Next Article in Journal
The Steric Effect in Green Benzylation of Arenes with Benzyl Alcohol Catalyzed by Hierarchical H-beta Zeolite
Next Article in Special Issue
Screening and Comparative Characterization of Microorganisms from Iranian Soil Samples Showing ω-Transaminase Activity toward a Plethora of Substrates
Previous Article in Journal
Overcoming Stability Problems in Microwave-Assisted Heterogeneous Catalytic Processes Affected by Catalyst Coking
Previous Article in Special Issue
Comparative Analysis and Biochemical Characterization of Two Endo-β-1,3-Glucanases from the Thermophilic Bacterium Fervidobacterium sp.
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Catalysts
Volume 9
Issue 10
10.3390/catal9100868
catalysts-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Review

Cofactor F420-Dependent Enzymes: An Under-Explored Resource for Asymmetric Redox Biocatalysis

by
Mihir V. Shah
1,†,
James Antoney
1,2,†,
Suk Woo Kang
1,2,
Andrew C. Warden
1,
Carol J. Hartley
1,
Hadi Nazem-Bokaee
1,
Colin J. Jackson
1,2 and
Colin Scott
1,*
1
CSIRO Synthetic Biology Future Science Platform, Canberra 2601, Australia
2
Research School of Chemistry, Australian National University, Canberra 2601, Australia
*
Author to whom correspondence should be addressed.
†
M.V.S. and J.A. contributed equally to this review.
Catalysts 2019, 9(10), 868; https://doi.org/10.3390/catal9100868
Submission received: 20 September 2019 / Revised: 8 October 2019 / Accepted: 10 October 2019 / Published: 20 October 2019
(This article belongs to the Special Issue Novel Enzyme and Whole-Cell Biocatalysts)
Browse Figures
Versions Notes

Abstract

:
The asymmetric reduction of enoates, imines and ketones are among the most important reactions in biocatalysis. These reactions are routinely conducted using enzymes that use nicotinamide cofactors as reductants. The deazaflavin cofactor F420 also has electrochemical properties that make it suitable as an alternative to nicotinamide cofactors for use in asymmetric reduction reactions. However, cofactor F420-dependent enzymes remain under-explored as a resource for biocatalysis. This review considers the cofactor F420-dependent enzyme families with the greatest potential for the discovery of new biocatalysts: the flavin/deazaflavin-dependent oxidoreductases (FDORs) and the luciferase-like hydride transferases (LLHTs). The characterized F420-dependent reductions that have the potential for adaptation for biocatalysis are discussed, and the enzymes best suited for use in the reduction of oxidized cofactor F420 to allow cofactor recycling in situ are considered. Further discussed are the recent advances in the production of cofactor F420 and its functional analog FO-5′-phosphate, which remains an impediment to the adoption of this family of enzymes for industrial biocatalytic processes. Finally, the prospects for the use of this cofactor and dependent enzymes as a resource for industrial biocatalysis are discussed.

1. Introduction

Enzymes that catalyze the asymmetric reduction of activated double bonds are among the most important in biocatalysis, allowing access to chiral amines from imines (C=N), sec-alcohols from ketones C=O), and enantiopure products derived from enoates (C=C). To date, the reduction of imines, ketones and enoates has been achieved largely using enzymes that draw their reducing potential from the nicotinamide cofactors NADH and NADPH; e.g., imine reductases, ketoreductases and Old Yellow Enzymes [1,2,3,4]. However, there has been recent interest in an alternative reductive cofactor, cofactor F420 (8-hydroxy-5-deazaflavin) [5,6].
Cofactor F420 is a deazaflavin that is structurally similar to flavins (Figure 1), with a notable difference at position 5 of the isoalloxazine ring, which is a nitrogen in flavins and a carbon in deazaflavins. Additionally, while C-7 and C-8 are methylated in riboflavin, they are not in cofactor F420: C-7 is hydroxylated and C-8 is unsubstituted. These structural differences cause significant differences in the electrochemical properties of cofactor F420 and flavins: a −360–340 mV the redox mid-point potential of cofactor F420 is not only lower than that of the flavins (−205 mV to −220 mV), but it is also lower than that of the nicotinamides (−320 mV) [7]. Additionally, as a consequence of the substitution of N-5 for a carbon, cofactor F420 cannot form a semiquinone (Figure 1), which means that unlike other flavins, cofactor F420 can only perform two-electron reductions.
Cofactor F420 was originally described in methanogenic archaea, where it plays a pivotal role in methanogenesis [8,9]. Cofactor F420 has since been described in a range of soil bacteria supporting a range of metabolic activities, including catabolism of recalcitrant molecules (such as picric acid) and the production of secondary metabolites, such as antibiotics [7]. A comprehensive review of the biochemistry and physiological roles of cofactor F420 was recently published by Greening and coworkers [7]. This review considers the potential of F420-dependent enzymes in industrial biocatalysis, focusing on the enzyme families relevant to biocatalytic applications and the reactions that they catalysis. Cofactor recycling strategies and cofactor production are also discussed, with a focus on the prospects for achieving low-cost production at scale in the latter case.

2. Families of F420-Dependent Enzymes Relevant to Biocatalysis

With respect to their prospective biocatalytic applications, the two most important families of F420-dependent enzymes are the Flavin/Deazaflavin Oxidoreductase (FDOR) and Luciferase-Like Hydride Transferase (LLHT) families, albeit F420-dependent enzyme from other families have also been shown to have catalytic activities of interest (e.g., TomJ, the imine reducing flavin-dependent monooxygenase or OxyR, the tetracycline oxidoreductase) [10,11]. The FDOR and LLHT families are large and contain highly diverse flavin/deazaflavin-dependent enzymes. In both families, there are enzymes with preferences for flavins, such as flavin mononucleotide (FMN) and flavin adenine dinucleotide (FAD), as well as those that use cofactor F420 [12,13]. Moreover, there are F420-dependent FDORs that have been shown to be able to promiscuously bind FMN and use it in oxidation reactions [14]. In this section, the FDOR and LLHT families and the classes of reaction that they catalyze are discussed.

2.1. The FDOR Superfamily

The FDOR superfamily (PFAM Clan CL0336) can be broadly divided into two groups: the FDOR-As (which includes a sub-group called the FDOR-AAs) and the FDOR-Bs. The FDOR-As are restricted to Actinobacteria and Chloroflexi and to date no FDOR-As have been described that use cofactors other than F420 [7,12]. The FDOR-Bs are found in a broader range of bacterial genera than the FDOR-A enzymes, and in addition to F420-dependent enzymes, this group also includes heme oxygenases, flavin-sequestering proteins, pyridoxine 5’ oxidases and a number of proteins of unknown function [12,15,16,17]. Both groups of FDOR are highly diverse, with many homologs often found within a single bacterial genome (e.g., Mycobacterium smegmatis has 28 FDORs) [18]. In addition, the majority of the enzymes of this family are yet to be characterized with respect to either their biochemical or physiological function, and therefore the FDORs represent a currently under-explored source of enzymes for biocatalysis.
The FDOR enzymes share a characteristic split β-barrel fold that forms part of the cofactor-binding pocket. The majority of the protein sequences of enzymes currently identified as belonging to this family are small single-domain proteins. The topologies of the two FDOR subgroups are broadly similar (Figure 2), with the split-barrel core composed of 7–8 strands and with 4–5 helices interspersed. All FDOR-Bs studied so far have been demonstrated to be dimeric, with stands β2, β3, β5 and β6 making up the core of the dimer interface (Figure 2). In structures of full-length FDOR-As solved to date, the N-terminal helix (if present) lies on the opposite face of the beta sheet to that in FDOR-Bs. Thus, the N-terminus occupies part of the dimer interface region and prevents interaction between the sheets of adjacent monomers. In contrast to the FDOR-Bs, the oligomerization state of the FDOR-As is more varied. While a number of FDOR-As have been determined to be monomeric [18], the deazaflavin-dependent nitroreductase (DDN) from M. tuberculosis forms soluble aggregates through the amphipathic N-terminal helix [19]. DDN and the FDOR-AA subgroup have been shown to be membrane-associated [20,21,22], and FDOR-AAs have been associated with fatty acid metabolism [12]. No structures of FDOR-AAs have been solved to date.

