Inhibition and Cofactor Targeting of Hypoxia-Sensing Proteins

University of Massachusetts Amherst ScholarWorks@UMass Amherst

Doctoral Dissertations Dissertations and Theses

August 2015

Cornelius Y. Taabazuing University of Massachusetts Amherst

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Recommended Citation Taabazuing, Cornelius Y., "Inhibition and Cofactor Targeting of Hypoxia-Sensing Proteins" (2015). Doctoral Dissertations. 409. https://doi.org/10.7275/6956835.0 https://scholarworks.umass.edu/dissertations_2/409

This Open Access Dissertation is brought to you for free and open access by the Dissertations and Theses at ScholarWorks@UMass Amherst. It has been accepted for inclusion in Doctoral Dissertations by an authorized administrator of ScholarWorks@UMass Amherst. For more information, please contact [email protected]. INHIBITION AND COFACTOR TARGETING OF HYPOXIA-SENSING PROTEINS

A Dissertation Presented

CORNELIUS TAABAZUING

Submitted to the Graduate School of the University of Massachusetts Amherst in partial fulfillment of the requirements for the degree of

DOCTOR OF PHILOSOPHY

May 2015

Department of Chemistry

INHIBITION AND COFACTOR TARGETING OF HYPOXIA-SENSING PROTEINS

A Dissertation Presented

CORNELIUS TAABAZUING

Approved as to style and content by:

______

Michael J. Knapp, Chair

______

Michael J. Maroney, Member

______

Nathan A. Schnarr, Member

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Scott C. Garman, Outside Member

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Craig T. Martin, Department Head

Department of Chemistry

DEDICATION

To my mom, Barbara Soonyime, thank you for the sacrifices you have made to make it possible to pursue my dreams.

To my wife Rachelle Taabazuing, thank you for providing me with your love, motivation, and patience.

To the Taabazuings, thank you for your constant support and inspiration.

ACKNOWLEDGMENTS

I would especially like to thank my advisor, Professor Michael J. Knapp for his mentorship and contributions to my research. Thank you for training me well and allowing me to explore my interests.

I would like to thank my undergraduate thesis advisor, Professor Nathan Schnarr, who is now one of my committee members. Thank you for training me as an undergrad and for getting me excited about research.

I would also like to thank the rest of my committee members, Professors Michael Maroney and Scott Garman for their valuable insight and guidance on coordination chemistry and crystallography.

Thank you to the Hardy lab, especially Scott Eron and Derek Macpherson for all their help when I was getting started with crystallography. Thank you Professor Hardy for letting me tag along on the beam trips.

Thank you Heidi for all your help with XAS and fitting all that data.

Thank you Professor Justin Fermann for performing the calculations.

Thank you to past Knapp lab members Dr. Evren Saban and Dr. Shannon Coates Flagg for training me.

Thank you Dr. John Hangasky for all your work with the D201X mutants and valuable discussions about my research.

Thank you Tina for all your Mass Spec analysis work and great discussions.

Thank you Vanessa for purifying D201G, helping with the undergrads, and proof reading this document.

Thanks to all the undergrads who I’ve worked with, Jackie, Kate, Jack, Ben, and Mike.

Thanks to all my mentors and friends who have helped me keep my sanity throughout this incredible journey.

Lastly, thank you Tina Johnson for helping me format this document.

ABSTRACT

INHIBITION AND COFACTOR TARGETING OF HYPOXIA-SENSING PROTEINS

MAY 2015

CORNELIUS TAABAZUING, B.S., UNIVERSITY OF MASSACHUSETTS

AMHERST Ph.D., UNIVERSITY OF MASSACHUSETTS AMHERST

Directed by: Professor Michael J. Knapp

Hypoxia Inducible Factor (HIF) is a transcription activator considered to be the main regulator of O2 homeostasis in humans. The transcriptional ability of HIF is regulated by the Fe2+/αKG-dependent enzyme, Factor Inhibiting HIF (FIH). FIH uses molecular oxygen to catalyze hydroxylation of an asparagine residue (Asn803) in the C- terminal transactivation domain (CTAD) of the HIFα subunit, abrogating HIF target gene expression. The mechanism of FIH and other αKG-dependent oxygenases involves the ordered sequential binding of αKG, substrate, and O2, which becomes activated to form a reactive ferryl intermediate that hydroxylates the substrate. The key to understanding

FIH’s function as an oxygen sensor, and the chemistry of the broader class of enzymes, is to elucidate steps coincident with O2 binding and activation. These steps are shared amongst this class of enzymes and are the least understood.

In order to understand structural factors that influence O2 binding and reactivity, we solved the crystal structure of FIH in the presence of NO, an O2 mimic. Our data suggests that the sterics of the target asparagine residue modestly influences O2 binding, but plays a major role in stimulating O2 reactivity by orienting bound gas productively for

vi decarboxylation chemistry. The increased O2 reactivity in the presence of CTAD would ensure tight coupling of O2 binding and activation to hydroxylation. The reorientation of bound O2 may be the mechanism utilized by the broader class of αKG oxygenases to activate molecular oxygen.

The active site of the Fe2+/αKG-dependent enzymes is highly conserved. The Fe cofactor is typically ligated by 2 His residues and 1 carboxylate residue. A class of αKG- dependent enzymes that catalyze halogenation chemistry ligate the Fe cofactor with only

2 His residues, and bind a halide ion in place of the carboxylate ligand. In order to understand the structural factors that control the incorporation of alternative ligands such as halides, we mutated Asp201 in FIH to Glu (D201E), and Gly (D201G). The D201G variant activity was dependent on chloride, and the crystal structure suggested that chloride was bound at the active site. XAS data indicated that chloride was bound, implicating D201 in controlling ligand access. Other non-native ligands such as azide, and cyanate were able to access the active site Fe cofactor of D201G, consistent with relaxed ligand selectivity upon mutation of the facial triad carboxylate. The identification of exogenous ligands targeting the Fe cofactor raises the possibility that FIH can be engineered for alternative rebound chemistry.

Recently, there have been many reports of “gasotransmitters” such as NO and

H2S impacting HIF controlled gene expression in cells, suggesting that the HIF hydroxylases are potential physiological targets. We tested the effect of H2S on the activity of FIH. Our data suggests that H2S binds to the Fe cofactor of FIH to inhibit activity. H2S was also demonstrated to reduce inactive ferric FIH back to the ferrous

vii state, restoring activity. The dual regulation of FIH activity provides a rationale for some of the observed physiological responses to H2S treatment.

Taken together, our data suggests that the active site of FIH relies on several structural motifs to direct chemistry. The active site carboxylate controls the access of O2 and other ligands to the Fe cofactor. The target asparagine residue stimulates O2 reactivity by provided the proper steric environment to orient O2 productively for chemistry. However, certain ligands such as H2S are able to access the Fe cofactor to inhibit chemistry, implicating them in the hypoxia sensing pathway in cells.

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TABLE OF CONTENTS

Page

ACKNOWLEDGMENTS ...... v

ABSTRACT ...... vi

LIST OF TABLES ...... xii

LIST OF FIGURES ...... xiii

LIST OF SCHEMES...... xvii

LIST OF ABBREVIATIONS ...... xviii

CHAPTER

1. AN OVERVIEW OF OXYGEN SENSING AND THE KEY PLAYERS ...... 1

1.1 Introduction ...... 1 1.2 Acute Hypoxia Sensing by Mammalian Tissue ...... 2

1.2.1 Neuroepithelial Bodies (NEBs) and Type 1 Glomus Cells ...... 2 1.2.2 Smooth Muslce Cells (SMCs) ...... 4 1.2.3 Heme Oxygenase (HO) ...... 5

1.2.4 H2S as an O2 Sensor ...... 7

1.4 Hypoxia Inducible Factor (HIF) and the HIF Hydroxylases (FIH and PHD) ...... 12 1.5 Conclusion ...... 19 1.6 References ...... 20

2. CRYSTAL STRUCTURE OF THE HYPOXIA SENSING ENZYME FIH COMPLEXED TO FE, NOG, AND NO ...... 28

2.1 Introduction ...... 28 2.2 Materials and Methods ...... 30

2.2.1 Protein Purification ...... 30 2.2.2 CTAD Purification ...... 30 2.2.3 Crystallography and Data Refinement ...... 31 2.2.4 Electronic Absorption Spectroscopy ...... 32 2.2.5 EPR Spectroscopy ...... 33 2.2.6 DFT Calculations ...... 34

2.3 Results ...... 34

2.3.1 Crystal Structure of FIH in Complex with Fe and NO ...... 34 2.3.2 Electronic Absorption Spectroscopy ...... 37 2.3.3 Electron Paramagnetic Resonance Spectroscopy ...... 40 2.3.4 DFT Calculations of FIH-{Fe-NO}7 ...... 42

2.4 Discussion ...... 46

2.4.1 CTAD Binding Has a Modest Effect on Gas Affinity ...... 47 2.4.2 CTAD Binding Introduces Steric Barriers that Reorient Gas for Productive Reaction ...... 48 2.4.3 The Gas Binding Pocket of FIH ...... 51

2.5 Conclusion ...... 54 2.6 Appendix ...... 55

2.6.1 Supplemental ...... 55 2.6.2 DEANO Decomposition in 50 mM Hepes pH 7.00 ...... 55

2.7 References ...... 58

3. CONTRIBUTIONS OF THE FACIAL TRIAD CARBOXYLATE TO LIGAND ACCESS IN FACTOR INHIBITING HIF (FIH) ...... 64

3.1 Introduction ...... 64 3.2 Materials and Methonds ...... 67

3.2.1 Protein Expression and Purification ...... 67 3.2.2. CTAD Purification ...... 67 3.2.3 Crystallography ...... 68 3.2.4 UV-Vis Spectroscopy ...... 69 3.2.5 Initial Screen and Dose Response Assays ...... 69 3.2.6 X-ray Absorption Spectroscopy Sample Preparation ...... 70 3.2.7 X-ray Absorption Spectroscopy Data Collection and Analysis ...... 71

3.3 Results ...... 73

3.3.1 Initial Screen and Dose Response Assays ...... 73

3.3.2 UV-Vis Absorption Spectroscopy ...... 76 3.3.3 Crystal Structure of (Fe+NOG+Cl)D201G ...... 79 3.3.4 X-Ray Absorption Spectroscopy (XAS) ...... 83

3.4 Discussion ...... 87

3.5 Conclusion ...... 93 3.6 Appendix ...... 94

3.6.1 Supplemental...... 94

3.7 References ...... 121

4. HYDROGEN SULFIDE INHIBITION AND RESCUE OF THE HUMAN HYPOXIA SENSING ENZYME FIH ...... 128

4.1 Introduction ...... 128 4.2 Methods ...... 131

4.2.1 Protein Purification ...... 131 4.2.2 CTAD Purification ...... 131 4.2.3 Dose Response Assays ...... 131 4.2.4 EPR Spectroscopy ...... 132 4.2.5 Electronic Absorption Spectroscopy ...... 132 4.2.6 Rescue Assays ...... 133 4.2.7 Dynamic Light Scattering (DLS) ...... 134

4.3 Results ...... 134 4.4 Discussion ...... 143 4.5 Supplemental Figures...... 150 4.6 References ...... 152

BIBLIOGRAPHY ...... 160

LIST OF TABLES

Table Page

2.1: Summary of EPR data and NO dissociation constants ...... 42

2.2: Comparison of calculated and experimental geometric parameters for FIH- {Fe-NO}7 and related enzymes ...... 46

2.3: Data collection and refinement statistics. Values in parentheses represent the highest resolution shell ...... 56

3.1: Summary of IC50 and electronic absorption spectroscopy data...... 78

3.2: EXAFS Analysis of (Fe+NOG+Cl)D201G ...... 86

3.3: Data collection and refinement statistics for D201G. Values in parentheses represent the highest resolution shell ...... 94

3.4: EXAFS Analysis of (Fe+NOG+Cl)D201G with 1.75 Cl ...... 95

3.5: Table of EXAFS fits...... 96

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LIST OF FIGURES

Figure Page

1.1: Proposed mechanism for ion channel O2 sensing in NEBs. NADPH oxidase - mediates the hypoxic response in NEBs via O2 or H2O2...... 3

1.2: Acute hypoxia sensing in mammalian tissues. Acute hypoxia causes inhibition of K+ channels in type 1 glomus cells and NEBs. The depolarization increases intracellular [Ca2+], leading to neurotransmitter release and improved ventilation. Acute hypoxia causes KATP channels to open in systemic SMCs, inhibiting calcium influx and causing vasodilation. Inhibition of K+ channels in pulmonary artery SMCs causes depolarization and calcium influx that results in vasoconstriction ...... 5

1.3: H2S production and oxidation in the cytosol and mitochondria. Key compounds are cysteine (Cys), homocysteine (hCys), methionine (Met), and glutathione persulfide (GSSH)...... 9

1.4: Regulation of HIF-1α by FIH and PHD2. Posttranslational regulation of HIF-1α by FIH and PHD2 control HIF-1α transcriptional activity and stability under normoxic conditions. During hypoxia, HIF-1α forms a transcription complex with HIF-1β and p300, and initiates target gene expression ...... 15

2.1: Structure of (Fe+αKG)FIH and (Fe+NOG+NO)FIH. Iron is shown in magenta, water in red. (A) View of the (Fe+αKG)FIH active site with the 2Fo – Fc map contoured to 1σ. (B) (Fe+NOG+NO)FIH active site and the 2Fo – Fc map contoured to 1σ. (C) (Fe+NOG+NO)FIH active site with the Fo(Fe+NOG+NO)FIH – Fo(Fe+αKG)FIH map contoured to 1σ. (D) (Fe+NOG+NO)FIH active site measurements. PEG is colored yellow ...... 36

2.2: Electronic absorption spectra of FIH. FIH (250 µM), FeSO4 (250 µM), αKG (500 µM), DEANO (2 mM) and CTAD or CTADN803A (500 µM) at 23°C in anaerobic Hepes (50 mM pH 7.00)...... 38

2.3: Binding isotherm of (Fe+NO+αKG)FIH, (Fe+NO+αKG)FIH/N803A and (Fe+NO+αKG)FIH/CTAD. FIH (0.25 mM), FeSO4 (0.24 mM), αKG (0.5 mM), CTAD (0.5 mM), N803A (0.5 mM) and DEANO (.013 - 2 mM)...... 39

2.4: Experimental and simulated X-Band EPR spectra of {FeNO}7 FIH complexes: (A) (Fe+αKG+NO)FIH/CTAD (B) (Fe+αKG+NO)FIH/N803A (C) (Fe+αKG+NO)FIH (D) (Fe+NO)FIH. FIH (0.10mM) , FeSO4 (0.10mM), αKG (0.50mM), CTAD (0.50mM), N803A (0.50mM), DEANO (0.50 mM), 9.624 GHz frequency, 2.0mW power, 10G modulation amplitude, 100 GHz modulation frequency, 163 ms time 3+ constant, 4K. (*) = Fe with geff = 4.30, 4.25, present for all spectra...... 41

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2.5: Rigid scanned potential energy surface for rotation about the Fe-NO axis in the FIH-{FeNO}7 complex (inside) schematic diagram showing the relative orientations of NO with respect to the C2 carbon of αKG. Minima are denoted with letters...... 44

2.6: Optimized geometries of A, B, and C used in DFT calculations as models for FIH-{FeNO}7 ...... 45

2.7: Example of the first order decay plot used to calculate the rate of DEANO decomposition in 50 mM Hepes pH 7.00 at 23°C...... 55

2.8: Crystal of (Fe+NOG+NO)FIH grown in the presence of DEANO used for data collection ...... 56

2.9: UV-Vis data used to construct binding isotherm (top left) (Fe+NO+αKG)FIH/CTAD (top right) (Fe+NO+αKG)FIH (bottom left) (Fe+αKG+NO)FIH/N803A: FIH (0.25 mM), αKG (0.50 mM), FeSO4 (0.25 mM), CTAD (0.50 mM), N803A (0.50 mM), DEANO (0.015 – 2 mM),) in 50 mM Hepes pH 7.00 at 23°C ...... 57

