Functional Divergence of Heme-Thiolate Proteins: A Classification

Review pubs.acs.org/CR

Functional Divergence of Heme-Thiolate Proteins: A Classification Based on Spectroscopic Attributes Aaron T. Smith,†,⊥ Samuel Pazicni,‡,⊥ Katherine A. Marvin,§,⊥ Daniel J. Stevens,∥ Katherine M. Paulsen,∥ and Judith N. Burstyn*,∥ †

Department Department § Department ∥ Department ‡

of of of of

Molecular Biosciences, Northwestern University, 2205 Tech Drive, Evanston, Illinois 60208, United States Chemistry, University of New Hampshire, 23 Academic Way, Durham, New Hampshire 03824, United States Chemistry, Hendrix College, 1600 Washington Avenue, Conway, Arkansas 72032, United States Chemistry, University of Wisconsin−Madison, 1101 University Avenue, Madison, Wisconsin 53706, United States 7.1. Heme-Thiolate CO Adducts Exhibit Distinctive Optical Spectra and rR Frequencies 8. Ferric/Ferrous Nitrosyl Adducts 8.1. Three Distinct Nitrosyl Adducts Are Observed in Heme-Thiolate Proteins 9. Functional Characteristics of Type-1 Heme-Thiolates 9.1. The Type-1 Heme-Thiolates Form HighValent Intermediates during Catalytic Cycles 9.1.1. Cytochrome P450 9.1.2. Chloroperoxidase 9.1.3. Nitric Oxide Synthase 9.1.4. Nitric Oxide Reductase 9.2. The Type-1 Heme-Thiolates Use a “ThiolatePush” Mechanism 9.3. Does the Thiolate Ligand Facilitate Multiple Reaction Pathways? 9.4. The Heme-Thiolate Affects the Basicity of Compound II 10. Functional Characteristics of the Type-2 HemeThiolates 10.1. Type-2 Heme-Thiolates Are (Frequently) Components of Signaling Pathways 10.1.1. CooA 10.1.2. NPAS2 10.1.3. RcoM 10.1.4. E75, DHR51, and Rev-erb 10.1.5. eIF2α Kinase 10.1.6. CBS 10.1.7. Nitrophorin 10.2. Type-2 Heme-Thiolates Destabilize Thiolate Coordination in the Ferrous State 10.3. Conversion of Type-1 Heme-Thiolates to Type-2 Heme-Thiolates Results in Loss of Activity and Unique Spectral Characteristics 11. Unclassified Heme-Thiolates 12. Conclusion Author Information Corresponding Author Author Contributions Notes Biographies

CONTENTS 1. Introduction 2. Classification of Heme-Thiolate Proteins 3. Spectroscopic Properties of Heme-Thiolate Proteins 4. Ferric Heme-Thiolates 4.1. Electronic Absorption Spectroscopy Distinguishes between Type-1 and Type-2 HemeThiolates 4.2. Magnetic Circular Dichroism (MCD) Spectra of Low-Spin Heme-Thiolates Exhibit Multiple Charge-Transfer Bands, the Positions of Which Vary between Type-1 and Type-2 Proteins 4.3. A Distinctive Fe−S Stretching Mode Is Observed in Resonance Raman (rR) spectra 4.4. Low- and High-Spin Fe(III) Heme-Thiolates Exhibit Unique Electron Paramagnetic Resonance (EPR) Spectra 5. Ferrous Heme-Thiolates 5.1. Low-Spin Fe(II) Heme-Thiolates Exhibit Coordination-Dependent Spectral Characteristics 5.2. High-Spin Fe(II) Heme-Thiolates Exhibit Atypical Optical and rR Spectra 6. Ferrous Dioxygen Adducts 6.1. The Existence of Two Distinct Dioxgen Adducts Is a Function of the OxyhemeThiolate Environment 6.2. Heme-Thiolate Compound I Is a Ferryl Species with a Porphyrin-Based Radical 6.3. Type-2 Heme-Thiolates, When Reduced, Reoxidize in the Presence of Dioxygen 7. Ferrous Carbonyl Adducts

© 2015 American Chemical Society

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Received: January 30, 2014 Published: March 12, 2015 2532

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when reduced. The high-spin, five-coordinate Fe(II) heme is functionally essential in small-molecule activation. Type-2 heme thiolates are always low-spin with two axial ligands; however, the thiolate ligand is replaced upon heme reduction. A second ligand replacement at the low-spin, six-coordinate Fe(II) heme enables function in small-molecule sensing. The differences in spin and coordination states between type-1 and type-2 heme thiolates give rise to their distinctive spectroscopic properties, as described in sections 4−8, and correlate with their functions in smallmolecule activation or sensing, as described in sections 9−11. Because the spectroscopic signatures arise from variations in electronic structure at the heme cofactor, it follows that each type of Cys(thiolate)-ligated hemoprotein displays distinct reactivity. This difference in reactivity is then manifested in the two classes of thiolate-ligated hemoproteins: type-1 that retains its Cys(thiolate) ligand throughout changes in the heme iron oxidation state and type-2 with a propensity to lose its Cys(thiolate) upon reduction of the heme iron (Figure 2). It is the differences in coordination behavior that give rise to the distinct spectroscopic signatures that allow their classification. Importantly, these two types of reactivity allow heme-thiolate proteins to play disparate roles in nature: as catalysts (type-1) or small-molecule sensors/ transporters (type-2). The purpose of this Review is to identify, separate, and classify these two types of heme-thiolate proteins on the basis of their distinct spectroscopic and functional properties. Because the classification system is based on spectroscopic attributes, and because the function is dependent on the distinctive reactivity arising from the heme electronic structure, spectroscopic and reactivity differences are presented before functional differences. This Review draws on nearly six decades of spectroscopic and functional data from well-studied and well-reviewed hemethiolate monooxygenases, but its new classification system is inspired by the emergence of new families of thiolate-ligated hemoproteins that function in numerous signaling pathways. Amazingly, these divergent and independent functions are possible despite the fact that each protein uses the exact same cofactor (heme b) and a common thiolate ligand in the firstcoordination sphere of the heme iron. Such diversity is made possible by utilizing the protein scaffold to modulate how the heme cofactor changes spin and coordination states upon substrate binding, gas binding, and/or changes in the redox state of the heme iron (Figure 2). This Review attempts to explain how nature utilizes the versatility of heme-thiolate proteins to play multiple indispensible roles in life processes.

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1. INTRODUCTION Heme proteins are among the most versatile players in the biological milieu; their functions range widely and include electron transfer, catalysis, and small-molecule sensing and transport.1 This broad functional diversity may be attributed to the protein environment that constitutes the heme cofactor’s binding site, particularly those amino acid residues that serve as axial ligands to the heme iron atom. For example, cytochromes, globins, heme oxygenases, and the majority of peroxidases employ the imidazole moiety of a histidine (His) residue as an axial ligand. Certain cytochromes are also coordinated by the thioether of methionine (Met) or, rarely, the polypeptide Nterminal amino group.2 Catalases utilize a tyrosine (Tyr) phenolate.3 An ever-growing class of heme proteins employs a cysteine (Cys) thiolate as an axial ligand to heme b (ironprotoporphyrin IX, Figure 1). Differences in solvent exposure,

Figure 1. (A) Iron protoporphyrin IX (heme b) consists of one iron atom coordinated by the tetradentate protoporphyrin IX dianion. (B) Two additional ligands may coordinate the heme iron, at positions above (X) and below (Y) the plane of the porphyrin macrocycle (represented as a parallelogram with nitrogen atoms at the corners).

2. CLASSIFICATION OF HEME-THIOLATE PROTEINS On the basis of their distinctive spectroscopic and functional attributes, we propose that thiolate-ligated heme proteins may be separated into two groups: type-1 and type-2 (Table 1). The type-1 heme-thiolates are centers of biological reactions in which small molecules, for example, O2, H2O2, or NO, are activated. The type-1 heme centers are catalytically competent only in a five-coordinate, high-spin state with an available vacant (or labile) sixth iron coordination site. The distinguishing reactivity characteristic of the type-1 centers is a cysteine(thiolate) ligand that is stable to ligand exchange upon reduction (nonlabile). Thus, the cysteine(thiolate) residue is retained during the catalytic cycle as well as upon binding exogenous ligands (Figure 2A). In contrast, the type-2 heme-thiolates are centers of small-molecule sensing or transport. The type-2 cysteine(thiolate) ligand is labile, which is fundamental to its function. Upon reduction of the type-2 heme-thiolates, the

hydrophobicity, iron atom oxidation and spin state, and the identity of the iron atom axial ligand(s) give rise to a wide variety of spectroscopic characteristics. These spectroscopic signatures provide useful fingerprinting handles with which to classify Cys(thiolate)-ligated hemoproteins. Two distinct classes of heme thiolate proteins emerge, which cluster according to their biological functions. We call these two classes the type-1 and type-2 heme thiolates. Herein, we demonstrate how type-1 and type-2 heme thiolate proteins differ in coordination numbers, spin states, and propensities to undergo ligand switching when reduced. Type1 heme thiolates have a vacant or labile axial coordination site trans to the thiolate ligand, and they retain the thiolate ligand 2533

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Figure 2. Substrate-, redox-, and effector-dependent coordination changes in the type-1 and type-2 heme-thiolates. (A) In the resting state of the type-1 heme-thiolates, the Fe(III) heme is ligated by a cysteine(thiolate) ligand and (commonly) a water ligand. The cysteine(thiolate) ligand is retained during substrate (sub) binding and heme iron reduction; during any gas-binding event, the gas molecule (CO in this example) binds trans to the cysteine(thiolate) ligand. (B) In the resting state of the type-2 heme-thiolates, the Fe(III) heme is ligated by a cysteine(thiolate) ligand and another neutral donor ligand (L), commonly a His residue. Heme reduction is accompanied by a ligand-switching event in which the cysteine(thiolate) is replaced by another neutral donor (L′); during any gas-binding event, the gas molecule (CO in this example) binds trans to a neutral donor ligand that is not the cysteine(thiolate) ligand.

Table 1. Coordination States and Functions of Heme-Thiolate Proteins type-1 heme-thiolate chloroperoxidase (C. fumago) cytochromes P450 nitric oxide reductase nitric oxide synthase

Fe(III) Cys(S−) Cys(S−)/H2O Cys(S−)/H2O Cys(S−)

Fe(II) Cys(S−) Cys(S−) Cys(S−) Cys(S−)

Fe(II)−CO Cys(S−)/CO Cys(S−)/CO Cys(S−)/CO Cys(S−)/CO

heme function peroxidation oxygen activation NO activation oxygen activation

type-2 heme-thiolate CooA (R. rubrum) human cystathionine β-synthase heme-regulated eIF2α kinase nitrophorin (C. lectularius) NPAS2 RcoM-1,-2 (B. xenovorans) E75 (D. melanogaster) Rev-erbβ (H. sapiens) DHR51 (D. melanogaster)

Fe(III) Cys(S−)/Pro(N) Cys(S−)/His Cys(S−)/His Cys(S−) Cys(S−)/His Cys(S−)/His Cys(S−)/His Cys(S−)/His Cys(S−)/His

Fe(II) His/Pro(N) Cys(S−)/Hisa His/His NRb His/His Met/His His/His His/His Ld/His

Fe(II)−CO His/CO His/CO His/CO Cys(SH)/COc His/CO His/CO His/CO His/CO Ld/CO

heme function CO sensing unknown NO sensing NO storage/transport CO sensing CO sensing CO and NO sensing unknown unknown

unclassified DGCR8

Fe(III) Cys(S−)/Cyse

Fe(II) Lysf/Ld

Fe(II)−CO Ld/CO

heme function unknown

a

Fe(II) CBS was found to retain its His/Cys coordination only upon reduction at low temperatures. bNot reported. cAn X-ray crystal structure of the C. lectulatius nitrophorin revealed a cysteine residue opposite CO. However, the CO adduct of this protein exhibits a Soret band at 420 nm. Therefore, we tentatively assign coordination as a cysteine(thiol). dUnknown. eExact protonation state unknown. fPutative.

transition-metal model complexes, have contributed significantly to an understanding of the characteristics that define the two proposed categories of heme-thiolate proteins. First identified in the extensive cytochrome P450 family,4 type-1 heme-thiolate coordination has now been characterized in chloroperoxidase (CPO) from Caldariomyces fumago,5 the fungal nitric oxide reductases (NORs),6 and the nitric oxide synthases (NOSs).7 The type-2 heme-thiolates include CooA from Rhodospirillum rubrum,8 human cystathionine β-synthase (hCBS),9 hemeregulated eukaryotic initiation factor 2α (eIF2α) kinase,10 neuronal PAS domain protein 2 (NPAS2),11 the Cimex lectularius nitrophorin (cNP),12 the regulators of CO metabolism

cysteine(thiolate) ligand may become protonated and is replaced either by a protein-derived ligand (termed a “ligand switch”) or by an exogenous ligand. Type-2 heme-thiolates are generally sixcoordinate and low-spin, and the cysteine(thiolate) is bound to the iron opposite a neutral donor, most often a histidine imidazole, in the Fe(III) state. Because of the propensity for type2 heme-thiolates to undergo a ligand switch upon reduction, the low spin, six-coordinate state is preserved when the heme is converted to the ferrous state (Figure 2B). X-ray crystallographic, spectroscopic, and biochemical studies of native proteins, complemented by computational investigations, studies of protein-based heme-ligand adducts, and 2534

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(RcoM-1 and RcoM-2) from Burkholderia xenovorans,13 the nuclear receptors E75 from Drosophila melanogaster,14 human Rev-erbα and Rev-erbβ,14b and DHR51 from Drosophila melanogaster.15 This Review outlines the distinctive spectroscopic and functional attributes of the two types of heme-thiolate protein and draws upon historical knowledge and recent advances in heme-thiolate chemistry to gain insight into how the catalytic and small-molecule sensing/transport functions are supported and distinguished by the heme-thiolate moiety.

3. SPECTROSCOPIC PROPERTIES OF HEME-THIOLATE PROTEINS Although cysteine(thiolate) coordination is a unifying feature of heme-thiolate proteins, type-1 and type-2 heme centers exhibit distinctly different spectral features either in the same oxidation states or in the presence of exogenous ligands. The type-1 and type-2 heme-thiolates are excellent examples of how the protein environment of a heme cofactor influences both the function and the spectroscopic characteristics of the heme. The unique spectroscopic signatures of heme-thiolate ligation were first observed over 50 years ago in studies of a CO-binding pigment from liver microsomes that contained iron protoporphyrin IX and exhibited an “anomalous” red-shifted electronic absorption band at 450 nm.16 Stern and Peisach later proposed sulfur coordination of this cytochrome “P450” CO adduct based on their model compound studies.17 Despite compelling spectroscopic evidence, including X-ray absorption data,18 the identity of the axial ligands remained in question until the X-ray crystal structure of soluble camphor-hydroxylating cytochrome P450 from Pseudonomas putida (cytochrome P450CAM) confirmed the presence of a thiolate-bound heme cofactor.19 The spectroscopic “fingerprints” that have been used to identify essentially all other heme-thiolate proteins originate from studies on cytochrome P450, its derivatives, and related model complexes. Sections 4−8 summarize the lines of spectroscopic evidence that distinguish the type-1 and type-2 heme-thiolates.

Figure 3. Electronic absorption spectrum of Fe(III) BxRcoM-2, a typical low-spin type-2 heme-thiolate. Fe(III) RcoM-2 displays a wellresolved δ band (354 nm), a sharp Soret (γ) band (423 nm), a broad α−β absorption envelope (500−600 nm), and two low-energy visible LMCT transitions (640 nm, 730 nm). Reprinted with permission from ref 13. Copyright 2008 American Chemical Society.

coordinate Fe(III) heme-thiolates is a pair of broad, low-energy (and low intensity) visible absorption LMCT bands (S(p) → Fe(III)), which are observed at approximately 650 and 750 nm.23 The five-coordinate, high-spin Fe(III) state with a sole cysteine(thiolate) heme ligand is accessed only by the type-1 heme-thiolates and exhibits spectral features distinct from those of the type-2 heme-thiolates. The resting states of isolated CPO and NOR are predominantly high-spin while the six-coordinate, low-spin resting states of P450 and NOS are converted to the high-spin state by the binding of substrate.5b,21,24 In contrast to low-spin heme-thiolate spectra, high-spin Fe(III) heme-thiolate absorption spectra are dominated by a broadened, blue-shifted Soret band at 390−400 nm that obscures the strong S(p) → Fe(III) LMCT transition. The visible bands are observed between 500 and 600 nm, as well as a characteristic porphyrin → Fe(III) LMCT transition at approximately 650 nm.25

4. FERRIC HEME-THIOLATES

4.2. Magnetic Circular Dichroism (MCD) Spectra of Low-Spin Heme-Thiolates Exhibit Multiple Charge-Transfer Bands, the Positions of Which Vary between Type-1 and Type-2 Proteins

4.1. Electronic Absorption Spectroscopy Distinguishes between Type-1 and Type-2 Heme-Thiolates

Electronic absorption spectroscopy provides a straightforward and useful means to identify the difference between type-1 and type-2 Fe(III) heme-thiolates. The visible and near-ultraviolet regions (300−800 nm) of heme absorption spectra are dominated by porphyrin (π → π*) electronic transitions.20 In type-2 heme-thiolates, the six-coordinate, low-spin Fe(III) state is common; the P450 cytochromes and NOSs also exhibit sixcoordinate, low-spin Fe(III) resting states, in which the sixth axial position is coordinated by water at physiological pH.7a,21 A classical Fe(III) low-spin heme absorption spectrum consists of (in order of decreasing energy) the δ (or “n”) band, the Soret (γ or “B”) band, and the α and β (visible or “Q”) bands (Figure 3).20 The position of the low-spin heme-thiolate Fe(III) Soret band typically ranges from 415 to 430 nm, while the α−β bands appear at 535−575 nm and may be resolved (typical for H2O or the Nterminal amino group trans to the thiolate) or consist of a broad absorption envelope (typical for histidine trans to thiolate). An intense, well-resolved δ band with a strong oscillator strength consistently appears at 355−365 nm; this band may be attributed to mixing of thiolate character into the porphyrin π → π* transition and serves as a hallmark for low-spin Fe(III) Cys(thiolate) ligation.22 A second distinctive feature of six-

