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Electron flow through biological molecules: does hole hopping protect proteins from oxidative damage?

Published online by Cambridge University Press:  16 July 2015

Jay R. Winkler  and
Harry B. Gray
Jay R. Winkler
Affiliation:
Beckman Institute, California Institute of Technology, Pasadena, CA 91125, USA
Harry B. Gray*
Affiliation:
Beckman Institute, California Institute of Technology, Pasadena, CA 91125, USA
*
* Author for correspondence: Beckman Institute, MC 139-74, California Institute of Technology, 1200 E. California Boulevard, Pasadena, CA 91125, USA. Tel: +626-395-6500; Fax: +626-449-4159; Email: [email protected]
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Abstract

Biological electron transfers often occur between metal-containing cofactors that are separated by very large molecular distances. Employing photosensitizer-modified iron and copper proteins, we have shown that single-step electron tunneling can occur on nanosecond to microsecond timescales at distances between 15 and 20 Å. We also have shown that charge transport can occur over even longer distances by hole hopping (multistep tunneling) through intervening tyrosines and tryptophans. In this perspective, we advance the hypothesis that such hole hopping through Tyr/Trp chains could protect oxygenase, dioxygenase, and peroxidase enzymes from oxidative damage. In support of this view, by examining the structures of P450 (CYP102A) and 2OG-Fe (TauD) enzymes, we have identified candidate Tyr/Trp chains that could transfer holes from uncoupled high-potential intermediates to reductants in contact with protein surface sites.

Keywords

electron transferprotein radicalhole hoppingazurincytochrome P450
Type
Perspective
Information
Quarterly Reviews of Biophysics , Volume 48 , Issue 4 , November 2015 , pp. 411 - 420
Creative Commons
Creative Common License - CCCreative Common License - BY
This is an Open Access article, distributed under the terms of the Creative Commons Attribution licence (http://creativecommons.org/licenses/by/3.0/), which permits unrestricted re-use, distribution, and reproduction in any medium, provided the original work is properly cited.
Copyright
Copyright © Cambridge University Press 2015

Background

Many vital biological transformations involve the incorporation of one (monooxygenases) or two (dioxygenases) O-atoms from molecular oxygen into organic substrates. Enzymes that utilize oxygen must coordinate the delivery of four protons and four electrons to O2 in order to prevent the formation of harmful molecular oxidants (O2 −, HO2 • H2O2, and HO•), collectively known as reactive oxygen species (ROS). It is our view that the risks posed by reactive intermediates are so great that oxygen-utilizing enzymes have protection mechanisms to help them avoid inactivation when the primary electron/proton transfer mechanism is disrupted.

The mechanism of O2 reduction by cytochrome c oxidase illustrates some of the challenges facing these enzymes (Wikström, Reference Wikström2012; Yu et al. Reference Yu, Egawa, Shinzawa-Itoh, Yoshikawa, Yeh, Rousseau and Gerfen2011, Reference Yu, Egawa, Shinzawa-Itoh, Yoshikawa, Guallar, Yeh, Rousseau and Gerfen2012). Reaction of the fully four-electron reduced enzyme (CuA II,I, FeII-heme a, FeII-heme a 3, and CuB I) with O2 generates an intermediate designated as PR. When the two-electron reduced, mixed valence enzyme (CuA II,II, FeIII-heme a, FeII-heme a 3, and CuB I) reacts with O2, the PM intermediate is formed. The O–O bond has been cleaved in both PR and PM to produce FeIV(O)-heme a 3 and CuB II in the binuclear site. The difference between PR and PM is in the source of the fourth electron: PM is thought to have a Tyr244 radical (bovine numbering), whereas the fourth electron in PR is provided by FeII-heme a. When PM is prepared using H2O2, the hole on (TyrO•)244 is believed to migrate through (Trp•+)236 to (TyrO•)129; the latter residue is suggested to participate in proton pumping (Yu et al. Reference Yu, Egawa, Shinzawa-Itoh, Yoshikawa, Guallar, Yeh, Rousseau and Gerfen2012). The key point is that Tyr244 is available to fill the gap when the fourth electron required for O2 reduction cannot be supplied by FeII-heme a (Wikström, Reference Wikström2012; Yu et al. Reference Yu, Egawa, Shinzawa-Itoh, Yoshikawa, Guallar, Yeh, Rousseau and Gerfen2012).

In many oxygenases, including the cytochromes P450 (P450) and the 2-oxo-glutarate-dependent nonheme iron oxygenases (2OG-Fe), the four electrons required for O2 reduction have different origins (Fig. 1). Typically, two electrons are delivered from a reductase (P450) or co-substrate (2OG), and the remaining two electrons are provided by the organic substrate (Denisov et al. Reference Denisov, Makris, Sligar and Schlichting2005; Hausinger, Reference Hausinger2004; Whitehouse et al. Reference Whitehouse, Bell and Wong2012). In the consensus mechanism for iron oxygenases, the first two electrons induce O–O bond cleavage, producing a powerfully oxidizing ferryl species. The ferryl complex abstracts a hydrogen atom from the substrate and HO• rebound leads to hydroxylated product (Denisov et al. Reference Denisov, Makris, Sligar and Schlichting2005; Hausinger, Reference Hausinger2004; Whitehouse et al. Reference Whitehouse, Bell and Wong2012). For enzymes with broad substrate specificities, or when operating in the presence of xenobiotic compounds, the fidelity of substrate oxidation is less than 100%, with potentially damaging consequences (Chen et al. Reference Chen, Comeaux, Herbst, Saban, Kennedy, Maroney and Knapp2008; De Matteis et al. Reference De Matteis, Ballou, Coon, Estabrook and Haines2012; Denisov et al. Reference Denisov, Baas, Grinkova and Sligar2007a ; Grinkova et al. Reference Grinkova, Denisov, McLean and Sligar2013; Saban et al. Reference Saban, Flagg and Knapp2011; Staudt et al. Reference Staudt, Lichtenb and Ullrich1974). This circumstance is manifested as an increased molar ratio of O2 consumption to substrate hydroxylation (uncoupling). We think it likely that organisms have evolved protection mechanisms to guard against deactivation of oxygenase enzymes in the event of uncoupled O2 consumption. In particular, we suggest that radical transfer pathways are employed to deliver strongly oxidizing holes (E°~1 V versus NHE) from ferryl complexes in active sites to less fragile regions of oxygenases.

Fig. 1. Schematic representation of the catalytic mechanisms of P450 and 2OG-Fe oxygenases: RH, substrate; 2OG, 2-oxoglutarate; Suc, succinate. Black arrows indicate the functional substrate hydroxylation pathways. Blue arrows indicate oxidase uncoupling pathways.

In this perspective, we will advance the hypothesis that there are potentially protective radical chains in P450 and 2OG-Fe; but first we will review what we know about the factors controlling hopping through aromatic amino acids in multistep electron tunneling constructs designed in azurin, a prototypal cupredoxin.

Radical transfer pathways in azurin

Azurin is a robust cupredoxin (128 residues) that is amenable to site-directed mutagenesis and surface-labeling with photosensitizers (Farver & Pecht, Reference Farver and Pecht2011; Gray & Winkler, Reference Gray and Winkler2010; Reece & Nocera, Reference Reece and Nocera2009; Wilson et al. Reference Wilson, Yu and Lu2013). Oxidized radicals of Trp and Tyr are substantially stronger acids than their neutral precursors (Trp, pK a > 14; Trp•+, pK a = 4; TyrOH, pK a = 10; TyrOH•+, pK a = −1) (Aubert et al. Reference Aubert, Vos, Mathis, Eker and Brettel2000; Bonin et al. Reference Bonin, Costentin, Louault, Robert, Routier and Saveant2010; Costentin et al. Reference Costentin, Louault, Robert and Saveant2009; Harriman, Reference Harriman1987; Jovanic et al. Reference Jovanic, Harriman and Simic1986); management of the acidic proton is a critically important factor controlling radical formation with these amino acids. Proton management is particularly challenging for buried amino acids and, thus far, we have not succeeded in detecting buried Trp or Tyr radicals as electron transfer (ET) intermediates. Our kinetics data indicate that surface exposed Trp•+ and NO2TyrO• radicals can, in appropriate constructs, accelerate CuI oxidation by distant Re- and Ru-diimine complexes (Shih et al. Reference Shih, Museth, Abrahamsson, Blanco-Rodriguez, Di Bilio, Sudhamsu, Crane, Ronayne, Towrie, Vlcek, Richards, Winkler and Gray2008; Warren et al. Reference Warren, Herrera, Hill, Winkler and Gray2013a ).