2.10: Structure of (Fe+αKG)FIH and (Fe+NOG+NO)FIH. (A) Overlay of (Fe+αKG)FIH (green) and (Fe+NOG+NO)FIH (cyan) showing the conserved β-strand jelly roll core...... 57

3.1: Surface representation of FIH in the absence of substrate showing the solvent exposed active site (PDB: 1H2N, 3P3P, 1H2L) ...... 66

3.2: Initial inhibitor screen for FIH (top left) and the facial triad D201E (top right) and D201G (bottom) variants. Assays were performed in 50 mM Hepes containing 100 mM NaCl at 37°C; FIH (0.5 µM), D201G (2.2 µM), D201E (5 µM), αKG (500 μM), FeSO4 (25 μM), CTAD (100 μM), Ascorbate (2 mM). All inhibitors were 2 mM except for CO, which was 750 µM...... 74

3.3: Dose response assays varying NO (left) and N3 (right) of FIH and the D201E and D201G variants. Assays were performed in 50 mM Hepes containing 100 mM NaCl at 37°C; FIH (0.5 µM), D201G (2.2 µM), D201E (5 µM), αKG (500 μM), FeSO4 (25 μM), CTAD (100 μM), Ascorbate (2 mM), NO - (0 – 0.8 mM) N3 (0 – 5 mM)...... 75

3.4: Anaerobic UV-Vis spectra showing the MLCT for WT-FIH and the D201 variants in 50 mM Hepes pH 7.00 (100 mM NaCl). (A) WT-FIH, D201E, - - and D201G treated with NO. (B) D201G treated with N3 and OCN . (C) - - - D201E treated with N3 . (D) WT-FIH treated with N3 and OCN . FIH (400 uM), D201E (400 µM), D201G (400 µM), FeSO4 (375 µM), αKG (400 - - µM), NO (2 mM), OCN ( 2mM), N3 (2mM)...... 77

xiv

3.5: Crystal structure of (Fe+NOG+Cl)D201G. (A) The active site 2Fo – Fc map contoured to 1σ. (B) Active site of D201G showing the metal chloride bond lengths. (C) The 2Fo – Fc map contoured to 1σ showing the second coordination sphere residues that help position the target residue. (D) D201G active site second coordination sphere distances to N803...... 80

3.6: Structure of (Fe+NOG+Cl)D201G (green), (Zn+αKG)D201G (cyan), and (Fe+αKG+Cl)SyrB2 (white). (A) (Zn+αKG)D201G active site distances. (B) Comparison of (Zn+αKG)D201G (cyan) and (Fe+NOG+Cl)D201G (green). (C) Alternate view of (Zn+αKG)D201G (cyan) and (Fe+NOG+Cl)D201G (green) overlaid. (D) (Fe+αKG+Cl)SyrB2 active site distances. (E) Overlay of (Fe+αKG+Cl)SyrB2 (white) and (Fe+NOG+Cl)D201G (green). The Fe and Cl- for SyrB2 are colored grey and green respectively and the Fe and Cl- for D201G are colored magenta and yellow respectively. Waters are colored red (F) Overlay of (Fe+αKG+Cl)SyrB2 (white) and (Zn+αKG)D201G (cyan). The Fe and Cl- for SyrB2 are colored grey and green respectively and the Zn for D201G is colored light grey. Waters are colored red and the ones derived from SyrB2 are labelled...... 82

3.7: Fe K-edge XANES spectrum of (Fe+NOG+Cl)D201G in 50mM Hepes pH 7.00...... 85

3.8: EXAFS analysis of (Fe+NOG+Cl)D201G in 50mM Hepes pH 7.00. Fourier transform (FT) XAS data with data in black and fit in red (see fit details in Table 3.2). Inset: Unfiltered k3-weighted EXAFS spectra for the fit (red) and data (black). Fits were performed in r-space (Δk=2 – 14Å-1; Δk=1 – 4.0Å) as described in materials and methods...... 87

3.9: EXAFS analysis of (Fe+NOG+Cl)D201G for 1 and 1.75 Cl in 50mM Hepes pH 7.00. Fourier transform (FT) window of 2 – 14 Å-1 with data in black and fits in red (1Cl) and blue (1.75 Cl). Fit details for 1 Cl are in Table 3.2 and details for 1.75 Cl are in Table 3.4). Inset: Unfiltered k3-weighted EXAFS spectra for the fits and data. Fits were performed in r-space (Δk=2 – 14Å-1; Δk=1 – 4.0Å) as described in materials and methods...... 95

4.1: pH dependent inhibition of FIH (0.5 µM) in 50 mM Hepes pH 6.00 – 8.00 at 37°C; αKG (25 μM), FeSO4 (25 μM), CTAD (100 μM), sulfide (0 – 520 μM), Ascorbate (2 mM). The inset is an example of a dose response curve at pH 7.40...... 135

4.2: (Left) Steady state kinetics of FIH in the presence of sulfide (Right) Regression analysis of steady state kinetics data. Assays were performed at 37°C with FIH (0.5 µM), ascorbate (2 mM), αKG (500 µM), FeSO4 (25 µM), and CTAD (24 – 288 µM) in 50 mM Hepes pH 7.40...... 137

4.3: Electronic absorption spectra of aerobic and anaerobic (Fe+αKG)FIH treated with sulfide. (black) Anaerobic (red) Aerobic. FIH (1 mM), FeSO4 (1 mM), αKG (2 mM), and sulfide (2 mM)...... 138

4.4: Electronic absorption spectra of (Fe+NOG+Sulfide)FIH in 50 mM Hepes pH 7.40. (left) Spectra in the absence and presence of sulfide before and after purification with SEC; FIH (467 µM), Fe2+ (467 µM), NOG (2 mM), sulfide (0, 467 µM). After SEC, [FIH] = 424 µM. (Right) Difference spectra calculated by subtracting the spectrum of (Fe+NOG)FIH...... 139

4.5: DLS of (Fe+αKG)FIH and (Fe+αKG+Sulfide)FIH. FIH (50 μM), FeSO4 (49 μM), αKG (250 μM), Sulfide (50 µM) in 50 mM Hepes pH 7.40...... 140

4.6: Rescue of auto-hydroxylated FIH activity by reductants. FIH (4.9 μM), Fe2+ (4.9 μM), αKG (185 μM), CTAD (100 µM), reductant (20 µM) in 50 mM Hepes pH 7.40...... 141

4.7: X-Band EPR of auto hydroxylated FIH samples. FIH (100 μM), FeSO4 (100 μM), αKG (500 μM), Sulfide (0 – 1 mM) in aerobic 50 mM Hepes pH 7.40. 9.609 GHz, 6.0 mW power, 10 G modulation amplitude, 100 GHz, 77 K...... 142

4.8: Concentration dependence for sulfide rescue of auto-hydroxylated FIH activity. FIH (5 μM), FeSO4 (5 μM), αKG (185 μM), Sulfide (0 - 600 µM) in 50 mM Hepes pH 7.40. EC50 = 333 ± 32 µM...... 143

4.9: IC50 measurement at high αKG levels FIH (0.5 μM) in 50 mM Hepes pH 7.00 at 37°C; αKG (500 μM), FeSO4 (25 μM), CTAD (100 μM), Sulfide (0 – 120 μM), Ascorbate (2 mM)...... 151

4.10: Anaerobic (black) and aerobic (red) UV-vis samples of Fe (400 µM), NOG/αKG (2 mM), and HS- (400 µM) in 50 mM Hepes pH 7.40...... 151

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LIST OF SCHEMES Scheme Page

1.1: The consensus mechanism of αKG-dependent oxygenases ...... 14

2.1: Proposed mechanism for coupling in FIH ...... 29

4.1: Proposed consensus mechanism of FIH and αKG-dependent oxygenases ...... 129

4.2: Kinetic mechanism of FIH...... 136

4.3: Proposed model for sulfide interaction with FIH ...... 149

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LIST OF ABBREVIATIONS

NO – Nitric oxide

DEANO – Diethylammonium (Z)-1-(N,N-diethylamino)diazen-1-ium-1,2-diolate

αKG – α-Ketoglutarate

HIF – Hypoxia Inducible Factor

FIH – Factor Inhibiting HIF

CTAD – C-terminal Transactivation Domain

CAS – Clavaminate synthase

TauD – Taurine dioxygenase

HPPD – 4-Hydroxyphenylpyruvate dioxygenase

MCD – Magnetic Circular Dichroism

CD – Circular Dichroism

NOG – N-oxalylglycine

ESEEM – Electron Spin Echo Envelope Modulation

5C – 5 Coordinate

6C – 6 Coordinate

– H2S/HS – Hydrogen sulfide

PHD – Prolyl Hydroxylase

PH4 – Prolyl 4 Hydroxylase

CcO – Cytochrome c oxidase

SOD – Superoxide dismutase

EPR – Electron Paramagnetic Resonance

MLCT – Metal to Ligand Charge Transfer

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LMCT - Ligand to Metal Charge Transfer

MALDI-MS – Matrix Assisted Laser Desorption Ionization Mass Spectrometry

SEC – Size Exclusion Chromatography

DLS – Dynamic Light Scattering

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CHAPTER 1

AN OVERVIEW OF OXYGEN SENSING AND THE KEY PLAYERS

This chapter has previously been published as: Taabazuing, C. Y.; Hangasky, J. A.; Knapp, M. J. Oxygen Sensing Strategies in Mammals and Bacteria. J. Inorg. Biochem. 2014, 133, 63–72.

1.1 Introduction

Oxygen is the terminal electron acceptor used during aerobic respiration, making it essential for the survival of nearly all organisms. During respiration, the reduction of

O2 into H2O is coupled to the production of cellular energy in the form of ATP. Too little oxygen results in inefficient respiration and a switchover to anaerobic metabolism or cell death if the proper adaptive response is not elicited. Conversely, too much oxygen can lead to oxidative stress and the production of reactive oxygen species (ROS) that can damage the cellular machinery and cause cellular death. As a result of the necessity to regulate O2 availability, organisms have evolved complex and elegant systems for sensing and maintaining O2 homeostasis. O2 sensing pathways typically utilize metalloproteins to either reversibly bind O2 or to catalyze irreversible O2-dependent reactions, leading to transcriptional and metabolic changes in response to varied pO2.

The atmosphere is composed of ~ 21 % oxygen, leading to an appreciable

1 concentration of dissolved O2 in water under ambient conditions (288 µM at 20 °C).

However, due to the limits of diffusion and metabolic demands, intracellular O2 levels are often less than 10% of this ambient value in mammalian tissues. Further deviations from this value are possible depending on the tissue, cell type, and developmental stage of the

2 mammal. It is estimated that O2 concentrations ([O2]) are ~5% of the ambient level in typical adult tissue.3 As a result, the body relies on number of finely tuned responses to variations in [O2] that work over a narrow range of pO2.

In relation to biology, environments with sufficient [O2] are considered to be normoxic while conditions in which [O2] is deficient are considered hypoxic. Hypoxia has a significant effect on the pathogenesis of many human diseases, including chronic heart and lung disease, myocardial ischemia, pulmonary hypertension, and cancer, all of which are major contributors to the mortality rate worldwide.3–7 Therefore, a comprehensive understanding of O2 sensing can potentially improve therapeutics for treating diseases related to poor or over oxygenation.

1.2 Acute Hypoxia Sensing by Mammalian Tissue

Mammals follow distinct sensing strategies for acute hypoxia (time scale of minutes) and chronic hypoxia (time scale of hours to days). Acute hypoxia sensing starts at the tissue level with specialized cells responding to changes in pO2 through the effects of potassium and calcium ion channels on the membrane potential of the cells.2,8,9 The two dominant physiological responses to acute hypoxia are hypoxic systemic vasodilation and hypoxic pulmonary vasoconstriction, which change vascular tension to modify blood flow. Vasodilation is a response that increases the perfusion of blood to the affected tissue while vasoconstriction is a regulatory mechanism that balances perfusion to ventilation. Although these acute responses are crucial for tissues experiencing hypoxia, and are thought to be mediated within the cell by ion channel signaling pathways, the specific molecular mechanisms linking changes in pO2 to ion channel activity are not well understood.

1.2.1 Neuroepithelial Bodies (NEBs) and Type 1 Glomus Cells

The first step in acute hypoxia response is facilitated by specific chemosensory cells termed neuroepithelial bodies (NEBs), and type 1 glomus cells of the carotid body.

NEBs are the first responders to pO2 as they line the mucosa of mammalian airways.

NADPH oxidase (NOX) has been experimentally demonstrated to mediate the hypoxic response in NEBs via ROS generation.10 Activation of NOX by protein kinase C (PKC)

- + leads to superoxide (O2 ) production, which may be the signal to close K channels.

- 11 Counter intuitively, hypoxia results in increased production of O2 by NOX. Whether

- - the chemical signal is O2 or H2O2 is ambiguous as O2 can dismutate to H2O2 and direct

+ 12 addition of H2O2 boosts the whole cell K channel current in NEBs (Figure 1.1). The shift in membrane potential due to K+ channel closure causes an influx of Ca2+, triggering neurotransmitter release (serotonin and neuropeptides) from NEBs (Figure 1.2).13

Although activation of NOX by PKC led to hypoxia sensitive K+ channel closure in the immortalized model NEB H146 cell line14 NOX inhibitors failed to fully suppress this response, suggesting that multiple mechanisms may be utilized by these cells to sense

15 O2.

Figure 1.1: Proposed mechanism for ion channel O2 sensing in NEBs. NADPH oxidase - mediates the hypoxic response in NEBs via O2 or H2O2.

Type 1 glomus cells are specialized chemoreceptor cells belonging to the carotid body, and are similar to NEBs. In type 1 glomus cells, hypoxia results in closure of K+ channels, which depolarizes the cellular membrane. The resultant membrane depolarization triggers an influx of Ca2+ and the release of neurotransmitters (dopamine) to the carotid-sinus nerve that relays the information to the central nervous system and stimulates ventilation (Figure 1.2).9,16 In contrast to the role NOX plays in NEBs, NOX

3 suppresses K+ channel closure in type 1 glomus cells,11,17 indicating that the mechanism for O2-sensing is subtly tweaked from tissue to tissue. As the same ion channel types can be either potentiated or inhibited depending on cell types, this would allow individual tissues to fine tune the hypoxic response.

1.2.2 Smooth Muscle Cells (SMCs)

Smooth muscle cells (SMCs) work in concert with NEBs and type 1 glomus cells to facilitate tissue-specific responses to acute hypoxia. In pulmonary artery SMCs, acute hypoxia closes the K+ channels, which leads to an increase in intracellular [Ca2+], triggering neurotransmitter release and vasoconstriction.18 In contrast, the hypoxic response in many other tissues is vasodilation due to the function of systemic SMCs. In

+ systemic SMCs, hypoxia causes ATP sensitive K (KATP) channels to open and hyperpolarize the membrane, inhibiting Ca2+ influx and causing muscle cell relaxation.18,19 These findings and the observation that inhibiting oxidative phosphorylation drastically influences O2 sensing in NEBs and SMCs led to the model that the energy demands of the cell ultimately govern the response to hypoxia, however the scientific literature contains conflicting data regarding this model. For example, myocite relaxation can occur at pO2 levels that don’t compromise energy metabolism, suggesting that decreased energy metabolism is not a requirement for hypoxia sensing by ion channels.8 For a more in depth discussion of this evidence the reader is directed to more focused reviews on that subject.2,8

Figure 1.2: Acute hypoxia sensing in mammalian tissues. Acute hypoxia causes inhibition of K+ channels in type 1 glomus cells and NEBs. The depolarization increases intracellular [Ca2+], leading to neurotransmitter release and improved ventilation. Acute hypoxia causes KATP channels to open in systemic SMCs, inhibiting calcium influx and causing vasodilation. Inhibition of K+ channels in pulmonary artery SMCs causes depolarization and calcium influx that results in vasoconstriction.

