogenase, a 2-oxopent-4-enoate hydratase, and a predicted fumarylacetoacetate hydrolase.

Figure 2.

Figure 2

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Proposed DMS degradation gene cluster from A. globiformis NBRC 12137. (A) Gene organization of ORFs. Orf171 corresponding to the putative dmoB gene from this work is colored blue and orf465 corresponding to the putative dmoA gene is colored red. (B) Proposed functions of ORFs in the DMS gene cluster.

DmoB Protein Expression and Purification

We set out to biochemically characterize the flavin reductase (orf171, accession ID WP_003800025.1) on the A. globiformis 12137 operon recombinantly expressed in E. coli, termed DmoB herein. Interestingly, sequence alignment between DmoB from A. globiformis has low sequence identity to the two flavin reductases encoded on the dmo operon of H. sulfonivorans [orf136, accession ID ADL39573 (24.3%) and orf176, accession ID WP_083509410 (25.9%)]. The highest sequence identity of A. globiformis DmoB is between PrnF from Pseudomonas protegens (accession ID WP_011061886) and EmoB from EDTA-degrading bacterium BNC1 (accession ID WP_011581165, both at 28%) (Figure S3). Because of its close proximity to the monooxygenase gene, we tested the purified DmoB from A. globiformis with isolated DmoA protein from H. sulfonivorans also recombinantly expressed in E. coli. Our data suggest that the DmoB flavin reductase from A. globiformis can drive DMS conversion (Figure S9).

Codon-optimized, C-terminally His6-tagged DmoB from A. globiformis expresses in BL21(DE3) E. coli and is purified to >95% at a yield of ∼60 mg of protein per liter of culture. Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) gel bands are present around 19 kDa (Figure 3A), which corresponds to the Mr = 19,385 Da based off the amino acid sequence. The native molecular weight of DmoB was determined to be 39.5 kDa as estimated by gel filtration calibrated with protein standards (Figures 3B and S4). These data are consistent with other flavin reductases in two-component FMNH2-dependent monooxygenases that also exist as dimers, such as SsuE, EmoB, and PrnF.21−23 Purified DmoB does not contain bound flavin cofactor as the UV–vis spectrum exhibits no peak at 450 nm.

Figure 3.

Figure 3

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SDS-PAGE gel and gel filtration chromatogram of the DmoB flavin reductase from A. globiformis expressed in E. coli. DmoB has an Mr of 19,385 Da based off the amino acid sequence. Gel filtration data suggest that DmoB from A. globiformis is a dimer in its native conformation (A) lanes: 1, protein molecular weight marker (Fisher Scientific); 2, E. coli lysate; 3, protein after purification on the Cobalt-Talon resin; 4, protein after purification on the Superdex 200 gel filtration column. (B) Gel filtration chromatograph of pure A. globiformis DmoB. The retention time, 78 mL, was used to determine the Kav and subsequent molecular weight of 39.5 kDa.

DmoB Flavin Reductase Enzyme Activity

Oxidation of NAD(P)H to NAD(P)+ was monitored by the decrease in absorbance at 340 nm with various flavin and pyridine nucleotide cosubstrates. DmoB from A. globiformis has preference for the substrates FMN and NADH as evidenced by their low Km and high catalytic efficiency values (Table 1 and Figure S5). We were unable to obtain accurate Vmax values in the presence of NADPH because of the saturation of the detector. This is sufficient evidence to indicate that DmoB has preference for NADH as its nicotinamide source. When alternate flavins were tested, neither FAD or riboflavin had a large effect on Vmax. The Km, however, had 5- and 41-fold increases when tested with FAD and riboflavin, respectively, compared to FMN.

Table 1. Substrate Specificities of the Flavin Oxidoreductase DmoB from A. globiformisa.

substrate second substrate Vmax (units/mg) Km (μM) kcat/Km (min–1 μM–1)
NADH FMN 149 ± 11 2.3 ± 0.5 1256
NADPH FMN N.D.b N.D.b  
FMN NADH 207 ± 18 18 ± 5 223
FAD NADH 153 ± 8 84 ± 12 35
riboflavin NADH 130 ± 4 142 ± 7 17
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a

The first substrate concentration is varied and the second substrate concentration remains constant

b

N.D. = not determined.