Multistep ET through Trp and Tyr radicals in azurin

We have used Pseudomonas aeruginosa azurin as a test bed for mechanistic investigations of Trp and Tyr radical formation in protein ET reactions (Blanco-Rodriguez et al. Reference Blanco-Rodriguez, Di Bilio, Shih, Museth, Clark, Towrie, Cannizzo, Sudhamsu, Crane, Sykora, Winkler, Gray, Zalis and Vlcek2011; Shih et al. Reference Shih, Museth, Abrahamsson, Blanco-Rodriguez, Di Bilio, Sudhamsu, Crane, Ronayne, Towrie, Vlcek, Richards, Winkler and Gray2008; Takematsu et al. Reference Takematsu, Williamson, Blanco-Rodríguez, Sokolová, Nikolovski, Kaiser, Towrie, Clark, Vlček, Winkler and Gray2013; Warren et al. Reference Warren, Ener, Vlček, Winkler and Gray2012, Reference Warren, Herrera, Hill, Winkler and Gray2013a ). Our initial investigation revealed that CuI oxidation by a photoexcited ReI–diimine complex (ReI(CO)3(4,7-dimethyl-1,10-phenanthroline)) covalently bound at His124 on a His124Gly123Trp122Met121 β-strand (ReHis124Trp122CuI-azurin) occurs in a few nanoseconds, fully two orders of magnitude faster than documented for single-step electron tunneling at a 19-Å donor–acceptor distance, owing to a two-step hopping mechanism involving a Trp•+ radical intermediate (Shih et al. Reference Shih, Museth, Abrahamsson, Blanco-Rodriguez, Di Bilio, Sudhamsu, Crane, Ronayne, Towrie, Vlcek, Richards, Winkler and Gray2008).

Our work on multistep ET in sensitizer-modified azurin is informed by semiclassical ET theory (Marcus & Sutin, Reference Marcus and Sutin1985). Given a particular spatial arrangement of redox cofactors, we can predict driving-force dependences of the relative time constants for single-step (τ ss = 1/k ss) and multistep (τ hop) electron transport (Warren et al. Reference Warren, Ener, Vlček, Winkler and Gray2012). Alternatively, given the redox and reorganization energetics, we can predict the hopping propensity for different cofactor arrangements (Warren et al. Reference Warren, Herrera, Hill, Winkler and Gray2013a ). We considered three Ru(2,2′-bipyridine)2(imidazole)(HisX)-labeled azurins (RuHis107, RuHis124, and RuHis126) and examined the hopping advantage (τ ss/τ hop) for a protein with a generalized intermediate (Int) situated between a diimine-RuIII oxidant and CuI (Warren et al. Reference Warren, Herrera, Hill, Winkler and Gray2013a ). In all cases, the greatest hopping advantage occurs in systems where the Int–RuIII distance is up to 5 Å shorter than the Int–CuI distance. The hopping advantage increases as systems orient nearer a linear Donor–Int–Acceptor configuration, owing to minimized intermediate tunneling distances. The smallest predicted hopping advantage is in RuHis124 azurin, which has the shortest Ru–Cu distance of the three proteins. The hopping advantage is nearly lost as ΔG° for the first step (RuIII ← Int) rises above +0.15 eV. Isoergic initial steps provide a wide distribution of arrangements, where advantages as great as 104 are possible (for a fixed donor–acceptor distance of 23.7 or 25.4 Å). A slightly exergonic Int → RuIII step provides an even larger distribution of arrangements for productive hopping, which will be the case as long as the driving force for the first step is not more favorable than that for overall transfer.

We tested these predictions experimentally in three Ru–His-labeled azurins using nitrotyrosinate (NO2TyrO–) as a redox intermediate (RuHis107(NO2TyrOH)109; RuHis124(NO2TyrOH)122; and RuHis126−(NO2TyrOH)122; E°′(NO2TyrO•/–) ≈ 1.02 V versus NHE) (Fig. 2) (Warren et al. Reference Warren, Herrera, Hill, Winkler and Gray2013a ). The first two systems have cofactor placements that are close to the predicted optimum; the last system has a larger first-step distance, which is predicted to decrease the hopping advantage. The phenol pK a of 3-nitrotyrosine (7.2) permitted us to work at near-neutral pH, rather than high pH (>10) required for hopping with tyrosinate. ET via nitrotyrosinate avoids the complexities associated with the proton-coupled redox reactions of tyrosine. We found specific rates of CuI oxidation more than 10 times greater than those of single-step ET in the corresponding azurins lacking NO2TyrOH, confirming that NO2TyrO– accelerates long-range ET. The results are in excellent agreement with hopping maps developed using semiclassical ET theory and parameters derived from our body of protein ET measurements (Gray & Winkler, Reference Gray and Winkler2010; Warren et al. Reference Warren, Ener, Vlček, Winkler and Gray2012, Reference Warren, Herrera, Hill, Winkler and Gray2013a ).

Fig. 2. (a) Space-filling structural model of RuHis107NO2TyrOH109Cu-azurin. (b) Space filling models of the residues comprising the hole-hopping pathway from Cu to RuHis107.

Potential radical transfer pathways in iron oxygenases

The cytochromes P450 are members of a superfamily of heme oxygenases that perform two broad functional roles: xenobiotic metabolism and biosynthesis (Denisov et al. Reference Denisov, Makris, Sligar and Schlichting2005; Johnson & Stout, Reference Johnson and Stout2013; Nebert et al. Reference Nebert, Wikvall and Miller2013; Orr et al. Reference Orr, Ripp, Ballard, Henderson, Scott, Obach, Sun and Kalgutkar2012; Whitehouse et al. Reference Whitehouse, Bell and Wong2012). The oxygenation chemistry catalyzed by some P450 enzymes is tightly coupled to substrate hydroxylation: one mole of product is produced for each mole of O2 consumed. In many enzymes, however, particularly the eukaryotic proteins with broad substrate specificities, hydroxylation is much less efficiently coupled to O2 consumption (frequently less than 10%) (Denisov et al. Reference Denisov, Baas, Grinkova and Sligar2007a ; Grinkova et al. Reference Grinkova, Denisov, McLean and Sligar2013; Staudt et al. Reference Staudt, Lichtenb and Ullrich1974). When the enzyme does not transfer an O-atom to substrate, it can produce ROS (O2 −, H2O2) or a second H2O molecule (Puntarulo & Cederbaum, Reference Puntarulo and Cederbaum1998). The production of ROS can lead to rapid degradation of the enzyme and other harmful chemistry. In the case of oxidase chemistry (formation of 2H2O from O2), two reducing equivalents must be delivered by sources other than the substrate. When a CYP enzyme binds a refractory substrate, ferryl formation is likely to proceed, but substrate hydroxylation is inhibited. Under these circumstances, chains of redox-active Tyr, Trp, Cys, and/or Met residues can direct the oxidizing hole to the protein periphery where it can react with intracellular antioxidants such as glutathione.

Enzymes from the 2OG-Fe superfamily use 2-oxoglutarate as a 2-electron donating co-substrate, Fe2+ as a cofactor, and O2 to effect the hydroxylation of organic substrates (Fig. 1). The 2OG-Fe enzymes exhibit a wide array of biological functions including collagen biosynthesis, lysyl hydroxylation of RNA splicing proteins, DNA repair, RNA modification, chromatin regulation, epidermal growth factor-like domain modification, hypoxia sensing, and fatty acid metabolism (Mantri et al. Reference Mantri, Zhang, McDonough and Schofield2012; Rose et al. Reference Rose, McDonough, King, Kawamura and Schofield2011). The 2OG-Fe oxygenase enzymes have conserved double-stranded β-helix folds with octahedral Fe-binding sites with the HXD/E…H triad providing two His imidazole ligands and one monodentate carboxylate ligand. The remaining three coordination sites in the resting enzyme are occupied by O-donors from 2OG and a water ligand.