Adaptation to hypoxia may involve vasodilation or vasoconstriction, depending on the affected tissue; however, both result from changes in membrane channels that cause myocite contraction or relaxation. In this manner, adaptation to acute hypoxia relies on a carefully orchestrated response that involves neuroepithelial cells, type 1 glomus cells, and smooth muscle cells. The cellular machinery that play a role in the acute hypoxic response are just now emerging and understanding of the molecular details awaits further research.

1.2.3. Heme Oxygenase (HO)

The challenges with distinguishing O2-sensing processes from those that merely consume O2 are exemplified by heme oxygenase (HO). One isoform of HO (HO-2) has been proposed to sense hypoxia based on data from genetic knockouts in mice, but the literature is conflicting. Heme oxygenases oxidatively cleave hemes into CO and

20 biliverdin, consuming O2 and NADPH. There are three isoforms of HO identified, of which the first two have been well characterized and reported to have a high affinity for

2,21 O2 (P50(O2) values: HO-1 = 0.036 μM , HO-2 = 0.013 μM). HO-1 is an inducible heme oxygenase that is expressed in response to stresses such as hypoxia,22 whereas HO-2 is constitutively expressed and is proposed to modulate Ca2+ activated K+ channels in the

23 membranes of glomus cells. While the extremely low P50(O2) of HO-1 and -2 indicates that these enzymes will be saturated with respect to [O2] over the physiological range of

O2, and incapable of a response proportionate to changes in pO2, HO-2 has intriguing connections to cellular hypoxia responses.

HO-2 was proposed to respond to hypoxia due to the HO-2 produced CO increasing K+ channel activity.23 This model requires that hypoxia leads to decreased

[CO], thereby closing the K+ channels and leading to Ca2+ influx and neurotransmitter

24 release. While the very low P50(O2) value for HO-2 appears to eliminate a direct link between CO production and physiologically relevant changes in [O2], a number of observations suggest a connection between HO-2 and responses to hypoxia. At the

organismal level, HO-2 deficient mice were reported to be hypoxaemic (decreased pO2 in the blood) and exhibited signs of vasoconstriction activity.25 Cell-based assays showed

+ that both HO-2 activity and CO could open K channels, implicating HO-2 in the O2 sensing pathway for pulmonary artery SMCs.25,26 In contrast, other cell-based assays produced conflicting data on the role of HO-2 in glomus cells,25,27 which may be due to the use of cells from different species in these studies. These findings underscore the need to resolve the mechanisms linking hypoxia to ion channel function, in order to have a more comprehensive understanding of O2 sensing in mammalian tissue.

An important aspect of acute hypoxia sensing that remains elusive is a molecular

+ view connecting decreased pO2 with K channel activity. The proposals described above are further muddied by patch-clamp experiments indicating that K+ ion channels may sense O2 directly as there is an absence of detectable modifications to the cytosolic environment, such as changes in pH, ATP levels or [Ca2+].8 Although NOX regulates ion channels,10–12 the specific molecular players connecting NOX to K+ channels remain to

- be clarified. Whether CO, O2 , or the recently proposed H2S (see below), the chemistry underneath acute hypoxia sensing promises to be a fertile field for investigation.

1.3 H2S as an O2 Sensor

A recent proposal for hypoxia sensing in higher organisms is that hydrogen sulfide (H2S), or some other sulfur species, is the direct sensor for acute hypoxia in many tissues of higher organisms.28,29 While controversial, there are compelling correlations between O2 and H2S biochemistry, suggesting a connection between these gases. H2S elicits responses similar to those caused by hypoxia in many tissues,29 and the molecular

- players are more fully identified than for the CO and O2 models discussed above. The key features of this hypothesis are: the O2-sensitive speciation of sulfur into reduced and oxidized pools to signal changes in pO2; and the transduction of this signal by an unknown mechanism into cellular responses to hypoxia.

At a very basic level, the speciation of sulfur into reduced (H2S) and oxidized

(SOx) pools depends on the availability of O2, leading to a correlation between hypoxia

30 and elevated [H2S] within cells. While a simplified view suggests that this is due to the balance between the cytosolic metabolism of S-containing compounds to produce H2S

2- 2- and the mitochondrial oxidation of H2S to SSO3 and SO4 (Figure 1.3), the story is somewhat more complex. In particular, the distribution of various enzymes involved in

7 sulfur metabolism may be more varied than previously thought. As oxidation to form

2- 2- SSO3 and SO4 are slowed under conditions of low pO2, the reduced sulfur pool increases under hypoxic conditions. But other factors, such as H2S consumption by

31–33 34 ROS and H2S production promoted by elevated glutathione levels, indicate that H2S levels do not respond solely to changes in pO2. This interplay between various redox pools, pO2 and [H2S], combined with the challenges in measuring different sulfur species,31,35,36 makes it difficult to establish a clear causal link between hypoxia and elevated levels of reduced sulfur species.

A simplified view of the production of H2S centers on the transsulfuration

31,32,37,38 pathway and on cysteine catabolism. In the transsulfuration pathway, H2S is liberated from cysteine, homocysteine, and cystathionine by the pyridoxal phosphate

(PLP)-dependent enzymes cystathione β-synthase (CBS) and cystathione γ-lyase (CSE), which are typically cytosolic enzymes.39 However, data suggests that CBS and CSE translocate to mitochondria under cellular stress, which may account for cysteine

40,41 metabolism within the mitochondria. H2S is also produced from Cys by the sequential action of the enzyme cysteine aminotransferase (CAT), which uses αKG as a co-substrate, and mercaptopyruvate sulfotransferase (MST); while CAT and MST are predominantly cytosolic, MST is also found within the mitochondrion.42 Whereas

2- oxidation of mitochondrial H2S to SO4 leads to excretion, the fate of cytosolic H2S is less clear (Figure 1.3). Connections between the metabolism of glutathione (GSH),

31 cysteine, and H2S (as reviewed by Gojon ), imply that cytosolic H2S equivalents are fed into the mitochondria for oxidation. Indeed, it was recently proposed that H2S oxidation

43 rather than H2S production is the crucial step in hypoxia sensing.

Mitochondrial enzymes oxidize H2S within the mitochondrial matrix, connecting

37 H2S catabolism to respiration through the quinone pool in the inner membrane. The membrane enzyme sulfide quino-oxidoreductase (SQR) oxidizes S2- to S0 using the quinone pool, transferring the S0 equivalent to glutathione to form a persulfide (GSSH) and sending electrons into the respiratory redox chain via catechols. The S0 equivalent is then oxidized to the S4+ oxidation state (sulfite) by the enzyme persulfide dioxygenase

(ETHE1)37,38,44 with subsequent conversion either by sulfite oxidase to sulfate, or by rhodanese, which combines a S0 equivalent (RSSH) with sulfite to form thiosulfate

2- (SSO3 ). The non-enzymatic reduction of thiosulfate by GSH has been shown to rapidly produce H2S under hypoxic conditions using purified reagents or mammalian tissues, suggesting that this may be a reaction that will rapidly increase the H2S level under acute hypoxia.34

Figure 1.3: H2S production and oxidation in the cytosol and mitochondria. Key compounds are cysteine (Cys), homocysteine (hCys), methionine (Met), and glutathione persulfide (GSSH).

Cells can both produce and consume H2S in proportion to varied pO2, with GSH, thiosulfate, and sulfite as key players in H2S metabolism. Notably, the rate of H2S

30 consumption decreases when [O2] drops below ~ 20 µM in bovine tissues, suggesting that acute hypoxia leads to elevated H2S. Thiosulfate has been proposed to serve as a pool for the rapid reductive release of H2S by reducing agents such as glutathione (GSH),

34 which could lead to the production of H2S under hypoxic conditions. The mitochondrial enzyme persulfide dioxygenase (ETHE1) catalyzes a key O2-consuming step in the

44 catabolism of H2S, producing sulfite. Precisely how the concentrations of these molecular species vary with pO2 is crucial for the H2S-centered model for acute hypoxia sensing. Intriguingly, ETHE1 is a non-heme Fe(II)-dependent oxygenase,44 which binds to the Fe(II) cofactor using a His2Asp facial triad; this is similar to the cofactor structure for the well-established hypoxia sensors FIH and PHD, which control the transcriptional activity of HIF, the hypoxia inducible factor (see below). If H2S is a transient hypoxia sensor, it would be a remarkable development if yet another non-heme Fe(II) dioxygenase were to sense O2 in cells. This, however, remains an open question at present.

Exogenously administered H2S (or N-acetyl cysteine) mimics the hypoxic response in a variety of tissues, such as vasoconstriction or vasodilation depending on the

30 organism. However, the responses to elevated [H2S] and low [O2] may be complementary rather than identical, with much cross-talk between the two stressors.32

While it has been asserted that this indicates that H2S and hypoxia are mediated by the

28 same effector pathways within cells, the molecular pathways linking H2S to tissue responses have not yet been identified for testing. An intriguing study from 2011 found that while HIF-1 was required for the transcriptional effects of chronic H2S in C. elegans, the identity of genes regulated by chronic H2S administration were distinct from those

10 regulated by hypoxia.45 It is possible that better discrimination of responses to acute hypoxia from those due to chronic hypoxia would help to clarify the specific effector pathways induced by H2S.

Although the proposal that H2S levels report on acute hypoxia is consistent with a number of correlations, the molecular basis for signal transduction is unclear. Two key questions moving forward are: How is sulfur speciation transduced as a signal? How does the mitochondrial sulfur pool connect with the cytosolic sulfur pool? It has been suggested that post-translational protein modification via sulfhydration is the molecular

46 basis for signaling increased [H2S]. In sulfhydration, a protein-bound persulfide (R-

SSH) is formed by one of several possible mechanisms.31,47 There is an oxidant-

+ promoted reaction (RSH + H2S → RSSH + 2 H + 2 e-), or sulfane transfer from an endogenous persulfide (GSSH + RSH → GSH + RSSH). Notably, this signaling mode

0 47 suggests that sulfane sulfur (S ) may be the sulfur signaling agent rather than H2S. We were unable to find any tests of sulfhydration connected to hypoxia sensing at present.

The connections between cytosolic and mitochondrial sulfur speciation are not completely understood. Under some conditions the persulfide form of the cytosolic enzyme MST may be imported into the mitochondria, thereby importing S0 equivalents in limited concentrations. However, in order to excrete the sulfide formed within the cytosol, there must be bulk mitochondrial import, thereby leading to oxidation to sulfate.

Despite these gaps, the proposal that H2S senses acute hypoxia is much richer in molecular detail than the proposals connecting K+ ion channels to hypoxia via CO and

- O2 signaling.

1.4 Hypoxia Inducible Factor (HIF) and the HIF Hydroxylases (FIH and PHD)

The response to acute hypoxia is largely a matter of vascular tension to modify blood delivery to tissues, however chronic hypoxia leads to responses within cells at the transcriptional level. Hypoxia Inducible Factor (HIF) is the most important monitor of pO2 as it controls the expression of hundreds of genes over a wide range of physiological

48 O2 tensions. HIF also links acute and chronic responses to hypoxia by controlling a number of adaptations to pO2 including erythropoiesis, angiogenesis, glucose metabolism, cell proliferation, and glycolysis.3,49 Several isoforms of HIF are known, with HIF-1 being the dominant player in the human hypoxia response.

After the discovery of HIF-1 in the nuclear extracts of hypoxic cells in 1992,50 it was soon realized that HIF-1 was a heterodimeric protein consisting of α and β subunits,51 and that HIF-1α levels and transcriptional activity increased with decreasing

52 cellular O2 concentrations. Three isoforms of the HIFα transcription factor have been identified, HIF-1α -3α, which are evolutionarily conserved in all metazoans53 but absent from bacteria, yeast, and plants. While the abundance of HIF-α increase under hypoxia,49,51 HIF-1β is a constitutively expressed protein that is also known as the aryl hydrocarbon receptor nuclear translocator (ARNT). Although both HIF-1α and HIF-2α are capable of initiating transcriptional activity, HIF-1α is believed to be the key regulator of the hypoxic response as the other two isoforms have varied expression levels depending on cell types.3 HIF-3α, which is incapable of initiating hypoxia induced transcriptional activity, is not closely related to HIF-1α or HIF-2α and appears to serve no direct role in hypoxic sensing.48,54

The transcriptional activity of HIF-1α decreases with increasing pO2 due to the competition between nuclear translocation and the post-translational hydroxylation of

HIF-1α. HIF-1α accumulates during hypoxia, permitting it enough time to translocate into the nucleus. Once there, it can dimerize with HIF-1β to bind the transcriptional co- activator protein p300 at the promoter regions of genes that allow the cell to survive hypoxia. HIF-1α undergoes O2-dependent post-translational modifications that either marks it for proteasomal degradation or prevents its interaction with p300, halting HIF-1 mediated gene expression (Figure 1.4). These post-translational modifications to HIF-1α take the form of hydroxylations to specific Pro and Asn residues, the rates of which are proportional to [O2]. Consequently, the transcriptional activity of HIF-1α decreases when the hydroxylation rate exceeds the rate of nuclear localization.

The regulatory enzymes that control HIF-1 activity via posttranslational hydroxylation of asparagine and proline residues are the proteins factor inhibiting HIF

(FIH) and prolyl hydroxylase (PHD). Only one isoform of FIH is currently known, but there are 3 isoforms of PHD (PHD1-3) identified to date, of which PHD2 is recognized as

55 56 the main regulator of HIF-1α. Both PHD2 (KM(O2) = 250 μM) and FIH (KM(O2) = 90

49 μM) are O2 sensors as they have an absolute requirement for O2 in order for chemistry to occur, and their activity is proportional to variations in pO2 over the physiological range. The dual regulation of HIF-1α likely ensures strict control of the hypoxic response over a wide range of pO2 levels.

FIH and PHD2 belong to a large superfamily of non-heme Fe(II)/αKG-dependent oxygenases. FIH and PHD2 are proposed to follow an ordered sequential mechanism in which the co-substrate αKG binds first, then substrate (a domain of HIF-1α) binds second, followed by O2 (Scheme 1.1). Substrate binding stimulates O2 reactivity, which is typically attributed to the creation of an open coordination site due to aquo release.

Oxidative decarboxylation generates a high-valent ferryl intermediate that abstracts hydrogen from an un-activated carbon, followed by rebound chemistry to hydroxylate the substrate (Scheme 1.1). Although hydroxylation is the typical reaction for the αKG oxygenases, a number of different products can be found for this class of enzymes. For more comprehensive reviews on the chemistry and structural conservation of αKG oxygenases the reader is directed to recent reviews.57,58

Scheme 1.1: The consensus mechanism of αKG-dependent oxygenases

PHD2 and FIH hydroxylate specific residues in the O2-dependent degradation domain (ODD) and the C-terminal transactivation domain (CTAD) of HIF-1α respectively, which leads to decreased transcriptional activity for HIF. Hydroxylation of either of two proline residues (Pro402, Pro564) in the ODD by PHD2 is a prerequisite of

HIF-1α recognition by the von Hippel-Lindau protein (pVHL) (Figure 1.4),59 with

14 hydroxylation at Pro564 being ten-fold faster than at Pro402.60–62 pVHL is the recognition component of an E3 ubiquitin-protein ligase that targets HIF-1α for rapid proteasomal degradation after successive rounds of ubiquitinylation.63 Hydroxylation of

Asn803 in the CTAD by FIH blocks p300 from binding, stopping expression of HIF-1α

64 target genes (Figure 1.4). O2 mediated regulation of HIF-1α varies slightly between

65 different cells, depending on the O2 requirements in specific cell types.

Figure 1.4: Regulation of HIF-1α by FIH and PHD2. Posttranslational regulation of HIF- 1α by FIH and PHD2 control HIF-1α transcriptional activity and stability under normoxic conditions. During hypoxia, HIF-1α forms a transcription complex with HIF-1β and p300, and initiates target gene expression.