2.2. The LLHT Family:

The LLHT family form part of the Luciferase-Like Monooxygenase family (PFAM PF00296). They adopt an (α/β)8 TIM-barrel fold with three insertion regions, IS1–4 (Figure 3). IS1 contains a short loop and forms part of the substrate cleft. IS2 contains two antiparallel β-strands, and IS3 contains a helical bundle at the C-terminus of the β-barrel and contains the remainder of the substrate-binding pocket (Figure 3). All structures solved to date from the LLHT family contain a non-prolyl cis peptide in β3 [23,24,25,26]. Recent phylogenetic reconstructions have shown that the F420-dependent LLHTs form two clades: the F420-dependent reductases and the F420-depented dehydrogenases [27]. The F420-reductases contain methylenetetrahydromethanopterin reductases (MERs), which catalyze the reversable, ring-opening cleavage of a carbon-nitrogen bond during the biosynthesis of folate in some archaea [28,29,30]. The F420-dependent dehydrogenases can be further divided into three subgroups. The first contains F420-dependent secondary alcohol dehydrogenases (ADFs) and the hydroxymycolic acid reductase from M. tuberculosis [31]. The second contains the F420-dependent glucose-6-phosphate dehydrogenases (FGDs) from Mycobacteria and Rhodococcus, while the third appear to be more general sugar-phosphate dehydrogenases [27]. In contrast to the heterodimeric structure of bacterial luciferase, the F420-dependent dehydrogenases form homodimers with the dimer interface burying a relatively large portion of the surface area of the monomers (≈2000 Å2, roughly 15% of the total surface area) [24,25,26]. A number of enzymes involved in the F420-dependent degradation of nitroaromatic explosives, such as picrate and 2,4-dinitroanisole, appear to belong to the LLHT family as well [32,33].

2.3. Cofactor F420-Dependent Reactions with Relevance to Biocatalysis

From the perspective of biocatalysis, cofactor F420-dependent enzymes catalyze a number of key reductions including the reduction of enoates, imines, ketones and nitro-groups (Table 1; Figure 4).
For enoate reductions, a small number of FDORs have been studied. However, the substrate range for most of these enzymes is yet to be fully elucidated. The ability of the mycobacterial FDORs to reduce activated C=C double bonds was first identified when DDN was shown to be responsible for activating the bicyclic nitroimidazole PA-824 in M. tuberculosis. These enzymes were then shown to also reduce enoates in aflatoxins, coumarins, furanocoumarins and quinones [6,12,14,16,34,35,36,37,38]. Recent studies have shown that these enzymes are promiscuous and can use cyclohexen-1-one, malachite green and a wide range of other activated ene compounds as substrates [35]. However, there have been a few FDOR studies to date that have examined their kinetic properties and stereospecificity. In one of these studies, FDORs from Mycobacterium hassiacum (FDR-Mha) and Rhodococcus jostii RHA1 (FDR-Rh1 and FDR-Rh2) were shown to reduce a range of structurally diverse enoates with conversions ranging from 12 to >99% and e.e. values of up to >99% [6]. Interestingly, it has been proposed that both the hydride and proton transfer from F420H2 in these reactions was directed to the same face of the activated double bond (Figure 5), which results in the opposite enantioselectivity compared to that of the FMN-dependent Old Yellow Enzyme family of enoate reductases [6]. This suggests that the F420-dependent FDORs may provide a stereocomplementary enoate reductase toolbox. However, other studies suggest that protonation of the substrate is mediated by solvent or an enzyme side-chain (as it is in Old Yellow Enzyme) [37]. Further structure/function studies are needed to fully understand the mechanistic diversity of this family of enzymes.
The LLHT family contains several enzymes with alcohol oxidase or ketoreductase activity (Table 1; Figure 4). The F420-dependent glucose-6-phosphate dehydrogenases of several species have been investigated [25,26,39]. Although an extensive survey of their substrate ranges has yet to be conducted, it has been demonstrated that glucose is a substrate for the Rhodococcus jostii RHA1 enzymes [26]. An F420-dependent alcohol dehydrogenase (ADH) from Methanogenium liminatans has been shown to catalyze the oxidation of the short chain aliphatic alcohols 2-propanol, 2-butanol and 2-pentanol (85, 49 and 23.1 s−1 kcat, 2.2, 1.2 and 7.2 mM KM respectively) [40], but it was unable to oxidize primary alcohols, polyols or secondary alcohols with more than five carbons. It is unclear whether these alcohol oxidations are reversible, but in the oxidative direction, these reactions provide enzymes that can be used to recycle reduced cofactor F420 (see Section 4). Alcohol oxidation can also be used to produce ketones as intermediates in biocatalytic cascades that can then be used in subsequent reactions, such those catalyzed by transaminases or amine dehydrogenases in chiral amine synthesis [1,41,42,43] or by ketoreductases or alcohol dehydrogenases in chiral sec-alcohol synthesis (i.e., deracemization or stereoinversion of sec-alcohols). This approach can be achieved in a one pot cascade if different cofactors are used for the oxidation and reduction (Figure 6) [44].
At least one F420-dependent ketoreductase has been described. The mycobacterial F420-dependent phthiodiolone ketoreductase catalyzes a key reduction in the production of phthiocerol dimycocerosate, a diacylated polyketide found in the mycobacterial cell wall [45]. Although the physiological role of this enzyme has been elucidated, biochemical studies of the catalytic properties and substrate range are required to assess this enzymes’ potential for use as a biocatalyst.
F420-dependent enzymes have also been shown to reduce imines (Table 1; Figure 4). An FDOR fromr Streptomyces tateyamensis (TpnL) is responsible for the reduction of dehydropiperidine in the piperidine-containing series a group of thiopeptide antibiotics produced in this bacterium (Figure 4). TpnL was identified as the F420-dependent dehydropiperidine reductase responsible for the reduction of dehydropiperidine ring in thiostrepton A to produce the piperidine ring in the core macrocycle of thiostrepton A [45]. TpnL activity was affected by substrate inhibition at concentrations higher than 2 μM of thiostrepton A, preventing the measurement of the KM, but its kcat/KM was measured at 2.80 × 104 M−1 S−1 [45]. The substrates for phthiodiolone ketoreductase and TpnL are large secondary metabolites and, as yet, it is unclear if it will accept smaller substrates or substrates with larger/smaller heterocycles (e.g., dehydropyrroles).
Another F420-dependent imine reductase (TomJ) has been described from Streptomyces achromogenes that reduces the imine in 4-ethylidene-3,4-dehydropyrrole-2-carboxylic acid during the production of the secondary metabolite tomaymycin, which has been shown to have potentially interesting pharmaceutical properties [11]. Additionally, the reduction of a prochiral dihydropyrrole to a pyrrole is a reaction with a number of biocatalytic applications [5].
Nitroreductases have the potential application in the reduction of a prochiral nitro group to form a chiral amine [46]. The LLHT family F420-dependent nitroreductase Npd from Rhodococcus catalyzes the two-electron reduction of two nitro groups in picric acid during catabolism of the explosive TNT (Table 1; Figure 4) [47]. While this stops short of reducing the nitro group to an amine, this catalytic activity may contribute to a reductive cascade that achieves this conversion.
The final class of reaction for consideration in this review is the unusual, reversable ring-opening/closing reaction catalyzed by the MERs (Figure 4; Table 1). This reaction is required for folate biosynthesis in some archaea [23,28,29,30]. However, ring-closing reactions of this type could be used for producing N-containing heterocycles, which are intermediates in the synthesis of numerous pharmaceuticals [48,49]. The promiscuity of the MERs has not yet been investigated, and so the potential to re-engineer these enzymes is not fully understood.