Magnetic circular dichroism (MCD) spectroscopy is a powerful probe of spin state, oxidation state, and axial ligand identity that readily discriminates between type-1 and type-2 heme-thiolates. This technique has been used successfully to compare synthetic heme model complexes to heme proteins of unknown axial ligation, particularly in studies of the P450 cytochromes.26 Whereas the electronic absorption transitions of many heme proteins are broad and nondescript, the circularly polarized MCD transitions are frequently oppositely signed; this increase in fine structure gives superior fingerprinting capability to MCD spectroscopy.27 In most Fe(III) heme proteins that are sixcoordinate and low-spin, a characteristic temperature-dependent (C-term) peak/crossover/trough pattern in the Soret region dominates the MCD spectrum at He(l) temperatures. The peak/ crossover/trough position, which is typically red-shifted in type-2 six-coordinate, low-spin Fe(III) heme-thiolate proteins, is used as a marker for Cys coordination (Table 2, Figure 4).28 In contrast, type-1 high-spin Fe(III) heme-thiolates display an asymmetric MCD spectral pattern that spans the visible and UV regions with an intense negative feature between 390 and 400 nm (Table 2, Figure 4).28a,d,29 2535

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Table 2. Positions of MCD Soret (γ) Peak/Crossover/Trough (nm) and LMCT Transitions (nm), Spin States, and Temperatures of Data Collection for Selected Heme-Thiolate Proteins spin state HSb LSd HS HS

T (K) RTc NRe NR 277

ref 5b 30 30 24a

675a

spin state LS LS HS LS LS LS

T (K) εβ). The Fe(II) type-2 heme-thiolates (except Fe(II) hCBS; vide infra) display these spectral characteristics, due to their propensity to have the Cys(thiolate) ligand replaced by a neutral donor ligand upon reduction (Figure 8). The type-1 Fe(II) heme-thiolates

Figure 9. Electronic absorption spectra of Fe(II) cytochrome P450 with various exogenous ligands. “Hyperporphyrin” type-1 Fe(II) hemethiolate electronic absorption spectra (top panel) are generated upon the addition of π-acceptor ligands such as CO and nitrosoalkanes, whereas “normal” type-1 Fe(II) heme-thiolate electronic absorption spectra (bottom panel, solid) are generated upon addition of pure σdonor ligands such as metyrapone. Fe(II) cytochrome b5 (bis His ligation; bottom panel, dashed) is included for comparison. Reprinted with permission from ref 61. Copyright 1983 ASBMB. Figure 8. Thermally induced ligand switching occurs at the Fe(II) heme in CBS. Upon reduction of the heme in CBS, the Cys(thiolate) ligand is retained, as evidenced by the red-shifted Soret band at 450 nm (solid line). Upon heat treatment, a ligand-switching event occurs in which the Fe(II) Cys(thiolate) ligand is replaced by an unknown neutral donor ligand; the final species has a Soret band at 424 nm (dashed line). Reprinted with permission from ref 93b. Copyright 2005 American Chemical Society.

features arise only in low-spin heme-thiolates with a sixth strong π-acceptor ligand, for example, CO or nitrosoalkanes, which enables orbital mixing of the Soret and charge transfer transitions between the iron and axial ligand π orbitals.63 In contrast, “regular” low-spin Fe(II) heme-thiolate centers are observed with pure σ-donors, for example, dimethylsulfide, n-phenylimidazole, and metyrapone.61 MCD spectroscopy distinguishes among low-spin hyperporphyrin Fe(II) type-1, regular Fe(II) type-1, and type-2 hemethiolates. Hyperporphyrin MCD spectra are dominated by transitions in the Soret region. A strong negative δ band and an intense, derivative-shaped temperature-independent A-term in the Soret region are hallmarks of hyperporphyrin Fe(II) low-spin type-1 heme-thiolates (S = 0). In contrast, the most prominent feature of regular low-spin Fe(II) type-1 and type-2 hemethiolate MCD spectra is an intense A-term (“α-band”; dominant at T > 77 K, but easily visible in the absence of B- and C-terms at T < 77 K) in the visible region (Figure 10). The α-band feature is present in most low-spin Fe(II) heme MCD spectra, and its position is indicative of the identity of the axial heme ligands. An α-band crossover position between 562 and 576 nm is typical for low-spin Fe(II) type-1 heme-thiolates; an α-band position between 550 and 560 nm is typical for low-spin Fe(II) type-2

retain their thiolate ligand upon reduction and are usually encountered in the five-coordinate, high-spin (S = 2) state. The low-spin (S = 0) Fe(II) type-1 heme-thiolates are usually accessed by adding exogenous ligands, such as carbon monoxide,56 nitric oxide,57 nitrosoalkanes,58 thiols and thioethers,59 and amines.46a,60 The various low-spin ligand adducts of cytochrome P450CAM universally exhibit an extremely red-shifted Soret absorption maximum due to retention of the thiolate ligand. The ∼450 nm Soret, first observed in CO-bound Fe(II)P450 cytochromes, is now a hallmark diagnostic of lowspin, Fe(II) type-1 heme-thiolate centers. Elegant absorption and MCD characterization of Fe(II) P450CAM ligand adducts by Dawson and co-workers61 divide the low-spin Fe(II) heme-thiolates into two groups: (1) hyperporphyrins,20,62 which exhibit a prominent extra absorption band in the 320 < λ < 400 nm region (called the “split Soret” or “blue 2539

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Figure 10. (A) Electronic absorption spectra of Fe(III) (thin line) and Fe(II) (thick line) of BxRcoM-2. (B) MCD spectrum of Fe(II) BxRcoM-2 at 4 K and 7 T. RcoM-2 exhibits prototypical behavior of a type-2 heme-thiolate in which Cys(thiolate) ligation is lost upon heme reduction; the Cys(thiolate) ligand is replaced by another neutral donor (Met104), maintaining a six-coordinate, low-spin heme iron center. Reprinted with permission from ref 13. Copyright 2008 American Chemical Society. Figure 11. MCD spectra of five-coordinate Fe(II) high-spin thiolateligated heme complexes. Fe(II) P450-LM2 (top, solid), Fe(II) protoheme-thiolate model complex in dimethylacetamide (top, dotted), and Fe(II) CPO (bottom, solid) all display similar MCD spectra. Reprinted with permission from ref 95. Copyright 1987 American Chemical Society.

heme-thiolate proteins because the thiolate is no longer coordinated to the Fe(III) heme.61 5.2. High-Spin Fe(II) Heme-Thiolates Exhibit Atypical Optical and rR Spectra

A high-spin, thiolate-coordinated Fe(II) state is only accessed by the type-1 heme-thiolates, which present spectral features distinct from both low-spin type-1 and type-2 heme-thiolate proteins. Biologically, this spin and ligation state is most relevant to the catalytic cycle of the P450 (and P450-like) enzymes. Fivecoordinate, high-spin Fe(II) heme-thiolates typically exhibit Soret and unresolved visible absorption maxima at 410 and 540 nm, respectively.30 High-spin Fe(II) heme-thiolates (S = 2) are also characterized by a temperature-dependent MCD C-term in the Soret region (Figure 11), which dominates the Soret region of the spectrum at T < 77 K. The red-shifted Soret region MCD term bears a strong negative band at 420 nm with a plateau of positive intensity and a weak negative feature at 570 nm in the visible region.64 However, an equilibrium mixture of high and low spin states is commonly encountered in the MCD spectra. Resonance Raman spectroscopy reveals further disparities between the type-1 high-spin and type-1 and type-2 low-spin heme-thiolate centers. The oxidation state marker bands (ν4) of Fe(II) P450 cytochromes,39b,65 CPO,39b,66 and NOS67 occur at unusually low energies as compared to other Fe(II) high-spin heme centers. High-spin Fe(II) heme-thiolates exhibit ν4 in the range of 1341−1348 cm−1; the analogous band for high-spin

Fe(II) heme centers with a lone histidine ligand as well as lowspin Fe(II) heme centers with two neutral ligands occurs between 1355 and 1360 cm−1. Similarly, the spin and coordination state marker band ν3 consistently appears near 1466 cm−1 for high-spin Fe(II) heme-thiolates; ν3 occurs at 1470−1475 cm−1 for histidine-coordinated high-spin Fe(II) heme proteins and at 1485−1490 cm−1 for low-spin Fe(II) heme proteins with two neutral donor ligands. The anomalously low frequencies seen for thiolate-ligated heme proteins are attributed to the strong π-donor influence of the cysteine(thiolate) ligand, which is sensed by the Raman-active porphyrin π* orbitals via back-donation from the dπ orbitals of the Fe(II) atom.68 Because a cysteine(thiolate) is typically replaced by a neutral donor in the Fe(II) type-2 systems, their resonance Raman spectra are indistinguishable from other Fe(II) heme systems that bear two neutral donors. 2540

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6. FERROUS DIOXYGEN ADDUCTS 6.1. The Existence of Two Distinct Dioxgen Adducts Is a Function of the Oxyheme-Thiolate Environment

The most relevant reaction of the type-1 heme-thiolate oxygenases involves interaction with dioxygen at the Fe(II) high-spin heme center to form a reversible complex with triplet O2. This diamagnetic, low-spin adduct is generally described as a Fe(II)−dioxygen (Fe(II)−O2) complex, a Fe(III)−superoxo (Fe(III)−O2•−) complex, or a resonance hybrid of the two.69 Ishimura et al. reported the first dioxygen complex of cytochrome P450CAM, which exhibits an absorbance spectrum with two prominent maxima at 355 and 418 nm.70 While the absorption features are superficially similar to those of oxymyoglobin,71 the MCD spectrum of oxy-cytochrome P450CAM is distinct from that of oxymyoglobin, suggesting a different electronic structure.72 The νO−O stretch observed (1140 cm−1) in the rR spectrum of oxy-cytochrome P450CAM73 is of nearly identical frequency (1140 cm−1) to that of the oxy-thiolate-bound “picket fence” porphyrin complex74 and an elaborate cytochrome P450 model (1137 cm−1) containing a hydrogen-bonded dioxygen ligand.75 The relatively low νO−O of oxyheme-thiolate centers suggests that there is a weaker O−O bond than in oxyhemoglobin, where a multiplet of bands is detected at higher average energy.76 This decrease in νO−O for heme-thiolates relative to imidazole-ligated heme centers likely originates from greater π back-bonding from the more electron-rich thiolate-ligated iron into the π* orbitals of oxygen, weakening the O−O bond and facilitating its cleavage.27h Alternatively, the presence of the electron-rich thiolate ligand may preferentially stabilize the Fe(III) oxidation state over the Fe(II) oxidation state, thus facilitating formation of the Fe(III) superoxo adduct. The variability in the spectral features of oxy-heme-thiolates, including the sensitivity of the Soret band position to solvent polarity, has been well-documented for synthetic model complexes.77 Furthermore, several crystal structures exist that reveal differences in hydrogen bonding at the Cys(thiolate) ligand among the P450s and CPOs; these secondary interactions may differentially attenuate the donor capability of the thiolate ion and thus modify the electronics at the oxy-heme center.5a,78 This difference may be one way of controlling the reactivity of P450 and CPO toward O2. Support for this argument comes from electronic absorption and MCD studies on thiolate-ligated forms of the H93G Mb mutant, which recapitulate trends seen in the oxy-ferrous form of putidaredoxin-reduced P450CAM.79 These observations likely explain why the absorption and MCD spectra of oxy-CPO are distinct from those of oxycytochrome P450CAM despite the extensive spectroscopic similarities between other adducts of the P450 cytochromes and CPO.80 Interestingly, the transitory oxo-adducts of the NOSs, which have been observed with stopped-flow techniques, are two distinct temperature-dependent species termed hemeoxy I and heme-oxy II, which parallel the respective dioxygen adducts of CPO and cytochrome P450.81

Figure 12. Fe(IV) or “ferryl” species attained in the catalytic cycles of the type-1 heme-thiolates. (A) The ferryl porphyrin π-cation radical complex known as Compound I. (B) The ferryl complex known as Compound II, which is one electron reduced from Compound I.

above the Fe(III) resting state, one equivalent being associated with a ferryl center [Fe(IV)O]2+ and the other with the porphyrin-based radical that is delocalized onto the Cys(thiolate) ligand. Compound 1 acts as a two-electron oxidant in cytochrome P450 and a one-electron oxidant in the peroxidase mode of CPO. Compound II, produced by one-electron reduction of Compound I, is inherent to the peroxidase cycle of CPO and may be best described as a protonated Fe(IV) hydroxide species (vide infra).83 Compound I of CPO (CPO-I) is the most fully characterized heme-thiolate Compound I and has been spectroscopically characterized by electronic absorption,84 resonance Raman,84a,85 EPR and Mössbauer,86 as well as X-ray absorption spectroscopies.87 The CPO-I absorption spectrum exhibits an extremely blue-shifted asymmetric Soret band (364 nm) and a distinct feature in the visible region (689 nm), which are comparable to peak positions calculated by DFT for a model complex.88 The EPR properties (geff 1.61, 1.72, 2.00) are interpreted as arising from the lowest-energy Kramers doublet of a strongly antiferromagnetically coupled S = 1 ferryl and a spin S′ = 1/2 porphyrin radical.86,89 Egawa et al. identified a ν4 band for CPO-I that is downshifted and weak as compared to other species in the CPO catalytic cycle,85b which is compatible with ν4 observed in model systems upon porphyrin π cation radical formation.85a,90 The presence of a ferryl core was verified by X-ray absorption spectroscopy.87 Recent studies have identified and characterized the spectroscopic and kinetic properties of Compound I generated from reaction of m-chloroperbenzoic acid with a P450 from the thermophilic bacterium Sulfolobus acidocaladarius (CYP119I).89,91 Despite being superficially similar, CYP119-I is substantially more reactive than CPO-I and is capable of reacting with unactivated C−H bonds, whereas CPO-I will not; it is suggested that this difference is attributable to electronics, not substrate access.89,92 Indeed, while electronic absorption (Figure 13) and Mössbauer spectroscopies suggest a similarity between CPO-I and CYP119-I, their EPR spectra are different. The fitted g values reported for CYP119-I are geff 1.86, 1.96, and 2.00, which are distinct from those of CPO-I; furthermore, fitted values of |J|/ D, a measure of the ratio of the antiferromagnetic coupling to that of the axial ZFS, suggest that there is stronger coupling for CYP119-I.89 This stronger coupling is thought to arise from the shorter Fe−S bond in CYP119-I as compared to CPO-I.89

6.2. Heme-Thiolate Compound I Is a Ferryl Species with a Porphyrin-Based Radical

6.3. Type-2 Heme-Thiolates, When Reduced, Reoxidize in the Presence of Dioxygen

Reactive oxygen intermediates are responsible for the biological transformations inherent to the type-1 heme-thiolate oxygenases. By analogy to intermediates formed in other heme-containing oxidases, the putative heme-thiolate-based oxidants are thought to be an Fe(IV)−oxo porphyrin radical cation species (Compound I) and an Fe(IV)−oxo porphyrin species (Compound II) (Figure 12).82 Compound I is doubly oxidized

In contrast to the type-1 heme-thiolates, which form dioxygen adducts as part of their catalytic cycle, the type-2 heme-thiolates react with dioxygen by an outer-sphere redox mechanism, that is, via autoxidation. To date, no oxygenated adducts of type-2 hemethiolates have been identified. Oxygen reactivity has been 2541

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Figure 13. Electronic absorption spectrum of Sulfolobus acidocaladarius cytochrome P450 (CYP119) Compound 1 (dashed) generated by mixing equal molar amounts of Fe(III) CYP119 and m-chloroperbenzoic acid (m-CPBA). Reprinted with permission from ref 89. Copyright 2010 AAAS Publishers.

investigated for RrCooA,8a BxRcoM-2,13 eIF2α kinase,10 E75 and Rev-erbβ,14b and hCBS.93 Thiolate coordination is retained in Fe(II)hCBS at high pH (Figure 8), and exposure of the protein to air results in oxidation to Fe(III)CBS without changes in heme iron coordination.93 The redox-mediated ligand switch of RrCooA is reversed when exposed to air.8a A reasonable explanation for this behavior is that type-2 Fe(II) heme-thiolates exist as low-spin, spin singlet species (S=0), and the reaction of triplet (S = 1) O2 with singlet heme iron to yield a singlet oxyheme adduct is a spin-forbidden process. In contrast, the highspin, quintet (S = 2) ground state of Fe(II) type-1 heme-thiolates binds O2 quite readily (vide supra). DFT calculations have suggested that little activation enthalpy is necessary for the associated spin-crossover transition that occurs upon reaction of the high-spin Fe(II) type-1 heme-thiolate center with O2, thus facilitating oxygen binding.94 It may be that low-lying quintet states are inaccessible for the low-spin Fe(II) type-2 hemethiolates due to the strong ligand field, thereby eliminating any potential reactivity with O2 beyond the observed autoxidation chemistry.

symmetric, derivative-shaped temperature-independent A-terms in the Soret and visible regions.13,14b,31,61,95 Resonance Raman spectroscopy further distinguishes the CO adducts derived from type-1 and type-2 heme-thiolates. HemeCO adducts exhibit two characteristic stretching modes attributed to Fe−CO (νFe−CO) and C−O (νC−O) vibrations in the 460−540 and 1900−2000 cm−1 regions, respectively. An inverse linear correlation exists between the frequencies of the νC−O and νFe−CO stretching modes of heme proteins and model compounds (Figure 14).96 This correlation arises from the

7. FERROUS CARBONYL ADDUCTS 7.1. Heme-Thiolate CO Adducts Exhibit Distinctive Optical Spectra and rR Frequencies

Figure 14. Back-bonding correlation diagram for Fe(II)−CO adducts derived from type-1 and type-2 heme-thiolates. CO adducts of type-1 heme-thiolates, which retain the cysteine(thiolate) ligand, obey the νC−O/νFe−CO correlation indicated by the dashed (- - - - -) line. CO adducts of type-2 heme-thiolates, in which CO is bound opposite a neutral donor, obey the νC−O/νFe−CO correlation indicated by the solid (−) line. Figure adapted from ref 10.

Although only relevant to the functions of certain type-2 hemethiolates, the differences in the spectroscopic attributes of carbon monoxide (CO) adducts of the type-1 and type-2 heme-thiolates provide further evidence for the distinctiveness of the two classes. Now a common probe of heme centers, CO(g) was first reacted with the Fe(II) heme-thiolate cytochrome P450 to reveal the anomalous Soret absorption maximum at ∼450 nm; this strongly red-shifted Soret maximum is attributed to CO binding trans to cysteine(thiolate) in type-1 heme-thiolates.62 Because the type-2 cysteine(thiolate) is replaced either by a redox-mediated ligand switch or by the CO molecule itself, the type-2 heme-thiolate CO adducts display a Soret band at 420 nm and equivalent α−β peaks at approximately 560 and 540 nm, respectively. In further contrast to type-1 heme-thiolate CO adducts (vide supra, section 5.1), MCD spectra for the type-2 CO adducts are dominated by

degree of back-bonding of the Fe(II) heme atom to the CO molecule, the extent of which is sensitive to the ligand (or lack thereof) trans to CO. Strong donor ligands, such as cysteine(thiolate), cause two different phenomena: (1) an inherently weaker Fe−C bond, due to stronger polarization of the dz2 orbital toward the thiolate, and (2) a strong inverse correlation due to π back-bonding from the electron-rich Fe atom to the π* CO molecular orbital.97 Because of the retention of the cysteine(thiolate) ligand, the frequencies of νC−O/νFe−CO for type-1 2542

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Figure 15. Characteristic X-band EPR spectra of several ferrous nitrosyl adducts. The EPR spectrum of the ferrous 14NO adduct of horseradish peroxidase (left panel, top) displays a characteristic 9-line g2 splitting pattern indicative of a His ligand bound trans to the NO molecule, whereas the EPR spectrum of the ferrous 14NO chloroperoxidase (left panel, bottom) displays only a 3-line g2 splitting pattern, consistent with retention of the Cys(thiolate) ligand at the ferrous heme of chloroperoxidase. Reprinted with permission from ref 105a. Copyright 1975 American Chemical Society. The EPR spectrum of the ferrous 14NO adduct of RrCooA (right panel, top) displays a characteristic 3-line g3 splitting pattern that collapses to a 2-line splitting pattern upon 15NO substitution (right panel, bottom), indicative of a ferrous five-coordinate nitrosyl adduct without endogenous ligation. Reprinted with permission from ref 49. Copyright 2000 American Chemical Society.

with loss of both endogenous ligands. Thus, reaction of NO with both Fe(III) and Fe(II) CooA yields the five-coordinate {FeNO}7 species.49 Both hCBS101 and eIF2α kinase10 form five-coordinate NO adducts, although reaction with NO has only been investigated in the Fe(II) state of these two enzymes. Reaction of Fe(II) BxRcoM-213 and HsRev-erbβ14b with NO results in six-coordinate adducts, while reaction of Fe(II) DmE7514b with NO results in a five-coordinate NO adduct. Similar to globin proteins with NO bound opposite a neutral donor ligand, the six-coordinate NO adducts exhibit a sharp Soret band in the 415−425 nm range with a broad α−β absorption envelope centered at about 560 nm.71 EPR spectroscopy, which is capable of detecting nuclearelectronic spin coupling of a coordinated NO• radical with a trans ligand, readily distinguishes between the five- and six-coordinate {FeNO}6 and {FeNO}7 type-1 and type-2 heme-thiolate adducts. In Fe(III) type-1 heme-thiolate {FeNO}6 species, the binding of NO forces a spin-state crossover; the only unpaired electron on the Fe(III) atom then couples to the unpaired electron on the bound NO, resulting in a spin-paired, EPR-silent species.102 The electronic structure of {FeNO}6 may be best described as Fe(II)−NO+;103 this complex is isoelectronic with that of the diamagnetic Fe(II)−CO complex and may be formed in the presence of a variety of anionic axial ligands.104 In contrast to the diamagnetic {FeNO}6 species, the paramagnetic type-1 heme-thiolate {FeNO}7 adducts exhibit a rhombic EPR signal whose central resonance (g2) displays a well-resolved triplet hyperfine structure due to coupling of the unpaired electron of NO to the nuclear spin of 14N (I = 1) (Figure 15, left panel, bottom).26c,105 The analogous type-2 {FeNO}7 adducts typically display classic 9-line rhombic EPR spectra in which g2 is split into hyperfine and superhyperfine signals that arise from coupling of the NO electron with the nuclear spin of the trans nitrogen atom of the coordinated residue (usually His) (Figure 15, left panel, top).54d,105 The five-coordinate {FeNO}7 adducts that are observed in the heme-thiolates exhibit only axially symmetric 3-line EPR spectra (Figure 15, right panel).105b,106

heme-thiolate CO adducts correlate on a distinct line from those of type-2 heme-thiolate CO adducts: the latter correlating with Fe−CO adducts bearing neutral donors trans to CO. Thus, type1 and type-2 heme-thiolates can be distinguished by which νC−O/ νFe−CO correlation their respective CO-adducts obey.

8. FERRIC/FERROUS NITROSYL ADDUCTS 8.1. Three Distinct Nitrosyl Adducts Are Observed in Heme-Thiolate Proteins

The type-1 and type-2 heme-thiolates exhibit quite different reactivity and spectral features when coordinated by nitric oxide (NO•). Unlike other exogenous heme ligands, such as CO and O2, NO is able to bind to both Fe(III) and Fe(II) heme centers. These species have been designated in the nomenclature of Feltham and Enemark as {FeNO}6 and {FeNO}7, respectively.98 Addition of NO to Fe(III) type-1 heme-thiolates, regardless of heme iron spin state, results in a six-coordinate thiolate-ligated {FeNO}6 species characterized by absorption maxima at approximately 430−440 nm (Soret), 540 nm (β), and 570 nm (α).80,99 A six-coordinate {FeNO}7 type-1 heme-thiolate adduct is formed by addition of NO to the Fe(II) protein, and is characterized by a hyperporphyrin absorption spectrum with a split Soret band (370, 440 nm) and a broad visible absorption at approximately 560 nm.80,99 Under certain conditions, such as denaturation, nitrosyl adducts of type-1 heme-thiolates convert to a distinct five-coordinate {FeNO}7 species, in which both protein-derived axial ligands have been lost.57a The type-2 heme-thiolates often form five-coordinate {FeNO}7 adducts; this species may be formed due to the strong σ trans effect of NO polarizing the dz2 orbital and weakening the opposing ligand.100 Electronic absorption spectra of the fivecoordinate {FeNO}7 adducts are characterized by broad, lowintensity Soret bands at 400 nm and visible bands at 540 nm (β) and 570 nm (α). However, some recently characterized type-2 heme-thiolates have exhibited a six-coordinate {FeNO}7 species. In all published accounts involving addition of NO to Fe(III) type-2 heme-thiolates, autoreduction of the heme center occurs 2543

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Significant differences among the five- and six-coordinate {FeNO}6 and {FeNO}7 adducts in type-1 and type-2 hemethiolates are also observed in their corresponding rR spectra. Type-1 heme-thiolate {FeNO}6 adducts exhibit vibrational Fe− NO (νFe−NO) and N−O (νN−O) stretching frequencies in the 520−540 and 1800−1880 cm−1 ranges, respectively.104a,107 While the 560 cm−1 band of the six-coordinate thiolate-bound {FeNO}7 adduct (previously identified as νFe−NO) is only modestly elevated from that of the {FeNO}6 species, the νN−O frequency in the type-1 {FeNO}7 adducts downshifts considerably to about 1600 cm−1 as compared to that of the {FeNO}6 species.107c The isotope-sensitive bands of the type-2 sixcoordinate {FeNO}7 adducts with a trans neutral ligand are observed at frequencies in the 560−570 cm−1 range and in the 1630−640 cm−1 range.13,14b,49,108 Type-2 rR spectra of fivecoordinate {FeNO}7 systems typically show isotope-sensitive modes in the 520−525 and 1660−1680 cm −1 ranges, respectively.10,49 Importantly, detailed studies utilizing nuclear resonance vibrational spectroscopy (NRVS) conclusively identified νFe−NO in the {FeNO}7 six-coordinate adducts of model compounds as well as various hemoproteins in the range of 440−450 cm−1, while the Fe−N−O bending frequency (δFe−N−O) was identified in the range 520−590 cm−1;109 one prior study made the same band assignments using isotopeedited rR spectroscopy.110 While NRVS has not been performed on any of the type-2 {FeNO}7 systems, in light of these data, it seems likely that the previously identified νFe−NO frequencies may actually correspond to the isotope-sensitive δFe−N−O mode. Whether all hemoproteins that form six-coordinate {FeNO}7 adducts (including the type-1 and type-2 heme-thiolates) also follow a roughly linear inverse correlation of νFe−NO/νN−O similar to the Fe(II)CO νFe−CO/νC−O correlation is controversial.108a,109a,111

Figure 16. Catalytic cycle of the P450 cytochromes. Proposed structures of putative intermediates, including Compound I, are indicated in brackets. The substrate is represented by RH, and the product is represented by R(O)H. Figure adapted from ref 95.

9. FUNCTIONAL CHARACTERISTICS OF TYPE-1 HEME-THIOLATES

state expels a heme-bound water (at physiological pH) molecule to generate the five-coordinate high-spin Fe(III) state. This spinstate transition causes a significant positive increase in the redox potential of the heme iron, which is then reduced by electron transfer from NADH or NADPH via a redox partner protein. To activate O2, the P450 cytochromes utilize the high-spin Fe(II) state of the heme-thiolate. The rate-limiting addition of a second electron to the oxy−Fe(II) complex is proposed to initiate a cascade, yielding the putative ferryl porphyrin π-cation radical species (Compound I) that serves as the reactive oxygenating species. Compound I of cytochrome P450 is predicted to undergo oxygen-atom transfer by an “oxygen rebound” mechanism, in which hydrogen atom abstraction from the substrate by Compound I is followed by radical recombination to form the hydroxylated product.114 9.1.2. Chloroperoxidase. Chloroperoxidase (CPO) from the marine fungus Caldariomyces fumago is a versatile type-1 heme-thiolate enzyme that possesses typical peroxidase (eq 2) and catalase (eq 3) functions. In addition, CPO has the unique ability to catalyze halogenation (chloride, bromide, and iodide) of organic substrates using H2O2 as the oxidant (eq 4)115 and catalyzes dehalogenation of certain halophenols.116 CPO is regarded as a “hybrid” of the cytochromes P450 and heme peroxidases, as it contains a polar, peroxidase-like heme pocket housing a thiolate-ligated cytochrome P450-like heme center.5a

9.1. The Type-1 Heme-Thiolates Form High-Valent Intermediates during Catalytic Cycles

As catalytic centers for biological reactions, the type-1 hemethiolate centers generally activate small molecules for substrate transformations.112 This chemistry is accomplished via reaction of O2 or H2O2 directly with the heme-thiolate center to form highly reactive oxo adducts, [(P)Fe(IV)O)] species known generally as “ferryls” (Figure 12). The heme-thiolate ferryl species are very similar in nature to those employed by the histidine-ligated oxygenases, as demonstrated by the peroxidase function of CPO. An obligatory requirement for small-molecule activation is the availability of an open or labile coordination site at the heme iron, which is a defining characteristic in the classification of type-1 heme-thiolate proteins. 9.1.1. Cytochrome P450. The cytochromes P450 are a ubiquitous family of type-1 heme-thiolate enzymes that perform a variety of reactions including the activation of carbon centers for catabolism, steroid metabolism, and detoxification of xenobiotics.113 P450-catalyzed monooxygenase reactions (eq 1) are performed with a remarkable degree of stereo- and regioselectivity. XH + *O2 + AH 2 → X(*O)H + H 2*O + A

(1)

The catalytic cycle of the P450 cytochromes is presumed to involve a high-valent iron-oxo intermediate (Figure 16).1c Substrate binding to a nearby cavity in the low-spin resting 2544

2RH + H 2O2 → R−R + 2H 2O

(2)

2H 2O2 → O2 + 2H 2O

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an unoxidized porphyrin macrocycle. Compound II is competent to oxidize an additional equivalent of substrate. 9.1.3. Nitric Oxide Synthase. The nitric oxide synthases (NOSs) catalyze the oxidation of L-arginine into L-citrulline and NO using a high-valent oxo-intermediate equivalent to other Compounds I (Figure 12). Three mammalian isoforms of NOS have been identified: (1) endothelial (eNOS), (2) inducible (iNOS, expressed in stimulated macrophages), and (3) neuronal (nNOS, from brain).117 All isoforms have essentially identical crystallographic structures and chemistry.118 The homodimeric NOSs contain a reductase and an N-terminal catalytic oxygenase domain (NOSoxy) in each subunit. Each NOSoxy domain binds a type-1 heme-thiolate moiety and (6R)-5,6,7,8-tetrahydro-Lbiopterin (H4B), which is adjacent to the heme and provides reducing equivalents.7a,118a,119 Electrons from NADPH are provided by the flanking C-terminal reductase domains, which also bind flavin mononucleotide (FMN) and flavin adenine dinucleotide (FAD). The substrate, L-arginine, binds directly above the heme-thiolate center, and oxidation of L-arginine to Lcitrulline and NO occurs via two consecutive oxygenase reactions.120 Insertion of oxygen into one of the terminal guanidine N−H bonds of L-arginine, using an activated ferryl porphyrin π-cation radical species, produces Nω-hydroxy-Larginine (NOHA, Figure 18) with conversion of the heme back to the Fe(III) state. In the second step, a hemehydroperoxide adduct acts as the reactive oxygen species, initiating a nucleophilic attack on the guanidino carbon to form a heme-bound peroxy-substrate that breaks down to form Lcitrulline (Figure 18) and NO. The nature of the second step of this reaction (the breakdown of Nω-hydroxy- L-arginine via a ferric-peroxo species rather than a ferryl species) has been probed via pH, metalloporphyrin substitution, site-directed mutagenesis, and product distribution of iNOS,121 as well as EPR and ENDOR spectroscopies of cryoreduced NOS from Geobacillus stearothermophilus (gsNOS).122 9.1.4. Nitric Oxide Reductase. The nitric oxide reductase (NOR) from Fusarium oxysporum was the first isolated hemethiolate NOR and remains the most thoroughly characterized NOR.123 Genomic analyses show that NOR enzymes occur frequently in fungi and are employed in denitrification processes.124 The NORs are unique among P450-like enzymes due to their ability to catalyze the reduction of NO to N2O according to eq 5; they are the only known type-1 heme-thiolates that function outside the realm of oxygen activation.125

(4)

The enzymatic cycles of CPO (Figure 17) involve both transient Compound I and II intermediates.95 High-spin Fe(III)

Figure 17. Catalytic cycles of chloroperoxidase. Proposed intermediates are bracketed. Reaction of the high-spin resting form of CPO (blue) with H2O2 generates the highly reactive Compound I (green). From CPO-I, the peroxidase function (top) converts two molecules of substrate (AH) to radical products; halogenation (bottom) is performed; or the catalase function (middle) catalyzes the disproportionation of H2O2. Figure adapted from ref 95.

CPO reacts with H2O2 to form the ferryl porphyrin π-cation radical Compound I (eqs 1 −3). In the peroxidase reaction cycle, Compound I serves as a one-electron oxidant that converts organic substrates (AH) into radical products (A•) with reduction of Compound I to Compound II, a ferryl heme with

Figure 18. Two-step reaction catalyzed by the nitric oxide synthases. In the first step, L-arginine is hydroxylated to form Nω-hydroxy-L-arginine (NOHA), utilizing one molecule of O2 and two electrons from NADPH. In the second step, NOHA is converted to L-citrulline and nitric oxide, using another molecule of O2 and one electron from NADPH. 2545

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Figure 19. Proposed reaction mechanism for the catalytic cycle of the reduction of NO by the fungal NORs. The resonance structures describe the delocalization of electron density over the Fe−N−O unit and thus more realistic electron configurations for the Fe(III)−NO complex and the intermediate I. Figure adapted from ref 6.

2NO + NADH + H+ → N2O + NAD+ + H 2O

H bond cleavage.129 In 2002, Ogliaro and co-workers used density functional theory (DFT) calculations that placed the “thiolate push” in a transition state at about 29 kcal/mol relative to the ground state.130 A more recent comparative computational study by Kamachi et al. further elucidated how the anionic thiolate ligand influences reactivity; their calculations suggested that strong donor−acceptor interactions occur between the axial ligand and the Fe−O bond, and the strong electron donation from the thiolate ligand is essential in the transition state to weaken and activate the Fe−O bond.131 Recent work by Green and co-workers has demonstrated the ability of the thiolate ligand to influence the pKa of the ferryl species, thus creating an extremely potent and basic oxidant that facilitates H atom abstraction.83c,132 The protein environment, via hydrogen-bonding interactions with the cysteine(thiolate) ligand, also influences the reactivity of the resulting intermediate species. This effect has been elegantly demonstrated by work on P450CAM and NOS variants that alter the hydrogen-bonding capabilities at the thiolate ligand, both in vitro and in silico.133 While a definitive consensus on the role of secondary interactions at the type-1 heme-thiolate Cys residue is still not achieved, several groups have suggested that hydrogen bonding at the Cys(thiolate) ligand modulates the electron density at the iron center by controlling the electron-donating capability of the sulfur atom, thus attenuating the thiolate “push” effect. This regulation of the “push” effect alters the rate of ferryl formation,133c the overall rate of reaction,133f and even enzymatic coupling.133a,e These results illuminate the intricacy by which nature employs second-sphere interactions at the Cys(thiolate) moiety to regulate catalysis by fine-tuning electron density at the heme iron atom.

(5)

The electrons required for NO reduction by the fungal NORs are directly transferred from NADH. A 1.0 Å X-ray crystal structure of FoNOR revealed a solvent-exposed distal heme pocket, which is consistent with the much more negative reduction potential than that observed for the P450 cytochromes.125,126 The chemistry of FoNOR has been nicely reviewed by Daiber et al.,6 and evidence for the catalytic mechanism, involving the Fe(II)NOR−HNO intermediate (I) after hydride transfer from NADH, has been documented (Figure 19).107b,125a,127 9.2. The Type-1 Heme-Thiolates Use a “Thiolate-Push” Mechanism

Although the nature of the highly reactive species inherent to the chemistry of type-1 heme-thiolates is a subject of ongoing investigation, a long-standing hypothesis (the “thiolate-push” formalism) concerning the ability of the heme-thiolate moiety to stabilize these intermediates remains widely accepted. This formalism, first proposed by Dawson in 1976,26a suggests that the cysteine(thiolate) ligand serves as a strong internal electron donor that facilitates O−O bond cleavage by pushing electron density onto the trans position via the heme iron atom as a conduit, weakening the O−O bond and stabilizing the resulting high-valent oxo-ferryl species. Early comparative experiments on P450 and myoglobin revealed that ligands bound trans to histidine in myoglobin had greater acidity (a measure of electron deficiency) than those trans to the cysteine(thiolate) in cytochrome P450.26c,128 Ohno et al. investigated model systems that demonstrated enhanced reactivity promoted by thiolate coordination to iron, which stabilizes the transition state during hydroxylation and lowers the activation energy for homolytic C− 2546

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Figure 20. Proposed structures of Compounds I and II based on EXAFS and DFT calculations that predict bond lengths and a protonated rebound intermediate (Compound II). Reprinted with permission from ref 138. Copyright 2006 American Chemical Society.

9.3. Does the Thiolate Ligand Facilitate Multiple Reaction Pathways?

reduction potential of Compound I and the pKa of Compound II.132b,138 The ability of cysteine(thiolate) to promote hydrogen abstraction in this situation could be essential. The π-donating ability of the thiolate ligand imparts a partial negative charge upon the oxo unit to facilitate protonation of the oxygen atom. This proposal assumes that the rebound mechanism is operative in P450 hydroxylations and that basic ferryls are a general and unique feature of cysteine(thiolate)-coordinated hemes.

The existence of type-1 heme-thiolate complexes in nonmonooxygenase reactions catalyzed by enzymes such as CPO and NOR suggests an additional role(s) of the thiolate ligand in catalysis. On the basis of studies of the P450 cytochromes, CPO, NOR, and related enzymes in combination with DFT calculations, Ullrich suggested that the thiolate may perform different functions in the pure monooxygenases versus nonmonoxygenases.134 In this proposal, the monooxygenase enzymes utilize the thiolate ligand’s electron density to restrict π donation from oxygen to the iron, which maintains the “active oxygen” complex in the transition state. The ferryl (Compound I, FeIVO) ground state exists with an electron hole delocalized about the porphyrin and the thiolate (thiyl). As part of the differentiated function utilized in nonmonoxygenase-like reactions, Compound II acts as a very rapid electron acceptor to assist in the facile oxidation of the substrate radical and subsequent reaction with the poised nucleophile. In this proposal, the “oxygen rebound” is abandoned as a final step in the reaction mechanism, and Compound II is an atypical ferryl in which the thiolate interaction may also be invoked to assist in electron transfer.

10. FUNCTIONAL CHARACTERISTICS OF THE TYPE-2 HEME-THIOLATES 10.1. Type-2 Heme-Thiolates Are (Frequently) Components of Signaling Pathways

The labile nature of the cysteine(thiolate) ligand in type-2 hemethiolates makes these heme centers ideally suited for functions in small-molecule sensing and transport. A role for the type-2 hemethiolates in redox sensing stems from the almost universal replacement of the cysteine(thiolate) ligand upon reduction of the Fe(III) heme-thiolate complex. An increase in the flexibility and exposure to solvent of the heme pocket is observed upon reduction of the Fe(III) heme-thiolate.140 Such flexibility allows the repositioning of another protein residue as a sixth ligand in the Fe(II) heme complex and results in local movement of the protein structure. In the type-2 heme-thiolates, gas binding requires replacement of at least one of the heme protein ligands; similar to the ferric/ferrous ligand-switch, gas-induced ligand replacement may result in additional changes within the protein structure, as gas binding alters intraprotein interactions and forms new ligand−protein contacts. In the type-2 heme-thiolates, mounting evidence links these local changes in heme coordination to global conformational and dynamical changes, especially of the protein functional domains.112,141 Thus, it is the lability of the heme ligands and the flexibility of the type-2 heme thiolate heme pocket that enable these proteins to function allosterically as small-molecule sensors. 10.1.1. CooA. The CO oxidation activator (CooA) from the purple photosynthetic bacterium Rhodospirillum rubrum represents the paradigm for gas-sensing type-2 heme-thiolate proteins.142 The coo regulon contains two operons, regulated by and encoding CooA, whose products are required for R. rubrum to anaerobically oxidize CO to CO2.143 Like other members of the cAMP receptor protein (CRP) and fumarate and nitrate reductase (FNR) superfamily of transcriptional regulators, RrCooA is arranged into distinct regulatory and DNAbinding domains. In the Fe(III) state, the regulatory domain heme is coordinated by Cys75 and the N-terminal Pro2 of the opposing monomer; reduction replaces Cys75 with His77, while CO binding replaces Pro2 on the opposing side (Figure 21).8,31,144 Coordination of NO results in a five-coordinate

9.4. The Heme-Thiolate Affects the Basicity of Compound II

Recent work has probed the unique protonation state of Compound II in the type-1 systems.132b,135 Type-1 hemethiolate Compounds II are different from the analogous Compounds II in histidine-ligated oxygenases.84a,136 Chloroperoxidase Compound II (CPO-II) was the only example of a highvalent heme peroxidase intermediate for which a νFe−O stretching frequency had not yet been reported, and this absence was originally attributed to an abnormally weak iron(IV)−oxo bond.84a,85b X-ray absorption measurements of CPO-II by Green et al. yielded an Fe−O bond length of 1.82 Å, a value much longer than typical ferryl iron−oxo distances (1.65 Å), suggesting that the oxo moiety is protonated and is best described as an FeIV−hydroxide species (pKa ≥ 8.2) (Figure 20).83a Green et al. combined DFT and Mössbauer spectroscopy to detect a FeIV−hydroxide species in CPO-II,137 and Stone et al. reported two species in CPO-II that were assigned as FeIV−oxo and FeIV−OH.138 The νFe−O for CPO-II was then identified at 565 cm−1, which is consistent with the presence of an FeIV−OH species. 83b Using DFT and Mö ssbauer spectroscopy of cytochrome P450, Behan et al. have also detected an FeIV−OH species for P450-II at physiological pH.132b Several studies by Mayer and co-workers have established that the strength of the O−H bond formed during H atom abstraction scales with the ability of the metal−oxo species to abstract hydrogen,139 and that O−H bond formation is controlled by the one-electron 2547

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Figure 21. Heme coordination states of Fe(III), Fe(II), and Fe(II)CO RrCooA. The cysteine(thiolate) ligand is replaced by a histidine upon reduction of the Fe(III) heme-thiolate. CO binding then replaces the distal proline ligand to yield the type-2 CO adduct. Similar behavior is hypothesized for the other type-2 heme-thiolates such as hCBS, eIF2α kinase, NPAS2, the BxRcoMs, and Drosophila E75.

NO adduct, which is inactive for DNA binding.49 Complementing compelling spectroscopic evidence, a 2.6 Å X-ray crystal structure of Fe(II)CooA by Lanzilotta et al. verified that Cys75 is replaced by His77 via a redox-mediated ligand switch.145 Biochemical and spectroscopic studies demonstrated that COspecific ligand replacement initiates the global conformational change that activates RrCooA to bind DNA (Figure 22).142b,146

coordinates to the Fe(III) PAS-A heme, and Cys170 is replaced by His171 upon reduction.11,151 Recent studies on a longer construct with PAS-A and bHLH domains (bHLH-PAS-A) have suggested that the Fe(III) ligation scheme is an equilibrium between Cys/His and His/His ligation; this equilibrium is speculated to aid in signal transduction in NPAS2.152 rR data suggest that CO binds opposite a neutral nitrogen donor at the Fe(II) bHLH-PAS-A heme, as is characteristic of type-2 hemethiolates;150 formation of a similar CO adduct inhibits heterodimerization of the full length NPAS2 with BMAL1.148 Binding of NO to the Fe(II) bHLH-PAS-A heme results in a fivecoordinate nitrosyl adduct; however, no functional characteristics of this complex have been reported.150 The spectral features exhibited by the PAS-B domain alone (i.e., the PAS-B domain expressed without the remainder of the protein) are also consistent with thiolate coordination in the Fe(III) state, but not in the Fe(II) state, suggesting the presence of a second type-2 heme-thiolate in NPAS2.151 Although CO is a likely candidate for the native signal of NPAS2, putative sensing of other small molecules has not been conclusively ruled out. 10.1.3. RcoM. The aerobic PCB-degrading bacterium Burkholderia xenovorans has been shown to express two transcription factors named the regulators of CO metabolism, BxRcoM-1 and BxRcoM-2, which are type-2 heme-thiolate proteins.13,153 Aerobic CO-dependent transcriptional activation in vivo has been observed.153 The BxRcoM proteins couple an Nterminal PAS-fold heme domain to a C-terminal LytTR DNAbinding domain, of which little is known. Structural modeling and mutagenic analyses implicate His74 and Met104 as heme iron ligands in the Fe(II) state. Recent spectroscopic data for BxRcoM-2 revealed that, similar to RrCooA, the Fe(III) state is low-spin, coordinated by a cysteine(thiolate) (Cys94) and a sixth neutral ligand, likely His74.13,153,154 In RcoM-2, a redox-mediated ligand switch occurs in which the Fe(III) heme ligand Cys94 is replaced with a neutral donor ligand, likely Met104, in the Fe(II) state. Binding of CO and NO in BxRcoM-2 occurs via replacement of one of the axial ligands to the Fe(II) heme by the gas molecule; resonance Raman and EPR spectral features are indicative of a histidine trans to both the CO and the NO ligands, consistent with replacement of the more weakly coordinated Met104 (Figure 23).13 10.1.4. E75, DHR51, and Rev-erb. The nuclear receptors (NRs) Drosophila melanogaster E75, Drosophila melanogaster DHR51, and Homo sapiens Rev-erbβ are transcriptional silencers that are now known to possess type-2 heme-thiolate centers.14,15 The NRs comprise a superfamily of eukaryotic transcription factors that coordinate inter- and intracellular signaling processes

Figure 22. Representation of the putative conformational changes induced by reduction and CO binding in RrCooA, which activates the protein for DNA binding. The inactive, “resting state” Fe(III)CooA structure is unknown. The inactive, “ready off” Fe(II)CooA structure (left) was modeled from PDB file 1FT9. The active, “Fe(II)CO CooA” structure is represented by cAMP-bound CRP (right), a close relative of CooA, and was modeled from PDB file 1I5Z.

The allosteric signal from the effector (CO)-binding domain to the DNA-binding domain is thought to be transmitted via two αhelices at the dimer interface of the protein that form a leucinezipper motif.147 10.1.2. NPAS2. The transcription factor neuronal PAS domain protein 2 (NPAS2) was the first CO-responsive type-2 heme-thiolate protein to be identified in mammals.148 NPAS2 forms a heterodimeric complex with BMAL1 to activate the expression of per and cry genes, which are negative regulatory components of the circadian clock. NPAS2 is a member of the basic helix−loop−helix (bHLH)-PAS family of transcription activators; the N-terminal bHLH DNA binding domain is coupled to two PAS domains, termed PAS-A and PAS-B, which bind heme.148,149 Formation of a complex between the NPAS2/ BMAL1 heterodimer and DNA is directly coupled to the levels of heme present; a study of a protein fragment bearing the PAS-A and bHLH domains (bHLH-PAS-A) found that specific binding to the E-box DNA sequence occurs only in the presence of heme.150 Independent characterization of a construct bearing only the PAS-A domain (PAS-A) revealed that Cys 170 2548

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nuclear receptor has been deemed adventitious,161 which emphasizes that caution must be taken to avoid overinterpretation of in vitro studies on isolated, fragmentary domains. 10.1.5. eIF2α Kinase. The heme-regulated eIF2α kinase (HRI) is a multidomain heme protein with two distinct hemebinding sites, one of which is a type-2 heme-thiolate that functions as an NO sensor.162 HRI is a member of a family of kinases that regulate initiation of protein synthesis in eukaryotic cells.163 The eIF2α kinases catalyze phosphorylation of eIF2α, which then binds eIF2β and inhibits the guanine nucleotide exchange activity required for protein synthesis. HRI specifically regulates globin synthesis in reticulocytes in response to available heme levels.164 Several environmental and chemical stimuli, including NO binding to heme, activate HRI.162a Metal cations, including Hg2+, Fe2+, Cu2+, Zn2+, and Pb2+, have been reported to inhibit HRI function due to competitive binding of the Cys(thiolate) heme ligand; the strong inhibition by Hg2+ (and only Hg2+) is fully reversed by NO.162c,163b,165 The hemebinding domains of HRI consist of a kinase insertion domain, which binds heme reversibly and regulates HRI activity in response to heme concentration, and an N-terminal domain, which binds a type-2 heme-thiolate complex.162c,166 Site-directed mutagenesis on the HRI N-terminal heme domain (HRI-NTD) suggested coordination of the heme by two histidine residues.167 Subsequent examination of full-length HRI by absorption, EPR, and rR spectroscopies revealed a thiolate-bound heme center.10 Interrogation via mutagenesis and spectroscopic studies on the full-length HR implicated a cysteine(thiolate) of the kinase insertion domain and a histidine of the N-terminal domain as axial ligands.168 Upon reduction, the spectral characteristics of cysteine(thiolate) coordination are lost, suggesting that either a ligand switch or protonation to cysteine(thiol) occurs.169 NO binds to the type-2 heme-thiolate center with loss of both endogenous ligands and a concomitant upregulation of kinase activity. Studies have also demonstrated that the activation of HRI kinase activity is specific for NO, as CO suppresses activity in HRI.162a,170 These findings suggested that the N-terminal heme domain adopts a conformation that inhibits HRI kinase activity; NO binding may induce the correct conformational change to release heme domain inhibition of HRI kinase activity.162a 10.1.6. CBS. Human cystathionine β-synthase (hCBS) is a pyridoxal 5′-phosphate (PLP)-dependent enzyme that also binds a type-2 heme-thiolate cofactor of unknown function. Human CBS catalyzes the condensation of serine and homocysteine to yield cystathionine via a β-replacement reaction performed by the PLP cofactor. A member of the fold type-II family of PLPdependent enzymes, hCBS is unique in that it also bears a heme, which is noncovalently bound within a N-terminal hydrophobic pocket.9b,171 The thiolate of Cys52 and the Nε2 atom of His65, both located on the N-terminal loop, axially coordinate the Fe(III) heme (Figure 24). This N-terminal heme-binding region in hCBS is absent in CBS isolated from Saccharomyces cerevisiae and Trypanosoma cruzi, suggesting that heme is an evolutionarily recent acquisition.172 Studies have demonstrated that the hCBS heme serves no direct catalytic role; the β-replacement reaction catalyzed by CBS follows a typical PLP-based mechanism, and the heme-free enzyme is catalytically competent, albeit with lowered activity, and heme-domain deletion variants have substantially decreased activity.173 Furthermore, CBS expressed in heme-free systems results in inactive enzyme; activity may be rescued by expression of CBS in the presence of chemical

Figure 23. Proposed coordination states of Fe(III), Fe(II), and Fe(II)CO BxRcoM-2. The cysteine(thiolate) ligand is replaced by a methionine upon reduction of the Fe(III) heme-thiolate. CO binding then replaces the proximal methionine ligand to yield the type-2 CO adduct. Reprinted with permission from ref 154. Copyright 2012 Springer.

and regulate development, growth, and reproduction, cell proliferation and death, and overall homeostasis.155 NRs are modular proteins with a highly conserved DNA binding domain (DBD) coupled to a ligand-binding domain (LBD) via a flexible hinge region.156 NRs may function as transcriptional activators or repressors, and binding of a small, lipophilic ligand within the LBD typically acts as a signaling event for these functions. E75, Rev-erbα, and Rev-erbβ are NR homologues that belong to the NR1D subfamily of transcriptional silencers, which regulate growth and development, energy metabolism, and the circadian clock through complicated feedback mechanisms.157 In 2005, Reinking et al. reported that Drosophila E75 bound a redox- and gas-sensitive heme cofactor within the LBD, which also regulated proper folding and association with its heterodimeric partner protein DHR3;158 they tentatively identified two conserved histidines (His574 and His364) as potential heme ligands. Subsequently, De Rosny and co-workers showed that the Fe(III) heme iron was coordinated by histidine and cysteine(thiolate) using EPR.14a The human Rev-erbα and Rev-erbβ NRs were reported to bind heme reversibly, which mediated corepressor binding. Spectroscopic and structural characterization of Reverbβ revealed a six-coordinate heme environment with cysteine(thiolate) (Cys 384 ) bound opposite a histidine residue (His568).14b,159 Recent reports suggest that a thiol−disulfide redox switch may be one means by which Rev-erbβ may control the binding of ferric heme.160 Under reducing conditions, spectroscopic data for E75 and Rev-erbβ indicate that the Fe(III) thiolate ligand is replaced by a neutral donor ligand.14b,160 Reinking et al. demonstrated that binding of CO and NO, forming six- and five-coordinate heme complexes, respectively, results in inhibition of E75 repressor activity.158 Current published reports for similar redox- or CO- and NO-binding dependence for the Rev-erb NRs are inconclusive.158−160 De Rosny and co-workers also identified the Drosophila homologue of the human photoreceptor and cell-specific nuclear receptor DHR51 as a heme-thiolate protein. Using electronic absorption and EPR data, they showed that the DHR51-LBD binds heme in a six-coordinate fashion and utilizes a Cys(thiolate)/neutral donor (likely His) motif, nearly identical to E75; furthermore, they demonstrated that DHR51 binds CO and NO to form sixand five-coordinate species, respectively, and speculate that DHR51 may be involved in gas sensing.15 Similar to NPAS2, spectroscopic and biochemical studies on these proteins have only been performed on fragmentary LBD constructs; thus, the functional implications of the observed heme coordination states are unknown. In at least one case, the binding of heme to a 2549

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characteristics of both type-1 and type-2 heme-thiolates. A 1.75 Å X-ray crystal structure of Fe(III) cNP shows a solvent-exposed heme pocket with a weakly associated water molecule trans to thiolate, although EPR spectroscopy revealed a predominantly high-spin, five-coordinate Fe(III)cNP species.12 Unlike the type1 heme-thiolates, the cNP heme is unable to form a stable complex with imidazole but does form stable NO and CO adducts.181 As in other type-1 heme-thiolates, NO binds opposite the thiolate moiety of Fe(III)cNP to form a sixcoordinate NO adduct. Consistent with its role as an NO transporter, however, cNP reversibly binds two molecules of NO. The second NO equivalent binds via heme-assisted S-nitrosylation of the thiolate (Cys60), generating an Fe(II) heme−NO and a cysteine(thiyl) radical. This S-nitrosylation occurs at higher NO concentrations, and whether this effect is physiologically relevant is unclear;182 however, higher local concentrations of NO may be experienced under certain circumstances in vivo. This novel mechanism of binding and transporting NO is consistent with the labile nature of cysteine(thiolate) in type-2 heme-thiolates. The cNP CO adduct exhibits a Soret maximum at 420 nm, although this result conflicts with a 1.45 Å X-ray crystal structure, which places CO opposite the cysteine ligand. Elegant work by Perera et al. has demonstrated that neutral cysteine(thiol) coordination to a heme center yields spectroscopic features very similar to those of neutral nitrogen coordination.169 A reasonable, if unverified, explanation for this apparent conflict may be that CO is bound opposite a neutral cysteine(thiol) in cNP. We have classified cNP as a type-2 hemethiolate due to greater apparent spectroscopic and functional similarities to this class of heme-thiolate proteins.

Figure 24. Location of key residues that interact with the heme and PLP in hCBS. Highlighted are the Fe(III) heme ligands Cys52 and His65; the cysteine(thiolate) hydrogen-bonding partners Arg266 and the amide backbone of Trp54; the PLP phosphate hydrogen-bonding partners Thr257 and Thr260; and Lys119 that forms the PLP internal aldimine. Structure modeled from PDB file 1JBQ. Reprinted with permission from ref 45. Copyright 2012 American Chemical Society.

chaperones or exogenous metalloporphyrins.174 In one example, expression of hCBS in the presence of CoCl2 resulted in the incorporation of Co(III)PPIX in place of heme. Not only did the Co(III)PPIX maintain the native type-2 thiolate ligation motif, but the enzyme was fully active; these results have suggested heme may play a structural role by stabilizing the N-terminus.175 Other studies have suggested that the hCBS heme behaves in a regulatory fashion as seen in other type-2 heme-thiolates;176 this proposal is very attractive, as it would connect the redox activity of CBS to supplies of Cys (produced in a downstream pathway), which is necessary for glutathione production. Binding of NO, CO, or cyanide, as well as thiolate-bond disruption by HgCl2 (in both Fe and Co hCBS), have a strong negative effect on hCBS activity, implying communication between the heme and active site.101,175b,177 Reduction of the CBS heme destabilizes thiolate coordination at near-physiological temperature and pH, ultimately resulting in a redox-mediated ligand switch similar to that of other type-2 heme-thiolates (Figure 8).93b However, at lower temperatures and pH, a thiolate-bound Fe(II) state forms transiently before reoxidizing to the native Fe(III) hemethiolate.32 Because hCBS activity was observed to decrease 2fold upon reduction of the heme, Taoka et al. proposed that the hCBS heme is a redox sensor.178 However, we have shown that formation of the ligand-switched species (CBS424, Figure 8) under assay conditions causes the observed decrease in activity; the enzyme is deactivated by conversion to CBS424, not by heme reduction alone.179 Recent results have suggested that replacement of the Cys(thiolate) ligand by CO, formed under physiological conditions, may function to control CBS activity;176c however, the lengthy amount of time (2 h) required to form the inactive PLP tautomer suggests that this mode of regulation is unlikely (at least as studied in vitro). A molecule or environment that is putatively sensed by the hCBS heme remains elusive, as does the physiological significance of the unusual redox behavior of the heme. 10.1.7. Nitrophorin. Cimex lectularius (the bedbug) makes use of a salivary heme-thiolate protein, termed a nitrophorin, to ensure a robust blood meal.12 Nitrophorins are a class of heme proteins from blood-sucking insects that bind and transport NO; dilution and pH changes inside the host lead to release of NO, which results in vasodilation.180 The nitrophorin of C. lectularius (cNP) is unique in that Cys60 serves as a ligand to the heme; all other nitrophorin hemes are coordinated by a histidine imidazole.12 The heme-thiolate of the cNP also exhibits

10.2. Type-2 Heme-Thiolates Destabilize Thiolate Coordination in the Ferrous State

Type-2 heme-thiolates generally exist as low-spin, six-coordinate species in the Fe(III) state, and the presence of the sixth ligand may labilize the cysteine(thiolate) upon reduction, a defining characteristic in the classification of the type-2 heme thiolates. The negative charge on the cysteine(thiolate) ligand, coupled with strong electron donation from the sixth ligand (most often a neutral nitrogen donor), are expected to stabilize the Fe(III) state of the heme. In addition, studies on Cys-ligated variants of yeast cytochrome c and microperoxidase-8 imply that entropic factors also stabilize the Cys(thiolate)-ligated Fe(III) heme.140 Upon reduction, the type-2 heme-thiolates may be intrinsically unstable unless the heme center compensates by decreasing the large amount of charge density donated by the cysteine(thiolate). This compensation may take the form of a weakened Fe−S interaction, and thus a labile cysteine(thiolate) ligand that is susceptible to replacement by a nearby residue if there is flexibility in the Fe(II) heme pocket.140 A similar propensity to lose thiolate coordination during heme reduction is seen in X-toCys variants of hemoproteins that naturally bear nitrogenous- or oxygen-containing axial ligands such as myoglobin or heme oxygenase.40,79a,169 The low-spin resting states of the type-1 P450 cytochromes and NORs need no such compensation when reduced, because the trans ligand (H2O) is a very weak donor in comparison to the neutral nitrogen donors trans to the type-2 cysteine(thiolate). Thus, the discriminating factor between type1 and type-2 heme-thiolates may be the effect of trans coordination to the heme-thiolate. This argument, however, does not include cNP, which contains a high-spin heme-thiolate that nonetheless becomes labile upon NO coordination/heme reduction, and which we therefore classify as type-2. The unusual 2550

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11. UNCLASSIFIED HEME-THIOLATES Recent spectral work in our lab has been used to identify a heretofore unseen native bis(cysteine) ligation motif. The homodimeric double-stranded RNA-binding protein Di George Critical Region 8 (DGCR8) (also called Pasha in worms and flies) is a part of the Microprocessor complex. DGCR8 acts together with its RNase partner Drosha. These two proteins are responsible for the first step in micro RNA (miRNA) processing: cleavage of primary miRNAs (pri-miRNAs) to form precursor miRNAs (pre-miRNAs).189 The absorbance spectrum of the purified, overexpressed DGCR8 displayed a hyperporphyrin spectrum with a red-shifted Soret at approximately 450 nm (Figure 25); this spectrum was originally attributed to either an

temperature- and pH-dependent effects seen in hCBS also deviate from expected type-2 behavior. These discrepancies may be rationalized by invoking second-sphere interactions. The type-1 cysteine(thiolate) ligands have been shown crystallographically to engage in a number of hydrogen-bonding interactions with surrounding residues.78b−d,183 These interactions stabilize the heme iron upon reduction by drawing thiolate electron density away from the heme iron, resulting in a nonlabile heme-thiolate moiety. Unfortunately, relatively few crystal structures of the type-2 heme-thiolates (hCBS, DmCBS, and cNP) exist with which to make comparisons.9b,12,171c While the cNP heme-thiolate engages in few hydrogen-bonding interactions, the hCBS heme-thiolate engages in several hydrogen bonds (Figure 24). This observation is not wholly inconsistent with the observed labile behavior of these two type-2 heme-thiolates, as hCBS undergoes a ligand switch at elevated temperatures. Furthermore, studies on the R266K variant of hCBS, in which one of the heme Cys(thiolate) hydrogenbonding partners has been altered, have shown that the thiolate ligand is more labile in the reduced form, and becomes more like a “classical” type-2 heme-thiolate by ligand-switching almost instantaneously at physiological temperature.45 Thus, in addition to differences in coordination number and resulting spin states that characterize the type-1 and type-2 heme thiolates, we propose that second-sphere hydrogen-bonding interactions and ligand trans effects modulate the functional and behavioral differences between and within these classes. 10.3. Conversion of Type-1 Heme-Thiolates to Type-2 Heme-Thiolates Results in Loss of Activity and Unique Spectral Characteristics

The type-1 heme-thiolates cytochrome P450 and CPO can be converted to inactive, type-2 heme-thiolate species. The cytochromes P450 undergo a universal transition to a stable but inactive species known as cytochrome P420. The term “P420” is derived from the 420 nm Soret absorption maximum exhibited by the ferrous−CO adduct of cytochrome P420. Numerous chemical and physical techniques,184 including increased temperature, increased pressures, the addition of salts, denaturants, or even organic solvents, convert cytochrome P450 to cytochrome P420.26b,95,185 Conversion to P420, a sixcoordinate, low-spin heme-thiolate, is also accompanied by an increase in cysteine(thiolate) lability, as CO replaces the thiolate ligand in Fe(II) cytochrome P420.185,186 Similarly, a change in CPO heme coordination at alkaline pH converts the CPO heme to a type-2 heme-thiolate, rendering the enzyme inactive. This species was termed C420 as it also forms a CO adduct with a Soret absorption maximum at 420 nm.186 Moreover, unlike native CPO, the heme-thiolate of C420 is susceptible to thiolspecific modifying agents, such as p-mercuribenzoate.187 The C420 Fe(III) heme appears to be coordinated by the native cysteine(thiolate) (Cys29) and an additional neutral donor. When reduced, C420 loses its cysteine(thiolate) ligand via replacement with a second neutral donor ligand. Thus, according to the classification scheme presented in this work, P420 and C420 are type-2 heme-thiolates. Interestingly, studies of NOS have suggested that its P420 form results from the protonation of the proximal thiolate to a neutral thiol, rather than ligand switching.188 Formation of low-spin, six-coordinate hemethiolate centers in CPO and cytochrome P450 explains the lack of catalytic activity, as an open coordination site at the heme no longer exists in the Fe(III) or Fe(II) forms to bind and activate peroxide or oxygen.

Figure 25. Electronic absorption spectrum of Fe(III) DGCR8. The Fe(III) heme in DGCR8, which is ligated by a unique bis(Cys) ligation motif, displays a highly split hyperporphyrin spectrum. Figure adapted from ref 33.

Fe(II)-thiolate or CO-Fe(II)-thiolate ligated heme.190 Using electronic absorption, MCD, rR, and EPR spectroscopies, we have shown that this unique spectrum arises from an Fe(III) heme b that is ligated by Cys352 from each DGCR8 monomer.33 While several forms of Fe(III) P450 and Fe(III) CPO are known to bind exogenous thiols and display strongly split hyperporphyrin spectra,48,95,128a,191 DGCR8 is the first known protein to bind heme using two endogenous Cys ligands. The presence of the hyperporphyrin spectrum suggests that both axial ligands are thiolate in nature, as the mixed thiol/thiolate state does not generate a hyperporphyrin spectrum.26c That the Fe(III) form of DGCR8 does not bear spectral similarities to native forms of Fe(III) type-1 or type-2 heme-thiolates places it in a category of its own; thus, DGCR8 may represent the first member of a type-3 class of heme-thiolates. Similar to type-2 heme-thiolates, Fe(II) DGCR8 loses thiolate-coordination upon reduction. The spectral characteristics of Fe(II) DGCR8 are most similar to Fe(II) heme proteins that have two neutral donors. Electronic absorption and MCD spectral changes that occur upon pH titration of the Fe(II) form of DGCR8 are inconsistent with retention of a protonated thiol as a ligand to the Fe(II) heme.192 Therefore, we propose that DGCR8 undergoes a dual ligand-switch in which both Cys352 ligands are replaced upon heme reduction. Intriguingly, DGCR8 loses its ability to support pri-miRNA processing once reduced to the Fe(II) state. Apo-DGCR8 exhibits lowered RNA binding 2551

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cooperativity; the functional activity may be fully rescued by reconstitution with Fe(III) heme (restoring the characteristic hyperporphyrin spectrum), but not by reconstitution with Fe(II) heme.192 Thus, it is clear that the Fe(III) DGCR8-heme complex is necessary for full activity; however, the exact nature of how Fe(III) heme supports pri-miRNA processing is unknown. Fe(II) DGCR8 will interact with CO and NO to form six- and five-coordinate species, respectively, but the implications for processing activity are unknown. Although only one protein has been identified that bears a bis(Cys) ligation motif at the heme iron center, the type-3 hemethiolates are likely functionally distinct from type-1 and type-2 heme-thiolates. In DGCR8, there is clearly no need to invoke a catalytic role for the heme macrocycle. While the functional aspects of gas-binding have yet to be explored in Fe(II) DGCR8, the slow (and likely unfavorable) reduction process may implicate that the Fe(II) state is physiologically irrelevant for DGCR8, ruling out possible regulation by CO or NO. We currently favor the theory that DGCR8 may use its unusual ligation motif in a specific manner to function as a heme sensor. 33,192 DGCR8 may be the first of many novel hemoproteins yet to be discovered in the type-3 category that use the specificity of a bis(Cys) ligation motif to respond to the presence of heme as a metabolic signal in vivo. Work from other laboratories has implicated the presence of several other thiolate-ligated heme proteins. Examples of these proteins include arginyl-tRNA synthetase193 and arginyl-tRNA transferase194 (both part of the “N-end rule” pathway), the nuclear retinoid X receptor α (RXRα),195 glutamyl-tRNA synthetase,196 glutamyl-tRNA reductase,197 and the hemesensing protein PpsR.198 Unfortunately, limited spectroscopic data exist for these proteins, prohibiting their classification. Further studies are necessary to group these proteins into their respective heme-thiolate classes.

speculate that this category may grow as more hemoproteins with this unconventional ligation motif are discovered.

AUTHOR INFORMATION Corresponding Author

*Tel.: (608) 262-0328. Fax: (608) 262-6143. E-mail: burstyn@ chem.wisc.edu. Author Contributions ⊥

These authors contributed equally.

Notes

The authors declare no competing financial interest. Biographies

Aaron T. Smith grew up in Springfield, IL, and received his B.A. in chemistry from Boston University in 2007. In 2012 he received his Ph.D. in chemistry from the University of Wisconsin−Madison under the supervision of professor Judith N. Burstyn. In her laboratory, he studied the role of cysteine(thiolate) ligation in metalloporphyrin-containing proteins involved in such diverse functions as transcriptional regulation, sulfur amino acid metabolism, and microRNA maturation. Aaron is currently a postdoctoral fellow in the laboratory of professor Amy C. Rosenzweig at Northwestern University. There he is investigating active transition metal transport in a superfamily of membrane-bound ATPases.

12. CONCLUSION This Review has presented a new classification scheme for cysteine(thiolate)-coordinated heme proteins. Nearly all known heme-thiolate centers can be described as having a nonlabile or a labile thiolate ligand, type-1 and type-2, respectively, according to the nomenclature introduced in this work. The type-1 hemethiolates are catalytic centers that engage in diverse reactions with common themes; a defining characteristic that allows these proteins to function catalytically is the presence of an open or labile coordination site on the heme iron opposite a stably coordinated cysteine(thiolate). The electron-donating ability of the cysteine(thiolate) ligand is fundamental in the formation and stabilization of very reactive, high-valent catalytic intermediates in these systems. The type-2 heme-thiolates are generally smallmolecule sensors and transporters; defining characteristics that allow these proteins to function as sensors are a coordinatively saturated heme with a labile cysteine(thiolate) and a flexible heme pocket that allows facile ligand switching. The labile nature of the cysteine(thiolate) is exploited by the sensor heme to ensure that a signal cascade is transmitted from the heme to the region of activity. Thus, two distinct chemical characteristics of the heme-thiolate, excellent electron-donating ability and lability in the reduced state, are distinguished and employed independently of each other in type-1 and type-2 heme-thiolate centers. This Review has also postulated the existence of a type-3 heme-thiolate center in which an unusual bis(Cys) ligation motif is utilized to recognize heme with high degree of specificity; we

Samuel Pazicni grew up in Grindstone, PA, and received a B.A. with majors in chemistry and music from Washington and Jefferson College in 2001. In 2006 he received his Ph.D. in chemistry from the University of Wisconsin−Madison under the supervision of Professor Judith N. Burstyn. In her laboratory, Sam investigated the role of heme in human cystathionine β-synthase. From 2006−2009, he completed a postdoctoral appointment with Profs. James Penner-Hahn and Brian Coppola at the University of Michigan where he utilized X-ray 2552

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absorbance spectroscopy to interrogate biological metal centers and performed chemistry education research. Sam is currently an assistant professor in the Department of Chemistry at the University of New Hampshire. There, he continues to pursue research interests in bioinorganic chemistry and in chemistry education.

Katherine M. Paulsen grew up in Erie, PA, where she completed her B.S. in chemistry from Gannon University in 2007. Katherine received her Ph.D. in chemistry from the University of Wisconsin−Madison under the supervision of Professor Judith N. Burstyn in 2013. In the Burstyn laboratory, the broad scope of her research examined the interaction of exogenous and native metal ions with proteins either to disrupt protein structure or to provide structural stability for proper protein function. Katherine is currently working for Thermo Fisher Scientific as a Senior Applications Scientist supporting the sales of molecular spectroscopy instrumentation.

Katherine A. Marvin grew up in Albany, IN. She received her B.S. in chemistry from Ball State University in 2001. She was employed at Sherry Analytical Laboratory as an analytical chemist in Muncie, IN, for a year before returning to graduate school. In 2008, she received her Ph.D. in chemistry from the University of Wisconsin−Madison under the supervision of professor Judith N. Burstyn. In her research, Katherine examined the redox and gas-sensing behavior of several hemoproteins with cysteine(thiolate) ligation using a variety of spectroscopic techniques. From 2008−2010, Katherine was employed as a Research Specialist in an undergraduate program developed at the University of Texas at Austin known as the Freshman Research Initiative. Currently, she is a visiting assistant professor in the chemistry department at Hendrix College in Conway, AR.

Judith N. Burstyn grew up in Boston, London, Pittsburgh, and New Jersey. The anchor of her childhood was summers in Woods Hole, MA, attending the Children’s School of Science. Judith received her degrees in chemistry: B.A. from Cornell in 1980, Ph.D. from University of California−Los Angeles in 1986. Her post-doctoral training was in pharmacology, molecular biology, and chemistry at UCLA School of Medicine and MIT. Judith’s research interests have centered on bioinorganic chemistry, in metal-mediated hydrolysis and metal centers in gas sensing and response. Her recent work has focused on how heme functions as a site for allosteric regulation of function in diverse protein families, including CO and NO sensors.

ACKNOWLEDGMENTS This work was supported by NIH grant HL-065217 and NSF grant CHE-1213739 to J.N.B. We thank John Fornoff for his assistance with table of contents graphic for this manuscript and Matthew Dent for checking the references.

Daniel Stevens grew up in Amery, WI. He received his B.S. in Chemistry from the University of Minnesota in 1998 and Ph.D. in Biochemistry from the University of California−Santa Cruz in 2008. Working with Prof. Glenn Millhauser, his work was focused on copper binding to the prion protein. He was a postdoctoral researcher with Judith Burstyn at

ABBREVIATIONS ARNT vertebrate aryl hydrocarbon receptor nuclear translocator BMAL1 brain and muscle Arnt-like protein-1

the University of Wisconsin−Madison, where he studied hemoprotein allostery. Dan is a currently a scientist in the Biophysics Instrumentation Facility in the UW−Madison Department of Biochemistry. 2553

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M.; Roach, M. P.; Coulter, E. D.; Dawson, J. H. Chem. Rev. 1996, 96, 2841. (2) Martinez, S. E.; Huang, D.; Szczepaniak, A.; Cramer, W. A.; Smith, J. L. Structure 1994, 2, 95. (3) Murthy, M. R. N.; Reid, T. J.; Sicignaro, A.; Tanaka, N.; Rossmann, M. G. J. Mol. Biol. 1981, 152, 465. (4) Poulos, T. L.; Johnson, E. F. In Cytochrome P450: Structure, Mechanism, and Biochemistry, 3rd ed.; Ortiz de Montellano, P. R., Ed.; Plenum Publishers: New York, 2005. (5) (a) Sundaramoorthy, M.; Terner, J.; Poulos, T. L. Structure 1995, 3, 1367. (b) Dawson, J. H.; Trudell, J. R.; Barth, G.; Linder, R. E.; Bunnenberg, E.; Djerassi, C.; Chiang, R.; Hager, L. P. J. Am. Chem. Soc. 1976, 98, 3709. (6) Daiber, A.; Shoun, H.; Ullrich, V. J. Inorg. Biochem. 2005, 99, 185. (7) (a) Li, H.; Poulos, T. L. J. Inorg. Biochem. 2005, 99, 293. (b) Sari, M.-A.; Booker, S.; Jaouen, M.; Vadon, S.; Boucher, J.-l.; Pompon, D.; Mansuy, D. Biochemistry 1996, 35, 7204. (8) (a) Shelver, D.; Thorsteinsson, M. V.; Kerby, R. L.; Chung, S.-Y.; Roberts, G. P.; Reynolds, M. F.; Parks, R. B.; Burstyn, J. N. Biochemistry 1999, 38, 2669. (b) Reynolds, M. F.; Shelver, D.; Kerby, R. L.; Parks, R. B.; Roberts, G. P.; Burstyn, J. N. J. Am. Chem. Soc. 1998, 120, 9080. (9) (a) Hasegawa, T.; Sadano, H.; Omura, T. J. Biochem. 1984, 96, 265. (b) Meier, M.; Janosik, M.; Kery, V.; Kraus, J. P.; Burkhard, P. EMBO J. 2001, 20, 3910. (c) Omura, T.; Sadano, H.; Hasegawa, T.; Yoshida, Y.; Kominami, S. J. Biochem. 1984, 96, 1491. (10) Igarashi, J.; Sato, A.; Kitagawa, T.; Yoshimura, T.; Yamauchi, S.; Sagami, I.; Shimizu, T. J. Biol. Chem. 2004, 279, 15752. (11) Uchida, T.; Sato, E.; Sato, A.; Sagami, I.; Shimizu, T.; Kitagawa, T. J. Biol. Chem. 2005, 280, 21358. (12) Weichsel, A.; Maes, E. M.; Andersen, J. F.; Valenzuela, J. G.; Shokhireva, T. K.; Walker, F. A.; Montfort, W. R. Proc. Natl. Acad. Sci. U.S.A. 2005, 102, 594. (13) Marvin, K. A.; Kerby, R. L.; Youn, H.; Roberts, G. P.; Burstyn, J. N. Biochemistry 2008, 47, 9016. (14) (a) de Rosny, E.; de Groot, A.; Jullian-Binard, C.; Gaillard, J.; Borel, F.; Pebay-Peyroula, E.; Fontecilla-Camps, J. C.; Jouve, H. l. n. M. Biochemistry 2006, 45, 9727. (b) Marvin, K. A.; Reinking, J. L.; Lee, A. J.; Pardee, K.; Krause, H. M.; Burstyn, J. N. Biochemistry 2009, 48, 7056. (15) de Rosny, E.; de Groot, A.; Jullian-Binard, C.; Borel, F.; Suarez, C.; Le Pape, L.; Fontecilla-Camps, J. C.; Jouve, H. M. Biochemistry 2008, 47, 13252. (16) (a) Klingenberg, M. Arch. Biochem. Biophys. 1958, 75, 376. (b) Omura, T.; Sato, R. J. Biol. Chem. 1962, 237, 2370. (17) Stern, J. O.; Peisach, J. J. Biol. Chem. 1974, 249, 7495. (18) Hahn, J. E.; Hodgson, K. O.; Andersson, L. A.; Dawson, J. H. J. Biol. Chem. 1982, 257, 10934. (19) Poulos, T. L.; Finzel, B. C.; Gunsalus, I. C.; Wagner, G. C.; Kraut, J. J. Biol. Chem. 1985, 260, 16122. (20) Gouterman, M. In The Porphyrins; Dolphin, D., Ed.; Academic Press: New York, 1978; Vol. III. (21) (a) Sligar, S. G. Biochemistry 1976, 15, 5399. (b) Raag, R.; Poulos, T. L. Biochemistry 1989, 28, 917. (c) Haines, D. C.; Tomchick, D. R.; Machius, M.; Peterson, J. A. Biochemistry 2001, 40, 13456. (22) Dawson, J. H.; Andersson, L. A.; Sono, M. J. Biol. Chem. 1982, 257, 3606. (23) McKnight, J.; Cheeseman, M. R.; Thomson, A. J.; Miles, J. S.; Munro, A. W. Eur. J. Biochem. 1993, 213, 683. (24) (a) Sono, M.; Stuehr, D. J.; Ikeda-Saito, M.; Dawson, J. H. J. Biol. Chem. 1995, 270, 19943. (b) Stuehr, D. J.; Ikeda-Saito, M. J. Biol. Chem. 1992, 267, 20547. (25) Galinato, M. G. I.; Spolitak, T.; Ballou, D. P.; Lehnert, N. Biochemistry 2011, 50, 1053. (26) (a) Dawson, J. H.; Holm, R. H.; Trudell, J. R.; Barth, G.; Linder, R. E.; Bunnenberg, E.; Djerassi, C.; Tang, C. S. J. Am. Chem. Soc. 1976, 98, 3707. (b) Collman, J. P.; Sorrell, T. N.; Dawson, J. H.; Trudell, J. R.; Bunnenberg, E.; Djerassi, C. Proc. Natl. Acad. Sci. U.S.A. 1976, 73, 6. (c) Sono, M.; Andersson, L. A.; Dawson, J. H. J. Biol. Chem. 1982, 257, 8308.

BxRcoM

the transcriptional regulator of CO metabolism from Burkholderia xenovorans CBS cystathionine β-synthase cNP nitrophorin from Cimex lectularius CO carbon monoxide Co hCBS cobalt CBS from Homo sapiens CoPPIX cobalt PPIX CPO chloroperoxidase CPO-I chloroperoxidase Compound I CPO-II chloroperoxidase Compound II DFT density functional theory DGCR8 DiGeorge critical region 8 protein DHR51 hormone receptor 51 from Drosophila melanogaster DmCBS CBS from Drosophila melanogaster DmE75 the nuclear receptor E75 from Drosophila melanogaster eIF2α the eukaryotic initiation factor 2α kinase eNOS endothelial nitric oxide synthase EPR electron paramagnetic resonance FAD flavin adenine dinucleotide FePPIX iron PPIX FMN flavin mononucleotide FoNOR nitric oxide reductase from Fusarium oxysporum HasA heme acquisition system protein A H4B (6R)-5,6,7,8-tetrahydro-L-biopterin HRI heme-regulated eukaryotic initiation factor 2α HsRev-erbβ the nuclear receptor Rev-erbβ from Homo sapiens iNOS inducible nitric oxide synthase LMCT ligand-to-metal-charge-transfer MCD magnetic circular dichroism miRNA microRNA NADH reduced from of nicotinamide adenine dinucleotide NADPH reduced form of nicotinamide adenine dinucleotide phosphate nNOS neuronal nitric oxide synthase NO nitric oxide NOR nitric oxide reductase NPAS2 the transcriptional regulator neuronal PAS domain protein 2 NRVS nuclear resonance vibrational spectroscopy P420 an inactive form of cytochrome P450 P450-I cytochrome P450 Compound I P450-II cytochrome P450 Compound II P450CAM camphor hydroxylating cytochrome P450 from Pseudomonas putida PAS a domain structure named from the proteins in which the motif was first identified (PER, ARNT, and SIM) PER the Drosophila period clock protein PLP pyridoxal-5′-phosphate PPIX protoporphyrin IX pre-miRNA precursor miRNA pri-miRNA primary miRNA rR resonance Raman RrCooA a CO-sensing transcriptional regulator from Rhodospirillum rubrum SIM Drosophila single-minded protein

REFERENCES (1) (a) Bertini, I.; Gray, H. B.; Lippard, S. J.; Valentine, J. S. Bioinorganic Chemistry; University Science Books: Mill Valley, CA, 1994. (b) GillesGonzalez, M.-A.; Gonzalez, G. J. Inorg. Biochem. 2005, 99, 1. (c) Sono, 2554

DOI: 10.1021/cr500056m Chem. Rev. 2015, 115, 2532−2558

Chemical Reviews

Review

(27) (a) Holmquist, B. In The Porphyrins; Dolphin, D., Ed.; Academic Press: New York, 1978; Vol. 3. (b) Cheek, J.; Dawson, J. H. In The Porphyrin Handbook; Kadish, K. M., Smith, K. M., Guilard, R., Eds.; Academic Press: San Diego, CA, 2000; Vol. 7. (c) Sutherland, J. C. Methods Enzymol. 1995, 246, 110. (d) Thomson, A. J.; Cheesman, M. R.; George, S. J. Methods Enzymol. 1993, 226, 199. (e) Cheesman, M. R.; Greenwood, C.; Thomson, A. J. Adv. Inorg. Chem. 1991, 36, 201. (f) Piepho, S. B.; Schatz, P. N. Group Theory in Spectroscopy with Applications to Magnetic Circular Dichroism; Wiley & Sons: New York, 1983. (g) Stephens, P. J. Adv. Chem. Phys. 1976, 35, 197. (h) Anzenbacher, P.; Dawson, J. H.; Kitagawa, T. J. Mol. Struct. 1989, 214, 149. (28) (a) Shimizu, T.; Iizuka, T.; Shimada, H.; Ishimura, Y.; Nozawa, T.; Hatano, M. Biochim. Biophys. Acta 1981, 670, 341. (b) Dawson, J. H.; Dooley, D. M. In Iron Porphyrins; Lever, A. B. P., Gray, H. B., Eds.; VCH Publishers: New York, 1989; Vol. 3. (c) Walker, F. A. Coord. Chem. Rev. 1999, 185−186, 471. (d) Svastits, E. W.; Alberta, J. A.; Kim, I.-C.; Dawson, J. H. Biochem. Biophys. Res. Commun. 1989, 165, 1170. (29) (a) Shimizu, T.; Nozawa, T.; Hatano, M.; Imai, Y.; Sato, R. Biochemistry 1975, 14, 4172. (b) Andersson, L. A.; Johnson, A. K.; Peterson, J. A. Arch. Biochem. Biophys. 1997, 345, 79. (30) Vickery, L.; Salmon, A.; Sauer, K. Biochim. Biophys. Acta 1975, 386, 87. (31) Dhawan, I. K.; Shelver, D.; Thorsteinsson, M. V.; Roberts, G. P.; Johnson, M. K. Biochemistry 1999, 38, 12805. (32) Pazicni, S.; Lukat-Rodgers, G. S.; Oliveriusova, J.; Rees, K. A.; Parks, R. B.; Clark, R. W.; Rodgers, K. R.; Kraus, J. P.; Burstyn, J. N. Biochemistry 2004, 43, 14684. (33) Barr, I.; Smith, A. T.; Senturia, R.; Chen, Y.; Scheidemantle, B. D.; Burstyn, J. N.; Guo, F. J. Biol. Chem. 2011, 286, 16716. (34) (a) Thomson, A. J.; Gadsby, P. M. A. J. Chem. Soc., Dalton Trans. (19721999) 1990, 1921. (b) Gadsby, P. M. A.; Thomson, A. J. J. Am. Chem. Soc. 1990, 112, 5003. (c) Cheesman, M. R.; Watmough, N. J.; Pires, C. A.; Turner, R.; Brittain, T.; Gennis, R. B.; Greenwood, C.; Thomson, A. J. Biochem. J. 1993, 289, 709. (35) Simpkin, D.; Palmer, G.; Devlin, F. J.; McKenna, M. C.; Jensen, G. M.; Stephens, P. J. Biochemistry 1989, 28, 8033. (36) Spiro, T. G. In Iron Porphyrins; Lever, A. B. P., Gray, H. B., Eds.; Addison-Wesley Publishing Co.: Reading, MA, 1983; Vol. 2. (37) (a) Spiro, T. G.; Stong, J. D.; Stein, P. J. Am. Chem. Soc. 1979, 101, 2648. (b) Abe, M.; Kitagawa, T.; Kyogoku, Y. J. Chem. Phys. 1978, 69, 4526. (38) Unno, M.; Christian, J. F.; Benson, D. E.; Gerber, N. C.; Silgar, S. G.; Champion, P. M. J. Am. Chem. Soc. 1997, 119, 6614. (39) (a) Champion, P. M.; Stallard, B. R.; Wagner, G. C.; Gunsalus, I. C. J. Am. Chem. Soc. 1982, 104, 5469. (b) Champion, P. M.; Gunsalus, I. C.; Wagner, G. C. J. Am. Chem. Soc. 1978, 100, 3743. (40) Liu, Y.; Moënne-Loccoz, P.; Hildebrand, D. P.; Wilks, A.; Loehr, T. M.; Mauk, A. G.; Ortiz de Montellano, P. R. Biochemistry 1999, 38, 3733. (41) Bangcharoenpaurpong, O.; Champion, P. M.; Hall, K. S.; Hager, L. P. Biochemistry 1986, 25, 2374. (42) Ishikawa, H.; Nakagaki, M.; Bamba, A.; Uchida, T.; Hori, H.; O’Brian, M. R.; Iwai, K.; Ishimori, K. Biochemistry 2011, 50, 1016. (43) Giroud, C.; Moreau, M.; Mattioli, T. A.; Balland, V.; Boucher, J.L.; Xu-Li, Y.; Stuehr, D. J.; Santolini, J. J. Biol. Chem. 2010, 285, 7233. (44) Green, E. L.; Taoka, S.; Banerjee, R.; Loehr, T. M. Biochemistry 2001, 40, 459. (45) Smith, A. T.; Su, Y.; Stevens, D. J.; Majtan, T.; Kraus, J. P.; Burstyn, J. N. Biochemistry 2012, 51, 6360. (46) (a) Jefcoate, C. R. E.; Gaylor, J. L. Biochemistry 1969, 8, 3464. (b) Bayer, E.; Hill, H. A. O.; Röder, A.; Williams, R. J. P. J. Chem. Soc., Chem. Commun. 1969, 3, 109. (c) Tang, S. C.; Koch, S.; Papaefthymiou, G. C.; Foner, S.; Frankel, R. B.; Ibers, J. A.; Holm, R. H. J. Am. Chem. Soc. 1976, 98, 2414. (d) Collman, J. P.; Sorrell, T. N.; Hoffman, B. M. J. Am. Chem. Soc. 1975, 97, 913. (e) Koch, S.; Tang, S. C.; Holm, R. H.; Frankel, R. B.; Ibers, J. A. J. Am. Chem. Soc. 1975, 97, 916. (f) Röder, A.; Bayer, E. Eur. J. Biochem. 1969, 11, 89. (47) Blumberg, W. E.; Peisach, J. Adv. Chem. Ser. 1971, 100, 271.

(48) Sono, M.; Hager, L. P.; Dawson, J. H. Biochim. Biophys. Acta 1991, 1078, 351. (49) Reynolds, M. F.; Parks, R. B.; Burstyn, J. N.; Shelver, D.; Thorsteinsson, M. V.; Kerby, R. L.; Roberts, G. P.; Vogel, K. M.; Spiro, T. G. Biochemistry 2000, 39, 388. (50) Nastainczyk, W.; Ruf, H. H.; Ullrich, V. Eur. J. Biochem. 1975, 60, 615. (51) He, C.; Nishikawa, K.; Erdem, Ö . F.; Reijerse, E.; Ogata, H.; Lubitz, W.; Knipp, M. J. Inorg. Biochem. 2013, 122, 38. (52) Caillet-Saguy, C.; Turano, P.; Piccioli, M.; Lukat-Rodgers, G. S.; Czjzek, M.; Guigliarelli, B.; Izadi-Pruneyre, N.; Rodgers, K. R.; Delepierre, M.; Lecroisey, A. J. Biol. Chem. 2008, 283, 5960. (53) Pazicni, S. University of Wisconsin−Madison, 2006. (54) (a) Hollenberg, P. F.; Hager, L. P.; Blumberg, W. E.; Peisach, J. J. Biol. Chem. 1980, 255, 4801. (b) Tsai, R.; Yu, C. A.; Gunsalus, I. C.; Peisach, J.; Blumberg, W.; Orme-Johnson, W. H.; Beinert, H. Proc. Natl. Acad. Sci. U.S.A. 1970, 66, 1157. (c) Peisach, J.; Blumberg, W. E. Proc. Natl. Acad. Sci. U.S.A. 1970, 67, 172. (d) Palmer, G. In Iron Porphyrins; Lever, A. B. P., Gray, H. B., Eds.; Addison-Wesley: Reading, MA, 1983; Vol. II. (55) Peisach, J.; Blumberg, W. E.; Ogawa, S.; Rachmilewitz, E. A.; Oltzik, R. J. Biol. Chem. 1971, 246, 3342. (56) (a) Omura, T.; Sato, R. J. Biol. Chem. 1964, 239, 2370. (b) Peterson, J. A.; Griffin, B. W. Arch. Biochem. Biophys. 1972, 151, 427. (57) (a) O’Keeffe, D. H.; Ebel, R. E.; Peterson, J. A. J. Biol. Chem. 1978, 253, 3509. (b) Ebel, R. E.; O’Keeffe, D. H.; Peterson, J. A. FEBS Lett. 1975, 55, 198. (c) Stern, J. O.; Peisach, J. FEBS Lett. 1976, 62, 364. (58) (a) Mansuy, D.; Beaune, P.; Chottard, J. C.; Bartoli, J. F.; Gans, P. Biochem. Pharmacol. 1976, 25, 609. (b) Mansuy, D.; Gans, P.; Chottard, J.-C.; Bartoli, J.-F. Eur. J. Biochem. 1977, 76, 607. (59) Ullrich, V.; Nastainczyk, W.; Ruf, H. H. Biochem. Soc. Trans. 1975, 3, 803. (60) Werringloer, J.; Estabrook, R. W. Arch. Biochem. Biophys. 1975, 167, 270. (61) Dawson, J. H.; Andersson, L. A.; Sono, M. J. Biol. Chem. 1983, 258, 13637. (62) Hanson, L. K.; Eaton, W. A.; Sligar, S. G.; Gunsalus, I. C.; Gouterman, M.; Connell, C. R. J. Am. Chem. Soc. 1976, 98, 2672. (63) (a) Jung, C.; Ristau, O. Chem. Phys. Lett. 1977, 49, 103. (b) Jung, C. Chem. Phys. Lett. 1985, 113, 589. (64) (a) Greschner, S.; Sharonov, Y. A.; Jung, C. FEBS Lett. 1993, 315, 153. (b) Yarmola, E. G.; Sharonov, Y. A. FEBS Lett. 1994, 355, 279. (65) Ozaki, Y.; Kitagawa, T.; Kyogoku, Y.; Shimada, H.; Iizuka, T.; Ishimura, Y. J. Biochem. 1976, 80, 1447. (66) (a) Remba, R. D.; Champion, P. M.; Fitchen, D. B.; Chiang, R.; Hager, L. P. Biochemistry 1979, 18, 2280. (b) Champion, P. M.; Remba, R. D.; Chiang, R.; Fitchen, D. B.; Hager, L. P. Biochim. Biophys. Acta 1976, 446, 486. (67) Wang, J.; Stuehr, D. J.; Ikeda-Saito, M.; Rousseau, D. L. J. Biol. Chem. 1993, 268, 22255. (68) (a) Champion, P. M. In Biological Applications of Raman Spectroscopy; Spiro, T. G., Ed.; Wiley: New York, 1988; Vol. 3. (b) Anzenbacher, P.; Evangelista-Kirkup, R.; Schenkman, J.; Spiro, T. G. Inorg. Chem. 1989, 28, 4491. (69) Momenteau, M.; Reed, C. A. Chem. Rev. 1994, 94, 659. (70) Ishimura, Y.; Ullrich, V.; Peterson, J. A. Biochem. Biophys. Res. Commun. 1971, 42, 140. (71) Antonini, E.; Brunori, M. Hemoglobin and Myoglobin in Their Reactions with Ligands; North-Holland Publishing Co.: Amsterdam, 1971. (72) (a) Dawson, J. H.; Cramer, S. P. FEBS Lett. 1978, 88, 127. (b) Vickery, L.; Nozawa, T.; Sauer, K. J. Am. Chem. Soc. 1976, 98, 351. (73) Bangcharoenpaurpong, O.; Rizos, A. K.; Champion, P. M.; Jollie, D.; Sligar, S. G. J. Biol. Chem. 1986, 261, 8089. (74) Chottard, G.; Schappacher, M.; Ricard, L.; Weiss, R. Inorg. Chem. 1984, 23, 4557. (75) Matsu-ura, M.; Tani, F.; Nakayama, S.; Nakamura, N.; Naruta, Y. Angew. Chem., Int. Ed. 2000, 39, 1989. 2555

DOI: 10.1021/cr500056m Chem. Rev. 2015, 115, 2532−2558

Chemical Reviews

Review

(76) Alben, J. O. In The Porphyrins; Dolphin, D., Ed.; Academic Press: New York, 1978; Vol. 3, Part A. (77) (a) Okubo, S.; Nozawa, T.; Hatano, M. Chem. Lett. 1981, 10, 1625. (b) Budyka, M. F.; Khenkin, A. M.; Shteinman, A. A. Biochem. Biophys. Res. Commun. 1981, 101, 615. (78) (a) Kühnel, K.; Blankenfeldt, W.; Terner, J.; Schlichting, I. J. Biol. Chem. 2006, 281, 23990. (b) Cupp-Vickery, J.; Poulos, T. L. Nat. Struct. Biol. 1995, 2, 144. (c) Ravichandran, K. G.; Boddupalli, S. S.; Hasemann, C. A.; Peterson, J.; Deisenhofer, J. Science 1993, 261, 731. (d) Hasemann, C. A.; Kurumbail, R. G.; Boddupalli, S. S.; Peterson, J.; Deisenhofer, J. Structure 1995, 3, 41. (79) (a) Roach, M. P.; Pond, A. E.; Thomas, M. R.; Boxer, S. G.; Dawson, J. H. J. Am. Chem. Soc. 1999, 121, 12088. (b) Glascock, M. C.; Ballou, D. P.; Dawson, J. H. J. Biol. Chem. 2005, 280, 42134. (80) Sono, M.; Eble, K. S.; Dawson, J. H.; Hager, L. P. J. Biol. Chem. 1985, 260, 15530. (81) (a) Marchal, S.; Gorren, A. C. F.; Sørlie, M.; Andersson, K. K.; Mayer, B.; Lange, R. J. Biol. Chem. 2004, 279, 19824. (b) Ledbetter, A. P.; McMillan, K.; Roman, L. J.; Masters, B. S. S.; Dawson, J. H.; Sono, M. Biochemistry 1999, 38, 8014. (82) (a) Dawson, J. H. Science 1988, 240, 433. (b) Hiner, A. N. P.; Raven, E. L.; Thorneley, R. N. F.; García-Cánovas, F.; Rodríguez-López, J. N. J. Inorg. Biochem. 2002, 91, 27. (83) (a) Green, M. T.; Dawson, J. H.; Gray, H. B. Science 2004, 304, 1653. (b) Stone, K. L.; Behan, R. K.; Green, M. T. Proc. Natl. Acad. Sci. U.S.A. 2006, 103, 12307. (c) Green, M. T.; Dawson, J. H.; Gray, H. B. Science 2004, 304, 1653. (84) (a) Egawa, T.; Proshlyakov, D. A.; Miki, H.; Makino, R.; Ogura, T.; Kitagawa, T.; Ishimura, Y. J. Biol. Inorg. Chem. 2001, 6, 46. (b) Palcic, M. M.; Rutter, R.; Araiso, T.; Hager, L. P.; Dunford, H. B. Biochem. Biophys. Res. Commun. 1980, 94, 1123. (85) (a) Hosten, C. M.; Sullivan, A. M.; Palaniappan, V.; Fitzgerald, M. M.; Terner, J. J. Biol. Chem. 1994, 269, 13966. (b) Egawa, T.; Miki, H.; Ogura, T.; Makino, R.; Ishimura, Y.; Kitagawa, T. FEBS Lett. 1992, 305, 206. (86) Rutter, R.; Hager, L. P.; Dhonau, H.; Hendrich, M.; Valentine, M.; Debrunner, P. Biochemistry 1984, 23, 6809. (87) Stone, K. L.; Behan, R. K.; Green, M. T. Proc. Natl. Acad. Sci. U.S.A. 2005, 102, 16563. (88) Harris, D.; Loew, G.; Waskell, L. J. Inorg. Biochem. 2001, 83, 309. (89) Rittle, J.; Green, M. T. Science 2010, 330, 933. (90) (a) Czarnecki, K.; Kincaid, J. R.; Fujii, H. J. Am. Chem. Soc. 1999, 121, 7953. (b) Kitagawa, T.; Mizutani, Y. Coord. Chem. Rev. 1994, 135− 136, 685. (91) Rittle, J.; Younker, J. M.; Green, M. T. Inorg. Chem. 2010, 49, 3610. (92) Zaks, A.; Dodds, D. R. J. Am. Chem. Soc. 1995, 117, 10419. (93) (a) Kim, I.-C.; Deal, W. C., Jr. Biochemistry 1976, 15, 4925. (b) Pazicni, S.; Cherney, M. M.; Lukat-Rogers, G. S.; Oliveriusová, J.; Rodgers, K. R.; Kraus, J. P.; Burstyn, J. N. Biochemistry 2005, 44, 16785. (94) Jensen, K. P.; Ryde, U. J. Biol. Chem. 2004, 279, 14561. (95) Dawson, J. H.; Sono, M. Chem. Rev. 1987, 87, 1255. (96) (a) Yu, N.-T.; Kerr, E. A. In Biological Applications of Raman Spectroscopy; Spiro, T. G., Ed.; John Wiley & Sons: New York, 1988; Vol. 3. (b) Ray, G. B.; Li, X.-Y.; Ibers, J. A.; Sessler, J. L.; Spiro, T. G. J. Am. Chem. Soc. 1994, 116, 162. (c) Spiro, T. G.; Wasbotten, I. H. J. Inorg. Biochem. 2005, 99, 34. (97) Li, X.-Y.; Spiro, T. G. J. Am. Chem. Soc. 1987, 110, 6024. (98) (a) Traylor, T. G.; Sharma, V. S. Biochemistry 1992, 31, 2847. (b) Enemark, J. H.; Feltham, R. D. Coord. Chem. Rev. 1974, 13, 339. (99) Quaroni, L. G.; Seward, H. E.; McLean, K. J.; Girvan, H. M.; Ost, T. W. B.; Noble, M. A.; Kelly, S. M.; Price, N. C.; Cheesman, M. R.; Smith, W. E.; Munro, A. W. Biochemistry 2004, 43, 16416. (100) (a) Lehnert, N.; Sage, J. T.; Silvernail, N.; Scheidt, W. R.; Alp, E. E.; Sturhahn, W.; Zhao, J. Inorg. Chem. 2010, 49, 7197. (b) Decatur, S. M.; Franzen, S.; DePillis, G. D.; Dyer, R. B.; Woodruff, W. H.; Boxer, S. G. Biochemistry 1996, 35, 4939. (101) Taoka, S.; Banerjee, R. J. Inorg. Biochem. 2001, 87, 245.

(102) Yonetani, T.; Yamamoto, H.; Erman, J. E.; Leigh, J. S.; Reed, G. H. J. Biol. Chem. 1972, 247, 2447. (103) (a) Averill, B. A. Chem. Rev. 1996, 96, 2951. (b) Praneeth, V. K. K.; Paulat, F.; Berto, T. C.; George, S. D.; Näther, C.; Sulok, C. D.; Lehnert, N. J. Am. Chem. Soc. 2008, 130, 15288. (104) (a) Paulat, F.; Lehnert, N. Inorg. Chem. 2007, 46, 1547. (b) Xu, N.; Goodrich, L. E.; Lehnert, N.; Powell, D. R.; Richter-Addo, G. B. Angew. Chem., Int. Ed. 2013, 52, 3896. (105) (a) Chiang, R.; Makino, R.; Spomer, W. E.; Hager, L. P. Biochemistry 1975, 14, 4166. (b) Praneeth, V. K. K.; Haupt, E.; Lehnert, N. J. Inorg. Biochem. 2005, 99, 940. (106) Kon, H. Biochim. Biophys. Acta 1975, 379, 103. (107) (a) Xu, N.; Powell, D. R.; Cheng, L.; Richter-Addo, G. B. Chem. Commun. 2006, 19, 2030. (b) Obayashi, E.; Tsukamoto, K.; Adachi, S.-i.; Takahashi, S.; Nomura, M.; Iizuka, T.; Shoun, H.; Shiro, Y. J. Am. Chem. Soc. 1997, 119, 7807. (c) Hu, S.; Kincaid, J. R. J. Am. Chem. Soc. 1991, 113, 9760. (108) (a) Soldatova, A. V.; Ibrahim, M.; Olson, J. S.; Czernuszewicz, R. S.; Spiro, T. G. J. Am. Chem. Soc. 2010, 132, 4614. (b) Maes, E. M.; Walker, F. A.; Montfort, W. R.; Czernuszewicz, R. S. J. Am. Chem. Soc. 2001, 123, 11664. (109) (a) Goodrich, L. E.; Paulat, F.; Praneeth, V. K. K.; Lehnert, N. Inorg. Chem. 2010, 49, 6293. (b) Zeng, W.; Silvernail, N. J.; Wharton, D. C.; Georgiev, G. Y.; Leu, B. M.; Scheidt, W. R.; Zhao, J.; Sturhahn, W.; Alp, E. E.; Sage, J. T. J. Am. Chem. Soc. 2005, 127, 11200. (c) Silvernail, N. J.; Barabanschikov, A.; Pavlik, J. W.; Noll, B. C.; Zhao, J.; Alp, E. E.; Sturhahn, W.; Sage, J. T.; Scheidt, W. R. J. Am. Chem. Soc. 2007, 129, 2200. (d) Paulat, F.; Berto, T. C.; George, S. D.; Goodrich, L. E.; Praneeth, V. K. K.; Sulok, C. D.; Lehnert, N. Inorg. Chem. 2008, 47, 11449. (110) Benko, B.; Yu, N. T. Proc. Natl. Acad. Sci. U.S.A. 1983, 80, 7042. (111) Vogel, K. M.; Kozlowski, P. M.; Zgierski, M. Z.; Spiro, T. G. J. Am. Chem. Soc. 1999, 121, 9915. (112) Omura, T. Biochem. Biophys. Res. Commun. 2005, 338, 404. (113) (a) Guengrich, F. P. Chem. Res. Toxicol. 2001, 14, 611. (b) Guengrich, F. P. In Cytochrome P450: Structure, Mechanism, and Biochemistry; Ortiz de Montellano, P. R., Ed.; Kluwer Academic/ Plenum: New York, 2005. (114) (a) White, R. E. Pharmacol. Ther. 1991, 49, 21. (b) Groves, J. T. J. Chem. Educ. 1985, 62, 928. (c) Groves, J. T.; Haushalter, R. C.; Nakamura, M.; Nemo, T. E.; Evans, B. J. J. Am. Chem. Soc. 1981, 103, 2884. (115) (a) Frew, J. E.; Jones, P. In Advances in Inorganic and Bioinorganic Mechanisms; Sykes, A. G., Ed.; Academic Press: Orlando, FL, 1984; Vol. 3. (b) Neidleman, S. L.; Geigert, J. Biohalogenation: Principles, Basic Roles and Applications; Halsted Press: New York, 1986. (116) Osborne, R. L.; Coggins, M. K.; Terner, J.; Dawson, J. H. J. Am. Chem. Soc. 2007, 129, 14838. (117) (a) MacMicking, J.; Xie, Q.-w.; Nathan, C. Annu. Rev. Immunol. 1997, 15, 323. (b) Lincoln, J.; Hoyle, C. H. V.; Burnstock, G. Nitric Oxide in Health and Disease; Cambridge University Press: Cambridge, UK, 1997. (c) Cooke, J. P.; Dzau, V. J. Annu. Rev. Med. 1997, 48, 489. (118) (a) Alderton, W. K.; Cooper, C. E.; Knowles, R. G. Biochem. J. 2001, 357, 593. (b) Fischmann, T. O.; Hruza, A.; Niu, X. D.; Fossetta, J. D.; Lunn, C. A.; Dolphin, E.; Prongay, A. J.; Reichert, P.; Lundell, D. J.; Narula, S. K.; Weber, P. Nat. Struct. Biol. 1999, 6, 233. (119) Raman, C. S.; Martasek, P.; Masters, B. S. S. In The Porphyrin Handbook; Kadish, K. M., Smith, K. M., Guilard, R., Eds.; Academic Press: New York, 2000. (120) Rousseau, D. L.; Li, D.; Couture, M.; Yeh, S.-R. J. Inorg. Biochem. 2005, 99, 306. (121) Woodward, J. J.; Chang, M. M.; Martin, N. I.; Marletta, M. A. J. Am. Chem. Soc. 2009, 131, 297. (122) Davydov, R.; Sudhamsu, J.; Lees, N. S.; Crane, B. R.; Hoffman, B. M. J. Am. Chem. Soc. 2009, 131, 14493. (123) (a) Kudo, T.; Tomura, D.; Liu, D. L.; Dai, X. Q.; Shoun, H. Biochimie 1996, 78, 792. (b) Usuda, K.; Toritsuka, N.; Matsuo, Y.; Kim, D. H.; Shoun, H. Appl. Environ. Microbiol. 1995, 61, 883. (c) Zhang, L.; Takaya, N.; Kitazume, T.; Kondo, T.; Shoun, H. Eur. J. Biochem. 2001, 2556

DOI: 10.1021/cr500056m Chem. Rev. 2015, 115, 2532−2558

Chemical Reviews

Review

268, 3198. (d) Tsuruta, S.; Takaya, N.; Zhang, L.; Shoun, H.; Kimura, K.; Hamamoto, M.; Nakase, T. FEMS Microbiol. Lett. 1998, 168, 105. (124) Shoun, H.; Kano, M.; Baba, I.; Takaya, N.; Matsuo, M. J. Bacteriol. 1998, 180, 4413. (125) (a) Shiro, Y.; Fujii, M.; Iizuka, T.; Adachi, S.-i.; Tsukamoto, K.; Nakahara, K.; Shoun, H. J. Biol. Chem. 1995, 270, 1617. (b) Nakahara, K.; Tanimoto, T.; Hatano, K.-i.; Usuda, K.; Shoun, H. J. Biol. Chem. 1993, 268, 8350. (126) Shimizu, H.; Park, S.-Y.; Shiro, Y.; Adachi, S.-i. Acta Crystallogr., Sect. D 2002, D58, 81. (127) (a) Silaghi-Dumitrescu, R. Eur. J. Inorg. Chem. 2003, 6, 1048. (b) Harris, D. L. Int. J. Quantum Chem. 2002, 88, 183. (c) Shimizu, H.; Obayashi, E.; Gomi, Y.; Arakawa, H.; Park, S.-Y.; Nakamura, H.; Adachi, S.-i.; Shoun, H.; Shiro, Y. J. Biol. Chem. 2000, 275, 4816. (d) Kudo, T.; Takaya, N.; Park, S.-Y.; Shiro, Y.; Shoun, H. J. Biol. Chem. 2000, 276, 5020. (e) Obayashi, E.; Takahashi, S.; Shiro, Y. J. Am. Chem. Soc. 1998, 120, 12964. (f) Lehnert, N.; Praneeth, V. K. K.; Paulat, F. J. Comput. Chem. 2006, 27, 1338. (g) Riplinger, C.; Neese, F. ChemPhysChem 2011, 12, 3192. (128) (a) Sono, M.; Dawson, J. H. J. Biol. Chem. 1982, 257, 5496. (b) LoBrutto, R.; Scholes, C. P.; Wagner, G. C.; Gunsalus, I. C.; Debrunner, P. G. J. Am. Chem. Soc. 1980, 102, 1167. (129) Ohno, T.; Suzuki, N.; Dokoh, T.; Urano, Y.; Kikuchi, K.; Hirobe, M.; Higuchi, T.; Nagano, T. J. Inorg. Biochem. 2000, 82, 123. (130) Ogliaro, F.; de Visser, S. P.; Shaik, S. J. Inorg. Biochem. 2002, 91, 554. (131) Kamachi, T.; Kouno, T.; Nam, W.; Yoshizawa, K. J. Inorg. Biochem. 2006, 100, 751. (132) (a) Green, M. T. Curr. Opin. Chem. Biol. 2009, 13, 84. (b) Behan, R. K.; Hoffart, L. M.; Stone, K. L.; Krebs, C.; Green, M. T. J. Am. Chem. Soc. 2006, 128, 11471. (133) (a) Brunel, A.; Wilson, A.; Henry, L.; Dorlet, P.; Santolini, J. J. Biol. Chem. 2011, 286, 11997. (b) Yoshioka, S.; Takahashi, S.; Ishimori, K.; Morishima, I. J. Inorg. Biochem. 2000, 81, 141. (c) Yoshioka, S.; Tosha, T.; Takahashi, S.; Ishimori, K.; Hori, H.; Morishima, I. J. Am. Chem. Soc. 2002, 124, 14571. (d) Lang, J.; Driscoll, D.; Gélinas, S.; Rafferty, S. P.; Couture, M. J. Inorg. Biochem. 2009, 103, 1102. (e) Hannibal, L.; Somasundaram, R.; Tejero, J.; Wilson, A.; Stuehr, D. J. J. Biol. Chem. 2011, 286, 39224. (f) Couture, M.; Adak, S.; Stuehr, D. J.; Rousseau, D. L. J. Biol. Chem. 2001, 276, 38280. (g) Usharani, D.; Zazza, C.; Lai, W.; Chourasia, M.; Waskell, L.; Shaik, S. J. Am. Chem. Soc. 2012, 134, 4053. (h) Lang, J.; Santolini, J.; Couture, M. Biochemistry 2011, 50, 10069. (i) Mak, P. J.; Yang, Y.; Im, S.; Waskell, L. A.; Kincaid, J. R. Angew. Chem. 2012, 51, 10403. (134) Ullrich, V. Arch. Biochem. Biophys. 2003, 409, 45. (135) Behan, R. K.; Green, M. T. J. Inorg. Biochem. 2006, 100, 448. (136) Nakajima, R.; Yamazaki, I.; Griffin, B. W. Biochem. Biophys. Res. Commun. 1985, 128, 1. (137) Green, M. T. J. Am. Chem. Soc. 2006, 128, 1902. (138) Stone, K. L.; Hoffart, L. M.; Behan, R. K.; Krebs, C.; Green, M. T. J. Am. Chem. Soc. 2006, 128, 6147. (139) (a) Gardner, K. A.; Kuehnert, L.; Mayer, J. M. Inorg. Chem. 1997, 36, 2069. (b) Gardner, K. A.; Mayer, J. M. Science 1995, 269, 1849. (c) Mayer, J. M. Acc. Chem. Res. 1998, 31, 441. (140) Zhong, F.; Lisi, G. P.; Collins, D. P.; Dawson, J. H.; Pletneva, E. V. Proc. Natl. Acad. Sci. U.S.A. 2014, 111, E306. (141) Igarashi, J.; Kitanishi, K.; Martinkova, M.; Murase, M.; Iizuka, A.; Shimizu, T. Acta Chim. Slov. 2008, 55, 67. (142) (a) Roberts, G. P.; Youn, H.; Kerby, R. L. Microbiol. Mol. Biol. Rev. 2004, 68, 453. (b) Roberts, G. P.; Kerby, R. L.; Youn, H.; Conrad, M. J. Inorg. Biochem. 2005, 99, 280. (143) Kerby, R. L.; Ludden, P. W.; Roberts, G. P. J. Bacteriol. 1995, 177, 2241. (144) (a) Aono, S.; Ohkubo, K.; Matsuo, T.; Nakajima, H. J. Biol. Chem. 1998, 273, 25757. (b) Clark, R. W.; Youn, H.; Parks, R. B.; Cherney, M. M.; Roberts, G. P.; Burstyn, J. N. Biochemistry 2004, 43, 14149. (c) Yamamoto, K.; Ishikawa, H.; Takahashi, S.; Ishimori, K.; Morishima, I.; Nakajima, H.; Aono, S. J. Biol. Chem. 2001, 276, 11473.

(145) Lanzilotta, W. N.; Schuller, D. J.; Thorsteinsson, M. V.; Kerby, R. L.; Roberts, G. P.; Poulos, T. L. Nat. Struct. Biol. 2000, 7, 876. (146) (a) Vogel, K. M.; Spiro, T. G.; Shelver, D.; Thorsteinsson, M. V.; Roberts, G. P. Biochemistry 1999, 38, 2679. (b) He, Y.; Shelver, D.; Kerby, R. L.; Roberts, G. P. J. Biol. Chem. 1996, 271, 120. (147) Kerby, R. L.; Youn, H.; Thorsteinsson, M. V.; Roberts, G. P. J. Mol. Biol. 2003, 325, 809. (148) Dioum, E. M.; Rutter, J.; Tuckerman, J. R.; Gonzalez, G.; GillesGonzalez, M.-A.; McKnight, S. L. Science 2002, 298, 2385. (149) (a) Reick, M.; Garcia, J. A.; Dudley, C.; McKnight, S. L. Science 2001, 293, 506. (b) Zhou, Y.-D.; Barnard, M.; Tian, H.; Li, X.; Ring, H. Z.; Francke, U.; Shelton, J.; Richardson, J.; Russell, D. W.; McKnight, S. L. Proc. Natl. Acad. Sci. U.S.A. 1997, 94, 713. (150) Mukaiyama, Y.; Uchida, T.; Sato, E.; Sasaki, A.; Sato, Y.; Igarashi, J.; Kurokawa, H.; Sagami, I.; Kitagawa, T.; Shimizu, T. FEBS J. 2006, 273, 2528. (151) Koudo, R.; Kurokawa, H.; Sato, E.; Igarashi, J.; Uchida, T.; Sagami, I.; Kitagawa, T.; Shimizu, T. FEBS J. 2005, 272, 4153. (152) Uchida, T.; Sagami, I.; Shimizu, T.; Ishimori, K.; Kitagawa, T. J. Inorg. Biochem. 2012, 108, 188. (153) Kerby, R. L.; Youn, H.; Roberts, G. P. J. Bacteriol. 2008, 190, 3336. (154) Smith, A. T.; Marvin, K. A.; Freeman, K. M.; Kerby, R. L.; Roberts, G. P.; Burstyn, J. N. J. Biol. Inorg. Chem. 2012, 17, 1071. (155) Gronemeyer, H.; Laudet, V. Protein Profile 1995, 2, 1173. (156) Renaud, J. P.; Moras, D. Cell. Mol. Life Sci. 2000, 57, 1748. (157) (a) Triqueneaux, G.; Thenot, S.; Kakizawa, T.; Antoch, M. P.; Safi, R.; Takahashi, J. S.; Delaunay, F.; Laudet, V. J. Mol. Endocrinol. 2004, 33, 585. (b) Woo, E.-J.; Jeong, D. G.; Lim, M.-Y.; Kim, S. J.; Kim, K.-J.; Yoon, S.-M.; Park, B.-C.; Ryu, S. E. J. Mol. Biol. 2007, 373, 735. (c) Preitner, N.; Damiola, F.; Lopez-Molina, L.; Zakany, J.; Duboule, D.; Albrecht, U.; Schibler, U. Cell 2002, 110, 251. (d) Torra, I. P.; Tsibulsky, V.; Delaunay, F.; Saladin, R.; Laudet, V.; Fruchart, J. C.; Kosykh, V.; Staels, B. Endocrinology 2000, 141, 3799. (158) Reinking, J.; Lam, M. M. S.; Pardee, K.; Sampson, H. M.; Liu, S.; Yang, P.; Williams, S.; White, W.; Lajoie, G.; Edwards, A.; Krause, H. M. Cell 2005, 122, 195. (159) (a) Pardee, K. I.; Xu, X.; Reinking, J.; Schuetz, A.; Dong, A.; Liu, S.; Zhang, R.; Tiefenbach, J.; Lajoie, G.; Plotnikov, A. N.; Botchkarev, A.; Krause, H. M.; Edwards, A. PLoS Biol. 2009, 7, e43. (b) Raghuram, S.; Stayrook, K. R.; Huang, P.; Rogers, P. M.; Nosie, A. K.; McClure, D. B.; Burris, L. L.; Khorasanizadeh, S.; Burris, T. P.; Rastinejad, F. Nat. Struct. Mol. Biol. 2007, 14, 1207. (c) Yin, L.; Wu, N.; Curtin, J. C.; Qatanani, M.; Szwergold, N. R.; Reid, R. A.; Waitt, G. M.; Parks, D. J.; Pearce, K. H.; Wisely, G. B.; Lazar, M. A. Science 2007, 318, 1786. (160) Gupta, N.; Ragsdale, S. W. J. Biol. Chem. 2011, 286, 4392. (161) Airola, M. V.; Du, J.; Dawson, J. H.; Crane, B. R. Biochemistry 2010, 49, 4327. (162) (a) Uma, S.; Yun, B.-G.; Matts, R. L. J. Biol. Chem. 2001, 276, 14875. (b) Chefalo, P. J.; Oh, J.; Rafie-Kolpin, M.; Kan, B.; Chen, J.-J. Eur. J. Biochem. 1998, 258, 820. (c) Ishikawa, H.; Yun, B.-G.; Takahashi, S.; Hori, H.; Matts, R. L.; Ishimori, K.; Morishima, I. J. Am. Chem. Soc. 2002, 124, 13696. (163) (a) Denver, T. E. Trends Biochem. Sci. 1999, 24, 398. (b) Chen, J.-J.; London, I. M. Trends Biochem. Sci. 1995, 20, 105. (164) Chen, J.-J. In Translational Control of Gene Expression; Soneberg, N., Hershey, J. W. B., Matthews, M. B., Eds.; Cold Spring Harbor Laboratory: Cold Spring Harbor, NY, 2000. (165) Martinkova, M.; Igarashi, J.; Shimizu, T. FEBS Lett. 2007, 581, 4109. (166) (a) Rafie-Kolpin, M.; Chefalo, P. J.; Hussain, Z.; Hahn, J.; Uma, S.; Matts, R. L.; Chen, J.-J. J. Biol. Chem. 2000, 275, 5171. (b) Uma, S.; Matts, R. L.; Guo, Y.; White, S.; Chen, J.-J. Eur. J. Biochem. 2000, 267, 498. (167) Inuzuka, T.; Yun, B.-G.; Ishikawa, H.; Takahashi, S.; Hori, H.; Matts, R. L.; Ishimori, K.; Morishima, I. J. Biol. Chem. 2004, 279, 6778. (168) (a) Igarashi, J.; Murase, M.; Iizuka, A.; Pichierri, F.; Martinkova, M.; Shimizu, T. J. Biol. Chem. 2008, 283, 18782. (b) Miksanova, M.; Igarashi, J.; Minami, M.; Sagami, I.; Yamauchi, S.; Kurokawa, H.; 2557

DOI: 10.1021/cr500056m Chem. Rev. 2015, 115, 2532−2558

Chemical Reviews

Review

Shimizu, T. Biochemistry 2006, 45, 9894. (c) Hirai, K.; Martinkova, M.; Igarashi, J.; Saiful, I.; Yamauchi, S.; El-Mashtoly, S.; Kitagawa, T.; Shimizu, T. J. Inorg. Biochem. 2007, 101, 1172. (169) Perera, R.; Sono, M.; Sigman, J. A.; Pfister, T. D.; Lu, Y.; Dawson, J. H. Proc. Natl. Acad. Sci. U.S.A. 2003, 100, 3641. (170) Yun, B.-G.; Matts, J. A. B.; Matts, R. L. Biochim. Biophys. Acta 2005, 1725, 174. (171) (a) Kery, V.; Bukovska, G.; Kraus, J. P. J. Biol. Chem. 1994, 269, 25283. (b) Taoka, S.; Lepore, B. W.; Kabil, O.; Ojha, S.; Ringe, D.; Banerjee, R. Biochemistry 2002, 41, 10454. (c) Koutmos, M.; Kabil, O.; Smith, J. L.; Banerjee, R. Proc. Natl. Acad. Sci. U.S.A. 2010, 107, 20958. (172) (a) Maclean, K. N.; Janosik, M.; Oliveriusova, J.; Kery, V.; Kraus, J. P. J. Inorg. Biochem. 2000, 81, 161. (b) Jhee, K.-H.; McPhie, P.; Miles, E. W. J. Biol. Chem. 2000, 275, 11541. (c) Nozaki, T.; Shigeta, Y.; SaitoNakano, Y.; Imada, M.; Kruger, W. D. J. Biol. Chem. 2001, 276, 6516. (173) (a) Kery, V.; Poneleit, L.; Kraus, J. P. Arch. Biochem. Biophys. 1998, 355, 222. (b) Evande, R.; Ojha, S.; Banerjee, R. Arch. Biochem. Biophys. 2004, 427, 188. (c) Oliveriusová, J.; Kery, V.; Maclean, K. N.; Kraus, J. P. J. Biol. Chem. 2002, 277, 48386. (d) Bruno, S.; Schiaretti, F.; Burkhard, P.; Kraus, J. P.; Janosik, M.; Mozzarelli, A. J. Biol. Chem. 2001, 276, 16. (e) Kery, V.; Elleder, D.; Kraus, J. P. Arch. Biochem. Biophys. 1995, 316, 24. (174) Majtan, T.; Singh, L. R.; Wang, L.; Kruger, W. D.; Kraus, J. P. J. Biol. Chem. 2008, 283, 34588. (175) (a) Majtan, T.; Freeman, K. M.; Smith, A. T.; Burstyn, J. N.; Kraus, J. P. Arch. Biochem. Biophys. 2011, 508, 25. (b) Smith, A. T.; Majtan, T.; Freeman, K. M.; Su, Y.; Kraus, J. P.; Burstyn, J. N. Inorg. Chem. 2011, 50, 4417. (176) (a) Banerjee, R.; Zou, C.-G. Arch. Biochem. Biophys. 2005, 433, 144. (b) Weeks, C. L.; Singh, S.; Madzelan, P.; Banerjee, R.; Spiro, T. G. J. Am. Chem. Soc. 2009, 131, 12809. (c) Kabil, O.; Weeks, C. L.; Carballal, S.; Gherasim, C.; Alvarez, B.; Spiro, T. G.; Banerjee, R. Biochemistry 2011, 50, 8261. (177) (a) Taoka, S.; Green, E. L.; Loehr, T. M.; Banerjee, R. J. Inorg. Biochem. 2001, 87, 253. (b) Taoka, S.; West, M.; Banerjee, R. Biochemistry 1999, 38, 2738. (178) Taoka, S.; Ohja, S.; Shan, X.; Kruger, W. D.; Banerjee, R. J. Biol. Chem. 1998, 273, 25179. (179) Cherney, M. M.; Pazicni, S.; Frank, N.; Marvin, K. A.; Kraus, J. P.; Burstyn, J. N. Biochemistry 2007, 46, 13199. (180) (a) Valenzuela, J. G.; Walker, F. A.; Ribeiro, J. M. J. Exp. Biol. 1995, 198, 1519. (b) Champagne, D. E.; Nussenzveig, R. H.; Ribeiro, J. M. C. J. Biol. Chem. 1995, 270, 8691. (c) Ribeiro, J. M.; Walker, F. A. J. Exp. Med. 1994, 180, 2251. (d) Valenzuela, J. G.; Ribeiro, J. M. J. Exp. Biol. 1998, 201, 2659. (e) Ribeiro, J. M.; Hazzard, J. M.; Nussenzveig, R. H.; Champagne, D. E.; Walker, F. A. Science 1993, 260, 539. (181) Walker, F. A. J. Inorg. Biochem. 2005, 99, 216. (182) Lehnert, N.; Berto, T. C.; Galinato, M. G. I.; Goodrich, L. E. In Handbook of Porphyrin Science; Kadish, K. M., Smith, K., Guilard, R., Eds.; World Scientific: Singapore, 2012; Vol. 14. (183) (a) Bird, L. E.; Ren, J.; Zhang, J.; Foxwell, N.; Hawkins, A. R.; Charles, I. G.; Stammers, D. K. Structure 2002, 10, 1687. (b) Crane, B. R.; Arvai, A. S.; Ghosh, D. K.; Wu, C.; Getzoff, E. D.; Stuehr, D. J.; Tainer, J. A. Science 1998, 279, 2121. (c) Pant, K.; Bilwes, A. M.; Adak, S.; Stuehr, D. J.; Crane, B. R. Biochemistry 2002, 41, 11071. (184) (a) Hui Bon Hoa, G.; Di Primo, C.; Dondaine, I.; Sligar, S. G.; Gunsalus, I. C.; Douzou, P. Biochemistry 1989, 28, 651. (b) Fisher, M. T.; Scarlata, S. F.; Sligar, S. G. Arch. Biochem. Biophys. 1985, 240, 456. (c) Yu, C. A.; Gunsalus, I. C. J. Biol. Chem. 1974, 249, 102. (d) Imai, Y.; Sato, R. J. Biochem. 1967, 62, 464. (185) Martinis, S. A.; Blanke, S. R.; Hager, L. P.; Sligar, S. G.; Hui Bon Hoa, G.; Rux, J. J.; Dawson, J. H. Biochemistry 1996, 35, 14530. (186) Hollenberg, P. F.; Hager, L. P. J. Biol. Chem. 1973, 248, 2630. (187) Blanke, S. R.; Martinis, S. A.; Sligar, S. G.; Hager, L. P.; Rux, J. J.; Dawson, J. H. Biochemistry 1996, 35, 14537. (188) Sabat, J.; Stuehr, D. J.; Yeh, S.-R.; Rousseau, D. L. J. Am. Chem. Soc. 2009, 131, 12186. (189) (a) Kim, V. N.; Han, J.; Siomi, M. C. Nat. Rev. Mol. Cell Biol. 2009, 10, 126. (b) Denli, A. M.; Tops, B. B. J.; Plasterk, R. H. A.; Ketting,

R. F.; Hannon, G. J. Nature 2004, 432, 231. (c) Gregory, R. I.; Yan, K.-p.; Amuthan, G.; Chendrimada, T.; Doratotaj, B.; Cooch, N.; Shiekhattar, R. Nature 2004, 432, 235. (190) Faller, M.; Matsunaga, M.; Yin, S.; Loo, J. A.; Guo, F. Nat. Struct. Mol. Biol. 2007, 14, 23. (191) Sono, M.; Dawson, J. H.; Hager, L. P. J. Biol. Chem. 1984, 259, 13209. (192) Barr, I.; Smith, A. T.; Chen, Y.; Senturia, R.; Burstyn, J. N.; Guo, F. Proc. Natl. Acad. Sci. U.S.A. 2012, 109, 1919. (193) Yang, F.; Xia, X.; Lei, H.-Y.; Wang, E.-D. J. Biol. Chem. 2010, 285, 39437. (194) Hu, R.-G.; Wang, H.; Xia, Z.; Varshavsky, A. Proc. Natl. Acad. Sci. U.S.A. 2008, 105, 76. (195) Gotoh, S.; Ohgari, Y.; Nakamura, T.; Osumi, T.; Taketani, S. Gene 2008, 423, 207. (196) (a) Levicán, G.; Katz, A.; de Armas, M.; Núñez, H.; Orellana, O. Proc. Natl. Acad. Sci. U.S.A. 2007, 104, 3135. (b) Katz, A.; Banerjee, R.; de Armas, M.; Ibba, M.; Orellana, O. Biochem. Biophys. Res. Commun. 2010, 398, 51. (197) (a) Jones, A. M.; Elliott, T. FEMS Microbiol. Lett. 2010, 307, 41. (b) de Armas-Ricard, M.; Levicán, G.; Katz, A.; Moser, J.; Jahn, D.; Orellana, O. Biochem. Biophys. Res. Commun. 2011, 405, 134. (198) Yin, L.; Dragnea, V.; Bauer, C. E. J. Biol. Chem. 2012, 287, 13850.

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DOI: 10.1021/cr500056m Chem. Rev. 2015, 115, 2532−2558