HIF controls the expression of several genes that help mediate the adaptive

66 22 7 55 response to fluctuating pO2, such as NOX, HO-1, GLUT-1, and PHD2 , placing it at the center of O2–sensing and homeostasis. Although the transcriptional activity of HIF is

15 linked to both acute and chronic hypoxia sensing, the true sensors are FIH and PHD2, which use molecular oxygen to hydroxylate HIF-1α. Interestingly, HIF controls the expression of PHD2, which is induced under hypoxia. As the KM(O2) for PHD2 is high in

49,56 relation to physiological pO2 within cells, PHD2 has low activity during hypoxia, but its expression would ensure that the proper balance to HIF-1α levels are restored upon reoxygenation of the cell. Whilst the critical players (FIH and PHD2) in HIF mediated pO2 sensing have been identified, key questions that remain to be answered include: How is the chemistry of O2 activation stimulated by enzyme/HIF-1α binding? Are there physiological effectors of PHD2/FIH activity, including other gases such as H2S, CO, and

NO? How can we selectively inhibit or increase PHD2/FIH activity? Resolving these questions would facilitate therapeutic targeting of these enzymes, as well as fundamental insight into the chemistry of this superfamily of non-heme Fe(II) αKG-dependent hydroxylases.

67 O2-activation by FIH and PHD2 is significantly stimulated by substrate binding, as seen for other αKG hydroxylases. This is crucial to their role as O2-sensors because

68 O2-activation in the absence of HIF leads to enzyme inactivation. The best supported model for substrate-stimulated O2-activation in these enzymes focuses on the coordination geometry of the Fe(II) cofactor. The Fe(II) switches from predominantly 6- coordinate to 5-coordinate upon substrate binding due to aquo release, as shown by electronic spectroscopy and computational results for several enzymes from the αKG-

67,69,70 dependent hydroxylase family. By creating a coordination site on Fe(II), O2 binding is then forced to follow substrate binding. Although the chemical mechanism of O-O

16 bond cleavage is unclear, back bonding from αKG to the metal likely plays a role in

69,70 stimulating O2 activation at Fe(II).

Despite the coordination changes observed under non-turnover conditions, certain features of the above model need to be refined. For example, the kinetics of ligand exchange from the Fe(II) cofactor is not known – unless aquo exchange from the Fe(II) is very slow, O2 would be able to access the metal prior to substrate binding. Further, the redox potential of the Fe(II) has been proposed to shift upon substrate binding, thereby

71 facilitating its reaction with O2. These two points suggest that substrate binding may, in fact, simply increase the equilibrium binding affinities for O2 relative to that of H2O.

Another broader question about this model that remains under study is how substrate binding alters contacts within the active site to stimulate O2 activation. Answering these questions will elucidate much about the specific chemistry involved in O-O cleavage as related to hypoxia sensing.

Recent work on the HIF hydroxylases has provided some insight into the molecular mechanism of pO2 activation by the HIF hydroxylases. The mechanical linkage between HIF-1α binding and O2-activation remains unclear in FIH, however this is central to O2 sensing. Although FIH will inactivate in vitro through an auto-oxidation

68,72 reaction in the absence of HIF-1α, HIF-1α binding stimulates the O2 reactivity of FIH many-fold. MCD and CD data of FIH in solution revealed that the Fe(II) cofactor geometry shifts from 6-coordinate to a mixture of 5/6-coordinate upon HIF-1α binding,67 in agreement with the X-ray crystal structure of HIF-1α bound FIH73 and suggesting that

74 aquo release may limit the O2 reactivity. As the consensus mechanism for this family of

58 enzymes requires that the Fe(II) be 5-coordinate prior to binding O2, the MCD data

17 showing only partial formation of the 5-coordinate Fe(II) suggests that O2 binding or activation may be sluggish in FIH. Further support for this notion comes from a suite of mechanistic data, which points to rate-limiting O2 activation for FIH under accessible

75 pO2, which would lead to a direct correlation between enzyme activity and pO2.

Notably, modeling and mutation studies demonstrate that the second coordination sphere

67,76 residues are pivotal for directing O2 reactivity in FIH. The picture that emerges is one in which HIF-1α binding induces structural changes near the Fe(II), including aquo release, which stimulates O2-activation.

PHD2 activity also appears to be limited by the rate of O2-activation, consistent with its role as an O2-sensor. Pre-steady state kinetics indicated that PHD2 reacted with

O2 very slowly when compared to other αKG oxygenases, and related freeze-quench experiments failed to isolate the putative ferryl intermediate,77 which was consistent with

O2-activation as the rate-limiting step in PHD2. Steady-state kinetics further supports

78 rate-limiting O2-activation in PHD2. In order for the HIF hydroxylases to be good O2 sensors, O2 binding or a step subsequent to O2 binding should be rate limiting as this would lead to proportionate changes in activity with pO2. The KM(O2) being higher than the physiologically relevant pO2 is also consistent with the notion of the HIF hydroxylases being O2 sensors as this would allow the rate of their chemistry be proportionate to pO2 near physiological levels.

A number of endogenously produced molecules, such as NO, CO, and H2S, are intriguing as putative effectors of the HIF hydroxylases as they may bind in place of O2 and are known to regulate pathways related to hypoxia responses. Nitric oxide (NO) regulates vascular tone, and has been shown to inhibit PHD279 and FIH,80 hinting that this

18 and other transient species could be physiological effectors of the HIF hydroxylases.

Although CO binding to the HIF hydroxylases has yet to be demonstrated, endogenous

CO has been shown to exhibit anti-apoptotic, anti-inflammatory, and protective responses

27 against hyperoxia-induced lung injury. Unpublished results show that H2S inhibits FIH

(IC50 ~ 30 µM) (Taabazuing and Knapp, unpublished) implicating H2S in the overall hypoxia response.

Methods to inhibit the HIF hydroxylases would be highly desired, as they would permit therapeutic targeting of the hypoxia sensing machinery. Although a number of synthetic compounds have been reported to inhibit these enzymes,81–83 challenges include

81 improving both selectivity and the relatively low affinity of the inhibitors (IC50 > 1µM).

Inhibitors for FIH and PHD2 are typically structural mimics of αKG, able to bind to the

Fe(II) cofactor via one or two ligation points, opening up the likelihood of cross- reactivity with other αKG oxygenases. Inclusion of a chiral center near the Fe(II) binding group improved inhibitor specificity in one study,84 suggesting that further improvements in inhibitors may be eminently achievable with further synthesis and screening efforts.

1.5 Conclusion

Acute and chronic changes in pO2 lead to distinct adaptive responses in mammals, ranging in scale from vascular changes through transcriptional changes. While several biochemical pathways for pO2 sensing are known with varied levels of detail, a number of outstanding research questions remain that focus on the chemical interactions that underlay sensing. Tissue level responses to changed pO2 appear to be mediated by ion- channels, however the molecular mechanisms for transducing pO2 into ion channel activity remain to be defined. Similarly, the proteins that control the transcriptional

19 activity of HIF through post-translational hydroxylation (FIH and PHD enzymes) are well defined, leading to questions that largely center on the chemical mechanisms of O-O bond cleavage and the identification of enzyme regulators/inhibitors. Emerging data suggests that the speciation of sulfur compounds report on hypoxia, with two large questions being the connections between cytosolic and mitochondrial sulfur pools, and the mechanisms for transducing the signal. Continuing research in these areas holds promise in treatments for diseases including ischemia and cancer, where vasculature development and anaerobic metabolism play significant roles in cell growth.

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CHAPTER 2 CRYSTAL STRUCTURE OF THE HYPOXIA SENSING ENZYME FIH COMPLEXED TO FE, NOG, AND NO

2.1 Introduction

The Fe2+/αKG-dependent enzymes constitute the largest family of mononuclear

1 non-heme oxygenases. These enzymes activate molecular oxygen (O2) and couple the oxidative decarboxylation of the αKG co-substrate to important biological transformations, such as repair of alkylated DNA and RNA, antibiotic synthesis, and protein modifications.2–5 These enzymes appear to utilize a sequential mechanism in which O2 is activated to form a reactive ferryl intermediate that subsequently hydroxylates the primary substrate. The mechanical linkage between substrate binding and oxidative decarboxylation reflects a combination of increased O2 affinity and

5–9 increased reactivity of bound O2. However, the details of how substrate binding promotes O2 binding and activation, which leads to coupled hydroxylation, are not well understood for this class of enzymes.

Factor Inhibiting HIF (FIH) is an Fe2+/αKG-dependent enzyme that regulates the activity of the HIF transcriptional activator, which regulates O2 homeostasis in human

10,11 cells. A key feature of FIH function is that O2 activation is stimulated by the binding of the CTAD domain of the HIF transcription factor, which ensures that O2 activation is tightly coupled to CTAD hydroxylation (Scheme 2.1).12–14 Recently, it was shown that

CTAD binding leads to a mixture of 6-coordinate (6C) and 5-coordinate (5C) Fe2+ in

7 FIH/CTAD, suggesting that O2 access to a coordination site is not the principal model to explain the increase in O2 reactivity in FIH. Understanding the factors governing the

28 control of O2 binding and reactivity in FIH is key to understanding its function as the main hypoxia sensor in human cells, and will provide fundamental insight into O2 binding and reactivity in the broader class of αKG oxygenases.

In order to gain insight into the structural factors that influence productive gas binding in the αKG oxygenases, we used NO as an O2 mimic to study the changes in the

O2 binding site of FIH. Nitric oxide (NO) is commonly used as an O2 surrogate to learn about intermediates and the chemistry of non heme O2 activating enzymes as NO adopts

15–17 2+ a similar geometry to that of O2. NO is capable of reversibly binding to Fe within enzyme active sites, altering the electronic properties to allow characterization by conventional EPR and electronic absorption spectroscopy.18–20 Metal-nitrosyl complexes are generally described as {MNO}n, where n represents the sum of the metal d and NO

π* electrons; Fe2+ bound NO is {Fe-NO}7.21

Scheme 2.1: Proposed mechanism for coupling in FIH

Although gas binding is crucial to defining the chemistry in non heme enzymes, little geometrical data exists. We are aware of only one structure of NO bound to an

Fe2+/αKG-dependent oxygenase, clavaminate synthase (CAS).17 The structure of CAS with NO bound revealed a pseudo octahedral metal center with NO oriented above αKG

29 in the presence of substrate.17 DFT calculations in a related enzyme, taurine dioxygenase

(TauD), indicate that NO preferentially adopts two conformations, one in which the O- atom is directed towards αKG and another pointing over the active site carboxylate moeity.16 As only the conformation in which NO is oriented above αKG is expected to be reactive based on the consensus mechanism, factors governing the switch in NO orientation may be central to coupled turnover in αKG oxygenases.

Here we report our use of crystallography, electronic spectroscopy, and DFT calculations to investigate factors that lead to productive gas binding in FIH. CTAD binding, specifically the target residue (Asn803), was tested for its role in forming the proper gas binding geometry which leads to substrate hydroxylation. This is particularly important as evidence suggests that the gas binding geometry may be the fundamental process controlling turnover in the αKG-dependent oxygenases.

2.2 Materials and Methods

2.2.1 Protein Purification

Unless noted otherwise, all reagents were purchased and used as received from commercial vendors. FIH expression and purification was carried out as previously

14,22 described. Briefly, the N-terminal His6 tagged protein was purified using a Ni-NTA column followed by removal of the tag via thrombin digestion. Exogenous metals were removed with EDTA treatment. FIH was further purified using size-exclusion chromatography to afford the active dimer. Protein concentrations were calculated by

-1 -1 22 absorption spectroscopy (ɛ280 = 48800 M cm ).

2.2.2 CTAD Purification

A 39 amino acid peptide substrate corresponding to the c- terminal transactivation domain of HIFα788-826 (CTAD) containing a Cys800 → Ala substitution with sequence

DESGLPQLTSYDAEVNAPIQGSRNLLQGEELLRALDQVN (Asn803 in bold) was ordered from EZBiolabs as a desalted synthetic peptide with free N and C termini.

Another 39 residue peptide also with a Cys800 → Ala point mutation containing the target residue Asn803 → Ala point mutation with sequence

DESGLPQLTSYDAEVAAPIQGSRNLLQGEELLRALDQVN (Ala803 in bold) was also ordered from EZBiolabs as a desalted peptide with free N and C termini. The peptides were dissolved in 25% acetonitrile and purified by reverse-phase HPLC as previously reported.14 After two rounds of purification, the samples were >95% pure as determined using matrix assisted laser desorption ionization mass spectrometry (MALDI-MS). The concentrations were established by measuring the UV-absorption at 293 nm in 0.1 M

NaOH (ε = 2400 M-1 cm-1).22

2.2.3 Crystallography and Data Refinement

Crystals of (Fe+NOG+NO)FIH were grown anaerobically using conditions similar to those previously reported in the literature.23 Crystals were grown in an anaerobic glovebox by the hanging drop vapor diffusion method by mixing 3 µL of a 20 mg/ml protein solution containing 0.55 mM FeSO4, and 2mM αKG in 50 mM Hepes pH

7.00 with 1 µL of the reservoir buffer containing 0.1 M Hepes, 1.2 M (NH4)2SO4, and 3

% polyethylene glycol (PEG) 400, pH 7.50. After setting up trays, the nitric oxide donor,

Diethylammonium (Z)-1-(N,N-diethylamino)diazen-1-ium-1,2-diolate (DEANO) was added only to the reservoir solution to facilitate NO release into the headspace during

31 crystallization. 100 µL of a 0.35 M stock of DEANO was added to the reservoir solutions

(total volume = 500 µL), such that 70 mM DEANO was present, and the coverslips were quickly resealed. Crystals were allowed to grow in the glovebox at 23°C with NO filling the headspace in the wells. Yellow crystals grew within 4 days (Figure 2.8), at which time they were harvested on the bench top within 1 minute of the coverslip being flipped to minimize gas diffusion and flash frozen in reservoir solution containing 24% glycerol.

Clear, colorless crystals of (Fe+αKG)FIH were grown anaerobically in the same fashion without the addition of DEANO to the reservoir solution.

Data was collected at the Advanced Photon Source facility in Chicago at 100 K using radiation of λ = 0.979 Å. 150 frames were collected at a crystal to detector distance of 345 mm with 1.5° oscillation and 1 second exposure per image for

(Fe+NOG+NO)FIH. For (Fe+αKG)FIH, 150 frames were captured with 2° oscillations at a crystal to detector distance of 380 nm and 1 second exposure per image. The data sets were processed using iMOSFILM.24 Coordinates from PDB 3D8C23 were employed for molecular replacement using Phaser25 and the model building and refinement done using

COOT26 and Refmac527 in the CCP428 software package. During refinement, 5% of the data was withheld and used to obtain an Rfree value. Data collection and crystallographic analysis statistics are reported in the supplementary material (Table 2.3).

2.2.4 Electronic Absorption Spectroscopy

For anaerobic experiments, stocks of FIH, αKG, NOG, and Hepes buffer were sealed with rubber septa and degassed by using argon gas. FeSO4 was brought into the glovebox as a solid and prepared using degassed H2O while DEANO stocks were prepared in degassed 10 mM NaOH and the concentration verified by its published

-1 - extinction coefficient and characteristic UV absorbance at 250 nm (ɛ250 = 6500 M cm

1).29,30 DEANO decomposes to form 1.5 equivalents of NO with a rate constant of 3.75 x

10-3 ± 8 x 10-5 s-1 under our conditions (Figure S1). Anaerobic UV-Vis samples contained

FIH (0.25 mM), FeSO4 (0.25 mM), αKG (0.5 mM), CTAD (0.5 mM) and DEANO (0.13 –

2 mM) in 50mM Hepes pH 7.00. Samples were incubated in the glovebox at 23°C for 20 minutes to permit NO release and binding. The samples were capped and then UV-Vis absorption spectra were collected immediately after removal from the glovebox on an HP

8453 UV-Visible spectrophotometer from 300nm to 900nm in 1 cm path length quartz cuvettes. Data used to construct the binding isotherm is reported in the supplementary material (Figure 2.9).