3. Cofactor Recycling for Cofactor F420

Cofactor recycling is essential for the practical application of the F420-dependent enzymatic processes in biocatalysis. There are various strategies for cofactor regeneration for NADH and NADPH, including enzymatic, chemical, electrochemical and photochemical methods [52]. In this section, the potential enzymes for the regeneration of cofactor F420 are discussed. As most of the industrially relevant F420-dependent reactions are asymmetric reductions, F420-dependent oxidases are required for cofactor regeneration. Figure 7 shows the characterized enzymes that catalyze F420-dependent oxidations that could be applied in cofactor F420 reduction.
Emulating methods developed for nicotinamide cofactors, both formate dehydrogenase (FDH) and glucose 6-phosphate dehydrogenase (G6PD) enzymes are attractive enzymatic routes for cofactor reduction both in vitro [53,54,55,56] and in vivo [57,58]. Fortunately, F420-dependent G6PDs and FDHs have been identified and characterized. The F420-dependent G6PD from Mycobacteria (FGD) is one potential cofactor F420-recycling enzyme. FGD is the only enzyme in these bacteria known to reduce oxidized cofactor F420. The intracellular concentration of G6P in Mycobacteria is up to 100-fold higher than it is in E. coli, which provides a ready source of reducing power for F420-dependent reduction reactions [59]. FGD from Rhodococcus jostii and Mycobacterium smegmatis have been studied and expressed in E. coli, both the enzymes were stable in in vitro assays [26,39,60]. Both FGDs have been expressed in engineered E. coli producing cofactor F420 together with FDORs [38,59] FGDs have been shown to efficiently regenerate reduced cofactor F420 both in vivo and in vitro. However, the cost of the glucose-6-phosphate and the need to separate reaction products from the accumulated FGD byproduct (6-phosphoglucono-d-lactone) may prove to be impediments for the adoption of FGD as a recycling system for cofactor F420 in the in vitro biotransformations.
Formate is an excellent reductant for cofactor recycling, with FDH-dependent cofactor reduction yielding carbon dioxide, a volatile byproduct that can be easily removed from the reaction mixture, thereby simplifying the downstream processing of the product of interest. Additionally, formate is a low-cost reagent, leading to favorable process economics. Most methanogens have the capability to use formate as sole electron donor using F420-dependent formate dehydrogenase [61]. The soluble F420-dependent FDH from Methanobacterium formicium has been expressed in E. coli [62], purified and studied in vitro with the reduction of 41.2 μmol of F420 min−1 mg−1 of FDH, with non-covalently bound FAD required for optimal activity [8]. Methanobacterium ruminantium FDH reduces cofactor F420 at a much slower rate than M. formicium: 0.11 μmol of F420 min−1 mg−1 of FDH [8]. As yet, the use of F420-dependnt FDHs for in vitro cofactor recycling has been sparsely studied. However, as these enzymes are soluble and can be heterologously expressed, they represent a promising system for use in cofactor F420-dependent biocatalytic processes.
Another potential recycling system for cofactor F420 is the F420:NADPH oxidoreductase (Fno), which couples the reduction of cofactor F420 with oxidation of NADPH. Methanogenic archaea use this enzyme to transfer reducing equivalents from hydrogenases to produce NADPH via F420, while in bacteria it functions in the opposite direction, that is, to provide the cell with reduced F420 via NADPH [63]. Fno is also required for the production of reduced F420 for tetracycline production in Streptomyces [63]. The Fno enzymes from the thermophilic bacteria Thermobifida fusca and the thermophilic archaeon Archeoglobus fulgidus have been expressed in E. coli [64,65]. These enzymes are thermostable, with their highest activity observed at 65 °C. As the redox midpoint potentials of NADP and cofactor F420 are very similar, it is perhaps unsurprising that pH has a significant influence on the equilibrium of the reaction, with the reduction of NADP+ favored at high pH (8–10) and the reduction of F420 favored at low pH (4–6) [64,65]. The Fno Streptomyces griseus has also been purified and characterized, and also displayed pH-dependent reaction directionality [66]. Fno may be an excellent enzyme for the in vivo reduction of cofactor F420, where NADPH would be provided from central metabolism. However, for its use as a cofactor F420 recycling enzyme in vitro, Fno would need to be coupled with an NADPH regenerating enzyme, such as an NADPH-dependent formate dehydrogenase [67]. This added complexity and cost may limit the use of Fno-dependent cofactor F420 recycling in vivo.
Hydrogenotrophic archaea, including methanogens and sulfate-reducing archaea, possess an essential, cofactor F420-dependent hydrogenase (FhrAGB) [68,69,70,71]. These nickel/iron enzymes could potentially be used in vivo to allow the direct H2-dependent reduction of cofactor F420. However, as these heterododecameric enzymes have complex cofactor requirements (four [4Fe 4S] clusters, and NiFe center and FAD), are oxygen-sensitive and tend to aggregate [71], it is unclear if they can be made suitable for in vitro use.

4. Cofactor Production

The lack of a scalable production system for cofactor F420 has been noted as a major impediment to the adoption of F420-dependent enzymes by industry [5]. Cofactor F420 is available as a research reagent (http://www.gecco-biotech.com/), but its production at scale is not yet economic. In fact, most research laboratories with an interest in cofactor F420-dependent enzymes synthesize and purify the cofactor themselves using slow-growing F420 producing microorganisms, most commonly methanogens and actinobacteria (Table 2). The economic production of cofactor F420 at large scale is not feasible using natural producers as they are ill-suited to industrial fermentation and generally lack the genetic tools required to improve cofactor F420 yield.
Recently, there have been significant advances towards the scalable production of the cofactor for F420-dependent enzymes. M. smegmatis has been engineered to overexpress the biosynthetic genes for cofactor F420 production, leading to a substantial improvement in yields (Table 2) [72]. However, M. smegmatis is not ideally suited as a fermentation organism as it is slow growing, forms clumps during cultivation and is not recognized as GRAS (generally regarded as safe). More recently, the biosynthetic pathway for cofactor F420 has been successfully transplanted to E. coli [59], allowing the heterologous production of the cofactor at levels similar to those of the natural F420 producers (Table 2) [59], accumulated to 0.38 μmol of F420 per gram of dry cells.
There is scope to further improve the production of F420 in E. coli. Cofactor F420 does not appear to be toxic to E. coli [59], which suggests that there is little interaction between F420 and the enzymes E. coli (although this is yet to be confirmed experimentally). The thermodynamics of cofactor F420 production are favorable (Appendix A), suggesting that there are no major thermodynamic impediments to improving yield. Interestingly, the first dedicated step of cofactor F420 production (catalyzed by CofC/FbiD) is not energetically favorable and may consequently be sensitive to intracellular metabolite concentrations. In addition to the engineering considerations that this may impose, it may also be responsible for the biochemical diversity of this step in different microorganisms. In different microbes, the CofC/FbiD-dependent step uses 2-phospholactate [75], 3-phosphoglycerate [76] or phosphoenolpyruvate [59] as a substrate, which may reflect the relative abundance of those metabolites in various bacteria and archaea and the thermodynamic constraints on this step.
Another recent advance is the production of a synthetic analog of cofactor F420, called FO-5′-phosphate (FOP). FOP was derived from FO, the metabolic precursor of cofactor F420, which is phosphorylated using an engineered riboflavin kinase [38]. FOP has also been shown to function as an active cofactor for cofactor F420-dependent enzymes activities, albeit there is a penalty in the rates of these reactions [38]. Drenth and coworkers prepared FO by chemical synthesis, using a method developed by Hossain et al. [77]. However, it is likely that the engineered kinase for the phosphorylation of FO could be introduced into an organism that over-produces FO allowing for the production of FOP by fermentation. This semisynthetic pathway would have the advantage that it needs only two biosynthetic steps, instead of the four steps needed for cofactor F420 production, and demands less metabolic input from the native host metabolism (e.g., no glutamate is required) [38]. The production of FOP also opens the possibility of making deazaflavin analogs of FMN and FAD, which would be electrochemically more like F420 than flavins, but may still bind FMN and FAD- dependent enzymes and potentially allow access to new chemistry with already well-characterized enzymes.

5. Prospects

Reduced cofactor F420 is electrochemically well suited for biocatalytic applications, and the small number of F420-dependent enzymes characterized to date show promise as potential biocatalysts (as discussed above). However, before these enzymes can be widely and effectively used as biocatalysts, further research is needed to better characterize them as the biochemistry of cofactor F420-dependent enzymes remains under-explored. The LLHT and FDOR families are a rich source of highly diverse enzymes with considerable potential for biocatalysis, albeit much of the research to date has focused on the physiological roles of these enzymes, rather than their in vitro enzymology. Although some of these enzymes have been shown to have small molecule substrates, those involved with secondary metabolite biosynthesis tend to act on high molecular weight substrates and it is not yet clear whether they will accept lower molecular weight molecules.
To be cost competitive, cofactor F420 needs to have effective recycling systems. The enzymes for cofactor recycling have already been identified, although there have been a few studies investigating their performance in this role. Moreover, alternative cofactor recycling strategies, such electrochemical or photochemical recycling, have not yet been investigated for cofactor F420. The production of cofactor F420 at scale and at low cost remains a roadblock for the use of these enzymes by industry. However, considerable progress has been made on this front in the last few years and it is likely that low cost cofactor F420, or F420 surrogates, will soon be available. Additionally, the availability of F420-producing bacteria with tools for facile genetic manipulation, along with a growing number of empirically determined protein structures, opens up the prospect of improving this class of enzymes using in vitro evolution and rational design. It is notable that there is still some uncertainty concerning the mechanistic detail of F420-dependent reactions, which need to be addressed through a detailed structure/function analysis to enable a rational design of these enzymes.

Supplementary Materials

The following are available online at https://www.mdpi.com/2073-4344/9/10/868/s1.

Author Contributions

Writing—original draft preparation, M.V.S., J.A., S.W.K., H.N.-B., C.J.J. and C.S.; writing—review and editing, M.V.S., J.A., S.W.K., H.N.-B., A.C.W., C.J.H., C.J.J. and C.S.