Flavin reductases can be classified into several classes depending on their amino acid sequence and biochemical properties. Class C flavin reductases are part of a two-component monooxygenase system encoded by separate genes that provide reducing equivalents to a flavin that is ultimately transferred to a monooxygenase subunit to drive substrate conversion upon reaction with molecular oxygen.8 Class C flavin reductases have varying degrees of substrate specificity and are not isolated with bound flavin. Comparing the flavin reductases from other two-component systems that exhibited the closest sequence identity, EmoB and PrnF, reveals that EmoB and DmoB are both specific to FMN and NADH, though DmoB is 4.5-times more efficient in its reductive chemistry.22,23 PrnF prefers binding FAD and NADH, though enzyme turnover was consistent with all nicotinamide and flavin substrates.22 The preference of DmoB in A. globiformis for FMN and NADH in optimal activity is the same as the results observed in DMS C-monooxygenase expressed natively in H. sulfonivorans.7 The natively expressed protein contains both subunits, so there is no data on the biochemical properties of the DmoB subunit independently. The observed cosubstrate specificity provides further support in our assignment that this gene cluster contains an uncharacterized DMS C-monooxygenase.

DmoB Fluorometric Flavin-Binding Affinity and Stoichiometry

To determine the binding affinity and stoichiometry of FMN to DmoB, fluorescence quenching experiments were performed (Figures 4 and S8). FMN exhibits intrinsic fluorescence emission at 540 nm when excited at 420 nm. Previous studies have reported fluorescence quenching of the flavin when bound to protein.22,24,25 Flavin quenching was reported on two instruments. The first is a commercial HORIBA fluorometer. The second instrument is a custom-built fluorometer consisting of a 488 nm laser source and a spectrometer. In each experiment, FMN (6 μM) was titrated with increasing concentrations of DmoB (0.6–48 μM) and the change in fluorescence intensity was recorded. A plot of fractional intensity changes at 510 nm as a function of the [DmoB]/[FMN] ratio is shown in Figures 2 and S8. The data were fit according to eq 1.24 A 1:1 binding stoichiometry (n = 1.10 ± 0.25) of FMN to DmoB was determined from the fit and a Kd = 1.11 ± 0.71 μM was calculated using the HORIBA fluorometer and these values are higher than reported values of PrnF (Kd = 76 nM), which prefers FAD, and EmoB (Kd = 0.42 μM), which prefers FMN.22,26 Both PrnF and EmoB bind their preferred flavin substrate at a 1:1 stoichiometry with enzymes.

Figure 4.

Figure 4

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Fluorometric titration of FMN with the DmoB enzyme. FMN (6 μM) was titrated with concentrations of the DmoB enzyme ranging from 0.6 to 48 μM (A,B). (A) Emission intensities at 540 nm were measured after excitation at 420 nm and quenching is observed with increasing DmoB enzyme concentrations. (B) Change in fluorescence intensities as a result of FMN quenching was plotted against the ratio of [DmoB]/[FMN] and fit to eq 1.

DmoB from A. globiformis Can Donate Electrons to DmoA from H. sulfonivorans for DMS Degradation

We were able to show reduction of FMN by the DmoB flavin reductase, but it is important to also determine if DMS degradation can occur when coupled with the monooxygenase subunit, DmoA. The DmoA candidate protein on the A. globiformis operon has not been expressed and purified to date. Based off the cofactor specificities and the phylogenetic analysis, we tested the DmoB from A. globiformis with a recombinantly expressed DmoA from H. sulfonivorans available in the laboratory. Codon-optimized C-terminally His6-tagged DmoA from H. sulfonivorans expresses in E. coli BL21(DE3) cells and is purified to >95% at a yield of ∼30 mg of protein per liter of culture (Figure S7). The monomeric Mr = 54,182 Da based off the amino acid sequence is observed in the SDS-PAGE gel. The native molecular weight as determined by gel filtration is 203,000 Da, indicating that the DmoA subunit from H. sulfonivorans exists as a tetramer under the conditions tested.