Several 2OG-Fe enzymes have been reported to undergo autocatalyzed oxidative modifications of aromatic amino acids. In the taurine-2OG dioxygenase that catalyzes the conversion of taurine to bisulfite, EPR data indicate the transient formation of a Tyr73-based radical that converts to an FeIII-catecholate (Mantri et al. Reference Mantri, Zhang, McDonough and Schofield2012). In 2,4-dichlorophenoxyacetate oxygenase (TfdA) and factor-inhibiting hypoxia-inducible factor (FIH) there is evidence for Trp hydroxylation when substrate is unavailable (Mantri et al. Reference Mantri, Zhang, McDonough and Schofield2012). These aromatic amino acid oxidations lead to inactivation of the enzyme. As with P450, we suggest that radical chains of Trp, Tyr, Cys, and/or Met residues in 2OG-Fe hydroxylases protect the enzymes from damage in the event of slow or unsuccessful substrate hydroxylation by diverting the powerfully oxidizing hole from FeIV(O) to the protein surface, where it can react with intracellular reductants (e.g. glutathione). This diversion of oxidizing equivalents would extend the functional lifetime of an enzyme.

When considering the many remarkable transformations catalyzed by natural enzymes, it is easy to be left with the impression that these macromolecules are perfect catalysts that, after millions of years of tinkering, have solved the riddle of simultaneously maximizing speed, selectivity, and specificity. Upon closer inspection, however, heme and nonheme oxygenases are far from perfect catalysts, yet manage to accomplish their primary functions. Indeed, in many oxygenases, the coupling between oxygen consumption and substrate hydroxylation is extremely low. The most abundant P450 in human liver, CYP3A4, is a case in point (Denisov et al. Reference Denisov, Grinkova, McLean and Sligar2007b ; Grinkova et al. Reference Grinkova, Denisov, McLean and Sligar2013). For enzyme incorporated into nanodiscs (Grinkova et al. Reference Grinkova, Denisov and Sligar2010), the coupling of substrate hydroxylation to NADH consumption was ≤16% for testosterone as a substrate, ≤10% for bromocriptine, and 2% for tamoxifen (Grinkova et al. Reference Grinkova, Denisov, McLean and Sligar2013). It is fair to say that, although the primary CYP3A4 function may be substrate hydroxylation, the primary enzyme activity is distributed more or less equally between H2O2 and H2O production (Grinkova et al. Reference Grinkova, Denisov, McLean and Sligar2013). Indeed, it would not be inaccurate to characterize CYP3A4 as a flawed oxidase that occasionally oxygenates organic substrates. More importantly, unless the enzyme was protected from damage in the event of uncoupled turnover, CYP3A4 would function not as a catalyst but as a stoichiometric reagent. A similar situation exists for uncoupled turnover in the 2OG-Fe enzymes.

The active sites of heme and nonheme oxygenases often are deeply buried within a polypeptide matrix. Consequently, powerfully oxidizing active site holes cannot efficiently migrate in single-step tunneling reactions to the enzyme surface for reduction by external reagents (Winkler & Gray, Reference Winkler and Gray2014a , Reference Winkler and Gray b ). We have shown that multistep tunneling reactions can be hundreds to thousands of times faster than their single-step counterparts (Shih et al. Reference Shih, Museth, Abrahamsson, Blanco-Rodriguez, Di Bilio, Sudhamsu, Crane, Ronayne, Towrie, Vlcek, Richards, Winkler and Gray2008; Warren et al. Reference Warren, Ener, Vlček, Winkler and Gray2012, Reference Warren, Herrera, Hill, Winkler and Gray2013a , Reference Warren, Winkler and Gray b ). Radical transfer pathways composed of Tyr, Trp, Cys, and Met residues are ideally suited to deliver active-site oxygenase holes to enzyme surfaces when reaction with substrate is disrupted.

A biologically useful Fe-oxygenase protection mechanism requires that a fine balance be struck between substrate reaction and hole migration to the surface. Overly efficient hole migration would lower enzyme hydroxylation activity, while a sluggish pathway would be ineffective at protecting the enzyme. Active-site hole scavenging in P450 by the natural reductase may be possible, but the timing of this reaction would be extremely variable, owing to fluctuations in reductase concentration. In the 2OG-Fe enzymes, there is no reductase that could protect the enzyme. An intraprotein radical transfer mechanism can be tuned to provide the proper balance between enzyme protection and substrate reaction. We suggest that the first step in the hole-migration pathway is the critical determinant of ferryl survival time. Once a radical forms on the first residue in the pathway (the gateway residue), further migration to the surface is rapid. In the potential pathways that we have identified, the distance from the active site to the first pathway residue is often longer than subsequent steps. In addition to the longer distance, proton coupling and enzyme conformational changes could contribute to limiting the rate of the first step in the transfer chain.

CYP102A1

CYP102A1 from Bacillus megaterium (also known as P450 BM3) is a rare example of a bacterial Class II cytochrome P450 enzyme in which both reductase and heme domains are contained within a single polypeptide chain (Miura & Fulco, Reference Miura and Fulco1974; Narhi & Fulco, Reference Narhi and Fulco1986). The enzyme catalyzes the remarkably rapid hydroxylation of long-chain fatty acids using NAD(P)H and O2, without the presence of any other proteins or cofactors (Narhi & Fulco, Reference Narhi and Fulco1986). The full-length enzyme (CYP102A1HR) has been expressed in Escherichia coli, as have independent heme (CYP102A1H) and reductase (CYP102A1R) domains (Boddupalli et al. Reference Boddupalli, Estabrook and Peterson1990, Reference Boddupalli, Oster, Estabrook and Peterson1992; Li et al. Reference Li, Darwish and Poulos1991a ; Narhi et al. Reference Narhi, Wen and Fulco1988; Oster et al. Reference Oster, Boddupalli and Peterson1991). The individual domains, as well as an assembly between the heme domain and a flavin-containing reductase domain, have been structurally characterized (Girvan et al. Reference Girvan, Seward, Toogood, Cheesman, Leys and Munro2007; Sevrioukova et al. Reference Sevrioukova, Immoos, Poulos and Farmer2000; Warman et al. Reference Warman, Roitel, Neeli, Girvan, Seward, Murray, McLean, Joyce, Toogood, Holt, Leys, Scrutton and Munro2005). The soluble, 119 kDa CYP102A1H enzyme serves as a convenient model system for the more complex membrane-bound enzyme assemblies (Whitehouse et al. Reference Whitehouse, Bell and Wong2012).

Uncoupled substrate, O2, and NAD(P)H consumption in P450 catalysis is a well-recognized and relatively common phenomenon (De Matteis et al. Reference De Matteis, Dawson, Pons and Pipino2002, Reference De Matteis, Ballou, Coon, Estabrook and Haines2012; Denisov et al. Reference Denisov, Baas, Grinkova and Sligar2007a ; Grinkova et al. Reference Grinkova, Denisov, McLean and Sligar2013; Puntarulo & Cederbaum, Reference Puntarulo and Cederbaum1998; Staudt et al. Reference Staudt, Lichtenb and Ullrich1974). If two reducing equivalents are not delivered to O2 by the substrate, then alternative sources are necessary to avoid ROS production and/or enzyme degradation. In some cases, the extra equivalents can be delivered by NAD(P)H, leading to NAD(P)H : O2 molar consumption ratios greater than 1 (De Matteis et al. Reference De Matteis, Ballou, Coon, Estabrook and Haines2012). Exogenous reductants such as bilirubin and uroporphyrinogen have been shown to contribute reducing equivalents during NAD(P)H/O2 CYP102A1 turnover in the presence of halogenated (perfluorolaurate) substrates (De Matteis et al. Reference De Matteis, Ballou, Coon, Estabrook and Haines2012). Although it is possible that an active site hole could tunnel to the protein surface in a single step, a multistep radical transfer mechanism would be far more efficient. There are two attractive radical transfer pathways from the CYP102A1 heme to the protein surface (Fig. 3) (Girvan et al. Reference Girvan, Seward, Toogood, Cheesman, Leys and Munro2007). Pathway I is comprised of heme–Trp96–Trp90–Tyr334; pathway II is heme–Cys156–Tyr115–Met112–Tyr305.

