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Flavin-N5-oxide intermediates in dibenzothiophene, uracil, and hexachlorobenzene catabolism
CHAPTER SEVENTEEN
Flavin-N5-oxide intermediates in dibenzothiophene, uracil, and hexachlorobenzene catabolism Sanjoy Adak, Tadhg P. Begley* Department of Chemistry, Texas A&M University, College Station, TX, United States *Corresponding author: e-mail address: [email protected]
Contents 1. Introduction 2. General protocols for enzyme overexpression and purification 3. Analytical methods 3.1 HPLC parameters 3.2 LC-MS parameters 4. Chemical methods 4.1 Synthesis of FMN-N5-oxide standard 4.2 Photochemical reduction of flavin 4.3 Reaction procedure for flavin-N5-oxide detection 5. Enzyme studies 5.1 Mechanistic studies on the DszA-catalyzed reaction 5.2 Mechanistic studies on the RutA catalyzed reaction 5.3 Mechanistic studies on the HcbA1-catalyzed reaction 5.4 General guidelines for the detection of flavin-N5-oxide intermediates Funding References Further reading
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Abstract Flavin-N5-oxide is a new intermediate in flavoenzymology. Here we describe the identification of DszA (dibenzothiophene catabolism), RutA (uracil catabolism) and HcbA1 (hexachlorobenzene catabolism) as flavin-N5-oxide-utilizing enzymes. Mechanistic analysis of these reactions suggests a model for the identification of other examples of this catalytic motif.
Methods in Enzymology, Volume 620 ISSN 0076-6879 https://doi.org/10.1016/bs.mie.2019.03.020
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2019 Elsevier Inc. All rights reserved.
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1. Introduction The long-held idea that all flavin-dependent oxygenases use flavinC4a-hydroperoxide as the oxygenating species was challenged when Bradley Moore and Bruce Palfey identified EncM on the enterocin biosynthetic pathway as a flavin-N5-oxide-utilizing-enzyme (Fig. 1) (Saleem-Batcha et al., 2018; Teufel, 2017, 2018; Teufel, Agarwal, & Moore, 2016; Teufel et al., 2013, 2015). The identification of this intermediate raised the question as to its generality in flavin-dependent oxygenases. In this chapter, we describe the identification of flavin-N5-oxide intermediates in dibenzothiophene, uracil, and hexachlorobenzene catabolism.
2. General protocols for enzyme overexpression and purification The DszA gene from Rhodococcus erythropolis was cloned into the pTHT vector (Adak & Begley, 2016) (derivative of pET28b with a TEV protease cleavage site after the N-terminal His-tag). The DszA-pTHT plasmid was transformed into electrocompetent Escherichia coli BL21 (DE3) by electroporation. A single colony was inoculated in 90 mL of LB medium containing 40 μg/mL of kanamycin. This starter culture was grown at 37 °C overnight with agitation (220 rpm). LB medium (9 L, Lennox, 20 g/L) containing 40 μg/mL of kanamycin was inoculated with 90 mL of starter culture. The cells were grown at 37 °C with shaking (220 rpm) until the culture reached an OD600 of 0.6 (3–3.5 h). After incubating the flasks
Fig. 1 The structures of relevant flavins discussed in the chapter.
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at 4 °C for 1 h, without shaking, the cultures were induced by adding isopropyl-β-D-thiogalactoside (IPTG) to a final concentration of 0.4 mM. For overproduction of the protein, the cultures were further incubated at 15 °C for 15 h with shaking (180 rpm). The cells were then harvested by centrifugation at 10,000 g for 10 min at 4 °C. Typical yields were 20–25 g cell pellet (wet weight) from the 9 L of culture. Cells from 9 L of culture were re-suspended in 30 mL of binding buffer (50 mM KH2PO4, 150 mM NaCl, 10 mM imidazole, pH 8.0) and approximately 5 mg of lysozyme was added. The cells were then lysed by five cycles of sonication on ice (each cycle consisted of a 1 s pulse followed by a 1 s delay repeated 15 times with 5 min of cooling between cycles. The cell debris was removed by centrifugation at 39,000g for 40 min at 4 °C. The clarified supernatant was loaded onto a 5 mL Ni-NTA-affinity column pre-equilibrated with binding buffer kept at 4 °C. The Ni-NTAaffinity column was then washed with 50 mL wash buffer (50 mM KH2PO4, 150 mM NaCl, 20 mM imidazole, pH 8.0). The protein was eluted from the column with elution buffer (50 mM KH2PO4, 150 mM NaCl, 200 mM imidazole, pH 8.0) at 4 °C. The fractions containing protein were pooled and concentrated to 3 mL using YM-10 Amicon centrifugal ultrafilters at 5000g. The concentrated sample was buffer exchanged into 100 mM phosphate buffer at pH 7.5 containing 100 mM NaCl and 15% glycerol using an Econo-Pac 10DG desalting column. Protein concentration was determined from its absorbance at 280 nm (A280) with an extinction coefficient calculated using the ProtParam tool of the ExPASy proteomics Server. A typical yield was 6 mg/L. The RutA gene encoded in the pCA24N overexpression plasmid (Mukherjee, Zhang, Abdelwahed, Ealick, & Begley, 2010) was transformed into Escherichia coli BL21 (DE3) by electroporation. For protein overexpression, a starter culture was grown overnight in 90 mL of LB medium containing 20 μg/mL of chloramphenicol at 37 °C. LB medium (9 L, 20 g/L) containing 20 μg/mL of chloramphenicol was inoculated with this starter culture. RutA overexpression and purification were carried out as described above for DszA. A typical yield was 20 mg/L. The HcbA1 gene from Nocardioides sp. strain PD653 was cloned into the pTHT vector. Electrocompetent Escherichia coli BL21 (DE3) were transformed with HcbA1-pTHT by electroporation. HcbA1 overexpression and purification were carried out as described above for DszA. A typical yield was 10 mg/L.