2.2.5 EPR Spectroscopy

Anaerobic EPR samples containing FIH (0.10 mM), FeSO4, (0.10 mM), αKG

(0.50mM), CTAD (0.50 mM), Asn803 → Ala peptide (0.50 mM) and DEANO (0.50 mM) in 50 mM Hepes pH 7.00 were prepared and aged anaerobically at 23°C for 20 minutes.

The samples were capped then flash frozen in liquid nitrogen (LN2) immediately upon removal from the glovebox. X-Band EPR data was collected using a Bruker Elexsys E-

500 EPR equipped with a DM4116 cavity and a Bruker ER 4118CF-O LHe/LN2 cryostat at 9.624 GHz frequency, 2.0mW power, 10G modulation amplitude, 100 GHz modulation frequency, 163 ms time constant at 4K.

The EPR data was analyzed by means of the standard spin Hamiltonian (HFe =

β.B.g.S + S.D.S) where β is the Bohr magneton, B is the applied field, g is the electron

Zeeman term, S is the electron spin operator, and D is the zero field splitting tensor.

Simulations of the EPR spectra were calculated using the XSophe software package

33 version 1.1.14.31 As the S = 3/2 {FeNO}7 spin is in the weak field limit (D>> hν), D was fixed at 10 cm-1, corresponding to the measured and calculated values of D for {FeNO}7 adducts of related enzymes 4-hydroxyphenylpyruvate dioxygenase (HPPD) and

TauD.15,16

2.2.6 DFT Calculations

Geometry optimizations were carried out on a truncated active site starting from the geometry of the crystal structure of FIH (PDB IH2L).32 Methyl imidazoles were used for His199 and His279, propionic acid for Asp201, 2 oxopropionate for αKG, and Asn803 was truncated and contained a methyl group from the adjacent amino acid (Figure 2.6). The

{Fe-NO}7 S = 3/2 spin state was generated using the spin-coupling scheme of a high spin

3+ - Fe (SFe = 5/2) antiferromagnetically coupled to two electrons from the NO π* orbitals

(SNO = 1) as this represents the proper electronic description of ferrous NO complexes.15,19 Using the geometry optimized structures, a rigid potential energy surface scan (B3LYP/6-311+g(2d,p)) was performed to investigate the energy barrier for rotating the bent NO ligand in the axial coordination position of FIH. The DFT calculations were completed with and without the target Asn803 residue in order to study the influence of the target residue on NO rotational barriers.

2.3 Results

2.3.1 Crystal Structure of FIH in Complex with Fe and NO

To gain insight into the changes in active site geometry upon O2 binding, we solved the crystal structures of (Fe+αKG)FIH and (Fe+NOG+NO)FIH. The structure of

(Fe+αKG)FIH was refined to 2.4 Å with Rfactor = 20.0% and Rfree = 24.8% while the structure of the (Fe+NOG+NO)FIH complex was refined to 2.1 Å resolution (Rfactor =

19.0% and Rfree = 23.9%). The backbones were nearly identical when residues 9-349 were aligned using PyMol (rmsd = 0.153 Å) (Figure 2.10). For both (Fe+αKG)FIH and

(Fe+NOG+NO)FIH, the active site metal retained pseudo octahedral geometry and Fe2+ was coordinated by His199, His279, Asp201 and αKG/NOG with the C1 carboxylate of

αKG/NOG bound trans to His199 as previously reported for structures of (Fe+αKG)FIH33 and (Fe+NOG)FIH.32

For (Fe+αKG)FIH, the 2Fo – Fc map displayed electron density at the coordination position trans to His279, indicating that the active site Fe of FIH was 6- coordinate as previously suggested by spectroscopic studies on FIH7 and other Fe2+/αKG oxygenases.9,34 The density was too small to accommodate a diatomic molecule such as

O2 or two aquo ligands and refined best with one water molecule in this position (Figure

2.1A). Previously reported structures of (Fe+αKG)FIH at resolutions of 2.84 Å and 2.4 Å contained unresolved density above the metal centers thought to be water molecules.32,33

Although the resolution of our structure was also modest (2.4 Å), the aquo ligand was

2 reasonably well ordered with a refined B-factor of 48.09 Å and a reasonable Fe-H2O bond length (2.0 Å). For comparison, the B-factors for the other atoms bonded to Fe

2 2 2 2 2 279 199 were, 39.8 Å , 39.8 Å , 40 Å , 34.2 Å , and 25.5 Å for His , His , O(C1αKG), O(C2αKG) and Asp201 respectively.

For (Fe+NOG+NO)FIH, the 2Fo – Fc map displayed electron density at the diffusible ligand site above the metal center which was approximately twice the density observed in the (Fe+αKG)FIH structure. This density was consistent with the presence of a diatomic molecule as a single water molecule did not adequately refine into the density.

Additionally, the Fo(Fe+NOG+NO)FIH – Fo(Fe+αKG)FIH difference Fourier map also displayed

35 corresponding electron density at nearly the same site, the size and shape of which was consistent with a diatomic molecule (Figure 2.1C). We refined this as NO due to the size of the electron density, the spectroscopic data indicative of {FeNO}7 adduct formation, and the yellow crystals that formed.

Figure 2.1: Structure of (Fe+αKG)FIH and (Fe+NOG+NO)FIH. Iron is shown in magenta, water in red. (A) View of the (Fe+αKG)FIH active site with the 2Fo – Fc map contoured to 1σ. (B) (Fe+NOG+NO)FIH active site and the 2Fo – Fc map contoured to 1σ. (C) (Fe+NOG+NO)FIH active site with the Fo(Fe+NOG+NO)FIH – Fo(Fe+αKG)FIH map contoured to 1σ. (D) (Fe+NOG+NO)FIH active site measurements. PEG is colored yellow.

The active site metal was pseudo octahedral with NO bound at the axial ligand site trans to His279, displacing water (Figure 2.1B). The O-atom of NO pointed away from the distal O-atom of Asp201 and formed van der Waals interactions with a PEG

36 molecule at the active site (Figure 2.1D). This PEG molecule was present at nearly the same position in the (Fe+αKG)FIH structure. Although it cannot be interpreted from the density that NO ligated with the nitrogen atom, chemically, this is observed in model complexes and calculations.15,16,35 The Fe-N bond length (1.83 Å) and the Fe-N-O angle

(132°) (Table 2.2), were within range of calculations and experimental data on Fe-N bond lengths (~1.70 Å – 1.88 Å) and Fe-N-O angles (~ 120° - 169°) of related enzymes and model complexes.15–17,35–37 The short Fe-NO bond length is attributed to a highly covalent Fe3+-NO- bond resulting from strong orbital overlap due to charge donation from an NO σ* orbital to Fe3+.19 The B-factors for NO were 50.4 Å2 for the N atom and 67.1 Å2 for the O atom, indicating that the N atom was reasonably well ordered while the O atom has some degree of mobility. The N atom of NO possessed a similar B-factor to that of the well-ordered N atoms on His199 (38.4 Å2) and His279 (37.3 Å2) in the

(Fe+NOG+NO)FIH structure, further supporting its location.

2.3.2 Electronic Absorption Spectroscopy

As the crystallography offered direct evidence of NO binding to Fe2+ in FIH, we sought to investigate the effect of NO binding on the electronic environment. The interaction of NO with Fe2+ at the active site of non-heme enzymes is known to cause fairly weak (ɛ < 1000 M-1 cm-1) absorption bands near 440 nm and broad transitions between 500 – 700 nm, arising from limited NO π* donor interaction into the Fe t2g orbtals.19,20,38

Addition of DEANO to (Fe+αKG)FIH led to a bright yellow solution with the expected electronic absorption bands (Figure 2.2) for {FeNO}7 non-heme iron centers.15,18,20,38,39 Upon addition of CTAD, the spectra increased in intensity, suggesting

37 either tighter gas binding as proposed by the consensus mechanism or a geometric change resulting in stronger orbital overlap. Recent computational work indicates that these transitions are formally spin-forbidden15 leading to the relatively small extinction coefficients.

Figure 2.2: Electronic absorption spectra of FIH. FIH (250 µM), FeSO4 (250 µM), αKG (500 µM), DEANO (2 mM) and CTAD or CTADN803A (500 µM) at 23°C in anaerobic Hepes (50 mM pH 7.00).

To test the effect of CTAD binding on the NO affinity of FIH, a binding isotherm was measured upon individual addition of aged [DEANO] to premixed samples (Figure

2.9). The dissociation constant (KD) for NO was obtained by fitting ΔA440 vs. [NO] to a saturation curve (equation 1) similar to previously reported for related extradiol

39 20 dioxygenases and phenylalanine hydroxylase in which Amax represents the A440 at saturation.

퐴 ∗[푁푂] 훥퐴 = 푚푎푥 (1) 퐾퐷+[푁푂]

The samples were aged for 20 minutes (~7 half-lives) to allow NO release and binding, by which point the absorbance had ceased increasing. The NO affinity of

(Fe+αKG)FIH (KD(NO) = 331 ± 24 μM) was moderate, and increased slightly for

(Fe+αKG)FIH/CTAD (KD(NO) = 203 ± 13 μM) (Table 2.1), suggesting that the gas binding pocket is only modestly influenced by CTAD such that substrate binding does not significantly promote gas binding. This is consistent with observations of a mixture of

5/6 C Fe in the presence of CTAD.7 In other αKG dependent enzymes, a much more noteworthy increase in NO affinity ranging from 5 fold to 2 orders of magnitude was observed in the presence of substrate.20,39

Figure 2.3: Binding isotherm of (Fe+NO+αKG)FIH, (Fe+NO+αKG)FIH/N803A and (Fe+NO+αKG)FIH/CTAD. FIH (0.25 mM), FeSO4 (0.24 mM), αKG (0.5 mM), CTAD (0.5 mM), N803A (0.5 mM) and DEANO (.013 - 2 mM).

To test if the steric and polar contacts created by CTAD binding promote gas

803 binding, we measured the KD(NO) using a mutant substrate containing an Asn Ala point mutation (N803A). For the (Fe+αKG+NO)FIH/N803A complex, the KD(NO) = 300

± 28 μM (Table 2.1), similar to the affinity for NO in the absence of CTAD (Figure 2.3).

8 FIH has comparable affinities for CTAD (KD = 80 µM) and N803A (KD = 180 µM), and using the method of Ramirez et al.,40 we calculated that FIH was ~84% bound with

CTAD and ~70% bound with the N803A substrate, signifying a modest contribution

39 from the substrate affinities on the gas binding affinity. This suggests that the steric environment produced by the Asn803 target residue modestly influences gas binding. A computational study on FIH suggested that the sterics of Asn803 help stimulate aquo release from the diffusible ligand site.7

2.3.3 Electron Paramagnetic Resonance Spectroscopy

To gain further insight into the geometry and electronic environment of the metal center, electron paramagnetic resonance (EPR) spectroscopy measurements were made after the addition of NO to samples of FIH. The electronic fine structure of {Fe-NO}7 is a sensitive reporter of minor geometric changes in the {Fe-NO}7 moeity.15,19,39,41–43 For an axial spin system (E/D ~ 0), the S =3/2 Ms = ± 3/2 and ±1/2 Kramers doublets are split by

2D; however, due to the large zero field splitting, the EPR signal arises solely from the

19 lower Ms = ±1/2 doublet with a positive D. As such, the effective g values are observed

19,39 at gx ≈ gy ≈ 4 and gz = 2. In light of the modest increase in NO affinity in the presence of CTAD, we tested the impact of CTAD binding on the geometry of the {Fe-NO}7 moiety in FIH. Addition of NO to (Fe)FIH produced a yellow color and complex EPR line-shape near g = 4 (Figure 4), and an S = 5/2 line-shape arising from oxidation of Fe2+ to Fe3+ as seen for related S = 3/2 {Fe-NO}7 sites.15,19,20,38,39

The EPR spectra of the (Fe+NO)FIH and (Fe+αKG+NO)FIH complex were nearly identical, implying that the geometry about the {Fe-NO}7 moiety was similar

(Figure 2.4A). The spectra of (Fe+NO)FIH displayed an axial S = 3/2 line-shape with geff

= 4.10, 3.93 (E/D = 0.014) (Figure 4). The holo FIH spectrum for the (Fe+αKG+NO)FIH sample also displayed a similar line-shape for the S = 3/2 species with geff = 4.11, 3.92

(E/D = 0.016) (Figure 2.4A). Notably, addition of CTAD containing the Asn803 target

40 residue led to the appearance of a new axial line-shape with geff = 4.06, 4.00 (E/D =

0.005) (Figure 2.4A), along with the species observed in the CTAD free complex. The change in line-shape is indicative of a change in gas binding geometry, likely due to a rotational change as Solomon et al showed experimentally and computationally that the

Fe-N-O bond length and angle do not change to any appreciable degree in the presence of substrate for related enzyme HPPD.15 This new axial line-shape corresponds to the active enzyme complex as CTAD containing the proper target residue was bound. With the addition of N803A substrate, there was no change in the gas binding environment from that of holo FIH as the sample (E/D = 0.014, geff = 4.10, 3.93) displayed no perturbation in the electronic fine structure (Figure 2.4A).

Figure 2.4: Experimental and simulated X-Band EPR spectra of {FeNO}7 FIH complexes: (A) (Fe+αKG+NO)FIH/CTAD (B) (Fe+αKG+NO)FIH/N803A (C) (Fe+αKG+NO)FIH (D) (Fe+NO)FIH. FIH (0.10mM) , FeSO4 (0.10mM), αKG (0.50mM), CTAD (0.50mM), N803A (0.50mM), DEANO (0.50 mM), 9.624 GHz

frequency, 2.0mW power, 10G modulation amplitude, 100 GHz modulation frequency, 3+ 163 ms time constant, 4K. (*) = Fe with geff = 4.30, 4.25, present for all spectra

We simulated the three different species present in the spectra of the

(Fe+NO+αKG)FIH/CTAD complex (Figure 2.4B) using XSophe.31 A summary of the simulation parameters and experimental geff values is shown in Table 1. The sum of the simulations overlaid well with the experimental spectra (Figure 4A), and showed that the spectra of Fe+αKG+NO)FIH/CTAD consisted of two S = 3/2 species and one S = 5/2 species (Figure 2.4B). By comparing the experimental and simulated spectra, we observed that the more axial S =3/2 signal with E/D = 0.005 is unique to CTAD bound

FIH in which the proper target residue is present as the N803A substrate did not exhibit this line-shape.

Table 2.1: Summary of EPR Data and NO Dissociation Constants

experimental simulation parameters

sample KD(NO) (µM) geff (gx, gy) g tensors (gx,y, gz) E/D

(Fe+NO+αKG)FIH/CTAD 203 ± 13 4.06, 4.00 2.015, 2.00 0.005

4.13, 3.91 2.015, 2.00 0.017

(Fe+NO+αKG)FIH/N803A 300 ± 28 4.10, 3.93 2.010, 2.00 0.014

(Fe+NO+αKG)FIH 331 ± 24 4.11, 3.92 2.010, 2.00 0.016

(Fe+NO) FIH ND 4.10, 3.93 2.010, 2.00 0.014

2.3.4 DFT Calculations of FIH-{Fe-NO}7

CTAD binding led to a new EPR line-shape which suggested that the geometry of

7 the {Fe-NO} moiety changed. As O2 would have to be oriented over the C2 keto group to facilitate oxidative decarboxylation, we reasoned that CTAD binding, specifically

Asn803, imposed steric constraints that changed the Fe-NO geometry to support oxidative decarboxylation. To gain further insight into the geometry of the {Fe-NO}7 moiety, we used DFT calculations to test the effect of CTAD-Asn803 sidechain on the Fe-NO geometry.

Coordinates for the 2-His 1-Carboxylate active site containing Fe2+, αKG, and

Asn803 were generated from the crystal structure of CTAD bound FIH (PDB IH2L).32

Calculations employed a formal high spin Fe3+ (S = 5/2) antiferromagnetically coupled to

NO- (S = 1).15,16,19 Following geometry optimization, a reference position defined by the

Fe-N-O-C2(αKG) vector was chosen with the Fe-N bond defining the z-axis, and NO was rotated around the equatorial plane along the Fe-N bond.