Fig. 3. (a) Space-filling structural model of the heme domain of CYP102A1 (PDB #2IJ2) highlighting the surface locations of terminal residues in pathways I (Tyr334) and II (Tyr305). (b) Space-filling model of the residues comprising CYP102A1 radical transfer pathways I and II. Blue spheres represent structurally resolved water molecules.

CYP102A1 radical transfer pathway I

The shortest direct distance between aromatic atoms of CYP102A1 Trp96 and the heme is 7.3 Å and Trp(Nε)96 is hydrogen bonded to the heme propionate (Girvan et al. Reference Girvan, Seward, Toogood, Cheesman, Leys and Munro2007). Sequence alignment (UniProtKB) in the P450 family suggests that Trp is conserved at this position in >75% of the members of this group. Interestingly, of the 698 sequences with Trp at this position, all but 5 derive from eukaryotic sources, whereas about half of the proteins with His at this position derive from bacterial or archaeal sources. In this regard, it is noteworthy that archaeal CYP119 does not have a Trp residue at this site and is the only P450 in which Cmpd-1 has been characterized (Park et al. Reference Park, Yamane, Adachi, Shiro, Weiss, Maves and Sligar2002; Rittle & Green, Reference Rittle and Green2010). The strong conservation of the Trp96 residue has been noted previously (Munro et al. Reference Munro, Malarkey, McKnight, Thomson, Kelly, Price, Lindsay, Coggins and Miles1994). To the best of our knowledge, no role other than structural has been reported for this highly conserved Trp residue in P450 (Whitehouse et al. Reference Whitehouse, Bell and Wong2012).

We suggest that Trp96 is the gateway residue for hole transfer from the heme to the protein surface during uncoupled turnover. Studies of the reactions of substrate-free P450cam (CYP101) with peracids revealed that a second intermediate (Cmpd-ES) forms as a result of ET from a Tyr residue to Cmpd-1 (Schünemann et al. Reference Schünemann, Lendzian, Jung, Contzen, Barra, Sligar and Trautwein2004; Spolitak et al. Reference Spolitak, Dawson and Ballou2005, Reference Spolitak, Dawson and Ballou2006, Reference Spolitak, Dawson and Ballou2008). A Cmpd-ES intermediate has been detected in CYP102A1 and Trp96 has been implicated as one of the residues hosting the oxidized radical (Raner et al. Reference Raner, Thompson, Haddy, Tangham, Bynum, Reddy, Ballou and Dawson2006). Addition of NADPH to Cmpd-ES of the CYP102HR holoenzyme regenerates the ferric resting state; and formation of these radicals may play a protective role during uncoupled P450 catalysis (Spolitak et al. Reference Spolitak, Dawson and Ballou2006). A combined computational/experimental investigation of CYP102A1 implicated buried Trp96, Trp90, His92, and Tyr334 residues as components of an ET pathway that could deliver reducing equivalents to Cmpd-1 from the protein surface (Vidal-Limon et al. Reference Vidal-Limon, Aguila, Ayala, Batista and Vazquez-Duhalt2013). The shortest aromatic contacts in this chain are: Trp96–Trp90, 8.4 Å; Trp90–Tyr334, 4.4 Å (Girvan et al. Reference Girvan, Seward, Toogood, Cheesman, Leys and Munro2007). The environment around Tyr334 appears well-suited for radical formation: the phenol hydroxyl group is hydrogen-bonded to both a carboxylate (Asp68) and a water molecule (HOH1215).

Our prior studies of P450 ET reactions are consistent with involvement of Trp96 in a radical transfer pathway to the heme (Ener et al. Reference Ener, Lee, Winkler, Gray and Cheruzel2010). We have found that RuII(bpy)2(phen•––Cys97) can deliver an electron across 24 Å to the FeIII-heme in 20 μs, and RuIII(bpy)2(phen–Cys97)CYP102A1H can oxidize the heme to a porphyrin radical in under 2 μs (Ener et al. Reference Ener, Lee, Winkler, Gray and Cheruzel2010). The latter reaction is particularly rapid given the low driving force (<200 meV) expected for the transformation. We have prepared a Trp96His mutant and found that RuIII(bpy)2(phen–Cys97)(His96)CYP102A1H does not promote photochemical heme oxidation to Cmpd-2. Electron transfer to the FeIII-heme from RuII(bpy)2(phen•–−Cys97)(His96), however, is unaffected by the Trp96His mutation.

CYP102A1 radical transfer pathway II

The second potential radical transfer pathway in CYP102A1, heme–Cys156–Tyr115–Met112–Tyr305, does not appear as favorable as pathway I, due largely to a long distance between the heme and the first step in the path. The distance from Cys(Sγ)156 to the closest heme aromatic carbon atom (10.8 Å) is slightly longer than the shortest aromatic–aromatic contact between the heme and Tyr115 (10.2 Å). If a radical is formed on Tyr115, then hole transport to the surface Tyr305 via Met(Sδ)112 could provide a secondary protection route.

Potential radical transfer pathways in 2OG-Fe oxygenases

TauD

The 2-oxoglutarate nonheme iron oxygenases catalyze substrate hydroxylation reactions in a fashion that is reminiscent of the cytochromes P450, but with some critical distinctions (Fig. 1). The consensus mechanism for catalysis involves Fe2+ binding to the apo-enzyme followed by 2OG incorporation. Substrate binding induces loss of the water ligand from Fe2+, creating a vacant coordination site for O2 binding. Oxidation of 2OG produces CO2, succinate, and an FeIV(O) center that is thought to hydroxylate substrate via the usual H-atom abstraction, hydroxyl rebound cycle (Mantri et al. Reference Mantri, Zhang, McDonough and Schofield2012; Rose et al. Reference Rose, McDonough, King, Kawamura and Schofield2011). The 2OG-Fe hydroxylases differ from the P450 enzymes in that substrate hydroxylation proceeds from the FeIV(O) oxidation level (equivalent to P450 Cmpd-2). The E. coli 2OG-Fe enzyme TauD is synthesized under conditions of sulfur deprivation (Hausinger, Reference Hausinger2004); large quantities of TauD have been prepared by over expression in E. coli BL21(DE3)(pME4141) cells (Eichhorn et al. Reference Eichhorn, van der Ploeg, Kertesz and Leisinger1997; Ryle et al. Reference Ryle, Padmakumar and Hausinger1999). The enzyme catalyzes the hydroxylation of taurine (2-aminoethanesulfonate), producing an unstable species that decomposes into sulfite and aminoacetaldehyde (Hausinger, Reference Hausinger2004). In the absence of taurine, the enzyme will slowly consume O2 and become inactivated: protein analysis indicates hydroxylation of Tyr73 (Koehntop et al. Reference Koehntop, Marimanikkuppam, Ryle, Hausinger and Que2006; Ryle et al. Reference Ryle, Liu, Muthukumaran, Ho, Koehntop, McCracken, Que and Hausinger2003). Although with deuterated substrates coupling between oxygen consumption and substrate hydroxylation is diminished, 2OG oxidation is not, suggesting that FeIV(O) continues to be formed in the presence of refractory substrates; and bis-Tris buffer, a potential reducing agent, decreases coupling between O2 activation and C–H hydroxylation (McCusker & Klinman, Reference McCusker and Klinman2009). We suggest that when FeIV(O) is unable to effect substrate hydroxylation, the oxidizing hole is directed to the protein surface where it can be reduced by external reagents.