Product formation in the reaction of DMS C-monooxygenase is the volatile gas MT. Because of this volatility, MT is amenable to headspace gas chromatography (HS-GC). A separation method was developed to detect DMS and MT and a six-point calibration curve for MT was generated (Figure S8). In HS vials, DmoB from A. globiformis (2.5 mg/mL), DmoA from H. sulfonivorans (2.5 mg/mL), NADH (1 mM), FMN (3 μM), (NH4)Fe(SO4)2 (5 μM), and DTT (5 μM) were mixed in 20 mM PIPES, pH 7.0, to a final volume of 1 mL.7 The reaction was initiated with the addition of DMS (1 mM) and allowed to incubate at 30 °C, shaking at ∼100 rpm, for 30 min. A control reaction containing no DmoB was also prepared. MT was detected only in the reactions containing all reaction components, not the negative control (Figure S9). A kcat of 3.5 min–1 was calculated, though more rigorous kinetic trials are underway. This kcat is vastly lower than the 5.4 s–1 previously reported for native DMS C-monooxygenase in H. sulfonivorans.7 It does, however, demonstrate that alternate reductases can provide the reducing equivalents necessary for DMS conversion to MT.

Protein–Protein Interactions

Possible DmoA/DmoB protein–protein interactions were probed by fluorometric titration and gel filtration. Reports of FAD-bound PrnF titrated with its monooxygenase subunit, PrnD, observed a reversal of the FMN fluorescence quenching.22 Similar results are observed upon FAD-SsuE titrated with SsuD.27 Reversal of the FMN-DmoB (6 μM) fluorescence quenching was tested with increasing H. sulfonivorans DmoA concentrations (6–24 μM). No change in fluorescence quenching was observed under these conditions.

In addition, gel filtration studies were used to observe potential protein–protein interactions between the two DmoA and DmoB subunits. A. globiformis DmoB (5 mg/mL) and H. sulfonivorans DmoA (5 mg/mL) were mixed at a 1:1 stoichiometry and applied to a Superdex 200 size exclusion column. Two peaks were observed in the chromatographic trace (Figure 5) with molecular weights corresponding to DmoB and DmoA.

Figure 5.

Figure 5

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Gel filtration chromatogram of DmoA from H. sulfonivorans and DmoB from A. globiformis incubated at a 1:1 molar ratio. The elution profile indicates that no static protein–protein interaction is occurring between the two subunits under the conditions tested.

One explanation of the results is that unlike the two-component systems SsuD/SsuE and PrnD/PrnF, there is no static protein–protein interaction occurring in the DmoB/DmoA system. If a static interaction were observed, the protein would coelute in a single peak on the gel filtration column at a molecular weight equaling the sum of the native molecular weights of the two proteins. We would also likely see a reversal of the FMN quenching with titration of the DmoA monooxygenase as well. Reversal of the FMN quenching would indicate transfer of the flavin out of the reductase as observed in other systems.22,27

An alternate explanation of the data is that no interaction would be observed under any condition as A. globiformis and H. sulfonivorans are soil bacteria that originate from differing phyla. Further studies are needed on a homogeneous DMS C-monooxygenase system to determine with more confidence if a transient protein–protein interaction is occurring. Indeed, this theory supports the low turnover numbers observed in the activity data and may be vastly increased with the two-component system’s native binding partner.

Conclusions

In summary, we have characterized a flavin reductase protein located on a proposed dmo operon of A. globiformis that is able to donate electrons necessary to reduce DMS. Previous reports of Arthrobacter species able to grow on DMS13,14 and the phylogenetic analysis reported in this work provide support in the assignment of this operon. The recombinant A. globiformis DmoB flavin reductase has specificity for FMN and NADH substrates as reported for the natively expressed DMS C-monooxygenase.7 It binds the flavin substrate with micromolar affinity at a stoichiometry of 1:1. Lastly, preliminary studies probing protein–protein interactions between the DmoB flavin reductase from A. globiformis and the DmoA monooxygenase from H. sulfonivorans indicate that a transient interaction may be occurring as the two proteins did not coelute on the gel filtration column and the FMN quenching was not reversed when DmoA from H. sulfonivorans was titrated in, though further studies are necessary to confirm this result. Future work to recombinantly express the DmoA protein from A. globiformis is underway with the penultimate goal to characterize new forms of the enzyme DMS C-monooxygenase. Gaining mechanistic insight into this family of enzymes is important in developing a clearer understanding of the enzymes involved in global sulfur cycling.

Methods and Materials

Materials

Unless specified, chemical reagents were purchased at the highest purity from Sigma-Aldrich, Fisher Scientific, or VWR.