Two well-defined minima were found for (Fe+αKG+NO)FIH that were nearly isoenergetic but differed in the orientation of NO with respect to αKG (Figure 2.5). In orientation A, NO was positioned near the C1 carboxylate group of αKG. Orientation B, which was ~ 4.5 kJ/mol more stable, depicted NO pointing away from the distal O atom of Asp201 between the π-acceptor groups of His199 and the C2 keto of αKG, as expected for {Fe-NO}7 complexes (Figure 2.6).16 This further supported the crystallographic refinement as the orientation in which NO faced away from the distal O atom of Asp201

(orientation B), represented the global minimum and was the same orientation seen in the

(Fe+NOG+NO)FIH crystal structure. The calculated Fe-N bond length was 2.29 Å and the Fe-N-O angle was 146° (Table 2.2). These results were in agreement with calculations of TauD-{Fe-NO}7.16 Two well-defined minima were also observed in the

TauD-{Fe-NO}7 complex with one orientation of NO positioned above the protein carboxylate ligand and another orientation that was ~5.4 kJ/mol more stable showing NO

43 oriented above αKG.16 Electron spin echo envelope modulation experiments (ESEEM) were integrated with crystallographic studies to model NO in TauD, corroborating the orientation of NO above αKG near the C1 π-acceptor group with an estimated Fe-N bond of 1.98 Å and an Fe-NO angle of 122°.44,45

Figure 2.5: Rigid scanned potential energy surface for rotation about the Fe-NO axis in the FIH-{FeNO}7 complex (inside) schematic diagram showing the relative orientations of NO with respect to the C2 carbon of αKG. Minima are denoted with letters.

In the presence of Asn803, a single energy minimum was found (orientation C)

(Figure 2.5), representing a conformation in which NO was oriented towards the C1 carboxylate of αKG (Figure 2.6), in agreement with the crystal structure of NO bound to

CAS. The dihedral angle in orientation C (55°) was nearly the same as that of orientation

A (56°). Although the Fe-N-O angle in the presence of Asn803 (143°), was nearly identical to that found in the absence of Asn803 (146°), there was a large energetic barrier to rotation imposed by Asn803. The presence of Asn803 led to a 4 fold higher energetic barrier than that of the facial triad aspartic acid (Figure 2.5).

Figure 2.6: Optimized geometries of A, B, and C used in DFT calculations as models for FIH-{Fe-NO}7

The geometry changed from pseudo octahedral (Fe+αKG+NO)FIH to distorted octahedral for (Fe+αKG+NO)FIH/Asn803. The Fe-N bond length shortened from 2.29 Å in the absence of Asn803 to 1.85 Å in the presence of Asn803. In addition, the Fe-His279,

199 Fe-His , and Fe-O(C2αKG) bonds lengthened from 2.08 Å, 2.12 Å, and 2.17 Å to 2.24 Å,

2.23 Å, and 2.35 Å respectively (Table 2). As a result of Asn803 being positioned above

201 199 803 Asp and His , bound NO was pushed away from Asn and the N(NO) – Fe – N(His199) and N(NO) – Fe – O(Asp201) angles increased from 93 and 87 to 102 and 100 respectively, placing the uncoordinated O atom of NO 0.5 Å closer to the equatorial ligands (Table

2.2). The presence of Asn803 caused NO to be pushed towards the equatorial ligand without altering the Fe-N-O angle when compared to the position in the absence of

Asn803. This suggests that the steric environment created by the target residue plays a significant role in directing O2 for nucleophilic attack on the keto position by placing bound gas closer to the equatorial ligands.

Table 2.2: Comparison of Calculated and Experimental Geometric Parameters for FIH-{Fe- NO}7 and related enzymes FIHa Bb Cc CASd HPPDe TauDf f ΔE(rotation) (kJ/mol) 34 147 9.4 Fe – N (Å) 1.83 2.29 1.85 1.79 1.76 1.88f (1.98) g N – O (Å) 1.20 1.43 1.17 1.16 1.17 1.17f (1.19)g Fe – N – O (deg) 133 146 143 146 149 144f (122)g │Fe – N – O – C2(αKG) (deg)│ 34.4 74 55 6.4 f Fe – O(C1-O αKG) (Å) 2.02 2.13 2.01 2.01 2.06 f Fe – O(C2=O αKG) (Å) 2.15 2.17 2.35 2.27 2.37 Fe – N(His199) (Å) 2.03 2.12 2.23 2.10 Fe – N(His279) (Å) 2.00 2.08 2.24 2.23 Fe – O(Asp201) (Å) 2.04 2.05 1.98 2.06 O(NO) – C2(αKG) (Å) 3.88 4.16 3.50 2.85 N(NO) – Fe – N(His199) (deg) 91 93 102 93 N(NO) – Fe – O(Asp201) (deg) 75 87 100 99 a Measured from the Fe+NOG+NO)FIH structure. b FIH-{FeNO}7 orientation B c FIH- {FeNO}7 orientation C. d Measured from PDB: 1GVG (reference 17). Values given correspond to residues in the same position as those for FIH. eTaken from reference 15. f Taken from reference 16. g Taken from reference 45 based on ESEEM data.

2.4 Discussion

A number of mechanistic and spectroscopic studies have been undertaken to study coupling in αKG-dependent enzymes,12,16,46–52 as uncoupled chemistry typically leads to

47,53 loss of function. Understanding factors governing O2 binding and reactivity are key to understanding the function of the Fe2+/αKG-dependent oxygenases as these factors ensure that ferryl formation leads to substrate hydroxylation. A comprehension of factors governing O2 reactivity may also provide novel targeting strategies for the αKG

2+ oxygenases. In light of the fact that primary substrate does not ligate directly to Fe , and the observation that the second coordination sphere residues greatly influence reactivity,14,54 it suggests that changes in steric and non-covalent interactions formed as a result of substrate binding stimulate O2 binding and reactivity. The present study integrated spectroscopic, computational, and crystallographic analysis to test the effect of the target residue of FIH on gas binding geometry. Our structure of (Fe+NOG+NO)FIH

46 offers the first direct insight into the O2 binding geometry in the hypoxia sensing enzyme

803 FIH. Our data suggests that Asn plays a pivotal role in directing O2 reactivity by providing steric constraints which orients O2 productively for oxidative decarboxylation, resulting in coupled turnover.

2.4.1 CTAD Binding has a Modest Effect on Gas Affinity

Spectroscopic studies on non-heme iron enzymes have revealed that the Fe2+ center typically converts from a 6C geometry to a 5C geometry upon substrate

34,55 binding, implying that access to a coordination site stimulates O2 reactivity. For FIH, a mixture of 5C and 6C geometries were observed,7 indicating that other structural factors may engender reactivity. In related enzymes, the coordinative switch leads to a large increase in NO binding affinity. For example, phenylalanine hydroxylase exhibits nearly a 5 fold increase in gas binding affinity (KD(NO) = 31 µM) while protocatechuate 4,5- dioxygenase has a 2 orders of magnitude increase in NO affinity upon substrate binding

20,39 (KD(NO) = 3 μM). Compared to related enzymes, FIH has a weak gas binding affinity as the affinity for NO increased only ~ 1.5 fold upon CTAD binding (KD(NO) = 203 µM), indicating that CTAD binding modestly induces gas binding. Mutation of Asn803 → Ala had minimal effect on gas binding (KD(NO) = 300 μM) and the NO affinity was similar to that of CTAD free FIH (KD(NO) = 331 μM), suggesting that the local environment was not responsive to the shortened target residue.

The weak binding of NO to FIH is congruent with the reported KM(O2) being on

56 the higher end of bioavailable O2 (KM(O2) = 90-150 µM). Along with earlier spectroscopic and mechanistic studies,7,12 our observations are consistent with the catalytic cycle of FIH being in accord with the proposed consensus mechanism for non-

47 heme O2 activating enzymes, however the coordinative switch is not the dominant model to explain the increased O2 reactivity in FIH as O2 binding is not gated by CTAD binding.

2.4.2 CTAD Binding Introduces Steric Barriers that Reorient Gas for Productive

Reaction

The rhombicity for (Fe+αKG+NO)FIH decreases from E/D = 0.016 to E/D =

0.005 upon binding CTAD. As the E/D ratio reflects the difference in π-symmetry

199 interactions between the Fe dxy and dyz orbitals and the π-acceptor orbitals on the His and αKG ligands, we attribute the decreased rhombicity to a stronger π-symmetry overlap with the αKG ligand due to rotation about the Fe-N-O-keto dihedral angle. As O2 attacks the keto position during turnover, such a geometric change would facilitate O2 activation by lining up the gas with the keto position. Calculations in TauD demonstrated that NO orients towards the π-acceptor (C=O) groups of the equatorial ligands,16 and calculations on FIH suggest that CTAD binding enhances H-bond contacts to αKG that lower the carbonyl LUMO, activating it for nucleophilic attack.7 This suggests that productive gas binding in FIH, and other αKG oxygenases is favored when primary substrate is bound.

We propose that the more rhombic S=3/2 species (E/D = 0.016) represents a 6C

Fe2+ center in which NO is bound in a less reactive orientation. The change to a more axial (activated) electronic fine structure is likely due to rotation about the Fe-NO bond axis as a result of CTAD binding. DFT calculations suggest that NO adopts at least 2 conformations in αKG oxygenases,16 and the “activated” conformer gives rise to the more axial (E/D = 0.005) S = 3/2 line-shape. A similar interpretation was applied to

Protocatechuate 4,5-dioxygenase and catechol 2,3-dioxygenase where protein conformational changes as a result of pH variations resulted in multiple S = 3/2 species.39

As the EPR line-shape is unchanged with addition of the Asn803 → Ala substrate, it suggests that the geometry at the active site is unperturbed and gas binding is unproductive without Asn803 providing the proper steric constraints. The CTAD bound form of the enzyme is the active enzyme complex, implying that the new axial electronic line-shape produced by CTAD binding represents an active site environment “activated” for chemistry to occur. Notably, FIH fails to hydroxylate the CTAD-Asn803 → Ala

803 substrate, but hydroxylates CTAD containing the proper target residue (Asn ) with kcat

-1 8,14 = 31 min and KM(CTAD) = 77 μM. Hence, the gas binding site is not created by CTAD

803 binding, but it is influenced by it such that the presence of Asn directs O2 activation, leading to coupled turnover. Such a mechanism provides rationale for the ordering of the reaction in FIH suggested by steady state kinetics.12

We note the presence of an S = 5/2 electronic fine structure in the EPR spectra

3+ with geff = 4.30, 4.25, which arises from formation of an S = 5/2 Fe species. In the absence of NO, (Fe+αKG)FIH generates an S = 5/2 EPR spectra with geff = 4.30, 4.25, suggesting that the S = 5/2 signal is due to a protein centered oxidation. The sum of the simulated EPR spectra was comprised of 10% S = 5/2 (E/D = 0.32), 75% S = 3/2 (E/D =

0.005) and 15% S = 3/2 (E/D = 0.017). Similarly, it was found that the S = 5/2 species in protocatechuate 4,5-dioxygenase accounted for 3% – 25% of the total Fe content in the

{FeNO}7 adduct.41

Further support for a reorientation of NO upon CTAD binding comes from our observation of increased intensity in the UV-Vis spectra for CTAD bound FIH. The charge transfer intensity is dependent on the ligand-metal overlap.19 For a linear binding mode, the charge transfer transitions are predicted to be nearly degenerate, but for a bent

49 binding mode, the charger transfer intensities are ordered as follows: NO- in-plane 2π* →

2 2 - - 19 Fe dx – y < NO in-plane 2π* → Fe dxz < NO out-of-plane π* → Fe dyz. The weak transitions are within reason based on electronic structure calculations of {FeNO}7 adducts (ɛ < 1000 M-1 cm-1), and suggest a limited π donor interaction into the Fe d

19 orbitals. We propose that NO primarily interacts with the Fe dxy orbitals in the absence

19,20 803 of CTAD, but in the presence of CTAD containing Asn , bound NO rotates to produce new ligand-metal overlap with the Fe dyz orbital, giving rise to a more intense charge transfer. If O2 were bound, stronger orbital overlap would result in a stronger Fe-

O2 bond, polarizing the bound O2 and giving the distal O-atom more nucleophilic character. Our calculations suggest that CTAD binding strengthens the Fe-N bond as the bond length shortened in the presence of Asn803.

Our calculations lend further credence to the bond axis rotation model as NO adopts a more stable orientation in the presence of CTAD that is different than the most stable orientation in the absence of CTAD. The Fe-N-O angle (~144°) in the presence or absence of CTAD remained relatively unchanged and are comparable to previous calculations of TauD and HPPD,15,16 indicating that the multiple EPR line-shapes observed are not due to changes in the Fe-N-O bond angle, but rather a change in NO orientation.

Combined with the EPR data which shows a unique electronic environment forming when the proper steric constraints are present, and the crystallographic data which depicts the NO orientation in the absence of CTAD, the calculations support a model in which productive gas binding is stimulated by substrate binding. When CTAD containing Asn803 binds, the expected changes in the gas geometry would be (i) bound

50 gas bends closer to the equatorial ligands (αKG) (ii) the gas rotates over αKG, and (iii) the gas bond becomes polarized such that the uncoordinated O atom gains more nucleophilic character, enhancing reactivity.

2.4.3 The Gas Binding Pocket of FIH

The active site of FIH was conserved between the (Fe+αKG)FIH and

(Fe+NOG+NO)FIH structures. There was a PEG molecule above the NO ligand that interacted with NO via van der Waals interactions. As the active site of FIH is solvent exposed, the presence of PEG, an artifact from the crystallization buffer, suggests that

FIH prefers to have peptide like constructs bound above the metal center. There was also a water molecule above NO that likely plays a stabilizing role and “locks” bound gas in this orientation. The energetic penalty to reorientation is approximately equal to the bond dissociation energy to break H-bonds, which is 23 kJ/mol per H-bond.57 The structure of

FIH/CTAD shows that the target asparagine residue occupies nearly the same space occupied by PEG and the water molecule,32 suggesting that CTAD removes stabilizing contacts to the bound ligand. These alterations likely trigger a rotation to a productive conformation for nucleophilic attack on the keto position of αKG. Similarly, slight shifts in nearby residues accommodated substrate and NO binding in the oxygen binding cavity in the extradiol dioxygenase BphC.36

Two well defined energy minima that differed in energy by ~ 4.5 kJ/mol were observed in the absence of Asn803. Despite the small difference in energy only the most stable conformer (orientation B) was observed in the crystal structure of

(Fe+NOG+NO)FIH. It’s possible that the difference in energy stabilizes B enough such that it is the only conformation observed crystallographically. In the proposed consensus

51 mechanism, nucleophilic attack on the C2 keto group of αKG initiates decarboxylation, leading to formation of the ferryl oxidant.2 In order for nucleophilic attack to commence,

O2 has to be in close proximity to the C2 keto position. The distance from the O atom of

NO to the C2 of NOG was ~ 3.6 Å. We interpret this orientation of NO to represent unproductive gas binding as the substrate is not present, and NO is not ideally positioned for oxidative decarboxylation. Support for this orientation being unproductive comes from observations that uncoupled chemistry in the absence of prime substrate occurs at a slow rate.58 Uncoupled chemistry has been reported to be a slow process in TauD as well.53

As the most accessible site in the gas binding pocket rests directly over αKG and the gas binding affinity was modestly influenced by CTAD binding, it suggests that steric

803 occlusion of NO due to the presence of Asn plays a more dominant role in directing O2 reactivity than increased gas binding affinity. The EPR line-shape for the Asn803 → Ala point mutation implied a malformed gas binding pocket owing to improper steric constraints, indicating that Asn803 is the major factor influencing gas orientation. FIH

803 -1 hydroxylates CTAD-Asn with kcat = 31 min but fails to hydroxylate the CTAD-

N803A substrate,8,14 implying that reorientation to the proper geometry greatly stimulates

O2 reactivity.

