Invited Review De Novo Design of Functional Proteins: Toward Artificial Hydrogenases Marina Faiella, Anindya Roy, Dayn Sommer, Giovanna Ghirlanda Department of Chemistry and Biochemistry, Arizona State University, Tempe, AZ Received 8 March 2013; revised 8 July 2013; accepted 18 September 2013 Published online 6 October 2013 in Wiley Online Library (wileyonlinelibrary.com). DOI 10.1002/bip.22420

ABSTRACT: Over the last 25 years, de novo design has proven to be a valid approach to generate novel, well-folded proteins, and most recently, functional proteins. In response to societal needs, this approach is been used increasingly to design functional proteins developed with an eye toward sustainable fuel production. This review surveys recent examples of bioinspired de novo designed peptide based catalysts, focusing in particular C 2013 Wiley Periodicals, Inc. on artificial hydrogenases. V

Biopolymers (Pept Sci) 100: 558–571, 2013. Keywords: de novo design; peptide; hydrogenase; ironsulfur clusters; catalysis

This article was originally published online as an accepted preprint. The “Published Online” date corresponds to the preprint version. You can request a copy of the preprint by emailing the Biopolymers editorial office at [email protected]

INTRODUCTION

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ne of the most important challenges facing our society is the development of sustainable energy sources. The current reliance on fossil fuels has unsustainable consequences on the environment; further, the unequal distribution of energy sources

Correspondence to: Dr. Giovanna Ghirlanda, Arizona State University, Chemistry and Biochemistry, 1711 S Rural rd, Tempe, Arizona 85287-1604, USA; e-mail: [email protected] Contract grant sponsor: U.S. Department of Energy Contract grant sponsor: Office of Science Contract grant sponsor: Office of Basic Energy Sciences Contract grant number: DE-SC0001016 C 2013 Wiley Periodicals, Inc. V

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throughout the world results in conflicts and societal instability. Nature uses solar energy to power biological processes through photosynthesis, storing solar radiation in energy rich molecules that in turn are used to drive cellular metabolism. This process indirectly powers most life on the planet, as it maintains oxygen levels in the atmosphere, generates key organic molecules, and supports the growth of biomass that in turns is used as food. Fossil fuels are also ultimately derived from biomass resulting from photosynthetic solar energy conversion. To address our society’s energy needs, research efforts are focusing on developing new renewable based sources of energy to generate storable fuels. Hydrogen in particular has emerged as an attractive, carbon-free candidate for use as fuel.1 One of the approaches followed for sustainable hydrogen production is inspired by nature, and aims to capture solar energy by reproducing the fundamental steps of photosynthesis through dedicated functional modules interfaced in a device.2–5 In such a device, the reductive equivalents needed for hydrogen generation are obtained through the same reaction at the basis of photosynthesis, oxidation of water into O2 and H1, also known as water splitting, which is driven by photoinduced charge separation in the light harvester. In turn, molecular hydrogen is generated through appropriate catalysts capable of handling multielectron redox chemistry. In nature, hydrogen metabolism is mediated by specialized enzymes called hydrogenases found in Bacteria, Archea, and Eucarya.6–8 Based on the composition of their active sites, natural hydrogenases are divided in [FeFe], [NiFe], and [Fe] only, with the latter group involved in methanogenesis.9,10 The active site of hydrogenases also contains unusual smallmolecule ligands as part of the first coordination sphere. [NiFe] and [FeFe] hydrogenases, in particular are of interest in this review, as they catalyze the reversible reduction of protons to hydrogen following the reaction in Scheme 1.1,6,7,11–14

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SCHEME 1 hydrogen.

Reversible reduction of protons to molecular

Despite no phylogenetic relationship, extensive structural and spectroscopic characterization revealed that both contain binuclear active sites that use unique, nonproteinogenic CO, and CN ligands.11,15–21 Hydrogenases can also be classified in terms of function: uptake hydrogenases, generally present in the periplasm or associated with the membrane, utilize molecular hydrogen as a source of electrons. Conversely, hydrogenases localized in the cytoplasm catalyze proton reduction as a way to dispose excess electrons.7 Both functions are relevant to technological applications, in the sustainable production of hydrogen, and in the efficient utilization of molecular hydrogen in fuel cells.1,22 The utilization of hydrogenases in devices is hindered by the complexity of their biosynthesis and structural architecture, which makes heterologous expression technically challenging, and by their high oxygen sensitivity. The complex pathway required for active site assembly and hydrogenase maturation has prevented the application of well-established methods to optimize natural enzymes, such as directed evolution, which could in principle be used to ameliorate oxygen sensitivity.23–25 A complementary approach to developing robust catalysts for use in devices uses organometallic complexes. Over the years, several groups have developed models with the dual purpose of elucidating the mechanistic requirements of the active site and applying them as a robust alternative to the hydrogenases. These complexes follow two broad philosophies: on one hand, structural mimicry of the natural enzymes, using similar ligands and metals to reproduce the enzyme’s active site; however, functional mimics aim to replicate the function using Earth-abundant metals and artificial ligands to form complexes. With a few notable exceptions,26–31 however, organometallic complexes present challenges that prevent their utilization in technological applications; they use non-earth abundant metals, or carry out proton reduction either at rates much slower than the natural enzymes, or with unfavorable energetics (thus requiring high overpotentials for the reaction), and generally in much harsher conditions (strong acids and=or organic solvents) than the natural enzymes. It is generally thought that such differences between the natural enzymes and the organometallic complexes result from the complexity of the second sphere and outer sphere of coordination in the natural enzyme. For these reasons, artificial model proteins are emerging as bridging ground between organometallic complexes and the natural proteins. Peptide based catalysts present the advantage Biopolymers (Peptide Science)

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of being inherently water soluble. They also allow for the design of second shell coordination sphere and long-range interactions with much ease compared to organometallic complexes. Compared with the natural protein, they can be easily prepared by synthesis or by expression without the need for maturation enzymes. The following sections briefly describe the natural enzymes and organometallic model systems, and discuss work in our lab and in others towards artificial bioinspired hydrogenases. More space is dedicated to the [FeFe] hydrogenase, given the success of peptide-based mimics in catalyzing hydrogen production. We will discuss also models of the electron transfer domain, and recent advances in catalyzing oxygen evolution.

CATALYTIC SITE: HYDROGENASE MIMICS [FeFe] Hydrogenases [FeFe] Hydrogenases are generally very efficient, with turnover numbers as high as 104 H2 molecules per second and are usually biased towards hydrogen production.32 These enzymes function strictly in the absence of oxygen, with exposure resulting in irreversible inhibition.14 Investigation of the mechanism, maturation, and structure of [FeFe] hydrogenases from different organisms revealed a functional organization into dedicated domains, with some domains, variable in type and number, containing a chain of redox active factors that ferry electrons to=from external donors=acceptors to the catalytic site, and associated with the domain that contains the active site.33,34 The [FeFe] hydrogenase found in Desulfovibrio desulfuricans has been characterized by X-ray crystallography, and provides an excellent example of the general architecture of the hydrogenases.17 Its large subunit comprises a ferredoxin-like domain that contains two [4Fe4S] clusters and functions as electron conduit to rapidly transfer the two electrons involved in the reaction, and a neighboring domain that contains the catalytic site, termed the H-cluster. The assembly in vivo of the H-cluster requires a number of specific maturation enzymes.33,35 The H-cluster is composed by a [4Fe4S] cluster and by a [FeFe] site coordinated by a non-proteinogenic dithiolate bridging ligand as well as carbon monoxide (CO) and cyanide (CN) ligands, with the [FeFe] site anchored to the protein through a cysteine (Cys) that bridges the iron-sulfur cluster and one iron, designated as proximal (Figure 1); depending on the redox states sampled during the catalytic cycle, one of the CO ligands bridges the two metals.16,17,36 The central atom in the bridging thiolate ligand has been proposed to be nitrogen, and to be instrumental in assisting proton transfer.37 Cleavage of the H-H bond has been shown to proceed heterolytically,

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FIGURE 1 Structure of [FeFe] hydrogenase from Desulfovibrio desulfuricans (PDB 1HFE; Reproduced from Ref. 17). The ferrodoxin-like domain and the H-cluster are highlighted on the left and on the right, respectively. All molecular figures were created with PyMOL [DeLano, W.L. (2002). The PyMOL Molecular Graphics System. (http:==www.pymol.org)].