Gene Cluster Identification

The protein sequence of DmoA from H. sulfonivorans S1 (JGI, 2503692000, NCBI: E9JFX9) was used to search for other gene candidates against the IMG database28 (DOE Joint Genome Institute, Walnut Creek, California, USA), using the pBLAST29 (National Center for Biotechnology Information, Bethesda, Maryland, USA) algorithm, which identifies genes even in unannotated regions. Gene hits of e < 1E – 30 were manually examined for neighboring reductase partners. Gene annotation was confirmed via the Pfam domain and TIGRFAM databases.30 Multiple sequence alignments were performed on these candidates using MAFFT,19 visualized using Jalview,31 and organized using the BLOSUM62 model. Phylogenetic analysis was conducted using MEGAX construction software.20 A maximum likelihood tree was constructed using the WAG model on our amino acid substitutions and (G + I) rates among sites. A partial deletion of gaps was allowed and the branch swap filter was set to strong. The major classes of bacteria represented are from Proteobacteria and Actinobacteria. Candidates from the Arthrobacter family were chosen based off previous literature demonstrating NADH-dependent oxygen uptake in the presence of DMS.13,14A. globiformis NBRC 12137 (JGI gene id GAB13127, JGI, 2520163479, NCBI accession: WP_003800031) was chosen based off its whole genome being known.

Molecular Cloning

Gene id GAB13125 (JGI, 2520163477) from A. globiformis NBRC 12137 was synthesized and codon-optimized for E. coli expression (GenScript). The 537-bp gene was cloned into a pET21b(+) vector (Novagen) between 5′-HindIII and 3′-XhoI restriction sites, generating a C-terminal hexahistidine tag. The correct gene sequence was verified by DNA sequencing (MWG operon Eurofins).

Expression and Purification of His6-Tagged DmoB Flavin Reductase from A. globiformis

Plasmids containing the dmoB gene were transformed into chemically competent E. coli BL21(DE3) cells (Lucigen) and transferred onto an LB-Agar plate containing 100 μg/mL of ampicillin and incubated overnight at 37 °C. A single transformant was selected and inoculated into 100 mL of LB (100 μg/mL of ampicillin in all LB cultures) for ∼16 h at 37 °C with shaking at 200 rpm; 10 mL of overnight culture was used to inoculate 5 × 1 L cultures (in a 1.8 L Fernbach flask) of LB media and incubated at 37 °C with shaking at 200 rpm. At OD600 ≈ 0.5, the cells were incubated at 4 °C for at least 1 h. After 1 h, the cells were induced with isopropyl-β-d-thiogalactopyranoside (IPTG) to a final concentration of 1 mM, followed by overnight incubation at 16 °C while shaking at 220 rpm. Cells were harvested via centrifugation at 4200 rpm at 4 °C, resuspended in 50 mM Tris, 500 mM NaCl, pH 8.0 (buffer A), and stored at −20 °C until needed.

The resuspended cells were thawed at room temperature and protease inhibitors (cOmplete Tablets, Mini EDTA-free) and lysozymes (final concentration ∼100 μg/mL) were added based on the total volume. The suspension was sonicated (QSonica) at an amplitude of 40% for 10 min with cycles of 10 s on/30 s off. After sonication, the suspension was centrifuged at 16,600g at 4 °C for 1 h. The lysate was loaded on to a 5 mL Co-TALON column (GE Biosciences) at a flow rate of 1.0 mL/min previously equilibrated with 10 column volumes (CV) of buffer A, followed by a wash with 20 CVs of buffer A. The protein was eluted with 20 CVs of 50 mM Tris, 500 mM NaCl, 500 mM imidazole (buffer B) by a linear gradient increase from 0% to 100%, followed by a 5 CV wash containing 100% buffer B. Fractions were analyzed by SDS-PAGE and concentrated using an Amicon Ultra 15 10 kDa MWCO centrifugal filter.

Concentrated DmoB was further purified by gel filtration chromatography using a HiLoad 16/600 Superdex 200 pg column (GE Healthcare) at a flow rate of 1.0 mL/min previously equilibrated with 50 mM Tris, 150 mM NaCl, pH 8.0 (buffer C). The sample eluted over 1.5 CV and 1.5 mL fractions were collected and analyzed by SDS-PAGE. Fractions containing DmoB were collected, concentrated down as stated above, and the concentration was determined by Bradford Assay.32 A protein yield of ∼60 mg per L of culture was obtained for the DmoB protein.