Funding

M.V.S. and H.N.-B. are funded by the CSIRO Synthetic Biology Future Science Platform. J.A. is funded by the Australian Government Research Training Program (AGRTP) and S.W.K. is funded by the Korean Institute of Science and Technology and the AGRTP.

Conflicts of Interest

The authors declare no conflict of interest.

Abbreviations

5AD: 5′-Deoxyadenosine; 5ARPD: 5-amino-6-(d-ribitylamino)uracil; 5ARPD4HB: 5-amino-5-(4-hydroxybenzyl)-6-(d-ribitylimino)-5,6-dihydrouracil; dF420-0: Dehydro coenzyme F420-0 (oxidized); EPPG: Enolpyruvyl-diphospho-5′-guanosine; Fo: 7,8-Didemethyl-8-hydroxy-5-deazariboflavin; F420-0: Coenzyme F420-0 (oxidized); F420-1: Coenzyme F420-1 (oxidized); F420-2: Coenzyme F420-2 (oxidized); F420-3: Coenzyme F420-3 (oxidized); FMN: Flavin mononucleotide (oxidized); FMNH2: Flavin mononucleotide (reduced); GDP: Guanosine diphosphate; GMP: Guanosine monophosphate; Glu: l-Glutamate; GTP: Guanosine triphosphate; H+: Proton; ImiAce: 2-iminoacetate or Dehydroglycine; Met: l-Methionine; NH4: Ammonium; PEP: Phosphoenolpyruvate; Pi: Phosphate; PPi: Diphosphate; SAMe: S-Adenosyl-l-methionine; Tyr: l-Tyrosine.

Appendix A. Thermodynamics of F420 Biosynthesis

The thermodynamic properties of each of the steps in cofactor F420 biosynthesis were estimated to evaluate the feasibility of increasing the production of the cofactor in an engineered microorganism. The pathway assembled by Bashiri et al. [59] in E. coli was used (i.e., PEP was used as substrate for CofC). The standard transformed Gibbs free energy (ΔrGt) of each step were calculated under the physiological conditions (25 °C, pH 7, and ionic concentration of 0.25 M) as described elsewhere [78,79]. The overall Gibbs free energy (ΔGt) was then calculated by summing up all individual ΔrGt (Table A1). The Gibbs free energy of metabolite formation (ΔfG) for each metabolite in the pathway was obtained (Supplementary Information) from comprehensive lists of metabolites whose ΔfG were estimated using a group contribution method [80,81]. The ΔfG for each metabolite was then converted into its transformed type (ΔfGt) method of Alberty [78]. The data were collected from relevant biochemical databases and the literature for any metabolite with missing ΔfG [82,83,84]. Owing to possessing different protonation states, the inconsistencies in ΔfG of certain metabolites such as the glutamates in F420-n among databases and the literature are inevitable. Thus, ΔrGt for reactions containing metabolites with varying ΔfG were calculated considering the differences in their ΔfG leading to the generation of a total of four sets of ΔrGt. Finally, the mean and standard deviations were calculated for these sets to yield the variation in each reaction as well as in the overall pathway (Table A1).
The data shown in Table A1 confirms that the overall cofactor F420 biosynthesis pathway is thermodynamically feasible under the given conditions. However, certain steps in this pathway impose a thermodynamic barrier with respect to the physiological conditions examined. For example, CofC seems to be one of the major thermodynamically unfavorable steps in the whole pathway possibly due to the energy-dependent synthesis of EPPG, one of the precursors for making F420. CofG/H combined appears to be the most thermodynamically favorable step in the whole pathway driving the biosynthesis of F0, the other key precursor for F420 biosynthesis. Interestingly, the formation of F420-2 molecule seems to be the most favorable step among other F420 molecules downstream of the pathway. It should be noted that the thermodynamic calculations were only performed up to three steps of F420 molecule production (i.e., F420-3) largely because of the high levels of inconsistencies of the data available for ΔfG of higher F420 molecules.
Table
Table A1. Standard transformed Gibbs free energy of reaction (ΔrGt), for the F420 biosynthesis pathway, calculated based on Gibbs free energy of metabolite formation (ΔfGt) calculated at 25 °C, pH of 7, and ionic concentration of 0.25 M.
Table A1. Standard transformed Gibbs free energy of reaction (ΔrGt), for the F420 biosynthesis pathway, calculated based on Gibbs free energy of metabolite formation (ΔfGt) calculated at 25 °C, pH of 7, and ionic concentration of 0.25 M.
EnzymeReaction aΔrGt (kJ) b
CofC/FbiDPEP + GTP → EPPG + PPi d+71.27(±67)
CofG/FbiC5ARPD + Tyr + SAMe → 5ARPD4HB + ImAcet + Met + 5AD−1192.39(±0) c
CofH/FbiC5ARPD4HB + SAMe → FO + NH4+ + Met + 5AD+71.90(±36) c
CofD/FbiAFO + EPPG → dF420-0 + GMP−31.3(±128)
CofX/FbiBdF420-0 + FMNH2 → F420-0 + FMN−74.59(±87)
CofE/FbiBF420-0 + GTP + Glu → F420-1 + GDP + Pi−7.50(±24)
CofE/FbiBF420-1 + GTP + Glu → F420-2 + GDP + Pi−39.44(±35)
CofE/FbiBF420-2 + GTP + Glu → F420-3 + GDP + Pi−21.99(±38)
OverallPEP + 5ARPD + Tyr + (2) SAMe + FMNH2 + (3) Glu + (4) GTP−1224.05(±82)
→ F420-3 + (2) Met + (2) 5AD + ImAcet + NH4+ + FMN + (3) GDP + (3) Pi + GMP + PPi
a For simplicity, protons were omitted in these equations and subsequent calculations as the ΔfGt of a proton under the set conditions is ~0.08 kJ. However, all ΔrGt calculations are based on a balanced equation. b The mean values of four sets and their standard deviations in parenthesis shown for each reaction. c ΔfG of 5ARPD4HB has only been reported in MetaCyc inferred by computational analysis. Including it in the calculations of ΔrGt for CofG and CofH results in −225.88(±0) and −894.62(±36), respectively. d Hydrolysis of PPi (H3P2O73− + H2O → 2 HPO42− + H+) yields a ΔrGt of ~17 kJ/mole, resulting in less than 2% change in the overall ΔrGt.