Interestingly, the calculated orientation in the presence of Asn803 appeared to be non-optimal for nucleophilic attack as it was 3.5 Å from the C2 keto group of αKG and the dihedral angle to the C2 carbon was 55°. However, the barrier to rotation over the C2 carbon of αKG was minimal, ~ 10 kJ/mol. In the structure of CAS with NO bound, NO was poised over αKG and was in a fairly optimal geometry for nucleophilic attack as the

NO to C2 carbon of αKG was 2.85 Å and the dihedral angle to the C2 carbon was 6.4°, suggesting productive gas binding in the presence of substrate.17 The moderately high B- factor (~60 Å2) for the distal O atom of NO in FIH suggests that it has some degree of mobility.

Our calculations showed that in the presence of Asn803, NO adopted a conformation in which it was positioned near the C1 π-acceptor group of αKG, consistent with the observed orientation of NO in substrate bound CAS,17 suggesting that this orientation may depict the lowest energy state in the αKG-dependent oxygenases upon substrate binding. Paradoxically, NO binding to Fe2+ in CAS displaced the bound aquo ligand concomitant with rearrangement of the C1 carboxylate of αKG to the position previously occupied by water, suggesting that ferryl formation would be incorrectly positioned for substrate oxidation.17 Despite this implication, NO was oriented towards

αKG as if poised to initiate nucleophilic attack, supporting the notion that altered local contacts which result from substrate binding help direct productive gas binding. It is possible that the ferryl may rearrange in CAS but at present there is no evidence to support this. In contrast to CAS, NO binding in FIH displaced water at the diffusible ligand site, leading to a geometry in which ferryl formation would be positioned appropriately for coupled substrate oxidation. Unfortunately, we were unable to obtain a crystal of NO bound in the presence of CTAD that diffracted well, therefore it is unclear if such a rearrangement also occurs in FIH, but our DFT calculations and those of related enzymes suggests that a rearrangement is unlikely.16

In the case of FIH, CTAD binding imposes steric constraints on NO which would be minimized by NO rotating away from His199 towards the most stable orientation

53 observed in our calculations. This path poses the least barrier to rotation, but importantly,

CTAD binding causes stronger orbital overlap, making the distal O-atom of NO more nucleophilic and the carbonyl carbon of αKG more electrophilic.7,19 NO, which is angled closer to the equatorial plane, encounters the C2 keto π acceptor group during rotation before any other π acceptor groups.

Taken together, the crystal structure of the CAS-{Fe-NO}7 complex and calculations of the TauD-{Fe-NO}7 complex suggest that gas reorientation may be a general mechanistic strategy employed by the αKG-dependent oxygenases whereby substrate binding stimulates O2 reactivity thereby ensuring tight coupling to substrate oxidation. Although speculative, if we consider the ratio (~ 6) of the mole fraction of the unproductive to productive orientations based on their relative energies, multiplied by the

~ 2 fold increase in gas binding affinity, CTAD binding could account for approximately a 12 fold increase in O2 reactivity.

2.5 Conclusion

In summation, we have provided evidence that suggests the coordinative switch model is not the dominant model to explain increased O2 reactivity in FIH. NO is able to access the active site of FIH, however, the orientation, which is stabilized by a network of

H-bonds, is non-optimal for decarboxylation. Based on our data, we propose that CTAD binding stimulates O2 reactivity via rotation about the Fe-O2 bond axis, thereby positioning “activated” O2 over the keto position of αKG for oxidative decarboxylation.

The sterics of the target residue and changes in local contacts are important for forming the proper gas binding pocket which leads to coupled turnover. Substrate induced rotation

54 about the Fe-O2 moiety may be a strategy utilized by the broader class of αKG oxygenases to facilitate O2 reactivity.

2.6 Appendix

2.6.1 Supplemental

2.6.2 DEANO Decomposition in 50 mM Hepes pH 7.00

The rate of DEANO decomposition to form NO in 50 mM Hepes pH 7.00 was determined in accordance with a previously described method.59 Hepes was degasses by bubbling argon gas through the solution before use in the glovebox. DEANO was prepared in 10 mM NaOH and 2.6 µL of a 15.4 mM stock was added to 197.4 µL of 50 mM Hepes pH 7.00. The cuvette was capped and the disappearance at 250 nm was monitored at 23°C on an HP 845x UV-Visible spectrophotometer for 20 minutes. Under our conditions, the rate of NO release followed first order exponential decay kinetics with time (Figure S1). The rate constant (k = 3.75 x 10-3 ± 8.46 x 10-5 s-1) was determined from the average of 5 runs. The half-life (3 mins) was calculated using the relationship: t1/2 =

0.693(1/k) as previously described.30

Figure 2.7: Example of the first order decay plot used to calculate the rate of DEANO decomposition in 50 mM Hepes pH 7.00 at 23°C.

Figure 2.8: Crystal of (Fe+NOG+NO)FIH grown in the presence of DEANO used for data collection

Table 2.3: Data collection and refinement statistics. Values in parentheses represent the highest resolution shell

Data set (Fe2++αKG)FIH (Fe2++NOG+NO)FIH λ (Å) 0.979 0.979 Space group P41212 P41212 Unit cell (Å) a = b = 86.6, c = 147.2 a = b = 86.6, c = 147.7 α = β = γ = 90° α = β = γ = 90° Resolution (Å) 74 – 2.42 (2.51 – 2.42) 61 – 2.1 (2.17 – 2.1) Rmerge (%) 13.9 (121) 9.7 (109) Rpim (%) 3 (29) 2.5 (36.9) Mean I/σ (I) 18.2 (2.9) 16 (1.9) Completeness 100 (100) 99.6 (96.8) Multiplicity 22 (18.6) 15.9 (13.6) Refinement Resolution range (Å) 74 – 2.42 61 – 2.1 Number of Reflections 22043 33489 Average B factor (Å2) 55 54.05 Rwork/Rfree (%) 20.0/24.8 19.0/23.9 R.m.s deviations Bond lengths (Å) 0.015 0.020 Bond angles (Å) 1.73 1.91 Ramachandran plot, residues in (%) Preferred region 94.40 94.4 Allowed region 5.60 5.60 Disallowed region 0 0

Figure 2.9: UV-Vis data used to construct binding isotherm (top left) (Fe+NO+αKG)FIH/CTAD (top right) (Fe+NO+αKG)FIH (bottom left) (Fe+αKG+NO)FIH/N803A: FIH (0.25 mM), αKG (0.50 mM), FeSO4 (0.25 mM), CTAD (0.50 mM), N803A (0.50 mM), DEANO (0.015 – 2 mM),) in 50 mM Hepes pH 7.00 at 23°C

Figure 2.10: Structure of (Fe+αKG)FIH and (Fe+NOG+NO)FIH. Overlay of (Fe+αKG)FIH (green) and (Fe+NOG+NO)FIH (cyan) showing the conserved β-strand jelly roll core.

2.7 References

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(39) Arciero, D. M.; Orville, a. M.; Lipscomb, J. D. [17O]Water and Nitric Oxide Binding by Protocatechuate 4,5-Dioxygenase and Catechol 2,3-Dioxygenase. Evidence for Binding of Exogenous Ligands to the Active Site Fe2+ of Extradiol Dioxygenases. J. Biol. Chem. 1985, 260, 14035–14044.

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(46) Flagg, S. C.; Giri, N.; Pektas, S.; Maroney, M. J.; Knapp, M. J. Inverse Solvent Isotope Effects Demonstrate Slow Aquo Release from Hypoxia Inducible Factor- Prolyl Hydroxylase (PHD2). Biochemistry 2012, 51, 6654–6666.

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CHAPTER 3

CONTRIBUTIONS OF THE FACIAL TRIAD CARBOXYLATE TO LIGAND ACCESS IN FACTOR INHIBITING HIF (FIH)

3.1 Introduction

The Fe2+/αKG-dependent oxygenases are a functionally diverse family of enzymes that catalyze a wide array of chemical transformations, including epoxidation, chlorination, desaturation, and most commonly, hydroxylation.1–3 These enzymes typically functionalize unactivated aliphatic carbons, a major hurdle in synthetic chemistry that still remains elusive to chemists. It has been shown that the chlorinating enzyme, SyrB2, can also catalyze bromination,4 hydroxylation,5 and more recently azidation and nitration chemistry,6 suggesting that these enzymes can be engineered to catalyze alternative reactions.

2+ Factor inhibiting HIF (FIH), is an O2 sensing Fe /αKG-dependent enzyme that, under normoxic conditions, catalyzes the hydroxylation of an asparagine residue (N803) in the Hypoxia Inducible Factor (HIF) transcription activator, thereby repressing HIF gene expression. Installing functional groups on specific aliphatic carbons, especially during late stages of a synthetic sequence, is a challenging task. As the natural substrate for FIH is a peptide, FIH is an attractive target for redesigning alternative chemistry.

Elucidating factors that control ligand access to the Fe cofactor is the first step in understanding how to engender FIH with alternative chemistry such as azidation, which could open novel avenues for “biorthogonal” conjugation chemistry.

The active site of the Fe2+/αKG-dependent enzymes is comprised of a highly conserved motif known as the 2-His-1-carboxylate facial triad motif.7–9 The facial triad provides three protein derived ligands for iron binding while the other coordination sites

64 are occupied by αKG in a bidentate fashion, and a water molecule at the diffusible ligand site (Figure 3.1).10–13 A subset of the Fe2+/αKG-dependent enzymes that catalyze halogenation chemistry have evolved to bind the Fe cofactor with only two protein derived His residues, and a halide ion in place of the carboxylate ligand.14,15 FIH affords a unique opportunity to understand how the protein active site differentiates between halides and other ligands as it was recently demonstrated to be active with a mutated facial triad carboxylate.16

The consensus mechanism for Fe2+/αKG-dependent oxygenases involves the

2 sequential binding of αKG, prime substrate, then O2. A combination of crystallography and VTVH MCD and CD spectroscopy have shown that substrate binding leads to

10,11,17–19 release of the aquo ligand in the axial coordination site, permitting O2 binding.

Oxidative decarboxylation of αKG to succinate releases CO2 and generates a reactive ferryl intermediate, which has been observed in the halogenases CytC320 and SyrB2,21 and the hydroxylases prolyl hydroxylase 4 (PH4)22 and taurine dioxygenase (TauD),22,23 and identified as the species responsible for hydrogen atom transfer (HAT). Following

HAT, radical rebound chemistry leads to the desired chemical product. For halogenases, halogen rebound chemistry ensues while hydroxide rebound occurs in the hydroxylases such as FIH.

A key feature of the consensus mechanism is that substrate binding stimulates O2 binding and reactivity, ensuring formation of the reactive ferryl intermediate is tightly coupled to substrate modification. The crystal structures of (Fe+αKG)FIH reveal that the active site is solvent exposed (Figure 3.1), and when primary substrate (CTAD) binds, there is a channel that opens up at the diffusible ligand site.11 As the active site is solvent

65 exposed, the Fe cofactor would be susceptible to binding by endogenously produced molecules that are similar to O2, such as NO, and CO, suggesting that mechanisms may exist to differentiate between different ligands. In the structures where the aquo ligand is resolved, the uncoordinated O-atom hydrogen bonds (H-bonds) to the coordinated water, and forms new H-bond contacts to the backbone amide of CTAD upon substrate binding

(Figure 3.1).11,13 Recent studies have suggested that the facial triad carboxylate plays a role in stabilizing the axial aquo ligand at the diffusible ligand site, implicating this residue in controlling ligand access.19

Figure 3.1: Surface representation of FIH in the absence of substrate showing the solvent exposed active site. PDB IDs: 1H2N11, 3P3P13, 1H2L11.

We reasoned that the facial triad carboxylate regulates the access of exogenous ligands, and disruption of the H-bond to the axial aquo ligand would facilitate exogenous ligand access to the Fe cofactor. In the present work, the facial triad carboxylate of FIH

(Asp201) was mutated to Glu (D201E) and Gly (D201G) in order to study the effect of the facial triad carboxylate in ligand access. Our data indicates that exogenous ligands are able to access the active site of FIH in the D201E and D201G variants, but not WT-FIH.

This data implicates the facial triad carboxylate in controlling ligand access, and suggests that FIH can be engineered for alternate rebound chemistry.

3.2 Materials and Methods

3.2.1 Protein Mutagenesis and Purification

Site-direct mutagenesis was carried out to insert point mutations in the pET28a-

FIH construct. The following primers were used to construct the desired mutations:

Asp201Gly forward, 5’-GTGACACCTGCTCACTATGGCGAGCAGCAGAAC-3’; reverse, 5’-GTTCTGCTGCTCGCCATAGTGAGCAGGTGTCAC-3’; Asp201Glu forward, 5’-GACACCTGCTCACTATGAGGAGCAGCAGAACTTTTTT-3’; reverse,

5’-AAAAAAGTTCTGCTGCTCCTCATAGTGAGCAGGTGTC-3’. The final constructs were sequenced by Genewiz to confirm that only the desired mutations were present before transforming into BL21(DE3) cells. WT-FIH and the variants were expressed and purified according to a previously described protocol.24 The N-terminal

His6 tagged WT-FIH and variants were purified using a Ni-NTA column. The His6 tag was removed by thrombin digestion and the eluents were treated with EDTA to remove exogenous metals, then further purified using size-exclusion chromatography to yield the active dimer ˃95% pure as assessed by SDS-PAGE. The cleaved protein contained 3 extra residues (GSH) at the N-terminus. Protein concentrations were determined by

-1 -1 24,25 absorption spectroscopy (ɛ280 = 48800 M cm ).

3.2.2 CTAD Purification

A 19 residue amino acid peptide substrate corresponding to the C- terminal

788-807 800 transactivation domain (CTAD19mer) of HIFα containing a Cys → Ala mutation with sequence DESGLPQLTSYDAEVNAPI (N803 bolded) was ordered from EZBiolabs as a desalted synthetic peptide with free N and C-termini and used as received without

67 further purification for co-crystallization studies. For steady state kinetics, a 39 amino acid peptide of HIFα788-826 also containing a Cys800 → Ala mutation with sequence

DESGLPQLTSYDAEVNAPIQGSRNLLQGEELLRALDQVN (N803 bolded) was ordered from EZBiolabs as a desalted synthetic peptide with free N and C-termini. The

39-residue peptide was dissolved in 25% acetonitrile and purified to >95% purity by reverse-phase HPLC as previously described24,25 The concentration was established by measuring the UV-absorption at 293 nm in 0.1 M NaOH (ε = 2400 M-1 cm-1).24

3.2.3 Crystallography

- Crystallization of (Fe+NOG+Cl )D201G/CTAD19mer was carried out as previously described in the literature with slight modifications.16 Crystals were grown aerobically by the hanging drop vapor diffusion method by mixing 2 µL of a 20 mg/ml protein solution containing 0.55 mM FeSO4, 2mM NOG, and 2 mM CTAD19mer in 50 mM Hepes pH 7.00

(100 mM NaCl) with 1 µL of the reservoir buffer containing 0.1 M Hepes, 1.2 M

(NH4)2SO4, and 3 % polyethylene glycol (PEG) 400, pH 7.50. The crystals were grown at

20°C and were harvested after 9 days and flash frozen in a cryoprotectant comprised of the reservoir solution containing 24% glycerol.

The data was collected at the Brookhaven National Synchrotron Radiation facility in New York using radiation of λ = 1.08 Å. 120 frames were collected at a crystal to detector distance of 300 mm with 1° oscillation and 3 second exposure per image. The data set was processed using HKL2000.26 Coordinates from PDB 3D8C16 were used for molecular replacement and the refinement completed using COOT27 and Refmac528 in the CCP429 software package. 5% of the data was withheld during refinement and used to obtain an Rfree value.

