New Structural and Mechanistic Insights Into Functional Roles of Cytochrome b ₅₅₉ in Photosystem II

Photosystem II is the site of photosynthetic water oxidation and contains 20 subunits with a total molecular mass of 350 kDa. The structure of photosystem II has been reported at resolutions from 3.8 to 2.9 Å. These resolutions have provided much information on the arrangement of protein subunits and cofactors but are insufficient to reveal the detailed structure of the catalytic centre of water splitting. Here we report the crystal structure of photosystem II at a resolution of 1.9 Å. From our electron density map, we located all of the metal atoms of the Mn(4)CaO(5) cluster, together with all of their ligands. We found that five oxygen atoms served as oxo bridges linking the five metal atoms, and that four water molecules were bound to the Mn(4)CaO(5) cluster; some of them may therefore serve as substrates for dioxygen formation. We identified more than 1,300 water molecules in each photosystem II monomer. Some of them formed extensive hydrogen-bonding networks that may serve as channels for protons, water or oxygen molecules. The determination of the high-resolution structure of photosystem II will allow us to analyse and understand its functions in great detail. ©2011 Macmillan Publishers Limited. All rights reserved Show more

Photosynthesis converts light energy into biologically useful chemical energy vital to life on Earth. The initial reaction of photosynthesis takes place in photosystem II (PSII), a 700-kilodalton homodimeric membrane protein complex that catalyses photo-oxidation of water into dioxygen through an S-state cycle of the oxygen evolving complex (OEC). The structure of PSII has been solved by X-ray diffraction (XRD) at 1.9 ångström resolution, which revealed that the OEC is a Mn4CaO5-cluster coordinated by a well defined protein environment. However, extended X-ray absorption fine structure (EXAFS) studies showed that the manganese cations in the OEC are easily reduced by X-ray irradiation, and slight differences were found in the Mn-Mn distances determined by XRD, EXAFS and theoretical studies. Here we report a 'radiation-damage-free' structure of PSII from Thermosynechococcus vulcanus in the S1 state at a resolution of 1.95 ångströms using femtosecond X-ray pulses of the SPring-8 ångström compact free-electron laser (SACLA) and hundreds of large, highly isomorphous PSII crystals. Compared with the structure from XRD, the OEC in the X-ray free electron laser structure has Mn-Mn distances that are shorter by 0.1-0.2 ångströms. The valences of each manganese atom were tentatively assigned as Mn1D(III), Mn2C(IV), Mn3B(IV) and Mn4A(III), based on the average Mn-ligand distances and analysis of the Jahn-Teller axis on Mn(III). One of the oxo-bridged oxygens, O5, has significantly longer distances to Mn than do the other oxo-oxygen atoms, suggesting that O5 is a hydroxide ion instead of a normal oxygen dianion and therefore may serve as one of the substrate oxygen atoms. These findings provide a structural basis for the mechanism of oxygen evolution, and we expect that this structure will provide a blueprint for the design of artificial catalysts for water oxidation. Show more

1 Introduction Redox reactions play important roles in almost all biological processes, including photosynthesis and respiration, which are two essential energy processes that sustain all life on earth. It is thus not surprising that biology employs redox-active metal ions in these processes. It is largely the redox activity that makes metal ions uniquely qualified as biological cofactors and makes bioinorganic enzymology both fun to explore and challenging to study. Even though most metal ions are redox active, biology employs a surprisingly limited number of them for electron transfer (ET) processes. Prominent members of redox centers involved in ET processes include cytochromes, iron–sulfur clusters, and cupredoxins. Together these centers cover the whole range of reduction potentials in biology (Figure 1). Because of their importance, general reviews about redox centers 1−77 and specific reviews about cytochromes, 8,24,78−90 iron–sulfur proteins, 91−93 and cupredoxins 94−104 have appeared in the literature. In this review, we provide both classification and description of each member of the above redox centers, including both native and designed proteins, as well as those proteins that contain a combination of these redox centers. Through this review, we examine structural features responsible for their redox properties, including knowledge gained from recent progress in fine-tuning the redox centers. Computational studies such as DFT calculations become more and more important in understanding the structure–function relationship and facilitating the fine-tuning of the ET properties and reduction potentials of metallocofactors in proteins. Since this aspect has been reviewed extensively before, 105−110 and by other reviews in this thematic issue, 2000,2001,2002 it will not be covered here. Figure 1 Reduction potential range of redox centers in electron transfer processes. 2 Cytochromes in Electron Transfer Processes 2.1 Introduction to Cytochromes Cytochromes are a major class of heme-containing ET proteins found ubiquitously in biology. They were first described in 1884 as respiratory pigments (called myohematin or histohematin) to explain colored substances in cells. 81,111 These colored substances were later rediscovered in 1920 and named “cytochromes”, or cellular pigments. 112 The intense red color combined with relatively high thermodynamic stability makes cytochromes easy to observe and to purify. As of today, more than 70 000 cytochromes have been discovered. 78 In addition, due to their small size, high solubility, and well-folded helical structure and the presence of the heme chromophore, cytochromes are one of the most extensively studied classes of proteins spanning several decades. 79 Cytochromes are present mostly in the inner mitochondrial membrane of eukaryotic organisms and are also found in a wide variety of both Gram-positive and Gram-negative bacteria. 113,114 Cytochromes play crucial roles in a number of biological ET processes associated with many different energy metabolisms. Additionally, cytochromes are involved in apoptosis in mammalian cells. 115 Further description of the latter role of cytochromes is beyond the scope of this review, which is solely focused on the role of cytochromes in ET. For a similar reason, another family of cytochromes, the cyts P450 (CYP), which catalyze the oxidation of various organic substrates such as metabolites (lipids, hormones, etc.) and xenobiotic substances (drugs, toxic chemicals, etc.), will not be discussed in this review either. A number of books and reviews have appeared in the literature describing the role of cytochromes as ET proteins. 8,24,78−90 Here we summarize studies on both native and designed cytochromes and their roles in biological ET processes. 2.2 Classification of Cytochromes Cytochromes are classified on the basis of the electronic absorption maxima of the heme macrocycle, such as a, b, c, d, f, and o types of heme. More specifically, these letter names represent characteristic absorbance maxima in the UV–vis electronic absorption spectrum when the heme iron is coordinated with pyridine in its reduced (ferrous) state, designated as the “pyridine hemochrome” spectrum (Figure 2). Figure 2 Representative pyridine hemochromogen spectra of hemin cofactors: (A) heme b, (B) heme a, and (C) heme d 1. The spectrum of pyridine ferrohemochrome c is similar to that of heme b. Reprinted with permission from ref (116). Copyright 1992 Springer-Verlag. Table 1 shows the maximum peak positions and their corresponding extinction coefficients of the pyridine hemochrome spectra of various classes of cytochromes. These differences arise from different substituents at the β-pyrrole positions on the periphery of the heme. Table 1 UV–Vis Spectral Parameters of Pyridine Hemochrome Spectra of Various Types of Cytochromesa   pyridine hemochromogen       heme position of α peak (nm) εmM (at α peak) α peak (nm) of reduced protein example ref protoheme IX (b) 557 34.4 557–563 cyt b 6 f complex (117) heme c 550 29.1 549–561 cyt c (118) heme a 587 26 587–611 cyt aa 3 oxidase (117) heme d 613   630–635 cyt bd oxidase (116) heme d 1 620 24 625 cyt cd 1 nitrite reductase (116) heme o 553   560 cyt bo 3 oxidase (119) a Adapted with permission from ref (116). Copyright 1992 Springer-Verlag. The word “heme” specifically describes the ferrous complex of the tetrapyrrole macrocyclic ligand called protoporphyrin IX (Figure 3). 81 It is the precursor to various types of cytochromes through different peripheral substitutions. Figure 3 shows a schematic of these various types of hemes. Figure 3 Different types of heme found in cytochromes. The b-type cytochromes have four methyl substitutions at positions 1, 3, 5, and 8, two vinyl groups in positions 2 and 4, and two propionate groups at positions 6 and 7, resulting in a 22-π-electron porphyrin. Hemes a and c are biosynthesized as derivatives of heme b. In heme a, the vinyl group at position 2 of the porphyrin ring of heme b is replaced by a hydroxyethylfarnesyl side chain while the methyl group at position 8 is oxidized to a formyl group. These substituents make heme a more hydrophobic as well as more electron-withdrawing than heme b due to the presence of farnesyl and formyl groups, respectively. Covalent cross-linking of the vinyl groups at β-pyrrole positions 2 and 4 of heme b with Cys residues from the protein yields heme c, where the vinyl groups of heme b are replaced by thioether bonds. The covalent cross-linking of the two Cys residues from the protein to the porphyrin ring occurs at the highly conserved -Cys-Xxx-Xxx-Cys-His- sequences (Xxx=any amino acid). This cross-linking covalently attaches heme c to the protein. The histidine residue in the conserved sequence serves as an axial ligand to the heme iron. In heme d, two cis-hydroxyl groups are inserted at positions 5 and 6 on the β-pyrrole, which renders heme d as a 20-π-electron chlorin. Heme d 1 contains two ketone groups in place of the vinyl groups at positions 2 and 4, while two acetate groups are added to positions 1 and 3 of the tetrapyrrole macrocycle, resulting in 18-π-electron isobacteriochlorins. The hemes f is similar to heme c, with the difference in the ligands that coordinate to the heme iron at the axial position (called axial ligands) make hemes c and f spectroscopically distinct. Common axial ligands found in cytochromes are shown in Figure 4. With the exception of cytochromes c′ (cyts c′), all cytochromes with ET function contain 6-coordinate low-spin (6cLS) hemes axially ligated to amino acids such as His or an N-terminal amine group. Two axial His residues act as ligands to the heme iron in b-type cytochromes. The only example of bis-Met axial coordination to heme b is observed in the iron storage protein bacterioferritin. 120,121 A common axial His ligand is found in all cyts c, where the axial His is a part of the conserved -Cys-Xxx-Xxx-Cys-His- sequence, through which the heme is covalently attached to the protein. The most commonly encountered second axial ligand in c-type cytochromes is Met with the exception of multiheme c-type cytochromes, which generally display bis-His axial ligation of the heme iron (section 2.3.6). 80 In most cases, the His ligands are coordinated to the heme iron by their Nε atom. However, an example of Nδ coordination has been reported. 122 The f-type cytochromes contain the same type of heme with one axial His ligand, as in cyts c; the only exception is in the nature of the second axial ligation in that the second axial ligand is the NH2 group of an N-terminal tyrosine instead of the most commonly found Met or His as the second axial ligand. 123 Not surprisingly, the variation in the axial ligation makes each heme type electronically unique, resulting in different out-of-plane distortions of the heme iron from the heme plane (Figure 4) as well as different spectroscopic features (Table 1). Figure 4 Commonly found heme axial ligands in various cytochromes. (A) Class I cyts c (PDB ID 3CYT) uses His/Met axial ligation. (B) Cyts b and multiheme cyts c contain bis-His ligation (bovine liver cyt b 5, PDB ID 1CYO). (C) An unusual His/amine ligation is found only in cyt f (PDB ID 1HCZ). (D) Bis-Met ligation is encountered in bacterioferritin (PDB ID 1BCF). For c-type cytochromes the conserved -Cys-Xxx-Xxx-Cys-His- ligation and its covalent linkage to the heme via Cys residues are shown. 2.3 Native Cytochromes c 2.3.1 Functions of Cytochromes c Cytochromes c are involved in biological ET processes in both aerobic and anaerobic respiratory chains. In aerobic respiration, they are involved in the mitochondrial respiratory chain to produce the energy currency ATP by transferring electrons from the transmembrane bc 1 complex to cyt c oxidase. 85,86 In addition, cyts c have also been recently discovered to play a crucial role in programmed cell death (apoptosis), where they activate the protease involved in cell death, caspase 3. 124−126 Other examples where c-type cytochromes are involved in ET include the reduction of sulfate to hydrogen sulfide, conversion of nitrogen to ammonia in nitrogen fixation, reduction of nitrate to dinitrogen in denitrification, in phototrophs that use light energy to carry out various cellular processes, and in methylotrophs that use methane or methanol as the carbon source for their growth. Detailed descriptions of the roles of cyts c in these cases will be discussed in the following sections. As cyts c are involved in numerous crucial biological processes, they have been used extensively as a hallmark system to study biological ET by site-directed mutagenesis, which have elucidated the regions of the protein that are critical for their ET properties as well as fine-tuning the reduction potentials. 87,127−131 In addition, various inorganic redox couples have been covalently appended to surface sites of cyts c to study intraprotein ET pathways. 24,132,133 Various complexes of cyts c with other protein partners have also been prepared to study interprotein ET pathways. 134−149 2.3.2 Classifications of Cytochromes c Cytochromes c generally contain ∼100–120 amino acids. Biosynthesis of cyts c involves the formation of two thioether bonds between two Cys residues and the two vinyl groups of heme b by post-translational modification. 150,151 Primary amino acid sequence alignment shows that the residue identity of cyts c is 45–100% among eukaryotes. The electronic spectra of cyts c are dominated by the allowed porphyrin π → π* transitions that are mixed together with interelectronic repulsions that give rise to an intense band at ∼410 nm (called the Soret or γ band) and two weaker signals in the 500–600 nm range (the α and β bands). The reduced form of the protein shows a Soret band at 413 nm and sharp α and β bands at 550 nm (ε = 29.1 mM–1 cm–1) and 521 nm (ε = 15.5 mM–1 cm–1), respectively, with a ratio of α to β bands of 1.87 (Table 1). The electronic spectra of cyts c from other sources are very similar to that of horse heart cyt c. Originally classified by Ambler, 89,152 cyts c have been divided into four major classes on the basis of the number of hemes, position and identity of the axial iron ligands, and reduction potentials (Table 2). Table 2 Axial Ligand Types and Reduction Potentials of Various Cytochromesa cytochrome axial ligand heme type E (mV)b mutant E (mV) ref Nitrosomonas europaea diheme cyt c peroxidase His/Met class I 450     (153, 154) Rhodocyclus tenuis THRC cyt c   class IV 420     (155) HP1 His/Met   420       HP2 His/Met   110       LP1 bis-His   60       LP2 His/Met           Rhodopseudomonas viridis THRC cyt c   class IV 380     (156,157) H1 (c 559) His/Met   330       H3 (c 556) His/Met   20       H2 (c 552) bis-His   –60       H4 (c 554) His/Met           Rhodobacter capsulatas cyt c 2 His/Met class I 373 Gly29Ser 330 (158−160)         Pro30Ala 258           Tyr67Cys 348           Tyr67Phe 308   Chlamydomonas reinhardtii cyt f His/Ntr-Tyr cyt f 370 Tyr1Phe 369 (161)         Tyr1Ser 313           Val3Phe 373           Phe4Leu 348           Phe4Trp 336           Tyr1Phe/Phe4Tyr 370           Tyr1Ser/Phe4Leu 289           Val3Phe/Phe4Trp 342   Rhodospirillum rubrum cyt c 2 His/Met class I 324     (156) Pseudomonas aeruginosa cyt c nitric oxide reductase His/Met class I 310     (162)   bis-His cyt b 345       Pseudomonas aeruginosa cyt c peroxidase His/Met class I 320     (163) Arthrospira maxima cyt c 6 His/Met class I 314     (164) Saccharomyces cerevisiae iso-2-cyt c His/Met class I 288 Asn52Ile 243 (130) Saccharomyces cerevisiae iso-1-cyt c His/Met class I 272 Arg38Lys 249 (131, 165−173)       285 Arg38His 245         290 Arg38Gln 242           Arg38Asn 238           Arg38Leu 231           Arg38Ala 225           Asn52Ala 257           Asn52Ile 231           Tyr67Phe 234           Phe82Leu 286           Phe82Tyr 280           Phe82Ile 273           Phe82Trp 266           Phe82Ala 260           Phe82Ser 255           Phe82Gly 247   Pseudomonas aeruginosa cyt c 551 His/Met class I 276     (156) horse cyt c His/Met class I 262 Met80Ala 82 (158, 174)         Met80His 41           Met80Leu –42           Met80Cys –390   rat cyt c His/Met class I 260 Pro30Ala 258           Pro30Val 261           Tyr67Phe 224   Rhodopseudomonas palustris cyt c 556 His/Met class II 230     (80) Escherichia coli cyt b 562 His/Met cyt b (class II) 168 Phe61Gly 90 (175, 176)         Phe65Val 173           Phe61Ile/Phe65Tyr 68           His102Met 240           Arg98Cys/His102Met 440   Alicycliphilus denitrificans cyt c′ His/Met class II 132     (80) Rhodopseudomonas palustris cyt c′ His/Met class II 102     (80) cytochrome b 5 bis-His cyt b   form A 80 (177)         form B –26   Desulfovibrio vulgaris cyt c 553 His/Met class I 37 Met23Cys 29 (156, 178)       20 ± 5 Gly51Cys 28           Met23Cys/Met23Cys 88           Gly51Cys/Gly51Cys 105   bovine liver microsomal cyt b 5 bis-His cyt b 3 protoheme IX dimethyl ester 70 (179) Saccharomyces cerevisiae cyt b 2 bis-His cyt b –3     (156) Chromatium vinosum cyt c′ His class II –5     (80) rat liver microsomal cyt b 5 bis-His cyt b –7 ± 1     (129, 180) Rhodospirillum rubrum cyt c′ His/Met class I –8     (80) tryptic bovine hepatic cyt b 5 His/Met class I –10 ± 3 Val61Lys 17 (181)         Val61His 11           Val61Glu –25           Val61Tyr –33   Allochromatium vinosum triheme cyt c bis-His class III –20     (182)   His/Met   –200         His-Cys/Met   –220       Rhodobacter sphaeroides cyt c′ His/Asn cyt c –22     (183) cyt b 6 f complex bis-His cyt b –45     (184)       –150       Thermosynechococcus elongates PS cyt c 550 His/Met class I –80 in the absence of mediators 200 (185) MamP magnetochrome His/Met class I –76     (186) rat liver OM cyt b 5 bis-His cyt b –102 His63Met 110 (187, 188)         Val45Leu/Val61Leu –148           protoheme IX dimethyl ester –36   Desulfovibrio desulfuricans Norway cyt c 3 bis-His class III –132     (78)   bis-His   –255         bis-His   –320         bis-His   –360       Chlorella nitrate reductase cyt b 557 bis-His cyt b –164     (189, 190) Ectothiorhodospira shaposhnikovii cyt b 558 bis-His cyt b –210     (191) Azotobacter vinelandii bacterioferritin bis-His cyt b –225     (192) (in the presence of a nonheme iron core)     –475       Desulfovibrio vulgaris Hildenborough cyt c 3 bis-His class III –280     (192, 193)   bis-His   –320         bis-His   –350         bis-His   –380       Synechocystis sp. cyt c 549 bis-His   –250     (78) Arthrospira maxima cyt c 549 His/Met   –260     (164) a Adapted with permission from ref (78). Copyright 2004 Elsevier. b All reduction potentials listed in this review are versus standard hydrogen electrode (SHE) or normal hydrogen electrode (NHE). The class I cyts c include small (8–120 kDa) soluble proteins containing a single 6cLS heme moiety and display a range of reduction potentials from −390 to +450 mV versus standard hydrogen electrode (SHE) (Table 2). 78 On the basis of sequence and structural alignments, class I cyts c have further been partitioned into 16 different subclasses. 88 The majority of the subclasses include mitochondrial cyts c and purple bacterial cyts c. Examples of other subclasses represent a wide variety of different sources, including cyts c 551, cyts c 4, cyts c 5, and cyts c 6 from Pseudomonas, Chlorobium cyt c 555, Desulfovibrio (Dv.) cyts c 553, c 550 from cyanobacteria and algae, Ectothiorhodospira cyts c 551, flavocytochromes c, methanol dehydrogenase-associated cyt c 550 or c L, cyt cd 1 nitrite reductase, the cyt subunit associated with alcohol dehydrogenase, nitrite reductase-associated cyt c from Pseudomonas, and cyt c oxidase subunit II from Bacillus. 78 Class I cyt c domains are characterized by their signature cyt c fold and the presence of an N-terminal conserved -Cys-Xxx-Xxx-Cys-His- sequence containing cysteines for covalent cross-linking of the heme to the protein and the His, which acts as the axial ligand to the heme iron. The class I cyt c fold is recognized as having a total of five α-helices arranged in a unique tertiary structure. There are two helices, one each at the N- and C-termini, represented as α1 and α5, respectively. In between, there is a small helix, α3 (also called the 50s helix in mitochondrial cyts c), followed by two other helices, α4 and α5, which are known as the 60s helix and 70s helix, respectively, in mitochondrial cyts c. The 70s helix precedes a loop toward the C-terminus that contains the second axial ligand, Met, to the heme iron. There are examples where the second axial ligand is a residue other than Met, e.g., Asn or His, or is even absent. 79 In many cases, this core cyt c domain can be found fused to other membrane proteins. General features of the class I cyt c fold are shown in Figure 5. Figure 5 Schematic representations of various classes of cyts c. (A) Class I cyt c fold with His/Met heme axial ligands (PDB ID 3CYT). Mitochondrial designation of the helices is also shown. (B) Four-helix bundle cyt c′ belongs to class II cyt c having a 5c heme with His120 as the sole axial ligand (PDB ID 1E83). (C) Tetraheme cyt c 3 belongs to class III cyt c with bis-His ligation to all four hemes (PDB ID 1UP9). Hemes I and III are attached to the protein via the highly conserved -Cys-Xxx-Xxx-Cys-His- sequence, whereas hemes II and IV are covalently bound to the protein by a -Cys-Xxx-Xxx-Xxx-Xxx-Cys-His- motif. In (A)–(C) the covalent attachment of the heme to the protein via Cys residues is shown. (D) Tetraheme cyt c from the photosynthetic reaction center (RC) belongs to class IV cyt c. Hemes I, II, and III have His/Met axial ligands, while heme IV has bis-His axial ligation to the heme iron (PDB ID 2JBL). (E) Cyt c 554 from Nitrosomonas europaea belongs to a class of its own. Hemes I, III, and IV have bis-His-ligated heme iron, whereas heme II is 5c with His as the only axial ligand (PDB ID 1BVB). Heme numbering in (C)–(E) is according to their attachment occurring along the protein’s primary sequence. (F) Cyt f from chloroplast is unique from all other classes of cytochromes in that it mostly contains β-sheets and the heme is 6c with a His and N-terminal backbone NH2 group of a Tyr residue (PDB ID 1HCZ). It has been included as a subclass of cyt c because the heme is covalently bound to the protein via the highly conserved -Cys-Xxx-Xxx-Cys-His- signature motif for heme attachment ubiquitously found in c-type cytochromes. The class II cyts c consist of a c-type heme covalently attached to the highly conserved C-terminal -Cys-Xxx-Xxx-Cys-His- sequence, as in class I cyts c, with the Cys residues and the His as one of the axial ligands. 80 Four α-helices and a left-handed twisted overall structure represent this subclass of cyts c (Figure 5). The second axial ligand to the heme iron is variable. 194,195 The subclass cyt c′ is axially coordinated to a single His imidazole ligand, lacks the second axial ligand, and has a relatively small range of reduction potentials ranging from approximately −200 to +200 mV. 8,84,90 Members from this subclass represent a wide range of sources that include photosynthetic, denitrifying, nitrogen-fixing, methanotrophic, and sulfur oxidizing bacteria. This class has two subclasses based on the distinct spin states displayed by the heme. Subclass IIa of cyt c′ displays high-spin (HS) ferrous [Fe(II), S = 2] electronic configurations, while the ferric form shows either a HS S = 5/2 state or S = 3/2, S = 5/2 mixture of spin states. 196−202 The subclass IIa proteins, isolated from Rhodopseudomonas palustris, Rhodobacter (Rb.) capsulatus, and Chromatium (Ch.) vinosum, display a large amount of the S = 3/2 ground state in the spin-state admixture, ranging from 40% to 57% as determined from electron paramagnetic resonance (EPR) simulations. 196,201,203 The second subclass, IIb, includes cyt c 556 from Rp. palustris, 204 Rb. sulfidophilus, 205 and Agrobacterium tumefaciens(80) and cyt c 554 from Rb. sphaeroides, 206 which contain heme in the low-spin (LS) configuration. This subclass of proteins has a second axial ligand to the heme iron which is a Met residue located close to the N-terminus. Class II cyts display reduction potentials ranging from −5 to +230 mV (Table 2). Class III cyts c include proteins containing multiple hemes with bis-His ligation and display reduction potentials in the range of −20 to −380 mV (Table 2). 80,88,152,207−212 In some cases this class of cytochromes have up to 16 heme cofactors and display no structural similarity with other classes of cyts c. They are found as terminal electron donors in bacteria involved in sulfur metabolism. 213 These bacteria utilize sulfur or oxidized sulfur compounds as terminal electron acceptors in their respiratory chain. One of the best studied proteins in this class is cyt c 3 (∼13 kDa) (Figure 5) from Desulfovibrio, which acts as a natural electron acceptor and donor in hydrogenases and ferredoxins. 214 The overall protein fold containing two β-sheets and three to five α-helices is conserved among the known structures of cyts c 3 as well as the orientation of the four hemes which are located in close proximity to each other, with each of the heme planes being nearly perpendicular to the others. 88 Each heme displays a distinct reduction potential spanning a range from −200 to −400 mV. 215−219 Cyt c 555.1, also known as cyt c 7 (∼9 kDa, 70 amino acids), from Desulfuromonas acetoxidans is another class III cyt c that contains three hemes. 220 These proteins have been proposed to be involved in ET to elemental sulfur as well as in the coupled oxidation of acetate and dissimilatory reduction of Fe(III) and Mn(IV) as an energy source in these bacteria. 221 In cyt c 7, two of the hemes have a reduction potential of −177 mV and the third heme has a reduction potential of −102 mV. 222 Class IV cyts c fall into the category of large molar mass (∼35–40 kDa) cytochromes that contain other prosthetic groups in addition to c-type hemes such as flavocytochromes c and cyts cd. 152 One example of class IV cyts c is revealed by the X-ray structure of the photosynthetic reaction center (RC) from Rhodpseudomonas viridis, where light energy is harvested and converted to chemically useful energy. The cyt c in the RC consists of four c-type heme moieties covalently bound to subunit C of the RC. Three of the hemes have His/Met axial ligation, while the fourth heme is bis-His-ligated. The four hemes are oriented in two types of pairs. The porphyrin planes of hemes I/III and II/IV are orientated parallel to each other, while the porphyrin planes of each pair of hemes are mutually perpendicular to each pair’s porphyrin planes (Figure 5). 223 Cyt c 554 is another tetraheme cytochrome that is involved in the ET pathway of the biological nitrogen cycle in the oxidation of ammonia in Nitrosomonas europaea. 122,224 This family of cytochromes does not fall into either class III or class IV cytochromes and has been proposed to belong to a class of its own. A pair of electrons are passed from hydroxylamine oxidoreductase (HAO) to two molecules of cyt c 554 upon oxidation of hydroxylamine to nitrite. One of the hemes is HS, and the other three are 6cLS with reduction potentials of +47, +47, −147, and −276 mV, respectively. Porphyrin planes of hemes III and IV are oriented almost perpendicular to each other, while the heme pairs I/III and II/IV have parallel orientation (Figure 5). The sets of parallel hemes overlap at an edge, and such heme orientation has been observed in HAO and cyt c nitrite reductase. Cyt f is a high-potential (Table 2) electron acceptor of the chloroplast cyt b 6 f complex involved in oxygenic photosynthesis by passing electrons from photosystem II to photosystem I of the RC. 123,225 Cyt f accepts electrons from a Rieske-type iron–sulfur cluster and passes electrons to the copper protein plastocyanin. Cyt f consists of two domains primarily of β-sheets and is anchored to the membrane by a transmembrane segment, while most of the protein is located on the lumen side of the thylakoid membrane. The heme is also located on the lumen side at the interface of the two domains and is covalently attached to the protein via the signature sequence of cyts c, -Cys-Xxx-Xxx-Cys-His-. The β-sheet fold has not been observed in any other families of cytochromes and is thus unique to cyts f. Intriguingly, this family of cytochromes also contains an unusual second axial ligation to the heme iron, an N-terminal −NH2 group of a Tyr residue (Figure 5). Quite uniquely, the only exception to the bis-Cys covalent attachment of the c-type hemes via the conserved -Cys-Xxx-Xxx-Cys-His- motif in cyt c is found in eukaryotes from the phylum Euglenozoa, including trypanosome and Leishmania parasites. In the mitochondrial cyt c of these organisms, the heme is attached to the protein via a single Cys residue from the heme binding motif -Ala (Ala/Gly)-Gln-Cys-His-. 226−228 2.3.3 Conformational Changes in Class I Cytochromes c Induced by Changes in the Heme Oxidation State Many structural studies have been undertaken to determine whether there is any effect on the protein structure associated with different oxidation states of the heme iron. These studies include X-ray and NMR structures of oxidized and reduced cyts c from various sources, 229−235 which indicate that the oxidation state of the heme iron has a minimal effect on the tertiary structures of the proteins (Figure 6). The major changes are observed in the conformation of some amino acid residues located close to the heme pocket. Among these residues, Asn52, Tyr67, Thr78, and a conserved water (wat166) molecule show maximal changes in conformations depending on the oxidation state of the heme iron. These conserved residues, 236 along with the conserved water molecule, the axial ligand Met80, and heme propionate 7, form a hydrogen-bonding network around the heme site. The high-resolution X-ray structure of yeast iso-1-cytochrome c shows that in the reduced state the heme is significantly distorted from planarity into a saddle shape. The degree of heme distortion in the oxidized state is even more pronounced compared to that of the reduced state, suggesting that the planarity of the heme group is dependent on the oxidation state of the iron. The major change in the bond length of the heme iron ligands is observed in the case of axial Met80, which increases from 2.35 to 2.43 Å in going from the reduced to the oxidized state. On the contrary, the other axial ligand, His18, shows a minute change of 0.02 Å, from 1.99 to 2.01 Å. 230 Figure 6 Overall structural overlay of the reduced (cyan, PDB ID 1YCC) and oxidized (orange, PDB ID 2YCC) iso-1-cyt c (left). A close look at the heme site and the nearby residues is shown on the right along with some hydrogen bond interactions. In the reduced state of iso-1-cytochrome c, the conserved water molecule is hydrogen bonded to Asn52, Tyr67, and Thr78 (Figure 6). Upon oxidation wat166 undergoes a 1.7 Å displacement toward the heme, which results in the loss of the hydrogen bond to Asn52, but interactions with Tyr67 and Thr78 are retained. Figure 6 shows an overlay of the residues near the heme pocket between the reduced and oxidized states of iso-1-cytochrome c. 87 Further analysis suggested that wat166 plays a key role in stabilizing both oxidation states of the heme iron by reorienting the dipole moment, by changing the heme iron–wat166 distance, and by variations in the nearby H-bonding network. Another noticeable change is observed in the H-bonding between a conserved water, wat121, and heme propionate 7. In the reduced state, wat121 and Trp59 are hydrogen-bonded to O1A and the O2A oxygen of propionate 7, respectively. In the oxidized state, interaction between Trp59 and O2A of the heme propionate weakens, while that of O2A and the conserved Gly41 increases. Additionally, wat121 moves by 0.5 Å and causes a bifurcated hydrogen bond between both O1A and O2A of the propionate. 230 Thus, it appears that there are three major regions that show significant changes in conformation between the two oxidation states: heme propionate 7, wat166, and Met80. A conserved region that does not show mobility between oxidation states is the region encompassing residues 73–80 in iso-1-cytochrome c, which is linked to the three major regions of conformation change through Thr78. On the basis of this observation, it has been suggested that region 73–80 acts as a contact point with redox partners and triggers the necessary conformational changes in other parts of the protein that are required to stabilize both oxidation states of cyt c. 230 A contrasting observation from NMR studies is that wat166 moves 3.7 Å away from the heme iron when going from the reduced to the oxidized state, rather than moving toward the heme iron. 237,238 Similar to the changes of heme propionate observed in eukaryotes, cyts c 2(160,239−242) and c 6(220,243,244) from some prokaryotes also display conformational changes in the heme propionate between the reduced and oxidized states of the protein. In the cases of cyt c H (reduces methanol oxidase in methylotropic bacteria) from Methylobacterium extorquens and cyt c 552(245−247) (electron donor to a ba 3–cytochrome c oxidase) from T. thermophilus, there is no conserved water molecule in the heme pocket, suggesting that the water-mediated H-bonding network is not a critical requirement for ET. 248−250 2.3.4 Cytochromes c as Redox Partners to Other Enzymes In the following sections we summarize some specific examples of native enzymes that use cyts c as the native electron donor for performing various biochemical processes. 