References

  1. Patil, M.D.; Grogan, G.; Bommarius, A.; Yun, H. Oxidoreductase-catalyzed synthesis of chiral amines. ACS Catal. 2018, 8, 10985–11015. [Google Scholar] [CrossRef]
  2. Toogood, H.S.; Scrutton, N.S. New developments in ‘ene’-reductase catalysed biological hydrogenations. Curr. Opin. Chem. Biol. 2014, 19, 107–115. [Google Scholar] [CrossRef] [PubMed]
  3. Cosgrove, S.C.; Brzezniak, A.; France, S.P.; Ramsden, J.I.; Mangas-Sanchez, J.; Montgomery, S.L.; Heath, R.S.; Turner, N.J. Imine reductases, reductive aminases, and amine oxidases for the synthesis of chiral amines: Discovery, characterization, and synthetic applications. In Enzymes in Synthetic Biology; Scrutton, N., Ed.; Elsevier Academic Press Inc.: San Diego, CA, USA, 2018; Volume 608, pp. 131–149. [Google Scholar]
  4. Bai, D.Y.; He, J.Y.; Ouyang, B.; Huang, J.; Wang, P. Biocatalytic asymmetric synthesis of chiral aryl alcohols. Prog. Chem. 2017, 29, 491–501. [Google Scholar]
  5. Taylor, M.; Scott, C.; Grogan, G. F-420-dependent enzymes-potential for applications in biotechnology. Trends Biotechnol. 2013, 31, 63–64. [Google Scholar] [CrossRef]
  6. Mathew, S.; Trajkovic, M.; Kumar, H.; Nguyen, Q.-T.; Fraaije, M.W. Enantio- and regioselective ene-reductions using F420H2-dependent enzymes. Chem. Commun. 2018, 54, 11208–11211. [Google Scholar] [CrossRef]
  7. Greening, C.; Ahmed, F.H.; Mohamed, A.E.; Lee, B.M.; Pandey, G.; Warden, A.C.; Scott, C.; Oakeshott, J.G.; Taylor, M.C.; Jackson, C.J. Physiology, biochemistry, and applications of F420- and Fo-dependent redox reactions. Microbiol. Mol. Biol. Rev. 2016, 80, 451–493. [Google Scholar] [CrossRef]
  8. Tzing, S.F.; Bryant, M.P.; Wolfe, R.S. Factor 420-dependent pyridine nucleotide-linked formate metabolism of Methanobacterium ruminantium. J. Bacteriol. 1975, 121, 192–196. [Google Scholar]
  9. Eirich, L.D.; Vogels, G.D.; Wolfe, R.S. Distribution of coenzyme F420 and properties of its hydrolytic fragments. J. Bacteriol. 1979, 140, 20–27. [Google Scholar]
  10. Wang, P.; Bashiri, G.; Gao, X.; Sawaya, M.R.; Tang, Y. Uncovering the enzymes that catalyze the final steps in oxytetracycline biosynthesis. J. Am. Chem. Soc. 2013, 135, 7138–7141. [Google Scholar] [CrossRef]
  11. Li, W.; Chou, S.C.; Khullar, A.; Gerratana, B. Cloning and characterization of the biosynthetic gene cluster for tomaymycin, an SJG-136 monomeric analog. Appl. Environ. Microbiol. 2009, 75, 2958–2963. [Google Scholar] [CrossRef]
  12. Ahmed, F.H.; Carr, P.D.; Lee, B.M.; Afriat-Jurnou, L.; Mohamed, A.E.; Hong, N.-S.; Flanagan, J.; Taylor, M.C.; Greening, C.; Jackson, C.J. Sequence–structure–function classification of a catalytically diverse oxidoreductase superfamily in mycobacteria. J. Mol. Biol. 2015, 427, 3554–3571. [Google Scholar] [CrossRef] [PubMed]
  13. Selengut, J.D.; Haft, D.H. Unexpected abundance of coenzyme F420-dependent enzymes in Mycobacterium tuberculosis and other actinobacteria. J. Bacteriol. 2010, 192, 5788–5798. [Google Scholar] [CrossRef] [PubMed]
  14. Lapalikar, G.V.; Taylor, M.C.; Warden, A.C.; Onagi, H.; Hennessy, J.E.; Mulder, R.J.; Scott, C.; Brown, S.E.; Russell, R.J.; Easton, C.J.; et al. Cofactor promiscuity among F420-dependent reductases enables them to catalyse both oxidation and reduction of the same substrate. Catal. Sci. Technol. 2012, 2, 1560–1567. [Google Scholar] [CrossRef]
  15. Harold, L.K.; Antoney, J.; Ahmed, F.H.; Hards, K.; Carr, P.D.; Rapson, T.; Greening, C.; Jackson, C.J.; Cook, G.M. FAD-sequestering proteins protect Mycobacteria against hypoxic and oxidative stress. J. Biol. Chem. 2019, 294, 2903–2912. [Google Scholar] [CrossRef] [PubMed]
  16. Ahmed, F.H.; Mohamed, A.E.; Carr, P.D.; Lee, B.M.; Condic-Jurkic, K.; O’Mara, M.L.; Jackson, C.J. Rv2074 is a novel F420H2-dependent biliverdin reductase in Mycobacterium tuberculosis. Protein Sci. 2016, 25, 1692–1709. [Google Scholar] [CrossRef] [PubMed]
  17. Mashalidis, E.H.; Mukherjee, T.; Śledź, P.; Matak-Vinković, D.; Boshoff, H.I.; Abell, C.; Barry, C.E. Rv2607 from Mycobacterium tuberculosis is a pyridoxine 5′-phosphate oxidase with unusual substrate specificity. PLoS ONE 2011, 6, e27643. [Google Scholar] [CrossRef] [PubMed]
  18. Taylor, M.C.; Jackson, C.J.; Tattersall, D.B.; French, N.; Peat, T.S.; Newman, J.; Briggs, L.J.; Lapalikar, G.V.; Campbell, P.M.; Scott, C.; et al. Identification and characterization of two families of F420H2-dependent reductases from Mycobacteria that catalyse aflatoxin degradation. Mol. Microbiol. 2010, 78, 561–575. [Google Scholar] [CrossRef]
  19. Cellitti, S.E.; Shaffer, J.; Jones, D.H.; Mukherjee, T.; Gurumurthy, M.; Bursulaya, B.; Boshoff, H.I.; Choi, I.; Nayyar, A.; Lee, Y.S.; et al. Structure of DDN, the deazaflavin-dependent nitroreductase from Mycobacterium tuberculosis involved in bioreductive activation of PA-824. Structure 2012, 20, 101–112. [Google Scholar] [CrossRef]
  20. De Souza, G.A.; Leversen, N.A.; Målen, H.; Wiker, H.G. Bacterial proteins with cleaved or uncleaved signal peptides of the general secretory pathway. J. Proteom. 2011, 75, 502–510. [Google Scholar] [CrossRef] [Green Version]
  21. He, Z.; De Buck, J. Cell wall proteome analysis of Mycobacterium smegmatis strain mc2 155. BMC Microbiol. 2010, 10, 121. [Google Scholar] [CrossRef]
  22. Sinha, S.; Kosalai, K.; Arora, S.; Namane, A.; Sharma, P.; Gaikwad, A.N.; Brodin, P.; Cole, S.T. Immunogenic membrane-associated proteins of Mycobacterium tuberculosis revealed by proteomics. Microbiology 2005, 151, 2411–2419. [Google Scholar] [CrossRef] [PubMed]
  23. Aufhammer, S.W.; Warkentin, E.; Ermler, U.; Hagemeier, C.H.; Thauer, R.K.; Shima, S. Crystal structure of methylenetetrahydromethanopterin reductase (MER) in complex with coenzyme F420: Architecture of the F420/FMN binding site of enzymes within the nonprolyl cis-peptide containing bacterial luciferase family. Protein Sci. 2005, 14, 1840–1849. [Google Scholar] [CrossRef] [PubMed]
  24. Aufhammer, S.W.; Warkentin, E.; Berk, H.; Shima, S.; Thauer, R.K.; Ermler, U. Coenzyme binding in F420-dependent secondary alcohol dehydrogenase, a member of the bacterial luciferase family. Structure 2004, 12, 361–370. [Google Scholar] [CrossRef] [PubMed]
  25. Bashiri, G.; Squire, C.J.; Moreland, N.J.; Baker, E.N. Crystal structures of F420-dependent glucose-6-phosphate dehydrogenase FGD1 involved in the activation of the anti-tuberculosis drug candidate PA-824 reveal the basis of coenzyme and substrate binding. J. Biol. Chem. 2008, 283, 17531–17541. [Google Scholar] [CrossRef] [PubMed]
  26. Nguyen, Q.T.; Trinco, G.; Binda, C.; Mattevi, A.; Fraaije, M.W. Discovery and characterization of an F420-dependent glucose-6-phosphate dehydrogenase (Rh-FGD1) from rhodococcus jostii rha1. Appl. Microbiol. Biotechnol. 2017, 101, 2831–2842. [Google Scholar] [CrossRef] [PubMed]
  27. Mascotti, M.L.; Kumar, H.; Nguyen, Q.-T.; Ayub, M.J.; Fraaije, M.W. Reconstructing the evolutionary history of F420-dependent dehydrogenases. Sci. Rep. 2018, 8, 17571. [Google Scholar] [CrossRef] [PubMed]
  28. Ceh, K.; Demmer, U.; Warkentin, E.; Moll, J.; Thauer, R.K.; Shima, S.; Ermler, U. Structural basis of the hydride transfer mechanism in F420-dependent methylenetetrahydromethanopterin dehydrogenase. Biochemistry 2009, 48, 10098–10105. [Google Scholar] [CrossRef]