TauD radical transfer pathways

We have identified two possible radical transfer pathways in the structure of TauD: the most attractive pathway from Fe to the surface has four Trp residues: Fe–Trp248–Trp128–Trp240–Trp238; relevant distances are: Fe–Trp248, 4.8 Å; Trp248–Trp128, 3.1 Å; Trp128–Trp240, 3.7 Å; Trp240–Trp238, 3.7 Å (Fig. 4) (O'Brien et al. Reference O'Brien, Schuller, Yang, Dillard and Lanzilotta2003). The structure of this Trp chain compares favorably to that identified in E. coli DNA photolyase (4–5 Å separations) (Byrdin et al. Reference Byrdin, Eker, Vos and Brettel2003; Lukacs et al. Reference Lukacs, Eker, Byrdin, Villette, Pan, Brettel and Vos2006). The photolyase chain has just three Trp residues, and hole migration from FADH•* to Trp306 at the protein surface is complete in less than 10 ns (Byrdin et al. Reference Byrdin, Eker, Vos and Brettel2003; Lukacs et al. Reference Lukacs, Eker, Byrdin, Villette, Pan, Brettel and Vos2006). We anticipate that a hole injected by FeIV(O)-TauD into Trp248 should migrate to Trp238 at the surface in less than 1 μs. A secondary radical transfer pathway in TauD [Fe–Tyr73–Tyr164–(Trp174,Tyr162)] is of particular interest because hydroxylated Tyr73 has been found during turnover in the absence of taurine (Koehntop et al. Reference Koehntop, Marimanikkuppam, Ryle, Hausinger and Que2006; Ryle et al. Reference Ryle, Liu, Muthukumaran, Ho, Koehntop, McCracken, Que and Hausinger2003). Both Trp174 and Tyr162 are well-exposed at the enzyme surface and both (or just one) of these residues could be involved in a radical transfer pathway. Relevant distances are: Fe–Tyr73, 6.5 Å; Tyr73–Tyr164, 5.0 Å; Tyr164–Trp174, 4.2 Å; Tyr164–Tyr162, 7.6 Å; Trp174–Tyr162, 8.8 Å (O'Brien et al. Reference O'Brien, Schuller, Yang, Dillard and Lanzilotta2003).

Fig. 4. (a) Space-filling structural model of E. coli TauD (PDB #1OS7) highlighting the surface locations of terminal residues in postulated radical transfer pathways (Trp238, Trp174, and Tyr162). (b) Space-filling model of the residues comprising TauD radical transfer pathways.

Outlook

Functional radical transfer pathways have been identified in several enzymes, including ribonucleotide reductase (Argirevic et al. Reference Argirevic, Riplinger, Stubbe, Neese and Bennati2012; Holder et al. Reference Holder, Pizano, Anderson, Stubbe and Nocera2012; Offenbacher et al. Reference Offenbacher, Burns, Sherrill and Barry2013a , Reference Offenbacher, Minnihan, Stubbe and Barry b ; Sjöberg, Reference Sjöberg1997; Stubbe & van der Donk, Reference Stubbe and van der Donk1998; Stubbe et al. Reference Stubbe, Nocera, Yee and Chang2003; Worsdorfer et al. Reference Worsdorfer, Conner, Yokoyama, Livada, Seyedsayamdost, Jiang, Silakov, Stubbe, Bollinger and Krebs2013; Yokoyama et al. Reference Yokoyama, Smith, Corzilius, Griffin and Stubbe2011), photosystem II (Boussac et al. Reference Boussac, Rappaport, Brettel and Sugiura2013; Keough et al. Reference Keough, Zuniga, Jenson and Barry2013; Sjoholm et al. Reference Sjoholm, Styring, Havelius and Ho2012), DNA photolyase (Aubert et al. Reference Aubert, Mathis, Eker and Brettel1999, Reference Aubert, Vos, Mathis, Eker and Brettel2000; Byrdin et al. Reference Byrdin, Eker, Vos and Brettel2003; Kodali et al. Reference Kodali, Siddiqui and Stanley2009; Li et al. Reference Li, Heelis and Sancar1991b ; Lukacs et al. Reference Lukacs, Eker, Byrdin, Villette, Pan, Brettel and Vos2006; Sancar, Reference Sancar2003; Taylor, Reference Taylor1994; Woiczikowski et al. Reference Woiczikowski, Steinbrecher, Kubař and Elstner2011), and MauG (Davidson & Liu, Reference Davidson and Liu2012; Davidson & Wilmot, Reference Davidson and Wilmot2013; Geng et al. Reference Geng, Dornevil, Davidson and Liu2013; Yukl et al. Reference Yukl, Liu, Krzystek, Shin, Jensen, Davidson, Wilmot and Liu2013). If radical transfer pathways do indeed provide protection mechanisms for enzymes operating at high electrochemical potentials, then it is likely that they will be found in many more redox-active enzymes. A survey of oxidoreductases in the protein data bank reveals that nearly 80% of structurally characterized peroxidases, oxygenases, and dioxygenases (enzyme classes EC 1.11, 1.13, and 1.14; 587 structures with sequence identity less than 90%) contain chains of 2 or more redox-active residues (Tyr, Trp, heme, Fe, and Cu) separated by no more than 5 Å (Fig. 5). The fraction increases to almost 90% if the cutoff distance is increased to 8 Å. We think it very likely that hole hopping through these types of radical transfer chains greatly reduces the production of ROS that destroy enzymes and other molecules in living cells.

Fig. 5. Distributions of radical transfer chain lengths among structurally characterized oxidoreductases from enzyme sub-classes EC 1.11 (peroxidases, blue), 1.13 (oxygenases, green), and 1.14 (dioxygenases, red). Radical transfer chains are defined to be composed of Tyr, Trp, heme, Fe, and Cu residues. Tyr residues were included only if a carboxylate (Asp, Glu) oxygen atom, an imidazole (His) nitrogen atom, or a water molecule was within 4 Å of the Tyr hydroxyl oxygen atom.

Acknowledgments

We thank Maraia Ener, Jeff Warren, Lionel Cheruzel, Kana Takematsu, and Oliver Shafaat for helpful discussions.

Financial support

Research reported in this publication was supported by The National Institute of Diabetes and Digestive and Kidney Diseases of the National Institutes of Health under award number R01DK019038 to HBG and JRW. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health. Additional support was provided by the Arnold and Mabel Beckman Foundation.