2.3.4.1 Cytochrome c as a Redox Partner to Cytochrome c Peroxidases Cytochrome c peroxidases (CcPs) are a family of enzymes that catalyze the conversion of H2O2 to water and are found in both eukaryotes and prokaryotes. Eukaryotic CcPs are located in the inner mitochondrial membrane and contain a single heme cofactor, heme b, while prokaryotic CcPs are located in the periplasmic space and contain two covalently bound c-type hemes, 251,252 one of which is a low-potential (lp) heme and the other is a high-potential (hp) heme. In general, the physiological electron donors to bacterial CcPs are monoheme cyts c, although other donors such as azurin (Az) or pseudoazurin have also been found in some bacteria. 253 The hp heme is located at the C-terminal domain and has a more positive reduction potential than cyt c as it accepts electrons from cyt c. The reduction potential for the hp heme varies depending on the organism; e.g., the Ps. aeruginosa CcP hp site has a reduction potential of +320 mV, 163 the Rb. capsulatus CcP hp site a reduction potential of +270 mV, 254 and the N. europaea CcP hp site a reduction potential of +130 mV. 154 The electrons are then transferred from the hp heme to the lp heme of CcP. In some organisms, e.g., Ps. aeruginosa and Rb. capsulatus, the hp heme should be in the ferrous state for the enzyme to be active, 254,255 whereas in other cases the enzyme is fully functional even with the ferric state of the hp heme, e.g., in N. europaea. 154 The axial ligands for the hp heme are a His and a Met, similar to most c-type cytochromes. The lp heme is the site for H2O2 reduction. It is located at the N-terminal domain and has two His residues as axial ligands. The lp heme also displays a wide range of reduction potentials from as low as −330 mV in Ps. aeruginosa(163) to as high as +70 mV in N. europaea CcP. 154 Electron transfer between the hp and lp hemes, which are 10 Å apart, is thought to occur through tunneling. 255 Cyts c interact with CcP at a small surface patch of the enzyme which has a hydrophobic center and a charged periphery. 256 The small size of the surface patch suggests that the interaction of the enzyme with the electron donor is transient, but at the same time is highly specific, which ensures complex formation due to desolvation of the surface waters and binding of cyt c. The charged periphery has been shown to be important to guide the donor toward the surface site, but it does not increase the specificity of the interactions or the ET rate. 257 Mutagenesis studies in Rb. capsulatus CcP have shown that the interface at which the enzyme interacts with its electron donor cyt c 2 involves nonspecific salt bridge interactions, as the extent of the interaction is dependent on the ionic strength of the solution. 258 In contrast, in Ps. nautica CcP, the interaction surface between the enzyme and the electron donor cyt c is highly hydrophobic on the basis of studies which showed that the enzyme was active across a wide range of ionic strength of the solution. 259 Studies from Pa. denitrificans CcP have shown that two molecules of horse heart cyt c are able to bind to the enzyme surface. 260 Binding of an “active” and “waiting” cyt c in a ternary complex with the enzyme has been proposed to improve the ET rate. Structural studies of Pa. denitrificans CcP with the monoheme cyt c has shown that the heme of the donor binds above the hp heme of CcP, while the two molecules of horse heart cyts c bind between the two hemes of the enzyme. 261 2.3.4.2 Cytochrome c as a Redox Partner to Denitrifying Enzymes: Nitrite, Nitric Oxide, and Nitrous Oxide Reductases Denitrification is a stepwise process in the biological nitrogen cycle where nitrogen oxides act as electron acceptors and are sequentially reduced from nitrate to nitrite, nitrite to nitric oxide, nitric oxide to nitrous oxide, and finally nitrous oxide to nitrogen. These four steps of the nitrogen cycle are catalyzed by a diverse family of enzymes, viz., nitrate reductase, nitrite reductase, nitric oxide reductase, and nitrous oxide reductase, all of which are induced under anoxic conditions. 262−264 Various cyt c domains act as electron donors in the denitrification process. Reduction of nitrite to nitric oxide is catalyzed by one of the two structurally diverse enzymes that also have different catalytic sites: (a) cytochrome cd 1 nitrite reductase (cyt cd 1 NiR) 265,266 and (b) multicopper nitrite reductase (CuNiR). 267,268 Cyt cd 1 NiRs are periplasmic, soluble heterodimeric enzymes containing an ET cyt c domain and a catalytic cyt d 1 domain in each subunit, while multicopper nitrite reductases are homotrimeric enzymes containing T1Cu as an ET site and T2Cu as a catalytic site. Cyts c 552 are the putative electron donors of cyt cd 1. 269 Multicopper nitrite reductases have cupredoxin-like folds and use azurins and pseudoazurins as their biological redox partner, and as such are not expected to have cyt c domains. Contrary to this expectation, two instances have been found where a fusion of multicopper nitrite reductase and cyt c domains was discovered in the genomes of Chromobacterium violaceum and Bdellovibrio bacteriovorus, where in both cases the cytochrome c domain is present at the end of a ∼500-residue-long sequence. 79 These cyt c sequences are similar to those of the caa 3 oxidase sequences. Nitric oxide reductases (NORs) are integral membrane proteins that catalyze the two-electron reduction of nitric oxide to nitrous reductase. 270,271 A recent X-ray structure of the Gram-negative bacterium Ps. aeruginosa cyt c-dependent NOR (cNOR) (Figure 7) shows that the enzyme consists of two subunits. 272 The NorB subunit is the transmembrane subunit and contains the binuclear active site consisting of an HS heme b 3 and a nonheme iron (FeB) site. It also houses an LS ET cofactor heme b. NorC is a membrane-anchored cyt c and contains a c-type heme. Electrons are received from cyt c 552 or azurin to the heme c, which then passes the electrons to LS heme b and then to HS heme b 3 of the catalytic binuclear active site. The reduction potentials are +310, +345, +60, and +320 mV for heme c, heme b, heme b 3, and the FeB sites, respectively. 162 Figure 7 X-ray structure of cyt c-dependent NOR (cNOR) (PDB ID 3O0R) from Ps. aeruginosa. 2.3.4.3 Cytochromes c as Redox Partners to Molybdenum-Containing Enzymes Mononuclear molybdenum-containing enzymes constitute a group of enzymes that catalyze a diverse set of reactions and are found in both eukaryotes and prokaryotes. 273,274 The general function of these groups of enzymes is to catalytically transfer an oxygen atom to and from a biological donor or acceptor molecule, and these enzymes are thus referred to as molybdenum oxotransferases. These enzymes possess a Mo=O unit at their active site and an unusual pterin cofactor which coordinates to the metal via its dithiolene ligand moiety. These Mo-containing enzymes are generally classified into three families depending on their structures and the reactions that they catalyze. The first one is xanthine oxidase from cow’s milk, which has an LMoVIOS(OH) (L = pterin) catalytic core and generally catalyzes the hydroxylation of carbon centers. The second family includes sulfite oxidase from avian or mammalian liver with a core coordination consisting of an LMoVIO2(S–Cys) moiety that catalyzes the transfer of an oxygen atom to or from the substrate’s lone pair of electrons. The third family of oxotransferases shows diversity in both structure and function and uses two pterin ligands instead of only one used by the first two classes. The reaction occurs at the active site core containing L2MoVIO(X), where X could be Ser as in DMSO reductase or Cys as in assimilatory nitrate reductase. Xanthine oxidases have been reported to be coexpressed with three cyt c domains in Bradyrhizobium japonicum, Bordetella bronchiseptica, Ps. aeruginosa, and Ps. putida; however, the exact cause of this association is not well understood as these enzymes use flavins as their redox partners. 79 Sulfite oxidase catalyzes the oxidation of sulfite to sulfate using 2 equiv of oxidized cyt c as physiological oxidizing substrates (Scheme 1). 273 The molybdenum is reduced from the VI to the IV oxidation state, and the reducing equivalents are then transferred sequentially to the cyt c in the oxidative half-reaction. The assimilatory nitrate reductases (NRs) are found in algae, bacteria, and higher plants which uptake and utilize nitrate. 273 These enzymes contain a cyt b 557 and flavin adenine dinucleotide (FAD) in addition to the Mo center. Electrons flow from FAD to cyt b 557 to the Mo center under physiological conditions. The midpoint reduction potentials for FAD and cyt b 557 from Chlorella NR have been determined to be −288 and −164 mV, respectively. 189,190,275 The Mo center displays reduction potentials of +15 mV for the MoVI/V couple and −25 mV for the MoV/IV couple. These reduction potentials indicate that the physiological direction of electron flow is thermodynamically favorable. The cyt b 557 domain of NR is homologous to the mammalian cyt b 5, yeast flavo-cyt b 2, and cyt b domain of sulfite oxidase. 276 Scheme 1 Scheme Showing the Oxidation of Sulfite to Sulfate by Cyt c in Sulfite Oxidase Reprinted from ref (273). Copyright 1996 American Chemical Society. The DMSO reductase family consists of a number of enzymes from bacterial and archaeal sources that display remarkable sequence similarity. Respiratory DMSO reductases are periplasmic and use membrane-anchored multiheme cyts c as electron donors that transfer electrons from the quinine pool to the periplasmic space. These cytochromes are about 400 amino acids long and are encoded in the same operon as the enzyme. In some γ-proteobacteria, the tetraheme cyts c occur as a fusion to the C-terminal cyt c binding domain of the enzyme. On the other hand, in some ε-proteobacteria single-domain cyts c have been coexpressed with the DMSO reductase and act as electron donors to the enzyme. Nonetheless, the cyt c sequences from both types of proteobacteria are clustered together, suggesting that even though the mechanism of ET is different, they are functionally similar. 79 Even though these ET proteins in DMSO reductases are referred to as cyts c because they contain c-type hemes, their structural folds do not fall into the uniquely defined category of cyt c folds as mentioned in section 2.3.2. 2.3.4.4 Cytochrome c as a Redox Partner to Alcohol Dehydrogenase The type II quinohemoprotein alcohol dehydrogenases are periplasmic enzymes that catalyze the oxidation of alcohols to aldehydes and transfer electrons from substrate alcohols first to the pyrroloquinoline quinone (PQQ) cofactor, which subsequently transfers electrons to an internal heme group that is found in a cyt c domain. 277 This cyt c domain of about 100 residues contains three α-helices in the core cytochrome domain and is similar to the cyt c domain in p-cresol methylhydroxylase (PCMH) from Ps. putida(278) and the cyt c551i from Pa. denitrificans. 279 2.3.4.5 Involvement of Cytochromes c in Photosynthetic Systems Photosynthesis involves the conversion of light energy to useful chemical forms of energy, which is accomplished by two large membrane protein complexes, photosystem I (PSI) and photosystem II (PSII). 280 The catalytic cores of the two PSs are referred to as the reaction centers, which have [4Fe−4S] clusters and quinines as terminal electron acceptors for PSI and PSII, respectively. Like algae and higher plants, cyanobacteria also use PSI and PSII to convert light energy to chemical forms by producing oxygen from water oxidation. Even though cyanobacteria have a bis-His-coordinated PS-C550 cyt subunit in their PSII, apparently there is no redox role of this cytochrome. 281,282 Being located at the lumenal surface of the enzyme, PS-C550 cytochrome acts as an insulator of the catalytic core from reductive attack and contributes to structural stabilization of the complex. 283,284 The low midpoint reduction potentials of the soluble protein from −250 to −314 mV exclude any redox role of this class of cytochromes. 285−288 When complexed with PSII, more positive values of reduction potentials have been determined. 288,289 A reduction potential of +200 mV in PS-C550 cytochrome from Thermosynechococcus elongates has recently been reported, 185 which suggests a possible role of this cytochrome in ET in PSII, despite a long distance (∼22 Å) between the PS-C550 cytochrome and its nearest redox center, the Mn4Ca cluster. 290 In cyanobacteria, cyt c 6 is known to act interchangeably with the copper protein plastocyanin as an electron donor to PSI, depending on the availability of copper, 291−293 while in higher plants plastocyanin is the exclusive electron donor. On the basis of this observation, it has been proposed that cyt c 6 is the older ancestor, which has been replaced by plastocyanin during evolution due to the shortage of iron in the environment. 294 Another cytochrome, cyt c M, is found exclusively in cyanobacteria, but its role is ambiguous. It has been shown to be expressed under stress-induced conditions such as intense light or cold temperatures where the expression of both cyt c 6 and plastocyanin is suppressed. 295 Thus, it would be tempting to believe that cyt c M is a third electron donor to PSI in cyanobacteria under stress conditions, but experimental evidence goes against this hypothesis. 296 2.3.4.6 Cytochrome c as a Single-Domain Oxygen Binding Protein Sphaeroides heme protein (SHP) is an unusual c-type cytochrome which was discovered in Rb. sphaeroides. 183 SHP (∼12 kDa) has a single HS heme with a reduction potential of −22 mV and an unusual His/Asn axial heme coordination in the oxidized form. SHP is spectroscopically distinct from cyts c′, which also have a HS heme. SHP was shown to bind oxygen transiently during slow auto-oxidation of the heme. The Asn axial ligand was shown to swing away upon reduction of the heme or binding of small molecules such as cyanide or nitric oxide. The distal pocket of SHP shows marked resemblance to other heme proteins that bind gaseous molecules. 297 It has been suggested that SHP could be involved as a terminal electron acceptor in an ET pathway to reduce small ligands such as peroxide or hydroxylamine. 297 2.3.5 Cytochrome c Domains in Magnetotactic Bacteria Magnetotactic bacteria consist of a group of taxonomically and physiologically diverse bacteria that can align themselves with the geomagnetic field. 298 The unique property of these bacteria is due to the presence of iron-rich crystals inside their lipid vesicles forming an organelle, referred to as the magnetosome. From sequence analysis, three proteins, MamE, MamP, and MamT, in the Gram-negative bacterium Magnetospirillum magneticum AMB-1 that contribute to the formation of the magnetosome have been discovered to contain a double -Cys-Xxx-Xxx-Cys-His- motif, characteristic of cyts c. 186 All three proteins were expressed and purified in E. coli. Subsequent characterization of these proteins confirmed that MamE, MamP, and MamT indeed belong to c-type cytochromes, and they have been designated as “magnetochromes”. Midpoint reduction potentials were determined to be −76 and −32 mV for MamP and MamE, respectively. The presence of cyts c proteins in magnetotactic bacteria is intriguing and suggests that these proteins take part in ET, although the exact nature of their ET partners is not known. It has been hypothesized that the magnetochromes can either donate electrons to Fe(III) and participate in magnetite [mixture of Fe(III) and Fe(II)] formation or accept electrons from magnetite to maintain a redox balance, or they can act as redox buffers to maintain a proper ratio of maghemeite (all ferric irons) and magnetite. 2.3.6 Multiheme Cytochromes c Multiheme cyts c occur as both soluble and membrane-anchored ET proteins in many enzymes across diverse functionalities. 79,299 Triheme cyts c 7 from Geobacter sulfurreducens and Dm. acetoxidans are involved in ET for Fe(III) respiration, 207,300−303 although their exact roles are not known. These proteins have conserved secondary structural elements consisting of double-stranded β-sheet at the N-terminus followed by several α-helices. The protein displays a miniaturized version of the cyt c 3 fold where heme II and the surrounding protein environment are missing (Figure 8). The arrangement of hemes is conserved in cyts c 7 in terms of the distances between heme iron atoms and the angles between heme planes. Hemes I and IV are almost parallel to each other and are mutually perpendicular to heme III, which is in close contact with hemes I and IV. NMR and docking experiments suggest that heme IV is the region of interaction with similar physiological partners, while the other interacting partner would most likely interact through the region near hemes I and III. Such differences in interaction surfaces might play a role in choosing the right redox partners to perform different physiological functions. Figure 8 (A) X-ray structure of triheme cyt c 7 (PDB ID 1HH5). All the hemes are bis-His-ligated. Cyt c 7 is a minimized version of cyt c 3 where heme II is missing. (B) Spatial arrangement of the four hemes in flavocytochrome c 3 fumarate reductase (PDB ID IQO8). The heme irons of the heme pair II and III are in close proximity at 9 Å from each other, and the heme edges are 4 Å away. An unusual triheme cyt c is DsrJ from the purple sulfur bacterium Allochromatium vinosum that is a part of a complex involved in sulfur metabolism. 182,304 Sequence analysis suggested the presence of three distinct c-type hemes containing bis-His, His/Met, and a very unusual His/Cys axial ligation, respectively. Subsequent cloning and expression of DsrJ in E. coli indeed confirmed the presence of three hemes, and EPR data showed the presence of partial His/Cys coordination to one of the hemes (His/Met is another possibility). From redox titrations, reduction potentials of the hemes were determined to be −20, −200, and −220 mV, respectively. Although the exact role of DsrJ is still unknown, its involvement in catalytic functions rather than in ET has been hypothesized. 182 Other examples of multiheme cyts c include a tetraheme cyt c (NapC) involved in nitrate reductase from Pa. denitrificans, 305 an Fe(III)-induced tetraheme flavocytochrome c 3 (Ifc3) 306 in fumarate reductase (Fcc3) from Sh. frigidimarina, an HAO containing eight heme groups for hydroxylamine oxidation in N. europaea, 307 and a pentaheme nitrite reductase (NrfA) for nitrite reduction in Sulfurospirillum deleyianum. 308,309 A periplasmic flavocytochrome c 3 which is an isozyme of the soluble Fcc3 is also induced by Fe(III). 310−312 The X-ray structure of this protein shows that the tetraheme arrangement in Fcc3 includes an intriguing heme pair where the two irons are only 9 Å from one another and the closest heme edges are within 4 Å (Figure 8). The four hemes from Ifc3 and Fcc3 can be superimposed on four of the eight hemes in HAO. 307 All four hemes of Ifc3 overlay on four of the hemes from the pentaheme NrfA, 308 and all five hemes from NrfA overlay on five of the HAO hemes. Lastly, two hemes from Ifc3 overlay on two of the four hemes of cyt c 554(122) from N. europaea, all four hemes of which overlay on four hemes from HAO. Despite such similarities in heme arrangement, there is no resemblance in the primary sequence of these enzymes. Nevertheless, such similar heme arrangements in these proteins suggest that they share a common ancestor, but have evolved divergently to perform four different reactions, viz., Fe(III) reduction, fumarate reduction, hydroxylamine reduction, and nitrite reduction. 313 Some membrane-bound multiheme cytochromes, belonging to the NapC/NirT family, contain four heme binding sequences that have evolved due to gene duplication of diheme domains. 314 In NapC and CymA all four hemes are 6cLS with bis-His axial ligation and display reduction potentials of +10 and −235 mV, respectively. 305,313 Sh. oneidensis MR-1 is a facultative anaerobe that is capable of using many terminal electron acceptors such as DMSO or metal oxides such as ferrihydrite and manganese dioxide outside the outer cell membrane, accepting electrons from the quinol pool and the tetraheme protein CymA. 317−325 Electron transfer in Sh. oneidensis MR-1 is facilitated by two periplasmic decaheme cyts c, DmsE, which supplies electrons to DMSO, and MtrA, which is involved in ET to metal oxides (Figure 9). Both of these decaheme proteins have been proposed to be involved in a long-range ET across a ∼300 Å “gap”  326 (∼230 Å periplasmic gap and ∼40–70 Å thick outer membrane). Using protein film voltammetry, a potential window between −90 and −360 mV and an ET rate of ∼122 mV s–1 were measured for DmsE at pH 6. 315 The measured reduction potential window for DmsE is shifted ∼100 mV lower than what was observed in MtrA, 327−329 although the rate of ET is similar in both proteins. Although the MtrA and DmsE families of decaheme proteins facilitate long-range ET in Sh. oneidensis, it is not clear how ET is feasible across a 300 Å gap, especially given the fact that MtrA spans only 105 Å in length. 330 Clearly, the arrangement of hemes must play a crucial role; however, the exact mechanism of this ET process is yet to be determined. A recent NMR study proposes the presence of two independent redox pathways by which the ET occurs from the cytoplasm to electron acceptors on the cell surface across the periplasmic gap in MtrA, 331 one involving small tetraheme cyt c (STC) and the other involving FccA (flavocytochrome c). Both of these proteins interact with their redox partners CymA (donor) and MtrA (acceptor) through a single heme and show a large dissociation constant for protein–protein complex formation. Together, these facts suggest that a stable multiprotein redox complex spanning the periplasmic space does not exist. Instead, ET across the periplasmic gap is facilitated through the formation of transient protein–protein redox complexes. Figure 9 (A) Schematic model for DMSO reduction by DmsEFAB and iron reduction by MtrABC(DEF). Flows of electrons are shown with arrows. DmsE and MtrA(D) are proposed to accept electrons from the menaquinone pool via CymA. Multiheme groups in CymA, MtrACDF, and DmsE are shown. IM = inner membrane, and OM = outer membrane. (B) “Staggered-cross” orientation of the hemes in outer membrane decaheme MtrF (PDB ID 3PMQ). Heme numbering is shown as Roman numerals, heme–iron distances are shown in orange, and distances between heme edges are shown in blue. (A) Reprinted with permission from ref (315). Copyright 2012 Biochemical Society. (B) Adapted from ref (316) Copyright 2011 National Academy of Sciences. MtrF is a decaheme c-type cytochrome found in the outer membrane of Sh. oneidensis MR-1 (Figure 9) which has been proposed to transfer electrons to solid substrates through the outer membrane, like its homologue MtrC, with the help of periplasmic MtrA and a membrane barrel protein, MtrE, that facilitates ET by forming contact between MtrA and MtrF. 332,333 A recent crystal structure of MtrF shows that the protein consists of four domains, domains I and III containing β-sheets and domains II and IV being α-helices. 316 The arrangement of the 10 bis-His-ligated hemes is like a “staggered cross” where four hemes (I, II, VI, VII) are almost coplanar with each other and are almost perpendicular to a group of three hemes (III, IV, V and VIII, IX, X) that are parallel to each other (Figure 9). The reduction potentials of the hemes in MtrF lie in the range of 0 to −312 mV as determined by both solvated and protein film voltammetry. Unfortunately, reduction potentials of individual hemes have not been possible to assign due to their similar chemical nature. Molecular dynamics simulations show an almost symmetrical free energy profile for ET. Additionally, the computed reorganization energy range from 0.75 to 1.1 eV is consistent for partially solvent exposed heme cofactors capable of overcoming the energy barrier for ET. 334,335 Further molecular details of ET in MtrF are unknown. Multiheme cyts c also act as ET agents in the Fe(III)-respiring genus Shewanella. 299 However, due to the fact that Fe(III) is soluble only at pH 5 M GuHCl due to heme dissociation. Folding of the reduced state has been shown to be triggered by photoinduced ET to the oxidized form of the protein under 2–3 M GuHCl concentrations. A folding rate of 5 μs was extrapolated in the absence of denaturant, which is similar to the intrachain diffusion time scale of the polypeptide. 388 2.4 Designed Cytochromes In addition to studying native systems by a top-down approach, in recent decades, many groups have adopted a bottom-up approach of building minimal functional proteins that mimic natural ones. The theoretical simplicity and ubiquity of cytochromes has made them appealing targets for design, and a number of artificial cytochrome-mimicking proteins have been engineered, with varying levels of sophistication. In this issue of Chemical Reviews, Pecoraro and co-workers give a thorough review of protein design strategies and successes, including designed heme ET proteins. 3000 Here, we give a brief account focusing on the redox properties of designed 6-coordinate heme proteins mimicking ET cytochromes. 2.4.1 Designed Cytochromes in de Novo Designed Protein Scaffolds Two de novo heme proteins called VAVH25(S–S) and retro(S–S) 389 were designed to bind heme in a bis-His coordination, by strategically engineering His residues into the de novo cystine-cross-linked, homodimeric four-helix bundle called α2. 390−392 Both sequences yielded artificial cytochromes with dissociation constants for heme in the submicromolar range, and spectroscopic properties of these proteins were consistent with low-spin bisimidazole-ligated heme, with reduction potentials of −170 and −220 mV for each of the proteins. Although these potentials are nearly unchanged from the potentials of bisimidazole heme in aqueous solution, the success of incorporation demonstrated the power of rational de novo design and set the stage for rapid development of more complex and nativelike structures. Using an alternative tetrameric protein scaffold, consisting of two pairs of disulfide linked α-helices, a series of proteins mimicking the heme b domain of cytochrome bc 1 were also designed by strategic placement of histidine residues. The designed proteins incorporated either two or four hemes per bundle, 393 with potentials of the individual sites reported to range from −230 to −80 mV in the tetraheme construct. More impressively, the sites showed cooperative redox properties, with the presence of a second ferric heme site proposed to raise the potential of the first by ∼115 mV through electrostatic interactions (vide infra). 393,394 In a systematic study of the electronic properties of this scaffold, varying the heme, pH, and local charge could achieve a potential range of 435 mV (−265 to +170 mV), 395 over half the 800 mV range covered by native cytochromes. Interestingly, investigation of the more natural mutation of one of the His ligands with a Met resulted in only a 30 mV increase in reduction potential, and substitution of heme b with heme c gave no significant change. 396 Rational mutagenesis of several core residues, as well as incorporation of helix–turn–helix and asymmetric disulfide bonds, further improved the structural rigidity and uniqueness of the designed scaffolds. 397,398 Subsequently, this maquette system was extended in a variety of ways to achieve coupling to electrode surfaces, 399 incorporation of non-natural amino acid ligands, 400 and binding of two different hemes—which mimics the structure of ba 3 oxidases. 401 Particularly exciting is the demonstration of coupling of ET and protonation of carboxylate residues on the protein, 402−404 which is relevant for understanding and engineering proton pumping. On the basis of recent developments in structural understanding of cytochrome bc 1 and improvements in computational modeling, Ghirlanda et al. investigated designing a more structurally unique mimic of the bc 1 complex. The structure of the heme b binding portion of bc 1 was modeled as a coiled coil, and secondary coordination sphere interactions to the coordinating histidines, such as conserved Gly, Thr, and Ala residues, were added to stabilize the orientation of the His ligand and tune its electronic properties (Figure 13A). 405 The potentials were measured by cyclic voltammetry (CV) as −76 and −124 mV in the oxidative and reductive directions, respectively, at pH 8, significantly higher than the potential of aqueous bisimidazole heme and earlier bis-His-ligated designed proteins. The hysteresis in the potentials is attributed to conformational reorganization of the ligating His residues between the oxidized and reduced forms. The model was further improved by linking and expression as a single chain for more efficient structure determination studies, 408 as well as incorporation into a membrane. 409 Figure 13 Structural models of designed cytochrome models in de novo scaffolds. (A) A design model for a homodimeric four-helix tetraheme binding protein inspired by cyt bc 1. Remade from coordinates courtesy of G. Ghirlanda and W. F. DeGrado. 405 (B) Schematic representation of monomeric four-α-helix maquettes used to mimic ET cytochromes. Reprinted with permission from ref (406). Copyright 2013 Macmillan Publishers Ltd. (C) Crystal structure of Co(II) mimichrome IV (PDB 1PYZ). 407 Most recently, Dutton and co-workers have reported the design and thorough characterization of a monomeric, single-chain four-α-helix bundle maquette protein, which can bind up to two hemes (Figure 13B). It is particularly noteworthy for the subject of this review that the redox properties of this scaffold as a function of charge distribution were systematically analyzed. By raising the total charge uniformly from −16 to +11, the reduction potential of both hemes changed from −290 to −150 mV, as expected. Furthermore, the potentials of the hemes could be changed individually by only increasing the charge at one end of the protein; the potentials of the individual hemes were −240 and −150 mV. Finally, it was demonstrated that the reduced negatively charged protein could transfer an electron to native cytochrome c with rate constants approaching those of native photosynthetic and respiratory electron transport chains. Such a single-chain four-helix bundle was also used to build an artificial oxygen binding cytochrome c with an intramolecular B-type ET heme with a 60 mV lower reduction potential, mimicking a natural ET chain. 410 More rational computational protein design algorithms have also been brought to bear on the de novo design of artificial cytochromes. Xu and Farid used the algorithm named CORE 411 to design a nativelike four (27 amino acid)-helix bundle that binds two to four hemes in a bis-His fashion. 412 The α-helical character was confirmed by circular dichroism (CD), and the binding affinity for the first 2 equiv was determined to be in the micromolar range, while, due to negative cooperativity, the remaining sites had K d > 3 mM. The measured potentials for the diheme and tetraheme protein were −133 to −91 and −190 to −0110 mV, respectively. While the rationally guided design strategies described above have been very successful, the lack of a priori knowledge about the necessary structural features for design of functional metalloproteins limits the scope of sequence and structure space that is probed by the strategy. As a complementary approach, Hecht and co-workers have utilized a semirational “binary code” library generation method to produce 15 74-residue sequences that formed helical bundles and bound heme, 413 one with submicromolar affinity. Extending this scaffold further produced five 102-residue sequences with higher stabilities and more “nativelike” structures. 414 Analysis of a handful of these proteins revealed spectroscopic features typical of low-spin heme proteins and reduction potentials ranging from −112 to −176 mV. 415 Furthermore, it was demonstrated that at least one construct was electrically competent on an electrode. 416 A similar semirational combinatorial approach was utilized by Haehnel and co-workers, who combined it with template-assisted synthetic protein (TASP) methods, in which two sets of antiparallel helices are templated onto a polypeptide ring, to design and screen an impressive library of 399 cytochrome b mimicking four-helix bundles. 417,418 Using a colorimetric screen, the potentials were estimated to range from −170 to −90 mV. It was also demonstrated that the proteins could be incorporated onto electrodes 419,420 and achieved estimated ET rate constants comparable to those of native cytochromes. A number of smaller, water-soluble peptide-based cytochrome mimics have also been developed, utilizing one or two short α-helical peptides. Two groups independently developed heme compounds with covalently attached, short α-helix-forming peptides, with His ligands. In one case, peptide-sandwiched mesoheme (PSM) compounds were prepared by covalently attaching a 12-mer peptide to each of the two propionate groups of the heme via amide bonds with lysine groups on the peptide. 421 Although the helicity of the free peptide was low, upon ligatation of the heme, the helicity was seen by CD to increase to ∼50%, and the electronic spectra were consistent with bis-His heme ligation, similar to b-type cytochromes. 421,422 Further work suggested that aromatic side chain interaction with the heme, such as Phe and Trp, improves helix stability and heme binding, 423 and covalent linkage of the peptide termini via disulfide bonds resulted in further stabilization. 424 Studies of the redox properties of a PSM and a mutant with an Ala to Trp mutation, (called PSMW), highlight the importance of stability in determining reduction potential, with more stable helix binding in PSMW lowering the reduction potential by 56 mV (−281 to −337 mV), due to the increased ability of the His ligands to stabilize the Fe(III) state. 