suggesting that hydrogen production proceeds through an hydride intermediate, postulated to be located on the distal iron.38–40 The unusual CN and CO ligands play an important role in tuning the electronic properties of the diiron states, and in assisting the transfer of electrons to=from hydrogen.39,41–43 During the catalytic cycle, the distal iron is thought to undergo a conformational change to assume an inverted square pyramidal conformation, or “rotated” state, which is poised for transfer of electrons through the proximal iron to a coordinated proton.39,44 The protein matrix is thought to stabilize the catalytically active “rotated” conformation of the distal iron and to facilitate the reaction by rapidly transferring electrons and a proton to the active site.27,33,44 Phylogenetic analysis of over 800 putative hydrogenase sequences revealed that the cysteines involved in coordinating the cubane-type [4Fe4S] cluster and more broadly the hydrophobic environment surrounding the active site are highly conserved.17,40,45,46An analysis of the high-resolution X-ray structures of two [FeFe] hydrogenases reveals additional electrostatic interactions with the CO and CN ligands, in particular a hydrogen bond to a conserved lysine, and a proposed bond between conserved Cys and methionine and the central nitrogen atom in the bridging dithiolate ligand. These latter interactions are thought to modulate the basicity of the central amine and affect its protonshuffling function during the catalytic cycle.40 Mutations of these residues have been shown to abolish the catalytic activity.45 These second sphere interactions participate directly in assisting the catalysis, and thus need to be considered in the design of hydrogenase mimics. The natural hydrogenases, however, necessitate other accessory functions in order to carry out their function in the context of the living organism.33,47 Beyond the immediate environment of the H-cluster, other conserved residues are thought to participate in the proton=H2 transfer pathway or assist electron transfer through stabiliza-

tion of the accessory clusters; these aspects are beyond the scope of this review; for a thorough discussion, see Ref. 40. Organometallic Models. Structural mimics of the [FeFe] hydrogenase were first developed based on a (l-S(CH2)3S)[Fe2(CO)6] complex, whose similarity to the enzyme’s active site was initially recognized by Pickett, Darensbourg, and Rauchfuss (Figure 2).48–50 These mimics revealed features such as the need for a thermodynamically disfavored “rotated” state of the distal iron in the diiron cluster, which frees a position for catalysis, and electron-donating ligands on the same iron atoms; further second-sphere requirements include a pendant base that delivers protons to the active site, and a redox active group to ferry electrons during catalysis.44,51–60 A recent, elegant design by the Rauchfuss group utilizes an electron-rich chelating phosphine ligand on the distal iron to lock the complex at the “rotated” state, and a ferrocenyl moiety on the proximal iron as electron relay.27

Peptide-Based Structural Models: [FeFe]Hydrogenase A typical approach to the de novo design of metalloproteins utilizes the primary coordination sphere of the metal in natural

FIGURE 2 Schematic representation of (a) the H-cluster of [FeFe] hydrogenases; the X in Figure 1a is likely to be NH=NH21; (b) a minimalist structural mimic used as basis for the development of organometallic complexes (Reproduced from Ref.48–50).

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FIGURE 3 Schematic representation of structural models of [FeFe] hydrogenases: (a) Boc-protected L-Cys methyl ester forming the diiron hexacarbonyl complex developed by Sun and coworkers (Reproduced from Ref. 62); (b) Fe(C5H4CO–Cys–OMe)2 di-peptide employed by Metzler-Nolte and coworkers as a chelate for an iron– carbonyl complex (Reproduced from Ref. 63).

proteins, and builds a peptide scaffold around it.61 Hydrogenases, however, present a challenge because the catalytic site is formed by nonproteinogenic ligands, and is linked to the protein through a single Cys shared with the cubane cluster component of the H-cluster. The need for alternative ways of anchoring the active site to the peptide scaffold was recognized early on, and initial work focused on organometallic models of the [FeFe] hydrogenase based on a (l-S-(CH2)3-S)[Fe2(CO)6] center, which preserves the dithiolate ligation observed in the natural enzyme. Sun and colleagues first observed that cysteinyl ligands could replace the bridging l-S-(CH2)3-S ligands.62 They reported that Boc-protected L-Cys methyl ester forms the desired diiron hexacarbonyl complex 1 (Figure 3) upon treatment with an inorganic precursor, [Fe3(CO)12]. Upon refluxing in various solvents, though, the initial complex undergoes an intramolecular cyclization to generate a chiral two-carbon-bridged dithiolate diiron complex, with structural parameters similar to (l-S-(CH2)3-S)[Fe2(CO)6]. This work highlighted the need for structural preorganization of the cysteinyl ligands in a rigid scaffold. A clever solution to the problem utilizes a ferrocene moiety (dicyclopentadienyl iron, Cp2Fe) tethered to two Cys molecules via peptide bonds to present thiolates at the appropriate distance to form a diiron hexacarbonyl complex.63–65 In this design, the ferrocene group serves as structural scaffold to the thiolates, and also provides a nearby redox-active center: however, no electronic interactions were observed between the iron–carbonyl and the ferrocene cores. The design of cysteinyl ligands within secondary structure motifs of peptide and protein scaffolds is also conductive to preorganization of the thiolate ligands at distances compatible with the formation of a diiron active site. Initial work used CXXC motifs within a helical model peptide, so that the cysteinyl ligands are presented at i, i 1 3 positions on the same face of the helix.66 The design of the hydrogenase maquette, SynHyd1, is based on Baldwin’s alanine-rich helical peptides. Reaction with an inorganic precursor resulted in the formation Biopolymers (Peptide Science)

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of a structurally intact diiron hexacarbonyl cluster, as shown by UV–vis and FTIR spectroscopies, albeit no activity was reported for Fe2SynHyd1. Incorporation of the cluster was well tolerated by the peptide, which conserved a high helical content. The peptide component serves the dual function of preorganizing the cysteinyl ligands as well as providing water solubility to the complex. The approach was successfully extended to functional models in which the diiron hexacarbonyl cluster is tethered within natural proteins by exploiting existing CXXC motifs. Hayashi and colleagues used apo cytochrome c, which contains unique Cys at position 14 and 17.67 Remarkably, the diiron hexacarbonyl-functionalized apo Cyt c catalyzes efficiently the reduction of protons to hydrogen in aqueous solutions at mild pH (4.7), in the presence of Ru(bpy)3 as photosensitizer, and ascorbate as sacrificial electron donor; turnover numbers of 80 are observed over 2 h. Similarly to apo Cyt c, though, the reconstituted H cluster-apo Cyt c appears only partially folded. In a helical protein, formation of the diiron hexacarbonyl center through a CXXC motif imposes a strain on the structure. The energetically favored rotamers of Cys in an a-helix place the sulfur atoms at a distance of 6 A˚, considerably larger than the distance of 3.2 A˚ measured between the thiols in the hydrogenase bridging dithiolate ligand. Relaxing the requirement for structure at the backbone level opens up new avenues: in a follow-up work, the Hayashi group used the C-terminal sequence of cytochrome c556, which contains both a CXXC motif and a His moiety in close proximity, to preorganize a diiron hexacarbonyl moiety close to a coordinatively attached ruthenium photosensitizer (Figure 4).68 The 18-residue peptide is unfolded both in the apo state and in the functionalized state. Even though the reported turnover numbers for photocatalytic hydrogen production are low compared with other models, this design elegantly demonstrates intramolecular electron transfer between the photosensitizer and the catalytic site:

FIGURE 4 Schematic representation of the intramolecular photochemical system developed by Hayashi and coworkers: a Fe2(CO)6 cluster and a photoactive Ru(bpy)(tpy) complex (bpy 5 2,20 -bipyridine, tpy 5 2,20 :60 ,200 -terpyridine) are attached to a peptide fragment of cyt c556 containing the native CXXCH sequence (Reproduced from Ref. 68).

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a control, in which the photosensitizer is not attached to the peptide, but added externally, is inactive in the conditions tested. Conversely, the inorganic cluster can impart desired structural constrains to a peptide. For example, coordination to diiron hexacarbonyl was shown to replace a disulfide bond in the cyclic peptide octreotide, (Sandostatin), preserving structural integrity and retaining much of the binding affinity for the somastotin receptors.69 Preliminary studies show that the octeotride construct retains the active site’s ability to catalyze electrochemical proton reduction, albeit in DMF and with efficiently lower than other inorganic analogues. One possible approach to overcome negative constraints on peptide backbone structure uses unnatural amino acids to anchor the cluster to a peptide scaffold. Roy et al. used an onresin protocol to derivatized a Lys with a propanedithiol unit, and demonstrated incorporation of the cluster in a simple model heptapeptide; no activity was reported.70 The synthetic strategy used, however, poses restrictions in the use of the amino acid: the side chain of the lysine-propanedithiolate adduct is polar, long, and flexible, properties that preclude its incorporation within a structured scaffold and leave the cluster fully exposed to the solvent. Our group has pioneered the use of an artificial amino acid containing a 1,3-propanedithiol moiety, Dt, which shows helical propensity and polarity similar to Leu.71 As a proof of concept design, we used Dt to tether the hydrogenase diiron cluster within a helical peptide (Figure 5). The approach is general, as the Dt amino acid can be incorporated into any peptide sequence, regardless of its fold, and thus allows for the design of specific interactions with the diiron cluster. A similar amino acid containing a 1,2-dithiolane moiety was first synthesized in Boc-protected form by Morera et al. with the intent of using it as switchable redox module,72 and later used in its reduced 1,3-dithiol form by Apfel et al. to reconstitute a diiron hexacarbonyl cluster.73 In this latter work, the investigators used the isolated, Boc-protected form of two related amino acids; while reconstitution of the cluster was demonstrated, limited catalytic activity was observed by cyclic voltammetry. Attempts to use the Boc-deprotected version of the amino acid to use the backbone amine moiety, although, resulted in decomposition of the complex. Building on these concepts, we synthesized an Fmoc-compatible version of the amino acid to incorporate it into more complex peptides by solid phase peptide synthesis by swapping Boc with Fmoc, and changing the dithiolane S-S bond to the acetamidomethyl protecting group on the two thiols. As a proof of concept we designed peptide 1 (Figure 5), based on the alanine-rich peptides developed by Baldwin and coworkers,74 which serve as excellent background to evaluate the effect on alpha helix formation of natural and

FIGURE 5 Computer model of peptide 1-[Fe2(CO)6] developed by Ghirlanda and coworkers. A diiron hexacarbonyl cluster is tethered to a model helical peptide via an artificial dithiol amino acid (Reproduced from Ref. 71).

unnatural amino acids. Placing the Dt amino acid at position 16 in the sequence facilitates i, i 1 3 interactions with the Cterminal Lys 19. The [2Fe2S] cluster was incorporated into purified peptide 1 by direct reaction with Fe3(CO)12 under strictly anaerobic conditions48,75 to reconstruct the diiron containing active site of natural hydrogenases. We assessed the effect of Dt on helix formation and showed that introduction of the cluster stabilized the helix. The complex, 1-[Fe2(CO)6], catalyzes photoinduced hydrogen production in the presence of a photosensitizer and a sacrificial reducing agent in water with turnover numbers (TON) of about 85 after 2 h. Further investigation by cyclic voltammetry showed that, in analogy to related inorganic complexes, 1[Fe2(CO)6] catalyzes hydrogen production in a pH dependent manner, and with the advantage of operating in aqueous solvent. Having demonstrated the feasibility of this approach, we are now investigating the use of more elaborate organometallic models built around the bridging thiolate ligand and inspired by the numerous examples found in the literature, with the ultimate goal of recreating the asymmetric substitution and division of function in the diiron site found in the [FeFe] hydrogenase. Possible examples include exchanging one-two CO ligands with CN, or introducing chelating phosphine ligands.49,76,77 A second research thrust focuses on the design of more elaborate peptide-based structures containing the Dt Biopolymers (Peptide Science)

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FIGURE 6 Structure of [NiFe] hydrogenase from Desulfovibrio vulgaris (PDB 1WUJ) (Reproduced from Ref. 85), with the small subunit (green) showing a chain of iron-sulfur clusters and the large subunit (gray) showing the bimetallic center.

amino acid as anchor for hydrogenase mimics to investigate the effect of second sphere interactions on the activity of the diiron center. Designed electrostatic interactions replicating those observed in the hydrogenase (see above) could assist in stabilizing asymmetrically substituted derivatives of the diiron hexacarbonyl center. The placement of basic residues such as lysine or histidine in the proximity of the catalytic site could assist in proton transfer.

[NiFe] Hydrogenases [NiFe] hydrogenases comprise a large number of enzymes, with variable numbers of subunits and function spanning “uptake,” bidirectional, and hydrogen evolving hydrogenases; based on sequence analysis they can be divided phylogenetically in four groups.6 Although generally 10–100 times less efficient than [FeFe] hydrogenases, [NiFe] hydrogenases presents properties that make them attractive in applications; in contrast with [FeFe] hydrogenases, oxygen inhibition is generally reversible in [NiFe] hydrogenases, and some even function in the presence of oxygen.13,14,54,78–84 The first structures, belonging to periplasmic uptake hydrogenases found in bacteria, revealed a heterodimer composed by a large subunit containing the active site, and a second, smaller electron-transfer subunit containing one [3Fe4S] and two [4Fe4S] clusters (Figure 6).15,19 The heterobimetallic catalytic site contains one Ni(II) coordinated in a distorted square planar geometry by four cysteinyl ligands, provided by two conserved CXXC motifs; two cysteines bridge over to an iron Biopolymers (Peptide Science)

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atom (Figure 7a). The first coordination sphere of the iron atom is reminiscent of [FeFe] hydrogenases, and contains two cyanides and one carbonyl as ligands. The enzyme is found in several “states,” some of which are thought to be artifacts of purification54 and which have been characterized spectroscopically.85,86 Despite much effort, the mechanism of hydrogen oxidation is still under debate.54,87 The availability of several X-ray structures of the [NiFe] hydrogenases spurred the development of a large number of structural mimics that replicated the bimetallic site of the enzyme. Recent reviews narrate the progress towards catalytically active models;54,88,89 we will summarize here issues that will be relevant to the design of peptide-based catalysts. Briefly, early structural mimics of the [NiFe] enzyme were not catalytically active.90–92 The introduction of ligands capable of modulating the electronic properties of the iron atom resulted in the development of several functional bimetallic complexes, which mimic the mechanism of the natural [NiFe] hydrogenase by using a bridging hydride intermediate between the two metals. Recently, the Ogo group reported a nickel-iron complex that replicates the bidirectionality of the [NiFe] hydrogenase.28 Peptide-Based Structural Models: [NiFe]-Hydrogenase. Initial work in the Darensburg laboratory explored the use of a tripeptide with the sequence CGC, derived from acetyl-coA synthetase, to coordinate nickel (Figure 7b).93 In this case, the primary coordination sphere of the nickel is N2S2, where the N is provided by the backbone amide of Gly and Cys. Due to the electronic and steric properties of the sulfur ligand, the Ni(CGC)22 complex functions as a synthon for the formation of heterobimetallic species of the general formula [Ni(CGC)]M(CO)x, in which M can be tungsten or rhodium. The complexes were prepared both in solution and on TentaGel resin, demonstrating stabilization to complex aggregation on solid state support.93 Dutta et al.94 expanded on the concept by utilizing the N-terminal of nickel superoxide dismutase,

FIGURE 7 Schematic representation of (a) the active site of [NiFe] hydrogenases; (b) the heterobimetallic complex [Ni(CGC)][W(CO)4]22 developed by Darensbourg and coworkers (Reproduced from Ref. 93).

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which can coordinate nickel and retains the enzyme’s activity,95 to preorganize the first coordination sphere around the nickel in heterometallic complexes. Reaction of existing Cys ligands with labile ligands of heterometallic fragments resulted in the synthesis of two [NiFe] hydrogenase mimics containing NiRu and NiFe sites. These peptides were soluble in water and displayed spectral signatures similar to corresponding inorganic complexes.94 Even though activity has yet to be reported for mixed-metal complexes anchored to a peptide via a Ni coordination site, the approach can be used to screen rapidly a number of heterobimetallic sites formed via sharing of the thiolate ligands.

Functional Models of Hydrogenase A complementary approach to the design of hydrogenase mimics builds on so-called functional mimics of the enzymes, defined as organometallic complexes capable to carry out hydrogen production reversibly via a different catalytic site. Notable in this area are the Ni-phosphine complexes developed by DuBois,96–101 the Fe-phosphine complex recently presented by Bullock and DuBois,26 and the Ni-thiolate, the Codithiolene and the cobaloxime complexes developed by Artero, Eisenberg, and Holland.102–109 Recent reviews summarize these achievements.89,110 Although the existing literature on these complexes is extensive, very little work has been done to integrate them into peptide scaffolds. Most notably, DuBois and Shaw explored the use of the DuBois catalysts, consisting of mononuclear organometallic complexes based on nickel that contain chelating phosphine ligands and a bridging pendant tertiary amine that serves as proton donor with the general formula [Ni(PPh2NR2)2]21; an overview of this work can be found in Ref. 101.101,111–114 Aiming to introduce second coordination and outer coordination sphere interactions, the investigators functionalized the four pendant amines in the complex with amino acids and peptides. The synthetic protocol was modified by forming the cyclic phosphine ligands starting with functionalized precursor amines, and results in identical amino acids displayed onto each of the four amines in the final complex. The prototype glycylglycine adduct shows turnover frequencies (TOF) in the 200–1400 s21 range, depending on substitution, in the presence of water; for comparison the unsubstituted [Ni(PPh2NPh2)2]21 displays TOF of 700 s21 in similar conditions. Systematic analysis of the results showed that, surprisingly, polar amino acids had little or no effect on catalytic activity. The presence of charged side chains, however, modulates the activity of the complex by one order of magnitude, with Glu and Lys increasing activity the most.97,101 Further development of these models in structured scaffolds will need

to address the current synthetic scheme, which results in symmetrical substitution of the complexes, as well as the dynamics of the complex during the catalytic cycle.

Electron-Transfer Modules Hydrogenases generally contain an electron-transfer module containing iron-sulfur clusters in proximity of their redoxactive catalytic site. These cofactors are also found in other complex catalytic machinery such as hydrogenases, nitrogenases, and photosystem I (PSI), making them highly relevant in designing proteins for fuel production. A number of different groups have made recent advances in the incorporation of these electron transfer modules into peptides. Iron-Sulfur Clusters Iron-sulfur clusters are found in a wide range of forms in nature, including [2Fe2S], [3Fe4S], [4Fe4S] and higher order clusters; of these, [4Fe4S] cubane-type clusters are the most abundant.115 These moieties are formed in proteins via coordination of iron through Cys side chains with bridging inorganic sulfur atoms. In many biological systems, [4Fe4S] clusters are found aligned to create an electron conduit that shuffles electrons to and from the active site. Along with their ability to function as electron transfer modules, iron-sulfur clusters also play a role in other processes, such as catalysis, iron level regulation, and storage, as well as transport of ligands within the cell; they are also used in the multielectron chemistry of proton reduction, sulfite reduction, and elemental nitrogen reduction.116,117 Here, we will focus on the use of iron-sulfur clusters for the purposes of electron transfer. Despite containing isostructural [4Fe4S] clusters, ironsulfur proteins exhibit redox potentials ranging from 2700 to 1450 mV, and have been shown to use three different redox states: [4Fe4S]31=21=11.118 It is believed that the electrostatic environment of amino acids surrounding the cluster, the hydrogen bonding interactions of bridging S-atoms, and the accessibility of the cluster to the solvent influence the potential observed for a particular cluster in a given context.116 Due to their inherent structural complexity and to the necessity of a proteinogenic environment for existence in nature, a substantial amount of scientific effort has been dedicated to elucidating the structure-function relationship of iron-sulfur proteins in natural as well as de novo designed proteins.119–121 Work in natural proteins has not been able to elucidate fully the relationship between structure and iron-sulfur clusters redox potentials. Point mutagenesis work on the well studied ferredoxin proteins and the unique Rieske cluster has given some insight into the mechanisms by which nature has tuned iron-sulfur cluster redox potentials to particular functions.119,121,122 Generally, it has been seen that introduction of Biopolymers (Peptide Science)

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positive charges into the proximity of the iron-sulfur clusters shifts the redox potential more positive, while the opposite holds true for introduction of negative charges. While the charges in proximity to each cluster can greatly affect the redox potential, it is also believed that hydrogen bonding to the polar peptide backbone significantly alters the apparent redox potentials.121,123 It has been suggested that it is this hydrogen bonding that differentiates the two types of [4Fe4S] clusters, those that employ a 13 to 12 transition, and those that employ a 12 to 11 transition (HiPiP vs Ferredoxin clusters), as the more hydrogen bonds to Ferredoxin clusters allow for stabilization of higher electron density.124,125 Although natural systems have lead to these general a priori trends, natural systems are limited in the information that can be obtained from them, as often multiple variables are changed at once. De novo design offers reliable systems to study these trends, to determine the contribution of each of these components in determining the redox potential of iron-sulfur clusters. Iron-sulfur cluster binding sites have been incorporated into proteins that do not naturally bind this cofactor by using computational methods,126 or reconstituted into minimalistic sequence motifs derived from natural proteins. Early work on [4Fe4S] binding peptides was based on a conserved CXXCXXC sequence motif found in native cluster binding proteins such as the ferredoxins. It was shown that these peptides can generally incorporate iron sulfur clusters via in situ incorporation, in which the cluster is believed to form from inorganic precursors with b-mercaptoethanol ligands followed by an entropically driven exchange with the peptide, which acts as a chelating agent and releases the b-mercaptoethanol ligands. These early designs were reviewed recently.118 Briefly, sequence alignment studies, confirmed by experimental characterization, narrowed the consensus sequence to CIACGAC.127,128 In the quest for increased structural and functional complexity, sequences adapted from the minimalistic motif CXXCXXC have been used as loops in helical hairpins. The Dutton laboratory engineered heme binding maquettes to design a helixloop-helix motif that coordinates two hemes and one ironsulfur cluster, and suggested that this motif dimerized into a four-helix bundle. The redox potential of the cluster, determined via spectroelectrochemical experiments, was 2350 mV, which is comparable to that of natural ferredoxins; it appears that the cluster’s properties are not affected by the neighboring heme. A few possible models were put forward based on sequence information, and it is quite likely that the protein is a molten globule; no evidence was found for coupling between cofactors.129 This concept was adapted to reconstruct bridged metal assemblies based on the A-cluster of carbon monoxide dehyBiopolymers (Peptide Science)

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drogenase, which contains a cubane iron-sulfur cluster bridged to a Ni(II) site.130 Using the ferredoxin consensus sequence, the Holm laboratory designed a series of four peptides that explored different ligand coordination spheres around the nickel. All the peptides bound a preformed cubane cluster with apparent 1:1 stoichiometry, and two of the peptides also bound nickel; it was not possible though to infer structural information from the available spectroscopic data. Other efforts have focused on modeling the [4Fe4S] clusters found in PSI, which contains three clusters, deemed FX, FA, and FB. The Fx cluster is characterized by a pseudosymmetric arrangement in which the Cys ligands are contributed in pairs by the two subunits of the PsaA=PsaB heterodimer. Scott et al.131 inserted the 10-amino acid consensus sequence CDGPGRGGTC as loops in helical hairpins designed to pair up into a four-helix bundle, so that the two loops coordinated a cubane cluster at one end of the bundle. The resulting protein displays a redox potential of 2422 mV, reflecting the solvent exposed location of the cluster. In contrast, models of the FA and Fb clusters had remained elusive. Utilizing the consensus binding sequence of CxxCxxCxxxCP found in PSI, Lubitz et al. modeled the individual binding sites of both FA and FB onto 16 amino acid peptides.132,133 After cluster incorporation, the peptides showed similar redox potentials to the natural clusters, at 2440 and 2470 mV, and also irreversibly bound to PSI. This approach has been extended recently to the incorporation of the less abundant Fe3S4 cluster through mutation of a single, ligating Cys residue to a Ser; further mutations in the variable portion of the consensus sequence were shown to modulate the efficiency of cluster incorporation as well as the relative preference for [4Fe4S] vs. [3Fe4S].120 Recently, a computational methodology that starts from the geometric requirements of the metal was used to design a protein that incorporates an iron-sulfur cluster.134,135 The natural symmetry of [4Fe4S] clusters was extended to the four Cys making up the first coordination sphere, and then a four-helix bundle was built around the site (Figure 8). Notably, incorporation of the cluster into their artificial protein stabilizes the overall fold, leading to the conclusion that the designed ahelices are integral in cluster incorporation. The authors subsequently added to this peptide an empirically designed heme binding site. The new model successfully bound both the cluster and heme, although the latter with very low affinity. Throughout this series, although, oligomerization of the helical peptides was a large problem, affecting the functionality of the peptides. The model systems reviewed above were designed to contain a single cubane-type cluster. In nature, however, clusters are often organized in pairs or chains that can be used as

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seen for other clusters. It has been proposed that these clusters are capable of storing two electrons, via a proton-coupled electron transfer mechanism.139 Upon reduction of the cluster, the protein undergoes a conformational change during deprotonation of a backbone amide and removal of the bridging Cys residue.139 This two electron chemistry is of great interest to de novo peptide design for electron transfer, but has yet to be accomplished in a rationally designed peptide.

Toward Artificial Photosynthetic Systems: Powering Fuel Production

FIGURE 8 Computational model of a de novo designed fourhelix bundle coordinating a [4Fe4S] cluster (Reproduced from Ref. 135). Image courtesy of V. Nanda.

electron conduits. Towards the goal of generating an artificial molecular wire, our lab has recently designed a domainswapped dimer peptide that incorporates two [4Fe4S] clusters at the same time.136 Using the crystallographic coordinates of an existing de novo designed peptide as scaffold, we docked the primary coordination sphere of a natural iron-sulfur cluster into the hydrophobic core, mutating existing Leu to Cys. The dimeric nature of the design results in a symmetric coordination site reminiscent of the two-cluster bacterial ferredoxins (Figure 9). The peptide readily incorporates two clusters per dimer, and maintains high helical content as well as stability to thermal denaturation. EPR characterization demonstrated the incorporation of cubane-like iron-sulfur clusters. The presence of two clusters per dimer was verified using pulsed ELDOR techniques, which showed coupling between the clusters consistent with the expected distance of about 35 A˚. This is the first example of an artificial peptide that incorporates two clusters at once. Current work is focusing on incorporation of different clusters, as well as their positioning at closer reciprocal distances inside the structure, with the aim of obtaining electronically coupled metal centers, which could potentially act as a molecular wire. Recently, a new type of iron-sulfur cluster was found in the oxygen-tolerant, membrane-bound NiFe hydrogenases.137,138 These proximal [4Fe3S] clusters are ligated into the protein via 6 Cys residues, as opposed to the typical 4 or 3 Cys residues

A connection between solar power and fuel production is ultimately necessary to establish sustainability. Higher plants, algae and cyanobacteria use energy from sunlight to oxidize water into O2, protons and electrons through photosynthesis, a process powered by a unique pigment=protein complex called Photosystem II (PSII).140–143 Its essential elements are: (i) a strongly oxidizing multichlorophyll complex termed P680; (ii) a redox-active tyrosine termed YZ; (iii) a bound plastoquinone electron acceptor; and (iv) a metalloprotein containing a manganese, calcium cluster called oxygen evolving center (OEC). The structure of OEC, which is the heart of the wateroxidizing machinery of photosynthesis, has been recently resolved at 1.98 A˚ by Umena and coworkers, and can be

FIGURE 9 Computational model of DSD-[4Fe4S]2, a de novo designed dimeric three-helix bundle protein that coordinates two iron sulfur clusters within its hydrophobic core (Ref. 136).

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FIGURE 10 Structure of one dimeric subunit of E. coli BFR. This subunit has been used by Wydrzynski and coworkers to mimic the water oxidation centre (WOC) photoassembly. In the design, heme was replaced with the photoactive zinc–chlorin e6 (ZnCe6) (highlighted in the blue box), and the di-iron binding site with two manganese ions (highlighted in the magenta box). Upon illumination, the bound ZnCe6 initiates electron transfer, with concomitant oxidization of MnII to MnIII via a tyrosine radical formed during the process (Reproduced from Ref. 156).

described as a Mn4O5-Ca(H2O)4 complex bound within the protein center.144 Numerous small inorganic catalysts,145–149 nanomaterials,150–153 and molecular devices obtained by linking together pigments, quinones, amino acids, and metal complexes have been synthesized with the aim of oxidizing water through a biomimetic mechanism.3,154,155 However, some of these systems have shown low efficiencies and stability due to rapid back reactions and thermodynamic losses, and medium-tohigh operating overpotential. The balance between photon absorption, charge separation, and electron transfer events found in the natural PSII complex remains unmatched. This complexity (32 cofactors and more than 27 protein subunits), however, renders direct PSII application toward fuel production impractical. These challenges could be addressed by a miniaturization approach through de novo designed proteins although mimicking all the features of the water oxidase into a de novo designed multimeric structure presents its own challenges. To date, the only example of an artificial photoactive PSII model was put forward by Wydrzynski and coworkers, who used bacterioferritin (BFR) from Escherichia coli as protein scaffold to demonstrate photoinduced oxidation of a dimanganese site.156,157 BFR is particularly suitable for the Biopolymers (Peptide Science)

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design process because it tolerates mutations well, in this case allowing multiple cofactor incorporation at distances similar to those found in PSII. The authors extracted and replaced the naturally occurring heme with a photoactive chlorin,158 and replaced the diiron site with a dimanganese site able to reach high oxidation states, with concomitant oxygen consumption in presence of light; similarly to PSII, a tyrosine side chain acts as redox mediator in the electron transfer processes (Figure 10).156 While this model is far from the desired reactivity, it is noteworthy for the multifunction, multiunit approach followed. More recently, Meyer and coworkers159 have developed a water oxidation catalyst based on a triglycylglycine macrocyclic ligand (TGG4–) complex of Cu(II). [(TGG4–)CuII–OH2]2– efficiently catalyzes water oxidation in phosphate buffer at pH 11 at room temperature, with TOF of 33 s21 for oxygen evolution. In the proposed active specie Cu(II) is coordinated by three amide nitrogen and one primary amine (the N terminal) in a plane, with the C terminal carboxylate coordinating in an axial position; the second axial position is taken up by the water substrate. This system is impressive for its ease of synthesis, stability, and water solubility, although the reaction occurs at an overpotential of 0.52 V. Nevertheless, these properties may be optimized in second-generation catalysts that will include first (and second) shell interactions.

CONCLUSIONS Since the first de novo designed proteins were reported 25 years ago, the field has advanced tremendously. This progress is mainly tied to the continuing optimization of computational algorithms and methods, which have emerged as a reliable approach to the design of protein structures and functions.61,160–165 Indeed, relatively simple principles have been established for the development of stable and well-defined proteins,166–168 metalloproteins,169,170 and membrane proteins,171– 174 paving the way for the design of functional proteins.175,176 Recent success stories include the design of transferases,177 hydrolases,178–180 and oxidoreductases.181–183 Building on these achievements, the field is now moving towards end-use inspired applications, with energy production emerging as one of the societal needs. We reviewed here recent work inspired by natural proteins, in particular iron-sulfur cluster proteins and hydrogenases, and work that incorporates lessons learned through traditional organometallic approaches. Looking forward, the combination of organometallic chemistry and protein science holds special promise in the development of new protein-based catalysts for the sustainable production of fuels.

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The authors are supported by the Center for Bio-Inspired Solar Fuel Production, an Energy Frontier Research Center. MF wishes to thank L’Oreal Foundation for partial support.

REFERENCES 1. Cammack, R.; Frey, M.; Robson, R. Hydrogen As a Fuel: Learning From Nature; Taylor and Francis: London, 2001. 2. Amao, Y. ChemCatChem 2011, 3, 458–474. 3. Gust, D.; Moore, T. A.; Moore, A. L. Faraday Discuss 2012, 155, 9–26. 4. Gust, D.; Moore, T. A.; Moore, A. L. Acc Chem Res 2009, 42, 1890–1898. 5. Wydrzynski, T. J.; Hillier, W. Molecular Solar Fuels. RCS: Cambridge, UK, 2009. 6. Vignais, P. M.; Billoud, B. Chem Rev 2007, 107, 4206–4272. 7. Vignais, P. M.; Billoud, B.; Meyer, J. FEMS Microbiol Rev 2001, 25, 455–501. 8. Meyer, J. Cell Mol Life Sci 2007, 64, 1063–1084. 9. Shima, S.; Ermler, U. Eur J Inorg Chem 2011, 2011, 963–972. 10. Shima, S.; Thauer, R. K. Chem Rec 2007, 7, 37–46. 11. Fontecilla-Camps, J. C.; Volbeda, A.; Cavazza, C.; Nicolet, Y. Chem Rev 2007, 107, 4273–4303. 12. Vincent, K. A.; Parkin, A.; Armstrong, F. A. Chem Rev 2007, 107, 4366–4413. 13. Frey, M. ChemBioChem 2002, 3, 153–160. 14. De Lacey, A. L.; Fernandez, V. M.; Rousset, M.; Cammack, R. Chem Rev 2007, 107, 4304–4330. 15. Volbeda, A.; Charon, M.-H.; Piras, C.; Hatchikian, E. C.; Frey, M.; Fontecilla-Camps, J. C. Nature 1995, 373, 580–587. 16. Peters, J. W.; Lanzilotta, W. N.; Lemon, B. J.; Seefeldt, L. C. Science 1998, 282, 1853–1858. 17. Nicolet, Y.; Piras, C.; Legrand, P.; Hatchikian, C. E.; FontecillaCamps, J. C. Structure 1999, 7, 13–23. 18. Nicolet, Y.; de Lacey, A. L.; Verne`de, X.; Fernandez, V. M.; Hatchikian, E. C.; Fontecilla-Camps, J. C. J Am Chem Soc 2001, 123, 1596–1601. 19. Happe, R. P.; Roseboom, W.; Pierik, A. J.; Albracht, S. P. J.; Bagley, K. A. Nature 1997, 385, 126–126. 20. Bagley, K. A.; Duin, E. C.; Roseboom, W.; Albracht, S. P. J.; Woodruff, W. H. Biochemistry 1995, 34, 5527–5535. 21. Pierik, A. J.; Hulstein, M.; Hagen, W. R.; Albracht, S. P. J. Eur J Biochem 1998, 258, 572–578. 22. Hoffmann, P. Tomorrow’s Energy: Hydrogen, Fuel Cells, and Prospects for a Cleaner Planet; MIT Press: Cambridge, MA, 2012. 23. Bingham, A. S.; Smith, P. R.; Swartz, J. R. Int J Hydrogen Energy 2012, 37, 2965–2976. 24. Stapleton, J. A.; Swartz, J. R. PLoS One 2010, 5, 1–8. 25. Stapleton, J. A.; Swartz, J. R. PLoS One 2010, 5, e10554. 26. Liu, T.; DuBois, D. L.; Bullock, R. M. Nat Chem 2013, 5, 228– 233. 27. Camara, J. M.; Rauchfuss, T. B. Nat Chem 2012, 4, 26–30. 28. Ogo, S.; Ichikawa, K.; Kishima, T.; Matsumoto, T.; Nakai, H.; Kusaka, K.; Ohhara, T. Science 2013, 339, 682–684. 29. Wilson, A. D.; Newell, R. H.; McNevin, M. J.; Muckerman, J. T.; Rakowski DuBois, M.; DuBois, D. L. J Am Chem Soc 2005, 128, 358–366.

30. Smith, S. E.; Yang, J. Y.; DuBois, D. L.; Bullock, R. M. Angew Chem 2012, 124, 3206–3209. 31. Le Goff, A.; Artero, V.; Jousselme, B.; Tran, P. D.; Guillet, N.; Metaye, R.; Fihri, A.; Palacin, S.; Fontecave, M. Science 2009, 326, 1384–1387. 32. Cammack, R. Nature 1999, 397, 214–215. 33. Mulder, D. W.; Shepard, E. M.; Meuser, J. E.; Joshi, N.; King, P. W.; Posewitz, M. C.; Broderick, J. B.; Peters, J. W. Structure 2011, 19, 1038–1052. 34. Peters, J. W.; Broderick, J. B. Annu Rev Biochem 2012, 81, 429–450. 35. B€ ock, A.; King, P. W.; Blokesch, M.; Posewitz, M. C. In Adv Microb Physiol; K. P. Robert, Ed.; Academic Press: New York, 2006; p 1–225. 36. Nicolet, Y.; Lemon, B. J.; Fontecilla-Camps, J. C.; Peters, J. W. Trends Biochem Sci 2000, 25, 138–143. 37. Lemon, B. J.; Peters, J. W. Biochemistry 1999, 38, 12969– 12973. 38. Lespinat, P. A.; Berlier, Y.; Fauque, G.; Czechowski, M.; Dimon, B.; Le Gall, J. Biochimie 1986, 68, 55–61. 39. Darensbourg, M. Y.; Lyon, E. J.; Zhao, X.; Georgakaki, I. P. Proc Natl Acad Sci USA 2003, 100, 3683–3688. 40. Winkler, M.; Esselborn, J.; Happe, T. Biochim Biophys Acta, in press. 41. Kubas, G. J. Chem Rev 2007, 107, 4152–4205. 42. Bruschi, M.; Greco, C.; Bertini, L.; Fantucci, P.; Ryde, U.; Gioia, L. D. J Am Chem Soc 2010, 132, 4992–4993. 43. Liu, Z.-P.; Hu, P. J Am Chem Soc 2002, 124, 5175–5182. 44. Darensbourg, M. Y.; Bethel, R. D. Nat Chem 2012, 4, 11–13. 45. Knorzer, P.; Silakov, A.; Foster, C. E.; Armstrong, F. A.; Lubitz, W.; Happe, T. J Biol Chem 2012, 287, 1489–1499. 46. Pandey, A. S.; Harris, T. V.; Giles, L. J.; Peters, J. W.; Szilagyi, R. K. J Am Chem Soc 2008, 130, 4533–4540. 47. Posewitz, M. C.; Mulder, D. W.; Peters, J. W. Curr Chem Biol 2008, 2, 178–199. 48. Lyon, E. J.; Georgakaki, I. P.; Reibenspies, J. H.; Darensbourg, M. Y. Angew Chem Int Ed 1999, 38, 3178–3180. 49. Schmidt, M.; Contakes, S. M.; Rauchfuss, T. B. J Am Chem Soc 1999, 121, 9736–9737. 50. Le Cloirec, A.; C. Davies, S.; J. Evans, D.; L. Hughes, D.; J. Pickett, C.; P. Best, S.; Borg, S. Chem Commun 1999, 2285– 2286. € F.; Schwartz, L.; Stein, M.; Silakov, A.; Kaur51. Erdem, O. Ghumaan, S.; Huang, P.; Ott, S.; Reijerse, E. J.; Lubitz, W. Angew Chem Int Ed 2011, 50, 1439–1443. 52. Singleton, M. L.; Crouthers, D. J.; Duttweiler, R. P., III; Reibenspies, J. H.; Darensbourg, M. Y. Inorg Chem 2011, 50, 5015–5026. 53. Barton, B. E.; Olsen, M. T.; Rauchfuss, T. B. Curr Opin Biotechnol 2010, 21, 292–297. 54. Tard, C. d.; Pickett, C. J. Chem Rev 2009, 109, 2245–2274. 55. Ezzaher, S.; Gogoll, A.; Bruhn, C.; Ott, S. Chem Commun 2010, 46, 5775–5777. 56. Surawatanawong, P.; Tye, J. W.; Darensbourg, M. Y.; Hall, M. B. Dalton Trans 2010, 39, 3093–3104. 57. Wang, F.; Wang, W.-G.; Wang, X.-J.; Wang, H.-Y.; Tung, C.-H.; Wu, L.-Z. Angew Chem Int Ed 2011, 50, 3193–3197.

Biopolymers (Peptide Science)

De Novo Design of Functional Proteins 58. Wang, W.-G.; Wang, F.; Wang, H.-Y.; Tung, C.-H.; Wu, L.-Z. Dalton Trans 2012, 41, 2420–2426. 59. Singleton, M. L.; Reibenspies, J. H.; Darensbourg, M. Y. J Am Chem Soc 2010, 132, 8870–8871. € F.; Harman, S. D.; Singleton, M. L.; 60. Hsieh, C.-H.; Erdem, O. Reijerse, E.; Lubitz, W.; Popescu, C. V.; Reibenspies, J. H.; Brothers, S. M.; Hall, M. B.; Darensbourg, M. Y. J Am Chem Soc 2012, 134, 13089–13102. 61. DeGrado, W. F.; Summa, C. M.; Pavone, V.; Nastri, F.; Lombardi, A. Annu Rev Biochem 1999, 68, 779–819. 62. He, C.; Wang, M.; Zhang, X.; Wang, Z.; Chen, C.; Liu, J.; A˚kermark, B.; Sun, L. Angew Chem 2004, 116, 3655–3658. 63. de Hatten, X.; Bothe, E.; Merz, K.; Huc, I.; Metzler-Nolte, N. Eur J Inorg Chem 2008, 2008, 4530–4537. 64. de Hatten, X.; Cournia, Z.; Huc, I.; Smith, J. C.; Metzler-Nolte, N. Chem-Eur J 2007, 13, 8139–8152. 65. de Hatten, X.; Weyherm€ uller, T.; Metzler-Nolte, N. J Organomet Chem 2004, 689, 4856–4867. 66. Jones, A. K.; Lichtenstein, B. R.; Dutta, A.; Gordon, G.; Dutton, P. L. J Am Chem Soc 2007, 129, 14844–14845. 67. Sano, Y.; Onoda, A.; Hayashi, T. Chem Commun 2011, 47, 8229–8231. 68. Sano, Y.; Onoda, A.; Hayashi, T. J Inorg Biochem 2012, 108, 159–162. 69. Apfel, U.-P.; Rudolph, M.; Apfel, C.; Robl, C.; Langenegger, D.; Hoyer, D.; Jaun, B.; Ebert, M.-O.; Alpermann, T.; Seebach, D.; Weigand, W. Dalton Trans 2010, 39, 3065–3071. 70. Roy, S.; Shinde, S.; Hamilton, G. A.; Hartnett, H. E.; Jones, A. K. Eur J Inorg Chem 2011, 2011, 1050–1055. 71. Roy, A.; Madden, C.; Ghirlanda, G. Chem Commun 2012, 48, 9816–9818. 72. Morera, E.; Pinnen, F.; Lucente, G. Org Lett 2002, 4, 1139– 1142. 73. Apfel, U. P.; Kowol, C. R.; Morera, E.; Gorls, H.; Lucente, G.; Keppler, B. K.; Weigand, W. Eur J Inorg Chem 2010, 5079–5086. 74. Marqusee, S.; Robbins, V. H.; Baldwin, R. L. Proc Natl Acad Sci USA 1989, 86, 5286–5290. 75. Ellgen, P. C.; Gerlach, J. N. Inorg Chem 1973, 12, 2526–2532. 76. Gloaguen, F.; Lawrence, J. D.; Schmidt, M.; Wilson, S. R.; Rauchfuss, T. B. J Am Chem Soc 2001, 123, 12518–12527. 77. Justice, A. K.; Zampella, G.; De Gioia, L.; Rauchfuss, T. B.; van der Vlugt, J. I.; Wilson, S. R. Inorg Chem 2007, 46, 1655–1664. 78. Adams, M. W. W. Biochim Biophys Acta 1990, 1020, 115–145. 79. Shafaat, H. S.; R€ udiger, O.; Ogata, H.; Lubitz, W. Biochim Biophys Acta - Bioenerg 2013, 1827, 986–1002. 80. Abou Hamdan, A.; Liebgott, P.-P.; Fourmond, V.; GutierrezSanz, O.; De Lacey, A. L.; Infossi, P.; Rousset, M.; Dementin, S.; Leger, C. Proc Natl Acad Sci USA 2012, 109, 19916–19921. 81. Abou Hamdan, A.; Dementin, S.; Liebgott, P.-P.; GutierrezSanz, O.; Richaud, P.; De Lacey, A. L.; Rousset, M.; Bertrand, P.; Cournac, L.; Leger, C. J Am Chem Soc 2012, 134, 8368– 8371. 82. Sch€afer, C.; Friedrich, B.; Lenz, O. Appl Environ Microbiol 2013, 79, 5137–5145. 83. Fritsch, J.; Lenz, O.; Friedrich, B. Nat Rev Microbiol 2013, 11, 106–114. 84. Friedrich, B.; Fritsch, J.; Lenz, O. Curr Opin Biotechnol 2011, 22, 358–364.

Biopolymers (Peptide Science)

569

85. Ogata, H.; Hirota, S.; Nakahara, A.; Komori, H.; Shibata, N.; Kato, T.; Kano, K.; Higuchi, Y. Structure 2005, 13, 1635–1642. 86. Pandelia, M.-E.; Ogata, H.; Lubitz, W. ChemPhysChem 2010, 11, 1127–1140. 87. Pardo, A.; Lacey, A.; Fernandez, V.; Fan, H.-J.; Fan, Y.; Hall, M. J Biol Inorg Chem 2006, 11, 286–306. 88. Simmons, T. R.; Artero, V. Angew Chem Int Ed 2013, 52, 6143–6145. 89. Dawson, J.; Ghiotto, F.; McMaster, J.; Schroder, M. In Molecular Solar Fuels; The Royal Society of Chemistry: Cambridge, 2012, p 326–386. 90. Lai, C.-H.; Reibenspies, J. H.; Darensbourg, M. Y. Angew Chem Int Ed 1996, 35, 2390–2393. 91. Canaguier, S.; Artero, V.; Fontecave, M. Dalton Trans 2008, 315–325. 92. Li, Z.; Ohki, Y.; Tatsumi, K. J Am Chem Soc 2005, 127, 8950– 8951. 93. Green, K. N.; Jeffery, S. P.; Reibenspies, J. H.; Darensbourg, M. Y. J Am Chem Soc 2006, 128, 6493–6498. 94. Dutta, A.; Hamilton, G. A.; Hartnett, H. E.; Jones, A. K. Inorg Chem 2012, 51, 9580–9588. 95. Neupane, K. P.; Gearty, K.; Francis, A.; Shearer, J. J Am Chem Soc 2007, 129, 14605–14618. 96. Helm, M. L.; Stewart, M. P.; Bullock, R. M.; DuBois, M. R.; DuBois, D. L. Science 2011, 333, 863–866. 97. Jain, A.; Lense, S.; Linehan, J. C.; Raugei, S.; Cho, H.; DuBois, D. L.; Shaw, W. J. Inorg Chem 2011, 50, 4073–4085. 98. O’Hagan, M.; Ho, M. H.; Yang, J. Y.; Appel, A. M.; Rakowski DuBois, M.; Raugei, S.; Shaw, W. J.; DuBois, D. L.; Bullock, R. M. J Am Chem Soc 2012, 134, 19409–19424. 99. O’Hagan, M.; Shaw, W. J.; Raugei, S.; Chen, S.; Yang, J. Y.; Kilgore, U. J.; DuBois, D. L.; Bullock, R. M. J Am Chem Soc 2011, 133, 14301–14312. 100. Pool, D. H.; Stewart, M. P.; O’Hagan, M.; Shaw, W. J.; Roberts, J. A.; Bullock, R. M.; DuBois, D. L. Proc Natl Acad Sci USA 2012, 109, 15634–15639. 101. Shaw, W. J.; Helm, M. L.; DuBois, D. L. Biochim Biophys ActaBioenerg, 2013, 1827, 1123–1139. 102. Jacques, P.-A.; Artero, V.; Pecaut, J.; Fontecave, M. Proc Natl Acad Sci USA 2009, 106, 20627–20632. 103. Zhang, P.; Jacques, P.-A.; Chavarot-Kerlidou, M.; Wang, M.; Sun, L.; Fontecave, M.; Artero, V. Inorg Chem 2012, 51, 2115–2120. 104. Fourmond, V.; Canaguier, S.; Golly, B.; Field, M. J.; Fontecave, M.; Artero, V. Energy Environ Sci 2011, 4, 2417–2427. 105. Canaguier, S.; Fourmond, V.; Perotto, C. U.; Fize, J.; Pecaut, J.; Fontecave, M.; Field, M. J.; Artero, V. Chem Commun 2013, 49, 5004–5006. 106. McCormick, T. M.; Han, Z.; Weinberg, D. J.; Brennessel, W. W.; Holland, P. L.; Eisenberg, R. Inorg Chem 2011, 50, 10660–10666. 107. McNamara, W. R.; Han, Z.; Alperin, P. J.; Brennessel, W. W.; Holland, P. L.; Eisenberg, R. J Am Chem Soc 2011, 133, 15368– 15371. 108. McNamara, W. R.; Han, Z.; Yin, C. J.; Brennessel, W. W.; Holland, P. L.; Eisenberg, R. Proc Natl Acad Sci USA 2012, 109, 15594–15599. 109. Du, P.; Schneider, J.; Luo, G.; Brennessel, W. W.; Eisenberg, R. Inorg Chem 2009, 48, 4952–4962.

570

Faiella et al.

110. Eckenhoff, W. T.; McNamara, W. R.; Du, P.; Eisenberg, R. Biochim Biophys Acta - Bioenerg 2013, 1827, 958–973. 111. Shaw, W. J. Catal Rev—Sci Eng 2012, 54, 489–550. 112. Jain, A.; Lense, S.; Linehan, J. C.; Raugei, S.; Cho, H.; DuBois, D. L.; Shaw, W. J. Inorg Chem 2011, 50, 4073–4085. 113. Reback, M. L.; Ginovska-Pangovska, B.; Ho, M.-H.; Jain, A.; Squier, T. C.; Raugei, S.; Roberts, J. A. S.; Shaw, W. J. ChemEur J 2013, 19, 1928–1941. 114. Jain, A.; Reback, M. L.; Lindstrom, M. L.; Thogerson, C. E.; Helm, M. L.; Appel, A. M.; Shaw, W. J. Inorg Chem 2012, 51, 6592–6602. 115. Fontecave, M. Nat Chem Biol 2006, 2, 171–174. 116. Beinert, H.; Holm, R. H.; Munck, E. Science 1997, 277, 653– 659. 117. Klausner, R. D.; Rouault, T. A. Mol Biol Cell 1993, 4, 1–5. 118. Koay, M. S.; Antonkine, M. L.; Gartner, W.; Lubitz, W. Chem Biodivers 2008, 5, 1571–1587. 119. Brereton, P. S.; Verhagen, M. F.; Zhou, Z. H.; Adams, M. W. Biochemistry 1998, 37, 7351–7362. 120. Hoppe, A.; Pandelia, M. E.; Gartner, W.; Lubitz, W. Biochim Biophys Acta 2011, 1807, 1414–1422. 121. Klingen, A. R.; Ullmann, G. M. Biochemistry 2004, 43, 12383– 12389. 122. Beck, B. W.; Xie, Q.; Ichiye, T. Biophys J 2001, 81, 601–613. 123. Kolling, D. J.; Brunzelle, J. S.; Lhee, S.; Crofts, A. R.; Nair, S. K. Structure 2007, 15, 29–38. 124. Backes, G.; Mino, Y.; Loehr, T. M.; Meyer, T. E.; Cusanovich, M. A.; Sweeney, W. V.; Adman, E. T.; Sanders-Loehr, J. J Am Chem Soc 1991, 113, 2055–2064. 125. Backes, G.; Mino, Y.; Loehr, T.; Meyer, T.; Cusanovich, M. A.; Sweeney, W. V.; Adman, E. T.; Sanders-Loehr, J. J Am Chem Soc 1991, 113, 2055–2064. 126. Coldren, C. D.; Hellinga, H. W.; Caradonna, J. P. Proc Natl Acad Sci USA 1997, 94, 6635–6640. 127. Mulholland, S. E.; Gibney, B. R.; Rabanal, F.; Dutton, P. L. J Am Chem Soc 1998, 120, 10296–10302. 128. Mulholland, S. E.; Gibney, B. R.; Rabanal, F.; Dutton, P. L. Biochemistry 1999, 38, 10442–10448. 129. Gibney, B. R.; Mulholland, S. E.; Rabanal, F.; Dutton, P. L. Proc Natl Acad Sci USA 1996, 93, 15041–15046. 130. Laplaza, C. E.; Holm, R. H. J Am Chem Soc 2001, 123, 10255– 10264. 131. Scott, M. P.; Biggins, J. Protein Sci 1997, 6, 340–346. 132. Antonkine, M. L.; Breitenstein, C.; Epel, B.; Bill, E.; G€artner, W.; Lubitz, W. Photosynthesis Energy from the Sun; 14th International Congress on Photosynthesis; Allen, J. F.; Gantt, E.; Golbeck, J. H.; Osmond, B., Eds.; Springer: Dordrecht, 2008; p 1257–1260. 133. Antonkine, M. L.; Koay, M. S.; Epel, B.; Breitenstein, C.; Gopta, O.; G€artner, W.; Bill, E.; Lubitz, W. Biochim Biophys Acta 2009, 1787, 995–1008. 134. Grzyb, J.; Xu, F.; Nanda, V.; Luczkowska, R.; Reijerse, E.; Lubitz, W.; Noy, D. Biochim Biophys Acta 2012, 1817, 1256– 1262. 135. Grzyb, J.; Xu, F.; Weiner, L.; Reijerse, E. J.; Lubitz, W.; Nanda, V.; Noy, D. Biochim Biophys Acta 2010, 1797, 406–413. 136. Roy, A.; Sarrou, I.; Astashkin, A.; Ghirlanda, G. Biochemistry 2013, in press. DOI: 10.1021/bi401199.

137. Fritsch, J.; Scheerer, P.; Frielingsdorf, S.; Kroschinsky, S.; Friedrich, B.; Lenz, O.; Spahn, C. M. Nature 2011, 479, 249– 252. 138. Shomura, Y.; Yoon, K. S.; Nishihara, H.; Higuchi, Y. Nature 2011, 479, 253–256. 139. Grubel, K.; Holland, P. L. Angew Chem 2012, 51, 3308–3310. 140. Ferreira, K. N.; Iverson, T. M.; Maghlaoui, K.; Barber, J.; Iwata, S. Science 2004, 303, 1831–1838. 141. Loll, B.; Kern, J.; Saenger, W.; Zouni, A.; Biesiadka, J. Nature 2005, 438, 1040–1044. 142. Yano, J.; Kern, J.; Sauer, K.; Latimer, M. J.; Pushkar, Y.; Biesiadka, J.; Loll, B.; Saenger, W.; Messinger, J.; Zouni, A.; Yachandra, V. K. Science 2006, 314, 821–825. 143. Najafpour, M. M.; Moghaddam, A. N.; Allakhverdiev, S. I.; Govindjee. Biochim Biophys Acta 2012, 1817, 1110– 1121. 144. Umena, Y.; Kawakami, K.; Shen, J.-R.; Kamiya, N. Nature 2011, 473, 55–60. 145. Mullins, C. S.; Pecoraro, V. L. Coord Chem Rev 2008, 252, 416–443. 146. Cady, C. W.; Crabtree, R. H.; Brudvig, G. W. Coord Chem Rev 2008, 252, 444–455. 147. Najafpour, M. M.; Ehrenberg, T.; Wiechen, M.; Kurz, P. Angew Chem Int Ed 2010, 49, 2233–2237. 148. Yin, Q.; Tan, J. M.; Besson, C.; Geletii, Y. V.; Musaev, D. G.; Kuznetsov, A. E.; Luo, Z.; Hardcastle, K. I.; Hill, C. L. Science 2010, 328, 342–345. 149. Tagore, R.; Crabtree, R. H.; Brudvig, G. W. Inorg Chem 2008, 47, 1815–1823. 150. Toma, F. M.; Sartorel, A.; Iurlo, M.; Carraro, M.; Parisse, P.; Maccato, C.; Rapino, S.; Gonzalez, B. R.; Amenitsch, H.; Da Ros, T.; Casalis, L.; Goldoni, A.; Marcaccio, M.; Scorrano, G.; Scoles, G.; Paolucci, F.; Prato, M.; Bonchio, M. Nat Chem 2010, 2, 826–831. 151. Sartorel, A.; Carraro, M.; Toma, F. M.; Prato, M.; Bonchio, M. Energy Environ Sci 2012, 5, 5592–5603. 152. Zhao, Y.; Swierk, J. R.; Megiatto, J. D.; Sherman, B.; Youngblood, W. J.; Qin, D.; Lentz, D. M.; Moore, A. L.; Moore, T. A.; Gust, D.; Mallouk, T. E. Proc Natl Acad Sci USA 2012, 109, 15612–15616. 153. Kanan, M. W.; Nocera, D. G. Science 2008, 321, 1072–1075. 154. Moore, T. A.; Moore, A. L.; Gust, D. Philos Trans R Soc Lond, Ser B 2002, 357, 1481–1498. 155. Magnuson, A.; Styring, S.; Hammarstr€ om, L. In Photosystem II; Wydrzynski, T. J.; Satoh, K.; Freeman, J. A., Eds.; Springer: Netherlands, 2005, p 753–775. 156. Conlan, B.; Cox, N.; Su, J.-H.; Hillier, W.; Messinger, J.; Lubitz, W.; Dutton, P. L.; Wydrzynski, T. Biochim Biophys Acta 2009, 1787, 1112–1121. 157. Williamson, A.; Conlan, B.; Hillier, W.; Wydrzynski, T. Photosynth Res 2011, 107, 71–86. 158. Hay, S.; Wallace, B. B.; Smith, T. A.; Ghiggino, K. P.; Wydrzynski, T. Proc Natl Acad Sci USA 2004, 101, 17675–17680. 159. Zhang, M.-T.; Chen, Z.; Kang, P.; Meyer, T. J. J Am Chem Soc 2013, 135, 2048–2051. 160. Schmitz, C.; Vernon, R.; Otting, G.; Baker, D.; Huber, T. J Mol Biol 2012, 416, 668–677.

Biopolymers (Peptide Science)

De Novo Design of Functional Proteins 161. Koga, N.; Tatsumi-Koga, R.; Liu, G.; Xiao, R.; Acton, T. B.; Montelione, G. T.; Baker, D. Nature 2012, 491, 222–227. 162. Kang, S.-g.; Saven, J. G. Curr Opin Chem Biol 2007, 11, 329– 334. 163. Guntas, G.; Purbeck, C.; Kuhlman, B. Proc Natl Acad Sci USA 2010, 107, 19296–19301. 164. Kulp, D. W.; Subramaniam, S.; Donald, J. E.; Hannigan, B. T.; Mueller, B. K.; Grigoryan, G.; Senes, A. J Comput Chem 2012, 33, 1645–1661. 165. Fletcher, J. M.; Boyle, A. L.; Bruning, M.; Bartlett, G. J.; Vincent, T. L.; Zaccai, N. R.; Armstrong, C. T.; Bromley, E. H. C.; Booth, P. J.; Brady, R. L.; Thomson, A. R.; Woolfson, D. N. ACS Synth Biol 2012, 1, 240–250. 166. Samish, I.; MacDermaid, C. M.; Perez-Aguilar, J. M.; Saven, J. G. Annu Rev Phys Chem 2011, 62, 129–149. 167. Nanda, V.; Koder, R. L. Nat Chem 2010, 2, 15–24. 168. Suarez, M.; Jaramillo, A. J R Soc Interface 2009. 169. Peacock, A. F. A.; Iranzo, O.; Pecoraro, V. L. Dalton Trans 2009, 2271–2280. 170. Ghosh, D.; Pecoraro, V. L. Curr Opin Chem Biol 2005, 9, 97–103. 171. Senes, A. Curr Opin Struct Biol 2011, 21, 460–466. 172. Ghirlanda, G. Curr Opin Chem Biol 2009, 13, 643–651. 173. Shinde, S.; Cordova, J.; Woodrum, B.; Ghirlanda, G. J Biol Inorg Chem 2012, 17, 557–564. 174. Yin, H.; Slusky, J. S.; Berger, B. W.; Walters, R. S.; Vilaire, G.; Litvinov, R. I.; Lear, J. D.; Caputo, G. A.; Bennett, J. S.; DeGrado, W. F. Science 2007, 315, 1817–1822.

Biopolymers (Peptide Science)

571

175. Koder, R. L.; Anderson, J. L. R.; Solomon, L. A.; Reddy, K. S.; Moser, C. C.; Dutton, P. L. Nature 2009, 458, 305–309. 176. Reig, A. J.; Pires, M. M.; Snyder, R. A.; Wu, Y.; Jo, H.; Kulp, D. W.; Butch, S. E.; Calhoun, J. R.; Szyperski, T. G.; Solomon, E. I.; DeGrado, W. F. Nat Chem 2012, 4, 900–906. 177. Wilcoxen, K. M.; Leman, L. J.; Weinberger, D. A.; Huang, Z.-Z.; Ghadiri, M. R. J Am Chem Soc 2007, 129, 748–749. 178. Khare, S. D.; Kipnis, Y.; Greisen, P., Jr.; Takeuchi, R.; Ashani, Y.; Goldsmith, M.; Song, Y.; Gallaher, J. L.; Silman, I.; Leader, H.; Sussman, J. L.; Stoddard, B. L.; Tawfik, D. S.; Baker, D. Nat Chem Biol 2012, 8, 294–300. 179. Der, B. S.; Edwards, D. R.; Kuhlman, B. Biochemistry 2012, 51, 3933–3940. 180. Zastrow, M. L.; PeacockAnna, F. A.; Stuckey, J. A.; Pecoraro, V. L. Nat Chem 2012, 4, 118–123. 181. Faiella, M.; Andreozzi, C.; de Rosales, R. T. M.; Pavone, V.; Maglio, O.; Nastri, F.; DeGrado, W. F.; Lombardi, A. Nat Chem Biol 2009, 5, 882–884. 182. Faiella, M.; Maglio, O.; Nastri, F.; Lombardi, A.; Lista, L.; Hagen, W. R.; Pavone, V. Chem Eur J 2012, 18, 15890–15890. 183. Lichtenstein, B. R.; Farid, T. A.; Kodali, G.; Solomon, L. A.; Anderson, J. L.; Sheehan, M. M.; Ennist, N. M.; Fry, B. A.; Chobot, S. E.; Bialas, C.; Mancini, J. A.; Armstrong, C. T.; Zhao, Z.; Esipova, T. V.; Snell, D.; Vinogradov, S. A.; Discher, B. M.; Moser, C. C.; Dutton, P. L. Biochem Soc Trans 2012, 40, 561–566.