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Article

Inter-Species Redox Coupling by Flavin Reductases and FMN-Dependent Two-Component Monooxygenases Undertaking Nucleophilic Baeyer–Villiger Biooxygenations

by
Andrew Willetts
Curnow Consultancies Ltd., Helston, Cornwall TR13 9PQ, UK
Microorganisms 2023, 11(1), 71; https://doi.org/10.3390/microorganisms11010071
Submission received: 10 November 2022 / Revised: 19 December 2022 / Accepted: 23 December 2022 / Published: 27 December 2022
(This article belongs to the Special Issue Microbial Biocatalysis and Biodegradation)
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Abstract

:
Using highly purified enzyme preparations throughout, initial kinetic studies demonstrated that the isoenzymic 2,5- and 3,6-diketocamphane mono-oxygenases from Pseudomonas putida ATCC 17453 and the LuxAB luciferase from Vibrio fischeri ATCC 7744 exhibit commonality in being FMN-dependent two-component monooxygenases that promote redox coupling by the transfer of flavin reductase-generated FMNH2 by rapid free diffusion. Subsequent studies confirmed the comprehensive inter-species compatibility of both native and non-native flavin reductases with each of the tested monooxygenases. For all three monooxygenases, non-native flavin reductases from Escherichia coli ATCC 11105 and Aminobacter aminovorans ATCC 29600 were confirmed to be more efficient donators of FMNH2 than the corresponding tested native flavin reductases. Some potential practical implications of these outcomes are considered for optimising FMNH2-dependent biooxygenations of recognised practical and commercial value.