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3. Analytical methods 3.1 HPLC parameters An Agilent 1260 HPLC, equipped with a quaternary pump, was used. The system included a diode array UV–Vis detector and products were detected using absorbance at 254, 280, and 450 nm. Analysis was performed either on a ZORBAX Eclipse XDB-C18 column (15 cm 4.6 mm, 5 μm particles, Agilent Technologies) or a PFP column (150 4.6 mm, 2.6 μm particles, Kinetex). Data were processed using Agilent ChemStation ver. B.04.01. HPLC conditions for the analysis of the DszA reaction mixture (using a C18 column): A—Water B—10 mM ammonium acetate buffer, pH 6.6 C—Methanol HPLC method (flow rate 1 mL/min): 0 min—90% A 10% B, 2 min—90% A 10% B, 22 min—20% A 10% B 70% C, 27 min—20% A 10% B 70% C, 28 min—90% A 10% B, 36 min—90% A 10% B HPLC conditions for the analysis of the RutA reaction mixture (using a C18 column): A—Water B—100 mM potassium phosphate buffer, pH 6.6 C—Methanol HPLC method (flow rate 1 mL/min): 0 min—100% B, 5 min—10% A 90% B, 12 min—48% A 40% B 12% C, 14 min—50% A 30% B 20% C, 18 min—30% A 10% B 60% C, 20 min— 100% B, 25 min—100% B HPLC conditions for the analysis of the HcbA1 reaction mixture (using a C18 column): A—Water with 0.1% phosphoric acid B—Acetonitrile HPLC method (flow rate 1 mL/min): 0 min—10% A 90% B, 30 min—10% A 90% B HPLC conditions for separation of FMN and FMN-N5-oxide (using a PFP column): A—Water B—100 mM potassium phosphate buffer, pH 6.6 C—Methanol
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HPLC method (flow rate 0.4 mL/min): 0 min—90% A 10% B, 2 min—90% A 10% B, 22 min—20% A 10% B 70% C, 27 min—20% A 10% B 70% C, 28 min—90% A 10% B, 36 min—90% A 10% B
3.2 LC-MS parameters LC-ESI-TOF-MS was performed using an Agilent 1260 HPLC system equipped with a binary pump and a 1200 series diode array detector followed by a MicroToF-Q II mass spectrometer (Bruker Daltonics) using an ESI source either in negative mode or positive mode. Analysis was performed on an LC-18-T column (15 cm 3 mm, 3 μm particles, Supelco). Data were processed with DataAnalysis ver. 4.0 (Bruker Daltonics). LC conditions: A—5 mM Ammonium acetate buffer, pH 6.6 B—75% Methanol and 25% Water. LC method (for positive and negative mode on MS): 0 min—100% A, 2 min—100% A, 12 min—30% A 70% B, 17 min—30% A 70% B, 18 min—100% A, 30 min—100% A The mass spectrometer parameters: Capillary, 4500 V; end plate offset, 500 V; nebulizer gas, 3.0 bar; Dry gas, 10.0 L/min; dry gas temperature, 200 °C; funnel 1 RF, 200.0 Vpp; funnel 2 RF, 200.0 Vpp; ISCID, 0.0 eV; hexapole RF, 100 Vpp; quadrupole, Ion energy, 3.0 eV; low mass, 100 m/z; collision cell, collision energy, 8.0 eV; collision RF, 150.0 Vpp; transfer time, 80.0 μs; prepulse storage, 5.0 μs.
4. Chemical methods 4.1 Synthesis of FMN-N5-oxide standard The synthesis of FMN-N5-oxide has been previously described (Mansurova, Koay, & Gaertner, 2008; Teufel et al., 2013). One can therefore easily compare the putative flavin-N5-oxide formed in the enzymatic reaction mixture with a chemically synthesized authentic sample. The scheme for the synthesis is shown in Fig. 2. In this route, 2,3 dimethylaniline 6 is converted to riboaniline 7 which is then reacted sequentially with chlorouracil and acetic anhydride to give compound 8. Treatment of 8 with sodium nitrite in glacial acetic acid yields compound 9. Deprotection using ammonia in methanol gives riboflavin-N5-oxide 10. Finally, enzymatic phosphorylation, using riboflavin kinase, gives the desired FMN-N5-oxide 5 (Adak & Begley, 2016).
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Fig. 2 The chemical synthesis of FMN-N5-oxide.
4.2 Photochemical reduction of flavin Photoreduction of flavin is an effective way to produce reduced flavin that is free from excess reducing agent. The flavin is reduced by irradiating for 5 min under anaerobic conditions in a glove box in the presence of EDTA as the reducing agent (A21 LED white light bulb, 100-W) (Enns & Burgess, 1965; Heelis, 1982; McCormick, Koster, & Veeger, 1967).
4.3 Reaction procedure for flavin-N5-oxide detection A typical reaction procedure for the detection of flavin-N5-oxide is as follows. A solution containing FMN (500 μM) and EDTA (10 mM) is irradiated with white light (A21 LED light bulb, 100-W) for 5 min under anaerobic conditions. Once flavin is photo-reduced, substrate and enzyme are added. Typical final concentrations of reduced flavin, substrate and enzyme are 100 μM. The reaction mixture is incubated under anaerobic conditions for 30 min and then exposed to atmospheric oxygen for an additional 30 min. The reaction is quenched by heat-denaturation of the protein and the solution is filtered (10 kDa cut-off ). The samples are analyzed by HPLC (PEP column is effective for the separation of flavin and flavinN5-oxide) and LC-MS.
5. Enzyme studies 5.1 Mechanistic studies on the DszA-catalyzed reaction Sulfur dioxide, a known environment pollutant, is released during the combustion of sulfur-containing petroleum. While there are several methods
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for the catalytic desulfurization of petroleum, these are not effective for the desulfurization of compounds like dibenzothiophene, the most abundant sulfur-containing heterocycle found in crude petroleum (Soleimani, Bassi, & Margaritis, 2007). The search for a microbial desulfurization process has resulted in the identification of a dibenzothiophene catabolic pathway in Rhodococcus erythropolis (Fig. 3) (Denome, Oldfield, Nash, & Young, 1994; Denome, Olson, & Young, 1993; Gray et al., 1996; Piddington, Kovacevich, & Rambosek, 1995). The pathway involves thioether oxidation catalyzed by the DszC (Barbosa, Neves, Sousa, Ramos, & Fernandes, 2018; Gonzalez-Osorio, Luong, Jirde, Palfey, & Vey, 2016; Guan et al., 2015) flavoenzyme, oxidative C–S bond cleavage catalyzed by the DszA flavoenzyme and a final C–S bond cleavage catalyzed by DszB (Lee, Ohshiro, Matsubara, Izumi, & Tanokura, 2006). The DszA catalyzed C–S bond cleavage is a novel reaction meriting further mechanistic exploration. The current mechanism for the DszA-catalyzed reaction is shown in Fig. 4 (Adak & Begley, 2016, 2017a). In this mechanism, the flavinC4a-peroxide 4 is added to 13 to give 16; release of oxidized flavin gives hydroperoxide 17, reduction of 17 gives 18, which upon C–S bond cleavage and tautomerization gives the final product 14. This mechanism is consistent with the requirement of the reaction for molecular oxygen, and the incorporation of oxygen from molecular oxygen and not from water into the product 14. Since there are no cysteine residues in the DszA sequence, the reduction of peroxide 17 using peroxiredoxin type chemistry (Hall, Karplus, & Poole, 2009) is not possible. The possibility of flavin functioning as the reducing agent was therefore considered (Saleem-Batcha et al., 2018; Teufel, 2017, 2018; Teufel et al., 2016, 2013, 2015). Since flavin-N5-oxide 5, the
Fig. 3 The dibenzothiophene catabolic pathway in Rhodococcus erythropolis.
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Fig. 4 Mechanistic proposal for the DszA-catalyzed reaction involving flavin-N5-oxide formation.
expected flavin product, is readily converted to flavin by reducing agents, it was essential to develop reaction conditions free of reducing agent. Thus, instead of using Fre and NADH, reduced flavin was generated in the glove box by photoreduction using EDTA as the electron source. Then enzyme and substrate were added and the reaction mixture was incubated in the glove box for 30 min before exposure to oxygen to complete the reaction. Under these conditions it was possible to detect the formation of a new flavin-derived species which was identified as FMN-N5-oxide 5 based on LC-MS analysis and coelution with chemically synthesized FMN-N5oxide. When the reaction was carried out in the presence of 18O2, the predicted 2 Da increase in the FMN-N5-oxide mass was observed.