  29. Shima, S.; Warkentin, E.; Grabarse, W.; Sordel, M.; Wicke, M.; Thauer, R.K.; Ermler, U. Structure of coenzyme F420 dependent methylenetetrahydromethanopterin reductase from two methanogenic archaea. J. Mol. Biol. 2000, 300, 935–950. [Google Scholar] [CrossRef]
  30. Vaupel, M.; Thauer, R.K. Coenzyme F420 dependent N-5, N-10-methylenetetrahydromethanopterin reductase (MER) from Methanobacterium thermautotrophicum strain marburg: Cloning, sequencing, transcriptional analysis and functional expression in Escherichia coli of the mer gene. Eur. J. Biochem. 1995, 231, 773–778. [Google Scholar]
  31. Purwantini, E.; Mukhopadhyay, B. Rv0132c of Mycobacterium tuberculosis encodes a coenzyme F420-dependent hydroxymycolic acid dehydrogenase. PLoS ONE 2013, 8, e81985. [Google Scholar] [CrossRef]
  32. Fida, T.T.; Palamuru, S.; Pandey, G.; Spain, J.C. Aerobic biodegradation of 2,4-dinitroanisole by Nocardioides sp. Strain js1661. Appl. Environ. Microbiol. 2014, 80, 7725–7731. [Google Scholar] [CrossRef] [PubMed]
  33. Ebert, S.; Rieger, P.G.; Knackmuss, H.J. Function of coenzyme f420 in aerobic catabolism of 2,4,6-trinitrophenol and 2,4-dinitrophenol by Nocardioides simplex FJ2-1A. J. Bacteriol. 1999, 181, 2669–2674. [Google Scholar] [PubMed]
  34. Lapalikar, G.V.; Taylor, M.C.; Warden, A.C.; Scott, C.; Russell, R.J.; Oakeshott, J.G. F420H2-dependent degradation of aflatoxin and other furanocoumarins is widespread throughout the Actinomycetales. PLoS ONE 2012, 7, e30114. [Google Scholar] [CrossRef] [PubMed]
  35. Greening, C.; Jirapanjawat, T.; Afroze, S.; Ney, B.; Scott, C.; Pandey, G.; Lee, B.M.; Russell, R.J.; Jackson, C.J.; Oakeshott, J.G.; et al. Mycobacterial F420H2-dependent reductases promiscuously reduce diverse compounds through a common mechanism. Front. Microbiol. 2017, 8. [Google Scholar] [CrossRef]
  36. Gurumurthy, M.; Rao, M.; Mukherjee, T.; Rao, S.P.S.; Boshoff, H.I.; Dick, T.; Barry, C.E.; Manjunatha, U.H. A novel F420-dependent anti-oxidant mechanism protects Mycobacterium tuberculosis against oxidative stress and bactericidal agents. Mol. Microbiol. 2013, 87, 744–755. [Google Scholar] [CrossRef]
  37. Mohamed, A.E.; Ahmed, F.H.; Arulmozhiraja, S.; Lin, C.Y.; Taylor, M.C.; Krausz, E.R.; Jackson, C.J.; Coote, M.L. Protonation state of F420H2 in the prodrug-activating deazaflavin dependent nitroreductase (DDN) from Mycobacterium tuberculosis. Mol. BioSys. 2016, 12, 1110–1113. [Google Scholar] [CrossRef]
  38. Drenth, J.; Trajkovic, M.; Fraaije, M.W. Chemoenzymatic synthesis of an unnatural deazaflavin cofactor that can fuel F420-dependent enzymes. ACS Catal. 2019, 9, 6435–6443. [Google Scholar] [CrossRef]
  39. Purwantini, E.; Daniels, L. Molecular analysis of the gene encoding F420-dependent glucose-6-phosphate dehydrogenase from Mycobacterium smegmatis. J. Bacteriol. 1998, 180, 2212–2219. [Google Scholar]
  40. Bleicher, K.; Winter, J. Purification and properties of F420-and NADP+-dependent alcohol dehydrogenases of Methanogenium liminatans and Methanobacterium palustre, specific for secondary alcohols. Europ. J. Biochem. 1991, 200, 43–51. [Google Scholar] [CrossRef]
  41. Knaus, T.; Cariati, L.; Masman, M.F.; Mutti, F.G. In vitro biocatalytic pathway design: Orthogonal network for the quantitative and stereospecific amination of alcohols. Org. Biomol. Chem. 2017, 15, 8313–8325. [Google Scholar]
  42. Guo, F.; Berglund, P. Transaminase biocatalysis: Optimization and application. Green Chem. 2017, 19, 333–360. [Google Scholar] [CrossRef]
  43. Adams, J.P.; Brown, M.J.B.; Diaz-Rodriguez, A.; Lloyd, R.C.; Roiban, G.D. Biocatalysis: A pharma perspective. Adv. Synth. Catal. 2019, 361, 2421–2432. [Google Scholar] [CrossRef]
  44. Musa, M.M.; Hollmann, F.; Mutti, F.G. Synthesis of enantiomerically pure alcohols and amines via biocatalytic deracemisation methods. Cat. Sci. Technol. 2019, 9, 10–1039. [Google Scholar] [CrossRef]
  45. Ichikawa, H.; Bashiri, G.; Kelly, W.L. Biosynthesis of the thiopeptins and identification of an F420H2-dependent dehydropiperidine reductase. J. Am. Chem. Soc. 2018, 140, 10749–10756. [Google Scholar] [CrossRef]
  46. Miller, A.F.; Park, J.T.; Ferguson, K.L.; Pitsawong, W.; Bommarius, A.S. Informing efforts to develop nitroreductase for amine production. Molecules 2018, 23, 22. [Google Scholar] [CrossRef]
  47. Heiss, G.; Hofmann, K.W.; Trachtmann, N.; Walters, D.M.; Rouvière, P.; Knackmuss, H.J. Npd gene functions of Rhodococcus (opacus) erythropolis Hl PM-1 in the initial steps of 2,4,6-trinitrophenol degradation. Microbiology 2002, 148, 799–806. [Google Scholar] [CrossRef]
  48. Xu, J.; Green, A.P.; Turner, N.J. Chemo-enzymatic synthesis of pyrazines and pyrroles. Angew. Chem.-Int. Ed. 2018, 57, 16760–16763. [Google Scholar] [CrossRef]
  49. Busacca, C.A.; Fandrick, D.R.; Song, J.J.; Senanayake, C.H. The growing impact of catalysis in the pharmaceutical industry. Adv. Synth. Catal. 2011, 353, 1825–1864. [Google Scholar] [CrossRef]
  50. Heiss, G.; Trachtmann, N.; Abe, Y.; Takeo, M.; Knackmuss, H.J. Homologous npdgi genes in 2,4-dinitrophenol- and 4-nitrophenol-degrading Rhodococcus spp. Appl. Environ. Microbiol. 2003, 69, 2748–2754. [Google Scholar] [CrossRef]
  51. Purwantini, E.; Daniels, L.; Mukhopadhyay, B. F420H2 is required for phthiocerol dimycocerosate synthesis in Mycobacteria. J. Bacteriol. 2016, 198, 2020–2028. [Google Scholar] [CrossRef]
  52. Wichmann, R.; Vasic-Racki, D. Cofactor regeneration at the lab scale. In Technology Transfer in Biotechnology; Springer: Berlin, Germany, 2005; pp. 225–260. [Google Scholar]
  53. Tishkov, V.I.; Popov, V.O. Catalytic mechanism and application of formate dehydrogenase. Biochemistry 2004, 69, 1252. [Google Scholar] [CrossRef] [PubMed]
  54. Eguchi, T.; Kuge, Y.; Inoue, K.; Yoshikawa, N.; Mochida, K.; Uwajima, T. NADPH regeneration by glucose dehydrogenase from Gluconobacter scleroides for L-leucovorin synthesis. Biosci. Biotechnol. Biochem. 1992, 56, 701–703. [Google Scholar] [CrossRef] [PubMed]
  55. Demir, A.S.; Talpur, F.N.; Sopaci, B.; Kohring, G.-W.; Celik, A. Selective oxidation and reduction reactions with cofactor regeneration mediated by galactitol-, lactate-, and formate dehydrogenases immobilized on magnetic nanoparticles. J. Biotechnol. 2011, 152, 176–183. [Google Scholar] [CrossRef] [PubMed]
  56. Wong, C.-H.; Whitesides, G.M. Enzyme-catalyzed organic synthesis: NAD(P)H cofactor regeneration by using glucose-6-phosphate and the glucose-5-phosphate dehydrogenase from Leuconostoc mesenteroides. J. Am. Chem. Soc. 1981, 103, 4890–4899. [Google Scholar] [CrossRef]
  57. Lee, W.-H.; Park, J.-B.; Park, K.; Kim, M.-D.; Seo, J.-H. Enhanced production of ɛ-caprolactone by overexpression of NADPH-regenerating glucose 6-phosphate dehydrogenase in recombinant Escherichia coli harboring cyclohexanone monooxygenase gene. Appl. Microbiol. Biotechnol. 2007, 76, 329–338. [Google Scholar] [CrossRef] [PubMed]
  58. Berrios-Rivera, S.J.; Bennett, G.N.; San, K.Y. Metabolic engineering of Escherichia coli: Increase of NADH availability by overexpressing an NAD+-dependent formate dehydrogenase. Metab. Eng. 2002, 4, 217–229. [Google Scholar] [CrossRef]
  59. Bashiri, G.; Antoney, J.; Jirgis, E.N.M.; Shah, M.V.; Ney, B.; Copp, J.; Stuteley, S.M.; Sreebhavan, S.; Palmer, B.; Middleditch, M.; et al. A revised biosynthetic pathway for the cofactor F420 in prokaryotes. Nat. Commun. 2019, 10, 1558. [Google Scholar] [CrossRef]