425 The authors propose that this effect may also explain the difference in potential between mitochondrial and microsomal cyts b 5. Similarly, short α-helical peptides, based on the heme binding peptide fragment of myoglobin, have been covalently attached to deuterohem by a similar amide-bond attachment strategy, yielding compounds known as mimochromes. 426 It is noteworthy that the peptides retained their α-helical character even in the absence of heme binding. 426,427 The stability of the model was further improved in later revisions by enhancing the intramolecular interpeptide interactions through extending the peptide (mimochrome II) 428 or rational mutagenesis (mimochrome IV). 429 A crystal structure of the Co(II) derivative of mimochrome IV has been obtained and substantiates the designed structure (Figure 13C). 407 The reduction potential of Fe mimochrome (IV) at pH 7 is −80 mV, though it exhibits strong pH dependence over the range of pH from 2 to 10 (∼+30 to −170 mV). 429 The low-pH dependence is attributed to the His ligands unbinding from the heme, while the high-pH transition is proposed to be caused by deprotonation of a nearby arginine; however, this is surprising due to the 4 orders of magnitude higher apparent acidity and requires further investigation to be proven. Still, it is exciting that this simple mimic is well folded enough to be crystallized and has a potential in the range of those of native cytochromes. Intermediate between these covalently attached heme–peptide models and full polyhelical bundles described above, heme protein complexes consisting of heme ligated by designed short peptides that are not covalently attached have also been developed. 430−434 Studies on the binding of a variety 15-mer peptides showed a strong correlation between peptide–heme affinity and reduction potential (−304 to −218 mV), with lower potentials for more stable complexes, consistent with the results of studies on PSMs. 425,431 The overall low potential was attributed to the inability of the small peptides to reduce the strong dielectric constant of the solvent, as native proteins do (vide infra). To further improve the stability, two peptides were covalently linked at both ends by disulfide ligands, resulting in a series of cyclic dipeptide heme binding motifs, with reduction potentials ranging from −215 to −252 mV. 433 Interestingly, in a step away from the helix bundle paradigm, Isogai and co-workers were able to rationally design a series of de novo proteins that would fold into a globin fold, but with only ∼25% sequence identity to sperm whale myoglobin. 435,436 Although the proteins were designed for a 5-coordinate myoglobin-like heme binding site, the resulting proteins were consistent with 6-coordinate bis-His-ligated heme. In these scaffolds, the reduction potential was in the range of −170 to −200 mV, similar to that of aqueous bis-Im heme, which was attributed to higher solvent access to the heme due to the molten-globular state of the proteins. This was further supported by the re-engineering of a nonheme globin protein, phycocyanin, into a heme binding protein (vide infra), which had a more unique, hydrophobic, and nativelike core structure and 50 mV higher reduction potential. 437 2.4.2 Designed Cytochromes in Natural Scaffolds In addition to designing scaffolds for cytochromes de novo, an appealing alternative strategy is to make use of the diversity of natural proteins as scaffolds. One of the most straightforward approaches is to convert a non-cytochrome heme protein into a cytochrome by site-directed mutagenesis. Along these lines, various myoglobins have also been redesigned into bis-His cytochrome-like proteins, similar to b 5, by mutating the valine near the heme at position E11 to histidine (Figure 14A). 438−440 The spectroscopic features of reduced and oxidized forms of these mutants are consistent with low-spin bis-His-ligated heme, and the crystal structure confirms the ligation. 440 The mutations result in a 170 mV decrease in the reduction potential of myoglobin, from ∼60 to ∼−110 mV. Figure 14 Structural models of designed cytochrome models in native scaffolds. (A) X-ray crystallographic model of a pig myoglobin designed to have cytochrome-like bis-His ligation (PDB ID 1MNI). 440 (B) Molecular dynamics model of a histidine mutant of the membrane protein, glycophorin A, designed to bind heme in a cytochrome-like manner. 441 Coordinates provided by courtesy of G. Ghirlanda. Similarly, natural nonheme proteins can also be designed to bind heme in a manner consistent with the cytochrome binding motif. As briefly mentioned above, Isogai and co-workers introduced two histidines into the natural nonheme plant globin phycocyanin 437 to generate a heme binding site. Although the protein was designed as a myoglobin mimic, the spectral features were consistent with low-spin bis-His coordination, similar to that of cytochromes b, with a one-electron reduction potential of −120 mV. Heme binding sites have also similarly been designed into native α-helical bundle proteins that do not have native heme binding sites. Starting with the DNA binding protein rop, a specific bis-His heme binding protein was designed by removing surface histidines and introducing two internal histidine residues. 442 An alternative His/Met binding mode was also investigated. 443 Both proteins displayed electronic spectra characteristic of low-spin heme, with reduction potentials of −155 and −88 mV, respectively. A cytochrome-like heme binding site was also designed into the transmembrane protein glycophorin A (Figure 14B). 441,444 Each of the proteins bound heme with submicromolar affinity, and the presence of aromatic phenylalanine residues near the heme lowered the reduction potential from −128 to −172 mV. 2.4.3 Conversion of One Cytochrome Type to Another In addition to designing cytochrome sites in non-cytochrome proteins, several groups have investigated the conversion of one type of cytochrome into another. 445−449 Conversion of c-type to b-type cytochrome has been achieved in cytochrome c 552 by removing the Cys residues in the -Cys-Xxx-Xxx-Cys-His- heme binding motif with the Cys11Ala/Cys14Ala double mutation. 447 CD and NMR spectra confirmed that the structure of the protein and heme site was maintained. 447,450 However, it was found that the removal of the c-type heme binding motif destabilized the protein toward chemical and thermal denaturation. While the electron-withdrawing potential of the vinyl groups of heme b relative to the thioether groups of heme c would be expected to raise the potential, 80 the resulting protein had a reduction potential of 170 mV, 75 mV lower than that of the wild type, suggesting that the electronic structure of the porphyrin is not the major determinant of the reduction potential difference between cytochromes c and b (discussed in section 2.5). Conversion from cyt b 562 to c-type heme has been achieved by introducing the conserved -Cys-Xxx-Xxx-Cys-His- motif into the wild-type protein by means of two mutations (Arg98Cys and Tyr101Cys). 449,451 The resulting c-type cytochrome displayed enhanced stability toward chemical denaturants, maintaining the same protein fold and axial His ligation. c-type heme attachment has also been achieved in cytochrome b 5 by introducing a surface cysteine residue with the Asn57Cys mutation. 448 The resulting holoprotein was isolated in four forms, with distinct forms of heme, one of which contained covalently attached heme and a hemochrome α-band at 553 nm, intermediate between those of b-type (556 nm) and c-type (551 nm) heme, suggesting the presence of a single c-type thioether linkage. NMR further confirmed the stereochemical nature of this linkage, and the protein displayed a reduction potential of −19 mV, 23 mV lower than that of the wild-type b 5. 2.5 Structural Features Controlling the Redox Chemistry of Cytochromes Being involved in distinct ET pathways, each cytochrome has evolved its ET properties to match those of its redox partners. Therefore, reduction potentials of cytochromes span a range of almost 1 V, from −475 mV in bacterioferritin from Azotobacter vinelandii(192,452) to +450 mV in the heme c of diheme cytochrome c peroxidase of N. europaea(153,154) vs the SHE. 453 Through a variety of studies, many properties have been found to be important in determining the redox properties of heme proteins. As expected, the molecules in the first coordination sphere of the iron, namely, the four pyrrole groups of the porphyrin and the axially coordinating residues, are important in determining the baseline reduction potential, as they interact directly with the iron center. These interactions are also fine-tuned by the secondary coordination sphere—chemical moieties that interact with the primary coordination sphere ligands and adjust their properties. Secondary coordination sphere interactions, such as H-bonding, can cause strengthening or weakening of ligand–metal interactions. The overall charge as well as the electrostatic environment of the metal center, which is determined by the surrounding charge, dipole distribution, and solvent accessibility, also critically modulates the redox properties. 2.5.1 Role of the Heme Type It is known that c-type hemes tend to be found in cytochromes with more extreme potentials (much lower or much higher) relative to b-type hemes; however, it is unclear whether a direct causative relationship exists. One way to probe the role of the heme type in a way that is less dependent on other factors is to replace the heme in one protein with another. In studies of the de novo designed four-helix bundles, the strongest effect on reduction potential was attributed to the nature of the heme, 395 though unnatural hemes were used in the study. In the more natural protein cases, several groups have interconverted b- and c-type hemes. 445−449 It has been found, however, that this interconversion shows little inherent effect on the reduction potential 447,448 with no clear trend. For instance, it was found that converting the c-type heme in cyt c 552 into a b-type heme by mutating away the conserved Cys residues lowered the reduction potential by 75 mV. 447 On the other hand, introducing a thioether bond between heme in cytochrome b 5 and the protein, and therefore converting the b-type heme into a c-type heme, also lowered the potential by 23 mV. 448 It is clear that the choice of heme c over heme b has little effect on the reduction potential, and other effects, such as structural changes or solvent accessibility, may play a bigger role. If the choice of heme c or heme b does not play a significant role in determining the reduction potentials of cytochromes, one may wonder why organisms invest in the energetically expensive process of synthesizing c-type linkages. Though the exact reason that Nature has chosen c-type hemes in certain proteins remains to be fully understood, several hypotheses have been proposed. 454−456 It is suggested that multiheme cytochromes, such as c 3, with largely exposed hemes in close proximity may utilize heme anchoring as a strategy to ensure stable heme binding in the absence of well-defined hydrophobic interactions. 457 Similarly, the high-potential cyts c, with His/Met coordination, may use covalent anchoring as a strategy to prevent heme dissociation due to the relatively weaker binding of methionine to ferric heme. 457 Alternatively, it is proposed that covalent heme attachment may help in protein folding and stability 454,456 or may strengthen the Fe–His bond and help maintain a low-spin state. 456 Regardless, the choice of heme c over heme b likely does not itself directly tune the reduction potential in a significant or consistent way, but may allow the protein greater flexibility in achieving other functionality and tuning the potential by other means, such as solvent accessibility. In addition to hemes b and c, heme a is a unique heme used for ET in enzymes such as heme copper oxidases (HCOs). The heme incorporates two unique peripheral structural features, namely, a hydroxyethylfarnesyl group and a formyl group, and these functional groups have been suggested to play a role in tuning the reduction potential of the heme. While heme a has been replaced with other hemes in a native system, 458 detailed studies of how this substitution affects the redox chemistry of the protein have not been reported. Using their de novo designed scaffold (vide supra), Gibney and co-workers 459 have studied the redox properties of hemes a and b, as well as diacetyl heme, and found that the electron-withdrawing acyl groups increased the potential by ∼160 mV. This effect can be fully accounted for by the 200-fold lower affinity of the ligands for the oxidized form over the reduced form of the heme, and it is proposed that the hydrophobic farnesyl group serves to anchor the heme stably in the protein 460 to compensate for the lower affinity of the ferric state. 2.5.2 Role of Ligands In addition to the heme type, the identity of the axial ligands sets the baseline for the reduction potentials of cytochromes. 457 Between the two most common ligands (His and Met), it has been found that the Met ligation generally raises the potential of the heme by ∼100–150 mV, relative to the His ligation. 461−463 However, contrary to this theory, early work by Sligar and co-workers found that redesigning bis-His cyt b 5 into a His/Met cyt lowered the reduction potential by ∼240 mV. This opposite change in the reduction potential was attributed to the change in the spin state of the heme, from low-spin bis-His to high-spin His/Met cyt. 464 More consistent with the theory, it was demonstrated that conversion of bis-His to His/Met ligation in cyts c 3 results in a reduction potential increase of 160–180 mV. 192 Similarly, using a proteolytic fragment of cyt c, it was found that methionine ligation in cyts c contributes 130 mV to the energy. 386 Conversely, a 105 mV drop in the reduction potential was observed when the methionine in cytochrome c 551 was replaced with a histidine. 463 Interestingly, Hay and Wydrzynski 462 observed a 260 mV decrease in reduction potential when they substituted the native Met ligand in cyt b 562 with His, yielding a typical bis-His cyt. This decrease is greater than ∼150 mV, and the authors attribute it to destabilization of the fold and increased solvent exposure, which is known to significantly lower the potential (vide infra). In contrast, an Arg98Cys and His102Met double mutant of the same protein, cyt b 562, shows 6cLS bis-Met axial ligation at low pH, with a reduction potential of +440 mV, ∼180 mV higher than that of native His/Met cyt b 562. 465 The authors note that the effect of bis-Met ligation is likely to be slightly higher at ∼200 mV, as they expect the c-type thioether heme linkage to lower the potential. The stereochemical alignment of the axial methionine ligands results in an almost axial symmetry of the heme, caused by a 110° change in the torsion angle between the sulfur lone pairs. 466 The reduction potential of this protein is 665 mV higher than that of the only other known bis-Met axially ligated heme system in bacterioferritin (−225 mV) 176 in which the ground state of the oxidized form of the heme is highly rhombic in nature. 120,121,467 Therefore, factors other than the differences in the ligand coordination are most likely to be involved to account for the reduction potential difference. 78 In general, all else being equal, the preference of soft methionine thioether for the softer ferrous heme over the harder ferric heme contributes to a ∼100–200 mV increase in reduction potential over His ligation. 2.5.3 Role of the Protein Environment 2.5.3.1 Solvent Exposure Consistently, one of the most important factors in raising the reduction potentials of the cytochromes is the extent of heme burial in the protein or, alternatively, the extent of solvent exposure of the heme. 178,187,386,457,468−473 The basis for this effect lies in the lower dielectric constant of proteins relative to aqueous solution, which significantly destabilizes the charged ferric site over the neutral ferrous state of the heme. For instance, Tezkan et al. estimated that solvent exclusion accounts for ∼240 mV of the potential increase in cyt c. 386 Similarly, in a thorough computational study of heme proteins spanning an 800 mV range of potentials, Zheng and Gunner identified that heme solvent exclusion accounts for ∼20% of the reduction potential difference between proteins. 457 Interestingly, the same study found less correlation between the reduction potentials and the remaining individual factors or energy terms, yet the computation was able to faithfully reproduce and account for heme protein potentials over an 800 mV range. This study elegantly demonstrates that the reduction potential is determined by an intricate balance of numerous factors of comparable energy. 2.5.3.2 Secondary Coordination Sphere of the Ligand Although the nature of the ligand itself determines primary interaction energies with the heme, and therefore is the primary determinant of the reduction potential, the electronic character of the ligand can be further modulated by secondary noncovalent interactions, such as hydrogen bonds. These so-called secondary coordination sphere effects have been shown to be influential in determining the potentials of a number of heme proteins, including cytochromes. 230,472,474−477 For instance, in cyt c in particular, Bowman et al. demonstrated that strengthening the hydrogen bond between the proximal His ligand and a backbone carbonyl through peripheral mutations resulted in an almost 100 mV decrease in the reduction potential, attributable to increased imidazolate character. 474 Similarly, Berguis et al. show in three different mutants of yeast iso-1-cyt c that a disruption of the hydrogen bond from tyrosine 67 to the methionine ligand consistently decreases the potential by 56 mV, due to an increase in electron density on the Met sulfur, stabilizing the ferric form of the heme, 230,476 and Ye et al., found that the presence of hydrogen bonds between Gln64 and the axial Met ligand in Ps. aeruginosa and Hydrogenobacter thermophilus cyt c lowered the potential by 15–30 mV. 477 In addition, aromatic interactions with the axial ligand have also been implicated in tuning the heme reduction potentials. For instance, it was shown that Tyr43, which interacts with the π system of His 34, contributed a ∼35–45 mV decrease in reduction potential. 478 Therefore, although the identity of the ligand is a primary determinant of the reduction potential of the heme, the secondary coordination sphere interactions with it also play a role of similar magnitude in determining the reduction potential. 2.5.3.3 Local Charges and Electrostatics Another important means by which cytochromes have been found to modulate their reduction potentials is through the judicious use of charge and electrostatic interactions. For instance, by comparison and selective mutagenesis of the structurally homologous cyts c 6 and c 6A, it was demonstrated that the interaction of the positive dipole of the amide group of a carefully positioned glutamine (residues 52 in c 6 and 51 in c 6A) with the heme is a strategy used by Nature to raise the reduction potential by ∼100 mV. 479 Similarly, Lett et al. observed an increase in the reduction potential of cytochrome c by 117 mV through the Tyr48Lys mutation. 480 Tyr48 is involved in a H-bonding interaction with a heme propionate, and it is likely that introduction of lysine at this position stabilizes the propionate negative charge and destabilizes the ferric heme state. It has also been shown that replacement of a neutral residue in contact with the heme in myoglobin with a polar or negatively charged residue can reduce the potential by up to 200 mV. 481 Furthermore, a library screen of cytochrome b 562 mutants at four residues near the heme binding site identified mutations that could gradually tune the potential over a 160 mV range. 482 Even relatively distant surface electrostatic interactions have been shown to control the redox function of cytochromes. 483 These reports demonstrate the critical role of local charge in determining the reduction potential of the heme. In general, negative local charges stabilize the ferric state and lower the reduction potential, and the magnitude of this effect can be comparable to that of ligand substitution or ligand secondary coordination sphere effects. In addition to charge interactions, more subtle effects such as electrostatic interactions can also play an important role in determining redox properties. As discussed in section 5.2.2 below, a conserved aromatic residue in cyt b 6 f is found to be in contact with the heme f at position 4, and the identity of the aromatic residue differs between cyanobacteria and algae. Interconversion between Phe and Trp at this position accounts for about half of the 70 mV difference between these proteins. 161 The origin of this effect is attributed to differential interaction of the side chain electrostatic potentials with the porphyrin π system and the Fe orbitals. A similar effect has also been reported in cyt c 3, where a phenylalanine in contact with heme I is proposed to maintain its low potential by a π–π interaction with the porphyrin π system. 484 Since many charged residues around the heme, such as Glu, Asp, Lys, and Arg, as well as the heme propionate group itself, can be protonated or deprotonated depending on the pK a values of the residues and pH of the solution, protonation states of these groups will affect the reduction potential of the heme by preferentially stabilizing one redox state over the other. Therefore, the pH of the solution can have significant effects on the reduction potentials in various cytochromes. 342,485−490 For example, protonation of a heme propionate in cyt c contributed an increase of 65 mV to the reduction potential. 485 Similar effects of 60 and 75 mV have been reported in cyt c 551(491,492) and in cyt b 559, 490 respectively. In cyt c 2, pH-dependent reduction potentials covered a range of ∼150 mV, between pH 4 and pH 10. 493 In their de novo designed maquette, Dutton and co-workers observed a 210 mV range of reduction potentials over a pH range of 3.5–10, and such a change was attributed to the involvement of Glu residues near the heme site. 494 Furthermore, the role of the propionate charge has been investigated specifically by studies in which the carboxylate groups have been neutralized to their ester form. An increase of reduction potential by ∼60 mV was reported, 495,496 consistent with those obtained from the studies described above. A special case of the effect of local charges on reduction potential is the cooperativity between nearby hemes in multiheme cytochromes. 497 It is known that the presence of multiple hemes in various oxidation states greatly affects the macroscopic or observable reduction potentials of the hemes. For instance, it has been demonstrated in multiheme cyt c 3 that the interaction energy between hemes can shift the reduction potential by 50–60 mV. 498−500 It is suggested that this effect may be mediated by electrostatic interactions also involving local aromatic groups. 484 The cooperativity between hemes in multiheme cytochromes is proposed to be a major factor in their reduction potential regulation. In cyt c 3, the redox-Bohr effect can result in pK a differences of up to 2.8 pH units, and the coupling between protonation has been linked to cooperativity between the hemes, resulting in concerted two-ET steps. 340,501,502 On the other hand, the pH-dependent reduction potential difference, over a range of 10 pH units, can be ∼200 mV. 503 Such property is crucial for proper charge separation to generate a promotive force that drives ATP synthesis. 343,504 Similarly, this coupling of proton and ET plays a key role in the proton pumping mechanism of cytochrome c oxidase. Although there are several proposed mechanisms, they share the common theme that proton uptake to the heme sites and release into the P-side of the membrane are driven by charge compensation during ET events from the low-spin to high-spin heme. 505−507 It is clear that local electrostatic interactions at heme redox centers are of immense physiological importance. 2.5.3.4 Heme Distortion/Ruffling Another significant contributor to heme redox properties is the plasticity of the heme. It is now well-known that heme distortion or ruffling plays an important role in the electronic sturcture of the porphyrins, 508,509 due to decreased delocalization of the π electrons. 510−516 While the phenomenon has been described in many heme proteins, including cytochromes, 512,513,515,517,518 thorough investigation of how it affects redox properties is limited. Recently, Marletta and co-workers demonstrated that protein-induced heme distortion can account for up to a 170 mV increase in potential in the heme nitric oxide/oxygen binding protein. 513 Furthermore, a basic computational model was implemented by Senge and co-workers, and it was estimated that porphyrin distortion can account for 54 mV of the difference between hemes in a bacterial tetraheme cytochrome. 519 Further investigation is needed to gain a more detailed understanding of the role of heme distortion in the redox properties of typical cytochromes. 3 Fe–S Redox Centers in Electron Transfer Processes 3.1 Introduction to Fe–S Redox Centers Fe–S proteins are among the oldest metalloproteins on earth. The early atmosphere, under which both sulfur and iron were abundant, enabled the spontaneous assembly of these two elements into clusters, mainly containing four iron and four sulfur atoms. 91,520 Early life took advantage of the redox properties of these clusters and used them as redox centers. Despite the later shift to a more oxidizing environment on earth, the established Fe–S proteins continued to be used as electron carriers. Thus, these proteins are found ubiquitously throughout all kingdoms of life and play roles in crucial processes such as photosynthesis and respiration. The wide range of reduction potentials these proteins can accommodate and their diverse structural motifs allow them to interact with different redox partners, acting as electron carriers in a variety of biological processes. 91−93 The Fe–S proteins were first discovered in the 1960s on the basis of their unique g = 1.9 EPR signal that appears upon reduction and was not observed before for any metalloproteins. 9000−9002 This discovery was aided by the abundance of these proteins and their unique spectral features and often highly charged nature, which made them easier to purify and analyze. Studies of these proteins were further facilitated by advances in molecular biology and recombinant protein expression, allowing the use of site-directed mutagenesis to unravel important features of these proteins and their function. While the Fe–S centers are well-known for their function as electron carriers, they are also known to be involved in the active sites of many enzymes, performing several functions 521 such as reduction of disulfide bonds and initiation or stabilization of radical chain reactions, 523,525,529 or serving as Lewis acids. 524,527,543 In addition, the Fe–S centers can simply function as structural elements that stabilize the protein or another active site in the protein. 523,525,527,529 Furthermore, the sensitivity of the Fe–S centers to an oxidative environment and their range of redox states make them good candidates for sensing oxidative and metal stress and balancing the oxidative homeostasis of the cells. 93,525,526,527,530−533,543 Functions in DNA repair have also been reported for several Fe–S proteins. 532,534 Recently, a function for Fe-S proteins has been proposed in formation of FemoCo cluster. 522 Finally it has been shown that the Fe–S proteins can be used as a storage for sulfur or iron. 529,532 This review focuses exclusively on the ET function of the Fe–S proteins. 3.2 Classification of Fe–S Redox Centers and Their General Features The Fe–S clusters are often classified on the basis of the number of iron and sulfur atoms in the cluster, as suggested by the Nomenclature Committee of the International Union of Biochemistry (IUB) in 1989. 535 In this convention, the elements of the core cluster (iron and inorganic sulfur atoms) are placed in brackets with the oxidized level of the core cluster shown as a superscript outside the brackets (e.g., [2Fe–2S]2+). A comma or a slash in the superscript can show multiple possible oxidation states. A more expanded notation can be used to show the ligands and the overall charge of the whole cluster, including those ligands. Another common classification of Fe–S clusters, which is used in this review, is based on the protein type. This scheme classifies the Fe–S centers on the basis of not only the number of iron and sulfur atoms but also certain structural motifs and spectroscopic and electrochemical properties. In this classification, the Fe–S proteins are divided into major groups as follows: rubredoxins (Rd’s; [1Fe–4S]), ferredoxins (low-potential [2Fe–2S], [4Fe–4S], [3Fe–4S], [3Fe–4S][4Fe–4S], and [4Fe–4S][4Fe–4S]), Rieske proteins (which are high-potential [2Fe–2S] proteins), and high-potential iron–sulfur proteins (HiPIPs, which are high-potential [4Fe–4S] proteins) (Table 3). In addition, we will also describe more complex Fe–S proteins that contain multiple Fe–S cofactors or Fe–S cofactors coupled with other cofactors, such as heme. 92,93,523,526,529,536−540 Table 3 Classification of Fe–S Proteins Though certain structural elements may differ between them, members of each class of Fe–S proteins usually consist of a common structural motif. Between classes the overall structure is distinct. Despite these overall structural differences, however, the geometries of the Fe–S clusters are quite similar, especially within each cluster class. The iron cofactor has a distorted tetrahedral geometry in almost all the Fe–S proteins. In the case of proteins with more than one iron, the S–S distance is usually 1.3 times longer than the Fe–Fe distance. 523 Each iron atom is coordinated by a total of four ligands, typically cysteine or inorganic sulfurs, although other ligands have been observed. For instance, in Rieske proteins, two cysteine ligands have been replaced with histidines. In some [3Fe–4S] clusters, an aspartate serves as a ligand to iron. In certain enzymes such as aconitase, a hydroxyl group from the solvent is shown to be one of the ligands. 541 While the geometry of Fe and its coordinating cysteine/sulfur ligands is very similar in all Fe–S proteins, the amino acid sequences and peptide motifs that accommodate these clusters differ significantly even in a given class, resulting in further categorization of each group. Interestingly, the ligands of the Fe–S proteins usually reside within loop regions. This structural flexibility is important in accommodating the geometric requirement of the Fe–S clusters and thus minimizing the reorganization energy required for rapid ET. The iron site has large spin-polarization effects, strong Fe–S covalency, and spin coupling through inorganic sulfurs. 542 The strong covalency and the delocalization features of Fe–S proteins result in a low reorganization energy, mostly by lowering the inner sphere effects. Gas-phase DFT calculations give the following reorganization energies for different Fe–S proteins in vacuum: 0.41 eV (1Fe, Rd) Fe(IV) ≈ Fe(II) > Fe(I), so only two states are observable in the oxidized state of HiPIPs, which explains the presence of two electronic isomers observed in NMR and EPR. 884 NMR of the oxidized pair shows two downfield signals arising from the mixed-valence pair and two upfield signals (or extrapolated upfield, which is two downfield signals with anti-Curie temperature dependence) assigned to the ferric pair with inverted electron polarization. 895,931 1H 2D exchange spectroscopy (EXSY) NMR studies have analyzed self-exchange rates for HiPIP from Ch. vinosum and its aromatic mutants. An exchange rate of 2.3 × 104 M–1 S–1 was observed for the native protein at 298 K, with rates within 2-fold for the mutants. This study ruled out the role of aromatic residues in ET. 876 β protons from cysteine ligands of the cluster experience large contact shifts. Eight signals from +110 to −40 ppm can be assigned to eight protons from four β-CH2 Cys ligands. The assignment of protons that are involved in amide–S H-bonds is more difficult due to their broad features that overlap with other protons. 929,932 NMR experiments have also been used to assess water accessibility of the cluster and its mutants through analyzing the H2O/D2O exchange rates. 1H–13C heteronuclear correlation (HETCOR) NMR was used to show that the oxidized cluster has an overall shorter relaxation time than the reduced state. 933 EPR of HiPIPs shows a nearly axial signal with g values at 2.13 and 2.03 that result from an S = 1/2 ground state in the oxidized form. 934 In contrast to ferredoxins, HiPIPs are EPR-silent in their reduced state. Some HiPIPs show heterogeneous signals, probably due to sample preparation or dimerization of the cluster. 799 ENDOR studies confirmed the presence of two pairs of irons in the oxidized form of the protein. 935,936 EPR of most HiPIPs has shown at least two populations. Four species can be observed by EPR of HiPIPs with g ⊥ = 2.15–2.13, 2.13–2.11, 2.06–2.08, and maybe 2.09–2.11, with the first two often being the most dominant species. 872 Assignment of these two species can be performed by correlating the EPR data with room temperature 1H NMR. Zero-field Mössbauer studies of HiPIPs at temperatures above 100 K show a broad quadruple splitting, indicative of fast electronic relaxation, with δ = 0.29–0.33 mm/s and quadruple splitting values of 0.74–0.80 mm/s. At lower temperature (4.2 K) the spectra show two nonequivalent iron pairs, one of which increases quadruple splitting with increased applied field, whereas the other decreases quadruple splitting. The subsets are assigned to a ferric pair (δ = 0.27 mm/s, with a −0.87 mm/s splitting) and a ferric–ferrous pair (δ = 0.37 mm/s with a splitting value of −0.94 mm/s). 