1. Introduction

Historically, the isoenzymic enantiocomplementary diketocamphane monooxygenases (DKCMOs) from the (rac)-camphor-degrading bacterium Pseudomonas putida ATCC 17453 [1] and the LuxAB luciferase from the bioluminescent marine bacterium Vibrio fischeri ATCC 7744 [2], along with a small number of other known enzymes, were first grouped together on shared functional properties as NADH + FMN-dependent Type 2 Baeyer–Villiger monooxygenases (BVMOs) to distinguish them from the NADPH + FAD-dependent Type 1 BVMOs [3]. While the much more prevalent Type 1 BVMOs have always been recognised as single component true flavoprotein enzymes replete with an active site-bound FAD coenzyme [4], it is now acknowledged that the DKCMOs (2,5-DKCMO [EC 1.14.14.108] and 3,6-DKCMO [EC 1.14.14.155]), and LuxAB luciferase (EC 1.14.14.3) are structurally different, being archetypal members of the recently delineated FMN-dependent two-component monooxygenases (fd-TCMOs, [5]). The defining feature of the fd-TCMOs that excludes them from being classed as true flavoproteins is that they are dependent on deploying fully reduced FMN (FMNH2) as a cosubstrate rather than as an active site-bound coenzyme. Significantly, the FMNH2 cosubstrate is acquired from one or more distal flavin reductases (FRs [EC 1.5.1.x]). FRs are widely distributed enzymes, a number of which have been purified from various bacteria: the nomenclature and characteristics of some well-studied prokaryotic FRs confirmed to fully reduce FMN effectively are summarised in Table 1. Because reduced flavins can react both directly and indirectly with molecular oxygen to generate highly reactive oxygen species (ROS) that are capable of destroying DNA, lipids, and proteins [6,7,8,9], the mode of functioning of the fd-TCMOs poses a considerable challenge to aerobic organisms that consequently requires a close logistical relationship between the two participating activities [10].
Being similar enzymes, the LuxAB luciferase of V. fischeri ATCC 7744 and the isoenzymic DKCMOs of P. putida ATCC 17453 share a significant amount of directly equivalent biochemistry. Despite the highly specialised evolved role of dodecanal-deploying LuxAB luciferase in promoting bioluminescence [11], purified preparations of each enzyme can catalyse nucleophilic Baeyer–Villiger oxidations of various aliphatic and alicyclic abiotic ketones to corresponding biooxygenated products [12,13]. However, both isoenzymic DKCMOs do exhibit an important difference from the luciferase with respect to the facial diastereoselectivity of the hydride ion exchanges promoted by the FMNH2 cosubstrate (si-face versus re-face, respectively). This in turn can generate enantiodivergent chiral lactones from racemic and prochiral alicyclic ketone substrates. A number of these products have considerable proven value as synthons in the chemoenzymatic synthesis of sarkomycin A [14], clerodin [15], R-(+)-lipoic acid [16,17], azadirachtin [18], various carbocyclic nucleoside [19] and prostaglandin [20] analogues, plus a significant number of α,ω-dicarboxylic and α-aminocarboxylic acids [21], including the Nylon 6 monomer 6-aminohexanoic acid [22] and the Nylon 9 monomer 9-aminononanoic acid [23]. Another shared characteristic of the luciferase and the DKCMOs is that the requisite FMNH2 can be generated by multiple alternative native FRs (FRGVf and FreVf in V. fischeri, [24]; Frp1, Frp2, Fred, and putidaredoxin reductase (PdR) in P. putida ATCC 17453 [10]), which may represent an evolved solution to the problem posed by the relatively inefficient reduction of flavins by reduced pyridine nucleotides [25,26,27,28]. Whereas Frp1, Frp2, Fred, and FreVf are devoid of any bound flavin and sequentially deploy FMN and NADH as cosubstrates to generate FMNH2, putidaredoxin and FRGVf are flavoproteins that contain a bound flavin coenzyme and generate FMNH2 by a ping-pong reaction mechanism (Figure 1).
These two functionally different types of enzyme have been termed Class I and Class II FRs, respectively [26,28]. That the biochemistry common to the LuxAB luciferase of V. fischeri and the isoenzymic DKCMOs of P. putida ATCC 17453 may extend to the functional interchangeability of the relevant FMNH2-generating and FMNH2-oxidising activities of these particular fd-TCMOs was suggested by some preliminary data indicating that one or more of the FRs present in a commercial grade crude extract prepared from cells of V. fischeri (Boehringer Mannheim Co) can support the biooxygenation of alicyclic ketones to lactones by purified preparations of each DKCMO isoenzyme [13,29]. The possibility that this apparent compatibility results from the effective functional inter-changeability of discrete highly purified preparations of the relevant FMNH2-generating and FMNH2-oxidising activities of the DKCMOs and LuxAB luciferase remains to be proven. This was the specific initial goal of the present paper.
Significant in the context of the inter-species compatibility of different functional activities of fd-TCMOs is that the LuxAB luciferase of V. fischeri [30,31] and both DKCMOs of P. putida [32,33,34] have all proved to be highly active when expressed in recombinant bacteria that are deficient in any of the corresponding native FRs (FRGVf, and FreVf [V. fischeri]; Frp1, Frp2, Fred, and PdR [P. putida]). Consequently, the study has been widened to investigate the ability of purified preparations of the characterised FRs of some other known FMN-dependent two-component enzymes (Table 1) to support the nucleophilic biooxygenating activities of the DKCMOs from P. putida ATCC 17453 and the LuxAB luciferase from V. fischeri ATCC 7744. This extended aspect of inter-species biocompatibility became the second specific goal of the present paper.
By way of direct support for this wider study, it has been demonstrated that a purified preparation of FreEc, the major FR isolated from E. coli, supports bioluminescence in vitro with the LuxAB luciferase from V. harveyi [35]. Less well characterised is recognition that a commercial grade crude extract of FRs prepared from cells of V. fischeri (Boehringer Mannheim Co) can support corresponding biooxygenations by the FMNH2-dependent monooxygenase moieties of the characterised fd-TCMOs nitrilotriacetate monooxygenase [36], pristinamycin PIIA synthase [37], and EDTA monooxygenase [38]. A relevant related observation is that purified 2,5-DKCMO from ATCC 17453 has been shown to function effectively with FMNH2 generated by hydride donation from the synthetic nicotinamide coenzyme biomimetics 1-benzyl-1,4-dihydronicotinamide (BNAH), and 1-(2-carbamoylmethyl)-1,4-dihydronicotinamide (AmNAH) [39].

2. Materials and Methods

2.1. Bacterial Strains, Culture Maintenance, and Growth Conditions

P.putida ATCC 17453 was cultured at 30 oC using a mineral salts medium supplemented with 17.5 mM (rac)-camphor as the principal carbon source as fully described previously [27].
A. aminovorans ATCC 29600 was cultured at 30 °C using a mineral salts medium supplemented with 1 g L−1 of nitrilotriacetate as fully described previously [40].
Escherichia coli ATCC 11105 was cultured at 37 °C using LB medium (Sigma-Aldrich, Dorset, UK) as fully described previously [41].
Vibrio fischeri ATCC 7744 was cultured at 26 °C using Photobacterium medium (Difco Laboratories, Detroit, Mich., USA) as fully described previously [2].

2.2. Extract Preparation

Cultures were grown into mid log phase of growth and the biomass content harvested by centrifugation (10,000× g for 15 min at 5 °C [MSE Coolspin 2, MSE, Heathfield, East Sussex, UK]), washed with an equal volume of cold Tris-HCl buffer (0.1 M, pH 7.0) and then recentrifuged. The recovered cells were evenly suspended in 7.5 mL of the same buffer and subsequently sonicated (Soniprep150, MSE, Heathfield, UK) in ice for 3 × 2 min. The resultant homogenates were centrifuged (20,000× g for 15 min at 5 °C) to remove the cell debris.