5.2 Mechanistic studies on the RutA catalyzed reaction In 2006 Sydney Kustu’s group identified an operon in Escherichia coli K12 responsible for uracil catabolism (Loh et al., 2006). In this catabolic pathway, RutA catalyzes the conversion of uracil to 3-ureidoacrylic acid 21 (Fig. 5A). Although this reaction seems like a simple amide hydrolysis, a labeling experiment showed that molecular oxygen is the source of one of the carboxylate oxygens of 21; thus, RutA catalyzes an oxidative amide bond cleavage reaction instead of an amide hydrolysis (Mukherjee et al., 2010). Our current mechanistic proposal for this reaction is shown in Fig. 5B (Adak & Begley, 2017a, 2017b). The flavin-C4a-peroxide 4 is added to uracil to give the tetrahedral intermediate 23; C–N bond cleavage and release of oxidized flavin give the peracid intermediate 25; reduction of this peracid, by flavin, gives the final product 21. The absence of any active site cysteine residue suggests that oxidized flavin reduces peracid 25 to form 21 and a
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Fig. 5 (A) The uracil catabolic pathway. (B) The mechanistic proposal for the RutAcatalyzed reaction.
flavin-N5-oxide 5. Similar strategies to those used for the DszA system were followed to characterize this putative flavin-N5-oxide. To avoid a reducing environment in which the flavin-N5-oxide is unstable, the RutA-catalyzed reaction was carried out in the presence of photo-reduced FMN. Under these conditions the formation of FMN-N5-oxide was observed and confirmed by LC-MS analysis and coelution with an authentic standard. When the reaction was carried out in the presence of 18O2, the predicted 2 Da increase in the FMN-N5-oxide mass was observed.
5.3 Mechanistic studies on the HcbA1-catalyzed reaction Hexachlorobenzene (HCB, 34) is a well-known fungicide that has been extensively used in agriculture since the 1940s (Barber, Sweetman, van Wijk, & Jones, 2005). Although HCB is banned under the Stockholm convention on persistent organic pollutants, it is still widely distributed across the globe. As part of a bioremediation program, a gene cluster, responsible for the conversion of HCB to pentachlorophenol (PCP, 26), has been identified in Nocardioides sp. strain PD653 (Ito et al., 2017, 2018). This gene cluster consists of a putative flavin monooxygenase HcbA1, and two putative flavin reductases HcbA2 and HcbA3. Experiments with cell lysate provided evidence that HcbA1 and HcbA3 together can convert HCB to PCP and we have
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demonstrated that purified HcbA1 alone can convert HCB to PCP in the presence of a heterologous flavin reductase (Fig. 7A). The best-studied flavin-dependent dehalogenases are PcpB and iodotyrosine deiodinase (Hu, Chuenchor, & Rokita, 2015; Phatarphekar, Buss, & Rokita, 2014; Sun, Su, & Rokita, 2017; Thomas, McTamney, Adler, LaRonde-LeBlanc, & Rokita, 2009) (Fig. 6A and B). PcpB catalyzes the conversion of pentachlorophenol 26 to tetrachlorobenzoquinone 28. In this reaction, a flavin hydroperoxide functions as an electrophile mediating the hydroxylation of the para position of 26. Iodotyrosine deiodinase catalyzes the conversion of 29 to 33. In this reaction, the substrate undergoes a keto-enol tautomerization, followed by a single electron transfer from the reduced flavin. Loss of iodide followed by reduction of the resulting phenoxy radical completes the reaction. The current mechanism for the HcbA1-catalyzed reaction is shown in Fig. 7B. In this mechanism, the flavin peroxide is added to the benzene ring of 34 to give the Meisenheimer intermediate 35. Elimination of chloride to give 36 followed by elimination of flavin gives peroxide 37. Flavin-mediated peroxide reduction gives the flavin-N5-oxide 5 and the product 26. This mechanism is consistent with the requirement of the reaction for molecular oxygen, and the incorporation of oxygen from molecular oxygen and not from water into the product 26. A similar strategy to that used for DszA and RutA was used to test for flavin-N5-oxide 5 formation. Under anaerobic conditions, stoichiometric amounts of 34, HcbA1 and FMNH2 were mixed, incubated for 30 min and then exposed to atmospheric oxygen for flavin-C4a-peroxide
Fig. 6 Flavoenzyme catalyzed dehalogenation reactions: (A) PcpB catalyzes the conversion of pentachlorophenol 26 to tetrachlorobenzoquinone 28. (B) The reaction catalyzed by iodotyrosine deiodinase.
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Fig. 7 (A) The reaction catalyzed by HcbA1. (B) Mechanistic proposal for the HcbA1catalyzed reaction involving flavin-N5-oxide 5 formation.
4 formation and completion of the reaction. LC-MS analysis of the resulting reaction mixture demonstrated the formation of the flavin-N5-oxide 5. When the reaction was carried out in the presence of 18O2, the predicted 2 Da increase in its mass was also observed (Adak & Begley, 2019).
5.4 General guidelines for the detection of flavin-N5-oxide intermediates 1. Flavin-N5-oxide 5 is readily reduced to flavin. It is therefore essential to exclude reducing agents such as DTT, TCEP and reduced nicotinamide from the reaction mixture. Reduced flavin is prepared by photoreduction in the presence of EDTA rather than by enzymatic reduction using NAD(P)H as the reducing agent. 2. Although the UV–Visible spectra of oxidized flavin 1 and flavin-N5oxide 5 are similar, there is a clear distinction between the two species in the 450 nm region (λmax of flavin is 447 nm whereas λmax of flavin-N5oxide is 462 nm). This can be used as an initial indicator of the formation of a flavin-N5-oxide 5 intermediate. 3. Flavin-N5-oxide 5 can be detected by HPLC analysis of the enzymatic reaction mixture (single turnover) using synthesized FMN-N5-oxide 5 as an authentic standard.
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Fig. 8 Proposal for the use of flavin-N5-oxide intermediates by flavoenzymes catalyzing substrate hydroperoxide formation in the absence of an active site cysteine.
4. As of yet, protein amino acid sequence analysis has not generated a characteristic sequence motif that enables the identification of flavin-N5oxide-forming enzymes. However, mechanistic analysis has proven useful in the identification of such enzymes. The analysis of the DszA reaction mechanism suggested that flavin-N5-oxides are likely intermediates in any flavoenzyme involving substrate hydroperoxide formation in the absence of an active site cysteine (Fig. 8) (Hall et al., 2009). This model led to the identification of RutA and HcbA1 as possible FMN-N5-oxide generating enzymes, but does not include the catalytic motif shown in the EncM-catalyzed reaction. It is highly likely that many additional examples of flavin-N5-oxide-utilizing-flavoenzymes remain to be discovered.
Funding This research was supported by the Robert A. Welch Foundation Grant A-0034.