895 Mössbauer shows nondistinguishable iron atoms in reduced HiPIPs. Mössbauer studies of mutated Cys → Ser HiPIP have shown loss of covalent iron features due to replacement of S with O and a different spectrum of the Ser-bound iron in the reduced form, suggesting the importance of Cys residues in maintaining the mixed-valence state of the cluster. 937 Mössbauer analyses of partially unfolded HiPIPs have found a slight increase in Fe–S bond distances without significant changes in the core cluster, indicating that the cluster is not denatured in early steps of unfolding. 529,938 EXAFS analysis of the structure of the core cluster of HiPIPs and Fe–S distances has found a small temperature dependence. Analyses of Cys → Ser mutants reveal slight changes to the core structure and the Fe–S distances of intact cysteines, while the Fe–O bond is shortened, suggesting that the entire cluster is shifted toward the Ser ligand. 937 Ligand K-edge XAS studies have also elucidated some of the differences between HiPIPs and ferredoxins. 900 3.4.6 Complex Fe–S Centers 3.4.6.1 Hydrogenases 3.4.6.1.1 [NiFe] Hydrogenase Cluster [NiFe] hydrogenases catalyze interconversion of H2 and H+ in microorganisms and ultimately provide electrons for ATP synthesis. [NiFe] hydrogenases from different sources have a conserved large domain of ∼60 kDa, containing the binuclear Ni–Fe active site and a small Fe–S cluster domain for ET. [NiFe] hydrogenase from Dv. gigas contains two [4Fe–4S] clusters and one [3Fe–4S] cluster, supported by EPR, Mössbauer, 939 and crystallographic studies. 940,941 The reduction potentials are −70 mV for the [3Fe–4S]+,0 cluster and −290 and −340 mV for the two flanking [4Fe–4S]2+,1+ clusters. The fully oxidized state of the two clusters ([4Fe–4S]2+) gives an isomer shift of 0.35 mm/s and quadruple splitting of 1.10 mm/s. Upon reduction, the two clusters are separated. Cluster I gives an isomer shift of 0.525 mm/s and quadruple splitting of 1.15 mm/s, and cluster II gives 0.47 and 1.35 mm/s, respectively. The parameters of [3Fe–4S]1+ are δ = 0.47 mm/s and ΔE Q = 1.67 mm/s, and those of [3Fe–4S]0 are δ = 0.39 mm/s and ΔE Q = 0.38 mm/s. The three Fe–S clusters are arranged linearly in the 3-D structure, with one [4Fe–4S] cluster proximal to the Ni–Fe–S catalytic center, the other [4Fe–4S] cluster at the surface, and the [3Fe–4S] cluster in the middle of them (Figure 35), 940,941 suggesting the existence of an ET pathway. Figure 35 Proposed ET pathway in Dv. gigas [NiFe] hydrogenase. Selected distances are given in angstroms. PDB ID 1FRV. Color code: Fe, green; Ni, gray blue; C, cyan; S, yellow, O, red; N, blue. Reprinted with permission from ref (940). Copyright 1995 Macmillan Publishers Ltd. [NiSeFe] hydrogenase, a subclass of [NiFe] hydrogenases, contains three [4Fe–4S] clusters. 942,943 The crystal structure reveals that a cysteine residue near the middle cluster, as opposed to proline usually observed in [NiFe] hydrogenases, serves as an extra ligand and results in a [4Fe–4S] cluster instead of a [3Fe–4S] cluster . [NiFe] hydrogenase from Dv. fructosovorans is structurally similar to that from Dv. gigas. 944 On the basis of observations made with respect to [NiSeFe] hydrogenases, a Pro238Cys mutation has been made. The [3Fe–4S]1+,0 cluster was successfully converted to a [4Fe–4S]2+,1+ cluster and resulted in a 300 mV decrease of the reduction potential with little influence on activity, indicating that the [3Fe–4S]1+,0 cluster is not essential in the ET pathway of [NiFe] hydrogenase. Recently, a new type of [NiFe] hydrogenase was discovered. Unlike the usually air-sensitive members of the family, [NiFe] hydrogenases from the bacteria Ralstonia eutropha, Ralstonia metallidurans, Hydrogenovibrio marinus, and Aquifex aeolicus could tolerate O2 to a limited extent. 947 The oxygen tolerance arises from neither modification of the [Ni–Fe] active site nor limited access to O2. Crystal structures of the proteins have revealed a novel Fe–S cluster proximal to the Ni–Fe center (Figure 36a). 948,949 Instead of the normal proximal [4Fe–4S] cluster coordinated by four cysteines from the protein, this cluster is a plastic [4Fe–3S] cluster bound by six cysteines with a flexible glutamic acid residue nearby. Upon oxidation, the backbone amide of the coordinating Cys26 is deprotonated by the nearby glutamic carboxylate and replaces the bridging Cys25 (Figure 36b,c), analogous to the P cluster in nitrogenases. The negative charge of amide will help to stabilize the oxidized state. As a result, the [4Fe–3S] cluster could transfer two electrons in a window of 200 mV and remain stable in three oxidation states. 950 DFT calculations have revealed that the supernumerary coordination frame provided by the six cysteines and the flexible coordination sphere of the Cys26-bound Fe lead to plasticity of the unique proximal [4Fe–3S] cluster and, consequently, low reorganization energy in the reduced state. 945 Hence, the proximal cluster could not only transfer electrons efficiently from the active site during H2 oxidation, but also rapidly supply two electrons to the active sites upon O2 binding, which in combination with one electron from the middle [3Fe–4S] cluster would efficiently reduce O2 to H2O and prevent formation of an inactive [Ni3+– –OOH–Fe2+] cluster, the so-called Ni-A state, and overoxidation by O2. 951−953 Figure 36 (a) Crystal structure of O2-tolerant membrane-bound hydrogenase from Ralstonia eutropha (PDB ID 3RGW). Reprinted from ref (945). Copyright 2013 American Chemical Society. (b) Reduced [4Fe–3S] cluster from MBH (PDB ID 3AYX) (Reprinted with permission from ref (946). Copyright 2012 Wiley-VCH) and (c) oxidized [4Fe–3S] cluster from MBH (PDB ID 3AYZ). Reprinted with permission from ref (946). Copyright 2012 Wiley-VCH. Color code: Fe, green; C, cyan; S, yellow; N, blue; Ni, orange. 3.4.6.1.2 [FeFe] Hydrogenases [FeFe] hydrogenases share a conserved catalytic subunit binding metal cluster, called the H-cluster, as the catalytic site and have various Fe–S subunits harboring different Fe–S clusters for ET to and from the H-cluster. The Fe–S domains are usually located at the N-terminus of the catalytic domain and contain [4Fe–4S] or [2Fe–2S] binding motifs similar to those of ferredoxins. 954−956 For example, [FeFe] hydrogenase from Dv. desulfuricans ATCC 7757 possesses two [4Fe–4S] clusters for ET, 957 and the protein from Cl. pasteurianum contains one [2Fe–2S] cluster and three [4Fe–4S] clusters. 958 The Fe–S clusters in Cl. pasteurianum [FeFe] hydrogenase are separated by 8–11 Å, indicating potential ET pathways through covalent bonds or a H-bonding network (Figure 37). The FS4C and FS2 near the protein surface possibly function as the initial electron acceptors of external electron donors and transfer electrons to the FS4B at the junction position. The FS4A is 10 Å from cluster FS4B and 9 Å from the H-cluster and could mediate sequential ET to and from the catalytic site. Figure 37 (a) Location of Fe–S clusters in [FeFe] hydrogenase (PDB ID 1FEH). (b) Proposed ET pathways for [FeFe] hydrogenase. Reprinted with permission from ref (958). Copyright 1998 American Association for the Advancement of Science. 3.4.6.2 Molybdonum-Containing Enzymes 273 3.4.6.2.1 [4Fe–4S] Cluster and P-Cluster in Nitrogenase Four types of nitrogenases have been discovered: two containing Mo and Fe, one containing V and Fe, and one containing only Fe in the catalytic site in a large domain with a molar mass of 220–250 kDa. Among them, [FeMo] nitrogenase has been the most extensively studied (Figure 38a). Besides the active site, all nitrogenases contain an iron protein as α2 dimers with a molar mass of 60–70 kDa. It contains a single [4Fe–4S] cluster between the two monomers, which is coordinated by one conserved cysteine from each monomer and is exposed to water. 959 The cluster transfers electrons efficiently via a MgATP hydrolysis reaction at the larger domain containing a catalytic site, along with other functions, including involvement in biosynthesis and insertion of FeMoco into [FeMo] nitrogenase and regulation of biosynthesis in other nitrogenases. 960 Figure 38 (a) Overall structure of nitrogenase (PDB ID 1N2C). Cofactors are shown as spheres and denoted. Reprinted with permission from ref (965). Copyright 1997 Macmillan Publishers Ltd. (b) Reduced P cluster from nitrogenase (PDB ID 3U7Q) (Reprinted with permission from ref (946). Copyright 2012 Wiley-VCH.) and (c) oxidized P cluster from nitrogenase (PDB ID 2MIN). Reprinted with permission from ref (946). Copyright 2012 Wiley-VCH. Three oxidation states, +2, +1, and 0, have been observed for the [4Fe–4S] cluster, indicating that the cluster could transfer one or two electrons to the catalytic domain. The reduction potential to achieve an all-ferrous [4Fe–4S]0 cluster is −460 mV, and this is the first example of this oxidation state for [4Fe–4S] clusters, both in proteins and in model complexes. 961−963 EXAFS studies show that changes of the Fe–S and Fe–Fe distances are less than 0.02 Å from the [4Fe–4S]2+ cluster to the [4Fe–4S]1+ cluster. 964 The Fe protein can bind 2 equiv of MgATP or MgADP, each in a Walker A motif on one monomer. The Walker A binding site is 15–20 Å away from the [4Fe–4S] cluster with a series of salt bridges and H-bonds in between. However, the reduction potential of the [4Fe–4S] cluster decreases ∼100 mV upon binding of either nucleotide, possibly arising from protein conformational changes induced by binding and hydrolysis reactions. 965−970 The reduction potential change is proposed to be the driving force for ET. 968 UV–vis, resonance Raman, and EPR spectroscopic studies indicate that the [4Fe–4S] cluster could reversibly cycle between a regular [4Fe–4S] cluster in the reduced state and two [2Fe–2S] clusters in the oxidized state. 971 The [FeMo] domain contains the FeMoco cluster and a P-cluster. The FeMoco cluster is the catalytic center and will not be discussed here. The P-cluster is situated at the interface of the α and β subunits of the [FeMo] domain. It is an [8Fe–7S] cluster, with a 6-coordinate sulfur at the center. The structure of the P-cluster changes with the oxidation state. The dithionite reduced P cluster (PN) is bound by six cysteines from the protein, four of which coordinate a single iron, and the remaining two function as bridging ligands (Figure 38b). 972 After two-electron oxidation of PN, a form called Pox is obtained. In the Pox cluster, the coordination between the center 6-coordinate sulfur and two irons associated with the β subunit is replaced by the amide N of Cys88 of the α subunit and side chain hydroxyl of Ser188 of the β subunit (Figure 38c), similar to the changes of oxygen-tolerant [NiFe] hydrogenases mentioned above (see Figure 36). The changes are proposed to be related to the proton-coupled electron transfer process in nitrogenases. 972−974 3.4.6.2.2 Aldehyde Oxidoreductases Aldehyde oxidoreductase belongs to the molybdoflavoenzymes. It is a homodimer and usually requires Fe–S clusters, a molybdopterin or tungstopterin site, and sometimes an FAD cofactor for substrate oxidation. Aldehyde oxidoreductase from Dv. gigas is composed of four domains, including two small N-terminal domains binding two types of [2Fe–2S] clusters and two large domains containing the molybdopterin cofactors. 975,976 The first Fe–S domain (residue 1–76) is similar to that of spinach ferredoxins, and the [2Fe–2S] cluster is coordinated by Cys40, Cys45, Cys47, and Cys60. The second Fe–S domain (residues 84–156) is a four-helix bundle, and the [2Fe–2S] cluster is coordinated by Cys100, Cys103, Cys137, and Cys139. The molybdopterin is 15 Å from the surface and 14.9 Å from the Fe–S cluster of the second domain. Recently, the crystal structure of aldehyde oxidase of mouse liver has been reported. The overall fold is very similar to that from Dv. gigas, but that of the mammalian protein has an additional FAD domain. 977 EPR studies revealed two types of [2Fe–2S] clusters, named Fe–SI and Fe–SII. 978−981 Fe–SI is observable at 77 K with g values of 2.021, 1.938, and 1.919, while Fe–SII is only observable below 40 K with g values of 2.057, 1.970, and 1.900. The reduction potentials of Fe–SI and Fe–SII are −260 and −280 mV, respectively. In the presence of the substrate benzaldehyde, partial reduction of the Fe–S clusters has been detected in Mössbauer studies, indicating participation of the Fe–S clusters in the catalytic reaction and fast ET from the molybdopterin center. 982 3.4.6.3 Ni-Containing CO Dehydrogenase and Hybrid Cluster Protein 3.4.6.3.1 Ni-Containing CO Dehydrogenase CO dehydrogenases (CODHs) catalyze oxidation of CO to CO2 along with dehydrogenation of water and release of protons and electrons. It is important in the oxygen-based respiratory process in hydrogenogenic bacteria. There are two types of CODHs. One is Mo-based CODHs with a mono-Mo cofactor coordinated by dithiolene sulfurs of a pterin ligand found in aerobic organisms, which is beyond the scope of this review but has been reviewed extensively in other papers. 983,984 The other is Ni-containing CODHs with a Ni–Fe–S cluster as well as multiple Fe–S clusters found in anaerobic organisms 985−987 and will be discussed briefly below. Ni CODHs are β2 homodimers. 988,989 Each monomer contains a Ni–Fe–S cluster (cluster C) as the catalytic site and a [4Fe–4S] cluster (cluster B). In addition, another [4Fe–4S] cluster (cluster D) is situated at the interface of the two monomers and coordinated by residues from both monomers (Figure 39a). Clusters B and D transfer electrons between cluster C and external redox regents. They also bind acetyl-CoA synthases to form α2β2 bifunctional enzymes acetyl-CoA synthases/carbon monoxide dehydrogenases (ACSs/CODHs). 990 Two additional [4Fe–4S] clusters, E and F, have been found in an extra subunit of the ACS/CODH complex. 991 The crystal structure of Ni CODH from Carboxydothermus hydrogenoformans reveals that cluster C is a [Ni–4Fe–5S] cluster (Figure 39b). The geometries of the irons are approximately tetrahedral, and that of Ni is close to square planar. It is associated with the protein through four cysteines and one histidine. 988 On the other hand, the structures of Rhodospirillum rubrum Ni CODHs 989 and the M. thermoacetica ACS/CODH complex 991 show cluster C as [Ni–4Fe–4S], coordinated similarly by five cysteines and one histidine from the protein (Figure 39c). The Ni is also coordinated by an external nonprotein ligand. Figure 39 (a) Crystal structure of Rs. rubrum Ni CODH. Clusters are shown as spheres. PDB ID 1JQK. (b) [4Fe–5S–Ni] cluster C of Ca. hydrogenoformans Ni CODH. PDB ID 1SU8. (c) [4Fe–4S–Ni] cluster C of M. thermoacetica Ni CODH. PDB ID 1MJG. Reprinted with permission from ref (990). Copyright 2011 Elsevier. 3.4.6.3.2 Hybrid Cluster Proteins Hybrid cluster proteins (HCPs) are a type of Fe–S proteins with unknown functions. However, they have been detected in more than 15 bacteria and archaea. There are three categories of HCPs. The first is found in anaerobic bacteria such as Dv. vulgaris and Dv. desulfuricans or methanogen archeon Methanococcus jannaschii, with coordinating cysteines arranged in the sequence Cys-(Xxx)2-Cys-(Xxx)7–8-Cys-(Xxx)5-Cys. The second is found in facultative anaerobic Gram-negative bacteria such as E. coli, Morganella morganii, or Tb. ferrooxidans, with the sequence Cys-(Xxx)2-Cys-(Xxx)11-Cys-(Xxx)6-Cys. The third is found in (hyper)thermophilic bacteria or archaea, including Methanobacterium thermoautotrophicum, Pyrococcus abyssi, or Tt. maritima, with the same sequence arrangement as the first category but with smaller size due to residue deletion downstream of the N-terminal cysteine region. HCP from Dv. vulgaris contains three domains (Figure 40a). 992,993 A [4Fe–4S] cluster is bound to domain 1 by Cys3, Cys6, Cys15, and Cys21 from the N-terminal region, similar to the cubane cluster in ferredoxins except that no cysteine is from the C-terminal region. This Cys-(Xxx)2-Cys-(Xxx)8-Cys-(Xxx)5-Cys motif is conserved in all HCPs, and HCPs from both categories 1 and 3 contain a [4Fe–4S] cluster linked by this motif. HCPs from category 2, on the other hand, might instead have two [2Fe–2S] clusters at this position. 994 Figure 40 Hybrid clusters in HCP. (a) Overall structure of as-isolated Dv. vulgaris HCP. Metal clusters are shown as spheres. PDB ID 1W9M. (b) Superposition of Dv. vulgaris HCP (cyan) and NiCODH (red, PDB code 1SU7). (c) Hybrid cluster in the as-isolated oxidized form of Dv. vulgaris HCP prepared anaerobically. PDB ID 1W9M. (d) Hybrid cluster in the reduced form of Dv. vulgaris HCP. PDB ID 1OA1. Residue backbones are omitted for clarity. Bonds inside the cluster are shown as dotted lines, and bonds between residues and the cluster are shown as solid lines. Color code: Fe, green; C, cyan; S, yellow; O, red; N, blue. Reprinted with permission from ref (995). Copyright 2008 International Union of Crystallography. HCPs also contain a unique hybrid cluster, [4Fe–2S–3O], which was isolated in the oxidized form from Dv. vulgaris HCP (Figure 40c), 995 and [4Fe–3S] with a water molecule between Glu494 and His244 in the reduced form (Figure 40d). 996 In the former state, the cluster is linked to the protein by Cys12, Cys434, Cys459, thio-Cys406 (Cys with an additional S on the S(Cys), called Css406), His244, Glu268, and Glu494, and in the latter case Css406 is reduced to cysteine. The EPR signal of HCP is similar to that of the prismane model complex (Et4N)3[Fe6S6(SC6H4-p-Me)6]3+. 997 Therefore, the four oxidation states of the hybrid cluster are named analogously to those of the prismane complex as “3+”, “4+”, “5+”, and “6+”. The midpoint reduction potentials of the Dv. vulgaris HCP hybrid cluster range from −200 to +300 mV at pH 7.5. 998 It is noteworthy that HCPs demonstrate a high degree of similarity to Ni CODHs. 992,993,999 They not only share similar overall folding, but also exhibit similar cluster positions and structures inside the monomer (Figure 40b). The closest distance between the [4Fe–4S] cluster and hybrid cluster is 10.9 Å, with Tyr493, Thr71, Asn72, and Glu494 in between. In addition, two tryptophan residues, Trp292 and Trp293, are located between the hybrid cluster and the protein surface. The arrangements indicate possible ET pathways, yet no involvement in such processes has been detected so far. The protein can be reduced by NAD(P)H oxidoreductase, 994 but there is no genomic evidence for the existence of a similar redox partner in the sources from which HCP has been detected or isolated. 3.4.6.4 Siroheme Fe–S Proteins Siroheme is an iron-containing reduced tetrahydroporphyrin of the isobacteriochlorin class (Figure 41a). Siroheme proteins are a type of iron–sulfur protein containing a siroheme conjugated to a [4Fe–4S] cluster through a thiolate bridge. 1000 Siroheme is the catalytic center, and the [4Fe–4S] cluster serves as an electron trapping and storage site. Siroheme proteins includes sulfite reductases and nitrite reductases, and they are important in assimilation and dissimilation of sulfite and nitrite. 1001,1002 Figure 41 (a) Structure of siroheme. (b) Siroheme and the [4Fe–4S] cluster of spinach nitrite reductase. PDB ID 2AKJ. Color code: Fe, green; C, cyan; S, yellow; O, red; N, blue. 3.4.6.4.1 Nitrite Reductase NiR catalyzes the six-electron reduction of nitrite to ammonia. It exists in both eukaryotes and prokaryotes. There are two types of NiR categorized by the physiological electron donor: ferredoxin-dependent NiR in photosynthetic organisms and NAD(P)H-dependent NiR in most heterotrophic organisms. 276,1003−1005 Ferredoxin-dependent NiR contains a siroheme and a [4Fe–4S] cluster, while NAD(P)H-dependent NiR contains an additional FAD cofactor bound at an extended N-terminal region. 276 Spinach nitrite reductase is a type of ferredoxin-dependent NiR isolated from higher plants. It is composed of 594 amino acids divided into three α/β domains. The siroheme cofactor is situated at the interface of the three domains and bridged to the [4Fe–4S] cluster via Cys486 (Figure 41b). The [4Fe–4S] cluster is also coordinated by Cys441, Cys447, and Cys482. The midpoint reduction potentials are −290 mV for the siroheme and −365 mV for the [4Fe–4S] cluster. Although the two cofactors are magnetically coupled with a distance of 4.2 Å, they are independent in redox titration processes. 1006,1007 Spinach NiR can form a 1:1 complex with ferredoxin with electrostatic interactions between acidic residues from NiR and basic residues from ferredoxin. The interprotein ET chain has been established as from photoexcited photosystem I via the [2Fe–2S] cluster of ferredoxin to the [4Fe–4S] cluster of NiR followed by intraprotein transfer to the siroheme. 1006−1008 3.4.6.4.2 Sulfite Reductase Sulfite reductase catalyzes the six-electron reduction of sulfite to sulfide in biological systems and can be categorized as assimilatory sulfite reductase (aSiR) or dissimilatory sulfite reductase (dSiR). aSiR reduces sulfite directly to sulfide, while dSiR provides a mixture of sulfide, trithionate, and thiosulfate in in vitro experiments. 1009 The aSiRs are found in archaebacteria, bacteria, fungi, and plants. 1010,1011 Assimilatory ferredoxin-dependent sulfite reductases from plant chloroplasts and cyanobacteria are soluble monomeric proteins with molar masses of ∼65 kDa. They contain a siroheme linked to a [4Fe–4S] cluster structurally similar to those in nitrite reductase, and they undergo reduction by ferredoxin from photoreduced photosystem I as well. 1002 They can also catalyze the reduction of nitrite to ammonia, the reaction catalyzed by NiR, but with a higher K M for nitrite than sulfite, further demonstrating the significant similarity of the two types of enzymes. 1002,1012,1013 For maize sulfite reductase, the midpoint potentials of siroheme and the [4Fe–4S] cluster have been determined to be −285 ± 5 and −400 ± 5 mV, respectively, at pH 7.5 in Tris buffer by redox titrations. Although the E° of the [4Fe–4S] cluster is more negative than that of spinach nitrite reductase (E° = −375 ± 10 mV at pH 7.5 in Tris buffer), reduction by ferredoxin (E° = −430 mV) is still a thermodynamically favorable process. In the presence of cyanide, the E° of siroheme shifts positively to −155 ± 5 mV, while that of the [4Fe–4S] cluster shifts negatively to −455 ± 10 mV, possibly due to decreased affinity of the enzyme for cyanide upon reduction of the [4Fe–4S] cluster. Similar trends are observed in spinach nitrite reductase as well. 1014 The aSiR from E. coli is a 780 kDa hemeoflavoprotein with an α8β4 arrangement. The α subunit, known as sulfite reductase flavoprotein, contains FAD and FMN, while the β unit, named sulfite reductase hemoprotein, harbors the associated [4Fe–4S] cluster and siroheme. The ET pathway is in the FAD–FMN–[4Fe–4S]–siroheme sequence, with NADPH as the initial donor and sulfite as the terminal acceptor. 1015 dSiRs exist in sulfate reducing microorganisms. 1010,1011 dSiR is composed of two types of subunits, DsrA and DsrB, generally in a heterotetrametric α2β2 arrangement with similar overall folds for all dSiRs from different sources. 1016,1017 Some dSiRs form a complex with two additional subunits of DsrC and result in an α2β2γ2 arrangement. The dSiR contains eight [4Fe–4S] clusters together with four sirohemes or two sirohemes and two sirohydrochlorins (the metal-free form of siroheme) (Figure 42a,b), and only two of the four sites are catalytically active. In Dv. gigas, desulfoviridin, a subcategory of dSiR, a [3Fe–4S] cluster is associated with the siroheme instead of a [4Fe–4S] cluster in one active form, DsrII (Figure 42c). The relative position of siroheme and the [4Fe–4S] cluster is similar to that in aSiRs, and both the [4Fe–4S] clusters proximal to and remote from the siroheme are coordinated by four cysteines from the protein. 1018−1020 Figure 42 (a) Siroheme group and [4Fe–4S] cluster of DsrI. PDB ID 3OR1. (b) Sirohydrochlorin group and [4Fe–4S] cluster of DsrII. PDB ID 3OR2. (c) Siroheme group and [3Fe–4S] cluster of DsrII. PDB ID 3OR2. Color code: Fe, green; C, cyan; S, yellow; O, red; N, blue. 3.4.6.5 Respiratory Complex Chain The mitochondrial respiratory system is the main energy producer in eukaryotic cells. 1021,1022 It consists of five membrane complexes, complex I, 1023 complex II (succinate dehydrogenase), 1024,1025 complex III (cytochrome bc 1 complex), 1026−1029 complex IV (cytochrome c oxidase complex), 1030,1031 and complex V (ATPase). 1032 The first four complexes are located on the inner membrane and function by transferring electrons from electron donors, NADH and succinate, to the final electron acceptor, oxygen, and meanwhile pump protons across the membrane. This proton gradient is utilized by ATPase to generate ATP. 3.4.6.5.1 Respiratory Complex I Respiratory complex I (CI), also known as NADH:ubiquinone oxidoreductase or NADH dehydrogenase, is involved in one of the ET pathways of the respiratory chain. It is composed of the following steps: (1) NADH donates electrons through CI to reduce ubiquinone to ubiquinol. (2) Ubiquinol transfers electrons through complex III to cytochrome c. (3) Cytochrome c is oxidized by complex IV and transfers electrons to O2 to produce water. In this process, each electron transferred is associated with five protons pumped from the matrix to the inner membrane space. Although CI is the most complicated complex in the mitochondrial respiratory chain, important breakthroughs have been achieved, and multiple structures have been reported recently. 1023,1033−1036 Mammalian CI (∼980 kDa) is composed of up to 45 different subunits, including 7 subunits in hydrophilic parts harboring one FMN and eight Fe–S clusters, 7 subunits in transmembrane parts, and ∼30 accessory subunits. 1022,1037 Bacterial NADH dehydrogenase (∼550 kDa) only contains 13–16 subunits, which is sufficient for complete CI function as well. 1023,1038−1040 The crystal structure of the hydrophilic part of complex I from T. thermophilus(1023) reveals for the first time the main ET pathway of the protein as shown in Figure 43: electrons from NADH are transferred through FMN to N3, followed by N1b, N4, N5, N6a, and N6b sequentially, and finally through N2 to ubiquinone coupled with proton translocation. 1022 Figure 43 Crystal structure of mitochondrial respiratory complex I from T. thermophilus. PDB ID 4HEA. Cofactors involved in the ET pathway are shown on the right side with distances and directions denoted. Reprinted with permission from ref (1022). Copyright 2013 Elsevier. 3.4.6.5.2 Respiratory Complex II (Succinate Dehydrogenase) and Fumarate Reducatse Complex II in the respiratory chain (CII), also known as succinate dehydrogenase (SDH) or succinate:quinone reductase, is a membrane-bound protein involved in the citric acid cycle and the second ET pathway in the mitochondrial respiratory chain. In the mitochondrial respiratory chain, electrons are transferred from succinate to ubiquinone through complex II, then to cytochrome c through complex III, and finally to O2 through complex IV. This process is less efficient than the process associated with complex I, and each electron transferred will pump only three protons across the membrane. CII catalyzes oxidation of succinate to fumarate by a hydrophilic catalytic domain composed of a large flavoprotein (Fp; 65–79 kDa) with a covalently bound FAD cofactor and an iron–sulfur protein (Ip; 25–37 kDa) containing [2Fe–2S] (center S1), [4Fe–4S] (center S2), and [3Fe–4S] (center S3) clusters. 1024,1025,1041 The catalytic domain is anchored to the membrane by one or two hydrophobic domains (CybL, CybS) harboring usually b-type cytochromes (Figure 44). The [2Fe–2S] center is coordinated by four cysteines close to the N-terminus, and the [4Fe–4S] and [3Fe–4S] clusters are coordinated near the C-terminus by two cysteine-containing sequences: Cys-(Xxx)2-Cys-(Xxx)2-Cys-(Xxx)3-Pro and Cys-(Xxx)2-Xxx-(Xxx)2-Cys-(Xxx)3–Cys-Pro (Xxx = Ile, Val, Leu, or Ala), similar to 7Fe ferredoxins. The [4Fe–4S] cluster usually has a low reduction potential and functions as the energy barrier of the ET process to direct the electron flow and, consequently, the reaction pathway. 1042 The [3Fe–4S] cluster is involved in a direct ET process from the initial electron donor quinones. 1043−1045 The midpoint reduction potential of the [3Fe–4S]1+,0 cluster is in the range of +60 to +90 mV, and the potential of the initial electron donor ubiquinone is +65 mV. 1046 SDH from Sl. acidocaldarius contains a [4Fe–4S] center instead of a [3Fe–4S] center for cluster S2 and displays poor reactivity toward caldariella quinone. 1047 Figure 44 Crystal structure of mitochondrial respiratory complex II. FAD binding protein (Fp) is shown in blue, iron–sulfur protein (Ip) is shown in cream, hydrophobic domains are shown in pink and orange, and the putative membrane is shown in gray shading. PDB ID 1ZOY. Cofactors involved in the ET pathway are shown on the right side, with distances, reduction potential, and directions denoted. Reprinted with permission from ref (1024). Copyright 2005 Elsevier. It is noteworthy that heme b (E° = +35 mV) in the hydrophobic domain of SDH is not involved in the ET pathway mentioned above. It is proposed that heme b in SDH of E. coli functions as an electron sink and reduces ROS to protect FAD and Fe–S clusters. 1025 However, the reduction potential of heme b in SDH of porcine is −185 mV, 1048 much lower than that of E. coli. Therefore, the electron sink mechanism is less effective in this case and needs further investigation. Fumarate reductase is a member of the succinate–ubiquinone oxidoreductase superfamily as well. It catalyzes the reduction of fumarate to succinate, the reverse reaction of SDH. It is very similar to SDH in subunit composition and cofactors. 1049,1050 Its three iron–sulfur clusters are linked to the protein by cysteine residues in E. coli, which are conserved in other fumarate reductases too. The midpoint reduction potential is between −70 and −20 mV, and that of the initial electron donor menaquinol is −74 mV. 1046 3.5 Engineered Fe–S Proteins 3.5.1 Artificial Rubredoxins A rubredoxin-like [FeCys4] center has been constructed into thioredoxin by computational design. The first coordination sphere is composed of two cysteines, Cys32 and Cys35, which form a disulfide bond in wild-type thioredoxin, as well as two cysteines introduced by mutation, Trp28Cys and Ile75Cys. The resulting monoiron center resembles Rd in UV–vis and EPR spectra, and the mimic protein is able to undergo three cycles of air oxidation and β-mercaptoethanol reduction. 1051 The redox process of rubredoxin is not fully reversible due to the instability of the reduced form. Nanda et al. have constructed a minimal rubredoxin mimic, RM1, on the basis of computational design for a more restrained tertiary structure derived from PfRd. RM1 is a domain-swapped dimer fused with a highly stable hairpin motif tryptophan zipper and displays spectroscopic properties very similar to those of native Rd’s. Moreover, it shows a reduction potential of 55 mV vs SHE and maintains redox activity for up to 16 cycles under aerobic conditions. 1051 3.5.2 Artificial [4Fe–4S] Clusters There have been numerous studies focusing on making model compounds of ferredoxins 1052−1054 and using those models to elucidate features of natural Fe–S clusters using several methods. 803,1055,1056,1057 In addition to synthetic models of ferredoxins that are discussed in a review in this journal, 2007 protein and peptide models of ferredoxins have also been made. These models have been discussed in detail in another review in this thematic issue, 3000 and we will discuss them here only briefly. Almost all of these mimics are modeled after [4Fe–4S] clusters, usually made by placing the conserved motif within a scaffold. These model systems have been used for unraveling the minimal structures required for binding of Fe–S clusters. 732,1058,1059,1061 A 16 amino acid peptide has been modeled to incorporate a low-potential [4Fe–4S] cluster. More detailed sequence alignments resulted in design of peptides with better cluster binding features that mimic FA and FB of photosystem I. 705 Other peptide models have also been made to analyze reduction potential properties of different Fe–S clusters, including [4Fe–4S] clusters, [2Fe–2S] clusters, and rubredoxins. 717 Four-helix bundle models of [4Fe–4S] clusters are among the most common systems to build and study these clusters. Both a single [4Fe–4S] cluster and a [4Fe–4S] cluster together with a heme cofactor have been designed in such four-helix bundles. 1061,1062 Recently, a “metal first” approach has been taken to introduce a [4Fe–4S] cluster into a non-natural α-helical coiled coil structure. The design then went through further optimization and addition of secondary sphere interactions to stabilize the reduced form and prevent aggregation. Such designs that are independent of structural motifs can be used as a platform for the future design of multiclusters to be used as biological “wires” that transfer electrons through a chain of proteins. 1063 3.6 Cluster Interconversion Although the Fe–S clusters are mostly classified on the basis of the number of iron atoms in the center, there are several cases in which changing one cluster to another type has been observed. These cluster interconversions can happen through three types of processes: natural changes in the environment of the cluster, chemical treatments of the cluster, or amino acid replacements. One of the most common types of cluster interconversion is the change from a [4Fe–4S] cluster to a [2Fe–2S] cluster. This kind of conversion has been observed in hydrogenases and nitrogenases. While CD and MCD analyses show that MgATP/ADP binding to the [4Fe–4S] cluster of Fe hydrogenase does not result in conversion to a [2Fe–2S] cluster, 1064 addition of α,α′-dipyridyl to the [4Fe–4S] cluster of nitrogenase resulted in formation of a [2Fe–2S] cluster in the presence of MgATP. 1065,1066 The [4Fe–4S] to [2Fe–2S] cluster conversion has been observed in enzymes such as ribonucleotide reductase 1067 and pyruvate formate activating enzyme 1068 as well, usually upon oxidation in air or chemical treatment. A very well studied case of the role of [4Fe–4S] to [2Fe–2S] cluster conversion in regulating cellular responses is that of fumarate nitrate reduction transcription factor. It has been shown that this protein undergoes the conversion upon O2 stress. The excess oxygen will oxidize S ligands and generate disulfide cysteines. The formation of a disulfide Cys-ligated [2Fe–2S] cluster will result in a monomerization of the fumarate nitrite reduction transcription factor dimer, hence unbinding from DNA. 1069,1070 The conversion is composed of two steps: first, the [4Fe–4S] cluster undergoes a one-electron oxidation to form a [3Fe–4S]1+ intermediate after releasing an Fe2+. Second, the [3Fe–4S]1+ cluster converts to a [2Fe–2S] cluster and releases an Fe3+ and two sulfide ions. 1071,1072 Mutating Ser24 into Phe and shielding Cys23 could inhibit step 1. 