2.3. Purification of 2,5-DKCMO, 3,6-DKCMO, and LuxAB Luciferase

Samples of the highly purified biooxygenating subunits of both enantiocomplementary DKCMO isoenzymes were prepared at 4 °C using a BioLogic FPLC system (BioLogic 10, Bio-Rad, Hercules, CA, USA) deploying anion-exchange chromatography (Q-Sepharose, Sigma-Aldrich, Dorset, UK), followed by a Mono-Q column (Pharmacia, Stockholm, Sweden) eluted with a linear gradient of 0–0.6 M KCl in 21 mM phosphate buffer pH 7.1, as fully described previously [18]. The A280 trace from the Mono-Q column output is shown in Supplementary Figure S1A. The isolated 2,5-DKCMO activity comprises an approximately equimolar mixture [12] of 2,5-DKCMO-1 (coded for by the camE25-1 gene) and 2,5-DKCMO-2 (coded for by the camE25-2 gene). Purified DKCMO activities were assayed in the co-presence of 20mU of a commercial FR preparation (NAD(P)H: FMN oxidoreductase from V. fischeri, (Boehringer Mannheim, Indianapolis, Ind, USA) by measuring the rate of NADH-stimulated lactone formation from the corresponding diketocamphane substrate by GC (Shimadzu GC-14A, Shimadzu Europe) using a 10% Carbowax 20 M column [27]. Samples of LuxAB luciferase were prepared using the BioLogic FPLC system deploying anion-exchange chromatography (DEAE-Sephadex, Sigma-Aldrich, Dorset, UK) as fully described previously [42]. LuxAB luciferase activity was assayed in the co-presence of 20 mU of a commercial FR preparation (NAD(P)H: FMN oxidoreductase from V. fischeri (Boehringer Mannheim, Indianapolis, Ind) by measuring the rate of NADH-stimulated dodecanoic acid formation from 2-tridecanone by GC (Shimadzu GC-14A) using a BP1 column [13].

2.4. Purification of Frp1, Frp2, FreEc, FRGVf, FreVf, and FRDAa

Samples of highly purified Frp1 and Frp2 were prepared at 4 oC using a BioLogic FPLC system (BioLogic 10, Bio-Rad, Hercules, CA, USA) deploying successive anion-exchange (Mono-Q, Pharmacia, Stockholm, Sweden), affinity (Reactive Blue-4-agarose, Pharmacia), and gel filtration (HiLoad 16/60 Superose 12, Pharmacia) columns in the three-stage protocol fully described previously [27]. Samples of highly purified FreEc were prepared using the BioLogic FPLC system deploying a nickel affinity (Sigma-Aldrich) column as fully described previously [35]. Samples of highly purified FreVf and FRGVf were prepared using the BioLogic FPLC system deploying successive gel filtration (Sephadex G-100, Sigma-Aldrich), and anion-exchange (DEAE-Sephadex, Sigma Aldrich) columns as fully described previously [43]. Samples of highly purified FRGAa were prepared using the BioLogic FPLC system deploying successive anion-exchange (Q-Sepharose [Sigma-Aldrich]), followed by Mono-Q [Pharmacia, Stockholm, Sweden]) and gel filtration (Superdex 75 [Sigma-Aldrich]) columns as fully described previously [40]. SDS-PAGE showing purification of each of the FRs is shown in Supplementary Figure S1B: lane 1, FreVf; lane 2, FRGVf; lane 3, Frp1; lane 4, Frp2; lane 5, FreEc; lane 6, FRDAa; lane 7, 2,5-DKCMO; lane 8, 3,6-DKCMO; lane 9, Pharmacia low MW markers.

2.5. Single-Enzyme Kinetic Studies

Assays using 20 mU aliquots of highly purified FR enzyme preparations were conducted spectrophotometrically at 340 nm under anaerobic conditions by measuring the initial rate of enzyme-catalysed reduction of FMN by NADH as fully described previously [10].

2.6. Coupled-Enzyme Kinetic Studies

Assays using appropriate combinations of 20 mU aliquots of highly purified FR preparations and 200 mU aliquots of highly purified monooxygenase preparations plus 1 mM (rac)-bicyclo[3.2.0]hept-2-en-6-one as the biooxidisable substrate were conducted spectrophotometrically at 340 nm by measuring the initial rate of enzyme-catalysed reduction of FMN by NADH as fully described previously [10]. Each reaction mixture was supplemented with 60 U catalase (Sigma-Aldrich) to avoid accumulation of hydrogen peroxide resulting from substrate-independent oxygen consumption.