References

Argirevic, T., Riplinger, C., Stubbe, J., Neese, F. & Bennati, M. (2012). ENDOR spectroscopy and DFT calculations: evidence for the hydrogen-bond network within alpha 2 in the PCET of E. coli ribonucleotide reductase. Journal of the American Chemical Society 134, 17661–17670.Google Scholar
Aubert, C., Mathis, P., Eker, A. P. M. & Brettel, K. (1999). Intraprotein electron transfer between tyrosine and tryptophan in DNA photolysase from Anacystis nidulans . Proceedings of the National Academy of Sciences of the United States of America 96, 5423–5427.Google Scholar
Aubert, C., Vos, M. H., Mathis, P., Eker, A. P. M. & Brettel, K. (2000). Intraprotein radical transfer during photoactivation of DNA photolyase. Nature 405, 586–590.Google Scholar
Blanco-Rodriguez, A. M., Di Bilio, A. J., Shih, C., Museth, A. K., Clark, I. P., Towrie, M., Cannizzo, A., Sudhamsu, J., Crane, B. R., Sykora, J., Winkler, J. R., Gray, H. B., Zalis, S. & Vlcek, A. (2011). Phototriggering electron flow through ReI-modified Pseudomonas aeruginosa azurins. Chemistry – A European Journal 17, 5350–5361.CrossRefGoogle ScholarPubMed
Boddupalli, S. S., Estabrook, R. W. & Peterson, J. A. (1990). Fatty-acid monooxygenation by cytochrome-P-450BM-3. Journal of Biological Chemistry 265, 4233–4239.Google Scholar
Boddupalli, S. S., Oster, T., Estabrook, R. W. & Peterson, J. A. (1992). Reconstitution of the fatty-acid hydroxylation function of cytochrome-P-450BM-3 utilizing its individual recombinant hemoprotein and flavoprotein domains. Journal of Biological Chemistry 267, 10375–10380.Google Scholar
Bonin, J., Costentin, C., Louault, C., Robert, M., Routier, M. & Saveant, J. M. (2010). Intrinsic reactivity and driving force dependence in concerted proton-electron transfers to water illustrated by phenol oxidation. Proceedings of the National Academy of Sciences of the United States of America 107, 3367–3372.Google Scholar
Boussac, A., Rappaport, F., Brettel, K. & Sugiura, M. (2013). Charge recombination in SnTyrZ •QA −• radical pairs in D1 protein variants of photosystem II: long range electron transfer in the marcus inverted region. Journal of Physical Chemistry B 117, 3308–3314.Google Scholar
Byrdin, M., Eker, A. P. M., Vos, M. H. & Brettel, K. (2003). Dissection of the triple tryptophan electron transfer chain in Escherichia coli DNA photolyase: Trp382 is the primary donor in photoactivation. Proceedings of the National Academy of Sciences of the United States of America 100, 8676–8681.Google Scholar
Chen, Y. H., Comeaux, L. M., Herbst, R. W., Saban, E., Kennedy, D. C., Maroney, M. J. & Knapp, M. J. (2008). Coordination changes and auto-hydroxylation of FIH-1: uncoupled O2-activation in a human hypoxia sensor. Journal of Inorganic Biochemistry 102, 2120–2129.Google Scholar
Costentin, C., Louault, C., Robert, M. & Saveant, J. M. (2009). The electrochemical approach to concerted proton-electron transfers in the oxidation of phenols in water. Proceedings of the National Academy of Sciences of the United States of America 106, 18143–18148.Google Scholar
Davidson, V. L. & Liu, A. M. (2012). Tryptophan tryptophylquinone biosynthesis: a radical approach to posttranslational modification. Biochimica et Biophysica Acta – Proteins and Proteomics 1824, 1299–1305.Google Scholar
Davidson, V. L. & Wilmot, C. M. (2013). Posttranslational biosynthesis of the protein-derived cofactor tryptophan tryptophylquinone. Annual Review of Biochemistry 82, 531–550.Google Scholar
De Matteis, F., Ballou, D. P., Coon, M. J., Estabrook, R. W. & Haines, D. C. (2012). Peroxidase-like activity of uncoupled cytochrome P450. Studies with bilirubin and toxicological implications of uncoupling. Biochemical Pharmacology 84, 374–382.Google Scholar
De Matteis, F., Dawson, S. J., Pons, N. & Pipino, S. (2002). Bilirubin and uroporphyrinogen oxidation by induced cytochrome P4501A and cytochrome P4502B – role of polyhalogenated biphenyls of different configuration. Biochemical Pharmacology 63, 615–624.Google Scholar
Denisov, I. G., Baas, B. J., Grinkova, Y. V. & Sligar, S. G. (2007 a). Cooperativity in cytochrome P450 3A4 – Linkages in substrate binding, spin state, uncoupling, and product formation. Journal of Biological Chemistry 282, 7066–7076.Google Scholar
Denisov, I. G., Grinkova, Y. V., McLean, M. A. & Sligar, S. G. (2007 b). The one-electron autoxidation of human cytochrome p450 3A4. Journal of Biological Chemistry 282, 26865–26873.Google Scholar
Denisov, I. G., Makris, T. M., Sligar, S. G. & Schlichting, I. (2005). Structure and chemistry of cytochrome P450. Chemical Reviews 105, 2253–2277.CrossRefGoogle ScholarPubMed
Eichhorn, E., van der Ploeg, J. R., Kertesz, M. A. & Leisinger, T. (1997). Characterization of α-Ketoglutarate-dependent taurine dioxygenase from Escherichia coli . Journal of Biological Chemistry 272, 23031–23036.Google Scholar
Ener, M. E., Lee, Y. T., Winkler, J. R., Gray, H. B. & Cheruzel, L. (2010). Photooxidation of cytochrome P450-BM3. Proceedings of the National Academy of Sciences of the United States of America 107, 18783–18786.Google Scholar
Farver, O. & Pecht, I. (2011). Electron transfer in blue copper proteins. Coordination Chemistry Reviews 255, 757–773.Google Scholar
Geng, J. F., Dornevil, K., Davidson, V. L. & Liu, A. M. (2013). Tryptophan-mediated charge-resonance stabilization in the bis-Fe(IV) redox state of MauG. Proceedings of the National Academy of Sciences of the United States of America 110, 9639–9644.Google Scholar
Girvan, H. M., Seward, H. E., Toogood, H. S., Cheesman, M. R., Leys, D. & Munro, A. W. (2007). Structural and spectroscopic characterization of P450BM3 mutants with unprecedented P450 heme iron ligand sets – new heme ligation states influence conformational equilibria in P450BM3. Journal of Biological Chemistry 282, 564–572.CrossRefGoogle Scholar
Gray, H. B. & Winkler, J. R. (2010). Electron flow through metalloproteins. Biochimica Et Biophysica Acta – Bioenergetics 1797, 1563–1572.Google Scholar
Grinkova, Y. V., Denisov, I. G., McLean, M. A. & Sligar, S. G. (2013). Oxidase uncoupling in heme monooxygenases: human cytochrome P450 CYP3A4 in nanodiscs. Biochemical and Biophysical Research Communications 430, 1223–1227.Google Scholar
Grinkova, Y. V., Denisov, I. G. & Sligar, S. G. (2010). Functional reconstitution of monomeric CYP3A4 with multiple cytochrome P450 reductase molecules in nanodiscs. Biochemical and Biophysical Research Communications 398, 194–198.Google Scholar
Harriman, A. (1987). Further comments on the redox potentials of tryptophan and tyrosine. Journal of Physical Chemistry 91, 6102–6104.Google Scholar
Hausinger, R. P. (2004). Fe(II)/alpha-ketoglutarate-dependent hydroxylases and related enzymes. Critical Reviews in Biochemistry and Molecular Biology 39, 21–68.Google Scholar
Holder, P. G., Pizano, A. A., Anderson, B. L., Stubbe, J. & Nocera, D. G. (2012). Deciphering radical transport in the large subunit of class I ribonucleotide reductase. Journal of the American Chemical Society 134, 1172–1180.Google Scholar
Johnson, E. F. & Stout, C. D. (2013). Structural diversity of eukaryotic membrane cytochrome P450s. Journal of Biological Chemistry 288, 17082–17090.Google Scholar
Jovanic, S. V., Harriman, A. & Simic, M. G. (1986). Electron-transfer reactions of tryptophan and tyrosine derivatives. Journal of Physical Chemistry 90, 1935–1939.CrossRefGoogle Scholar
Keough, J. M., Zuniga, A. N., Jenson, D. L. & Barry, B. A. (2013). Redox control and hydrogen bonding networks: proton-coupled electron transfer reactions and tyrosine Z in the photosynthetic oxygen-evolving complex. Journal of Physical Chemistry B 117, 1296–1307.CrossRefGoogle ScholarPubMed
Kodali, G., Siddiqui, S. U. & Stanley, R. J. (2009). Charge redistribution in oxidized and semiquinone E. coli DNA photolyase upon photoexcitation: stark spectroscopy reveals a rationale for the position of Trp382. Journal of the American Chemical Society 131, 4795–4807.Google Scholar
Koehntop, K. D., Marimanikkuppam, S., Ryle, M. J., Hausinger, R. P. & Que, L. (2006). Self-hydroxylation of taurine/alpha-ketoglutarate dioxygenase: evidence for more than one oxygen activation mechanism. Journal of Biological Inorganic Chemistry 11, 63–72.Google Scholar
Li, H. Y., Darwish, K. & Poulos, T. L. (1991 a). Characterization of recombinant Bacillus megaterium cytochrome-P-450BM-3 and its 2 functional domains. Journal of Biological Chemistry 266, 11909–11914.Google Scholar
Li, Y. F., Heelis, P. F. & Sancar, A. (1991 b). Active site of DNA photolyase: tryptophan-306 is the intrinsic hydrogen atom donor essential for flavin radical photoreduction and DNA repair in vitro. Biochemistry 30, 6322–6329.Google Scholar
Lukacs, A., Eker, A. P. M., Byrdin, M., Villette, S., Pan, J., Brettel, K. & Vos, M. H. (2006). Role of the middle residue in the triple tryptophan electron transfer chain of DNA photolyase: ultrafast spectroscopy of a Trp→Phe mutant. Journal of Physical Chemistry B 110, 15654–15658.Google Scholar
Mantri, M., Zhang, Z. H., McDonough, M. A. & Schofield, C. J. (2012). Autocatalysed oxidative modifications to 2-oxoglutarate dependent oxygenases. FEBS Journal 279, 1563–1575.Google Scholar
Marcus, R. A. & Sutin, N. (1985). Electron transfers in chemistry and biology. Biochimica et Biophysica Acta 811, 265–322.CrossRefGoogle Scholar
McCusker, K. P. & Klinman, J. P. (2009). Modular behavior of tauD provides insight into the origin of specificity in alpha-ketoglutarate-dependent nonheme iron oxygenases. Proceedings of the National Academy of Sciences of the United States of America 106, 19791–19795.Google Scholar
Miura, Y. & Fulco, A. J. (1974). (Omega – 2) Hydroxylation of fatty-acids by a soluble system from Bacillus megaterium . Journal of Biological Chemistry 249, 1880–1888.Google Scholar
Munro, A. W., Malarkey, K., McKnight, J., Thomson, A. J., Kelly, S. M., Price, N. C., Lindsay, J. G., Coggins, J. R. & Miles, J. S. (1994). The role of tryptophan-97 of cytochrome-P450-BM3 from Bacillus megaterium in catalytic function – evidence against the covalent-switching hypothesis of P450 electron-transfer. Biochemical Journal 303, 423–428.CrossRefGoogle Scholar
Narhi, L. O. & Fulco, A. J. (1986). Characterization of a catalytically self-sufficient 119,000-dalton cytochrome-P-450 monooxygenase induced by barbiturates in Bacillus megaterium . Journal of Biological Chemistry 261, 7160–7169.Google Scholar
Narhi, L. O., Wen, L. P. & Fulco, A. J. (1988). Characterization of the protein expressed in Escherichia coli by a recombinant plasmid containing the Bacillus megaterium cytochrome-P-450BM-3 gene. Molecular and Cellular Biochemistry 79, 63–71.Google Scholar
Nebert, D. W., Wikvall, K. & Miller, W. L. (2013). Human cytochromes P450 in health and disease. Philosophical Transactions of the Royal Society B – Biological Sciences 368 (1612), 20120431.Google Scholar
O'Brien, J. R., Schuller, D. J., Yang, V. S., Dillard, B. D. & Lanzilotta, W. N. (2003). Substrate-induced conformational changes in Escherichia coli taurine/α-ketoglutarate dioxygenase and insight into the oligomeric structure. Biochemistry 42, 5547–5554.Google Scholar
Offenbacher, A. R., Burns, L. A., Sherrill, C. D. & Barry, B. A. (2013 a). Redox-linked conformational control of proton-coupled electron transfer: Y122 in the ribonucleotide reductase β2 subunit. Journal of Physical Chemistry B 117, 8457–8468.Google Scholar
Offenbacher, A. R., Minnihan, E. C., Stubbe, J. & Barry, B. A. (2013 b). Redox-linked changes to the hydrogen-bonding network of ribonucleotide reductase β2. Journal of the American Chemical Society 135, 6380–6383.Google Scholar
Orr, S. T. M., Ripp, S. L., Ballard, T. E., Henderson, J. L., Scott, D. O., Obach, R. S., Sun, H. & Kalgutkar, A. S. (2012). Mechanism-based inactivation (MBI) of cytochrome P450 enzymes: structure-activity relationships and discovery strategies to mitigate drug-drug interaction risks. Journal of Medicinal Chemistry 55, 4896–4933.Google Scholar
Oster, T., Boddupalli, S. S. & Peterson, J. A. (1991). Expression, purification, and properties of the flavoprotein domain of cytochrome-P-450BM-3 – evidence for the importance of the amino-terminal region for FMN binding. Journal of Biological Chemistry 266, 22718–22725.CrossRefGoogle ScholarPubMed
Park, S.-Y., Yamane, K., Adachi, S.-I., Shiro, Y., Weiss, K. E., Maves, S. A. & Sligar, S. G. (2002). Thermophilic cytochrome P450 (CYP119) from Sulfolobus solfataricus: high resolution structure and functional properties. Journal of Inorganic Biochemistry 91, 491–501.Google Scholar
Puntarulo, S. & Cederbaum, A. I. (1998). Production of reactive oxygen species by microsomes enriched in specific human cytochrome P450 enzymes. Free Radical Biology and Medicine 24, 1324–1330.Google Scholar
Raner, G. M., Thompson, J. I., Haddy, A., Tangham, V., Bynum, N., Reddy, G. R., Ballou, D. P. & Dawson, J. H. (2006). Spectroscopic investigations of intermediates in the reaction of cytochrome P450(BM3)-F87G with surrogate oxygen atom donors. Journal of Inorganic Biochemistry 100, 2045–2053.Google Scholar
Reece, S. Y. & Nocera, D. G. (2009). Proton-coupled electron transfer in biology: results from synergistic studies in natural and model systems. Annual Review of Biochemistry, 78, 673–699.Google Scholar
Rittle, J. & Green, M. T. (2010). Cytochrome P450 compound I: capture, characterization, and C–H bond activation kinetics. Science 330, 933–937.CrossRefGoogle ScholarPubMed
Rose, N. R., McDonough, M. A., King, O. N. F., Kawamura, A. & Schofield, C. J. (2011). Inhibition of 2-oxoglutarate dependent oxygenases. Chemical Society Reviews 40, 4364–4397.Google Scholar
Ryle, M. J., Liu, A., Muthukumaran, R. B., Ho, R. Y. N., Koehntop, K. D., McCracken, J., Que, L. & Hausinger, R. P. (2003). O-2- and alpha-ketoglutarate-dependent tyrosyl radical formation in TauD, an alpha-keto acid-dependent non-heme iron dioxygenase. Biochemistry 42, 1854–1862.Google Scholar
Ryle, M. J., Padmakumar, R. & Hausinger, R. P. (1999). Stopped-flow kinetic analysis of Escherichia coli taurine/α-ketoglutarate dioxygenase: interactions with α-ketoglutarate, taurine, and oxygen†. Biochemistry 38, 15278–15286.Google Scholar
Saban, E., Flagg, S. C. & Knapp, M. J. (2011). Uncoupled O2-activation in the human HIF-asparaginyl hydroxylase, FIH, does not produce reactive oxygen species. Journal of Inorganic Biochemistry 105, 630–636.Google Scholar
Sancar, A. (2003). Structure and function of DNA photolyase and cryptochrome blue-light photoreceptors. Chemical Reviews 103, 2203–2238.Google Scholar
Schünemann, V., Lendzian, F., Jung, C., Contzen, J., Barra, A. L., Sligar, S. G. & Trautwein, A. X. (2004). Tyrosine radical formation in the reaction of wild type and mutant cytochrome P450cam with peroxy acids – a multifrequency EPR study of intermediates on the millisecond time scale. Journal of Biological Chemistry 279, 10919–10930.Google Scholar
Sevrioukova, I. F., Immoos, C. E., Poulos, T. L. & Farmer, P. (2000). Electron transfer in the ruthenated heme domain of cytochrome P450BM-3. Israel Journal of Chemistry 40, 47–53.Google Scholar
Shih, C., Museth, A. K., Abrahamsson, M., Blanco-Rodriguez, A. M., Di Bilio, A. J., Sudhamsu, J., Crane, B. R., Ronayne, K. L., Towrie, M., Vlcek, A., Richards, J. H., Winkler, J. R. & Gray, H. B. (2008). Tryptophan-accelerated electron flow through proteins. Science 320, 1760–1762.CrossRefGoogle ScholarPubMed
Sjöberg, B. M. (1997). Ribonucleotide reductases – a group of enzymes with different metallosites and a similar mechanism. Structure and Bonding 88, 139–173.Google Scholar
Sjoholm, J., Styring, S., Havelius, K. G. V. & Ho, F. M. (2012). Visible light induction of an electron paramagnetic resonance split signal in photosystem II in the S-2 state reveals the importance of charges in the oxygen-evolving center during catalysis: a unifying model. Biochemistry 51, 2054–2064.Google Scholar
Spolitak, T., Dawson, J. H. & Ballou, D. P. (2005). Reaction of ferric cytochrome P450cam with peracids – kinetic characterization of intermediates on the reaction pathway. Journal of Biological Chemistry 280, 20300–20309.Google Scholar
Spolitak, T., Dawson, J. H. & Ballou, D. P. (2006). Rapid kinetics investigations of peracid oxidation of ferric cytochrome P450cam: nature and possible function of compound ES. Journal of Inorganic Biochemistry 100, 2034–2044.Google Scholar
Spolitak, T., Dawson, J. H. & Ballou, D. P. (2008). Replacement of tyrosine residues by phenylalanine in cytochrome P450cam alters the formation of Cpd II-like species in reactions with artificial oxidants. Journal of Biological Inorganic Chemistry 13, 599–611.Google Scholar
Staudt, H., Lichtenb, F. & Ullrich, V. (1974). Role of NADH in uncoupled microsomal monoxygenations. European Journal of Biochemistry 46, 99–106.Google Scholar
Stubbe, J., Nocera, D. G., Yee, C. S. & Chang, M. C. Y. (2003). Radical initiation in the class I ribonucleotide reductase: long-range proton-coupled electron transfer? Chemical Reviews 103, 2167–2201.Google Scholar
Stubbe, J. & van der Donk, W. A. (1998). Protein radicals in enzyme catalysis. Chemical Reviews 98, 705–762.Google Scholar
Takematsu, K., Williamson, H., Blanco-Rodríguez, A. M., Sokolová, L., Nikolovski, P., Kaiser, J. T., Towrie, M., Clark, I. P., Vlček, A., Winkler, J. R. & Gray, H. B. (2013). Tryptophan-accelerated electron flow across a protein–protein interface. Journal of the American Chemical Society 134, 15515–15525.Google Scholar
Taylor, J. S. (1994). Unraveling the molecular pathway from sunlight to skin cancer. Accounts of Chemical Research 27, 76–82.Google Scholar
Vidal-Limon, A., Aguila, S., Ayala, M., Batista, C. V. & Vazquez-Duhalt, R. (2013). Peroxidase activity stabilization of cytochrome P450(BM3) by rational analysis of intramolecular electron transfer. Journal of Inorganic Biochemistry 122, 18–26.CrossRefGoogle ScholarPubMed
Warman, A. J., Roitel, O., Neeli, R., Girvan, H. M., Seward, H. E., Murray, S. A., McLean, K. J., Joyce, M. G., Toogood, H., Holt, R. A., Leys, D., Scrutton, N. S. & Munro, A. W. (2005). Flavocytochrome P450BM3: an update on structure and mechanism of a biotechnologically important enzyme. Biochemical Society Transactions 33, 747–753.Google Scholar
Warren, J. J., Ener, M. E., Vlček, A., Winkler, J. R. & Gray, H. B. (2012). Electron hopping through proteins. Coordination Chemistry Reviews 256, 2478–2487.Google Scholar
Warren, J. J., Herrera, N., Hill, M. G., Winkler, J. R. & Gray, H. B. (2013 a). Electron flow through nitrotyrosinate in Pseudomonas aeruginosa azurin. Journal of the American Chemical Society 135, 11151–11158.Google Scholar
Warren, J. J., Winkler, J. R. & Gray, H. B. (2013 b). Hopping maps for photosynthetic reaction centers. Coordination Chemistry Reviews 257, 165–170.Google Scholar
Whitehouse, C. J. C., Bell, S. G. & Wong, L. L. (2012). P450BM3 (CYP102A1): connecting the dots. Chemical Society Reviews 41, 1218–1260.Google Scholar
Wikström, M. (2012). Active site intermediates in the reduction of O2 by cytochrome oxidase, and their derivatives. Biochimica et Biophysica Acta – Bioenergetics 1817, 468–475.Google Scholar
Wilson, T. D., Yu, Y. & Lu, Y. (2013). Understanding copper-thiolate containing electron transfer centers by incorporation of unnatural amino acids and the Cu-A center into the type 1 copper protein azurin. Coordination Chemistry Reviews 257, 260–276.Google Scholar
Winkler, J. R. & Gray, H. B. (2014 a). Electron flow through metalloproteins. Chemical Reviews 114, 3369–3380.Google Scholar
Winkler, J. R. & Gray, H. B. (2014 b). Long-range electron tunneling. Journal of the American Chemical Society 136, 2930–2939.Google Scholar
Woiczikowski, P. B., Steinbrecher, T., Kubař, T. & Elstner, M. (2011). Nonadiabatic QM/MM simulations of fast charge transfer in Escherichia coli DNA photolyase. Journal of Physical Chemistry B 115, 9846–9863.Google Scholar
Worsdorfer, B., Conner, D. A., Yokoyama, K., Livada, J., Seyedsayamdost, M., Jiang, W., Silakov, A., Stubbe, J., Bollinger, J. M. & Krebs, C. (2013). Function of the Diiron Cluster of Escherichia coli class Ia ribonucleotide reductase in proton-coupled electron transfer. Journal of the American Chemical Society 135, 8585–8593.Google Scholar
Yokoyama, K., Smith, A. A., Corzilius, B., Griffin, R. G. & Stubbe, J. (2011). Equilibration of tyrosyl radicals (Y-356•, Y-731•, Y-730•) in the radical propagation pathway of the Escherichia coli class Ia ribonucleotide reductase. Journal of the American Chemical Society 133, 18420–18432.Google Scholar
Yu, M. A., Egawa, T., Shinzawa-Itoh, K., Yoshikawa, S., Guallar, V., Yeh, S. R., Rousseau, D. L. & Gerfen, G. J. (2012). Two tyrosyl radicals stabilize high oxidation states in cytochrome c oxidase for efficient energy conservation and proton translocation. Journal of the American Chemical Society 134, 4753–4761.Google Scholar
Yu, M. A., Egawa, T., Shinzawa-Itoh, K., Yoshikawa, S., Yeh, S. R., Rousseau, D. L. & Gerfen, G. J. (2011). Radical formation in cytochrome c oxidase. Biochimica et Biophysica Acta – Bioenergetics 1807, 1295–1304.Google Scholar
Yukl, E. T., Liu, F. G., Krzystek, J., Shin, S., Jensen, L. M. R., Davidson, V. L., Wilmot, C. M. & Liu, A. M. (2013). Diradical intermediate within the context of tryptophan tryptophylquinone biosynthesis. Proceedings of the National Academy of Sciences of the United States of America 110, 4569–4573.Google Scholar
Figure 0

Fig. 1. Schematic representation of the catalytic mechanisms of P450 and 2OG-Fe oxygenases: RH, substrate; 2OG, 2-oxoglutarate; Suc, succinate. Black arrows indicate the functional substrate hydroxylation pathways. Blue arrows indicate oxidase uncoupling pathways.

Figure 1

Fig. 2. (a) Space-filling structural model of RuHis107NO2TyrOH109Cu-azurin. (b) Space filling models of the residues comprising the hole-hopping pathway from Cu to RuHis107.

Figure 2

Fig. 3. (a) Space-filling structural model of the heme domain of CYP102A1 (PDB #2IJ2) highlighting the surface locations of terminal residues in pathways I (Tyr334) and II (Tyr305). (b) Space-filling model of the residues comprising CYP102A1 radical transfer pathways I and II. Blue spheres represent structurally resolved water molecules.

Figure 3

Fig. 4. (a) Space-filling structural model of E. coli TauD (PDB #1OS7) highlighting the surface locations of terminal residues in postulated radical transfer pathways (Trp238, Trp174, and Tyr162). (b) Space-filling model of the residues comprising TauD radical transfer pathways.

Figure 4

Fig. 5. Distributions of radical transfer chain lengths among structurally characterized oxidoreductases from enzyme sub-classes EC 1.11 (peroxidases, blue), 1.13 (oxygenases, green), and 1.14 (dioxygenases, red). Radical transfer chains are defined to be composed of Tyr, Trp, heme, Fe, and Cu residues. Tyr residues were included only if a carboxylate (Asp, Glu) oxygen atom, an imidazole (His) nitrogen atom, or a water molecule was within 4 Å of the Tyr hydroxyl oxygen atom.