The native Mr of DmoB was estimated by gel filtration on a 16/600 Superdex 200 pg column (GE Healthcare) equilibrated with buffer C. The column void volume was determined using Blue Dextran and calibrated with appropriate known Mr protein standards: ferritin (440,000), aldolase (158,000), conalbumin (75,000), ovalbumin (43,000), and cytochrome c (12,327).

Expression and Purification of His6-Tagged DmoA Monooxygenase A Subunit from H. sulfonivorans

The DmoA subunit from H. sulfonivorans was expressed and purified using analogous methods as with the DmoB subunit from A. globiformis with these exceptions. Upon OD600 reaching ∼0.5, IPTG was immediately added to a final concentration of 1 mM and incubated for 3 h at 37 °C. After 3 h the cells were harvested, resuspended in buffer A, and stored at −20 °C until lysis. During purification, the elution gradient ranged from 0 to 50% buffer B over 15 CV, which varies slightly from the conditions above. Concentrated DmoA was further purified by gel filtration chromatography using a HiLoad 16/600 Superdex 200 pg column (GE Healthcare) at a flow rate of 1.0 mL/min previously equilibrated with 50 mM Tris, 150 mM NaCl, pH 8.0 (buffer C). The sample eluted over 1.5 CV and 1.5 mL fractions were collected and analyzed by SDS-PAGE. The native Mr of DmoA was estimated by gel filtration as described above. The protein concentration was determined using Beer–Lambert’s law and an ε = 74,050 M–1 cm–1 calculated using ExPASY.33 The protein yield for the DmoA subunit is typically ∼30 mg per L of culture.

NAD(P)H:FMN Oxidoreductase Assay

The flavin reductase activity was determined at 25 °C from the decrease of the absorbance at 340 nm (ε340 = 6.22 mM–1 cm–1) because of the oxidation of NAD(P)H. Initial NAD(P)H concentrations were determined spectrophotometrically prior to each run using an Agilent 8454. Standard reaction conditions (typical volume 100 μL) to measure flavin kinetics consisted of 20 mM PIPES, pH 7.4, 50 μM NADH, and various concentrations of FMN (0.1–52.3 μM), FAD (7.6–300 μM), or riboflavin (15.1–200 μM) per reaction. To measure nicotinamide cofactor specificity, reaction conditions comprised 20 mM PIPES, pH 7.4, 50 μM FMN, and various concentrations of the nicotinamide substrates, NADH (4.13–174.24 μM) and NADPH (28.1–400 μM). Each reaction was initiated by the addition of DmoB enzyme (final concentration between 20 ng and 16.7 μg). Protein concentrations were varied to achieve rate linearity. One unit of activity was defined as the amount of enzyme catalyzing the oxidation of 1 μmol of NADH/min under aerobic conditions.

Fluorometric Titration of FMN with DmoB

The concentration of stock FMN was determined prior to each fluorometric measurement using ε510 = 12,200 cm–1 M–1 on a Nanodrop 2000c. Fluorescence measurements were obtained on a HORIBA fluoromax-4 spectrofluorometer using FluorEssence software. The fluorescence emission spectrum was measured at 540 nm after excitation at 420 nm; slits for both excitation and emission were set at 2 nm.

Complementary fluorescence measurements were obtained on a custom-built system incorporating an Acton SP2300 Spectrometer (Princeton Instruments), a 460 nm Modu-LaserTM Stellar Pro excitation laser, and a Zeiss Objective LD A-Plan (63x/0,65) for ensuring consistency and repeatability of measurements and to confirm that systematic error is not a factor.

Binding of FMN to DmoB was determined by spectrofluorometric titrations. Fluorescence quenching and a decrease in fluorescence intensity of the flavin is observed upon binding to DmoB. A 6 μM FMN solution in 50 mM Tris, 150 mM NaCl, pH 8.0, was titrated with increasing concentrations of DmoB (0.6–48 μM). Fluorescence quenching was monitored by the emission intensity at 540 nm. A plot of fractional fluorescence changes at 540 nm as a function of [DmoB]/[FMN] is shown in Figure 2. Those data were fit using eq 1. Each data point represents a minimum of four independent trials. Similar results from the custom-built system are shown in Figure S6.

graphic file with name ao9b04489_m001.jpg 1

[P]o—enzyme concentration (before binding), [S]o—substrate concentration (before binding).