2.7. Longer-Term (120 min) Biocatalytic Reactions with Combinations of Highly Purified Enzymes

Biotransformations with the various combinations of highly purified FRs and monooxygenases were carried out in reaction mixtures (1 mL) containing Tris/HCl buffer (60 mM, pH 7.6), 0.1 mM NADH, 0.03 mM FMN, 60 U catalase, 30 mU formate dehydrogenase, 50 mM sodium formate, 20 mU FR, 200 mU monooxygenase, and 1 mM (rac)-bicyclo[3.2.0]hept-2-en-6-one. All biotransformation reaction mixtures were incubated at 25 °C for 120 min, and then assayed by chiral capillary GC using a Lipadex D fused silica column as fully described previously [44,45].

2.8. Reproducibility

Where indicated, procedures were repeated three times with equivalent purified enzyme preparations, and the resultant data presented graphically with corresponding standard deviation error bars (Supplementary Figure S2A–D).

3. Results and Discussion

As prior studies with both the isoenzymic DKCMOs from P. putida ATCC 17453 [32,33,34,46] and the LuxAB luciferase from V. fischeri ATCC 7744 [13] have confirmed that highly purified preparations of each enzyme can biooxygenate (rac)-bicyclo[3.2.0]hept-2-en-6-one to some extent, thereby generating a mixture of regioisomeric 2-oxa- and 3-oxa-lactones (2,5-DKCMO > 3,6-DKCMO > LuxAB luciferase), this bicyclic ketone was used as the consensus substrate of choice in the present study of these three fd-TCMO enzymes. An important value of this substrate is that the nature of the ensuing lactone products serves as a monitor of both the regio- and stereoselectivity of the three enzymes. In this respect, prior research with this abiotic bicyclic ketone has indicated that whereas the two enzymic DKCMOs exhibit stereochemical congruence, as clearly reflected by the equivalence of the predominant lactones formed [12], the LuxAB luciferase exhibits stereochemical divergence from both DKCMOs by generating corresponding lactones in the opposite enantiomeric series [13]. However, because both the luciferase and the 3,6-DKCMO isoenzyme exhibit some form of toxic response to the ketone and/or generated ROS which begins to be progressively significant 3–4 hours after initial exposure [13,17], all the current studies that deployed this substrate were conducted over the shorter time-frame of 120 minutes, when typically 70–90%, 40–60%, and 20–40% of the (rac)-ketone had been converted to a mixture of the corresponding 2-oxa and 3-oxa lactones by 2,5-DKCMO, 3,6-DKCMO and the LuxAB luciferase, respectively.
Prior to investigating any aspect of the interchangeability of FMNH2-generating activities between different fd-TCMOs, a series of relevant comparative kinetic studies were conducted to establish whether the transfer of FR-generated FMNH2 to the corresponding monooxygenase moiety of the isoenzymic DKCMOs of P. putida and the LuxAB luciferase of V. fischeri occurs by rapid free diffusion or involves the formation of some form of transitory complex [47]. The specific aim of these kinetic studies was to establish the apparent KmFMN values for representative FRs of P. putida ATCC 17453 (Fpr1, and Frp2), and V. fischeri ATCC 7744 (FRGVf and FreVf), when assayed either as a highly purified FR preparation catalysing a single-enzyme FMNH2-generating step, or as a coupled-enzyme combination promoting FMNH2-turnover in the co-presence of (rac)-bicyclic ketone and a 10-fold excess of highly purified preparations of the corresponding native monooxygenase. While the single-enzyme assays were conducted under anaerobic conditions, the corresponding monooxygenase-dependent coupled-enzyme reactions were assayed aerobically, but with the inclusion of catalase to decompose any hydrogen peroxide generated abiotically from FMNH2 by molecular oxygen [10]. In each case, FMN was tested over a range of concentrations up to 30 μM, which resulted in the corresponding v values asymptotically approaching a consistent Vmax value. The outcomes of the representative comparative single- vs. coupled-enzyme kinetic studies, when plotted as corresponding Michaelis–Menten plots (Supplementary Figure S2A–D), confirmed that in each case the apparent Vmax of each coupled-enzyme assay was approximately 2–3-fold lower than that of the corresponding single-enzyme assay, most likely reflecting known differences in the apparent turnover rates of the relevant enzymes [27,41,47]. However, most significantly, for each comparative single- vs. coupled-enzyme study, the corresponding calculated apparent KmFMN values derived in each case using the Michaelis–Menten equation and relevant mean values were remarkably similar, as summarised in Table 2.