  60. Purwantini, E.; Daniels, L. Purification of a novel coenzyme F420-dependent glucose-6-phosphate dehydrogenase from Mycobacterium smegmatis. J. Bacteriol. 1996, 178, 2861–2866. [Google Scholar] [CrossRef]
  61. Costa, K.C.; Wong, P.M.; Wang, T.; Lie, T.J.; Dodsworth, J.A.; Swanson, I.; Burn, J.A.; Hackett, M.; Leigh, J.A. Protein complexing in a methanogen suggests electron bifurcation and electron delivery from formate to heterodisulfide reductase. Proc. Natl. Acad. Sci. USA 2010, 107, 11050–11055. [Google Scholar] [CrossRef] [Green Version]
  62. Shuber, A.P.; Orr, E.C.; Recny, M.A.; Schendel, P.F.; May, H.D.; Schauer, N.L.; Ferry, J.G. Cloning, expression, and nucleotide sequence of the formate dehydrogenase genes from Methanobacterium formicicum. J. Biol. Chem. 1986, 261, 12942–12947. [Google Scholar]
  63. Novotná, J.; Neužil, J.; Hoš?álek, Z. Spectrophotometric identification of 8-hydroxy-5-deazaflavin: Nadph oxidoreductase activity in Streptomycetes producing tetracyclines. FEMS Microbiol. Lett. 1989, 59, 241–245. [Google Scholar] [CrossRef]
  64. Kumar, H.; Nguyen, Q.T.; Binda, C.; Mattevi, A.; Fraaije, M.W. Isolation and characterization of a thermostable F420:NADPH oxidoreductase from Thermobifida fusca. J. Biol. Chem. 2017, 292, 10123–10130. [Google Scholar] [CrossRef] [PubMed]
  65. Kunow, J.; Schwörer, B.; Stetter, K.O.; Thauer, R.K. A F420-dependent NADP reductase in the extremely thermophilic sulfate-reducing Archaeoglobus fulgidus. Arch. Microbiol. 1993, 160, 199–205. [Google Scholar]
  66. Eker, A.P.M.; Hessels, J.K.C.; Meerwaldt, R. Characterization of an 8-hydroxy-5-deazaflavin: NADPH oxidoreductase from Streptomyces griseus. Biochim. Biophys. Acta 1989, 990, 80–86. [Google Scholar] [CrossRef]
  67. Seelbach, K.; Riebel, B.; Hummel, W.; Kula, M.R.; Tishkov, V.I.; Egorov, A.M.; Wandrey, C.; Kragl, U. A novel, efficient regenerating method of NADPH using a new formate dehydrogenase. Tetrahedron Lett. 1996, 37, 1377–1380. [Google Scholar] [CrossRef]
  68. Alex, L.A.; Reeve, J.N.; Ormejohnson, W.H.; Walsh, C.T. Cloning, sequence determination, and expression of the genes encoding the subunits of the nickel-containing 8-hydroxy-5-deazaflavin reducing hydrogenase from Methanobacterium thermoautotrophicum delta-H. Biochemistry 1990, 29, 7237–7244. [Google Scholar] [CrossRef]
  69. Tersteegen, A.; Hedderich, R. Methanobacterium thermoautotrophicum encodes two multisubunit membrane-bound NiFe hydrogenases - transcription of the operons and sequence analysis of the deduced proteins. Eur. J. Biochem. 1999, 264, 930–943. [Google Scholar] [CrossRef]
  70. Hocking, W.P.; Stokke, R.; Roalkvam, I.; Steen, I.H. Identification of key components in the energy metabolism of the hyperthermophilic sulfate-reducing archaeon archaeoglobus fulgidus by transcriptome analyses. Front. Microbiol. 2014, 5, 20. [Google Scholar] [CrossRef]
  71. Vitt, S.; Ma, K.; Warkentin, E.; Moll, J.; Pierik, A.J.; Shima, S.; Ermler, U. The F-420-reducing NiFe -hydrogenase complex from methanothermobacter marburgensis, the first x-ray structure of a group 3 family member. J. Mol. Biol. 2014, 426, 2813–2826. [Google Scholar] [CrossRef]
  72. Bashiri, G.; Rehan, A.M.; Greenwood, D.R.; Dickson, J.M.; Baker, E.N. Metabolic engineering of cofactor F420 production in Mycobacterium smegmatis. PLoS ONE 2010, 5, e15803. [Google Scholar] [CrossRef]
  73. Isabelle, D.; Simpson, D.R.; Daniels, L. Large-scale production of coenzyme F420-5,6 by using Mycobacterium smegmatis. Appl. Environ. Microbiol. 2002, 68, 5750–5755. [Google Scholar] [CrossRef] [PubMed]
  74. Eker, A.P.M.; Pol, A.; van der Meyden, P.; Vogels, G.D. Purification and properties of 8-hydroxy-5-deazaflavin derivatives from Streptomyces griseus. FEMS Microbiol. Lett. 1980, 8, 161–165. [Google Scholar] [CrossRef]
  75. Grochowski, L.L.; Xu, H.M.; White, R.H. Identification and characterization of the 2-phospho-L-lactate guanylyltransferase involved in coenzyme F-420 biosynthesis. Biochemistry 2008, 47, 3033–3037. [Google Scholar] [CrossRef] [PubMed]
  76. Braga, D.; Lasta, D.; Hasan, M.; Guo, H.; Leichnitz, D.; Uzum, Z.; Richter, I.; Schalk, F.; Beemelmanns, C.; Hertweck, C.; et al. Metabolic pathway rerouting in Paraburkholderia rhizoxinica evolved long-overlooked derivatives of coenzyme F420. ACS Chem. Biol. 2019, 2088–2094. [Google Scholar] [CrossRef]
  77. Hossain, M.S.; Le, C.Q.; Joseph, E.; Nguyen, T.Q.; Johnson-Winters, K.; Foss, F.W. Convenient synthesis of deazaflavin cofactor Fo and its activity in F420-dependent NADP reductase. Organ. Biomol. Chem. 2015, 13, 5082–5085. [Google Scholar] [CrossRef]
  78. Alberty, R.A. Calculation of standard transformed formation properties of biochemical reactants and standard apparent reduction potentials of half reactions. Arch. Biochem. Biophys. 1998, 358, 25–39. [Google Scholar] [CrossRef]
  79. Alberty, R.A. Calculation of standard transformed gibbs energies and standard transformed enthalpies of biochemical reactants. Arch. Biochem. Biophys. 1998, 353, 116–130. [Google Scholar] [CrossRef]
  80. Benedict, M.N.; Gonnerman, M.C.; Metcalf, W.W.; Price, N.D. Genome-scale metabolic reconstruction and hypothesis testing in the methanogenic archaeon Methanosarcina acetivorans C2A. J. Bacteriol. 2012, 194, 855–865. [Google Scholar] [CrossRef]
  81. Jankowski, M.D.; Henry, C.S.; Broadbelt, L.J.; Hatzimanikatis, V. Group contribution method for thermodynamic analysis of complex metabolic networks. Biophys. J. 2008, 95, 1487–1499. [Google Scholar] [CrossRef]
  82. Henry, C.S.; DeJongh, M.; Best, A.A.; Frybarger, P.M.; Linsay, B.; Stevens, R.L. High-throughput generation, optimization and analysis of genome-scale metabolic models. Nat. Biotechnol. 2010, 28, 977–982. [Google Scholar] [CrossRef]
  83. Nazem-Bokaee, H.; Gopalakrishnan, S.; Ferry, J.G.; Wood, T.K.; Maranas, C.D. Assessing methanotrophy and carbon fixation for biofuel production by Methanosarcina acetivorans. Microb. Cell Fact. 2016, 15, 10. [Google Scholar] [CrossRef] [PubMed]
  84. Caspi, R.; Billington, R.; Fulcher, C.A.; Keseler, I.M.; Kothari, A.; Krummenacker, M.; Latendresse, M.; Midford, P.E.; Ong, Q.; Ong, W.K.; et al. The metacyc database of metabolic pathways and enzymes. Nucleic Acids Res. 2018, 46, D633–D639. [Google Scholar] [CrossRef] [PubMed]
Catalysts 09 00868 g001 550
Figure 1. The structures of NAD(P) (top), cofactor F420 and its synthetic analog FOP (center) and common flavins (riboflavin, FMN and FAD; bottom). The oxidized and reduced forms are shown, as is the flavin semiquinone. Dashed lines indicate the differences in the structures of FOP and cofactor F420, and riboflavin, FMN and FAD.
Figure 1. The structures of NAD(P) (top), cofactor F420 and its synthetic analog FOP (center) and common flavins (riboflavin, FMN and FAD; bottom). The oxidized and reduced forms are shown, as is the flavin semiquinone. Dashed lines indicate the differences in the structures of FOP and cofactor F420, and riboflavin, FMN and FAD.
Catalysts 09 00868 g001
Catalysts 09 00868 g002 550
Figure 2. Representative structures of F420-dependent FDOR-A (PDB: 3R5Z, panels A and C) and FDOR-B (PDB: 5JAB, panels B and D). Both are predominantly composed of a single β-sheet forming a split barrel. The N-terminal helices are spatially displaced between the two families, falling on opposite faces of the β-sheet.