1073 Chelators of both Fe2+ and Fe3+ could accelerate step 2 significantly. 1074 Another very common interconversion is [4Fe–4S] to [3Fe–4S] interconversion. The [4Fe–4S] clusters are very sensitive to air, and oxidation in air can remove one of the irons, resulting in a 3Fe cluster. 1075 The most well studied case of this interconversion is the enzyme aconitase. Aconitase has a [4Fe–4S] cluster in its active form, which is very sensitive to air. Aerobic purification of the protein causes formation of an inactive enzyme with a 3Fe cluster. Addition of extra Fe, however, can reverse the conversion and reactivate the enzyme. 1076 Exposure of the [3Fe–4S] aconitase to high pH (>9.0) will result in the formation of a purple species that has been attributed to a linear [3Fe–4S] cluster. This purple protein can be activated again through reduction in the presence of Fe. 1077 While more often clusters of higher iron number convert into clusters with fewer iron atoms, the reverse case has also been observed. In biotin synthase, there are two [2Fe–2S] clusters that can convert to a [4Fe–4S] cluster after reduction. UV–vis and EPR studies reveal that the conversion process occurs through dissociation of Fe from the protein followed by slow reassociation. 1078 Ferredoxin II of Dv. gigas has a [3Fe–3S] cluster that can convert into a [4Fe–4S] cluster through incubation with excess Fe, presumably through a non-Cys ligand. 1079 The [3Fe–4S]1+ and [2Fe–2S]2+ clusters in isolated pyruvate formate–lyase can both be converted to [4Fe–4S] clusters with mixed valences of +1 and +2 upon dithionite reduction. 1080 Interconversion between [4Fe–4S] and [3Fe–4S] clusters has been investigated through mutational studies. Removal of Cys ligands in [4Fe–4S] clusters results in the formation of [3Fe–4S] clusters. Replacement of the conserved Asp in [3Fe–4S] clusters with a ligating residue such as His or Cys causes formation of [4Fe–4S] clusters. 735,944,1081,1082 In [NiFe] hydrogenase, mutating a conserved Pro residue into Cys near the [3Fe–4S] cluster has successfully converted it to a [4Fe–4S] cluster accompanied by a 300 mV decrease in the reduction potential, 944 while in F420 reducing hydrogenase of Methanococcus voltae the [4Fe–4S] to [3Fe–4S] conversion has been achieved by replacing a Cys residue, producing a ∼400 mV increase in the reduction potential. 1081 Addition of other metal ions in place of the fourth iron into a [3Fe–4S] cluster is sometimes also called interconversion. There are multiple reports of the formation of such hybrid clusters with Zn, Tl, and other metal ions. 1083,1084 3.7 Structural Features Controlling the Redox Chemistry of Fe–S Proteins The Fe–S proteins cover a wide range of reduction potentials, mostly in the lower or negative end of the range. Several parameters are known to be important in the ability of Fe–S proteins to accommodate such a wide range of reduction potentials. Unique electronic structures of iron in different clusters and different protein environments are among the most important factors. The ability of each iron to go through 2+ to 3+ oxidation states will allow multiple states for the core cluster, each of which having a different reduction potential range. This factor is more evident in the case of HiPIPs vs ferredoxins. Solvent accessibility, H-bonding patterns around the cluster, the net charge of the protein, partial charges around the cluster, and the identity of the ligands are among the other features that contribute to fine-tuning the reduction potential. Detailed examples of the role of each feature are discussed in section 3.4.3.3.3, “Important Structural Elements”. Below is a summary of these features and their effects in different Fe–S proteins. 3.7.1 Roles of the Geometry and Redox State of the Cluster As with other redox-active metal centers, the primary coordination sphere of a metal ion plays an important role in its redox properties. The iron center(s) has the same distorted tetrahedral structure in almost all Fe–S proteins; however, it has been shown that slight changes in this structure will result in changes in the reduction potentials. Differences in the Fe–S–Cα–Cβ torsion angle 618,731,1085 and distortion of the cuboidal structure in some [3Fe–4S] clusters 1086 are examples of this distortion. Different geometries can lead to slight differences in electronic structures that will affect the redox properties of the protein. Another important feature that influences the reduction potential is the number of redox centers in the cluster and the redox state of the cluster. While rubredoxin has only one iron that simply switches between Fe2+ and Fe3+ states, the same transition differs significantly in a [4Fe–4S] cluster in an environment with three more irons and a mixed-valence state (e.g., 2Fe3+–2Fe2.5+ and Fe2.5+). Even the same cluster can undergo different redox transitions, as has been observed in the case of HiPIPs and ferredoxins. 719 3.7.2 Role of Ligands While sulfurs are the most dominant ligands in Fe–S proteins, it has been shown that other ligands can replace sulfurs in some cases and that these ligands play a prominent role in fine-tuning the reduction potential of the proteins. 541 Generally speaking, ligands that are less electron-donating than sulfur will increase the reduction potentials by selectively destabilizing the oxidized state. A well-established example of this principle is the increased reduction potential of [2Fe–2S] clusters in Rieske proteins compared to ferredoxins due to replacement of two of the Cys ligands with His residues. Mutational studies on Cys ligands, mostly replacement with Ser, have shown an increased reduction potential compared to that of the wild-type (WT) proteins. 721,750,773,1087 3.7.3 Role of the Cellular Environment As mentioned earlier in this review, some Fe–S proteins such as vertebrate ferredoxins and certain [3Fe–4S] clusters and Rieske proteins show pH-dependent redox behavior. This behavior can be due to the presence of a protonable residue such as Asp or His residue as a ligand or near the active site. 712,746,801 Therefore, proteins in the presence of different pH values in different cellular compartments should demonstrate different reduction potentials. Another effect of the environment is indirect through evolution: as shown in the case of ferredoxins, organisms subjected to extreme environments will undergo changes in the overall charges of proteins, which will affect the reduction potentials. 823 Peptide models of different Fe–S clusters have demonstrated the impact of solvent composition in ET features of the cluster. 717 3.7.4 Role of the Protein Environment Several studies have shown the importance of the protein environment in fine-tuning the reduction potentials of metal centers. The protein environment is one of the, if not the, most important factors determining the reduction potential in Fe–S proteins because the general geometry and primary coordination of iron are very similar in this family of proteins. The protein environment conveys its effect via several routes. 3.7.4.1 Solvent Accessibility/Cluster Burial Solvent accessibility has been shown to be a very important factor in the reduction potential for different metal centers, including Cu centers, hemes, and Fe–S clusters. As a general rule of thumb, the more buried a cluster, the higher or more positive the reduction potential will be. This is mainly due to the electrostatic destabilization of more positive charges in the clusters. Being more buried is proposed to be one of the most important reasons behind the difference between the reduction potentials of the [4Fe–4S] clusters in HiPIPs vs ferredoxins. 618,749,752 Hydration of the cluster can influence the covalency of Fe–S bonds, hence affecting the reduction potential. 901 Cluster burial can be accomplished through physical positioning of the cluster by covering it with more secondary structure elements or partially via more hydrophobic residues around the cluster. As discussed earlier, there are exceptions to this trend, and there are clusters that are significantly more solvent-exposed, but little reduction potential change is observed for them. 875 It should be noted that cluster burial is dependent on the size of the protein, the location of the cluster, and the extent of solvent interaction, so it is difficult to make a fair comparison of the effect of cluster burial among different proteins. 92 3.7.4.2 Secondary Coordination Sphere While ligands in the primary coordination sphere are very important in tuning the reduction potentials of the Fe–S centers, the role of secondary coordination sphere interactions cannot be ignored. A mounting number of studies support the essential roles of these interactions in fine-tuning the reduction potentials. 1088 In the case of Fe–S proteins, secondary coordination interactions are the major cause of differences in the reduction potentials within a class of proteins. 887 The number of backbone to amide H-bonds has been shown to be important in redox potential differences between HiPIPs and ferredoxins. 617,618 As described in each section, a conserved H-bonding pattern is observed in each subclass of ferredoxins, and this pattern differs from one subclass to another. 718,719 Removal of some conserved H-bonds from this pattern is shown to be one of the main causes of different reduction potentials between different types of ferredoxins. 718,719 Removal of conserved H-bonds in several cases resulted in a decrease in the reduction potential. 773,780 It is important to mention that although H-bonds are important, they are not the sole cause of differences in the reduction potentials. Moreover, their analyses are complicated in some cases due to ambiguity in their assignment and variation in their number based on the environmental condition. 92 3.7.4.3 Electrostatics and Local Charges Local charges can selectively stabilize either the reduced or oxidized form of the cluster and influence the reduction potential. Many studies of the Fe–S proteins showed that although these proteins usually have conserved charged residues (such as positive charges in ferredoxins), these charges are mainly important for interaction with the redox partner, and usually their mutations do not cause significant changes in the reduction potential. 749 In cases where these residues are very close to the cluster, unpredictable effects have been observed. 611 However, the total charge of the cluster has been suggested to be an important factor influencing the higher reduction potential of Rieske proteins compared to ferredoxins. 773 Mutational analysis on rubredoxins and thioredoxin-like ferredoxins confirmed an important role for the charges around the cluster in the reduction potential of the protein. There is convincing evidence for the role of backbone amides and partial positive charges in the reduction potential of Fe–S centers. 887 It has been proposed that the diploes induced by the these backbone amides can influence the reduction potential of different clusters, such as HiPIPs and ferredoxins. The net protein charge and the dipole induced from backbone amides have been shown to be important in determining the reduction potential of HiPIPs. 752,873,890 While all these features are important, it should be noted that none of them are the sole determinants of the reduction potential in Fe–S proteins, and it has been found that different features act as the major contributors to differences in the reduction potential between different classes of the Fe–S proteins. Even among members of a class, the same factor might not play the same role. 3.7.5 Computational Analysis of the Reduction Potentials of Fe–S Proteins To further understand factors influencing the reduction potentials, computational methods have been developed for calculating the reduction potential of Fe–S proteins on the basis of their structures. 591,887 One of these methods uses Gunner’s multiconformational continuum electrostatics method and has been calibrated using proteins with known structure and reduction potential. 780 In another method a combined quantum-chemical and electrostatic calculation was used to generate predictions for reduction potentials. Poisson–Boltzmann electrostatic methods in combination with QM/MM studies have also been used to analyze the reduction potentials of Fe–S proteins. 93 The PDLP method was applied to HiPIPs to analyze the effects of solvent accessibility on the reduction potentials of these proteins. 92,719 B3LYP density functional methods have been used in combination with broken symmetry to analyze factors that are important in tuning the reduction potential of Rieske proteins. 800 Broken symmetry in combination with hybrid density functional theory has also been used to characterize Rieske proteins. 1089 4 Copper Redox Centers in Electron Transfer Processes 4.1 Introduction to Copper Redox Centers Copper is the second most abundant transition metal in biological systems, next to iron. 1090 In addition to their critical role in electron transfer process, copper-containing proteins catalyze a variety of reactions. In this section, we focus on copper proteins that merely function as ET mediators, which include blue or type 1 (T1) copper and CuA centers. A number of reviews on these two centers have appeared in the literature. 94−104 Despite the lack of modern structural and computational methods, initial attempts to understand the structure and function of copper redox centers were very successful. This success was in part due to the strong colors and interesting magnetic properties displayed by these redox centers that allowed various spectroscopic studies. The blue copper proteins were so-named because they display an intense blue color, due to a strong absorption around 600 nm, first observed in the 1960s. 1091,1092 It was found that this T1 copper protein also displayed an unusual EPR spectrum with narrow hyperfine splittings, suggesting the presence of Cu in a different ground state compared to the normal copper complexes. 1093 The electronic structure of the blue copper center was further elucidated with low-temperature absorption, CD, MCD, single-crystal EPR, XAS, and computational studies. 96,99,1094,1095 The results of all these studies demonstrated that the 600 nm band is associated with a S → Cu charge transfer transition and that the highly covalent nature of the Cu–S bond is responsible for the narrow hyperfine splitting in the EPR spectra. The crystal structure of poplar plastocyanin later confirmed that T1 copper proteins contain a copper site with an unusual geometry. 1096 Although the existence of copper in cytochrome c oxidases (CcOs) has been known since the 1930s, the nature of the CuA centers was not established until much later due to the presence of heme cofactors that complicated interpretation of the spectroscopic results. 1097 EPR and elemental analyses have revealed that two copper-binding sites exist in CcOs. 1098−1100 MCD studies by Thomson and co-workers showed features at 475, 525, and 830 nm corresponding to a CuA center. 1101,1102 Kinetic measurement of reoxidation of reduced CcO, performed by a flow-flash technique, indicated that the CuA is the ET center in CcO. 1103,1104 From 1987 to 1993, Buse and co-workers performed chemical analysis of CcO with inductively coupled plasma atomic emission spectroscopy, leading to the conclusion that three copper atoms exist in one protein along with two hemes. 1105,1106 Later, resonance Raman, 1107 EXAFS, 1108 and finally crystal structures 1030,1109 revealed an unusual dinuclear copper structure for the CuA center, which will be discussed in detail in section 4.5. 4.2 Classification of Copper Proteins As a diverse family of proteins, copper proteins could be divided into several types according to ligand sets, spectroscopic features, and functions (Table 9). 1110,1111 Mononuclear T1 copper centers and dinuclear CuA centers are the two types which act only as ET mediators. T1 copper centers and CuA centers share several common features. First, both centers contain Cu–thiolate bond(s), which are highly covalent and display rich spectroscopic signatures. 99,1095,1112−1115 Second, both centers are located in a cupredoxin fold. 94,100,103 Finally, they are highly optimized for ET, showing low reorganization energies and high ET rate constants. These two types of copper proteins are collectively called cupredoxins, analogous to ferredoxin for Fe–S-based ET centers. 1116 Other types of copper proteins may also involve ET as part of their enzymatic reactions, including peptidylglycine α-hydroxylating monooxygenase and dopamine β-monooxygenase, 1117 but will not be discussed here. Table 9 Different Types of Copper Proteinsa   mononuclear dinuclear tetranuclear   type 1 type 2 type 3 CuA CuZ UV–vis spectrum strong absorption, ∼600 nm and (in some proteins) 450 nm weak absorption, ∼700 nm 300–400 nm strong absorption, ∼480 and 530 nm strong absorption, ∼640 nm EPR spectrum four-line (A ||  1000 1148 (redox-inactive)   ceruloplasmin       1Cys, 2His, 1Met 448 1149 (redox-active)   hephaestin mammals 1999 1150         Fet3p yeast 1994 1151 1ZPU 1Cys, 2His 427 1152   nitrite reductase plants, bacteria   1NIA 1Cys, 2His, 1Met, 1 carbonyl oxygen 260 1153   Figure 46 Crystal structures of the T1 copper proteins. The secondary structure (α-helix and β-sheet) is shown in cartoon format, copper is shown as a purple ball, and ligands are shown in stick format. The name of the protein and its PDB ID are given below each structure. Figure 47 Topology diagram showing the scheme of the secondary structure of azurin. β-Strands are shown as arrows, and the α-helix is shown as a cylinder. Copper ligands between β-strands 3 and 4 and between β-strands 7 and 8 are shown as blue polygons, while copper is shown as a purple circle. Most of the ligands to the T1 copper center resides at the C-terminal end of the cupredoxin fold. As shown in Figure 47, one of the His ligands is the first residue of the fourth β-strand and is referred to as N-terminal His. Carbonyl oxygen, the fifth ligand of azurin, is located in the loop between the third and fourth β-strands. Other ligands, including Cys, the second His on the trigonal plane, and the axial ligand, are located in or adjacent to the loop between the seventh and eighth β-strands, close to the C-terminus of the protein. Cys is the last residue of the seventh β-strand, while the second His is in the middle of the loop and is referred as the C-terminal His. Met is the first residue of the eighth β-strand. The three ligands are arranged in Cys-(Xxx) n -His-(Xxx) m -Met fashion, where n and m could vary between 2 and 4 in different T1 copper proteins. This variation in length and amino acid composition is important for the function of T1 copper proteins. In section 4.4.5 we discuss the implications of the variations based on loop-directed mutagenesis results. While X-ray crystallography could give a fairly good description of the overall structure, EXAFS is more accurate in determining the metal–ligand distance because it is sensitive to oxidation state of the metal ion. 1158 The short Cu–S distance was first revealed by EXAFS. 99,1159 By comparing data from oxidized and reduced plastocyanin and azurin, it was found that an average increase of ∼0.06 and ∼0.08 Å for Cu–N(His) and Cu–S(Cys), respectively, happens upon reduction. 99 These small changes upon reduction are consistent with data from crystallography and suggest a small reorganization energy for the redox process. 4.3.1.1 Copper Ligands Even though the amino acid sequences and overall structures vary among different T1 copper proteins, the ligand composition, ligand–metal distance, and geometry of the T1 copper centers are almost identical (Figure 48). 94,95,99 As the most conserved structural feature, T1 copper centers invariably contain two His residues and one Cys residue as equatorial copper ligands. In T1 copper proteins, the His coordinates with copper through Nδ, in contrast to Nε used by T2 and most other copper proteins. The Cu–His bond length is about 2.0 Å in T1 copper proteins, which is normal for such types of bonds. On the other hand, the Cu–Cys bond lengths range from 2.07 to 2.26 Å, which is short compared to those of normal copper complexes and other copper proteins (Table 11). The short Cu–S distance is key to the unique spectroscopic properties of T1 copper and is maintained through extensive H-bonding within the protein scaffold, as will be discussed later in this section. The 2N and 1S from His and Cys, respectively, form a pseudotrigonal plane, with average bond angles in the Cu(II) state being 101°, 117°, and 134° with RMS deviations of 2.5°, 4.1°, and 2.8°, calculated from crystal structures with resolution of 2.0 Å or higher. 1119 The Cu–Sγ–Cβ–Cα and Sγ–Cβ–Cα–N dihedral angles are also consistently close to 180°, making the Cu–Sγ bond coplanar with the Cys side chain and backbone. Figure 48 T1 copper centers in plastocyanin, azurin, plantacyanin, and amicyanin. Reprinted with permission from ref (1119). Copyright 2006 Wiley-VCH. Table 11 Distances (Å) between Cu or Other Substituted Metals and Ligands in T1 Copper Proteinsa P. aeruginosa azurin pH Cu–Nδ (His46)b Cu–S (Cys112)b Cu–Nδ (His117)b Cu–S (Met121)b Cu–O (Gly45)b resolution (Å) PDB ID ref Cu(II) 5.5 2.08(6) 2.24(5) 2.01(7) 3.15(7) 2.97(10) 1.9 4AZU (1160) Cu(I) 5.5 2.14(9) 2.29(2) 2.10(9) 3.25(7) 3.02(8) 2.0 1E5Y   Cu(II) 9.0 2.06(6) 2.26(4) 2.03(4) 3.12(7) 2.94(11) 1.9 5AZU (1160) Cu(I) 9.0 2.20(11) 2.30(23) 2.21(12) 3.16(9) 3.11(11) 2.0 1E5Z   T. ferrooxidans rusticyanin pH Cu–Nδ (His85) Cu–S (Cys138) Cu–Nδ (His143) Cu–S (Met148) – resolution (Å) PDB ID ref Cu(II) 4.6 2.04 2.26 1.89 2.88 – 1.9 1RCY (1161) Cu(I) 4.6 2.22 2.25 1.96 2.75 – 2.0 1A3Z   P. nigra plastocyanin pH Cu–Nδ (His37) Cu–S (Cys84) Cu–Nδ (His87) Cu–S (Met92) – resolution (Å) PDB ID ref Cu(II) 6.0 1.91 2.07 2.06 2.82 – 1.33 1PLC (1162) Cu(I) 7.0 2.13 2.17 2.39 2.87 – 1.80 5PCY (1163) P. denitrificans amicyanin pH Cu–Nδ (His53) Cu–S (Cys92) Cu–Nδ (His95) Cu–S (Met98) – resolution (Å) PDB ID ref Cu(II) 6.0 1.95 2.11 2.03 2.90 – 1.31 1AAC (1164) Cu(I) 7.7 1.95 2.12 unbound 2.91 – 1.30 2RAC (1165) C. sativus cucumber basic protein pH Cu–Nδ (His39) Cu–S (Cys79) Cu–Nδ (His84) Cu–S (Met89) – resolution (Å) PDB ID ref Cu(II) 6.0 1.93 2.16 1.95 2.61 – 1.80 2CBP (1166) C. sativus stellacyanin pH Cu–Nδ (His46) Cu–S (Cys89) Cu–Nδ (His94) – Cu–O (Gln89) resolution (Å) PDB ID ref Cu(II) 7.0 1.96 2.18 2.04 – 2.21 1.60 1JER (1167) a Adapted with permission from ref (104). Copyright 2012 Elsevier. b Average of distances for four molecules in the asymmetric unit. Errors are 1 standard deviation. The axial ligand in the T1 copper center is less conserved. A Met is present at 2.6–3.2 Å in this axial position in most proteins, while a Gln is found in stellacyanin and dicyanin. In the T1 center of fungal laccase and ceruloplasmin, a noncoordinating ligand such as Phe or Leu takes this axial position. In azurin, there is an additional backbone carbonyl oxygen at the opposite end of the axial position to Met, giving the T1 copper site a trigonal bipyrimidal geometry. 4.3.1.2 Secondary Coordination Sphere While the above mentioned ligands exert significant influence on the properties of T1 copper centers, the protein scaffold should not be viewed as a passive entity to hold the copper site. On the contrary, it can play important roles. First, it can shield the copper site from water, raising the reduction potential and lowering the reorganization energy for ET. More importantly, the extensive H-bond network surrounding it can fine-tune the properties of the T1 copper site. 94,98 As shown in Figure 49, the Cys112 in azurin forms two hydrogen bonds with adjacent backbone amide groups of Asn47 and Phe114 at ∼3.5 Å. Together with S–Cu and S–Cβ covalent bonds, these H-bonds form a tetrahedral geometry around Sγ of Cys (Figure 49A). Plastocyanin, pseudoazurin, and amicyanin have only one H-bond around the Cys as a Pro in the site eliminates the other amide bond. Additionally, cucumber basic protein has a very weak H-bond at 3.7–3.8 Å. These H-bonds modulate the electron density of S on Cys, which is crucial for the highly covalent nature of the Cu–S bond. Figure 49 H-bonding around Cys112 (A) and other ligands (B) of azurin. PDB ID 4AZU. In azurin, the N-terminal His coordinates with Cu through Nδ, whereas Nε is hydrogen-bonded to the carbonyl oxygen of Phe15. The same His is hydrogen-bonded to the Gln49 side chain in amicyanin, the side chain of Asn80 in rusticyanin, and a water molecule in phytocyanins. The C-terminal His is in a hydrophobic patch of the protein packed against other residues. The Nε of C-terminal His is hydrogen-bonded to a water molecule. The axial Met/Gln usually packs against aromatic side chains such as Phe15 in azurin (Figure 49). In azurin, the carbonyl oxygen is held in place by the secondary structure of the loop and packs with Phe114. There are more H-bonding interactions beyond the copper center. For example, an Asn close to the N-terminal His in the first ligand loop is hydrogen-bonded to residues from the other ligand loop. This interaction, acting like a zipper, further holds the copper site together. Extensive H-bonding around the copper site in T1 copper proteins has important functional implications, as we will address in section 4.4.2. 4.3.1.3 Comparison of Structures in Different States As suggested by the “rack mechanism”  1168,1169 or entatic state, 1170 the active site structure is predetermined by the protein scaffold. Thus, there is little change in the structures of T1 copper proteins at different oxidation states, with different metals, or even in the absence of metal ions. As shown in Table 11, compared to the same protein with Cu(II), the metal to ligand bonds elongated by 0.1 Å or less in protein containing Cu(I). Similar results were obtained by EXAFS, which provides a more accurate determination of the bond length. 99 The small change in bond length is crucial for the low reorganization energy of the T1 copper site and, thus, fast ET for its function. However, bond lengths in X-ray crystal structures should be interpreted with caution, as it has been shown that Cu(II) ions in protein undergo photoreduction during X-ray exposure. 1171,1172 It will be useful to conduct single-crystal microspectrophotometry concurrent with X-ray diffraction to make sure that the oxidized protein is not reduced during diffraction. 1173 On the other hand, the oxidation state of the Cu ion can be easily monitored at the edge and XANES regions of its X-ray absorption spectrum. Bond lengths derived from carefully designed and conducted EXAFS should reflect the actual bond lengths at the corresponding oxidation states. Besides structures with copper in oxidized or reduced states, crystal structures of apo and metal-substituted T1 copper proteins also shed light on how proteins interact with copper. Structures of apo forms of azurin, 1174,1175 plastocyanin, 1176 pseudoazurin, 1177 and amicyanin 1178 show little difference (0.1–0.3 Å) from that of the copper-bound form, confirming the entatic state hypothesis. Metal substitution is useful in spectroscopic studies, such as electronic absorption 1118,1179 and NMR. 1180 Due to the different sizes and ligand affinities of different metals, the bond length and overall geometry are changed upon substitution, but only to a small extent due to confinement of the protein scaffold. 1181−1183 4.3.2 Spectroscopy and Electronic Structure Intense (∼5000 M–1 cm–1) electronic absorption at ∼600 nm is the hallmark of T1 copper proteins (Figure 50). Solomon and co-workers attributed the origin of the ∼600 nm absorption to the S(Cys)pπ → Cu x 2–y 2 LMCT transition. 1094,1184,1185 Another feature at ∼450 nm is not prominent in plastocyanin or azurin, but is more pronounced in a perturbed T1 copper sites such as that of cucumber basic protein. This absorption is attributed to S(Cys)pπ → Cu x 2–y 2 LMCT. The geometry of the copper site is believed to be important for the ratio between the two peaks at ∼600 and ∼450 nm. 1095,1186 A series of weak absorption peaks from 650 to 1050 nm are attributed to a d → d transition or ligand field transition. 1184 Figure 50 Electronic absorption (A) and EPR (B) spectra of azurin. EPR provides a sensitive way to determine the copper site geometry. T1 copper proteins exhibit a distinctive small hyperfine splitting ( 150 × 10–4 cm–1). 1119 Through S K-edge XAS, Solomon and co-workers showed that the small hyperfine splitting is due to high covalency between Cu and S, which delocalizes unpaired electrons onto S, thus decreasing the electron density on Cu. 1187 Other spectroscopic techniques, such as resonance Raman spectroscopy and Cu L-edge and S K-edge XAS, have also been important in deciphering the electronic structures of T1 copper proteins. They are beyond the scope of this review, but there are excellent reviews elsewhere 1095,1119 and in this issue that cover more details about these techniques. 2000 4.3.3 Redox Chemistry of Type 1 Copper Protein As a class of proteins dedicated to ET, T1 copper proteins display various features for facile redox chemistry. 4.3.3.1 Redox Partner T1 copper proteins shuttle electrons between donor and acceptor proteins as redox partners. So far five T1 copper proteins with known physiological redox partners have been identified: plastocyanin, amicyanin, rusticyanin, pseudoazurin, and azurin. As an electron carrier in chloroplasts in plants, plastocyanin accepts electrons from cytochrome f of membrane-bound cytochrome b 6 f complex and transfers them to P700+ in photosystem I. 256,1188−1192 Amicyanin accepts electrons from methylamine dehydrogenase and transfers them to cytochrome c oxidase via a c-type cytochrome. 279,1193−1200 Rusticyanin is suggested to shuttle electrons between cytochrome c and cytochrome c 4. 1201,1202 Pseudoazurin reduces nitrite reductase, but its electron donor is not yet known. 1203−1207 Azurin is likely to interact with aromatic amine dehydrogenase in vivo, as suggested by coexpression, the kinetics of reduction, and the crystal structure. 1208−1210 Interaction between a T1 copper protein and its redox partner is generally weak and transient. NMR and crystallographic studies have revealed a structural basis for this interaction. Interactions between plastocyanin from various organisms and cyt f have been extensively studied by NMR spectroscopy (Figure 51). Chemical shift analysis and rigid-body structure calculations have demonstrated that the hydrophobic patch around His87, the C-terminal His ligand to copper, mediates the interaction between plastocyanin and cyt f. 1211,1212 Besides that, two acidic patches around Tyr83 have been shown to interact with positively charged residues of cyt f. 1213 Mutation of Tyr83 to Phe or Leu drastically decreases the ET rate between the two proteins, indicating that Tyr83 is involved in binding to cyt f and ET. 1214 The absence of acidic patches also demolishes ET activity at low ionic strength, showing they are involved in the interaction with cyt f. 1215,1216 However, interaction between acidic patches and cyt f is not very specific as small changes in acidic patches have a minimal effect on the interaction between two proteins. 1216,1217 Figure 51 Structures of plastocyanin (left) and the complex of plastocyanin and cyt f (right). Left: copper ion is represented as a purple ball, His87 and Tyr 83 are represented in licorice format, and residues in two acidic patches are represented as ball and stick models. Right: plastocyanin is colored cyan, and cyt f is orange. The copper ion and His87 from plastocyanin and heme from cyt f are also shown. Another demonstration of the interaction between the T1 copper proteins and their redox partners comes from X-ray crystallography. The structures of the amicyanin–methylamine dehydrogenase complex and methylamine dehydrogenase–amicyanin–cytochrome c 551 ternary complex have been determined. 279,1196 These structures further confirmed that the hydrophobic patch surrounding His95 (the C-terminal His ligand equivalent to His87 in plastocyanin and His117 in azurin) interacts with a hydrophobic patch on methylamine dehydrogenase. An ET pathway from Trp57 and Trp108 in methylamine dehydrogenase to His95 in amicyanin and eventually to copper has been proposed from these structures. Recently, the crystal structure of the azurin and aromatic amine dehydrogenase complex from Alcaligenes faecalis has been solved. 1208 In this structure, only one azurin molecule is present in complex with four molecules of aromatic amine dehydrogenase. The B factor of the azurin structure is high except for those residues in the interface. This result is consistent with the transient nature of the interaction between the T1 copper proteins and their redox partners. The interaction is very similar to the one between amicyanin and methylamine dehydrogenase. The T1 copper proteins show promiscuity in reacting with proteins other than their physiological redox partners, 64,1218 including small inorganic complexes such as [Fe(CN)6]3– and [Co(phen)3]3+, 31,44,1219 small molecules such as flavins and ascorbate, and the proteins themselves through electron self-exchange reactions. 100 Gray and co-workers have used Ru derivatives of T1 copper proteins as a model to study long-range ET in biological systems. 24,31,44,2005 4.3.3.2 Electron Transfer Rate T1 copper proteins are involved in long-range ET in vivo and in vitro. For a more detailed review of long-range ET, please refer to the review in this issue by Gray et al. 2005 The process can be described by the semiclassical Marcus equation: Marcus equation 1 In this equation, ΔE° is the difference in reduction potential between the donor and acceptor sites (also known as the driving force), H AB is the donor–acceptor electron coupling or electron matrix coupling element, and λ is the reorganization energy required for ET. Under the same driving force, the rate is maximized when H AB is large and λ is small. In long-range ET, there is little direct coupling between the donor and the acceptor. The coupling is mediated by intervening atoms via the superexchange mechanism. H AB is determined by the distance between the donor and acceptor and the covalency of the metal–ligand bond. 1220−1222 Electron transfer rates between T1 copper proteins and their redox partners have been measured by kinetic UV–vis spectroscopy or cyclic voltammetry. 1223−1226 The k ET between plastocyanin and cyt f has been determined to be 2.8–62 s–1, 1227−1229 while the constant between plastocyanin and P700+ has been determined to be 38–58 s–1. 1191,1192,1230,1231 Davidson and co-workers have used kinetic UV–vis spectroscopy to measure the k ET between amicyanin and methylamine dehydrogenase, which was determined to be ∼10 s–1. 1232,1233 Suzuki and co-workers have determined the k ET between pseudoazurin and nitrite reductase to be (0.8–7) × 105 M–1 s–1 by kinetic UV–vis spectroscopy or cyclic voltammetry. 1204,1224,1234−1236 As several studies have pointed out, the rate constant measurement for interprotein ET processes is complicated by other processes, such as multiple binding sites of the two proteins, transient formation of conformational intermediates, and protonation/deprotonation processes. 1225,1237 There are two methods to measure the ET rate in T1 copper proteins without involvement of a redox partner: pulse radiolysis and NMR. Pulse radiolysis 1238 uses a short pulse (typically 0.1–1 μs) of high-energy (2–10 MeV) electrons to excite and decompose solvent molecules. A typical reaction generates the CO2 •– radical: Radicals generated in solvent molecules trigger downstream reactions. In azurin, CO2 – can reduce either Cu(II) or the disulfide bond between Cys3 and Cys26 at a nearly diffusion-controlled rate. Molecules with a reduced disulfide bond (RSSR–) can further reduce Cu(II) in the same protein via intramolecular ET: 101 By monitoring absorbance changes at 410 nm (RSS•R–) and 625 nm (Cu(II)), a fast reduction process corresponding to reduction of Cu(II) or RSS•R– by CO2 •– and a slower process of intramolecular ET between RSSR and Cu(II) can be resolved. The ET rate and driving force (ΔG°) can be calculated from the kinetics of intramolecular ET. By running experiments at different temperatures, the activation enthalpy and activation entropy of the ET process can be calculated. Using this method, Farver and Pecht determined the rate constant of intramolecular ET of WT azurin to be 44 ± 7 s–1 at pH 7.0 and 25 °C with a driving force ΔG° = −68.9 kJ mol–1. The activation enthalpy and activation entropy were calculated to be 47.5 ± 4.0 kJ mol–1 and −56.5 ± 7.0 J K–1 mol–1. 1239 ET rates for azurin of different origins and mutations have been measured and reviewed by Farver and Pecht. 101 Electron self-exchange is an intrinsic property of all redox systems. 1240 Exchange of electrons happens to two molecules of the same complex at different oxidation states. Only one redox couple is involved, and there is no driving force for this reaction. Measuring electron self-exchange rate constants by NMR provides a more universal way to measure ET transfer activity as it is carried out in T1 copper centers 1241−1249 (reviewed in ref (100)) as well as in other redox centers. 1250−1252 Electron self-exchange rate constants (k SES) of T1 copper proteins range from 103 to 106 M–1 s–1 at moderate to low ionic strength. The electron self-exchange is thought to happen through a hydrophobic patch as the rate constant is affected by the presence of an acidic patch 1248 or basic residues 1253 close to the hydrophobic patch. 4.3.3.3 Reduction Potential T1 copper proteins have reduction potentials ranging from 183 to 800 mV (see Table 10). Compared to the aqueous Cu(I)/Cu(II) couple (which has a reduction potential of ∼150 mV), copper complexes, and other copper proteins, T1 copper proteins have unusually high reduction potentials. Their potentials also span a wide range (>600 mV), nearly half the range of biologically relevant potentials (Figure 1). Within the T1 copper proteins, groups of proteins are apparent when sorted on the basis of the midpoint reduction potential (E m). Nitrite reductases, 1153 stellacyanins, 1133 amicyanins, 1124 and pseudoazurins 1128 natively have substantially lower (∼100 mV) E m values as compared to azurin. 98 Azurin and umecyanins have moderate E m values natively around 200–300 mV vs SHE. On the other end of the scale, rusticyanins have E m values ∼400 mV higher than that of azurin. Understanding the origin of this variance and the structural features involved in tuning the reduction potential are of great importance. By comparing the native proteins with different axial ligands (Table 12), it is revealed that proteins with Gln as an axial ligand generally have lower reduction potentials (190–320 mV), proteins with Met axial ligands have higher potentials (183–670 mV), and proteins with a noncoordinating ligand in multicopper proteins have the highest potentials (354–800 mV). This trend is further confirmed by mutagenesis studies that are discussed in section 4.4.1. Table 12 Dependence of E° on the Axial Ligand in Blue Cu Proteinsa   E° (mV)     Phe/Leu/Thr Met Gln ref fungal laccase 770 680   (1254−1256) azurin 412 310 285 (1122, 1257) cuc. stellacyanin 500 420 260 (1139) nitrite reductase 354 247   (1258) rusticyanin 800 667 563 (1259) mavicyanin   400 213 (1260) amicyanin   250 165 (1261) a Reprinted from ref (99). Copyright 2004 American Chemical Society. Variation within proteins containing the same axial ligand indicates that there are more factors affecting the reduction potentials of the T1 copper center. These factors have been uncovered by mutagenesis studies and engineering of copper proteins and are discussed in section 4.4. 4.3.4 T1 Copper Center in Multicopper Proteins The T1 copper center exists not only in single-domain proteins, but also in multidomain proteins with multiple copper cofactors. These proteins include multicopper oxidases and nitrite reductases (Table 9). The former contain a T1 copper (blue copper), a type II copper (abbreviated as T2), and a pair of type III copper centers (Figure 53). 1262−1266 The latter contain T1 and T2 copper centers and are evolutionarily related to the multicopper oxidases. 1265−1267 As shown in Figure 52, multicopper oxidases and nitrite reductases are closely related and are composed of two, three, or six domains. 1265 In multicopper oxidases, the T1 copper center resides in the cupredoxin-like domain while the T2 and T3 copper centers are located between domains. Figure 52 Domain organization and copper center distribution in multicopper oxidases. Reprinted with permission from ref (1265). Copyright 2011 Wiley-VCH. Figure 53 Active site of the multicopper oxidases. Cu sites are shown as green spheres. Figure generated from the crystal structure of ascorbate oxidase (PDB ID 1AOZ). Reprinted from ref (1264). Copyright 2007 American Chemical Society. T1 copper centers in multicopper oxidases (MCOs) are very similar to those in single-domain T1 copper proteins. The copper ion is coordinated by one Cys residue and two His residues at its equatorial positions. In plant laccases, ascorbate oxidases, and nitrite reductases, axial Met coordinates with copper and forms a trigonal pyramidal geometry. In fungal laccase, ceruloplasmin, and Fet3p, the axial ligand is a noncoordinating Leu or Phe, leaving equatorial ligands and copper in a more trigonal geometry. 1262,1265,1266 One noticeable feature for T1 copper centers in MCOs is their high reduction potential compared with that of single-domain T1 copper proteins. Ceruloplasmin has the highest reduction potential 1148 (>1000 mV) reported in T1 centers, while TvLac has the second highest reduction potential 1141−1143 (778 mV). The high reduction potential is partially attributed to the more hydrophobic axial ligand, while other factors such as hydrogen bonding around the T1 Cu centers may contribute too. 1268 4.3.5 A Novel Red Copper Protein—Nitrosocyanin Recently, a mononuclear red copper protein, nitrosocyanin from N. europaea, an ammonia oxidizing bacterium, was isolated and structurally characterized (Figure 54). 1137,1269−1271 The crystal structure shows that the copper ion is coordinated by two His residues, one S(Cys), and a side chain O(Glu) and has an additional fifth water ligand in the oxidized form, but not in the reduced form. Nitrosocyanin shows a strong absorption band at 390 nm (ε = 7000 M–1 cm–1), a large hyperfine splitting value (147 × 10–4 cm–1) in the EPR spectrum, and a very low reduction potential of 85 mV (compared with those of the T1 copper proteins, which are in the range of 150–800 mV). 1137,1271 With an exogenous water ligand, the reorganization energy of this protein is calculated to be 2.2 eV, significantly higher than those of T1 copper proteins. 1271 Similar to T1 copper proteins, nitrosocyanin has copper–thiolate coordination and strong UV–vis absorbance. However, the water ligand in nitrosocyanin has not been observed in T1 copper proteins before. Its copper site geometry and absorption at ∼400 nm are also different from those of T1 copper proteins. Its EPR spectrum, reorganization energy, and reduction potential more closely resemble those of T2 copper proteins. Solomon and co-workers attribute these properties to the relative orientation of the Cu-N-N-S and Cu-S-Cβ planes, which in turn is due to “coupled distortion” between the axial ligand and the whole copper center. 1095,1186,1271 Figure 54 Crystal structures of (A) the oxidized red copper site in nitrosocyanin, (B) the oxidized T1 copper site in plastocyanin, and (C) the reduced red copper site in nitrosocyanin. Reprinted from ref (1271). Copyright 2005 American Chemical Society. The biological role of this protein, however, has not yet been identified. It has been proposed that it might be involved in ET or serve some as-yet-unknown catalytic function due to the presence of the open coordination site. 1269,1270 4.4 Structural Features Controlling the Redox Chemistry of Type 1 Copper Proteins Although the study of native proteins provides valuable information about the structure, spectroscopy, and function of T1 copper centers, it is difficult to draw any conclusion only by comparing copper centers from different scaffolds with low sequence homology. With the advancement of modern molecular biology, powerful tools such as mutagenesis are available to research groups, allowing the amino acid sequence to be modified at will. Methods of unnatural amino acid mutagenesis have further expanded the toolbox for bioinorganic chemists. 1272−1274 With these methods, not only amino acid residues directly coordinating to copper, but also residues beyond the first coordination sphere have been changed. Mutagenesis reveals how different components of the protein contribute to the structure, spectroscopy, and function, especially in reduction potential tuning. 4.4.1 Role of Axial Met The T1 copper center has highly conserved equatorial ligands, two His residues and one Cys residue. The axial position of the T1 copper center shows more variation, as Met, Gln, and noncoordinating residues can all be found in the native proteins. Mutagenesis of the axial ligand has been carried out in azurin, 1122,1275−1278 nitrite reductase, 1235,1258,1279 amicyanin, 1261 rusticyanin, 1259 pseudoazurin, 1234 laccase, 1256 and stellacyanin. 1139,1280,1281 Mutation of the axial ligand in different T1 copper proteins generally results in a protein that retains copper-binding ability but with a different reduction potential or altered spectroscopic properties. An early work replaced Met121 in azurin with all other 19 amino acids with minimal alteration of the T1 character of the copper center. 1276 While changing the axial ligand to hydrophobic ligands such as Ala, Val, Leu, or Ile increases the reduction potential by 40–160 mV, 1122 substitution with Glu or Gln decreases the reduction potential by 100–260 mV. 1122,1257 As the axial ligand is changed from Gln to Met to more hydrophobic residues, the reduction potential of the protein increases. Theoretical studies have suggested that the axial ligand is involved in tuning the potential. 1282,1283 To test the role of the axial ligand in tuning the reduction potential of the T1 copper protein, Lu and co-workers incorporated unnatural amino acid analogs of Met with different hydrophobicities at the axial position in azurin. 1284,1285 The reduction potential varied from 222 to 449 mV at pH 4.0. Such a replacement of Met with its iso-structural analogs allowed conclusive identification of hydrophobicity of the axial ligand as the major factor in tuning reduction potentials, because a linear correlation was found between the reduction potential and hydrophobicity of the axial ligand. Likewise, Dennison and co-worker mutated the axial Met of cucumber basic protein to Gln and Val. As the axial ligand was changed from Gln to Met to Val, the electron self-exchange rate increased by 1 order of magnitude, and the reduction potential increased by ∼350 mV. 1286 These studies have firmly established a correlation between hydrophobicity of the axial ligand and reduction potential, providing a better understanding of the role of the axial ligand in reduction potential tuning. Within T1 copper proteins, there are two classes of proteins with slightly different spectroscopic features. Typical T1 copper proteins, such as plastocyanin and azurin, have absorption at ∼600 nm and an axial EPR signal, whereas “perturbed” T1 copper proteins or green copper proteins have an additional ∼400 nm absorption peak in their UV–vis spectra, as well as rhombic EPR signals. At the same time, the perturbed T1 copper proteins have longer Cu–S(Cys) distances and shorter Cu–axial ligand distances. 1283 A more extreme case comes from the newly discovered protein nitrosocyanin, which has a cysteine ligand and dominant ∼400 nm absorption in its UV–vis spectrum, resulting in a red color. 1137,1271 Although the strong absorption and 1Cys/2His/1Glu ligand set resembles those of T1 copper proteins, nitrosocyanin has large hyperfine splittings (A || ≈ 150 × 10–4cm–1) in its EPR spectrum and a low reduction potential (85 mV), which falls into the range of T2 copper proteins. 1136,1137,1271 Solomon and co-workers proposed coupled distortion theory on the basis of a suite of spectroscopic studies in combination with theoretical calculations to explain the variance in electronic absorption and concomitant color change from blue to green to red in native proteins. This theory states that shorter Cu–axial ligand distances result in distortion of the T1 copper geometry toward tetragonal, which elongates the Cu–S(Cys) distance. 1283 This distortion renders the pσ(Cys)–Cu CT more favorable than pπ(Cys)–Cu CT, which causes an increase in the ∼400 nm absorption over the ∼600 nm absorption in the UV–vis spectrum. Mutational studies on the axial ligand of various T1 copper proteins have validated the coupled distortion theory. By changing a weak Met to a stronger His 1277,1287,1288 or Glu ligand, 1289−1291 the blue copper protein azurin can be converted to a green copper protein. By mutating Met to a weaker ligand such as Thr, the natively green copper protein, nitrite reductase, has been converted to a blue copper protein. 1292 Recently, the axial Met was mutated to Cys, a strong ligand, and then to the unnatural amino acid homocysteine (Hcy), a strong ligand with a longer side chain. The resulting Met121Cys azurin has an additional ∼450 nm absorption, while in Met121Hcy the ∼410 nm peak dominates over the ∼625 nm peak. Together with EPR evidence, it was shown that, within the same scaffold, blue copper protein azurin was converted to a green copper protein and then to a red copper protein. 1293 Interestingly, the engineered red copper protein, Met121Hcy azurin, has a low reduction potential (113 mV) similar to that of nitrosocyanin (85 mV). 4.4.2 Role of His Ligands Although equatorial His residues are highly conserved in T1 copper proteins, their mutation does not impair the copper binding ability of the protein. Canters and co-workers mutated two His residues into Gly separately, and the resulting protein still had T1 characteristics. 1294,1295 As His to Gly mutation creates extra space around copper, exogenous ligands such as halides, azides, and imidazoles could diffuse into His46Gly and His117Gly azurins and coordinate with copper. Depending on the type of external ligand, the mutants will be either T1 or T2 copper proteins. 1294−1296 His117Gly and His46Gly mutations also changed solvent exposure of the copper site. Without external ligands, His117Gly azurin has a reduction potential of 670 mV, much higher than that of WT azurin (310 mV). The high reduction potential is due to loss of a water ligand during reduction. Addition of external ligands will lower the reduction potential. 1297 The open coordination site of His117Gly makes it possible to study ET using imidazole-modified complexes. 1298,1299 The mutants generally exhibit a lower ET rate. As the properties of exogenous imidazoles affect the ET rate, it has been suggested that His is also important in the WT protein. 1300 4.4.3 Role of Cys Ligands As the Cu–S(Cys) bond defines the properties of type I copper sites, 99 mutation of Cys to other natural amino acids will dramatically alter the copper site in T1 copper proteins (Figure 55). Substitution of any other amino acid for Cys will result in loss of the intense LMCT bands, which is due to the interaction of the Cys S with copper. As an isostructural analogue of Cys, selenocysteine (SeC) can replace Cys without major structural perturbation. This strategy has been employed as a spectroscopic probe for T1 copper centers. 1301−1303 The protein Cys112SeC azurin showed a reduction potential similar to that of WT azurin (328 mV vs 316 mV at pH 4) and a red-shifted LMCT band at 677 nm. 1301 So far, only Cys112Asp mutation in azurin has been characterized. Mutation of Cys to Asp makes azurin a T2 copper protein, as evidenced by large hyperfine splitting (A || ≈ 152 × 10–4 cm–1) in the EPR spectrum and slow ET. 1304−1307 Addition of another mutation at the axial position, Met121Leu (Phe/Ile), results in a novel copper center called type 0 copper, which has the small parallel hyperfine splittings and rapid ET characteristic of T1 copper centers but no longer fits the classification of T1 copper due to the loss of the copper–thiolate interaction. 1308−1311 Moreover, there is only a slight increase of the reorganization energy to 0.9–1.1 eV compared with that of WT azurin, much less than that of T2 copper proteins. The ET rate of type 0 copper protein is 100-fold faster than that of the Cys112Asp mutant, a typical T2 protein. 1308,1309,1311 Figure 55 Active sites of type 2, type 1, and the newly constructed type 0 copper. In the center, a plot shows (in the shaded ovals) the typical values of two electron paramagnetic resonance spectroscopy parameters, A ∥ and g ∥, for type 1 (lower) and type 2 (upper) copper sites and the values of type 0 copper (green, red, and black points, right center), showing that type 0 copper does not fall into the typical ranges for these other kinds of sites. Reprinted with permission from ref (1308). Copyright 2009 Macmillan Publishers Ltd. 4.4.4 Role of Structural Features in the Secondary Coordination Sphere Copper ligands exert a significant influence on the spectroscopic features and reduction potentials of T1 copper proteins. However, copper ligands cannot fully account for variation in the reduction potentials of T1 copper proteins. Mutation of copper ligands usually results in loss of T1 characteristics or reduction of ET activity. For the limited mutations that maintain T1 characteristics and ET activity, the reduction potential is tuned over a 227 mV range by introducing Met analogues at the axial position, which is far less than the 600 mV range reported in native proteins. 1285 As discussed in section 4.3.3, the H-bonding network beyond the T1 copper center plays an important role in maintaining the structure and function of T1 copper centers. Mutagenesis studies focusing on changes of hydrogen bonds have revealed important information about how the reduction potential and other properties are tuned in T1 copper proteins. Rusticyanin has a higher potential relative to other T1 copper proteins. By sequence comparison, it is identified that there is a Ser in rusticyanin at the position corresponding to Asn that “zips” two ligand loops together. Asn has been proposed to decrease the E m by strengthening the H-bonding interactions between two ligand-containing loops. Mutating Ser86 in rusticyanin to Asn established such a hydrogen bond and lowered the E m by 77 mV. 1312 On the other hand, changing Asn in azurin to Ser eliminates one hydrogen bond between two loops (Figure 56) and results in a protein with a 131 mV higher reduction potential. 1293 Figure 56 X-ray structures of Az and selected variants. (a) Native azurin (PDB ID 4AZU). (b) N47S/M121L azurin (PDB ID 3JT2). (c) N47S/F114N azurin (PDB ID 3JTB). (d) F114P/M121Q azurin (PDB ID 3IN0). Copper is shown in green, carbon in cyan, nitrogen in blue, oxygen in red, and sulfur in yellow. Hydrogen-bonding interactions are shown by dashed red lines. Reprinted with permission from ref (1088). Copyright 2009 Macmillan Publishers Ltd. By comparing certain cupredoxins that natively have lower E m than the rest of the family, it is observed that they share a conserved Pro residue that is two residues after the copper-ligating Cys. 114,1313 The backbone amide in the equivalent residue in azurin hydrogen bonds to the thiolate of Cys112. 1160 Placing a Pro in this position converts this secondary amide to a tertiary amide, which is incapable of donating a hydrogen bond. The Phe114Pro mutant has a lower reduction potential. 114 It is proposed that deleting the hydrogen bond to the thiolate gives Cys112 more conformational freedom, and it allows for the electron density that was previously tied up in a hydrogen bond to contribute to the Cu–SCys interaction. 114 Another examination of cupredoxin crystal structures reveals the presence of backbone carbonyl oxygen from Gly45 near the copper ion in azurin, which is missing in other cupredoxins such as rusticyanin. 97,98,1314 This ionic interaction in azurin is proposed to result in higher electron density near the copper, preferentially stabilizing the Cu(II) form of the protein and, therefore, lowering the E m. 98,481,1315 Phe114Asn mutation was made to hydrogen bond with Gly45 backbone carbonyl and decrease the effect of carbonyl oxygen in Az. The mutant showed a 129 mV higher reduction potential compared to that of the wild type. 1088 With all of these individual factors in mind, Lu and co-workers combined mutations on both the copper ligands and residues in the secondary coordination sphere. These mutations showed an additive effect on the reduction potential in azurin. With different combinations, the reduction potential was tuned from 90 to 640 mV, which is beyond the reported range of native T1 copper proteins and their mutants (Figure 57). 1088 Figure 57 Reduction potentials for a number of Az mutants versus a measure of the hydrophobicity (log P), revealing the linear trend with respect to the axial position (residue 121). Reprinted with permission from ref (1088). Copyright 2009 Macmillan Publishers Ltd. Unlike mutations on the copper ligands, mutations of residues in the secondary coordination sphere are less likely to change the T1 characteristics according to UV–vis, EPR, 1293 and resonance Raman 1316 spectroscopy. DFT studies were able to separate the effects of covalent interaction and nonlocal electrostatic components; while the covalent and nonlocal electrostatic contributions can be significant and additive for active H-bonds, they can be additive or oppose one another for dipoles (Figure 58). 1317 Figure 58 Illustration of the experimentally derived covalent and nonlocal electrostatic contributions to E° for the variants of Az relative to WT Az and their comparison to calculations. Reprinted from ref (1316). Copyright 2012 American Chemical Society. Lower reorganization energies in the ET process generally increase the ET rate constants and efficiency. However, rational design of ET centers to lower the reorganization energy has so far not been demonstrated. Such a task is particularly challenging for ET proteins such as the blue copper protein azurin that have already been shown to possess very low reorganization energies in comparison to the majority of the other proteins. A study of intramolecular ET by pulse radiolytically produced disulfide radicals to Cu(II) in the above rationally designed azurin mutants showed that the reorganization energies of the designed mutants are lower than that of WT azurin, increasing the intramolecular ET rate constants almost 10-fold. 1317 More interestingly, analysis of structural parameters of these mutants suggested that this lowering in reorganization energy is correlated with increased flexibility of the copper center. 4.4.5 Role of Ligand Loop Besides directly mutating individual ligands, loop-directed mutagenesis containing the ligands to the copper center enables manipulation of copper center by changing the protein structure on a broader scale. T1 copper proteins and CuA domains in heme–copper oxidases share the same cupredoxin fold, with three ligands of T1 copper and four ligands of CuA residing in the so-called “ligand loop” (Figure 59). By careful design, it is possible to transplant the ligand loop of one protein into another, enabling interconversion between T1 copper and CuA and between different T1 copper proteins (section 4.5.3). Figure 59 Ligand and loop structure in different T1 copper proteins, CuA from T. thermophilus heme–copper oxidase, and red copper protein nitrosocyanin: (A) amicyanin (PDB ID 1AAC); (B) pseudoazurin (PDB ID 1PAZ); (C) plastocyanin (PDB ID 1PLC); (D) azurin (PDB ID 2AZA); (E) rusticyanin (PDB ID 1RCY); (F) CuA from T. thermophilus heme–copper oxidase (PDB ID 1CUA); (G) nitrosocyanin (PDB ID 1IBY). An early example of loop-directed mutagenesis comes from interconversion between different copper centers, as two research groups independently installed a ligand loop from the CuA domain of cytochrome c oxidases on amicyanin and azurin, converting the T1 copper proteins to a CuA protein, 1318,1319 discussed in detail in section 4.5. Recently, Berry and co-workers transplanted the ligand loop of nitrosocyanin, a newly discovered red copper protein, to azurin. 1320 The resulting protein, NCAz, has UV–vis and EPR features similar to those of nitrosocyanin despite having His instead of Glu as the fourth ligand. Although the T1 copper proteins have a conserved ligand set (section 4.3.1.1), the ligand loops from different proteins show variation in length and sequence (Figure 59). Loop-directed mutagenesis has been carried out between different T1 copper proteins. Ligand loops from azurin, pseudoazurin, plastocyanin, rusticyanin, and nitrite reductase were introduced into the amicyanin scaffold to create loop elongation mutants. 1321−1324 Later, the ligand loop from amicyanin, which is the shortest among T1 copper proteins, was introduced into azurin, pseudoazurin, and plastocyanin scaffolds to create loop contraction mutants. 1325,1326 The ligand loop from plastocyanin was introduced into the azurin scaffold as well. 1327 All of the loop-directed mutants maintain T1 copper spectroscopic characteristics, indicating a similar structure in the Cu(II) state. On the other hand, the loop length has been shown to affect the pK a of C-terminal His and the Cu(I)–N(His) distance. 1326,1327 It has been observed that introducing the short loop of amicyanin into pseudoazurin and plastocyanin increases the pK a of C-terminal His, probably due to an entropically favored Cu(I)–N(His) interaction with a longer, more flexible loop. 1324−1326 As expected, the reduction potentials of loop-directed mutants are between the reduction potentials of donors of the loops and scaffolds. Amicyanin has the second lowest reduction potential in T1 copper proteins (see Table 10). Introducing the amicyanin loop into other copper protein scaffolds decreases their reduction potentials by 30–60 mV. 1326 On the other hand, introducing loops of other T1 copper proteins into amicyanin increases its reduction potential. 1322−1324 The ET activity of loop-directed mutants has been measured by the electron self-exchange rate constant (k SES). The loop elongation mutants generally have 10-fold lower k SES values, while loop contraction has less influence on k SES. 1322,1323,1326 All the studies indicate that, T1 copper proteins can accommodate changes in loops and assume the same active site structure, consistent with the “rack” or entatic state of the T1 copper center. 95,1168,1170 4.5 CuA Centers 4.5.1 Overview of the CuA Centers The CuA is a binuclear copper center bridged by two cysteine ligands to form a Cu2S2 “diamond-core” structure, which has been found naturally in CcOs, 1030,1109,1328 nitrous oxide reductases (N2ORs), 1329,1330 the oxidase from Sl. acidocaldarius (SoxH), 1331 and a nitric oxide reductase (qCuANOR) 1332,1333 to date (Figure 60). Interestingly, all of these proteins are terminal electron acceptors of ET processes; e.g., CcO is the terminal electron acceptor in aerobic respiration, and N2OR is the terminal electron acceptor in anaerobic respiration. One of the most important features of the CuA center is that the two copper ions form a direct metal–metal bond. Therefore, the unpaired electron is delocalized between two copper ions, and the resting state of the CuA center is a Cu(+1.5)–Cu(+1.5) state rather than a Cu(+2)–Cu(+1) state. This is the first example of a metal–metal bond found in biology, which makes it unique compared to centers of other metalloproteins. In addition to the bridging Cys ligands, the copper ions are coordinated by a His from the equatorial position to form a trigonal NS2 coordination. There is a weak distal axial ligand on each copper ion. The axial ligands are a methionine at one copper and a backbone carbonyl at the other. Considering only each copper ion, the CuA center is very similar to the T1 blue copper center with an overall distorted tetrahedral geometry. Hence, the CuA center can be treated as two T1 copper centers joined together with a Cu–Cu bond in between, suggesting an evolutionary relationship between these two centers. Indeed, such a relationship has been proposed on the basis of three-dimensional structure comparison and construction of phylogenetic trees, indicating that T1 copper and CuA proteins share a common ancestor and are developed in part by divergent evolution. 1334,1335 Figure 60 Crystal structures of cytochrome c oxidase (PDB ID 3HB3) and nitrous oxide reductase (PDB ID 1FWX). The CuA sites are highlighted (copper is in green, sulfur is in yellow, nitrogen is in blue, and carbon is in cyan). The UV–vis absorption spectrum of CuA shows two intense absorbance bands at ∼480 and 530 nm that arise from S(Cys) → Cu charge transfer in the visible region and also a broad band at ∼760–800 nm that arises from Cu(+1.5)–Cu(+1.5) intervalence charge transfer. 860,1113−1115 The reduced Cu(I)–Cu(I) form is colorless because of the d 10 electronic configuration at each copper center. The more oxidized Cu(II)–Cu(II) state has not been observed in proteins to date. 1336,1337 Attempts to oxidize the CuA site normally give an irreversible anodic current at around 1 V, probably due to oxidation of the bridging dithiolate to disulfide. 1337,1338 Therefore, the CuA site acts as a one-ET center under physiological condition. 72 The Cu–Cu bond in CuA sites has been the subject of extensive debate. 1353 Later, the structure of the CuA site was confirmed by different spectroscopic methods. Blackburn et al. reported the extended EXAFS studies of the CuA binding domain of B. subtilis CcO, which showed a strong Cu–Cu interaction of ∼2.5 Å together with a short 2.2 Å Cu–S interaction. 1108 The Cu–Cu bond distance is nearly identical to the results from EXAFS studies of native CcO from bovine heart mitochondria, which is 2.46 Å. 1354 The dinuclear nature and the unusually short Cu–Cu distance of ∼2.55 Å were further established by X-ray crystal structures of CcO from Pa. denitrificans and bovine heart mitochondria, 1030,1109 as well as an engineered CuA center in CyoA. 1349 Similar structural features were also observed in the crystal structure of N2OR from Ps. nautica. 1329,1330 The most intense bands at 339, 260, and 138 cm–1 observed in resonance Raman spectroscopy of the Pa. denitrificans CcO CuA domain were assigned to symmetric stretches involving primarily the Cu–S(Cys), Cu–N(His), and Cu–Cu bonds, respectively. 1107 The Cu–Cu bond in the CuA site causes a valence delocalization between the two copper ions and produces a seven-line hyperfine splitting pattern in the EPR spectra. This unique EPR pattern can be explained by the delocalized unpaired electron coupled with two equivalent copper ions with a nuclear spin I = 3/2. 1112,1355,1356 Compared to centers in T1 blue copper proteins, CuA centers show even smaller A ∥ on the basis of EPR simulations, 1114,1339,1342,1345,1346,1357 reflecting greater covalent interaction and unpaired electron delocalization between the copper ions and the bridging Cys residues. 4.5.2 CuA Centers in Water-Soluble Domains Truncated from Native Proteins Historically, studying the biochemical role and probing the unique structure of CuA centers have not been easy due to complications arising by overlapping spectroscopic features of other metal centers present in the native proteins containing the CuA center. For instance, the CcO is a membrane protein containing two heme groups (heme a and heme a 3), two copper centers (CuA and CuB), a zinc ion, and a magnesium ion. To overcome these inherent difficulties in studying native CuA centers, two strategies are developed: producing truncates of native proteins containing CuA sites 742,1331,1339,1341,1342,1345,1352,1358−1361 and designing CuA centers into small, soluble proteins. 1318,1362,1363 In the first strategy, the sequence of the CuA subunit from CcO or SoxH was isolated and recombinantly expressed without the helices that normally anchor this domain to the membrane. This way, a water-soluble protein containing only the CuA site was obtained. Such truncates have been constructed for CcO from B. subtilis, 1345 Pa. dentrificans, 742,1339,1358,1361 Procambarus versutus, 1360 Synechocystis PCC 6803, 1352 and T. thermophilus(1341,1342,1359,1361) and for SoxH from Sl. acidocaldarius. 1331 The UV–vis, EPR, and EXAFS spectroscopic characterizations as well as the reduction potentials measurments for these soluble truncates are consistent with each other (Table 13). 742,1339,1358,1361 To date, only the truncate from T. thermophilus has been successfully crystallized. 1359 Table 13 Summary of the Spectroscopic Parameters of CuA Sites in Different Proteins CuA site containing protein organism λmax (nm) (extinction coefficient, M–1cm–1) reduction potential vs SHE (mV) ERP params (g x , g y , g z ) Cu–Cu distance (Å) ref subunit II of cytochrome c oxidase Paracoccus denitrificans 363 (1200), 480 (3000), 530, 808 (1600) (pH 7) 240 g x  = g y  = 2.03, g z  = 2.18, A z  = 3.5 mT 2.6 (742, 1109, 1339) subunit II of cytochrome ba 3 Thermus thermophilus 363 (1300), 480 (3100), 530 (3200), 790 (1900) 250 (pH 8.1), 240 (pH 8), 297 (pH 4.6) g x  = 1.99, g y  = 2.00, g z  = 2.17, A z  = 3.1 mT 2.43 (1337, 1341−1344) subunit II of caa 3-type cytochrome c oxidase Bacillus subtilis 365, 480, 530, 775–800   g x  = g y  = 1.99–2.03, g z  = 2.178, A z  = 3.82 mT 2.44 (1344, 1345) nitrous oxide reductase Paracoccus dentrificans 480, 540 (1700), 800       (1330) nitrous oxide reductase Pseudomonas stutzeri 480, 540   g x  = g y  = 2.03, g z  = 2.18, A z  = 3.83 mT 2.44 (1346) nitrous oxide reductase Achromobacter cycloclastes 350, 481 (5200), 534 (5300), 780 (2900)   g x  = g y  = 2.045   (1347) biosynthetic model in CyoA protein Escherichia coli 360, 538 (2000)   g x  = 2.03, g y  = 2.03, g z  = 2.18, A z  = 6.8, 5.3 mT 2.48 (1114, 1348, 1349) biosynthetic model in amicyanin   360, 483, 532, 790   g x  = g y  = 1.99–2.02, g z  = 2.18, A z  = 3.24 mT   (1318) biosynthetic model in azurin   360 (550), 485 (3730), 530 (3370), 770 (1640)   g x  = g y  = 2.06, g z  = 2.17, A z  = 5.5 mT 2.39 (1319, 1350) nitrous oxide reductase Pseudomonas nautica 617 480, 540, 800 260 g x  = g y  = 2.021, g z  = 2.178, A z  = 7 mT   (1351) subunit II of SoxM Sulfolobus acidocaldrius 361 (2300), 478 (3200), 538 (3700), 789 (2400) 237 g x  = g y  = 2.01, g z  = 2.20   (1331) subunit II of cytochrome c oxidase Synechocystis PCC 6803 359 (1580), 482 (2820), 535 (3080), 785 (1840) 216 (pH 7)     (1352) 4.5.3 Engineered CuA Centers The second strategy to study CuA sites is designing this site into other proteins, first accomplished in a quinol oxidase. 1362 The authors aligned subunit II of cytochrome c and quinol oxidases and found that the C-terminal of both proteins contained a subdomain in a Greek key β-barrel scaffold. This alignment suggested that both proteins contain a basic structural motif characteristic of cupredoxins. The CyoA lacked the putative ligands for the formation of the CuA in CcO. The CuA ligand set was thus introduced by extensive mutagenesis of the isolated cupredoxin domain. This engineered CyoA bound copper and showed two strong peaks at 358 and 536 nm, a shoulder at 475 nm, and a broad peak between 750 and 780 nm, as well as an EPR pattern similar to that observed in native CuA from CcO. Later, the crystal structure of CyoA was reported with 2.3 Å resolution. 1349 The distance between the two coppers is 2.5 Å. Shortly after the release of the purple CyoA study, two other research groups independently developed designed CuA centers in T1 copper proteins. 1318,1319 Dennison et al. replaced the C-terminal loop of the blue copper protein amicyanin, which contained three of the four T1 Cu-binding ligands, with a CuA binding loop. After copper binding, a purple protein was produced with UV–vis absorbance at 360, 483, and 532 nm and a broad absorption at approximately 790 nm, almost identical to that of the native CuA domain of CcO from B. subtilis. 1318 The EPR spectrum of the CuA amicyanin contained signals from two Cu(II) species; a distinctive T2 copper site, and a CuA center. 1364 Hay et al. constructed a CuA protein from a recombinant T1 copper protein, Ps. aeruginosa azurin, by replacing the loop containing the three ligands to the blue copper center with the corresponding loop of the CuA site in CcO from Pa. dentrificans. 1319 The UV–vis and EPR spectra of this protein (CuAAz) were remarkably similar to those of native CuA sites in CcO from Pa. dentrificans (Figure 61). The UV–vis absorption spectrum of CuAAz features two S(Cys) → Cu CT bands at 485 (ε ≈ 3700 M–1 cm–1) and 530 nm (ε ≈ 3400 M–1 cm–1), 1113,1350 compared to 480–485 and 530–540 nm for native CuA centers. 98 CuAAz also showed a broad band centered at 760–800 nm (ε ≈ 2000 M–1 cm–1), typical of the Cu–Cu ψ → ψ* transition, suggesting that CuAAz had reproduced the Cu–Cu bond. Additionally, the EPR spectrum of CuAAz displayed a seven-line hyperfine splitting pattern, demonstrating that this biosynthetic model duplicated the mixed-valence ground state of native CuA centers. 1319,1350 EXAFS, CD, MCD, and resonance Raman analyses of the CuAAz also suggested a high level of electronic and structural identity with CuA centers from CcO. 1113,1319,1350,1364,1366 The X-ray crystal structure of CuAAz showed a very similar arrangement of ligands around the copper ions and a Cu–Cu distance that was even slightly shorter than the native CuA center in CcO, confirming the presence of a Cu–Cu bond. 1367 Figure 61 (A) Crystal structure of the biosynthetic model of the CuA site in azurin (PDB ID 1CC3). (B) Comparison of UV–vis spectra between the soluble CuA domain in cytochrome c oxidase (green line), wild-type azurin (blue line), and the biosynthetic CuA model in azurin (purple line). (C) Comparison of X-band CW EPR between wild-type azurin (blue line) and the biosynthetic CuA model in azurin (purple line), four-line splitting vs seven-line splitting. Reprinted with permission from ref (1365). Copyright 2010 Springer-Verlag. 4.5.4 Mutations of the Axial Met The weaker axial methionine ligand has been investigated through mutagenesis in CcO from Pa. denitrificans and Rb. sphaeroides. The Met227Ile mutation in CcO from Pa. denitrificans resulted in a protein with unchanged stoichiometry of the metals. However, the two copper ions in the CuA site were no longer equivalent and converted from a delocalized Cu(+1.5)–Cu(+1.5) system to a localized Cu(+1)–Cu(+2) system on the basis of EPR and near-IR studies. 1368 The ET from cytochrome c to CuA was not affected, but the rate of ET to heme a was significantly diminished in the mutant protein compared with the wild-type protein due to an altered reduction potential of the CuA site. It was concluded that the weak axial Met was not essential for copper binding, but it was important for maintaining the mixed-valence electronic structure of the CuA site. The Met263Leu mutation in CcO from Rb. sphaeroides also showed the binding of two copper ions and proton pumping activity. Multifrequency EPR studies showed that the two copper ions in the CuA site were still electronically coupled. While all the other metals remained unchanged on the basis of UV–vis, EPR, and FTIR spectroscopy, the mutant only maintained 10% of the activity 1369 of the native enzyme. The kinetic analysis of ET showed a decrease of ET rate from heme c to CuA to 16 000 s–1 in the mutant, compared to 40 000 s–1 in the wild type. The rate constant for the reverse reaction was increased to 66 000 s–1, compared to 17 000 s–1 in the wild type. This result was attributed to an increased reduction potential of 120 mV relative to that of the native enzyme. 1370 The perturbation of the weak axial methionine ligand was also tested in the soluble CuA-containing subunit of cytochrome ba 3 from T. thermophilus. 1357 The mutants, Met160Gln and Met160Glu, affected the g z region of the EPR spectra and the Cu hyperfine became more resolved and larger in both mutants. Notably, the A z values of both mutants were increased from 3.1 to 4.2 mT, larger than most of characterized native CuA sites. The UV–vis spectra showed enhanced intensity and a blue shift relative to that of the wild type. The EPR and UV–vis data suggested that the axial ligand to copper interaction became stronger, moving from WT to Met160Gln and then to Met160Glu. The effects of both mutations were further studied by pulsed EPR/ENDOR spectroscopy. 1371 The results from this study showed an increase of A ∥, larger hyperfine coupling, and reduction in the isotropic hyperfine interaction and the axial g tensor. All these effects were associated with an increase in the Cu–Cu distance and changes in the geometry of the Cu2S2 core structure. The mutant Met160Gln was also studied by paramagnetic 1H NMR spectroscopy. 1372 The fast nuclear relaxation in this mutant suggested that a low-lying excited state had shifted to higher energies compared to that of the wild-type protein. Blackburn et al. reported a selenomethionine-substituted T. thermophilus cytochrome ba 3 and characterized it with Cu K-edge EXAFS. 1373 Interestingly, the optical and EPR spectra of the selenomethionine-substituted CuA site were essentially identical to those of the native CuA site as was the reduction potential. These data suggested that whatever role the S(Met) atom played in the electronic structure of the CuA site was also carried out by the Se(Met) atom. The axial Met in CuAAz was mutated to Asp, Glu, and Leu, covering the entire range of the hydrophobicity among the natural amino acids. The measured reduction potentials for these axial Met variants showed very little change, only about 20 mV, from that of the original CuAAz, despite some visible perturbation to the UV–vis and EPR spectra of these mutants. The significantly smaller effect of axial ligand in tuning reduction potential of CuAAz compared with WT-Az may reflect the resilience of the diamond core of CuA. In other words, the stability of the interactions making up the diamond core—the bridging Cys thiolates and copper–copper bond—may lead to greater resistance to perturbations arising from the axial position. 1374 Recently, a different set of axial Met mutants was generated in the truncated water-soluble CuA domain from T. thermophilus. 1375 By introducing Gln, His, Ser, Tyr, and Leu at the axial Met position, a change of about 200 mV in reduction potential was observed. The difference between the results from the truncated CuA domain and CuAAz was attributed to the difference in Cu–S(Met) bond lengths in these two systems: 2.47 Å in the truncated CuA domain vs 3.07 Å in CuAAz. Another explanation is that CuAAz contains the shortest Cu–Cu bond length (∼2.4 Å), hence enhances resistance of the diamond-core structure toward ligand changes. It is interesting to note that the reduction potentials of the native CuA site from the soluble fragment of subunit II of T. thermophilus ba 3 at different pH values showed no significant changes. 1376 However, the engineered CuA site in azurin exhibited strong pH dependence of the redox properties. This difference might be caused by protonation and dissociation of one of the histidine ligands in the engineered CuA center. 4.5.5 Mutations of the Equatorial His Ligands The equatorial His ligands bind to the copper ions with a bond length of ∼2.0 Å. In principle, mutations at this His position would result in a significant perturbation of the CuA site. This assumption has been proven to be true in the native system. The His260Asn mutant in cytochrome c oxidase from Rb. sphaeroides only exhibited 1% of the wild-type activity. 1369 The 850 nm band was shifted, and the extinction coefficient was diminished to around 1230 M–1 cm–1, compared with 1900 M–1 cm–1 in the wild type. No apparent hyperfine splitting pattern was observed in the EPR spectrum. The kinetic analysis of ET rates showed that the rate constant for ET from CuA to heme c was decreased to 11 000 s–1, compared to 40 000 s–1 in the wild type. The ET rate from CuA to heme c was decreased to 45 s–1, compared with 90 000 s–1 in the wild type. An increase of 90 mV in the reduction potential was also observed. 1370 However, dramatic differences were observed in the biosynthetic model of CuA in azurin. The mutation of His120 to Ala yielded a UV–vis spectrum similar to that of the original CuAAz, including the Cu–Cu ψ → ψ* band at ∼760 nm. 1377,1378 The EPR spectrum of His120Ala only showed a four-line hyperfine splitting pattern, suggesting that the active site had undergone a transformation to trapped valence, although a Q-band ENDOR study of His120Ala CuAAz showed evidence for the CuA site still being delocalized. 1379 Xie et al. applied a series of spectroscopic techniques, including EPR, UV–vis, MCD, resonance Raman, and XAS to both CuAAz and His120Ala CuAAz and correlated the results with DFT calculations. 1380 The surprising conclusion of this work was that a minute, 1% mixing of the 4s orbital of one copper ion into the ground-state spin wave function caused the collapse to a four-line hyperfine splitting pattern in the EPR spectrum of His120Ala, not a change from valence-delocalized to trapped valence. The resonance Raman and MCD spectra both demonstrated that the valence delocalization of the CuA center was still intact, although slightly perturbed, despite the loss of His120 as a ligand. The authors attributed the ability of CuA in azurin to remain valence-delocalized, even with the loss of such a strong ligand, to the large electronic coupling matrix element, which arises from the strong and direct Cu–Cu bond. Thus, the diamond core of CuA plays an immense role in the robust nature of this center. 4.5.6 Mutations of the Bridging Cys Ligands Mutagenesis studies of the CuA binding ligands in native CcO from Pa. denitrificans and N2OR from Ps. stutzeri have demonstrated that the cysteine ligands play an important role in the functions of the enzymes and the spectroscopic features of CuA. Mutating one of the two bridging cysteines to serine, Cys216Ser, in CcO from Pa. denitrificans resulted in a type 1 blue copper site with four-line EPR hyperfine splitting rather than the seven-line EPR signal observed in the CuA site, and only retained below 1% of the wild-type activity. The Cys216Ser mutant no longer exhibited the near-IR absorption in the optical spectrum, indicating the loss of the Cu–Cu bond. Mutation of the second cysteine, Cys220Ser, resulted in 5–10% of the wild-type activity. The higher activity in Cys220Ser is suggested to be due to the intact binuclear copper site on the basis of the metal/protein ratio and copper/iron ratio. 1381 The Cys618Asp mutant in N2OR resulted in almost complete loss of activity. The copper was bound only weakly and was hardly detectable on the gel filtration column. In contrast to the Cys618Asp mutant, the Cys622Asp mutant retained some copper binding ability and activity, although the characteristic multiline feature of the mixed-valence CuA was no longer resolved in EPR. 1382 Similar to the studies in the native system, the bridging Cys ligands were also individually mutated to Ser in the biosynthetic model of CuA in azurin. 1383 Although the resulting mutants still bound to the copper ions, the features of the Cu–Cu bond were completely lost in that the Cys112Ser mutant resulting in two T2 copper sites. The Cys116Ser mutation resulted in a T1 copper site. To account for the loss of symmetry in a single Cys to Ser mutant, a double Cys to Ser construct was made. 1384 At high pH, the double mutant indeed bound two coppers, but the EPR spectrum showed that the two copper ions were in two distinct T2 copper sites rather than a mixed-valence site with seven-line hyperfine splitting. 4.5.7 Tuning the CuA Center through Noncovalent Interactions The H-bonding and hydrophobic interactions around the active site of copper proteins can significantly tune the ET process. 1088 Two mutations, Asn47Ser and Glu114Pro, were made in CuAAz. 1385 Both the Asn47Ser and Phe114Pro mutations alter H-bonding interactions near the Cys112 ligated to a copper ion, but the Phe114Pro mutation decreases the reduction potential by deleting the hydrogen bond between Cys112 and the backbone NH group, 114 while the Asn47Ser mutation increases the reduction potential by affecting the rigidity of the copper binding site and most likely forming a direct hydrogen bonds between the protein backbone and Cys112 (Figure 62). 1088 Interestingly, by placing both CuA and T1 blue copper centers in the same scaffold of azurin, Lu and coworkers were able to demonstrate that the same mutations in the secondary coordination sphere resulted in similar decease or increase of the reduction potentials of the copper centers, but the magnitude of the effect is much smaller in CuA center, probably because its “diamond core” structure is more resistant to the perturbation (Figure 62). 1088 Figure 62 Tuning the reduction potential at blue copper azurin and CuA azurin by redesigning the second coordination sphere. The effects of these mutants are in the same direction, but the magnitude is smaller in the CuA site due to the electron delocalization between the two copper ions. Adapted with permission from ref (1385). Copyright 2012 The Royal Society of Chemistry. 4.5.8 Electron Transfer Properties of the CuA Center The CuA site is the point of entry of the electrons from cytochrome c. In CcO, the CuA receives electrons from cytochrome c and transfers them to cytochrome a. However, in N2OR, the CuA is believed to transfer electrons between cytochrome c and the catalytic site where nitrous oxide is reduced. The characterization of the ET between cytochrome c and cytochrome c oxidase has been a difficult problem. The stopped-flow method has been used to study the kinetics of electron transfer but does not have sufficient time resolution to monitor such a rapid ET process. The electron transfers between bovine cytochrome c oxidase and horse cytochrome c labeled with (dicarboxybipyridine)bis(bipyridine)ruthenium(II) were studied by laser flash photolysis. 1386 The electron was transferred from Lys25 ruthenium-labeled cytochrome c to the CuA site with a rate constant of 11 000 s–1. The CuA site then transferred an electron to cytochrome a with a rate constant of 23 000 s–1. Lys7, Lys39, Lys55, and Lys60 ruthenium-labeled derivatives showed nearly the same kinetics. The ET between the CuA site and heme a in bovine cytochrome c oxidase was measured by pulse radiolysis. 1387 The rate constant of ET was 13 000 s–1 from the CuA site to heme a, and 3700 s–1 for the reverse process. From this study a low activation barrier was observed, suggesting a small reorganization energy during the ET process. The method was also applied to study the electron transfer between the CuA site and heme a in cytochrome c oxidase from Pa. denitrificans. 1340 The ET rates were found to be 20 400 and 10 030 s–1 for the forward and reverse reactions, respectively. The T1 blue copper sites and CuA sites are commonly used as ET centers found in many biological systems. However, direct comparison between the ET rates of these two centers is difficult to achieve due to different protein scaffolds and redox partners. The engineered CuA site in azurin provides a great opportunity to eliminate the protein structure contribution to the ET process since the ET rates are measured in the same azurin scaffold. 1388 The authors first radiolytically reduced the disulfide bond within the azurin scaffold and then measured the long-range ET rate from the reduced disulfide bond to the oxidized CuA center. The rate constant of this intramolecular ET process in CuAAz is ∼650 s–1. Although CuAAz has a smaller driving force (0.69 eV for CuAAz vs 0.76 eV for blue copper azurin), the ET rate of CuAAz is almost 3-fold faster than for the same process in the WT-Az (∼250 s–1). The calculated reorganization energy of the CuA center is only ∼0.4 eV, which is 50% of that found for the blue copper azurin. The low reorganization energy of CuA was also observed in the truncated soluble CuA domain of CcO from T. thermophilus. 1337 Farver et al. studied the ET rates and reorganization energies of the mixed-valence CuAAz site and trapped-valence His120Ala CuAAz. 1389 They found that changing from the mixed-valence to the trapped-valence state increased the reorganization energy by 0.18 eV, but lowering the pH from 8.0 to 4.0 resulted in a ∼0.4 eV decrease in the reorganization energy, suggesting that the mixed-valence state only played a secondary role in controlling the ET property. 4.5.9 pH-Dependent Effects As an electron entry site for cytochrome c oxidase, the CuA center receives electrons from cytochrome c and transfers the electrons to the heme a site. The electrons are finally transferred to the heme a 3–CuB site where dioxygen reduction takes place. The reduction results in a proton gradient, which in turns drives the synthesis of ATP. For the CcO to function well, a regulator is needed for initiating and shutting down the whole ET process and dioxygen reduction reaction. A pH-dependent study on engineered CuAAz suggested that the CuA site may play such a role. 1390 CuAAz displayed a seven-line EPR hyperfine with a mixed-valence state. When the pH was decreased from 7.0 to 4.0, the absorption at 760 nm shifted to 810 nm; at the same time, a four-line EPR hyperfine was observed. The pH dependence was reversible, and the mixed-valence state was restored when the pH was increased back to 7.0. A dramatic increase in reduction potential was also observed from 160 to 340 mV when the pH was decreased from 7.0 to 4.0. It was identified that the protonation of C-terminal His120 caused such a pH-dependent transition, as the His120Ala mutation completely abolished this observation. A feedback mechanism was proposed to explain how the CuA site regulated the function of cytochrome c oxidase. The pumped proton may result in protonation of the C-terminal His and cause a different valence state of the CuA site. The increased reduction potential in the new state will stop the whole ET process and proton pumping (Figure 63). This hypothesis is further supported by ET studies in the His260Asn mutant in cytochrome c oxidase from Rb. sphaeroides which showed that protonation of the C-terminal histidine resulted in a change in the valence state and an increase of the reduction potential by 90 mV. 1370 The ET rate from the CuA site to heme a decreased by over 4 orders of magnitude. The His260 in cytochrome c oxidase corresponds to His120 in CuAAz. Figure 63 Schematic model of different states of the CuA center in cytochrome c oxidase: (A) mixed-valence form at neutral pH and (B) trapped-valence form at low pH. Subunit I is in light blue, and subunit II is in pink. Black arrows represent the flow of electrons, and orange arrows represent the flow of protons. Reprinted with permission from ref (1390). Copyright 2004 National Academy of Sciences. 4.5.10 Copper Incorporation into the CuA Center While numerous studies have established the structural features of CuA, the question of how copper ions are delivered into the CuA sites in vivo is still poorly understood. In the cytoplasm, copper levels are rigorously regulated, and free copper levels are extremely low and estimated to be at the attomolar level. 1391−1397 Although it has been proposed that a metallochaperone called Sco is responsible for metalation of the CuA site, delivering the copper ions to the CuA site in CcO by Sco proteins has not been demonstrated. 1398 Besides the delivery of copper ions by Sco proteins, another possibility is unmediated metalation. The CcOs from eukaryotes are located in mitochondrial membranes. 1399 In Gram-negative bacteria, CuA in CcO is exposed to the periplasmic space. However, in Gram-positive bacteria, CuA in CcO is exposed to the extracellular space. 1109,1393,1400,1401 In addition, the N2OR is a soluble protein also located in the periplasmic space. 1402 In periplasmic and extracellular spaces, copper levels are not regulated as rigorously as inside the cell, and the free copper ion concentration could be much higher. In fact, unmediated CuA metalation has been considered as a possibility for CuA metalation in N2OR. 1403−1405 From this view, the studies of free copper ion incorporation into CuA sites in vitro may provide important insights into this process, although they do not perfectly reflect the process in vivo. In an early study of CuAAz, the metalation of apo-CuAAz by adding a 10-fold excess of CuSO4 was studied by stopped-flow UV–vis spectroscopy. 1406 A single intermediate with intense absorbance at 385 nm was observed, which is characteristic of the Cys S → Cu CT bands of tetragonal T2 copper centers. 98,1095 This T2 copper intermediate formed with k obsd = 1.2 × 103 s–1 and subsequently decayed with k obs = 3.1 s–1; meanwhile the absorptions corresponding to the CuA site increased. An isosbestic point between the ∼385 nm band and the ∼485 nm band of the CuA site was observed, indicating the T2 copper intermediate was converted to CuA. Because only Cu(II) ion was added during metalation, a reducing agent must be supplied by the system itself to form a Cu(+1.5)–Cu(+1.5) site, indicating that the free thiols in apo-CuAAz were providing electrons by forming disulfide bonds. 1407−1409 Adding ascorbate or Cu(I) salt increased the yield of CuA center formation. Figure 64 Proposed mechanism of copper incorporation into the biosynthetic CuA model in azurin. Reprinted with permission from ref (103). Copyright 2012 Elsevier. A similar study was investigated in N2OR from Pa. denitrificans. 1410 Different from the previous study, two intermediates were observed upon adding Cu(II) salt. These two intermediates formed within a similar time scale and also decayed at the same time with simultaneous formation of CuA sites. Two isosbestic points were present between the absorption bands of both intermediates and the CuA absorption bands, strongly suggesting conversion of these intermediates to CuA. One of these two intermediates has spectral features typical of T2 copper centers with thiolate ligation, and another shows the characteristics of a T1 copper center. These observations suggested that the purple CuA site contained the essential elements of T1 and T2 copper centers and provided experimental evidence for a previously proposed evolutionary link between the cupredoxin proteins. 1334,1335 Guided by the observation of both T1 copper and T2 copper intermediates in the metalation of the CuA site in N2OR, the metalation of CuAAz was revisited by varying both the copper concentration and pH. 1411 When the CuAAz concentration was greater than the CuSO4 concentration, both T2 copper and T1 copper intermediates were observed, similar to the results obtained for N2OR. Global fitting of the UV–vis absorption kinetic data and time-dependent EPR together with previously studied mutants of CuAAz provided valuable information about the mechanism of copper incorporation where a new intermediate, I x , was observed. When Cys112 was mutated to Ser, a T2 copper site formed, with UV–vis and EPR spectra similar to those of the T2 copper intermediate. From this study it was inferred that the T2 copper intermediate is a capture complex with Cys116, which is also supported by the greater solution accessibility of this residue, compared to Cys112. Conversely, when Cys116 was changed to Ser, a T1 copper center formed, with UV–vis and EPR spectra nearly identical to those of the T1 copper intermediate (Figure 64). 1383 4.5.11 Synthetic Models of the CuA Center Another approach to study the CuA center is to synthesize small-molecule mimics of CuA. 1412 This has been proven to be a difficult task because of the formation of disulfide bonds between free thiols mediated by copper ions. 1338 Also, the most important feature in the CuA site, the diamond-core structure with Cu–Cu bond bridging by thiolates, is difficult to achieve. Besides the first coordination sphere, the second coordination sphere has also proven to be important in tuning the properties of the CuA site, which is even harder to mimic in small-molecule compounds. 1385 However, model compounds have met with varying degrees of success. 369,1413−1428 Houser et al. reported a fully delocalized mixed-valence dicopper complex with bis(thiolate) bridging which was the first closet small-molecule CuA mimic. The crystal structure of this model complex showed that the Cu2S2 core is planar with an average Cu–Cu distance of 2.92 Å. However, it is still longer than the Cu–Cu distance (2.46 Å by EXAFS 1354 and 2.55 Å by X-ray crystal structures 1030,1109 ) in native CuA centers. 1416 The EPR spectrum recorded at 4.2 K clearly showed the seven-line hyperfine splitting indicating the fully delocalized electronic structure. More recently, Gennari et al. reported a new bis(μ-thiolato)dicopper complex that mimicked most of the important spectroscopic features of the CuA site. 1429 Notably, this dicopper complex is the first CuA model with a Cu2S2 core that can be reversibly oxidized or reduced between the Cu(+1.5)–Cu(+1.5) state and the Cu(+1)–Cu(+1) state. However, the short Cu(+1)–Cu(+1) distance (2.64 Å) and long Cu(+1.5)–Cu(+1.5) distance (2.93 Å) significantly increased the reorganization energy of ET, which was much higher compared to the reorganization energy observed in the water-soluble CuA domain of T. thermophilus cytochrome ba 3. 1337 4.6 Structural Features Controlling the Redox Chemistry of the Cupredoxins 4.6.1 Role of the Ligands As the immediate residues that coordinate to the copper centers, the ligands exert a huge influence on the redox properties of cupredoxins. The strong Cu–thiolate bond(s) playd the dominant role in defining T1 Cu and CuA centers in both their electronic structures and ET functions. Except for a few unnatural amino acids, mutation of Cys will change the T1 copper character. The same happens in the CuA center in that mutation of Cys to Ser will result in either T1 or T2 center. The His residues are important for shielding the copper center from the solvent and for directing ET. C-terminal His is on a hydrophobic patch of T1 copper proteins. The hydrophobic patch directly interacts with redox partners of T1 copper proteins. Mutation of either His to Gly creates an open binding site, where external ligands could coordinate with copper and influence the properties of T1 copper proteins. Due to the open binding site, the His to Gly mutant exhibited a high reorganization energy and low ET rate. The axial Met is less conserved in T1 copper proteins. Besides Met, native T1 copper proteins could have the more hydrophilic Gln or the more hydrophobic, noncoordinating Leu/Phe at the axial position. There is a general trend that proteins with Gln as their axial ligand have the lowest reduction potentials, proteins with Met have intermediate reduction potentials, and proteins with Leu/Phe have the highest potentials. The reduction potential tuning role of the axial ligand has been further confirmed by mutagenesis studies. The correlation between the hydrophobicity of the axial ligand and the reduction potential has been established by incorporation of a series of Met analogues. The role of the highly conserved axial methionine ligand was performed by glutamate, aspartate, and leucine in the engineered CuAAz. 1374 In contrast to the same substitutions in the structurally related blue copper azurin, much smaller changes (∼20 mV) in reduction potential were observed, indicating that the diamond-core structure of the CuA site is much more resistant to variation in axial ligand interactions than the distorted tetrahedral structure of the blue copper protein. 4.6.2 Role of the Protein Environment The first coordination sphere directly affects the spectroscopic properties and ET of the T1 copper proteins. Beyond the first coordination sphere, the protein scaffold holds copper ligands together and forces trigonal geometry regardless of the oxidation state of copper, as suggested by the rack mechanism 1168 or the entatic state. 1170 Furthermore, the environment around the primary coordination sphere can fine-tune the electronic structure and redox properties of the copper centers by noncovalent interactions such as a H-bonding network to the copper ligands. 94,1119,1430 Through manipulation of H-bonding networks in the secondary coordination sphere, Marshall et al. managed to tune the reduction potential of azurin over the natural range while maintaining T1 character in the copper center. 1088 The same mutations that affected the noncovalent interactions in azurin were introduced to tune the reduction potentials of engineered CuAAz. 1385 The effects of these mutations were in the same direction, but with smaller magnitude in the CuA site due to dissipation of the effects by two copper ions rather than the single copper ion in blue copper proteins. All these findings are important in understanding the different roles of the two cupredoxins. Since the T1 blue copper proteins are used in a wide range of ET processes, the reduction potentials of the blue copper proteins need to be tuned to fit a wide range. Such a tuning is mainly achieved by changing the axial ligands and H-bonding network in the secondary coordination sphere. 95,1088 However, the CuA sites are only found in terminal electron acceptors with very small potential differences between redox partners where a wide range of reduction potentials is not preferred. The diamond-core structure of CuA sites decreases the reorganization energies and enables fast ET processes. 4.6.3 Blue Type I Copper Centers vs Purple Cu A Centers The type I blue copper centers are widely found as ET centers common in many biological systems. However, the CuA centers are only found in CcOs, N2ORs, and the oxidase from Sl. acidocaldarius (SoxH). Several key questions that have been raised regarding these sites are concerned with how such a mixed-valence binuclear copper site was selected, what the advantage of such a site compared to T1 blue copper sites is, and why the CuA sites are only found in terminal electron acceptors. To answer these questions, a direct comparison of the ET rates of these two centers is required. The engineered CuA site in azurin provides a great opportunity to eliminate the protein structure contribution to the ET process since the ET rates are measured in the same azurin scaffold. 1388 The CuAAz demonstrated that CuA is a more efficient ET site even with a smaller driving force between the reduced disulfide and CuA site than between the reduced disulfide and blue copper site. The calculated reorganization energy of the CuA site is only half that of the blue copper site, which is due to the rigid structure of the diamond core in the CuA site. Both CcOs and N2ORs are large enzymes that contain multiple ET sites. As the electrons are transferred along the chain, the difference in reduction potentials as the driving force must fall within a narrow range of values. In this case, the ET sites with lower reorganization energy would be preferred because the driving force might be small. 5 Enzymes Employing a Combination of Different Types of Electron Transfer Centers 5.1 Enzymes Using Both Heme and Cu as Electron Transfer Centers 5.1.1 Cytochrome c and CuA as Redox Partners to Cytochrome c Oxidases The CcO is a terminal protein complex in the respiratory electron transport chain located in the bacterial or mitochondrial membranes. This large protein complex receives four electrons from cyt c that are used to efficiently reduce molecular oxygen to water with the help of four protons from the aqueous phase without producing any reactive oxygen species such as superoxide and peroxide. In addition, it translocates four protons across the membrane, which establishes an electrochemical potential gradient used for ATP synthesis. Out of many different types of CcOs from various different organisms, the families involved in aerobic respiration that generally use cyt c as their biological electron donors are caa 3, aa 3, cbb 3, ba 3, co, bb 3, cao, and bd oxidases. 1431 Cyts caa 3 and cbb 3 oxidases contain a distinct cyt c domain integrated into the cyt c oxidase enzyme complex. Cyt aa 3 oxidase is the mitochondrial counterpart of cyt caa 3 except that it does not contain the cyt c domain at the C-terminal end of subunit II (Cox2) of the enzyme complex. Subunit II also contains the binuclear CuA center. Cyt cbb 3 oxidases do not contain the CuA center, but they contain both a monocytochrome c subunit (FixO or CcoO) and a dicytochrome c subunit (FixP or CcoP). 79,1432 Many facultative anaerobes use bo and bo 3 oxidases which use quinol as the substrate instead of cyts c. Depending on the organism, the cyts c are associated with the enzyme complex by either covalent or noncovalent interactions. 1433 For example, in the bacterium PS3, cyt c binds covalently to the protein complex at the C-terminal end of subunit II. 1434−1438 In Pa. denitrificans, the cyt c subunit is tightly bound to the oxidase subunit by covalent interactions and can be removed by treatment of a high concentration of detergent. In eukaryotes, cyts c bind to the cyt c oxidase loosely, which can be removed at high salt concentrations. Mammalian cyt c oxidases have been shown to bind one molecule of cyt c at a high-affinity site, which serves as the electron entry point. 1439−1441 There is evidence of the presence of a second low-affinity site, but the role of such secondary interactions between cyt c and the oxidase is not well-known. It has been shown that cyts c use a series of several (six or seven) positively charged lysines near the heme edge which form complementary electrostatic interactions with negatively charged carboxylates on the high-affinity site of subunit II of the oxidase. Such electrostatic interactions are important for placing the substrate in the correct orientation to bind to the oxidase complex. 1442,1443 Available data suggest that electrons are transferred from reduced cyt c, one at a time, to the oxidized CuA. 1444,1445 Then internal ET takes place from the reduced CuA to the LS heme a and to the binuclear active site consisting of HS heme a 3 and CuB where the dioxygen reduction takes place (Figure 65). It has been measured that the ET rate constant from CuA to heme a is 20 400 s–1 and the rate of the reverse process, from heme a to CuA, is 10 030 s–1 in Pa. denitrificans cytochrome c oxidase by pulse radiolysis. 1340 A similar study was also applied to cytochrome ba 3 from T. thermophilus, and the first-order rate constants are 11 200 and 770 s–1, respectively. 1340 Electron transfer from cyt c to CuA and CuA to heme a is fast, 1445,1446 while the ET from heme a to the heme a 3/CuB site is slow and has been proven to be the rate-limiting step of the reaction. 1447,1448 It has also been shown that the presence of CuA is not required for the oxidase activity as the deletion of the CuA gene from beef heart cyt c oxidase slows down the ET rate, but still maintains some oxidase activity. 1449,1450 Figure 65 Cyt c oxidase from Pa. denitrificans (PDB ID 3HB3). The ET pathway is shown with arrows. Binding of cyt c to the oxidase causes conformational changes in both the protein partners. 1451,1452 The major changes are observed upon reduction of the CuA and heme a centers. It has been proposed that the reduction of these two redox centers causes a conformational change of the binuclear active site from a closed to an open state that facilitates the intramolecular ET that couples the subsequent redox reaction and proton translocation. 1453−1456 NRVS on cyt c 552 from Hydrogenobacter thermophilus has indicated the presence of strong vibrational dynamic coupling between the heme and the conserved -Cys-Xxx-Xxx-Cys-His- motif of the polypeptide chain. 1457 Such vibrational coupling has been proposed to lower the energy barrier for ET by either transferring the vibration energy released upon protein–protein complex formation or by modulating the heme vibrations. A recent NMR study has shown that the hydrophobic residues near the heme of cyt c form hydrophobic interactions with cyt c oxidase and are major contributors to the complex formation, while the charged residues near the hydrophobic core dictate the alignment and orientation of cyt c with the enzyme to ensure efficient ET. 1458 The affinity of oxidized cyt c for complex formation with CcO is significantly lower, suggesting that ET is gated by the dissociation of oxidized cyt c from CcO. The rate of dissociation of oxidized cyt c is dictated by the affinity of oxidized cyt c for CcO that provides facile ET. 5.1.2 CuA and Heme b as Redox Partners to Nitric Oxide Reductases Although the NORs from Gram-negative bacteria use cyt c as the biological electron donor to the heme c, one NOR (qCuANOR) purified from the Gram-positive bacterium B. azotoformans shows the presence of a quinol binding site and uses the binuclear CuA site as an electron acceptor instead of heme c. 1332,1333 This family of NORs use melaquinol as the physiological electron donor to the CuA site instead of cyt c. Electrons are passed from melaquinol to the CuA site and are then transferred to the LS heme b and onto the binuclear active site consisting of a HS heme b 3 and a nonheme FeB site. 5.1.3 Cytochrome c and CuA as Redox Partners to Nitrous Oxide Reductases The N2OR is the last enzyme in the denitrification pathway which reduces nitric oxide to dinitrogen. 1329,1330,1459 N2ORs are homodimeric periplasmic enzymes containing the binuclear ET site CuA which receives electrons from cyt c and a tetranuclear catalytic site, CuZ. A unique N2OR has been reported from Wolinella succinogenes which has a C-terminal cytochrome c domain that is suggested to be the biological electron donor to the CuA center. 1460 5.2 Enzymes Using Both Heme and Iron–Sulfur Clusters as Electron Transfer Centers 5.2.1 As Redox Partners to the Cytochrome bc 1 Complex The coenzyme Q–cytochrome c oxidoreductase, also called the cytochrome bc 1 complex or complex III, is the third complex in the electron transport chain playing a crucial role in oxidative phosphorylation or ATP generation. The bc 1 complex is a multisubunit transmembrane protein complex located at the mitochondrial and bacterial inner membrane that catalyzes the oxidation of ubihydroquinone and the reduction of cyt c(1461) coupled to the proton translocation from the matrix to the cytosol. The catalytic core of the bc 1 complex consists of three respiratory subunits: (1) subunit cyt b that contains two b-type hemes, b L and b H, (2) subunit cyt c, containing a heme c 1, and (3) iron–sulfur protein subunit containing a Rieske-type [2Fe–2S] cluster (Figure 66). While in some α proteobacteria such as Paracoccus, Rs. rubrum, and Rb. capsulatus, this enzymatic core containing the three subunits is catalytically active, several additional (seven or eight) subunits are present in the mitochondrial cytochrome bc 1 complexes. 86,1462 Figure 66 Bovine cytochrome bc 1 complex (PDB ID 1BE3). Different ET domains and their cofactors are shown. bL = low-potential heme, bH = high-potential heme, and Q = ubiquinol. Electron transfer pathways are shown with arrows. Structures of the bc 1 complex from various resources such as yeast, chicken, 1029 rabbit, 1029 and cow 1026,1029,1463 show that the cyt b subunit consists of eight transmembrane helices designated as A–H. Hemes b L and b H are contained in a four-helix bundle formed by helices A–D and are separated by a distance of 8.2 Å. The axial ligands for both hemes are all His and are located in helices B and D. His83 and His182 are bound to heme b L, while His97 and His196 are axial ligands for heme b H. The cyt c subunit containing cyt c 1 is anchored to the membrane by a cytoplasmic domain and belongs to the Ambler type 1 cyt c based on the protein fold and the presence of the signature sequence -Cys-Xxx-Xxx-Cys-His-. Electron transfer has been proposed to occur through the exposed “front” face of the corner of the pyrrole II ring. 1029 One of the His residues that acts as a ligand to the [2Fe–2S] cluster is 4.0 Å from an oxygen atom of heme propionate-6 and 8.2 Å from the C3D atom of the heme edge of cyt c 1. Such proximity of the heme group and the Rieske-type cluster has been proposed to facilitate ET. Using this distance of 8.2 Å, a rough estimation of the ET rate from the iron–sulfur protein to cyt c 1 has been calculated to be 4.8–80 × 106 s–1. On the basis of the relative orientations of the prosthetic groups as discussed above, an ET pathway has been proposed where in round I an electron is transferred from a bound ubiquinol to the Rieske-type cluster into the cyt c 1 via its heme propionate-6 and out of cyt c 1 via its pyrrole II heme edge to the cyt c (not the same as cyt c 1). 78,1029 At the same time the low-potential heme (bL) pulls an electron from the ubiquinol and transfers it to the high-potential heme (bH), which is ultimately picked up by an oxidized ubiquinone. The same cycle is repeated in round II. Mitochondrial cyt c or bacterial cyt c 2 connects the bc 1 complex with the photosynthetic reaction center or cyt c oxidase. 80,1464 The mode of interaction between cyt c (or c 2) with its redox partners has been proposed to involve docking of cyt c with its solvent-exposed heme edge (called the “front” side). There are multiple dynamic H-bonding and salt bridge interactions between the cyt c and cyt c 1 of the bc 1 complex. 1465 The front side is composed of a ring of positively charged Lys residues near the exposed heme edge. The opposite side, called the “back” side, is composed of several negatively charged residues. This charge separation creates a dipole moment in both bacterial cyts c 2 and mitochondrial cyt c. 1466,1467 The positively charged front side forms complementary interactions with the negatively charged surface of its partner, which orients the electron donor in proper alignment for facile ET. EPR experiments with cyt c 2 from Rb. capsulatus have demonstrated that the dipolar nature of cyt c 2 influences its orientations, which facilitate ET to its partner under physiological conditions. 1468−1470 Rieske protein can accommodate three conformations in the complex: The first is the c1 position in which the His ligand is H-bonded to propionate of heme in cyt c, and fast ET (60 000 s–1) 1471 between the two will occur. 1026 At this state the cluster is far from the quinone binding site. The b position allows interaction between the cluster and quinone. This position was stabilized by interaction of H161 with the inhibitor stigmatellin that mimics the H-bond pattern of semiquinone. 223,1029 The final conformation is an intermediate state in which the Rieske protein cannot interact with either cytochrome or quinone. 865 The cycle starts from an intermediate state (Figure 67). Upon binding of reduced hydroquinone, the Rieske protein will move to state b and an electron will be transferred to hydroquinone, generating a semiquinone, which binds tightly to the Rieske protein. This tight interaction will become loose by transfer of a second electron from semiquinone to heme b L and generation of quinone. The thermodynamically disfavored reduction of heme b L by semiquinone is coupled to favorable oxidation of hydroquinone by the Rieske center. As a result the reduction potential of the Rieske center is of significant importance to the rate of reduction of heme b L. Reduction of the Rieske center and heme b L happens within a half-life of 250 μs as evident by freeze quench EPR. The semiquinone intermediate has a very high affinity for the Rieske protein. This tight binding will increase the reduction potential of the Rieske center by 250 mV. This binding mode and increased reduction potential will ensure that the Rieske center will not reduce cyt c before heme b L is reduced and quinone is formed. The reduced Rieske center will then move to its c 1 state and transfer an electron to cyt c. After complete transfer of both electrons, the Rieske protein will go back to its intermediate state for the second cycle. 773,787,865 The binding of quinone and Rieske protein is redox-dependent. While the kinetics of ET to cyt c is pH-dependent due to the pH dependence of the reduction potential, it has been proposed that the rate-limiting step in this reaction is mostly the transition from one state (e.g., state b) to another state (e.g., state c 1) of the Rieske center and not the ET, considering the same rate observed in mutants with different reduction potentials. 1078 Figure 67 Schematic cycle of Rieske positions in the bc 1 complex. Reprinted from ref (865). Copyright 2013 American Chemical Society. Although the mechanism of proton transfer is not very well understood in this system, evidence suggested that the two protons are bound to the Rieske center, one to each His in the reduced state. The oxidized state can have no protons, one proton, or two protons depending on the pH. It has been shown that removal or mutation of the Rieske cluster will result in a proton-permeable bc 1 complex, suggesting a role as a proton gate for the Rieske protein. 1472 NMR was used to calculate the pK a of His ligands in the T. thermophilus Rieske protein. In this study, residue-selective labeling was used to unambiguously assign the NMR shifts. The results were consistent with other pH-dependent studies of Rieske proteins, showing that one of the water-exposed His ligands that is close to quinone undergoes large redox-dependent ionization changes. Their system also supports proton-coupled ET in the Rieske–quinone system. 864 Analysis of driving forces using a Marcus–Bronsted method in mutants that had distorted H-bonding due to mutation of either conserved Ser or Tyr resulted in the proposal of a proton-first-then-electron mechanism in which the ET follows the transfer of a proton between hydroquinone and the imidazole ligand of the Rieske cluster. 814 5.2.2 As Redox Partners to the Cytochrome b 6 f Complex Cyt b 6 f (plastoquinol–plastocyanin or cyt c 6 oxidoreductase) is a protein complex belonging to a “Rieske–cytochrome b” family of energy-transducing protein complexes found in the thylakoid membrane in the chloroplasts of green algae, cyanobacteria, and plants and catalyzes ET from plastoquinol to plastocyanin or cyt c 6 (PSII to PSI) coupled with the proton translocation across the membrane for ATP generation. 282,1473−1476 It is located in between the PSII and PSI reaction centers in oxygenic photosynthesis (Figure 68). The b 6 f complex is analogous to the bc 1 complex of the mitochondrial electron transport chain. The b 6 f complex comprises seven subunits: a cyt b 6 with a low-potential (b p) and a high-potential (b n) heme, a cyt f, a Rieske iron–sulfur protein, subunit IV, and three low molar mass (∼4 kDa) transmembrane subunits. 1473 There are a total of seven prosthetic groups that are found in the b 6 f complex: cyt f, hemes b n and b p, a Rieske [Fe2–S2] cluster, chlorophyll a, β-carotene, and a c-type heme designated as c n, c x , or c i . This heme, located close to the quinone reductase site near the electronegative side of the membrane, is linked to the protein via a single thioether linkage, lacks any axial ligands, and has been shown to be critical for function of the b 6 f complex. 225,1478−1481 The cyt b 6 subunit contains two bis-His-ligated hemes, a high-potential heme (−45 mV) on the luminal side and a low-potential heme (−150 mV) on the stromal side of the thylakoid membrane. EPR and Mössbauer data reveal that both hemes are 6cLS and have His planes that are perpendicular. Cyt b 6 and subunit IV of the b 6 f complex are structurally similar to cyt c of the bc 1 complex, 184 while there is no structural similarity between cyt f and cyt c 1 even though they are functionally similar. 123,1029 The cyt b 6 f complex takes part in linear electron flow between PSII and PSI where it links the plastoquinone pool of PSII to plastocyanin or cyt c 6 to PSI as well as in cyclic electron flow within PSI (Figure 68). The linear electron flow path involves oxidation of quinol to quinone from PSII to PSI coupled to the generation of ATP and reduced ferredoxin, which reduces NADP+ to NADPH via an oxidoreductase FNR. Cyclic electron flow in PSI involves electron flow via the b 6 f complex back to the P700 reaction center of PSI. In both the cases two electrons are passed from plastoquinol at the quinol oxidation site (QP) near the lumenal, electropositive site of the membrane to the one-electron acceptor plastocyanin, which is coupled to the “Q-cycle”  1482,1483 involving proton translocation across the membrane. One of the electrons from plastoquinol is transferred to PSI via the high-potential chain, while the second electron is passed onto the low-potential, transmembrane chain on the electronegative side of the membrane where plastoquinone reduction takes place. Figure 68 Cyt b 6 f complex in the photosynthetic electron transport chain. P680 = reaction center chlorophylls of PSII, QA, QB = quinones of PSII, PQ/PQH2 pool = plastoquinone/plastoquinol pool, Fe–S = Rieske cluster, f = cyt f of the high-potential chains (blue arrows), Qp, Qn = plastoquinol oxidation and plastoquinone reduction sites, b p, b n, c n = hemes of the low-potential chain (red arrows), Fd = ferredoxin, and P700 = reaction center chlorophylls of PSI. The domain movement of the Rieske protein is shown by a two-sided arrow. The direction of proton translocation across the membrane is shown by proton arrows. The electronegative (cytoplasmic) (n) and electropositive (luminal) (p) sides of the membrane are labeled, and ET pathways are shown by arrows. A possible direct ET path from PSI to the cyt b 6 f complex is shown as the dashed line from Fd to the Qn site. Reprinted with permission from ref (1477). Copyright 2012 Springer Science+Business Media. On the His ligation side of the heme, a chain of five conserved water molecules oriented in an L-shaped manner have been identified from the X-ray structure, which form hydrogen bonds with ten amino acid residues from the protein, seven of which are conserved. 1473,1484,1485 These water molecules have been proposed to act as “proton wires” in coupling of the ET with proton transfer across the membrane. 1485,1486 The heme of cyt f is located in a hydrophobic environment and is protected from the solvent by Tyr1, Pro2, Ile3, and Phe4 (or Trp4 in cyanobacteria). 161 The side chain of residue 4 is located close to the heme edge and oriented almost perpendicular to the heme plane (Figure 69). 1485 This edge-to-face interaction of the Trp4 and the heme has been proposed to be responsible for tuning the reduction potential of the heme by interaction with the porphyrin π molecular orbitals. Such edge-to-face interactions have been observed in cyt b 5 (Phe58, Phe35), 141,366 cyt b 562 (Phe61), 382 and peptide-sandwich mesoheme model systems reported by Benson and co-workers (Trp or Phe). 423,1487 In these peptide mesoheme sandwich complexes the heme–Trp interaction has been shown to be important to stabilize the α-helical scaffold as well as the ferric state of the heme iron. 1488 Such interactions also stabilize the ferric state of the heme iron in the cyanobacterium cyt f. Figure 69 Environment around the heme of cyt f (PDB ID 1HCZ). Hydrophobic residues are shown as gray sticks. The “edge-to-face” interaction at 4 Å between Phe4 and the heme that is proposed to be important to tune the reduction potential of the heme iron is shown. The five conserved molecules that have been proposed to act as “proton wires” that couple ET with proton transfer are shown as red spheres. Residue numbering of waters is arbitrary. The chloroplast Rieske proteins work in the same way. It has been shown that the movement of these Rieske proteins will also function as a redox-state sensor that can balance the light capacity of the two photosystems. This state transition can also act as a switch between cyclic and linear electron flow. 1489 5.2.3 As Redox Centers in Formate Dehydrogenases Formate dehydrogenases (Fdh’s) catalyze decomposition of formate to CO2. They exist in both prokaryotes and eukaryotes. Fdh’s are mainly NAD+-dependent in aerobic organisms and NAD+-independent in anaerobic prokaryotes, donating electrons from formate to a terminal electron acceptor other than O2. 1490 Structural studies reveal that Fdh’s contain one to three subunits with either W or Mo in the active site. 1491−1493 Fdh-N from E. coli is among the most well studied Fdh’s. It is important in the nitrate respiratory pathway under anaerobic conditions. It is a membrane-bound trimer (α3β3γ3) with a molar mass of 510 kDa. It harbors a Mo-bis-MGD cofactor and a [4Fe–4S] cluster in the catalytic α subunit, four [4Fe–4S] clusters in the β subunit, and two heme b groups in the γ subunit (Figure 70). 1492 The β subunit transfers electrons between the α and γ subunits, similar to other membrane-bound oxidoreductases that bind four [4Fe–4S] clusters, such as nitrate reductases, [NiFe] hydrogenases, DMSO reductase, and thiosulfate reductase. 1494 Figure 70 Overall structure of Fdh-N from E. coli. Cofactors are displayed as spheres and denoted accordingly on the right. The putative membrane is shown in gray shading. PDB ID 1KQF. Reprinted with permission from ref (1492). Copyright 2002 American Association for the Advancement of Science. Fdh from Dv. desulfuricans is an αβγ protein with a molar mass of ∼150 kDa. It contains four different types of redox centers, including four heme c centers, two [4Fe–4S] clusters, and a molybdopterin. 1495 EPR studies showed the existence of two types of Fe–S clusters after reduction, i.e., center I with g values of 2.050, 1.947, and 1.896 and center II with g values of 2.071, 1.926, and 1.865. Midpoint reduction potentials of the two Fe–S clusters are −350 ± 5 mV for center I and −335 ± 5 mV for center II. Fdh from Dv. gigas is an αβ protein 1493 containing tungsten instead of molybdenum. It also possesses two [4Fe–4S] clusters similar to Fdh from Dv. desulfuricans. 981,1496 5.2.4 As Redox Centers in Nitrate Reductase NARs reduce nitrate to nitrite, a vital component in the nitrogen respiratory cycle. Most NARs isolated so far contain three subunits, NarG (112–140 kDa), NarH (52–64 kDa), and NarI (19–25 kDa). NarG harbors a Mo-bis-MGD cofactor and a [4Fe–4S] cluster, NarH contains one [3Fe–4S] cluster and three [4Fe–4S] clusters, and NarI immersed in the membrane binds two b-type hemes (Figure 71). 1497−1502 The overall folding and cofactor positions are strongly homologous to those of Fdh from E. coli. 1503 The eight redox centers are separated by 12–15 Å from each other and form an ET pathway about 90 Å long. NAR from Cupriavidus necator does not contain the NarH domain and harbors two c-type hemes in the small subunit. 1504 Figure 71 Overall three-dimensional structure of NarGHI from E. coli K12. PDB ID 1Q16. Subunit and cofactor names are denoted. Reprinted with permission from ref (1505). Copyright 2006 Elsevier. 6 Summary and Outlook This review summarizes three important classes of redox centers involved in ET processes. Although each class spans a wide range of reduction potentials, none of them can cover the whole range needed for biological processes. Together, however, they can cover the whole range, with cytochromes in the middle, Fe–S centers toward the lower end, and the cupredoxins toward the higher end (Figure 1). All three redox centers have structural features that make them unique, and yet they also show many similarities that make them excellent choices for ET processes. Because the redox-active iron is fixed into a rigid porphyrin that accounts for four of the iron’s six coordination sites, most of the electronic structure and redox properties remain similar between different cytochromes. In completing the primary coordination sphere of the iron, cytochromes typically use a combination of nitrogen and sulfur ligations from histidine or methionine side chains, respectively; terminal amine ligation has also been observed. In general, mutagenesis studies reveal that methionine ligation raises the reduction potential by 100–200 mV, relative to histidine ligation, primarily due to the lower affinity of thioether to the higher oxidation state of the heme, and that the effect is generally additive. 192,386,461−463,465 Heme puckering or flexing has been demonstrated to tune the reduction potentials by up to 200 mV. 513 Changes in the heme type between b and c would be expected to change the electronic properties of the heme; however, the effect on the reduction potential is small and varies depending on the systems studied. 446,448 It is clear, on the other hand, that the electron-withdrawing formyl group on heme a appears to be responsible for the increase in the reduction potential by ∼160 mV. 459,460 For iron–sulfur proteins, the reduction potential ranges are influenced to some extent by the number of irons because it affects the redox states and transitions. In the case of clusters with the same number of irons, the higher the redox pair, the higher the reduction potentials (e.g., HiPIPs have a [4Fe–4S]2+/3+ pair, while ferredoxins have a [4Fe–4S]1+/2+ pair). 719 In addition, the cluster geometry such as Fe–Sγ–Cα–Cβ torsional angles, the Fe–Fe distance, and covalency of Fe–S bonds also play important roles in some proteins. 618,901,1085,1506 Electron delocalization of the cluster and the net charge of the cluster are also important. For example, it has been shown that the net charge of the protein is the main factor determining the reduction potential within HiPIPs. Electrostatic effects of the charged residues in the secondary coordination sphere can influence the solvent accessibility and consequently the dielectric constant around the metal center. However, the effects are usually complicated and difficult to rationalize by just Coulomb’s law. For example, in rubredoxin from Cl. pasteurianum, replacement of a neutral surface residue by a positively charged Arg or a negatively charged Asp has led to an increase of reduction potentials in both cases. 611,612 Finally, the direct ligands to iron and H-bonding interactions with the direct ligands make significant contributions to the reduction potential. 541 When the common Cys thiolate ligand was replaced with a His imidazole ligand, naturally in the Rieske proteins, or with Ser by site-directed mutagenesis, the reduction potentials changed accordingly. 721,773,1087 The multiple NH···S H-bonding interactions in rubredoxin render the reduction potential of the [FeCys4] center to fall in the range of −100 to +50 mV, while reduction potential of the corresponding model complexes without the H-bonding networks is around 1 V. 92,588−590 The NH···S H-bonds have also been shown to be important in determining reduction potentials between different ferredoxins as well as ferredoxins vs HiPIPs. 617,618,718,719 For cupredoxins, the metal centers cannot be easily fixed like in either porphyrin or thermodynamically stable iron–sulfur clusters and proteins play a more prominent role in enforcing the unique trigonal geometry and strong copper–thiolate bond to maintain a low reorganization energy for the ET function. In this class of proteins, both the geometry and the ligands, particularly the strictly conserved Cys, play a dominant role in controlling the redox properties. In T1 copper protein azurin, changing axial Met to a stronger cysteine or homocysteine induced a geometry change and weakened the Cu–S bond. These changes in turn resulted in a >100 mV decrease in the reduction potential. 1293 Deleting the H-bonding to Cys, realized through the Phe114Pro mutation in azurin, affected the covalency of the Cu–S bond and lowered the reduction potential of azurin. 114,1088,1316 Despite the differences in the primary coordination spheres, all three redox centers employ noncovalent secondary coordination interactions in fine-tuning the redox properties. The first common feature is the control of the degree of solvent exposure; the deeper the redox centers are buried into the hydrophobic center of the protein, the higher the reduction potential and the smaller the changes in the reorganization energy due to influences by the solvent. For example, redox center burial is considered to be one of the main factors for differences in reduction potentials between different HiPIPs and ferredoxins. 618,719,749,752 Furthermore, a computational study of heme proteins over an 800 mV range has attributed the greatest correlation with the reduction potential to solvent exposure. 457 The second common feature is the electrostatic interactions. For example, the net charge of protein is shown to be the only factor that correlates with the reduction potentials of different HiPIPs. 715,752,890 The number of amide dipoles and not necessarily H-bonding is shown to be important in reduction potential determination in ferredoxins. 718,719 In myoglobin, Val68, which was in the van der Waals interaction distance with the heme group, was replaced by Glu, Asp, and Asn. A 200 mV decrease in reduction potential was observed for the Glu and Asp mutants compared to the wild type. 481 This study demonstrated that replacement of hydrophobic Val68 by charged and polar residues led to substantial changes in the reduction potential of the heme iron. In a number of different cytochromes, electrostatic polar and charged groups near the heme were shown to vary the potential by 100–200 mV. 169,479,481,482 For instance, in cyts c 6 and c 6A, the glutamine at positions 52 and 51, respectively, were shown to raise the potential ∼100 mV, 479 and in cyt c, the Tyr48Lys mutation raised the potential 117 mV; 480 all these effects can be attributed to charge compensation in the heme pocket. Similarly, replacing Met121 with Glu or Asp in T1 copper azurin resulted in 100 and 20 mV decreases in the reduction potentials, respectively. 1278,1289 Beyond copper ligands, mutating Met44 in azurin to Lys destabilizes Cu(II), causing a 40 mV increase of the reduction potential. 1507 The final common feature is the presence of a hydrogen-bonding network around the ligands to the metal center, especially those to the ligand that dominates the metal–ligand interactions. For example, the NHamide···Scys H-bonds are known to be important in different reduction potentials between rubredoxins, HiPIPs, and ferredoxins. 617,618,718,719 They are also shown to play a role in different reduction potentials of different ferredoxins. Other than backbone amide H-bonds, H-bonds from side chains are also important. A good example of such is H-bonds from conserved Ser and Tyr in Rieske proteins and a lack of thereof in Rieske-type proteins, hence differences in the reduction potential. 781 In cytochromes, H-bonding interactions with the axial ligands can tune the potential by up to 100 mV. 474,476,477,1508 For instance, increasing the imidazolate character of the axial His ligand in cyt c by strengthening H-bonding from the H to the Nε increased the potential by nearly 100 mV, 474 and disrupting the hydrogen bond donation from Tyr67 to the axial Met resulted in a 56 mV decrease in potential. 476,1508 Similarly, the H-bonding interactions to the Cys in cupredoxins are known be responsible for their reduction potential differences. 114 A test of how much we understand these structural features responsible for the redox properties is to start with a native redox center and use the above knowledge to fine-tune the redox properties. A pioneering work in this area is the demonstration of a ∼200 mV decrease in the reduction potential of myoglobin when a buried ionizable amino acid (Glu) was introduced into the distal pocket of the protein, and such a change has been attributed to electrostatic interactions. 481 Since then, not many examples have shown similar magnitude changes of reduction potentials by electrostatic interactions, perhaps due to the compensation effect by ions in the buffer or other ionizable residues nearby. Instead, hydrophobicity and H-bonding network have been shown to play increasing roles, and a combination of these effects has been shown to fine-tune the reduction potentials of T1 copper azurins by more than 700 mV, beyond its natural range. 1088 These features were further shown to be additive, making reduction potential tuning predictable. Such rational design also allowed the lowering of the reorganization energy of azurin, 1317 which is already known to be very low in comparison to those of other redox centers. With more such successful examples in other systems, we will be able to achieve a deeper understanding of ET reactivity in proteins and facilitate de novo design of ET centers for applications such as advanced energy conversions. Show more