This commonality of the derived kinetic outcomes indicates that 2,5-DKCMO, 3,6-DKCMO, and the LuxAB luciferase of V. fischeri each operate a common mode of action that involves the transfer of FR-generated FMNH2 by rapid free diffusion, as the alternative formation of a FR-monooxygenase transitory complex involving protein–protein interaction would have been signalled by significantly lower recorded KmFMN values for the coupled-enzyme reactions (≤1 order of magnitude [41]). Equivalent less rigorously supported proposals were made for the LuxAB luciferase of V. harveyi when functioning with the with FreEc, the major FR of E. coli [35], and for the transfer to 2,5-DKCMO of FMNH2 generated by the synthetic nicotinamide coenzyme biomimetic BNAH [39].
That rapid free diffusion of the FMNH2 cosubstrate from competent native donor FRs is the proven mode of action of all three tested fd-TCMOs may enable them to function effectively with non-native FMNH2-generating FRs. This possibility was examined by monitoring the outcomes of 120 min biotransformations of (rac)-bicyclo[3.2.0]hept-2-ene -6-one by highly purified preparations of both isoenzymic DKCMOs and the LuxAB luciferase in relevant coupled-enzyme assays with in each case a range of both corresponding native and non-native FRs (Figure 2). The respective relevant highly purified native FRs deployed were those tested previously (Table 2), whereas for all three monooxygenases the two additional non-native NADH-dependent FRs that were deployed were FRDAa and FreEc, which both deploy a sequential mechanism of hydride transfer (Table 1). Both FRDAa and FreEc are highly effective NADH-dependent FMN-dependent reductases as reflected by their reported KmFMN single-enzyme assay values of 1.0 µM [40] and 0.8 µM [35], respectively. Each biotransformation was supplemented with a formate/formate dehydrogenase NADH-regenerating system to enhance hydride ion availability, and catalase to avoid accumulation of hydrogen peroxide resulting from substrate-independent oxygen consumption. All the tested FRs are NADH-dependent enzymes which, with the exception of FRGVf, deploy a sequential reaction mechanism (Table 2).
Analysis of the stopped 120-minute reaction mixtures (Table 3) indicated a number of shared and monooxygenase-specific outcomes. Most significantly, with respect to redox coupling, the data confirm that the LuxAB luciferase and both isoenzymic DKCMOs can biooxygenate the bicyclic ketone by sourcing the requisite FMNH2 cosubstrate from both non-native as well as corresponding native FRs. Because equivalent aliquots of FR (20 mU) and highly purified monooxygenase (200 mU) were used in each case, the residual ketone remaining in the 120-minute stopped reaction mixtures can be taken as a reflexion of the resultant activity of the relevant coupled-enzyme reactions. Relevant in this respect is that the affinity of each monooygenase for FMNH2 will be an idiosyncratic constant [5]. The data indicate that for each of the tested fd-TCMOs, the relative order of effectiveness of the alternative highly purified FRs serving as suppliers of FMNH2 was FreEc > FRDAa > Frp2 > Frp1 = FreVf > FRGVf. This outcome, indicating that FreEc and FRDAa promote the highest recorded levels of ketone biotransformation with each set of coupled-enzyme reactions, correlates with the corresponding KmFMN values of the tested FRs calculated from relevant single-enzyme kinetic assays listed in Table 2, albeit the values for FreEc and FRDAa (0.8 μM, and 1.0 μM, respectively) are published precedents generated under slightly different reaction conditions. The relatively poor performance of FRGVf, the only tested FR that deploys a ping-pong rather than a sequential reaction mechanism to generate FMNH2 (Figure 1), may be related to the comparatively low Vmax recorded for this enzyme (Supplementary Figure S2A–D). It may also be relevant that for each coupled-enzyme partnership there will be an uncharacterised effect resulting from any proximity factor that may influence the logistics of the transfer of FMNH2 by rapid free diffusion between the two participating biocatalysts [5].
Previous studies of enzyme catalysed Baeyer–Villiger reactions have proposed that monooxygenase-dictated stereoelectronic effects serve as the principal determinant in controlling multiple aspects of the selectivity expressed by the relevant outcomes [48,49]. Detailed analysis of the lactones formed from (rac)-bicyclo-[3.2.0]hept-2-en-6-one in the currently analysed coupled-enzyme reactions (Table 3) shows that the relative amounts (and consequently the ratio) of the ketone enantiomers biooxidised, and both the absolute configuration and enantiomeric excess of the resultant predominant regioisomeric lactones are highly consistent with all six partner FRs tested in each individual monooxygenase-catalysed series. These data both concur with the proposed cardinal role of the monooxygenases in delineating all currently investigated aspects of their specificity, and conversely confirm a predictable absence of any relevant specificity-related influence(s) resulting from the deployment of the tested range of non-native as well as native coupled-enzyme partner FRs.
The particular outcomes of the biooxygenations catalysed by LuxAB luciferase are interesting in another respect. Because the (−)-(1S,5R)-2-oxa-lactone formed is an acknowledged synthon for the chemoenzymatic synthesis of various potentially useful prostaglandin analogues [50], it is significant that the calculated enantiomeric purities [45] of the (−)-2-oxa-lactone recorded with this monooxygenase in combination with each of the tested purified FRs are all higher than those reported previously for equivalent biotransformations undertaken by the Type 1 BVMOs 2-oxo-Δ3-4,5,5-trimethylcyclopentenylacetyl-CoA monooxygenase sourced from P. putida ATCC 17453 [4,45,51], and cyclohexanone monooxygenase sourced from either Acinetobacter TD63 [52], or Acinetobacter calcoaceticus NCIMB 9871 [53,54].
Clearly, under the reaction conditions deployed, using highly purified enzyme preparations supplemented with catalase to promote hydrogen peroxide scrubbing, NADH-dependent FreEc from E. coli and FRDAa from A. aminovorans are superior suppliers of FMNH2 than each of the corresponding native FRs for each of the tested monooxygenases. Interestingly, purified 2,5-DKCMO plus catalase has also been shown to function effectively in undertaking biooxygenation of (rac)-bicyclo[3.2.0]hept-2-en-6-one with FMNH2 generated by hydride donation from synthetic nicotinamide coenzyme biomimetics [39]. Although less extensively characterised, an equivalent outcome conducted without supplementary exogenous catalase addition was reported for flavin reductase-coupling with DszA and DszC, two fd-TCMOs involved in the 4S pathway of dibenzothiophene desulfurisation by Rhodococcus erythropilis D-1 [55]. Coupled-reactions by both monooxygenases with a non-native FR sourced from Paenibacillus polymyxa A-1 were reported to proceeded more efficiently than with the corresponding native FR DszD. Less well defined are a number of reports confirming relevant biooxidative activity in recombinant strains of FreEc-containing E. coli expressing 2,5-DKCMO [32], and 3,6-DKCMO [33,34], and co-expressing both 4S pathway monooxygenases (DszA and DszC) plus the FRGVh gene sourced from V. harveyi [56,57,58].
The major outcome of the study has been to use highly purified enzyme preparations to demonstrate the comprehensive inter-species compatability of the FR and monooxygenase componenents of three fd-TCMOs catalysing FMNH2-dependent Baeyer–Villiger-type nucleophilic biooxygenations of a bicyclic ketone to corresponding 2-oxa- and 3-oxa-lactones. This, along with another equivalent precedent involving an abiotic source of the requisit FMNH2 [39], suggests that FMN-dependent TCMOs are characterised by an evolved commonality of function in deploying the reduced form of the flavin cosubstrate irrespective of its biotic or abiotic source. For all three tested fd-TCMOs, it is most significant that the non-native FRs FreEc and FRDAa were confirmed to be more effective donators of FMNH2 than the corresponding native FRs when tested with highly purified enzyme preparations in the co-presence of exogenous catalase, as reflected consistently by all tested parameters of the corresponding coupled reactions. This confirms absolutely a promiscuous aspect to the functioning of these enzymes, as proposed previously for the functioning of both isoenzymic DKCMOs with multiple confirmed competent native FRs (Frp1, Frp2, Fred, and PdR [1]), and specifically for 2,5-DKCMO with the abiotic synthetic nicotinamide biomimetics BNAH and AmNAH [39]. Catalytic promiscuity as a concept has been proposed to serve an important role in enzyme evolution [59,60,61], which in the case of the tested fd-TCMOs may be related to the known inefficiency of FMN reduction by NAD(P)H2 [25,26,27]. While the implications of this particular outcome may have considerable potenial practical value for optimising fd-TCMO-catalysed biotransformation and bioremediation processes [3,12,16,17,62,63,64,65,66], this must be tempered with the recognition that plural detrimental toxic effects can result from the production of hydrogen peroxide and superoxide radicals generated by high levels of FR activity in the absence of corresponding levels of catalase [6,7,8,9,67].

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/microorganisms11010071/s1. Figure S1: Final step purifications of both tested DKCMO isoenzymes (A), and all tested FRs (B). Figure S2: Michaelis-Menton plots of the kinetic data for the studied complementary single-enzyme and coupled-enzyme assays.

Funding

This research received no external funding.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

The original contributions presented in the study are included in the article: further enquiries can be directed to the corresponding author.

Acknowledgments

A special acknowledgment must go to Raffaella Villa ([email protected]), who as a postgraduate research student in my laboratory was the first to discover that the LuxAB luciferase of V. fischeri ATCC 7744 could catalyse nucleophilic Baeyer–Villiger oxidations of abiotic alicyclic and aliphatic ketones to corresponding biooxygenated products [13]. The support of Curnow Consultancies Ltd in capitalising on those initial outcomes is gratefully acknowledged: ‘Mighty oaks from little acorns grow’ (14th century English proverb).

Conflicts of Interest

The author declares no conflict of interest.

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Microorganisms 11 00071 g001 550
Figure 1. The different types (bound vs. unbound flavin) and the different reaction mechanisms (sequential vs. ping-pong) of the flavin reductases of V. fischeri ATCC 7744 and P. putida ATCC 17453. E = flavin reductase with unbound flavin coenzyme (Frp1; Frp2; Fred; FreVf): F = FMN: EF* = flavin reductase with bound flavin coenzyme (FRGVf [FMN]; PdR [FAD]): FH2 = FMNH2.
Figure 1. The different types (bound vs. unbound flavin) and the different reaction mechanisms (sequential vs. ping-pong) of the flavin reductases of V. fischeri ATCC 7744 and P. putida ATCC 17453. E = flavin reductase with unbound flavin coenzyme (Frp1; Frp2; Fred; FreVf): F = FMN: EF* = flavin reductase with bound flavin coenzyme (FRGVf [FMN]; PdR [FAD]): FH2 = FMNH2.
Microorganisms 11 00071 g001
Microorganisms 11 00071 g002 550
Figure 2. Potential outcomes for the biooxygenation of (rac)-bicyclo-[3.2.0]hept-2-en-6-one to 2-oxa- and 3-oxa-lactones by fd-TCMOs dependent on FMNH2 sourced from an FR. Tested fd-TCMOs: 2,5-DKCMO; 3,6-DKCMO; LuxAB luciferase. Tested FRs: Frp1 (P. putida); Frp2 (P. putida); FreVf (V. fischeri); FRGVf (V. fischeri); FreEc (E. coli); FRDAa (A. aminovorans).
Figure 2. Potential outcomes for the biooxygenation of (rac)-bicyclo-[3.2.0]hept-2-en-6-one to 2-oxa- and 3-oxa-lactones by fd-TCMOs dependent on FMNH2 sourced from an FR. Tested fd-TCMOs: 2,5-DKCMO; 3,6-DKCMO; LuxAB luciferase. Tested FRs: Frp1 (P. putida); Frp2 (P. putida); FreVf (V. fischeri); FRGVf (V. fischeri); FreEc (E. coli); FRDAa (A. aminovorans).
Microorganisms 11 00071 g002
Table
Table 1. Characterised FMN-dependent FRs [5].
Table 1. Characterised FMN-dependent FRs [5].
Flavin Reductase:
Nomenclature and Reaction Mechanism		Microbial
Source		MW and
Structure		Bound or Unbound FMN
Frp1
Sequential		P. putida26.0 kDaUnbound
Frp2
Sequential		P. putida27.0 kDaUnbound
Fred
Sequential		P. putida2 × 18.0 kDaUnbound
PdR
Ping-pong		P. putida48.5 kDaBound
FreEc (FR-II)
Sequential		E. coli28.5 kDaUnbound
FRGVf (FRG-I)
Ping-pong		V. fischeri24.6 kDaBound
FreVf
Sequential		V. fischeri25.5 kDaUnbound
FRDVh
Sequential		V. harveyi26.7 kDaUnbound
FRPVh
Ping-pong		V. harveyi26.3 kDaBound
ActVB (FRD-II)
Ping-pong		Streptomyces coelicolor2 × 18.0 kDaBound
SnaC (FRD-I, PIIB)
Ping-pong		S. pristinaspiralis28.0 kDaBound
EmoB (cB’)
Ping-pong		Chelatovorans
multitrophicus		2 × 25.0 kDaBound
DszD (FRD-III)
Sequential		Rhodococcus
erythropolis		4 × 22.5 kDaUnbound
FRDAa (cB)
Sequential		Aminobacter
aminovorans		2 × 44.0 kDaBound
Table
Table 2. Calculated apparent KmFMN values for highly purified representative FRs from P. putida ATCC 17453 and V. fischeri ATCC 7744 tested both as single-enzyme assays (S) and coupled-enzyme assays with 2,5-DKCMO (C:+2,5-MO), 3,6-DKCMO (C:+3,6-MO), and LuxAB luciferase (C:+LuxAB) and, in each case, 1mM of the biooxidisable ketone (rac)-bicyclo[3.2.0]hept-2-en-6-one. The equivalent reported single-enzyme KmFMN values for FreEc and FRDAa are 0.8 μM [35] and 1.0 μM [40], respectively.
Table 2. Calculated apparent KmFMN values for highly purified representative FRs from P. putida ATCC 17453 and V. fischeri ATCC 7744 tested both as single-enzyme assays (S) and coupled-enzyme assays with 2,5-DKCMO (C:+2,5-MO), 3,6-DKCMO (C:+3,6-MO), and LuxAB luciferase (C:+LuxAB) and, in each case, 1mM of the biooxidisable ketone (rac)-bicyclo[3.2.0]hept-2-en-6-one. The equivalent reported single-enzyme KmFMN values for FreEc and FRDAa are 0.8 μM [35] and 1.0 μM [40], respectively.
Flavin Reductase
and Source		Apparent KmFMN (μM) Source of
Hydride Ion		Mechanism of
Hydride Ion
Transfer
Frp1 P. putida2.5 (S)
2.0 (C + 2,5-MO)
2.5 (C + 3,6-MO)		NADHSequential
Frp2 P. putida4.2 (S)
3.6 (C + 2,5-MO)
4.1 (C + 3,6-MO)		NADHSequential
FRGVf (V. fischeri)4.3 (S)
4.0 (C + LuxAB)		NADHPing-pong
FreVf (V. fischeri)2.5 (S)
2.6 (C + LuxAB)		NADHSequential
Table
Table 3. 120 min outcomes of the fd-TCMO-FR coupled-enzyme reactions with highly purified preparations of LuxAB luciferase, 2,5-DKCMO, and 3,6-DKCMO with FRs from P. putida (Frp1 and Frp2), V. fischeri (FRGVf and FreVf), A. aminovorans (FRDAa) and E. coli (FreEc). The structures of (+)k, (−)k, (+)2l, (−)2l, (+)3l, and (−)3l are shown in Figure 2. (n) = native FR, (n.n) = non-native FR.
Table 3. 120 min outcomes of the fd-TCMO-FR coupled-enzyme reactions with highly purified preparations of LuxAB luciferase, 2,5-DKCMO, and 3,6-DKCMO with FRs from P. putida (Frp1 and Frp2), V. fischeri (FRGVf and FreVf), A. aminovorans (FRDAa) and E. coli (FreEc). The structures of (+)k, (−)k, (+)2l, (−)2l, (+)3l, and (−)3l are shown in Figure 2. (n) = native FR, (n.n) = non-native FR.
fd-TCMO and Partner FR in Coupled-enzyme
Reaction		Residual Ketone
(mM) Remaining
after 120 min		Percentage and
Ratio of the Ketone
(k) Enantiomers Biotransformed		Predominant Regioisomeric Lactones (l) Formed Expressed as ee%
(+)k(−)k(+):(−)(+)2l(−)2l(+)3l(−)3l
LuxAB luciferase
Frp1 (n.n)0.7041.618.42.26:1–>98–16
Frp2 (n.n)0.6943.018.42.33:1–>98–18
FRGVf (n)0.7534.215.02.28:1–96–18
FreVf (n)0.7942.818.82.28:1–>99–16
FRDA (n.n)0.6252.222.82.29:1–>98–16
FreEc (n.n)0.6154.224.02.25:1–>99–18
2,5-DKCMO
Frp1 (n)0.2481.869.71.17:184–>99–
Frp2 (n)0.2084.374.81.13:186–>98–
FRGVf (n.n)0.2678.868.41.15:182–>98–
FreVf (n.n)0.2382.571.71.15:184–>98–
FRDAa (n.n)0.0610088.61.13:188–>99–
FreEc (n.n)0.0410092.01.09:190–>98–
3,6-DKCMO
Frp1 (n)0.5358.835.01.68:130–80–
Frp2 (n)0.5359.835.21.70:130–82–
FRGVf (n.n)0.5557.033.01.72:132–86–
FreVf (n.n)0.5261.035.41.72:132–84–
FRDAa (n.n)0.3877.645.01.72:134–82–
FreEc (n.n)0.3780.045.81.74:132–88–
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Willetts, A. Inter-Species Redox Coupling by Flavin Reductases and FMN-Dependent Two-Component Monooxygenases Undertaking Nucleophilic Baeyer–Villiger Biooxygenations. Microorganisms 2023, 11, 71. https://doi.org/10.3390/microorganisms11010071

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Willetts A. Inter-Species Redox Coupling by Flavin Reductases and FMN-Dependent Two-Component Monooxygenases Undertaking Nucleophilic Baeyer–Villiger Biooxygenations. Microorganisms. 2023; 11(1):71. https://doi.org/10.3390/microorganisms11010071

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Willetts, Andrew. 2023. "Inter-Species Redox Coupling by Flavin Reductases and FMN-Dependent Two-Component Monooxygenases Undertaking Nucleophilic Baeyer–Villiger Biooxygenations" Microorganisms 11, no. 1: 71. https://doi.org/10.3390/microorganisms11010071

APA Style

Willetts, A. (2023). Inter-Species Redox Coupling by Flavin Reductases and FMN-Dependent Two-Component Monooxygenases Undertaking Nucleophilic Baeyer–Villiger Biooxygenations. Microorganisms, 11(1), 71. https://doi.org/10.3390/microorganisms11010071

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