Figure 2. Representative structures of F420-dependent FDOR-A (PDB: 3R5Z, panels A and C) and FDOR-B (PDB: 5JAB, panels B and D). Both are predominantly composed of a single β-sheet forming a split barrel. The N-terminal helices are spatially displaced between the two families, falling on opposite faces of the β-sheet.
Catalysts 09 00868 g002
Catalysts 09 00868 g003 550
Figure 3. Structure of representative luciferase-like hydride transferase (LLHT) (PDB: 1RHC). (A) A 3D representation of the biologically relevant dimer (panel A). Monomer of an LLHT with insertion sequences IS1–4 highlighted, along with the helical bundle composed of α7–9 (panel B). Topology diagram showing (α/β)8 fold with insertion sequences highlighted: IS1, red; IS2, orange; IS3, light green, IS4, pink. The helical bundle of α7–9 is highlighted in purple (panel C).
Figure 3. Structure of representative luciferase-like hydride transferase (LLHT) (PDB: 1RHC). (A) A 3D representation of the biologically relevant dimer (panel A). Monomer of an LLHT with insertion sequences IS1–4 highlighted, along with the helical bundle composed of α7–9 (panel B). Topology diagram showing (α/β)8 fold with insertion sequences highlighted: IS1, red; IS2, orange; IS3, light green, IS4, pink. The helical bundle of α7–9 is highlighted in purple (panel C).
Catalysts 09 00868 g003
Catalysts 09 00868 g004 550
Figure 4. Representative cofactor F420-dependent oxidoreductions with the potential for adaptation to biocatalytic applications. Included are: nitroreduction, enoate reduction, ketoreduction and imine reduction (from top to bottom). For clarity, only the dehydropiperidine ring of the thiopeptide is shown and partial structures for biliverdin-Ixα and phthiodiolone dimycocerosates are shown.
Figure 4. Representative cofactor F420-dependent oxidoreductions with the potential for adaptation to biocatalytic applications. Included are: nitroreduction, enoate reduction, ketoreduction and imine reduction (from top to bottom). For clarity, only the dehydropiperidine ring of the thiopeptide is shown and partial structures for biliverdin-Ixα and phthiodiolone dimycocerosates are shown.
Catalysts 09 00868 g004
Catalysts 09 00868 g005 550
Figure 5. Enoate reduction by a flavin-dependent enzyme (Old Yellow Enzyme) and the proposed mechanism for cofactor F420-dependent reduction. Notably the mechanism of reduction yields trans-hydrogenation products for Old Yellow Enzyme and cis-hydrogenation products for the F420-dependent enzymes.
Figure 5. Enoate reduction by a flavin-dependent enzyme (Old Yellow Enzyme) and the proposed mechanism for cofactor F420-dependent reduction. Notably the mechanism of reduction yields trans-hydrogenation products for Old Yellow Enzyme and cis-hydrogenation products for the F420-dependent enzymes.
Catalysts 09 00868 g005
Catalysts 09 00868 g006 550
Figure 6. Proposed scheme for one-pot, enzyme cascades for deracemization/steroinversion of sec-alcohols (top) and chiral amine synthesis (bottom) using cofactor F420-dependent alcohol oxidation.
Figure 6. Proposed scheme for one-pot, enzyme cascades for deracemization/steroinversion of sec-alcohols (top) and chiral amine synthesis (bottom) using cofactor F420-dependent alcohol oxidation.
Catalysts 09 00868 g006
Catalysts 09 00868 g007 550
Figure 7. Cofactor F420-dependent oxidation reactions that could be exploited to produce reduced cofactor F420.
Figure 7. Cofactor F420-dependent oxidation reactions that could be exploited to produce reduced cofactor F420.
Catalysts 09 00868 g007
Table
Table 1. Characterized F420-dependent enzymes with activities that could be adapted for biocatalytic applications.
Table 1. Characterized F420-dependent enzymes with activities that could be adapted for biocatalytic applications.
ReactionFamilyReference
Enoate reduction
AflatoxinsFDOR[14,18,34]
CoumarinsFDOR[14,34,35]
QuinonesFDOR[36]
Biliverdin reductionFDOR[12,16]
NitroimidazolesFDOR[36]
CyclohexenonesFDOR[6,34,38]
Citral/Neral/GeranialFDOR[6]
CarvoneFDOR[6]
KetoisophoroneFDOR[6]
Alcohol oxidation/ketoreduction
Glucose-6-phosphateLLHT[26,50]
Phthiodiolone dimycocerosateLLHT[51]
IsopropanolLLHT[40]
Imine reductions
Dehydropiperidine (in thiopeptins)FDOR[45]
4-ethylidene-3,4-dihydropyrrole-2-carboxylic acidFlavin-dependent monooxygenase[11]
Nitroreductions
PicrateLLHT[47,50]
2,4-DNPLLHT[48,50]
Ring opening/closing
C-N bond cleavage/formation in methylenetetrahydromethanopterinLLHT[23,28,29,30]
Table
Table 2. Published production systems for cofactor F420.
Table 2. Published production systems for cofactor F420.
SourceF420 Yield (μmol/g Cell Weight)Growth ConditionsRef
Methanobacterium thermoautotrophicum0.42 a,cGrown at 60 °C using complex media in fermenter, under pressurized hydrogen[9]
Methanobacterium formicium0.27 a,cGrown at 37 °C using complex media in fermenters[9]
Methanospirillum hungatii0.41 a,cGrown at 37 °C using complex media in fermenters[9]
Methanobacterium strain M.o.H0.53 a,cGrown at 40 °C using complex media in fermenters[9]
Methanobacterium thermoautotrophicum1.7 eGrown using complex media in fermenters, under pressurized hydrogen gas[73]
Streptomyces flocculus0.62 eGrown using complex media in fermenters[73]
Streptomyces coelicolor0.04 eGrown using complex media in fermenters[73]
Streptomyces griseus0.008 a,cGrowth conditions not mentioned in the publication[74]
Rhodococcus rhodochrous0.11 eGrown using complex media in fermenters[73]
Mycobacterium smegmatis0.30 eGrown using complex media in fermenters[73]
Mycobacterium smegmatis3.0 dOverexpression of F420 pathway genes, cultivation in complex media at 37 °C in shake flasks[72]
Escherichia coli0.38 bOverexpressing F420 pathway genes, grown in minimal media at 30 °C in shake flasks.[59]
a Mol weight of F420 with 1 glutamate tail is 773.6 Da, which was used to convert values published as μg of F420, noting that micro-organisms produce mixture of F420 with different number of glutamates (1–9) attached. b Concentration estimated through absorbance at 420 nm and using extinction coefficient of 41.4 mM−1 cm−1 [73]. c F420 concentration per g of wet cell weight. d Concentration of F420 not mentioned in the publication, but F420 yield was stated to be 10 times higher than wild-type M. smegmatis. e Concentration estimated through absorbance at 400 nm and using extinction coefficient of 25.7 mM−1 cm−1 [74].

Share and Cite

MDPI and ACS Style

Shah, M.V.; Antoney, J.; Kang, S.W.; Warden, A.C.; Hartley, C.J.; Nazem-Bokaee, H.; Jackson, C.J.; Scott, C. Cofactor F420-Dependent Enzymes: An Under-Explored Resource for Asymmetric Redox Biocatalysis. Catalysts 2019, 9, 868. https://doi.org/10.3390/catal9100868

AMA Style

Shah MV, Antoney J, Kang SW, Warden AC, Hartley CJ, Nazem-Bokaee H, Jackson CJ, Scott C. Cofactor F420-Dependent Enzymes: An Under-Explored Resource for Asymmetric Redox Biocatalysis. Catalysts. 2019; 9(10):868. https://doi.org/10.3390/catal9100868

Chicago/Turabian Style

Shah, Mihir V., James Antoney, Suk Woo Kang, Andrew C. Warden, Carol J. Hartley, Hadi Nazem-Bokaee, Colin J. Jackson, and Colin Scott. 2019. "Cofactor F420-Dependent Enzymes: An Under-Explored Resource for Asymmetric Redox Biocatalysis" Catalysts 9, no. 10: 868. https://doi.org/10.3390/catal9100868

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop