Same Substrate, Many Reactions: Oxygen Activation in

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Cite This: Chem. Rev. 2018, 118, 1742−1769

Same Substrate, Many Reactions: Oxygen Activation in Flavoenzymes ́ ez Castellanos,‡ Giovanni Gadda,*,§ Marco W. Fraaije,*,† Elvira Romero,† J. Rubeń Gom ,‡ and Andrea Mattevi* †

Molecular Enzymology Group, University of Groningen, Nijenborgh 4, 9747AG Groningen, The Netherlands Department of Biology and Biotechnology “Lazzaro Spallanzani”, University of Pavia, Via Ferrata 9, 27100 Pavia, Italy § Departments of Chemistry and Biology, Center for Diagnostics and Therapeutics, and Center for Biotechnology and Drug Design, Georgia State University, Atlanta, Georgia 30302-3965, United States ‡

ABSTRACT: Over time, organisms have evolved strategies to cope with the abundance of dioxygen on Earth. Oxygen-utilizing enzymes tightly control the reactions involving O2 mostly by modulating the reactivity of their cofactors. Flavins are extremely versatile cofactors that are capable of undergoing redox reactions by accepting either one electron or two electrons, alternating between the oxidized and the reduced states. The physical and chemical principles of flavin-based chemistry have been investigated widely. In the following pages we summarize the state of the art on a key area of research in flavin enzymology: the molecular basis for the activation of O2 by flavin-dependent oxidases and monooxygenases. In general terms, oxidases use O2 as an electron acceptor to produce H2O2, while monooxygenases activate O2 by forming a flavin intermediate and insert an oxygen atom into the substrate. First, we analyze how O2 reaches the flavin cofactor embedded in the protein matrix through dedicated access pathways. Then we approach O2 activation from the perspective of the monooxygenases, their preferred intermediate, the C(4a)−(hydro)peroxyflavin, and the cases in which other intermediates have been described. Finally, we focus on understanding how the architectures developed in the active sites of oxidases promote O2 activation and which other factors operate in its reactivity.

CONTENTS 1. Introduction 2. Reaction of Flavins with Molecular Oxygen 3. How Does Molecular Oxygen Reach the Flavin Cofactor in Flavoenzymes? 4. Flavin-Dependent Monooxygenases 4.1. O2 Activation in Flavoprotein Monooxygenases 4.1.1. Aromatic Hydroxylation 4.1.2. Baeyer−Villiger Oxidation 4.1.3. Heteroatom Oxygenation 4.1.4. Light Emission 4.1.5. Halogenation and Decarboxylative− Halogenation 4.1.6. Dehalogenation 4.1.7. Denitrification 4.1.8. Hydroxylation−Dehydrogenation 5. Flavin-Dependent Oxidases 5.1. Mechanisms of Oxygen Activation 5.1.1. Preorganization of the Active Site 5.1.2. Choline Oxidase: Taking Advantage of a Charged Substrate 5.1.3. Pyranose 2-Oxidase and Bacterial NADH Oxidase: Deviating from the Canon 6. Self-Sacrificing Flavin

© 2018 American Chemical Society

7. Conclusions and Outlook Author Information Corresponding Authors ORCID Notes Biographies Acknowledgments Abbreviations References

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1. INTRODUCTION The atmosphere of the Earth is relatively rich in molecular oxygen (O2). This is the result of the photosynthetic abilities of plants and cyanobacteria which led to a steady rise of oxygen gas resulting in the so-called great oxygenation event about 2.5 billion years ago. Since then the abundance of oxygen gas has forced organisms to develop strategies to cope with this potent oxidant. While there are many organisms that thrive in anaerobic environments, the majority has found mechanisms to handle molecular oxygen. In fact, the high reduction potential of O2 makes it an excellent oxidizing agent in thermodynamic terms.1−3 Yet, the triplet ground state of

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Received: October 27, 2017 Published: January 11, 2018 1742

DOI: 10.1021/acs.chemrev.7b00650 Chem. Rev. 2018, 118, 1742−1769

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dioxygen, with two unpaired electrons, prevents spontaneous combustion of organic molecules. A redox catalyst can overcome this kinetic lack of reactivity. However, the uncontrolled reduction of O2 often generates toxic products, such as radicals that react nonspecifically with many cellular compounds.3 Therefore, a tight control of the reactions involving O2 must be enforced by oxygen-utilizing enzymes. Such biocatalysts can also accelerate reactions with O2 at the active site and control the fate of the formed activated oxygen intermediates so that only the specific target molecule is oxidized and formation of unwanted reactive oxygen species is prevented.4 A large variety of enzyme types that utilize dioxygen as substrate have been identified in recent decades. Many of them rely on a metal or a metal-containing cofactor for catalysis. For example, laccases are copper-containing enzymes that can oxidize aromatic compounds with concomitant reduction of dioxygen into water, whereas mononuclear iron dioxygenases and heme-containing P450 monooxygenases are effective in many oxygenation reactions.5,6 There is also a large group of oxygen-utilizing enzymes that are devoided of metals and rely on a flavin cofactor. Flavins are extremely versatile molecules that are capable of undergoing oxidation−reduction reactions by exchanging either one electron or two electrons. Flavins can thereby exist in oxidized (quinone; Flox), one-electron-reduced (semiquinone; Flsq), and two-electron-reduced (hydroquinone; Flred) states. Flavoenzymes typically contain either a flavin mononucleotide (FMN) or a flavin adenine dinucleotide (FAD) cofactor, both of which are synthesized in vivo from riboflavin, i.e., vitamin B2 (Figure 1).7−9 In the majority of flavoenzymes, the cofactor is tightly

Scheme 1. Catalytic Cycle of Flavoenzymes Typically Comprises Two-Half Reactionsa

a Oxidized flavin (Flox) is reduced through oxidation of the substrate in the reductive half-reaction (RHR). In the oxidative half-reaction (OHR), the reduced flavin (Flred) reacts with an electron acceptor. Depending on the enzyme, substrate, and pH or other environmental factors, the reaction can proceed via a ternary complex (bottom), a ping-pong (up) mechanism, or both.2

through a ternary complex, where the electron acceptor reacts with the enzyme−product complex. Alternatively, when the enzyme functions in a ping-pong mechanism, the electron acceptor reacts with the enzyme after product release.2 Flavoenzymes may also operate via both routes simultaneously as found for putrescine oxidase.24 There are three well-defined groups of flavoproteins based on the rate of reaction with O2 and the nature of the product. The first group is comprised of flavin-dependent dehydrogenases/reductases. These redox enzymes do not react at all or react very slowly with O2 to mainly form hydrogen peroxide (H2O2) or some superoxide anion (O2−•) as the reaction products.2,25,26 Flavoprotein dehydrogenases often may show an oxidase activity in vitro but act as genuine dehydrogenase in vivo where the preferred electron acceptor is in abundance and/or dioxygen is rather limited. For example, based on initial in vitro biochemical studies the plant flavoenzyme responsible for the oxidative degradation of cytokinins was originally described as a cytokinin oxidase. Yet, more detailed studies have revealed that the enzyme operates optimally by using quinones as electron acceptor, while the reduced enzyme reacts poorly with molecular oxygen.27 Medium-chain acyl CoA dehydrogenase is another classic representative of flavoprotein dehydrogenases that show some sluggish activity with dioxygen.28−33 An example of a flavoprotein dehydrogenase that is fully resilient toward dioxygen as electron acceptor is 34 D-arginine dehydrogenase from Pseudomonas aeruginosa. The second and third groups of dioxygen-reactive flavoproteins, the oxidases and monooxygenases, instead react rapidly with molecular oxygen to yield the oxidized flavoprotein.2,35−37 Flavin-dependent oxidases efficiently use O2 as an electron acceptor to produce, in most cases, H2O2, whereas flavoprotein monooxygenases generally activate dioxygen by forming a C(4a)−(hydro)peroxide with the flavin, which is used to insert an oxygen atom into the substrate (Scheme 2). Oxygen reactivity in flavoprotein oxidases and monooxygenases is considerably enhanced compared to free flavins in bulk solvent with second-order rate constants kcat/KM(O2) up to 106−107 M−1 s−1. This sets them apart from all other flavoenzymes that

Figure 1. Structures of flavin mononucleotide (FMN) and flavin adenine dinucleotide (FAD) cofactors, derived from riboflavin. (Inset) UV−vis absorption spectra of the oxidized, two-electron reduced, and C(4a)−peroxyflavin (orange, black broken, and black lines, respectively).

but noncovalently bound.10,11 However, in a subset of them (ca. 10%), the flavin is mono- or bicovalently attached to the polypeptide chain.12,13 In these cases, it has been observed that the covalent flavin allows catalysis of more thermodynamically challenging reactions by modulating the cofactor redox potential.14−23 With a few limited exceptions, the reaction of flavoenzymes typically uses two half-reactions by which the flavin alternates between the oxidized and the reduced states (Scheme 1).1,2 In the reductive half-reaction, the substrate reduces the flavin; then in the oxidative half-reaction the cofactor is oxidized in return by an electron acceptor. Depending on the enzyme, substrate, and pH or other environmental factors, the overall catalytic cycle can occur 1743


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Scheme 2. Conventional Pathways for the Reactions with O2 in Monooxygenases and Oxidasesa

a

The left structure highlights the positions of the reactive N(5) and C(4a) atoms, which are directly involved in the reaction with molecular oxygen.53

activation of molecular oxygen. While considerable advances have been made, how flavoproteins are able to delicately tune their reactivity toward this small, hydrophobic, and diradical molecule is still not fully understood. Here, we present an overview of the mechanistic and structural features of flavoenzymes that have evolved to utilize molecular oxygen: flavin-dependent oxidases and monooxygenases.

are poorly reactive or inert toward dioxygen as electron acceptor.2,3,25,38−40 In biological terms, the adjustable oxygen reactivity makes flavoenzymes useful biocatalysts for all kind of organisms. In addition to enabling efficient oxidations of target compounds, it also allows Nature to generate and exploit reduced oxygen species. In fact, it was recently found that the reactive oxygen species formed by action of flavoproteins serve important roles as signaling molecules.41,42 The tunability of the flavin is likely one of the major factors contributing to flavoenzymes as a whole being a widespread protein class. Currently, more than 100 000 protein sequences deposited in the NCBI database are classified as flavin-dependent enzymes and over 1800 FADand over 900 FMN-containing protein structures have been deposited in the Protein Data Base (as of August 2017).43 In this context, it is worth mentioning that also an oxygeninsensitive flavin cofactor is present in nature: the deazaflavin (FO or F420).44 Through the replacement of the N(5) atom by a carbon, these molecules exhibit hardly any oxygen reactivity and are utilized by microorganisms as cofactor for dehydrogenases and reductases. Intriguingly, it has been concluded that regular flavins and deazaflavins were already present in the last universal common ancestor.45 As deazaflavoenzymes are rather rare and restricted to archaea and bacteria, it indicates that regular flavins have become much more popular in nature. This may be partly explained by their catalytic versatility, which includes its reactivity with molecular oxygen. Flavins and the physical and chemical principles by which they perform catalysis have been the subject of intensive research for >60 years.46−48 A large part of the current understanding of the chemistries facilitated by flavoenzymes is based on seminal works performed in the 1970s−1980s by the teams of Thomas C. Bruice and Vincent Massey, which led to the current view of the reaction of reduced flavin with dioxygen in oxidases and monooxygenases.25,49−52 A wealth of mechanistic, kinetic, structural, and computational data provide consistent clues about the molecular, functional, and structural properties of flavoprotein oxidases and oxygenases. Yet, not all aspects of flavoenzyme-catalyzed reactions have been solved. One major flavoenzyme research area indeed focuses on the elucidation of the molecular basis for the

2. REACTION OF FLAVINS WITH MOLECULAR OXYGEN Reduced flavins display an intrinsic reactivity with dioxygen (Scheme 3). The oxidation of free flavin proceeds with an Scheme 3. Proposed Reaction Mechanism for the Oxidation of the Flavin Hydroquinone with O2 in Solutionq

After initial transfer of an electron from the reduced flavin to O2 generating O2−• and a neutral flavin semiquinone (1), spin inversion of the resulting radical pair (2) allows for further reaction to yield the oxidized flavin and hydrogen peroxide (3). The latter step may proceed through the formation of a transient C(4a)−hydroperoxyflavin (shown in Scheme 2) or through an outer-sphere second electron transfer (shown).35 q

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estimated rate of 250 M−1 s−1, leading to the accumulation of reactive species, in particular flavin radicals and O2−•.25,54 Thorough kinetic and chemical analyses suggest an initial electron transfer from singlet reduced flavin to triplet O2 to yield a caged radical pair. After spin inversion, this intermediate can collapse into a peroxyflavin which is unstable in water and dissociates heterolytically to H2O2 and the oxidized flavin.55 Evidence supporting this mechanistic route was provided by experiments in which the neutral flavin radical and O2−• were produced by pulse radiolysis of aerated solutions of riboflavin. The flavin radical was converted in a fast second-order reaction to the C(4a)−hydroperoxyflavin, which then decayed to the oxidized flavin at an O2−•independent rate.56 More recently, spectroscopic evidence for the formation of a reaction intermediate with absorption properties resembling those of a flavin semiquinone have been reported in the oxidative half-reaction of human glycolate oxidase.57 Of note, the reaction can also be triggered by lightmediated reduction of flavin in the presence of molecular oxygen. Such uncontrolled light-induced flavin-catalyzed formation of reactive oxygen species can have detrimental effects when occurring in living cells and in some industrial processes.58 In fact, there is evidence suggesting that the socalled dodecins, small flavin-binding proteins, are meant to scavenge phototoxic flavins to prevent flavin-triggered cellular damage.59 Given the nature of the reaction, it is essential to consider the redox potentials of the reactants. One obvious factor is the thermodynamic driving force, i.e., the difference in redox potentials between the flavin and the oxygen couples.54,60 The potential of the oxygen−hydrogen peroxide couple is +281 mV (pH 7.0),61 while the redox potentials for the O2/O2−• and O2−•/H2O2 couples are approximately −330 and +890 mV, respectively.62 These values must be compared to those of the flavin couples. Although the midpoint potential for the oxidized−fully reduced Flox/Flred couple in aqueous solution is −207 mV (pH 7.0),63 protein-bound flavins exhibit widely different potentials that range from −400 to +150 mV, reflecting the strong redox modulation exerted by the protein environment.64,65 Despite these variations, the two-electron potential of the bound flavins is lower than that of the oxygen− hydrogen peroxide (+281 mV). This feature strongly favors the drive toward the flavoenzyme-catalyzed conversion of oxygen to hydrogen peroxide, which is essentially irreversible. However, the one-electron redox potential for the oxygen− superoxide anion couple (−330 mV)61 is often lower than that of the fully reduced−one-electron-reduced flavin couple, which ranges from −400 to −50 mV among flavoproteins and is −101 mV in the free flavin.63 This indicates a weak driving force for the initial electron-transfer step. Consideration of the redox potential must be paired with the observed kinetic behavior for the oxidation reaction. The rate at which reduced flavins or flavoproteins react with molecular oxygen can be accurately determined using the stopped-flow technique by exploiting the highly different spectral properties of the different flavin intermediates. From the accumulated wealth of kinetic data on flavins and flavoproteins, a key observation is that dioxygen reacts with reduced flavin following second-order kinetic behavior in most flavoenzymes, without showing saturation of the observed rate constants with increasing concentration of dioxygen.25,52,66 From this one can conclude that the reaction involves a collision event between molecular oxygen and the reduced flavin cofactor without an

observable binding event of dioxygen to form a discrete enzyme−oxygen complex. In fact, the reaction seems similar to that between free flavin and molecular oxygen.55 Yet, the protein matrix in flavoenzymes is able to tune the rate at which dioxygen can react with a reduced flavoprotein with reoxidation rate constants that range from 1 × 106 M−1 s−1 to almost zero.2

3. HOW DOES MOLECULAR OXYGEN REACH THE FLAVIN COFACTOR IN FLAVOENZYMES? Even though molecular oxygen is a rather small molecule, it displays a different behavior when compared with the natural solvent, water. Several recent studies have addressed the general question of how the protein matrix of flavoproteins can promote or prevent the use of molecular oxygen as substrate. Typically, the reactive isoalloxazine moiety of the FAD and FMN cofactors is deeply bound in the protein core of a flavoenzyme. This provides such enzymes a means for filtering out which molecules can reach the active site. While relatively wide and water-filled tunnels allow substrate molecules to reach the catalytic center, these entries typically are not favorable for dioxygen, which is a hydrophobic, diradical molecule and will prefer more hydrophobic routes. It has been suggested in the past that, being such a small molecule, dioxygen may reach the active site through a process of passive diffusion, taking advantage of the dynamics of protein structures. However, in recent years it has become clear that certain oxidases and monooxygenases feature favorite accession tunnels or pathways for dioxygen as an electron-accepting substrate. Such mode of protein-guided dioxygen diffusion is in line with the observed dioxygen pathways elucidated for other oxygen-utilizing proteins (Figure 2).67

Figure 2. Binding of oxygen to flavoenzymes. General consensus is that dioxygen preferentially diffuses through tunnels that transiently form through connecting cavities. These tunnels converge to the reactive flavin N(5)−C(4a) locus where gating residues may operate to finely control the reaction of the flavin with dioxygen. The reader is referred to the movies published as additional online material in the articles in refs 71 and 73−77 for a view of the dynamic nature of these transiently forming oxygen tunnels as gathered from molecular dynamics simulations.

One of the first hints for dedicated molecular oxygen pathways in molecular oxygen-dependent flavoenzymes came from the analysis of the crystal structure of cholesterol oxidase from Brevibacterium sterolicum.68 The structure of this bacterial FAD-containing oxidase revealed a distinct and narrow tunnel connecting the surface with the active site of the enzyme. Two residues, a glutamate-arginine pair, were suggested to control 1745


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the opening and closing of the tunnel for controlling access of molecular oxygen to the flavin cofactor at the active site of the enzyme. A subsequent mutagenesis study on the gating residues provided corroborating evidence for this proposal.69 Also, for the flavoprotein monooxygenase 3-hydroxybenzoate hydroxylase from Comamonas testosteroni, a putative molecular oxygen tunnel has been identified upon elucidation of its crystal structure.70 In this case, the tunnel was suggested to be involved in guiding the diffusion of molecular oxygen based on the presence of a xenon atom in a pocket of the tunnel close to the flavin cofactor. Another mimic for molecular oxygen, the chloride ion, has revealed potential molecular oxygen binding pockets in the active sites of several flavoprotein oxidases.71,72 While such experimental evidence using molecular oxygen mimics may provide leads concerning preferred binding pockets for molecular oxygen, it does not fully disclose the complete pathway that molecular oxygen takes to reach the flavin cofactor starting from the exterior of a flavoprotein. Performing enhanced-statistics molecular dynamics simulations on various dioxygen-dependent flavoenzymes, i.e., alditol oxidase, p-hydroxyphenylacetate hydroxylase, histone demethylase LSD1, and sarcosine oxidase, revealed strategies for capturing and guiding molecular oxygen toward the active site.71,73−75 In these enzymes, multiple transient tunnels were found that guide molecular oxygen, from hydrophobic patches on the protein surface to the flavin cofactor. Intriguingly, these pathways all converge at a common entry point at the active site of the enzymes. By this funnel approach, molecular oxygen is efficiently and precisely guided to enter the active site to react with the C(4a)−N(5) locus of the flavin cofactor. The data also revealed that molecular oxygen and the organic substrate use distinct entry paths for reaching the flavin cofactor (Figure 2). The convergence of the molecular oxygen diffusion pathways at the entrance of the active site pocket was experimentally verified. The role of Ala105 in alditol oxidase as a gate keeper residue was assessed through the creation of mutant proteins. As predicted, by replacing alanine with glycine, the rate of oxidation of the reduced FAD in this oxidase was significantly increased.73 This confirmed the observation from the molecular dynamics data that Ala105 is blocking the access for dioxygen to reach the reduced flavin. On the basis of these insights, it was shown that a typical flavoprotein dehydrogenase could be switched into an reasonably effective flavoprotein oxidase.74 By replacing Ala113 in a plant L-galactono-γ-lactone dehydrogenase, which is structurally equivalent to Ala105 in alditol oxidase, a mutant enzyme (Ala113Gly) was created that acts as an L-galactono-γlactone oxidase. The mutation resulted in a 400-fold increase in the rate of flavin oxidation and confirms the ability of flavoproteins to shield their active site from small gaseous molecules such as dioxygen. The role of residues close to the flavin cofactor in tuning the rate of oxygen-mediated flavin reoxidation was also shown for a plant glucose dehydrogenase. This flavoprotein was converted into a very efficient oxidase by mutating a gatekeeper residue. The Ile153Val mutation transformed the enzyme into an efficient oxidase with a remarkable 60 000-fold increase in the oxygen reactivity (Figure 3).78 Likewise, by targeting a gatekeeper residue the rate by which dioxygen reacted with an aryl−alcohol oxidase was affected by 3−4 orders of magnitude. The crystal structure of this enzyme does not show open access to the active site.76 However, using the

Figure 3. Active site of the plant glucose dehydrogenase Phl p 4. Selection of residues and the flavin are shown in sticks for the wildtype enzyme (gold and yellow carbons, respectively; PDB code: 4PVE) as well as Val153 for the Ile153Val mutant (green carbons; PDB code: 4PWB). Superimposition of the crystal structure of Ile153Val/Asn158His mutant harboring a Br− atom (brown dots) in the active site (PDB code: 4PWC) and that of the Ile153Val mutant was carried out to show the position of the oxygen surrogate Br− in the cavity above the flavin C(4a) atom where dioxygen is likely accommodated in flavin-dependent oxidases.

Protein Energy Landscape Exploration (PELE) computational approach,79 it was possible to map the buried active site connected to the solvent by a hydrophobic funnel-shaped channel, with Phe501 and two other aromatic residues forming a narrow bottleneck that prevents direct access of alcohol substrates.80 Site-directed mutagenesis of Phe501 yielded a Phe501Ala variant with strongly reduced O2 reactivity. However, a variant with ∼120-fold increased reactivity was obtained with the Phe501Trp mutant. This shows that O2 reactivity can be increased by introducing, at the position contiguous to the flavin, a residue improving O2 positioning in the active-site cavity (Figure 4).80,81 The PELE approach was recently also exploited for deciphering the pathways for substrate, dioxygen, and hydrogen peroxide in vanillyl−alcohol oxidase. The calculations suggested that dioxygen and the formed hydrogen peroxide share the same pathway to enter or exit the active site of the oxidase.77

Figure 4. O2 at the aryl−alcohol oxidase active site.81 After migration from the solvent region to the active site of reduced enzyme, O2 adopts a catalytically relevant position near the flavin C(4a) atom (in yellow) and protonated His502 Hϵ atom (PDB code: 3FIM). 1746


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separate proteins to carry out the overall reaction; an NAD(P)H-dependent reductase delivers reduced FMN or FAD to the monooxygenase component, where the reaction with molecular oxygen and an organic substrate takes place. Current data indicate that in most cases the reduced flavin is transferred from the reductase to the oxygenase by a free diffusion mechanism.83,84 The rate of diffusion of the reduced flavin and binding to the oxygenase component must be larger than the reaction rate of free reduced flavin with molecular oxygen to minimize the waste of reduced flavin and the formation of hydrogen peroxide and other reactive oxygen species.85 As expected, the reductase component usually exhibits a lower Kd value for binding oxidized flavin than reduced flavin, and the opposite applies to the monooxygenase component.83 Finally, in the case of internal monooxygenases, the same substrate is oxygenated by the enzyme and also acts as electron donor in flavin reduction.86

The remaining sections of this review will discuss the catalytic mechanisms underlying the reaction of dioxygen once it has reached the reactive C(4a)−N(5) locus of the flavin. This will be discussed in connection to the enzymes that employ the dioxygen reactivity of this locus: the flavoprotein monooxygenases and oxidases.

4. FLAVIN-DEPENDENT MONOOXYGENASES Flavin-dependent monooxygenases utilize dioxygen to catalyze the incorporation of one oxygen atom into an organic substrate, reducing the other oxygen atom to water. Relevant industrial applications have been proposed for numerous flavin-dependent monooxygenases,37 which constitute the largest section within the class of flavin-dependent oxidoreductases.43 Flavin-dependent monooxygenases are classified into three categories based on the strategy used to reduce FMN or FAD (Figure 5): (I) one-component monooxyge-

4.1. O2 Activation in Flavoprotein Monooxygenases

In monooxygenases, reduction makes the flavin able to react with molecular oxygen, generally yielding a C(4a)−(hydro)peroxyflavin intermediate, which is formed with typical secondorder rate constants of 104−106 M−1 s−1.87−92 Only this catalytic step is dependent on molecular oxygen concentration (Schemes 2 and 3). The C(4a)−(hydro)peroxyflavin is responsible for the insertion of an oxygen atom into a substrate, yielding the second intermediate C(4a)−hydroxyflavin (Scheme 4). The oxidized cofactor is regenerated after elimination of the second oxygen atom from the flavin in the form of a water molecule. In the absence of an oxygen-acceptor substrate, the wasteful oxidation of NAD(P)H results in the formation of hydrogen peroxide instead of water. This process is known as uncoupling, and flavoproteins in general show much less wasteful cycles compared to other monooxygenases such as P450 enzymes.92,93 The versatile C(4a)−(hydro)peroxyflavin is able to perform very different chemical tasks, including hydroxylation, Baeyer−Villiger oxidation, sulfoxidation, epoxidation, halogenation, light emission, and oxidative decarboxylation reactions.82 Recently, the groups of Moore and Palfey have made the remarkable discovery of a monooxygenase, EncM, that does not operate through the formation of a C(4a)−(hydro)peroxyflavin. The catalytic mechanism of EncM involves the formation and use of a flavin N(5)−oxygen adduct (Scheme 4) as an oxygenating species instead of a flavin C(4a)−oxygen adduct. Details on this newly discovered mechanism of flavinmediated oxygenation will be discussed in section 4.1.8.86,94 In the following sections, we will present examples of the plethora of reaction types that are catalyzed by flavindependent monooxygenases to illustrate how these catalysts control the activation of dioxygen and modulate the reactivity of the flavin cofactor to perform specific chemical reactions. Their catalytic mechanism has been elucidated over the years taking advantage of the changes in the absorption and fluorescence properties of flavins which are associated with changes in their redox state and environment. Crystallographic studies have also been instrumental in elucidating the mechanism of flavin-dependent monooxygenases, mainly by reporting on important residues and conformational changes associated with catalysis. Additionally, structure-inspired enzyme engineering efforts have been instrumental to probe the role of active site residues in the catalytic mechanism of flavoproteins.

Figure 5. Categories of flavin-dependent monooxygenases (MOs) based on the strategy used to reduce FMN or FAD. There are three categories: one-component monooxygenase, two-component monooxygenases, and internal monooxygenases.

nases (groups A and B), (II) two-component monooxygenases (groups C−F), and (III) internal monooxygenases (groups G and H).37,82 One-component monooxygenases contain FAD as a prosthetic group, which is tightly bound to the enzyme active site during the whole catalytic cycle. They are structurally composed of a dinucleotide-binding domain for FAD binding. The group B flavoprotein monooxygenases also have a dinucleotide-binding domain for NAD(P)H, while group A members bind the reduced nicotinamide coenzyme in a less tight manner. These enzymes use NAD(P)H as a hydride donor; NAD(P)+ is released immediately upon flavin reduction (class A) or kept bound during the catalytic cycle (group B). Two-component flavin-dependent enzymes require two 1747


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Scheme 4. Proposed Flavin−Oxygen Adducts Reacting with a Substrate in Monooxygenasesa

a

R is ribityl adenosine diphosphate for FAD and phosphoribityl for FMN. S is substrate.

4.1.1. Aromatic Hydroxylation. Many microorganisms produce flavin-dependent hydroxylases to use aromatic compounds as a carbon and energy source as well as to synthesize natural products. The aromatic hydroxylations have been found to be catalyzed by members of two specific flavoprotein monooxygenase groups: group A one-component and group D two-component flavoprotein monooxygenases.82 Depending on the hydroxylase, NADH or NADPH is used as an electron donor and hydroxylation occurs at the position ortho or para to a substrate hydroxyl group (Scheme 5). Among the one-component ortho-hydroxylases, 4-hydroxybenzoate 3-hydroxylase (PHBH) from the soil bacterium Pseudomonas f luorescens is the best characterized (Schemes 5 and 6).95 PHBH exhibits a narrow substrate specificity, efficiently acting only on phenolic acids at the expense of NADPH.90 PHBH and other Group A aromatic hydroxylases contain only one Rossmann fold motif which is involved in binding of the ADP moiety of FAD.95 NADPH binds in contact with several residues located on the protein surface.96 The catalytic cycle of PHBH requires the formation of a ternary complex between the enzyme-bound oxidized FAD, the aromatic substrate, and NADPH (Scheme 6).90,97 The reduction of the enzyme is the next step in the turnover of PHBH, which is followed by an immediate release of NADP+. The PHBH reduction rate is 105-fold slower in the absence of a phenolic substrate because substrate deprotonation of the hydroxyl group, which is facilitated by a hydrogen-bond network that extends to the protein surface, plays an important role in catalysis.98 Specifically, substrate deprotonation makes the substrate a better nucleophile and triggers a conformational change that is required for the movement (∼7 Å) of the flavin isoalloxazine ring to a more solvent-accessible region in the proximity of the NADPH nicotinamide ring (see Figure 6).96,99 The subsequent reaction of the reduced flavin anion with molecular oxygen occurs after the flavin isoalloxazine ring returns to a solvent-free environment and the substrate is reprotonated by the hydrogen-bond network. As a result, an electrophilic C(4a)−hydroperoxyflavin intermediate is formed. The terminal oxygen of the hydroperoxide is transferred to the substrate, forming a nonaromatic molecule which rearranges to an aromatic structure, probably by transferring a proton to the flavin resulting in a C(4a)−hydroxyflavin intermediate.93 After release of the hydroxylated product, a water molecule is produced to yield the oxidized FAD. A key feature of this mechanism is the role played by substrate binding and protonation/deprotonation in the oxygen activation. Substrate binding favors the reaction of the reduced flavin with molecular oxygen, which becomes 10 times slower in the absence of the aromatic ligand. Furthermore, the reactivity

Scheme 5. Examples of Aromatic Hydroxylations Catalyzed by Flavin-Dependent Monooxygenasesa

a

PHBH, 4-hydroxybenzoate 3-hydroxylase; 3HB6H, 3-hydroxybenzoate 6-hydroxylase; MHPCO, 2-methyl-3-hydroxypyridine-5-carboxylic acid oxygenase; HPAH, 4-hydroxyphenylacetate 3-hydroxylase; MHPC, 2-methyl-3-hydroxypyridine-5-carboxylic acid; AAMS, E-2(acetamino-methylene)succinate.

with C(4a)−hydroperoxyflavin is enhanced by the deprotonation of the substrate hydroxyl group, whereas the subsequent reprotonation favors rearomatization to generate the final 3,4dihydroxybenzoate product.100 Other single-component hydroxylases catalyzing electrophilic aromatic substitutions have been studied, including 3hydroxybenzoate 6-hydroxylase, phenol hydroxylase, and 2methyl-3-hydroxypyridine-5-carboxylic acid oxygenase.88,101,102 These flavoprotein monooxygenases were all found to be similar to PHBH in that multiple conformations, including 1748


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Scheme 6. Catalytic Cycle of PHBH with 4-Hydroxybenzoate (4-OHB) as a Substratea

a

R is ribityl adenosine diphosphate for FAD. 3,4-Dihydroxybenzoate (3,4-DOHB) is produced. B represents a hydrogen-bond network that facilitates substrate and product deprotonation. In addition, this hydrogen-bond network may donate a proton to the carbonyl oxygen of the nonaromatic product. This catalytic cycle involves at least three enzyme conformations: “out” form, for enzyme reduction; “in” form, for substrate hydroxylation; and “open” form, for substrate binding and product release (see Figure 6).

Two-component flavin-dependent monooxygenases have been also mechanistically investigated.105 Several of these monooxygenases catalyze the hydroxylation of aromatic compounds, as exemplified by the well-characterized 4hydroxyphenylacetate 3-hydroxylase from Acinetobacter baumannii (Scheme 5).91,106−108 An outstanding property of this enzyme is that the C(4a)− hydroperoxyflavin intermediate is stabilized for minutes, having a slow hydrogen peroxide elimination rate (0.003 s−1) when no suitable substrate is present. Furthermore, the bimolecular rate constant for the intermediate formation is >100 times higher in the absence of the aromatic substrate. Accordingly, crystallographic studies suggested that molecular oxygen may have easier access to the isoalloxazine ring in the absence of the 4hydroxyphenylacetate substrate.109 Structural studies highlighted the cavity where the flavin C(4a)−hydroperoxide adduct can be accommodated (Figure 7).109 The C(4a)− hydroperoxyflavin is predicted to be stabilized within this cavity via a hydrogen bond between the Oγ atom of Ser171 and the flavin N(5)−H atom, since this interaction avoids the deprotonation at the flavin N(5)−H.85 Consistently, stoppedflow studies showed that the Ser171Ala mutant enzyme exhibits a 1400-fold increase for the rate constant of hydrogen peroxide elimination, compared to wild-type enzyme.85 A similar interaction is likely stabilizing the C(4a)−hydroperoxyflavin in other enzymes (section 5.1.3).85,110 Computational and experimental studies have shown that molecular oxygen is activated by the concomitant acceptance of an

Figure 6. Comparison of the solvent accessibility of the FAD in various PHBH crystal structures. During enzyme catalysis, the flavin isoalloxazine ring moves (∼7 Å) to a more solvent-accessible out position where it can be reduced by NADPH (out form, light blue surface and sticks, PDB code: 1DOD; in form, yellow surface and sticks, PDB code: 1PBE).

flavin movement, facilitate their complex catalysis.103,104 Structural studies demonstrate that regiospecificity in these aromatic hydroxylases is determined by differences in the arrangement of active-site residues that do not affect the overall mechanism for dioxygen activation and substrate monooxygenation/hydroxylation (Scheme 5). 1749


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as a prosthetic group. These enzymes preferentially utilize NADPH as coenzyme. On the contrary, type II BVMOs (Group C flavoprotein monooxygenases) are two-component enzymes consisting of NADH-dependent flavin reductase and oxygenase protomers. The first cloned BVMO-encoding gene was that of cyclohexanone monooxygenase from Acinetobacter sp. strain NCIMB 9871 in 1988.117 The catalytic mechanism of this and related BVMOs has been elucidated based on steady-state and rapid kinetics and computational studies (Scheme 8).87,129−131 The reaction starts when NADPH binds in the active site to reduce the FAD. The reduced prosthetic group then reacts with molecular oxygen forming a C(4a)−peroxyflavin intermediate, while NADP+ remains bound to the enzyme. The requirement in catalysis of a deprotonated flavin−molecular oxygen adduct was confirmed by studying the reaction at two different pH values.87 The C(4a)−peroxyflavin intermediate is able to perform a nucleophilic attack on the electron-poor carbonyl carbon of a ketone or aldehyde substrate, resulting in the formation of a tetrahedral intermediate. This mechanism is generally accepted in analogy to the mechanism postulated by Criegee for chemical Baeyer−Villiger reactions almost 70 years ago, which has been experimentally confirmed.132,133 However, the tetrahedral intermediate, known as Criegee intermediate, has never been observed experimentally in enzyme-catalyzed Baeyer−Villiger reactions. Mutagenesis data indicate that a highly conserved arginine side chain is responsible for its stabilization.134 The rearrangement of the Criegee intermediate to give the C(4a)−hydroxyflavin and an ester or lactone as product involves the migration of one of the carbon atoms adjacent to the substrate carbonyl group to the closest oxygen atom of the peroxide group. The more nucleophilic carbon, which in most cases is the more substituted one, generally migrates to give the so-called normal regioisomer (R3C > R2HC > phenyl > RH2C > methyl) (Scheme 8).131,135 Migration of the less nucleophilic carbon yielding the abnormal regioisomer has been also reported in several BVMOs.136−143 NADP+ release is the last step before the next catalytic cycle starts with binding NADPH. For cyclohexanone monooxygenase, a conformational change preceding the release of NADP+ was found to be the ratelimiting step in catalysis. A crucial feature of this mechanism is that the C(4a)−peroxyflavin intermediate is not observed during the reoxidation of the enzyme in the absence of NADP+. Accordingly, crystallographic studies consistently confirmed that NADP+ plays a crucial role in stabilizing the C(4a)−peroxyflavin intermediate by providing hydrogenbonding interactions (Figure 8).119,131,134,144−147 Thus, NADP(H) serves the dual role of being the flavin-reducing agent, while it is also an essential catalytic element for C(4a)− peroxyflavin stabilization. Interestingly, a BMVO was recently converted into an efficient NADPH oxidase by replacing only one residue close to the flavin C(4a)/N(5) locus (Cys65 in phenylacetone monooxygenase).145 The rate of decay of the C(4a)− peroxyflavin intermediate was significantly increased in the resulting mutant compared to the wild-type enzyme (25 and 0.01 s−1, respectively). The structural analysis indicated that the mutation perturbs the conformation of the enzyme-bound NADP+, pointing out the importance of NADP+ in the stabilization of the C(4a)−hydroperoxyflavin. It was shown that the engineered NADPH oxidase could be used as biocatalyst to support NADP+ regeneration required for an

Figure 7. Active site of 4-hydroxyphenylacetate 3-hydroxylase from A. baumannii. Selection of residues (gold carbons) and 4-hydroxyphenylacetate (HPA, cyan carbons) are shown in sticks (PDB code: 2JBT). Distance in Angstroms between the FMN N(5) atom and Ser171 and that between the FMN C(4a) atom and either His396 or 4-hydroxyphenylacetate (HPA) is indicated next to the broken lines. Superimposition of FMN in the crystal structure of 4-hydroxyphenylacetate 3-hydroxylase and the FAD C(4a)−oxygen adduct (C(4a)− O) in the crystal structure of pyranose dehydrogenase (PDB code: 4H7U) was carried out to infer the position of the cavity where the flavin C(4a)−hydroperoxide adduct is likely accommodated in 4hydroxyphenylacetate 3-hydroxylase.

electron from reduced FMN and a proton from a protonated histidine (His396, 4.6 Å from the flavin C(4a) atom).111 Subsequently, the nearly barrierless formation of a C(4a)− hydroperoxyflavin intermediate occurs. Scheme 7 shows a comparison of the proposed proton-coupled electron-transfer mechanism for molecular oxygen activation in 4-hydroxyphenylacetate 3-hydroxylase and previously described mechanisms for molecular oxygen activation in flavin-dependent monooxygenases and oxidases. A proton-coupled mechanism has been also suggested for a fungal aryl−alcohol oxidase, and future research will tell whether other flavoenzymes use this type of mechanism.81 4.1.2. Baeyer−Villiger Oxidation. In 1899, Baeyer and Villiger demonstrated the conversion of cyclic ketones into lactones using potassium monopersulfate (Caro’s acid) as oxidant.112 Currently, the Baeyer−Villiger reaction is one of the most important transformations in organic synthesis. Conversion of ketones into esters or cyclic ketones into lactones is generally carried out using percarboxylic acids.113,114 A safer and more environmentally friendly alternative is the implementation of flavin-dependent Baeyer−Villiger monooxygenases (BVMO).115 The most characterized BVMOs are from bacterial origin, a few from fungi, and two from photosynthetic organisms. 116−121 However, genes encoding putative BVMOs have also been found in Metazoa.116,122−126 Recently, it was also shown that a human flavin-containing monooxygenase (FMO5) primarily acts as a BVMO.124 In general, BVMOs proceed with excellent regio-, chemo-, and enantioselectivity using NAD(P)H and molecular oxygen as electron donor and acceptor, respectively.36 In addition to Baeyer−Villiger reactions, BVMOs perform oxidation of aldehydes and heteroatoms and epoxidation reactions.36,127 BVMOs are classified into three types, referred as type I, type II, and “atypical, odd, O, or type III” BVMOs.36,82,128 Type I BVMOs (Group B flavoprotein monooxygenases) and “O” BVMOs (Group A flavoprotein monooxygenases) are one-component enzymes that use FAD 1750


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Scheme 7. Mechanisms of Molecular Oxygen Activation and Flavin−Oxygen Adducts in Oxidases and Monooxygenasesa

a

Four mechanisms are shown: (i) OX, typical for oxidases; (ii) MO, typical for monooxygenases; (iii) PCET, proton-coupled electron transfer in the oxidases pyranose oxidase and a fungal aryl−alcohol oxidase and the monooxygenase 4-hydroxyphenylacetate 3-hydroxylase; and (iv) N(5)oxide, in the monooxygenase EncM. Neutral flavin semiquinone is formed in OX, MO, and PCET mechanisms. Anionic flavin semiquinone is formed in N(5)-oxide mechanism. Mechanism OX and MO involves superoxide anion radical, while mechanisms PCET and N(5)-oxide involve the protonated form of superoxide anion radical named perhydroxyl radical. To form the perhydroxyl radical in PCET mechanism, molecular oxygen simultaneously accepts an electron from the flavin and a proton from an active site residue. To form the perhydroxyl radical in the N(5)oxide mechanism, both the electron and the proton are transferred from the flavin to molecular oxygen. R is ribityl adenosine diphosphate for FAD and phosphoribityl for FMN.

protein sequence databanks together with the two Rossmann fold motifs.121,150,151 Interestingly, the FMO-conserved motif differs from the BVMO-conserved motif [FxGxxxHxxxW(P/ D)] by only one residue.121,151−154 Studies carried out with FMO1 have demonstrated that the mechanism of FMO is generally similar to that of BVMOs.155 As in BVMOs, NADP+ is essential for the stabilization of the C(4a)−hydroperoxyflavin, and it remains bound to the enzyme throughout the catalytic cycle, being the last product to be released from the active site of the enzyme. Structure−activity studies have suggested that, in addition to nucleophilicity, size and charge determine which compounds are able to access and react with the C(4a)−hydroperoxyflavin intermediate in FMOs.156

alcohol dehydrogenase-catalyzed oxidation reaction. This structure-inspired enzyme engineering result shows the value of structural information and mechanistic knowledge. 4.1.3. Heteroatom Oxygenation. Flavin-containing monooxygenases (FMOs) catalyze the oxidation of soft nucleophilic heteroatoms in a large variety of compounds. FMOs are one-component enzymes containing FAD as a prosthetic group, which is reduced by NADPH. In 1972, the first FMO was isolated from pig liver microsomes.148 Mammalian FMOs, which arose by divergent evolution from a single ancestral gene, are designated by an Arabic numeral (FMO1-FMO5).149 FMOs are sequence related to the type I BVMOs mentioned above. The FMO-conserved motif FxGxxxHxxx(Y/F) can be used to identify FMOs in the 1751


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intermediate lies in the proximity of NADP+, which remains bound to the enzyme throughout the catalytic cycle. It was found that the rate of formation of the C(4a)−hydroperoxyflavin intermediate in ornithine monooxygenase is greatly enhanced by L-arginine.164 Crystallographic studies showed that L-arginine binds to the active site in a similar position to that of L-ornithine (Figure 8).161 It was suggested that the positive charge of this compound may electrostatically facilitate the transfer of one electron from the anionic reduced flavin to molecular oxygen to form superoxide anion and flavin semiquinone, which rapidly collapse to form the C(4a)− hydroperoxyflavin (Scheme 7).161,164 Indeed, structurally similar compounds having uncharged side chains do not act as efficient effectors for the C(4a)−hydroperoxyflavin formation.164 Of note, not all family members feature a highly stable C(4a)−hydroperoxyflavin as indicated by the investigation of lysine hydroxylase of Mycobacterium smegmatis. This enzyme is specific for L-lysine; however, its reaction is highly uncoupled, producing high levels of superoxide and hydrogen peroxide besides hydroxylated L-lysine.165 Accordingly, a C(4a)−(hydro)peroxyflavin intermediate was not observed by reacting the reduced enzyme with molecular oxygen using rapid kinetic techniques.166 The physiological significance, if any, of such a pronounced leakage of reactive oxygen species remains unknown. Interestingly, structural data on the ornithine monooxygenase from Kutzneria sp. 744 suggest that there is a conformational change involving a movement of the reoxidized isoalloxazine ring (6.5 Å) to a solvent exposed environment, which facilitates NADP+ release at the end of each catalytic cycle. It remains to be elucidated whether this conformational change, to eject the NADP+ from the enzyme active site, takes place in other monooxygenases of this family.167 4.1.4. Light Emission. The enzymes that catalyze bioluminescence reactions in different organisms are called luciferases. Bacterial luciferase was the first enzyme shown to be a two-component flavin-dependent monooxygenase.168 All bacterial luciferases are found in Gram-negative bacteria, mainly from the Vibrionaceae family.169 Luciferase converts a long-chain aldehyde into an aliphatic carboxylic acid producing

Scheme 8. Catalytic Cycle of Cyclohexanone Monooxygenase with Cyclohexanone (Cyc) as a Substratea

a R is ribityl adenosine diphosphate for FAD. ε-Caprolactone (ε-Cap) is produced.119

N-Hydroxylating monooxygenases are sequence related to both BVMOs and FMOs.121 As for all BVMOs and FMOs, these enzymes are single-component FAD-containing enzymes and their sequences contain two Rossmann fold motifs. They have been found in bacteria and fungi, where they catalyze the hydroxylation of the terminal amino group of long-chain primary amines in the production of low-molecular-weight siderophores.157−159 The best characterized family members are the ornithine monooxygenases from Aspergillus f umigatus (named SidA) and P. aeruginosa.160−163 Their catalytic mechanism is similar to that described for FMOs and BVMOs in that a long-lived C(4a)−hydroperoxyflavin

Figure 8. Active sites of phenylacetone monooxygenase and ornithine monooxygenase (SidA). (A) In phenylacetone monooxygenase (PDB code: 2YLT) the active site is occupied by a molecule of 2-(N-morpholino)ethanesulfonic acid, a mimic of the Criegee intermediate (cyan carbons). Arg337 is a key residue in the architecture of the active site for the stabilization of the Criegee intermediate. (B) In SidA (PDB code: 4B68), Arg144 interacts with the NADP+ carbamide. L-Arginine ligand (purple carbons) binds in a similar position to that of the L-ornithine substrate. 1752


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Scheme 9. Catalytic Mechanism of Bacterial Luciferasea

a

R is phosphoribityl for FMN, and R is (CH2)nCH3 for the aldehyde substrate where n is 6-14. Three mechanisms to form the excited C(4a)− hydroxyflavin intermediate have been proposed: (A) Baeyer−Villiger reaction, (B) dioxirane intermediate, and (C) chemically initiated electron exchange luminescence (CIEEL). NADH-dependent FMN reductase (LuxG) delivers reduced FMN to the oxygenase component of bacterial luciferase (LuxAB).

blue-green light (λmax ≈ 490 nm). Tetradecanal (myristaldehyde) is the natural aldehyde substrate for bacterial luciferase, but many long-chain aldehydes ranging from octanal to hexadecanal are in vitro substrates for this enzyme.170 The general catalytic cycle of bacterial luciferases has been elucidated (Scheme 9).171 The α-subunit of luciferase contains the active site, while the β-subunit stabilizes the active conformation of the α-subunit.172 Reduced FMN binds to the α-subunit, where it reacts with molecular oxygen forming a C(4a)−peroxyflavin intermediate. 13C NMR studies on luciferase provided the first direct proof for addition of molecular oxygen at the C(4a) position of the enzyme-bound reduced flavin.173 When the nucleophilic C(4a)−peroxyflavin intermediate reacts with an aldehyde substrate, a flavinC(4a)−peroxyhemiacetal adduct is formed. This adduct decomposes to produce a carboxylic acid and the excited state of the C(4a)−hydroxyflavin intermediate, which emits blue-green light to return to the ground-state species. Finally, a water molecule is proposed to be released from the C(4a)− hydroxyflavin yielding oxidized FMN. In the absence of an

aldehyde substrate, a stable C(4a)−peroxyflavin intermediate is formed and its decay produces hydrogen peroxide without light emission. 4.1.5. Halogenation and Decarboxylative−Halogenation. Typical FAD-dependent halogenases are two-component monooxygenases. These enzymes introduce chloride and bromide into activated organic compounds. Depending on the flavin-dependent halogenase, the substrate is a free molecule or it is bound to an acyl carrier protein as a thioester. All sequences of the oxygenating component of flavin-dependent halogenases feature a dinucleotide-binding motif (GxGxxG) and a unique motif containing two tryptophan residues located near the flavin (GWxWxIP). Flavin-dependent halogenases are generally highly substrate specific and regioselective, in contrast to haloperoxidases.174 Dairi and co-workers identified the first gene coding for a flavin-dependent halogenase involved in the biosynthesis of the antibiotic 7-chlortetracycline.175 Subsequently, the first direct evidence for flavin-dependent halogenating activity was obtained in vitro for tryptophan 7-halogenase (PrnA) and 1753


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Scheme 10. Active Site and Catalytic Mechanism of the Halogenase PrnAa

a

PrnA catalyzes the chlorination or bromination of the tryptophan C(7) atom. Hypohalous acid (HOX) diffuses within the protein matrix to encounter the substrate-binding site. Whether hypohalite (−OX) forms a covalent haloamine intermediate or a tightly coordinated hypohalous acid is not clear. The inset shows the crystal structure of PrnA in complex with both the tryptophan substrate (S, cyan sticks) and the chloride ion (green sphere) showing important residues for catalysis (pink sticks) and the FAD (yellow sticks; PDB code: 2AQJ). Distance in Angstroms between the FAD C(4a) atom and the chloride ion and that between the chloride ion and the substrate C(7) atom is indicated next to the broken lines.

Scheme 11. Proposed Mechanism for the Decarboxylative−Brominase Reaction Catalyzed by Bmp5181

monodechloroaminopyrrolnitrin 3-halogenase (PrnC) using their natural substrates.176 Both enzymes participate in the biosynthesis of the antifungal antibiotic pyrrolnitrin in P. f luorescens. The determination of the first crystal structure for a flavin-dependent halogenase (PrnA) was a major contribution to the understanding of halogenation chemistry by flavindependent enzymes.177 The structure showed that the FAD and the substrate tryptophan are separated by a 10 Å tunnel, suggesting the participation of a diffusible halogenating agent in the reaction. On the basis of this and other studies, a catalytic mechanism for flavin-dependent two-component halogenases has been proposed (Scheme 10).178 The overall halogenation reaction requires a FAD reductase to provide reduced FAD, which diffuses to the halogenase active site, where it reacts with molecular oxygen forming a C(4a)− hydroperoxyflavin intermediate. Next, a halide anion makes a nucleophilic attack on the flavin peroxide yielding hypohalite and C(4a)−hydroxyflavin. Dehydration of the latter intermediate yields oxidized FAD. Both intermediates have been detected by stopped-flow spectroscopy.179 Halogenation of the substrate occurs by electrophilic aromatic substitution. The mechanism to facilitate the reaction and determine regioselectivity is not fully understood yet. A covalent haloamine

intermediate may take place in the reaction of hypohalous acid with a conserved Lys residue.180 A one-component flavin-dependent decarboxylative-brominase has been recently discovered during a study of the origin of polybrominated aromatic organic compounds in marine environments.182 This enzyme, called Bmp5, is produced by epiphytic marine bacteria of the genera Pseudoalteromonas and Marinomonas (Scheme 11). Interestingly, this novel decarboxylative-brominase shows the highest sequence identity with FMOs instead of with two-component flavin-dependent halogenases. Bmp5 performs the bromination of 4-hydroxybenzoate and the subsequent decarboxylative-bromination of the resulting 3-bromo-4-hydroxybenzoate yielding 2,4-dibromophenol. In a subsequent study, four bromocatechols were detected in the reaction of Bmp5 with 3,4-dihydroxybenzoic acid.181 Bmp5 is able to use iodide too, forming iodophenols, whereas chloride is not a substrate.182 4.1.6. Dehalogenation. There are flavin-dependent monooxygenases involved in dehalogenation reactions in which a chlorine atom placed at the para position of a phenolic ring is eliminated forming a quinone. Microbes producing these enzymes present potential use for detoxification of chlorophenols accumulated in the environment. 1754


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Scheme 12. Dehalogenation through Monooxygenationa

a C(4a)−hydroperoxyflavin of PcpB catalyzes the oxidative elimination of Cl − from pentachlorophenol to yield the unstable tetrachlorobenzoquinone.183

Scheme 13. Catalytic Mechanism of Nitronate Monooxygenase (NMO) with Propionate 3-Nitronatea

a Two possible mechanisms to form 3-peroxy-3-nitropropanoate have been proposed: (A) flavosemiquinone transfers an electron to molecular oxygen forming superoxide anion and reoxidized flavin, followed by reaction of superoxide anion with the substrate radical; (B) substrate radical reacts with molecular oxygen yielding 3-peroxy-3-nitropropanoate radical, which accepts an electron from the flavosemiquinone.

The best studied single-component flavin-dependent dechlorinase is PcpB from Sphingobium chlorophenolicum, which catalyzes the first step in the degradation of pentachlorophenol, a highly toxic compound that was widely used as biocide and wood preservative and is banned since 2004. As in PHBH and other aromatic hydroxylases, the rate of NADPH oxidation in the enzyme is significantly enhanced by the binding of the aromatic substrate and may involve similar conformational changes to those shown for PHBH.183 The reaction of the dechlorinase with NADPH, pentachlorophenol, and molecular oxygen gives a C(4a)−hydroperoxyflavin intermediate, which transfers a OH to pentachlorophenol (Scheme 12).183−187 The catalytic cycle is completed with the formation and dehydratation of C(4a)−hydroxyflavin to regenerate the oxidized flavin. The catalytic mechanism of a two-component dechlorinase, Rp-HadA, has been recently studied for the first time.188 RpHadA acts on various chlorophenols, preferring 4-chlorophenol and 2-chlorophenol rather than chlorophenols with multiple substituents. Rp-HadA binds reduced FAD, supplied by a flavin reductase. The reduced FAD then reacts with molecular oxygen to form a C(4a)−hydroperoxyflavin intermediate. By transferring the OH from the C(4a)− hydroperoxyflavin to the chlorophenol, Cl− elimination takes place. The hydroxylation step is the main rate-limiting step of the Rp-HadA reaction. In the absence of the chlorophenol, the

C(4a)−hydroperoxyflavin eliminates hydrogen peroxide very slowly (0.005 s−1, at 25 °C). 4.1.7. Denitrification. 3-Nitropropionate, which exists in equilibrium with propionate 3-nitronate, is found in plants and fungi.189 3-Nitropropionate serves in plants as a defense against herbivores by interrupting their Krebs cycle, and it also participates in nitrogen fixation in some legumes. Nitronate monooxygenase, previously known as 2-nitropropane dioxygenase, protects organisms against the toxicity of propionate 3nitronate. This FMN-dependent internal monooxygenase catalyzes the denitrification of propionate 3-nitronate with impressive speed (kcat > 103 s−1 and kcat/Km > 106 M−1 s−1), and it is also able to act on other nitronate analogues.190,191 Nitronate monooxygenase is one of the few flavin-dependent monooxygenases that do not form a C(4a)−(hydro)peroxyflavin intermediate (Scheme 13).192,193 Rather, the anionic flavosemiquinone corresponds to the first flavin absorption spectrum observed after mixing the enzyme with the substrate under anaerobic conditions using a stopped-flow instrument. Therefore, a very fast single-electron transfer (≥1900 s−1, at 7 °C) takes places within the mixing time of the instrument (i.e., 2.2 ms) yielding a radical species of propionate 3-nitronate. The flavosemiquinone may transfer an electron to molecular oxygen forming superoxide anion and oxidized flavin, followed by reaction of superoxide anion with the propionate 3-nitronate radical species to give 3-peroxy-31755


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Scheme 14. Catalytic Mechanism of EncM with a Polyketide Substratea

a EncM-bound flavin-N(5)-oxide transfers an oxygen atom to the substrate. The resulting product is then oxidized to regenerate EncM-bound reduced FAD. The electrophilic ketone moiety of the dehydrogenation product triggers a Favorskii-type rearrangement forming the lactone ring of enterocin. Schemes 4 and 7 show more details regarding the reaction of reduced enzyme with molecular oxygen and the dehydration step. R is ribityl adenosine diphosphate for FAD, and R is (CH2CO)4-S-EncC for the polyketide substrate.

transfers a hydrogen atom from the flavin N(5) atom to molecular oxygen yielding protonated superoxide and anionic flavin semiquinone. Alternatively, the formation of these species may be a two-step process involving one-electron transfer followed by proton transfer. The flavin-N(5)-peroxide is formed by the reaction of protonated superoxide and anionic flavin semiquinone. The high spin density at the N(5) atom of the anionic flavin semiquinone favors the formation of a N(5) adduct rather than a C(4a) adduct. After water elimination from the flavin-N(5)-peroxide, the N(5)−oxoammonium resonance form is expected to be formed. EncM catalyzes a key step in the biosynthesis of the antibiotic enterocin in Streptomyces maritimus using FAD covalently bound to a histidine as a prosthetic group. Presumably, the flavin-N(5)-oxide in EncM can hydroxylate a polyketide substrate bound to an acyl carrier protein (Scheme 14). Future research will tell whether a N(5)− nitroxyl radical, a nucleophilic flavin-N(5)-oxide, or an electrophilic oxoammonium resonance form of the N(5)oxide participates in the mechanism for monooxygenation with a flavin N(5)−oxygen (Scheme 4).194,195 Next, the resulting hydroxyl group is probably converted by EncM into an electrophilic ketone moiety, which would be able to trigger the subsequent Favorskii-type rearrangement forming the lactone ring of enterocin.86 The dehydrogenation reaction regenerates the EncM-bound reduced FAD, which reacts with molecular oxygen forming the flavin-N(5)-oxide to carry out the next catalytic cycle. EncM can be regarded as an internal monooxygenase, since both the hydroxylation and the dehydrogenation catalyzed by EncM take place in the absence of external reductants. The exciting discovery of a flavoenzyme that employs a flavin-N(5)-oxide has triggered a search for other flavindependent enzymes using a flavin-N(5)-oxide as the oxygen-

nitropropanoate (Scheme 13). Alternatively, the radical may react with molecular oxygen yielding 3-peroxy-3-nitropropanoate radical, which would then accept an electron from the flavosemiquinone. Subsequently, 3-peroxy-3-nitropropanoate decays either in the enzyme active site or after its release into solution. Products detected for the reaction of NMOs from different sources are malonic semialdehyde, nitrate, nitrite, and at least for one case hydrogen peroxide.189 4.1.8. Hydroxylation−Dehydrogenation. Recently, the participation of a flavin-N(5)-oxide in an enzyme-catalyzed reaction has been shown for the first time.86,94 UV−vis absorption spectroscopic, mass spectrometric, and 18O-labeling studies have provided convincing evidence to establish that a flavin-N(5)-oxide participates in the reaction of the monooxygenase EncM instead of a C(4a)−(hydro)peroxyflavin intermediate (Scheme 4). Interestingly, EncM has a vanillyl− alcohol oxidase fold, which is usually observed for flavindependent dehydrogenases/oxidases rather than oxygenases. The flavin-N(5)-oxide intermediate in EncM is remarkably stable (at least >7 days at 4 °C). The absorption spectrum of flavin-N(5)-oxide, both enzyme bound or chemically synthesized free in solution, presents a “bridge” in the 300−340 nm region, a lower (∼20%) extinction coefficient in the 450 nm region than oxidized flavin, and a red-shifted absorption maximum at 450 nm.86 These spectral properties clearly contrast with those observed for the C(4a)−(hydro)peroxyflavin intermediate, which has high absorbance around 360−400 nm and very low absorption in the 450 nm region. Mass analysis of the flavin−oxygen adduct in EncM is consistent with the FMN-N(5)-oxide structure.94 When EncM bound to reduced FAD was reacted with 18O2, the mass analysis showed the expected 2 Da mass increase.94 A mechanism for the formation of the flavin-N(5)-oxide in EncM has been proposed (Scheme 7).94 EncM-bound reduced FAD 1756


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large-scale fermentation processes (e.g., L-aspartate oxidase).203 There are several factors that contribute to their attractiveness as biocatalysts. The fact that they can be used without any special requirements (they merely require dioxygen for activity) makes them easy to use while they can catalyze highly attractive reactions.204 Comparison of the currently available sequences and structures allows one to group the flavoprotein oxidases into six distinct families (Table 1).204 The relatively large glucose-

ating agent. This has resulted in the identification of few other flavoenzymes, distinct in sequence from EncM, that may also rely on formation of a flavin-N(5)-oxide intermediate. Using similar approaches to those reported for EncM, the formation of a flavin-N(5)-oxide has been confirmed in the FMNdependent two-component monooxygenases DszA and RutA.196,197 DszA, a sulfone monooxygenase from Rhodococcus erythropolis, catalyzes an oxidative C−S bond cleavage reaction converting dibenzothiophene sulfone to 2-(2-hydroxyphenyl)benzenesulfinic acid in the dibenzothiophene catabolic pathway.196 In this case, the proposed mechanism involves transfer of oxygen from the substrate hydroperoxide to oxidized flavin to form the flavin-N(5)-oxide (Scheme 15).196 A similar

Table 1. Six Major Flavoprotein Oxidase Families204 family

cofactor

GMC

FAD

alcohols/aldehydes/carbon acids

AO VAO ACO SO HAO

FAD FAD FAD FAD FMN

amines alcohols/amines acyl-CoA thiols hydroxyacids

Scheme 15. Simplified Catalytic Mechanism of Flavin-N(5)Oxide Formation in DszA and RutAa

substrate types

prototype enzyme glucose oxidase/methanol oxidase/choline oxidase D-amino acid oxidase vanillyl−alcohol oxidase acyl-CoA oxidase sulfhydryl oxidase glycolate oxidase

methanol-choline (GMC) oxidoreductase flavoprotein superfamily205 contains a number of well-known flavin-containing oxidases, such as glucose oxidase, cholesterol oxidase, pyranose oxidase, methanol oxidase, aryl−alcohol oxidase, choline oxidase, formate oxidase, and 5-hydroxymethylfurfural oxidase. The protein sequence of each member of this family entails a conserved N-terminal FAD-binding domain with a Rossmann fold involved in binding the ADP moiety of the FAD.202 GMCtype oxidases typically act on alcohols. A similar N-terminal FAD-binding domain as found in GMC-type flavoprotein oxidases is also present in the FADcontaining amine oxidases. As the name suggests, most enzymes of this large oxidase family are solely active on amines as illustrated by sarcosine oxidase, monoamine oxidases, and D-amino acid oxidase.200,201,206 There seems to be a consensus that the catalytic reaction in GMC and amine oxidase enzymes starts with a hydride transfer from the organic substrate to the enzyme-bound FAD.207,208 Flavoproteins of the VAO family,209,210 named after the eponymous fungal vanillyl−alcohol oxidase, contain a distinct FAD binding domain in the N-terminal half of the protein.211 In the past decade it has become clear that a subfamily of VAO-type enzymes also contains a bicovalently linked FAD cofactor in which the FAD is linked to a cysteine and a histidine.65,212 All bicovalent flavoproteins discovered so far act as oxidases, which correlates well with the high redox potentials displayed by these enzymes. This seems logical as such high redox potentials limit the range of natural electron acceptors, leaving molecular oxygen as one of the few candidates. It has also been suggested that the second covalent bond between the flavin and the protein allows for an active site widely open to the bulk solvent while being able to hold the FAD cofactor in a suitable position for catalysis through the bicovalent anchoring. This is in line with the observation that the bicovalent flavoprotein oxidases characterized thus far accept rather bulky substrates.213 An interesting example of a bicovalent flavoprotein oxidase is reticuline oxidase, also known as berberine bridge enzyme. This plant enzyme can form a C−C bond upon oxidation of an N-methyl group in reticuline.212

a

Flavin-N(5)-oxide is formed by transferring oxygen from the substrate (S) hydroperoxide to oxidized flavin. R is phosphoribityl for FMN.

mechanism to form the flavin-N(5)-oxide has been proposed for Escherichia coli K12 RutA.197 RutA catalyzes an amide monooxygenation reaction resulting in pyrimidine ring opening in the uracil catabolic pathway.197

5. FLAVIN-DEPENDENT OXIDASES About 80 years ago the first flavoprotein oxidase was isolated from yeast and described as a “new yellow protein”.198 The respective protein was D-amino acid oxidase, and isolation of its yellow cofactor led to the elucidation of the molecular structure of the FAD cofactor. Since the discovery of D-amino acid oxidase, many more flavoprotein oxidases have been discovered and studied in detail.199 Several flavin-dependent oxidases are the focus for fundamental studies in order to understand their physiological role and catalytic mechanism. In fact, many human flavoprotein oxidases fulfill delicate roles in human cells with some serving as important pharmacological targets. Examples are mitochondrial monoamine oxidases, which are involved in the degradation of neurotransmitters, and the nuclear LSD1, which is involved in histone demethylation200,201 Yet, flavin-dependent oxidases are also studied for their potential as biotechnological tools. In fact, the most widely applied redox enzyme is an FAD-containing oxidase: glucose oxidase, which is used in commercially available glucose-monitoring devices.202 Many more oxidases are being applied or developed for their use in various applications ranging from their integration in biosensors to 1757


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Acyl-CoA oxidase represents the prototype oxidase of another well-characterized flavoprotein oxidase family, which employs a tightly bound FAD as redox cofactor. Oxidases of the acyl-CoA oxidase family catalyze the oxidation of fatty acids, with an active site glutamate promoting the deprotonation of the substrate at the Cα atom in order to trigger a hydride transfer from the Cβ to the N(5) of the FAD cofactor.32,214 The sulfhydryl oxidases are instrumental in the formation of disulfide bonds in proteins by oxidizing cysteine residues. Two sulfhydryl oxidase subfamilies have been identified: Erv-like and Ero-like sulfhydryl oxidases. Structural studies revealed a topology of five α-helices which fold around the FAD cofactor.215 Structural studies have shown that these flavoprotein oxidases often feature a dedicated dioxygendiffusion tunnel. Of interest, in rat Erv1 no such tunnel could be identified which is in line with its poor reactivity toward molecular oxygen. The inability of some Erv members to react with molecular oxygen suggests that these homologous flavoenzymes may utilize a different physiological electron acceptor for the oxidation of the reduced flavin.216,217 While all of the above flavoprotein oxidase families harbor a FAD as prosthetic group, the hydroxyacid oxidases (see Table 1) contain an FMN as the tightly bound cofactor. Glycolate oxidase can be regarded as prototype oxidase of this family and has been identified in plants but also in humans.218 Similar to the mechanism of alcohol oxidation by other flavoprotein oxidase types, the mechanism of oxidation of the organic substrate in this family generally involves a hydride transfer to the N(5) of the FMN cofactor.

Figure 9. Active site of glucose oxidase from A. niger (PDB code: 1CF3). Protonation of the His516 side chain dramatically enhances the oxidase activity of the protein. Dioxygen is expected to diffuse near the site occupied by a water (shown as red sphere).

tunneling effects in the flavoenzyme oxygen activation remains an open and debated question.238−243 The role of preorganized electrostatics in dioxygen activation is well illustrated by monomeric sarcosine oxidase.244,245 The N(5) position of FAD in this enzyme is hydrogen bonded to a lysine side chain (Lys265) via a bridging water208 (Figure 10). Mutating Lys265 to a neutral residue

5.1. Mechanisms of Oxygen Activation

5.1.1. Preorganization of the Active Site. Glucose oxidase from Aspergillus niger has been studied for many decades with a variety of techniques that include X-ray crystallography, NMR spectroscopy, electrochemistry, and steady-state and rapid reaction kinetics.202,219−231 With kcat/ KM(O2) values between 5.7 × 102 and 1.5 × 106 M−1 s−1 at high and low pH, respectively, glucose oxidase can be considered as the point of reference for the kinetics of the oxygen reaction in oxidases. It is notable that the rate constant measured at high pH is close to the rate constant of the reaction of free reduced flavins with O2 (250 M−1 s −1).55,232 In-depth and well-conceived mechanistic and mutagenesis investigations demonstrated that protonation of a histidine side chain (His516) is responsible for the enhanced oxygen reaction at low pH by favoring the rate-limiting electron transfer from the reduced flavin to O2 (Figure 9; Schemes 2 and 3).233−235 These data were further comprehensively evaluated in the context of the Marcus theory of electron transfer. The low redox potential of the oxygen−superoxide anion couple (−330 mV) inherently implies that the first electron-transfer step has a weak thermodynamic driving force. Conversely, the reaction is dominated by a relatively large (12−16 kcal mol−1) reorganization energy term (“the energy to transform the configuration of the reactant state to that of the product state without electron transfer”236,237). Such a predominant effect can arise from the electrostatic contribution exerted by the charged groups in the active site including the protonated His516. This analysis additionally hinted at a tunneling behavior of the reaction with dioxygen even if it involves a large oxygen nucleus. The presence and relevance of

Figure 10. Dioxygen activation site in monomeric sarcosine oxidase.71 Hydrogen bonds are indicated by dashed lines (PDB code: 2GB0). Dioxygen is proposed to react with the reduced flavin by binding in the position occupied by the water molecule which is located between Lys265 and the flavin N(5) atom in the oxidized enzyme.

causes a pronounced 8000-fold decrease in dioxygen reactivity, whereas a 250-fold decrease is observed with the Lys265Arg variant enzyme.246 This indicated that the FAD:N(5)-waterLys motif found in nearly all members of the amine oxidase family likely serves as a conserved structural module responsible for oxygen activation.75,247−249 Indeed, mutating the equivalent lysine to a neutral residue in N-methyltryptophan oxidase and fructosamine oxidase decreased the rate constant for the oxygen reaction by ∼1000−2500-fold.248,250 These data together with computational studies led to the conclusion that the bridging water that intercalates between lysine and the flavin N(5) atom effectively occupies the dioxygen site (see Figure 10). The implication is that dioxygen localizes in direct contact with the flavin N(5) at the edge of the cofactor ring, whereas the nearby Lys side chain likely 1758


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Figure 11. Interactions and conformation of the FAD cofactor in the active site of choline oxidase. (Left) Glycine betaine is shown as cyan sticks; FAD is shown as yellow sticks. Side chains of the residues close to glycine betaine are displayed in gold sticks; hydrogen bonds are highlighted with black dashes. (Right) Crystal structure of the Val464Ala mutant (PDB code: 4MJW and 3LJP).

Scheme 16. Reaction Mechanism with Molecular Oxygen in the Oxidative Half-Reaction of Pyranose Oxidase (P2O)53

the H atom from the reduced flavin and a proton from the solvent or a solvent exchangeable site being transferred in the same kinetic step, as demonstrated using multiple deuterium kinetic isotope effects (Scheme 7). Prior to reaction with dioxygen, the reduced enzyme undergoes an isomerization gated by the side chain of Phe357, as established with solvent viscosity effects and mutagenesis.257,258 The work on choline oxidase further demonstrated that other factors, in addition to electrostatics, operate in the reactivity with dioxygen. It was found that replacement of an active-site valine side chain (Val464, see Figure 11) with threonine or alanine causes a 50-fold decrease in the rate constant of the reaction of the reduced enzyme with dioxygen.259,260 Val464 is positioned in the proximity of the flavin and is part of the choline-binding site. Its role in dioxygen activation might be dual. First, it may sterically guide dioxygen toward the site of reactivity in the proximity of the N(5)−C(4a) locus. Second, it may also contribute to the binding of choline and its oxidation product in the position that most effectively allows the trimethylammonium moiety to electrostatically favor the reaction with dioxygen. These findings further add to the concept that each enzyme features specific mechanistic properties though in a context of electrostatically driven catalysis.261 5.1.3. Pyranose 2-Oxidase and Bacterial NADH Oxidase: Deviating from the Canon. Pyranose 2-oxidase catalyzes the oxidation of D-glucose and other aldopyranoses at the C(2) position, yielding 2-ketoaldose and H2O2 as products.262,263 This member of the GMC superfamily has been the subject of a wealth of mechanistic studies.53,264,265 The interest stemmed from the discovery that the reaction of reduced pyranose 2-oxidase with dioxygen involves a stable C(4a)−hydroperoxyflavin intermediate (Scheme 2). The enzyme can therefore be viewed as a system captured on the

provides a preorganized, positively charged environment to promote dioxygen reduction by the flavin.247 However, simply placing a positive charge near the N(5) atom of the flavin does not guarantee increased oxygen reactivity. Recent studies on dihydroorotate dehydrogenase revealed that mutating a flavinN(5) interacting lysine had little influence on the oxygen reaction of this enzyme. Clearly, an appropriate molecular and structural context is required to fully disclose the beneficial effects of electrostatics on dioxygen activation.250 5.1.2. Choline Oxidase: Taking Advantage of a Charged Substrate. Choline oxidase illustrates a further variation on the role of electrostatics in dioxygen activation.39,251−254 The enzyme belongs to the GMC superfamily, and its active site comprises a histidine residue (His466) that is homologous to the His516 of glucose oxidase (Figure 9).205,255,256 Nevertheless, the outcome of mutagenesis studies targeting this active-site histidine in choline oxidase did not conform to the pattern predicted from the previous work on glucose oxidase. Replacement of the histidine drastically decreased the rate constant for the turnover of the enzyme with choline. However, the effect was found to arise from a strongly diminished rate constant of choline oxidation, i.e., the reductive half-reaction (Scheme 1), whereas the reaction of the reduced flavin with oxygen was only minimally affected. Instead, it was found that the rate of flavin reoxidation decreases (80-fold) when a noncharged substrate analogue is used. Choline oxidase functions through a ternary complex mechanism, and the reaction with O2 takes place in the presence of the trimethylammonium-containing product (Scheme 1). Consistently, inspection of the enzyme crystal structure suggests that the bound positively charged trimethylammonium group is positioned to fully exert an electrostatic effect on the reaction of dioxygen with the flavin N(5)−C(4a) locus (Figure 11). Flavin oxidation occurs with 1759


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route converting a monooxygenase into an oxidase (Scheme 16).53,265 Remarkably, full conversion to a classic oxidase mechanism could be simply achieved by mutating Thr169, an active-site residue that interacts with the flavin N(5) atom (Figure 12). Thr169 mutants displayed high activity toward

Heme- and cytochrome-deficient bacteria (e.g., enterococci and streptococci) are equipped with a family of NADH oxidases which rely on the C(4a)−peroxyflavin to catalyze the four-electron reduction of O2 to water (Scheme 17). The redox center of these enzymes is formed by the juxtaposition of the flavin with the thiol group of a redox-active cysteine.268 The combination of these catalytic elements entails a fourelectron-reduced state which forms through the successive binding of two NADH molecules. In this FADH−:Cys-S− state, the enzyme reacts with dioxygen to generate the C(4a)− peroxyflavin, whose distal oxygen atom is then efficiently transferred to the adjacent catalytic cysteine. This remarkable “intra-enzymatic” oxygenation step generates two groups, C(4a)−hydroxyflavin and Cys-SOH, which both release a water molecule.269,270 Thus, by combining the flavin with a reactive thiol, this unique family of oxidases generates water rather than hydrogen peroxide as the end product of the reaction.

6. SELF-SACRIFICING FLAVIN At the end of this analysis, the bluB enzyme deserves to be discussed as a most unusual case of dioxygen reactivity. This bacterial flavoprotein synthesizes 5,6-dimethylbenzimidazole, a ligand of the cobalt atom of cobalamin (vitamin B12). Structural studies showed that this enzyme belongs to the well-known family of the NAD(P)H-flavin oxidoreductases.271 Despite this standard folding topology, the enzyme performs a unique reaction which uses FMN and dioxygen as substrates. The enzyme binds the two-electron-reduced flavin generated by a reductase. Next, the reduced flavin reacts with dioxygen to form the C(4a)−peroxyflavin. Differently, from the flavoprotein oxidases and monooxygenases, the intermediate undergoes a spectacular enzyme-catalyzed fragmentation that eventually generates 5,6-dimethylbenzimidazole as product together with D-erythrose 4-phosphate (Scheme 18). The mechanism underlying such a complex reaction remains elusive. Recent computational studies propose that the reaction may proceed through a C(4a)−C(10a)-peroxide, but this idea awaits experimental validation.272 The future will tell whether peroxyflavins are used as chemical precursors in other biosynthetic pathways or whether bluB will rather remain a sort of an exotic and unique case.

Figure 12. Active site of pyranose oxidase (PDB code: 1TT0). Dioxygen molecule was modeled in the active site in an orientation ready to attack the C(4a) atom of the flavin.

dioxygen but without a detectable formation of any intermediate.266,267 These mutagenesis data were combined with isotope-labeling experiments and computational studies.53,110 These comprehensive approaches support the view that the initial single electron transfer is coupled to a proton transfer that generates a diradical •OOH (Scheme 7). This species further reacts with the flavin to form the C(4a)− hydroperoxyflavin, whose oxygen atoms interact with a cluster of hydrogen-bonding side chains present in the active site (Thr169, His548, and Asn593). In agreement with the mutagenesis data, a key factor for C(4a)−hydroperoxyflavin stabilization was shown to be the interaction between Thr169 and the protonated N(5) atom of the flavin since the release of hydrogen peroxide from the intermediate requires the breakage of the N(5)−H bond (Schemes 7 and 16).46 It is fascinating to see how choline oxidase, pyranose 2-oxidase, and glucose oxidase, which belong to the same GMC superfamily of flavoenzymes with similar fold and the same type of chemistry for the oxidation of the organic substrate, exhibit substantial differences in the fine-tuned details of their dioxygen enzymology, thereby perfectly epitomizing the versatility of the flavin.265

7. CONCLUSIONS AND OUTLOOK Though in the context of amazing functional and mechanistic diversity, our survey indicates that flavoprotein oxidases generally (but not always) react with oxygen through an outer-sphere mechanism that does not involve a stable covalent intermediate but rather the stepwise transfer of two electrons

Scheme 17. Summary of the Overall Reaction Catalyzed by Bacterial Water-Forming NADH Oxidases Together with a Scheme for the Critical Reaction Step Leading to C(4a)−Hydroxyflavin and Cys-Sulphenic Acid (Cys-SOH) Formation

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Scheme 18. Reaction Catalyzed by bluBa

a

Reduced FMN reacts with dioxygen to form C(4a)−peroxyflavin that fragments into 5,6-dimethylbenzimidazole and D-erythrose 4-phosphate, possibly through a C(4a)−C(10a) adduct. R is phosphoribityl of the FMN.272

from the reduced flavin to oxygen. The three-dimensional structures show that this reaction often involves the binding of dioxygen at the edge of the flavin ring in direct contact with the N(5)−C(4a) atoms of the prosthetic group. The geometry of the active site of sarcosine oxidase perfectly recapitulates these notions (Figure 10). Conversely, monooxygenases activate oxygen through a covalent C(4a)−(hydro)peroxyflavin intermediate that inserts an oxygen atom into the organic substrate. Formation of the C(4a)−peroxyflavin poses more stringent stereochemical and geometrical requirements for oxygen binding compared to the oxidases. Indeed, monooxygenases typically feature small oxygen-binding cavities precisely located above the flavin ring and in direct contact with the flavin C(4a) atom. The active site of 4-hydroxyphenylacetate-3-hydroxylase nicely outlines these features (Figure 7). However, such classic paradigmmonooxygenases as the “peroxyflavin enzymes” and oxidases as the “no-intermediate enzymes”must be very substantially revised and expanded. The very exciting discovery that flavoenzymes can react with oxygen by forming a N(5)−O adduct has literally transformed this way of thinking. Moreover, the never-ending discovery of new flavoenzymes in all kingdoms of life continuously reveals unexpected reactions as documented by the astonishing diversity in the substrate specificities and reactivities discussed in this review. These advances raise several fascinating questions and pose fundamental challenges that remain to be conquered. (1) It remains to be established whether the newly discovered enzymes (e.g., EncM) that operate though a flavin-N(5)-oxide intermediate are the very first examples of a large and so far neglected flavoenzyme class. The actual catalytic potential of the N(5)-oxide remains mostly unexplored, and the hunting season for N(5)-oxide enzymes is obviously open. (2) An open challenge is to rationalize the elements that determine how flavoenzymes operate with regard to the outcome of their reaction with dioxygen. What are the structural bases for the selectivity of adduct formation? Why is dioxygen attacking the C(4a) atom rather than the N(5) atom depending on the flavoenzymes? Of note, flavoenzymes can share the same folding topology and similar active site and yet feature distinct modes of operation. (3) Monooxygenases can substantially differ for their ability to stabilize the C(4a)−(hydro)peroxyflavin intermediate that, depending on the enzyme, can last for minutes or be hardly detectable even in time-resolved stopped-flow kinetic experiments. The structural basis for such diversity remains poorly understood. (4) The initial one-electron transfer from the reduced flavin to dioxygen yielding a transient superoxide is the initial

reaction step that is common to oxidases and monooxygenases and can be enhanced by positively charged groups (Schemes 2 and 3). A key mechanistic question that remains to be fully investigated is whether this step is generally coupled to dioxygen protonation as suggested by recent computational studies (see Scheme 7). Is proton-coupled electron transfer a mechanistic feature shared by most oxygen-reacting flavoenzymes? (5) Why do certain flavoenzymes react so poorly with dioxygen (e.g., flavocytochrome b2,273 D-arginine dehydrogenase,34 and cellobiose dehydrogenase274)? Residues locally gating access to flavin N(5)−C(4a) locus might control the accessibility and, therefore, reactivity of the flavin with dioxygen, although other unidentified factors may contribute to the lack of reactivity in some instances.275 Furthermore, finely tuned electrostatics clearly play a role in dioxygen activation.35,231 However, it remains to be fully understood how these factors cooperate in the control and regulation of the oxygen reaction. In this context, the enzymes that use F420 (an oxygen-unreactive 5-deazaflavin cofactor) represent an unexplored and fertile territory for flavoenzyme discovery.44 (6) The goal of rationally creating tailor-made enzymes that can be used for biotechnological applications remains farfetched. The current level of knowledge on enzyme mechanisms has occasionally enabled efficient enzyme redesign. Success stories have been the conversion of BVMO and dehydrogenase enzymes into oxidases.145,276 Can a monooxygenase enzyme be engineered from an oxidase or a dehydrogenase? Can a true dehydrogenase be used toward this aim? These seem to be the next required steps that must be taken toward enabling future efficient enzyme redesign. The challenge here is way more complex because it requires one not only to fully understand and solve the problem of dioxygen accessing and reacting with the flavin but also to resolve the issue of how to stabilize the reaction intermediates required by the monooxygenases. A full and detailed understanding of the chemical and structural basis underpinning the distinct reactivities of flavoenzymes will be needed to transform engineering of oxygen-reacting enzymes into a fully rational and predictable exercise. The diversity and complexity of the reactions carried out by oxygen-reacting flavoenzymes is breathtaking and will never cease to fascinate chemists and biochemists as highlighted by the recent discovery of prenylated flavins.277 It is truly amazing that after 80 years of flavoenzymology new and unexpected reactions are being discovered. 1761


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AUTHOR INFORMATION

Pavia (Italy). There he started his studies on the role and mechanism of covalent flavinylation. In 1999, he became Assistant Professor in the Biotechnology group of Professor Dr. Dick B. Janssen at the University of Groningen (The Netherlands), where he extended his enzymology research by studying flavoenzymes in the context of biocatalysis. He is now leading his own Molecular Enzymology research group at the same university, and his research focuses on the discovery, engineering, and application of redox enzymes.

Corresponding Authors

*E-mail: [email protected]. *E-mail: [email protected]. *E-mail: [email protected]. ORCID

Giovanni Gadda: 0000-0002-7508-4195 Marco W. Fraaije: 0000-0001-6346-5014 Andrea Mattevi: 0000-0002-9523-7128

Andrea Mattevi is Professor and Member of the Structural Biology Program at the University of Pavia, Italy. He conducted his Ph.D. work with Professor W. G. Hol at the University of Groningen, The Netherlands, and, as EMBO long-term fellow, his postdoctoral research with Dr. J. Walker and A. W. Leslie at the MRC Laboratory of Molecular Biology, Cambridge, UK. His laboratory studies structural and molecular mechanisms of enzymes that use oxygen to either produce ROS in signaling networks, post-translationally modify histones, or transform metabolites and xenobiotics.

Notes

The authors declare no competing financial interest. Biographies Elvira Romero obtained her Ph.D. degree at the Center of Biological Research of the Spanish National Research Council (Spain), where she investigated fungal oxidoreductases involved in lignin decay under the supervision of Professor Marı ́a Jesús Martı ́nez. Her postdoctoral research in Professor Pablo Sobrado and Giovanni Gadda’s laboratories, at Virginia Tech and Georgia State Universities (USA), respectively, were mainly focused on elucidating the catalytic mechanism of flavin-dependent oxidoreductases with medical relevance. Currently, she is learning enzyme engineering and biotechnology by working as a Postdoctoral Associate in the Molecular Enzymology research group of Professor Marco W. Fraaije at Groningen University (The Netherlands).

ACKNOWLEDGMENTS Part of this work was supported by the EU project ROBOX (grant agreement no. 635734) under the EU’s Horizon 2020 Programme Research and Innovation actions H2020-LEIT BIO-2014-1 and a National Science Foundation grant CHE1506518. ABBREVIATIONS FMN flavin mononucleotide FAD flavin adenine dinucleotide PHBH 4-hydroxybenzoate 3-hydroxylase FMO flavin-containing monooxygenases BVMO Baeyer−Villiger monooxygenases GMC glucose-methanol-choline VAO vanillyl−alcohol oxidase

J. Rubén Gómez Castellanos is a postdoctoral fellow in the Group of Professor Andrea Mattevi at the University of Pavia working in the Structural Biology of oxidative biocatalysts. He completed his Ph.D. degree in Chemical Biology at the University of Oxford under the supervision of Professor Chris Schofield investigating the biosynthesis of carbapenems. He holds an M.Sc. degree in Drug Discovery from The School of Pharmacy of the University of London (2006) and a B.Sc. degree in Pharmaceutical and Biological Chemistry from La Salle University in Mexico City (2005). Before returning to academia, Rubén spent time working for pharmaceutical companies Eli Lilly and Company and Servier in clinical research in Mexico and Latin America.

REFERENCES (1) Ghisla, S.; Massey, V. Mechanisms of Flavoprotein-Catalyzed Reactions. Eur. J. Biochem. 1989, 181, 1−17. (2) Mattevi, A. To Be or Not to Be an Oxidase: Challenging the Oxygen Reactivity of Flavoenzymes. Trends Biochem. Sci. 2006, 31, 276−283. (3) Malmström, B. G. Enzymology of Oxygen. Annu. Rev. Biochem. 1982, 51, 21−59. (4) Palfey, B. A.; Ballou, D. P.; Massey, V. Active Oxygen in Biochemistry; Springer Netherlands: Dordrecht, 1995; pp 37−83. (5) Solomon, E. I.; Heppner, D. E.; Johnston, E. M.; Ginsbach, J. W.; Cirera, J.; Qayyum, M.; Kieber-Emmons, M. T.; Kjaergaard, C. H.; Hadt, R. G.; Tian, L. Copper Active Sites in Biology. Chem. Rev. 2014, 114, 3659−3853. (6) Poulos, T. L. Heme Enzyme Structure and Function. Chem. Rev. 2014, 114, 3919−3962. (7) Spencer, R.; Fisher, J.; Walsh, C. Preparation, Characterization, and Chemical Properties of the Flavin Coenzyme Analogues 5Deazariboflavin, 5-Deazariboflavin 5′-Phosphate, and 5-Deazariboflavin 5′-Diphosphate, 5′ → 5′-Adenosine Ester. Biochemistry 1976, 15, 1043−1053. (8) Manstein, D. J.; Pai, E. F. Purification and Characterization of FAD Synthetase from Brevibacterium Ammoniagenes. J. Biol. Chem. 1986, 261, 16169−16173. (9) Karthikeyan, S.; Zhou, Q.; Mseeh, F.; Grishin, N. V.; Osterman, A. L.; Zhang, H. Crystal Structure of Human Riboflavin Kinase Reveals a β Barrel Fold and a Novel Active Site Arch. Structure 2003, 11, 265−273. (10) Hefti, M. H.; Vervoort, J.; van Berkel, W. J. H. Deflavination and Reconstitution of Flavoproteins. Eur. J. Biochem. 2003, 270, 4227−4242.

Giovanni Gadda is a Distinguished University Professor of Chemistry at Georgia State University, with a dual appointment in the Departments of Chemistry and Biology. He is also a member of the Center for Diagnostics and Therapeutics and the Center for Biotechnology and Drug Design at GSU. He earned his B.Sc. degree in Biological Sciences and Ph.D. degree in Biochemistry at the Università degli Studi di Milano, Italy. He conducted postdoctoral research as an EMBO Fellow recipient at Konstanz Universität in Germany and at Texas A&M University, College Station, TX. His research interests are on the mechanistic enzymology of flavindependent oxidases, monooxygenases, dehydrogenases, and reductases. The long-term objectives are to understand how enzymes can influence the energetics of reaction intermediates and transition states and how the interactions of the flavin cofactor with the protein and substrate modulate flavin versatility and reactivity to an extent that the unproductive reactions are abated in favor of the specific reaction that needs to be catalyzed. Since 2010, he has been the organizer of the Annual Southeast Enzyme Conference in Atlanta, GA. Marco W. Fraaije obtained his Ph.D. degree in Biochemistry from Wageningen University (The Netherlands) in 1998, supervised by Professor Willem van Berkel. After that he obtained a long-term EMBO fellowship and worked as a postdoctoral fellow in the crystallography group of Professor Andrea Mattevi at the University of 1762


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