Fluorometric titration of FMN-bound A. globiformis DmoB with H. sulfonivorans DmoA. A 6 μM FMN-found DmoB sample in 50 mM Tris, pH 8.0, 150 mM NaCl was titrated with increasing concentrations of DmoA from H. sulfonivorans (6–24 μM). Emission intensity measures were recorded at 540 nm after excitation at a wavelength of 420 nm using a HORIBA fluorimeter.

DMS C-Monooxygenase Assay

Quantification of MT and DMS was performed by HS-GC coupled to a flame ionization detector. In headspace vials, DmoB from A. globiformis (2.5 mg/mL), DmoA from H. sulfonivorans (2.5 mg/mL), NADH (1 mM), FMN (3 μM), (NH4)Fe(SO4)2 (5 μM) and DTT (5 μM) were mixed in 20 mM PIPES, pH 7.4, to a final volume of 1 mL.7 The reaction was initiated with the addition of DMS (1 mM) and allowed to incubate at 30 °C, shaking at ∼100 rpm, for 30 min.

The compounds were separated on a Poraplot Q capillary column (25 m × 0.25 m × 08 μm, Agilent) installed on an Agilent 6850 series II GC coupled to an Agilent G1888 headspace autosampler. The flow rate of helium carrier gas was maintained at 1.5 mL/min. The initial oven temperature was 50 °C, which was ramped at 45 °C/min up to 200 °C. The oven was held at 3.67 min at 200 °C, giving a total run time of 7.00 min. Injections of the headspace were performed with an autosampler utilizing a sample loop; the headspace autosampler oven was maintained at 70 °C, the sample loop was at 85 °C, and the transfer line was held at 95 °C. The vial pressurization and loop fill were held for 0.15 min, the loop equilibration for 0.10 min, and the sample injection for 1.50 min. ChemStation software was used to integrate the chromatograms. The peak areas of MT were used to calculate product formation by regression analysis based off the six-point standard calibration curve of MT (200 μM–1 mM) in 20 mM PIPES, pH 7.4 buffer (Figure S8).

Acknowledgments

The authors would like to acknowledge Dr. Jennifer Cecile for the use of the HORIBA fluorometer and data discussion and Dr. Jennifer Burris for the use of the Home-built laser source. In addition, M.A.C. would like to acknowledge Dr. Grace E. Kenney for her thoughtful discussions on genome mining and phylogenetic analysis. M.A.C. would also like to acknowledge the Department of Chemistry and Fermentation Sciences for funding. B.C.H. acknowledges the Department of Physics and Astronomy for funding. Both M.A.C. and B.C.H. acknowledge funding through the Appalachian State University RIEEE-Concert award. Support for this research was provided by a grant from the National Institute of Environmental Health Sciences, National Institutes of Health, 5RO3ES026305.

Glossary

Abbreviations

CCN

cloud condensation nuclei

CDR

carbon dioxide reduction

DMS

dimethyl sulfide

FMN

flavin mononucleotide

MT

methanethiol

SRM

solar radiation management

VOSC

volatile organic sulfur compound.

Supporting Information Available

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acsomega.9b04489.

  • Phylogenetic tree comparing DmoA from H. sulfonivorans and Arthrobacter species, protein sequence alignment of DmoA from H. sulfonivorans and putative DmoA from A. globiformis, protein sequence alignments of DmoB from A. globiformis, gel filtration calibration curve, Michaelis–Menten curves of DmoB from A. globiformis, FMN fluorometric titration with DmoB from A. globiformis, SDS-PAGE gel on DmoA from H. sulfonivorans, calibration curve of MT, and GC activity between DmoA from H. sulfonivorans and DmoB from A. globiformis (PDF)

Accession Codes

Hyphomicrobium sulfonivorans DmoA: E9JFX9, Arthrobacter globiformis DmoB: JGI, 2520163477; NCBI, WP_003800025.1.

Author Contributions

The paper was written through contributions of all the authors. All the authors have given approval to the final version of the paper.

The authors declare no competing financial interest.

Supplementary Material

ao9b04489_si_001.pdf (1.3MB, pdf)

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Supplementary Materials

ao9b04489_si_001.pdf (1.3MB, pdf)

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