References Adak, S., & Begley, T. P. (2016). Dibenzothiophene catabolism proceeds via a flavin-N5oxide intermediate. Journal of the American Chemical Society, 138(20), 6424–6426. Adak, S., & Begley, T. P. (2017a). Flavin-N5-oxide: A new, catalytic motif in flavoenzymology. Archives of Biochemistry and Biophysics, 632, 4–10. Adak, S., & Begley, T. P. (2017b). RutA-catalyzed oxidative cleavage of the uracil amide involves formation of a flavin-N5-oxide. Biochemistry, 56(29), 3708–3709. Adak, S., & Begley, T. P. (2019). Hexachlorobenzene catabolism involves a nucleophilic aromatic substitution and flavin-N5-oxide formation. Biochemistry, 58(9), 1181–1183. Barber, J. L., Sweetman, A. J., van Wijk, D., & Jones, K. C. (2005). Hexachlorobenzene in the global environment: Emissions, levels, distribution, trends and processes. Science of the Total Environment, 349(1–3), 1–44. Barbosa, A. C. C., Neves, R. P. P., Sousa, S. F., Ramos, M. J., & Fernandes, P. A. (2018). Mechanistic studies of a flavin monooxygenase: Sulfur oxidation of dibenzothiophenes by DszC. ACS Catalysis, 8(10), 9298–9311. Denome, S. A., Oldfield, C., Nash, L. J., & Young, K. D. (1994). Characterization of the desulfurization genes from Rhodococcus sp. strain IGTS8. Journal of Bacteriology, 176(21), 6707–6716.
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Denome, S. A., Olson, E. S., & Young, K. D. (1993). Identification and cloning of genes involved in specific desulfurization of dibenzothiophene by Rhodococcus sp. strain IGTS8. Applied and Environmental Microbiology, 59(9), 2837–2843. Enns, K., & Burgess, W. H. (1965). The photochemical oxidation of ethylenediaminetetraacetic acid and methionine by riboflavine. Journal of the American Chemical Society, 87(24), 5766–5770. Gonzalez-Osorio, L., Luong, K., Jirde, S., Palfey, B. A., & Vey, J. L. (2016). Initial investigations of C4a-(hydro)peroxyflavin intermediate formation by dibenzothiophene monooxygenase. Biochemical and Biophysical Research Communications, 481(1–2), 189–194. Gray, K. A., Pogrebinsky, O. S., Mrachko, G. T., Xi, L., Monticello, D. J., & Squires, C. H. (1996). Molecular mechanisms of biocatalytic desulfurization of fossil fuels. Nature Biotechnology, 14(13), 1705–1709. Guan, L.-J., Lee, W. C., Wang, S., Ohshiro, T., Izumi, Y., Ohtsuka, J., et al. (2015). Crystal structures of apo-DszC and FMN-bound DszC from Rhodococcus erythropolis D-1. The FEBS Journal, 282(16), 3126–3135. Hall, A., Karplus, P. A., & Poole, L. B. (2009). Typical 2-Cys peroxiredoxins—Structures, mechanisms and functions. The FEBS Journal, 276(9), 2469–2477. Heelis, P. F. (1982). The photophysical and photochemical properties of flavins (isoalloxazines). Chemical Society Reviews, 11(1), 15–39. Hu, J., Chuenchor, W., & Rokita, S. E. (2015). A switch between one- and two-electron chemistry of the human flavoprotein iodotyrosine deiodinase is controlled by substrate. The Journal of Biological Chemistry, 290(1), 590–600. Ito, K., Takagi, K., Iwasaki, A., Tanaka, N., Kanesaki, Y., Martin-Laurent, F., et al. (2017). Identification of the hcb gene operon involved in catalyzing aerobic hexachlorobenzene dechlorination in Nocardioides sp. strain PD653. Applied and Environmental Microbiology, 83(19), e00824-17/1-e00824-17/17. Ito, K., Takagi, K., Matsushima, Y., Igimi, S., Ito, K., Takagi, K., et al. (2018). Identification of the novel hcbB operon catalyzing the dechlorination of pentachlorophenol in the Gram-positive bacterium Nocardioides sp. strain PD653. Journal of Pesticide Science, 43(2), 124–131. Lee, W. C., Ohshiro, T., Matsubara, T., Izumi, Y., & Tanokura, M. (2006). Crystal structure and desulfurization mechanism of 2’-hydroxybiphenyl-2-sulfinic acid desulfinase. The Journal of Biological Chemistry, 281(43), 32534–32539. Loh, K. D., Gyaneshwar, P., Papadimitriou, E. M., Fong, R., Kim, K.-S., Parales, R., et al. (2006). A previously undescribed pathway for pyrimidine catabolism. Proceedings of the National Academy of Sciences of the United States of America, 103(13), 5114–5119. Mansurova, M., Koay, M. S., & Gaertner, W. (2008). Synthesis and electrochemical properties of structurally modified flavin compounds. European Journal of Organic Chemistry, (32), 5401–5406. McCormick, D. B., Koster, J. F., & Veeger, C. (1967). Mechanisms of photochemical reductions of FAD and FAD-dependent flavoproteins. European Journal of Biochemistry, 2(4), 387–391. Mukherjee, T., Zhang, Y., Abdelwahed, S., Ealick, S. E., & Begley, T. P. (2010). Catalysis of a flavoenzyme-mediated amide hydrolysis. Journal of the American Chemical Society, 132(16), 5550–5551. Phatarphekar, A., Buss, J. M., & Rokita, S. E. (2014). Iodotyrosine deiodinase: A unique flavoprotein present in organisms of diverse phyla. Molecular BioSystems, 10(1), 86–92. Piddington, C. S., Kovacevich, B. R., & Rambosek, J. (1995). Sequence and molecular characterization of a DNA region encoding the dibenzothiophene desulfurization operon on Rhodococcus sp. strain IGTS8. Applied and Environmental Microbiology, 61(2), 468–475.
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Saleem-Batcha, R., Stull, F., Sanders, J. N., Moore, B. S., Palfey, B. A., Houk, K. N., et al. (2018). Enzymatic control of dioxygen binding and functionalization of the flavin cofactor. Proceedings of the National Academy of Sciences of the United States of America, 115(19), 4909–4914. Soleimani, M., Bassi, A., & Margaritis, A. (2007). Biodesulfurization of refractory organic sulfur compounds in fossil fuels. Biotechnology Advances, 25(6), 570–596. Sun, Z., Su, Q., & Rokita, S. E. (2017). The distribution and mechanism of iodotyrosine deiodinase defied expectations. Archives of Biochemistry and Biophysics, 632, 77–87. Teufel, R. (2017). Flavin-catalyzed redox tailoring reactions in natural product biosynthesis. Archives of Biochemistry and Biophysics, 632, 20–27. Teufel, R. (2018). Preparation and characterization of the Favorskiiase flavoprotein EncM and its distinctive flavin-N5-oxide cofactor. Methods in Enzymology, 604, 523–540. Teufel, R., Agarwal, V., & Moore, B. S. (2016). Unusual flavoenzyme catalysis in marine bacteria. Current Opinion in Chemical Biology, 31, 31–39. Teufel, R., Miyanaga, A., Michaudel, Q., Stull, F., Louie, G., Noel, J. P., et al. (2013). Flavin-mediated dual oxidation controls an enzymatic Favorskii-type rearrangement. Nature, 503(7477), 552–556. Teufel, R., Stull, F., Meehan, M. J., Michaudel, Q., Dorrestein, P. C., Palfey, B., et al. (2015). Biochemical establishment and characterization of EncM’s flavin-N5-oxide cofactor. Journal of the American Chemical Society, 137(25), 8078–8085. Thomas, S. R., McTamney, P. M., Adler, J. M., LaRonde-LeBlanc, N., & Rokita, S. E. (2009). Crystal structure of iodotyrosine deiodinase, a novel flavoprotein responsible for iodide salvage in thyroid glands. The Journal of Biological Chemistry, 284(29), 19659–19667.
Further reading Agarwal, V., Miles, Z. D., Winter, J. M., Eustaquio, A. S., El Gamal, A. A., & Moore, B. S. (2017). Enzymatic halogenation and dehalogenation reactions: Pervasive and mechanistically diverse. Chemical Reviews, 117(8), 5619–5674. Crawford, R. L., Jung, C. M., & Strap, J. L. (2007). The recent evolution of pentachlorophenol (PCP)-4-monooxygenase (PcpB) and associated pathways for bacterial degradation of PCP. Biodegradation, 18(5), 525–539. Hlouchova, K., Rudolph, J., Pietari, J. M. H., Behlen, L. S., & Copley, S. D. (2012). Pentachlorophenol hydroxylase, a poorly functioning enzyme required for degradation of pentachlorophenol by Sphingobium chlorophenolicum. Biochemistry, 51(18), 3848–3860.
Flavin-N5-oxide intermediates in dibenzothiophene, uracil, and hexachlorobenzene catabolism Sanjoy Adak, Tadhg P. Begley* Department of Chemistry, Texas A&M University, College Station, TX, United States *Corresponding author: e-mail address: [email protected]
Contents 1. Introduction 2. General protocols for enzyme overexpression and purification 3. Analytical methods 3.1 HPLC parameters 3.2 LC-MS parameters 4. Chemical methods 4.1 Synthesis of FMN-N5-oxide standard 4.2 Photochemical reduction of flavin 4.3 Reaction procedure for flavin-N5-oxide detection 5. Enzyme studies 5.1 Mechanistic studies on the DszA-catalyzed reaction 5.2 Mechanistic studies on the RutA catalyzed reaction 5.3 Mechanistic studies on the HcbA1-catalyzed reaction 5.4 General guidelines for the detection of flavin-N5-oxide intermediates Funding References Further reading
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Abstract Flavin-N5-oxide is a new intermediate in flavoenzymology. Here we describe the identification of DszA (dibenzothiophene catabolism), RutA (uracil catabolism) and HcbA1 (hexachlorobenzene catabolism) as flavin-N5-oxide-utilizing enzymes. Mechanistic analysis of these reactions suggests a model for the identification of other examples of this catalytic motif.
Methods in Enzymology, Volume 620 ISSN 0076-6879 https://doi.org/10.1016/bs.mie.2019.03.020
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2019 Elsevier Inc. All rights reserved.
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1. Introduction The long-held idea that all flavin-dependent oxygenases use flavinC4a-hydroperoxide as the oxygenating species was challenged when Bradley Moore and Bruce Palfey identified EncM on the enterocin biosynthetic pathway as a flavin-N5-oxide-utilizing-enzyme (Fig. 1) (Saleem-Batcha et al., 2018; Teufel, 2017, 2018; Teufel, Agarwal, & Moore, 2016; Teufel et al., 2013, 2015). The identification of this intermediate raised the question as to its generality in flavin-dependent oxygenases. In this chapter, we describe the identification of flavin-N5-oxide intermediates in dibenzothiophene, uracil, and hexachlorobenzene catabolism.
2. General protocols for enzyme overexpression and purification The DszA gene from Rhodococcus erythropolis was cloned into the pTHT vector (Adak & Begley, 2016) (derivative of pET28b with a TEV protease cleavage site after the N-terminal His-tag). The DszA-pTHT plasmid was transformed into electrocompetent Escherichia coli BL21 (DE3) by electroporation. A single colony was inoculated in 90 mL of LB medium containing 40 μg/mL of kanamycin. This starter culture was grown at 37 °C overnight with agitation (220 rpm). LB medium (9 L, Lennox, 20 g/L) containing 40 μg/mL of kanamycin was inoculated with 90 mL of starter culture. The cells were grown at 37 °C with shaking (220 rpm) until the culture reached an OD600 of 0.6 (3–3.5 h). After incubating the flasks
Fig. 1 The structures of relevant flavins discussed in the chapter.
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at 4 °C for 1 h, without shaking, the cultures were induced by adding isopropyl-β-D-thiogalactoside (IPTG) to a final concentration of 0.4 mM. For overproduction of the protein, the cultures were further incubated at 15 °C for 15 h with shaking (180 rpm). The cells were then harvested by centrifugation at 10,000 g for 10 min at 4 °C. Typical yields were 20–25 g cell pellet (wet weight) from the 9 L of culture. Cells from 9 L of culture were re-suspended in 30 mL of binding buffer (50 mM KH2PO4, 150 mM NaCl, 10 mM imidazole, pH 8.0) and approximately 5 mg of lysozyme was added. The cells were then lysed by five cycles of sonication on ice (each cycle consisted of a 1 s pulse followed by a 1 s delay repeated 15 times with 5 min of cooling between cycles. The cell debris was removed by centrifugation at 39,000g for 40 min at 4 °C. The clarified supernatant was loaded onto a 5 mL Ni-NTA-affinity column pre-equilibrated with binding buffer kept at 4 °C. The Ni-NTAaffinity column was then washed with 50 mL wash buffer (50 mM KH2PO4, 150 mM NaCl, 20 mM imidazole, pH 8.0). The protein was eluted from the column with elution buffer (50 mM KH2PO4, 150 mM NaCl, 200 mM imidazole, pH 8.0) at 4 °C. The fractions containing protein were pooled and concentrated to 3 mL using YM-10 Amicon centrifugal ultrafilters at 5000g. The concentrated sample was buffer exchanged into 100 mM phosphate buffer at pH 7.5 containing 100 mM NaCl and 15% glycerol using an Econo-Pac 10DG desalting column. Protein concentration was determined from its absorbance at 280 nm (A280) with an extinction coefficient calculated using the ProtParam tool of the ExPASy proteomics Server. A typical yield was 6 mg/L. The RutA gene encoded in the pCA24N overexpression plasmid (Mukherjee, Zhang, Abdelwahed, Ealick, & Begley, 2010) was transformed into Escherichia coli BL21 (DE3) by electroporation. For protein overexpression, a starter culture was grown overnight in 90 mL of LB medium containing 20 μg/mL of chloramphenicol at 37 °C. LB medium (9 L, 20 g/L) containing 20 μg/mL of chloramphenicol was inoculated with this starter culture. RutA overexpression and purification were carried out as described above for DszA. A typical yield was 20 mg/L. The HcbA1 gene from Nocardioides sp. strain PD653 was cloned into the pTHT vector. Electrocompetent Escherichia coli BL21 (DE3) were transformed with HcbA1-pTHT by electroporation. HcbA1 overexpression and purification were carried out as described above for DszA. A typical yield was 10 mg/L.
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3. Analytical methods 3.1 HPLC parameters An Agilent 1260 HPLC, equipped with a quaternary pump, was used. The system included a diode array UV–Vis detector and products were detected using absorbance at 254, 280, and 450 nm. Analysis was performed either on a ZORBAX Eclipse XDB-C18 column (15 cm 4.6 mm, 5 μm particles, Agilent Technologies) or a PFP column (150 4.6 mm, 2.6 μm particles, Kinetex). Data were processed using Agilent ChemStation ver. B.04.01. HPLC conditions for the analysis of the DszA reaction mixture (using a C18 column): A—Water B—10 mM ammonium acetate buffer, pH 6.6 C—Methanol HPLC method (flow rate 1 mL/min): 0 min—90% A 10% B, 2 min—90% A 10% B, 22 min—20% A 10% B 70% C, 27 min—20% A 10% B 70% C, 28 min—90% A 10% B, 36 min—90% A 10% B HPLC conditions for the analysis of the RutA reaction mixture (using a C18 column): A—Water B—100 mM potassium phosphate buffer, pH 6.6 C—Methanol HPLC method (flow rate 1 mL/min): 0 min—100% B, 5 min—10% A 90% B, 12 min—48% A 40% B 12% C, 14 min—50% A 30% B 20% C, 18 min—30% A 10% B 60% C, 20 min— 100% B, 25 min—100% B HPLC conditions for the analysis of the HcbA1 reaction mixture (using a C18 column): A—Water with 0.1% phosphoric acid B—Acetonitrile HPLC method (flow rate 1 mL/min): 0 min—10% A 90% B, 30 min—10% A 90% B HPLC conditions for separation of FMN and FMN-N5-oxide (using a PFP column): A—Water B—100 mM potassium phosphate buffer, pH 6.6 C—Methanol
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HPLC method (flow rate 0.4 mL/min): 0 min—90% A 10% B, 2 min—90% A 10% B, 22 min—20% A 10% B 70% C, 27 min—20% A 10% B 70% C, 28 min—90% A 10% B, 36 min—90% A 10% B
3.2 LC-MS parameters LC-ESI-TOF-MS was performed using an Agilent 1260 HPLC system equipped with a binary pump and a 1200 series diode array detector followed by a MicroToF-Q II mass spectrometer (Bruker Daltonics) using an ESI source either in negative mode or positive mode. Analysis was performed on an LC-18-T column (15 cm 3 mm, 3 μm particles, Supelco). Data were processed with DataAnalysis ver. 4.0 (Bruker Daltonics). LC conditions: A—5 mM Ammonium acetate buffer, pH 6.6 B—75% Methanol and 25% Water. LC method (for positive and negative mode on MS): 0 min—100% A, 2 min—100% A, 12 min—30% A 70% B, 17 min—30% A 70% B, 18 min—100% A, 30 min—100% A The mass spectrometer parameters: Capillary, 4500 V; end plate offset, 500 V; nebulizer gas, 3.0 bar; Dry gas, 10.0 L/min; dry gas temperature, 200 °C; funnel 1 RF, 200.0 Vpp; funnel 2 RF, 200.0 Vpp; ISCID, 0.0 eV; hexapole RF, 100 Vpp; quadrupole, Ion energy, 3.0 eV; low mass, 100 m/z; collision cell, collision energy, 8.0 eV; collision RF, 150.0 Vpp; transfer time, 80.0 μs; prepulse storage, 5.0 μs.
4. Chemical methods 4.1 Synthesis of FMN-N5-oxide standard The synthesis of FMN-N5-oxide has been previously described (Mansurova, Koay, & Gaertner, 2008; Teufel et al., 2013). One can therefore easily compare the putative flavin-N5-oxide formed in the enzymatic reaction mixture with a chemically synthesized authentic sample. The scheme for the synthesis is shown in Fig. 2. In this route, 2,3 dimethylaniline 6 is converted to riboaniline 7 which is then reacted sequentially with chlorouracil and acetic anhydride to give compound 8. Treatment of 8 with sodium nitrite in glacial acetic acid yields compound 9. Deprotection using ammonia in methanol gives riboflavin-N5-oxide 10. Finally, enzymatic phosphorylation, using riboflavin kinase, gives the desired FMN-N5-oxide 5 (Adak & Begley, 2016).
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Fig. 2 The chemical synthesis of FMN-N5-oxide.
4.2 Photochemical reduction of flavin Photoreduction of flavin is an effective way to produce reduced flavin that is free from excess reducing agent. The flavin is reduced by irradiating for 5 min under anaerobic conditions in a glove box in the presence of EDTA as the reducing agent (A21 LED white light bulb, 100-W) (Enns & Burgess, 1965; Heelis, 1982; McCormick, Koster, & Veeger, 1967).
4.3 Reaction procedure for flavin-N5-oxide detection A typical reaction procedure for the detection of flavin-N5-oxide is as follows. A solution containing FMN (500 μM) and EDTA (10 mM) is irradiated with white light (A21 LED light bulb, 100-W) for 5 min under anaerobic conditions. Once flavin is photo-reduced, substrate and enzyme are added. Typical final concentrations of reduced flavin, substrate and enzyme are 100 μM. The reaction mixture is incubated under anaerobic conditions for 30 min and then exposed to atmospheric oxygen for an additional 30 min. The reaction is quenched by heat-denaturation of the protein and the solution is filtered (10 kDa cut-off ). The samples are analyzed by HPLC (PEP column is effective for the separation of flavin and flavinN5-oxide) and LC-MS.
5. Enzyme studies 5.1 Mechanistic studies on the DszA-catalyzed reaction Sulfur dioxide, a known environment pollutant, is released during the combustion of sulfur-containing petroleum. While there are several methods
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for the catalytic desulfurization of petroleum, these are not effective for the desulfurization of compounds like dibenzothiophene, the most abundant sulfur-containing heterocycle found in crude petroleum (Soleimani, Bassi, & Margaritis, 2007). The search for a microbial desulfurization process has resulted in the identification of a dibenzothiophene catabolic pathway in Rhodococcus erythropolis (Fig. 3) (Denome, Oldfield, Nash, & Young, 1994; Denome, Olson, & Young, 1993; Gray et al., 1996; Piddington, Kovacevich, & Rambosek, 1995). The pathway involves thioether oxidation catalyzed by the DszC (Barbosa, Neves, Sousa, Ramos, & Fernandes, 2018; Gonzalez-Osorio, Luong, Jirde, Palfey, & Vey, 2016; Guan et al., 2015) flavoenzyme, oxidative C–S bond cleavage catalyzed by the DszA flavoenzyme and a final C–S bond cleavage catalyzed by DszB (Lee, Ohshiro, Matsubara, Izumi, & Tanokura, 2006). The DszA catalyzed C–S bond cleavage is a novel reaction meriting further mechanistic exploration. The current mechanism for the DszA-catalyzed reaction is shown in Fig. 4 (Adak & Begley, 2016, 2017a). In this mechanism, the flavinC4a-peroxide 4 is added to 13 to give 16; release of oxidized flavin gives hydroperoxide 17, reduction of 17 gives 18, which upon C–S bond cleavage and tautomerization gives the final product 14. This mechanism is consistent with the requirement of the reaction for molecular oxygen, and the incorporation of oxygen from molecular oxygen and not from water into the product 14. Since there are no cysteine residues in the DszA sequence, the reduction of peroxide 17 using peroxiredoxin type chemistry (Hall, Karplus, & Poole, 2009) is not possible. The possibility of flavin functioning as the reducing agent was therefore considered (Saleem-Batcha et al., 2018; Teufel, 2017, 2018; Teufel et al., 2016, 2013, 2015). Since flavin-N5-oxide 5, the
Fig. 3 The dibenzothiophene catabolic pathway in Rhodococcus erythropolis.
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Fig. 4 Mechanistic proposal for the DszA-catalyzed reaction involving flavin-N5-oxide formation.
expected flavin product, is readily converted to flavin by reducing agents, it was essential to develop reaction conditions free of reducing agent. Thus, instead of using Fre and NADH, reduced flavin was generated in the glove box by photoreduction using EDTA as the electron source. Then enzyme and substrate were added and the reaction mixture was incubated in the glove box for 30 min before exposure to oxygen to complete the reaction. Under these conditions it was possible to detect the formation of a new flavin-derived species which was identified as FMN-N5-oxide 5 based on LC-MS analysis and coelution with chemically synthesized FMN-N5oxide. When the reaction was carried out in the presence of 18O2, the predicted 2 Da increase in the FMN-N5-oxide mass was observed.
5.2 Mechanistic studies on the RutA catalyzed reaction In 2006 Sydney Kustu’s group identified an operon in Escherichia coli K12 responsible for uracil catabolism (Loh et al., 2006). In this catabolic pathway, RutA catalyzes the conversion of uracil to 3-ureidoacrylic acid 21 (Fig. 5A). Although this reaction seems like a simple amide hydrolysis, a labeling experiment showed that molecular oxygen is the source of one of the carboxylate oxygens of 21; thus, RutA catalyzes an oxidative amide bond cleavage reaction instead of an amide hydrolysis (Mukherjee et al., 2010). Our current mechanistic proposal for this reaction is shown in Fig. 5B (Adak & Begley, 2017a, 2017b). The flavin-C4a-peroxide 4 is added to uracil to give the tetrahedral intermediate 23; C–N bond cleavage and release of oxidized flavin give the peracid intermediate 25; reduction of this peracid, by flavin, gives the final product 21. The absence of any active site cysteine residue suggests that oxidized flavin reduces peracid 25 to form 21 and a
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Fig. 5 (A) The uracil catabolic pathway. (B) The mechanistic proposal for the RutAcatalyzed reaction.
flavin-N5-oxide 5. Similar strategies to those used for the DszA system were followed to characterize this putative flavin-N5-oxide. To avoid a reducing environment in which the flavin-N5-oxide is unstable, the RutA-catalyzed reaction was carried out in the presence of photo-reduced FMN. Under these conditions the formation of FMN-N5-oxide was observed and confirmed by LC-MS analysis and coelution with an authentic standard. When the reaction was carried out in the presence of 18O2, the predicted 2 Da increase in the FMN-N5-oxide mass was observed.
5.3 Mechanistic studies on the HcbA1-catalyzed reaction Hexachlorobenzene (HCB, 34) is a well-known fungicide that has been extensively used in agriculture since the 1940s (Barber, Sweetman, van Wijk, & Jones, 2005). Although HCB is banned under the Stockholm convention on persistent organic pollutants, it is still widely distributed across the globe. As part of a bioremediation program, a gene cluster, responsible for the conversion of HCB to pentachlorophenol (PCP, 26), has been identified in Nocardioides sp. strain PD653 (Ito et al., 2017, 2018). This gene cluster consists of a putative flavin monooxygenase HcbA1, and two putative flavin reductases HcbA2 and HcbA3. Experiments with cell lysate provided evidence that HcbA1 and HcbA3 together can convert HCB to PCP and we have
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demonstrated that purified HcbA1 alone can convert HCB to PCP in the presence of a heterologous flavin reductase (Fig. 7A). The best-studied flavin-dependent dehalogenases are PcpB and iodotyrosine deiodinase (Hu, Chuenchor, & Rokita, 2015; Phatarphekar, Buss, & Rokita, 2014; Sun, Su, & Rokita, 2017; Thomas, McTamney, Adler, LaRonde-LeBlanc, & Rokita, 2009) (Fig. 6A and B). PcpB catalyzes the conversion of pentachlorophenol 26 to tetrachlorobenzoquinone 28. In this reaction, a flavin hydroperoxide functions as an electrophile mediating the hydroxylation of the para position of 26. Iodotyrosine deiodinase catalyzes the conversion of 29 to 33. In this reaction, the substrate undergoes a keto-enol tautomerization, followed by a single electron transfer from the reduced flavin. Loss of iodide followed by reduction of the resulting phenoxy radical completes the reaction. The current mechanism for the HcbA1-catalyzed reaction is shown in Fig. 7B. In this mechanism, the flavin peroxide is added to the benzene ring of 34 to give the Meisenheimer intermediate 35. Elimination of chloride to give 36 followed by elimination of flavin gives peroxide 37. Flavin-mediated peroxide reduction gives the flavin-N5-oxide 5 and the product 26. This mechanism is consistent with the requirement of the reaction for molecular oxygen, and the incorporation of oxygen from molecular oxygen and not from water into the product 26. A similar strategy to that used for DszA and RutA was used to test for flavin-N5-oxide 5 formation. Under anaerobic conditions, stoichiometric amounts of 34, HcbA1 and FMNH2 were mixed, incubated for 30 min and then exposed to atmospheric oxygen for flavin-C4a-peroxide
Fig. 6 Flavoenzyme catalyzed dehalogenation reactions: (A) PcpB catalyzes the conversion of pentachlorophenol 26 to tetrachlorobenzoquinone 28. (B) The reaction catalyzed by iodotyrosine deiodinase.
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Fig. 7 (A) The reaction catalyzed by HcbA1. (B) Mechanistic proposal for the HcbA1catalyzed reaction involving flavin-N5-oxide 5 formation.
4 formation and completion of the reaction. LC-MS analysis of the resulting reaction mixture demonstrated the formation of the flavin-N5-oxide 5. When the reaction was carried out in the presence of 18O2, the predicted 2 Da increase in its mass was also observed (Adak & Begley, 2019).
5.4 General guidelines for the detection of flavin-N5-oxide intermediates 1. Flavin-N5-oxide 5 is readily reduced to flavin. It is therefore essential to exclude reducing agents such as DTT, TCEP and reduced nicotinamide from the reaction mixture. Reduced flavin is prepared by photoreduction in the presence of EDTA rather than by enzymatic reduction using NAD(P)H as the reducing agent. 2. Although the UV–Visible spectra of oxidized flavin 1 and flavin-N5oxide 5 are similar, there is a clear distinction between the two species in the 450 nm region (λmax of flavin is 447 nm whereas λmax of flavin-N5oxide is 462 nm). This can be used as an initial indicator of the formation of a flavin-N5-oxide 5 intermediate. 3. Flavin-N5-oxide 5 can be detected by HPLC analysis of the enzymatic reaction mixture (single turnover) using synthesized FMN-N5-oxide 5 as an authentic standard.
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Fig. 8 Proposal for the use of flavin-N5-oxide intermediates by flavoenzymes catalyzing substrate hydroperoxide formation in the absence of an active site cysteine.
4. As of yet, protein amino acid sequence analysis has not generated a characteristic sequence motif that enables the identification of flavin-N5oxide-forming enzymes. However, mechanistic analysis has proven useful in the identification of such enzymes. The analysis of the DszA reaction mechanism suggested that flavin-N5-oxides are likely intermediates in any flavoenzyme involving substrate hydroperoxide formation in the absence of an active site cysteine (Fig. 8) (Hall et al., 2009). This model led to the identification of RutA and HcbA1 as possible FMN-N5-oxide generating enzymes, but does not include the catalytic motif shown in the EncM-catalyzed reaction. It is highly likely that many additional examples of flavin-N5-oxide-utilizing-flavoenzymes remain to be discovered.
Funding This research was supported by the Robert A. Welch Foundation Grant A-0034.
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Denome, S. A., Olson, E. S., & Young, K. D. (1993). Identification and cloning of genes involved in specific desulfurization of dibenzothiophene by Rhodococcus sp. strain IGTS8. Applied and Environmental Microbiology, 59(9), 2837–2843. Enns, K., & Burgess, W. H. (1965). The photochemical oxidation of ethylenediaminetetraacetic acid and methionine by riboflavine. Journal of the American Chemical Society, 87(24), 5766–5770. Gonzalez-Osorio, L., Luong, K., Jirde, S., Palfey, B. A., & Vey, J. L. (2016). Initial investigations of C4a-(hydro)peroxyflavin intermediate formation by dibenzothiophene monooxygenase. Biochemical and Biophysical Research Communications, 481(1–2), 189–194. Gray, K. A., Pogrebinsky, O. S., Mrachko, G. T., Xi, L., Monticello, D. J., & Squires, C. H. (1996). Molecular mechanisms of biocatalytic desulfurization of fossil fuels. Nature Biotechnology, 14(13), 1705–1709. Guan, L.-J., Lee, W. C., Wang, S., Ohshiro, T., Izumi, Y., Ohtsuka, J., et al. (2015). Crystal structures of apo-DszC and FMN-bound DszC from Rhodococcus erythropolis D-1. The FEBS Journal, 282(16), 3126–3135. Hall, A., Karplus, P. A., & Poole, L. B. (2009). Typical 2-Cys peroxiredoxins—Structures, mechanisms and functions. The FEBS Journal, 276(9), 2469–2477. Heelis, P. F. (1982). The photophysical and photochemical properties of flavins (isoalloxazines). Chemical Society Reviews, 11(1), 15–39. Hu, J., Chuenchor, W., & Rokita, S. E. (2015). A switch between one- and two-electron chemistry of the human flavoprotein iodotyrosine deiodinase is controlled by substrate. The Journal of Biological Chemistry, 290(1), 590–600. Ito, K., Takagi, K., Iwasaki, A., Tanaka, N., Kanesaki, Y., Martin-Laurent, F., et al. (2017). Identification of the hcb gene operon involved in catalyzing aerobic hexachlorobenzene dechlorination in Nocardioides sp. strain PD653. Applied and Environmental Microbiology, 83(19), e00824-17/1-e00824-17/17. Ito, K., Takagi, K., Matsushima, Y., Igimi, S., Ito, K., Takagi, K., et al. (2018). Identification of the novel hcbB operon catalyzing the dechlorination of pentachlorophenol in the Gram-positive bacterium Nocardioides sp. strain PD653. Journal of Pesticide Science, 43(2), 124–131. Lee, W. C., Ohshiro, T., Matsubara, T., Izumi, Y., & Tanokura, M. (2006). Crystal structure and desulfurization mechanism of 2’-hydroxybiphenyl-2-sulfinic acid desulfinase. The Journal of Biological Chemistry, 281(43), 32534–32539. Loh, K. D., Gyaneshwar, P., Papadimitriou, E. M., Fong, R., Kim, K.-S., Parales, R., et al. (2006). A previously undescribed pathway for pyrimidine catabolism. Proceedings of the National Academy of Sciences of the United States of America, 103(13), 5114–5119. Mansurova, M., Koay, M. S., & Gaertner, W. (2008). Synthesis and electrochemical properties of structurally modified flavin compounds. European Journal of Organic Chemistry, (32), 5401–5406. McCormick, D. B., Koster, J. F., & Veeger, C. (1967). Mechanisms of photochemical reductions of FAD and FAD-dependent flavoproteins. European Journal of Biochemistry, 2(4), 387–391. Mukherjee, T., Zhang, Y., Abdelwahed, S., Ealick, S. E., & Begley, T. P. (2010). Catalysis of a flavoenzyme-mediated amide hydrolysis. Journal of the American Chemical Society, 132(16), 5550–5551. Phatarphekar, A., Buss, J. M., & Rokita, S. E. (2014). Iodotyrosine deiodinase: A unique flavoprotein present in organisms of diverse phyla. Molecular BioSystems, 10(1), 86–92. Piddington, C. S., Kovacevich, B. R., & Rambosek, J. (1995). Sequence and molecular characterization of a DNA region encoding the dibenzothiophene desulfurization operon on Rhodococcus sp. strain IGTS8. Applied and Environmental Microbiology, 61(2), 468–475.
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Saleem-Batcha, R., Stull, F., Sanders, J. N., Moore, B. S., Palfey, B. A., Houk, K. N., et al. (2018). Enzymatic control of dioxygen binding and functionalization of the flavin cofactor. Proceedings of the National Academy of Sciences of the United States of America, 115(19), 4909–4914. Soleimani, M., Bassi, A., & Margaritis, A. (2007). Biodesulfurization of refractory organic sulfur compounds in fossil fuels. Biotechnology Advances, 25(6), 570–596. Sun, Z., Su, Q., & Rokita, S. E. (2017). The distribution and mechanism of iodotyrosine deiodinase defied expectations. Archives of Biochemistry and Biophysics, 632, 77–87. Teufel, R. (2017). Flavin-catalyzed redox tailoring reactions in natural product biosynthesis. Archives of Biochemistry and Biophysics, 632, 20–27. Teufel, R. (2018). Preparation and characterization of the Favorskiiase flavoprotein EncM and its distinctive flavin-N5-oxide cofactor. Methods in Enzymology, 604, 523–540. Teufel, R., Agarwal, V., & Moore, B. S. (2016). Unusual flavoenzyme catalysis in marine bacteria. Current Opinion in Chemical Biology, 31, 31–39. Teufel, R., Miyanaga, A., Michaudel, Q., Stull, F., Louie, G., Noel, J. P., et al. (2013). Flavin-mediated dual oxidation controls an enzymatic Favorskii-type rearrangement. Nature, 503(7477), 552–556. Teufel, R., Stull, F., Meehan, M. J., Michaudel, Q., Dorrestein, P. C., Palfey, B., et al. (2015). Biochemical establishment and characterization of EncM’s flavin-N5-oxide cofactor. Journal of the American Chemical Society, 137(25), 8078–8085. Thomas, S. R., McTamney, P. M., Adler, J. M., LaRonde-LeBlanc, N., & Rokita, S. E. (2009). Crystal structure of iodotyrosine deiodinase, a novel flavoprotein responsible for iodide salvage in thyroid glands. The Journal of Biological Chemistry, 284(29), 19659–19667.
Further reading Agarwal, V., Miles, Z. D., Winter, J. M., Eustaquio, A. S., El Gamal, A. A., & Moore, B. S. (2017). Enzymatic halogenation and dehalogenation reactions: Pervasive and mechanistically diverse. Chemical Reviews, 117(8), 5619–5674. Crawford, R. L., Jung, C. M., & Strap, J. L. (2007). The recent evolution of pentachlorophenol (PCP)-4-monooxygenase (PcpB) and associated pathways for bacterial degradation of PCP. Biodegradation, 18(5), 525–539. Hlouchova, K., Rudolph, J., Pietari, J. M. H., Behlen, L. S., & Copley, S. D. (2012). Pentachlorophenol hydroxylase, a poorly functioning enzyme required for degradation of pentachlorophenol by Sphingobium chlorophenolicum. Biochemistry, 51(18), 3848–3860.