3.2.4 UV-Vis Spectroscopy

All reagents were degassed using argon gas and brought into an anaerobic glovebox except for FeSO4 and DEANO which were brought in as solids and dissolved in degassed water and 10 mM NaOH respectively. The ligands were also brought in as solids and dissolved in buffer. Samples contained FIH (400 µM) or the variants D201E

(400 µM) and D201G (400 µM), αKG (400 µM), FeSO4 (375 µM), and 2 mM of each

- - respective ligand (NO, NaCN, NaN3 , NaCO3, NaOCN , NaSCN, NaC2H3O2) in 50 mM

Hepes pH 7.00 containing 100 mM NaCl. Each ligand was added to an individual sample of (Fe+αKG)Protein and the spectra were collected from 300 – 900 nm on an Avantes

AvaSpec-2084 Fiber Optic Spectrometer set up in a glovebox. Difference spectra were generated from subtracting the respective (Fe)Protein only spectrum from each respective

(Fe+αKG)Protein and (Fe+αKG+Ligand)Protein spectrum.

3.2.5 Initial Screen and Dose Response Assays

30 All reactions were performed under ambient O2 (~220 μM) conditions. For initial screens, reactions containing ascorbate (2 mM), FeSO4 (25 µM), αKG (100 µM), and CTAD (100 µM), were pre-incubated for 1 minute with each ligand, which were present at 2 mM concentrations except for CO (750 µM) in 50 mM Hepes (100 mM

NaCl) pH 7.00 at 37°C. Reactions were initiated by adding either FIH (0.5 µM), D201G

(2.2 µM), or D201E (5 µM). 5 μL aliquots of the reaction were quenched after 2 minutes in 20μL of 75% acetonitrile with 0.2% trifluoroacetic acid saturated with the matrix, 3,5

Dimethoxy-4-hydroxycinnamic acid. Hydroxylation was monitored by determining the mole fraction of peptide converted to product based on measuring the ratio of the un-

69 hydroxylated peptide (M+H)+, m/z = 4255) and hydroxylated peptide (m/z = 4271) using

MALDI-MS as previously described.24,31

Dose response assays were carried out in a similar manner according to previous reports from the literature.31 Assays were carried out in 50 mM Hepes containing NaCl

(100 mM), ascorbate (2 mM), FeSO4 (25 µM), αKG (100 µM), CTAD (100 µM), and

- NaN3 (0 – 5 mM). The NO donor DEANO was prepared anaerobically in 10 mM NaOH and the extinction coefficient verified by its characteristic UV absorption at 250 nm (ɛ250

= 6500 M-1cm-1).32 Individual aliquots of DEANO were opened right before addition to the reaction mixtures. DEANO releases 1.5 moles of NO per mole of DEANO with a rate

-3 -1 constant of k = 9.77 x 10 s 37°C (t1/2 = 1.2 minutes) at 37°C pH 7.00, such that our concentration of NO ranged from 0 – 0.8 mM. Aliquots (5 μL) of the reaction were quenched in 20μL of 75% acetonitrile with 0.2% trifluoroacetic acid at designated time

OH OH OH points. Initial rates were determined by calculating the χ(CTAD ) = {I(CTAD ) / (I(CTAD ) +

31 I(CTAD))} * [CTAD].

3.2.6 X-ray absorption spectroscopy sample preparation

The XAS sample was prepared anaerobically in a glovebox by mixing 4 mM

D201G with 6 mM FeSO4 in 50 mM Hepes pH 7.00 containing 100 mM NaCl. The sample was concentrated using a 30 kDa molecular weight cutoff filter to remove unbound metal. The concentration was verified and NOG was added such that the final concentrations were 4.3 mM D201G, 5 mM NOG, 100 mM NaCl, in 50 mM Hepes pH

7.00. The sample was immediately flash frozen in liquid N2 upon removal from the glovebox and stored at -80°C until analysis. ICP measurements were made on the protein sample and showed that the concentration of Fe was 1.5 mM.

3.2.7 X-ray absorption spectroscopy data collection and analysis

Iron K-edge data were collected at Stanford Synchrotron Radiation Lightsource

(SSRL) beamline 7-3 with ring conditions of 3 GeV and 495 – 500 mA. Dedicated beamline optics for 7-3 was used, including a Si (220) double crystal monochromator with a rhodium-coated mirror upstream for focusing and harmonic rejection. The sample was cooled to ~10K during data collection with a liquid helium cryostat (Oxford

Instruments). X-ray fluorescence was collected with a 30-element germanium detector

(Canberra) and scattering was minimized with a 3mm Mn filter and Sӧller slits placed in- between the sample chamber and the detector. An X-ray transmission spectrum of a reference Fe foil was collected concurrently with fluorescence data and used for calibrating the K-edge energy of each spectrum.

X-ray absorption near edge structure (XANES) was analyzed by EXAFS123.33 as previously described.34 A cubic function was used to fit the baseline at the pre-edge region and the rising edge was fitted using a 75% Gaussian and 25% Lorenzian function.

A Gaussian function was used to fit the transitions occurring below the edge (pre-edge), where the area of the Gussians were taken to be peak areas. The position of the K-edge energy (E0) of the spectra is deduced from the first derivative of the XAS spectra.

Data processing and background subtraction was performed using ATHENA35 software using IFEFFIT.36 Bad detector channels were removed from each scan, and the

K-edge energy was calibrated to the first inflection point of Fe reference foil (7112.5 eV).

Background removal and normalization was performed using the Autobk algorithm35 with E0 set to 7125 eV and Rbkg set to 1.0 with a k-weight of 3. A linear pre-edge range of -150 to -30 eV and a quadratic polynomial post-edge of 150 to 900 eV relative to E0

71 was used with the edge jump normalized to 1. Data extraction was performed using a spline k range of 2 to 14 Å-1.

EXAFS fitting was performed using ARTEMIS35 software using the IFEFFIT

36,33 engine. First shell scattering paths were generated for reff of Fe-N at 2.0 Å, Fe-S at

2.2 Å, and Fe-Cl at 2.3 Å using FEFF6. Multiple-scattering paths for Fe-imidazoles were generated using metal-histidine coordinates from a high resolution protein crystal structure (PDB ID 1Q5Y)37 using FEFF6. Multiple-scattering paths for Fe-NOG were generated from metal-NOG coordinates from the protein crystal structure (PDB ID

4V2W)38 using FEFF6. The k3-weighted r-space data over a k-range of 2 – 14 Å-1 was

2 fitted. The amplitude reduction factor, S0 was set to 0.9 and the E0 was allow to vary for each model. For each single scattering path or set of multiple scattering paths that describe a ligand (imidazole or NOG), the distance between Fe and the scattering atom and the σ2 was allowed to vary.

Goodness of fit was compared between different models by minimizing three parameters: χ2 (Equation 1), reduced χ2, and R factor (Equation 2).

2 푁푖푑푝 푁 2 휒 = ∑푖=1{(푅푒(푓푖)) } (1) 푁ɛ2

푁 2 2 ∑푖=1{(푅푒(푓푖)) +(lm(푓푖)) } 푅 = 푁 2 2 (2) ∑푖=1{(푅푒(휒푑푎푡푎푖)) +(lm(휒푑푎푡푎푖)) }

2 Where Nidp is the number of independent points; Nϵ is the number of uncertainties to minimize; Re(fi) is the real part of the EXAFS fitting function; Im(fi) is the imaginary part of the EXAFS fitting function. Reduced χ2 = χ2/(Nind Nvarys) (where Nvarys = the number of refining parameters) and represents the degrees of freedom in the fit.

Increasing the number of adjustable parameters in the fit are expected to improve R, but reduced χ2 will go through a minimum with the increase indicating the model is

72 overfitting the data. The best fit models were deduced by minimizing both R and reduced

χ2 and deviation of σ2 values from typical values.

3.3 Results

The D201E and D201G variants of FIH were purified for steady state kinetic assays aimed at testing the role of the facial triad carboxylate in ligand access. A previous study demonstrated that the D201G had wild type like activity while the D201A and

D201E variants exhibited significant uncoupling.16 The crystal structure of

(Zn+αKG)D201G/CTAD revealed that in the absence of the coordinated carboxylate ligand, a water molecule was bound in the equatorial plane at the site normally ligated by

D201.16 A carbonate ion was bound in a bidentate fashion near this position in the structure of (Fe+αKG)D201A structure,16 suggesting that D201 may regulate exogenous ligand binding to the Fe cofactor. We previously found that chloride was necessary for hydroxylation chemistry in D201G (Hangasky unpublished results), implying that chloride and other exogenous ligands may interact with the Fe cofactor in D201G.

3.3.1 Initial Screen and Dose Response Assays

To test the role of D201 in facilitating exogenous ligand access, we performed an initial activity screen in the presence of 2 mM exogenous ligands under ambient [O2]

(~217 µM) and saturating concentrations of FeSO4 (25 µM), αKG (500 µM), ascorbate (2 mM) and chloride (100 mM) in 50 mM Hepes pH 7.00. In contrast to WT-FIH which was

- only inhibited by NO, D201G and D201E were both inhibited by NO and azide (N3 )

(Figure 3.2). NO is commonly used as an O2 mimic and is known to bind to non heme O2

2+ 39–42 - activating Fe /αKG-dependent enzymes. N3 has also been demonstrated to bind to the non heme Fe center of the chlorinating enzyme SyrB2.6 Although cyanide (CN-)

73 seemed to be a modest inhibitor in the initial screen, changes in the UV absorption spectrum after incubating (Fe+αKG)FIH with CN- led to the conclusion that ferricyanide formation caused metal sequestration, inhibiting activity. Interestingly, cyanate (OCN-) was an activator of D201G activity, suggesting that it interacted with the Fe cofactor and served as a counter ion in the same capacity as the missing carboxylate ligand.

Figure 3.2: Initial inhibitor screen for FIH (top left) and the facial triad D201E (top right) and D201G (bottom) variants. Assays were performed in 50 mM Hepes pH 7.00 containing 100 mM NaCl at 37°C; FIH (0.5 µM), D201G (2.2 µM), D201E (5 µM), αKG (500 μM), FeSO4 (25 μM), CTAD (100 μM), ascorbate (2 mM). All inhibitors were 2 mM except for CO which was 750 µM.

- As the screen identified NO and N3 as ligands that significantly inhibited WT-

FIH and the D201 variants, a full dose response curve was measured in order to

- determine the IC50 for both NO and N3 . Inhibition by NO was greatest for D201E (IC50 =

4 µM), followed by D201G (IC50 = 22 µM) and then WT (IC50 = 65 µM) (Table 3.1), suggesting that disruption of the stabilizing metal-aquo H-bond relaxed ligand selectivity.

NO has previously been demonstrated to inhibit the other hypoxia sensing enzyme prolyl

43 hydroxylase domain 2 (PHD2) with IC50 = 10 µM. NO has also been shown to inhibit

44 - FIH activity in cells. N3 significantly inhibited D201G activity (IC50 = 130 µM) compared to WT, and moderately inhibited D201E activity (IC50 = 280 µM) compared to

WT (Figure 3.3 and Table 3.1).

- Figure 3.3: Dose response assays varying NO (left) and N3 (right) for FIH and the D201E and D201G variants. Assays were performed in 50 mM Hepes pH 7.00 containing 100 mM NaCl at 37°C; FIH (0.5 µM), D201G (2.2 µM), D201E (5 µM), αKG (500 μM), - FeSO4 (25 μM), CTAD (100 μM), ascorbate (2 mM), NO (0 – 0.8 mM), and N3 (0 – 5 mM).

3.3.2 UV-Vis Absorption Spectroscopy

To test for binding of exogenous ligands to the Fe cofactor, UV-Vis absorption spectra were measured in the presence of 2 mM concentrations of the ligands (NO, OCN-,

- and N3 ) identified from the kinetic assays that affected FIH activity. Both the robustness and versatility of using spectral features to assess ligand binding has been well established.6,45,46 Bidentate chelation of αKG in the αKG-dependent enzymes leads to a metal to ligand charge transfer (MLCT) typically centered around 500 nm, attributed to

40,46 transitions from the Fe t2g(π) → αKG π* orbitals. (Fe+αKG)FIH exhibited the characteristic MLCT for αKG-dependent oxygenases previously observed in FIH47 centered at 500 nm (Figure 4.4D), indicative of αKG binding to Fe2+ at the active site.

(Fe+αKG)D201E exhibited the same MLCT as WT-FIH centered at 500 nm (Figure

4.4C), but (Fe+αKG)D201G exhibited a blue shifted MLCT centered at 485 nm (Figure

- 4.4B), suggesting stabilization of the Fe t2g orbitals. The extinction coefficient (~ 200 M

1cm-1) of the MLCT for WT-FIH and the variants were similar to a previous report for

(Fe+αKG)FIH (~ 240 M-1cm-1)47 and (Fe+αKG)TauD (~ 250 M-1cm-1).48

Figure 3.4: Anaerobic UV-Vis spectra showing the MLCT for WT-FIH and the D201 variants in 50 mM Hepes pH 7.00 (100 mM NaCl). (A) WT-FIH, D20E, and D201G - - - treated with NO. (B) D201G treated with N3 and OCN . (C) D201E treated with N3 . (D) - - WT-FIH treated with N3 and OCN . FIH (400 µM), D201E (400 µM), D201G (400 µM), - - FeSO4 (375 µM), αKG (400 µM), NO (2 mM), OCN ( 2mM), N3 (2mM).

Binding of NO to Fe2+ in non heme enzymes typically leads to an intense transition around 440 nm and a broad transition from 550 – 700 nm.39,40,49 The transitions are ligand to metal charge transfers (LMCT) arising from electronic transitions between

41 the NO π* and Fe t2g orbitals. The spectrum of (Fe+αKG+NO)FIH exhibited a well- defined peak at 450 nm and a broad transition from 550 – 700 nm as seen for related enzymes (Figure 4.4A).39,40 (Fe+αKG+NO)D201E also had a strong transition centered at

450 nm and a broad transition from 550 – 700 nm while (Fe+αKG+NO)D201G displayed

77 a strong transition centered at 435 nm and a broad transition from 550 – 700 nm, indicative of NO binding to Fe2+ in FIH and the variants. The intensity of the transition near 440 nm followed the same trend as the inhibition observed for NO, D201E > D201G

> WT-FIH.

- Binding of N3 to (Fe+αKG)D201G and (Fe+αKG)D201E resulted in a 15 nm

(λmax = 500 nm) and 10 nm (λmax = 500 nm) red shift respectively (Table 3.1), consistent

- - 2+ 6 with observations of a red shift for N3 and OCN binding to Fe in (Fe+αKG)SyrB2.

- Similarly, binding of OCN to (Fe+αKG)D201G (λmax = 500 nm) also led to a 15 nm red shift. The red shifts in the MLCT transitions can be attributed to destabilization of the Fe t2g orbitals or stabilization of the ligand π* orbitals upon binding. We did not observe any

- - spectral perturbations in WT-FIH in the presence of N3 or OCN , consistent with the

- - kinetic data which suggested N3 and OCN are unable to bind the Fe cofactor to inhibit activity.

Table 3.1: Summary of IC50 and electronic absorption spectroscopy data

IC50 (µM) MLCT and LMCT λ (nm)

WT D201G D201E Ligand WT D201G D201E

- - - αKG 500 485 500

65 ± 11 22 ± 2.0 4.0 ± 0.4 NO 450, 550-700 430, 550-700 450, 550-700

- > 2 mM 130 ± 15 280 ± 20 N3 500 500 510

- - - OCN- 500 500 -

3.3.3 Crystal Structure of (Fe+NOG+Cl)D201G

To gain insight into the metal geometry and examine the mode of interaction with the metal environment, crystallographic analysis was performed. Attempts were made to

- - - crystallize D201G in the presence of N3 , OCN , and Cl but unfortunately we were

- - unable to get well diffracting crystals of D201G in the presence of OCN and N3 .

Crystals were grown by the hanging drop vapor diffusion method in the presence of Fe2+,

N-oxalylglycine (NOG), 100 mM NaCl, and a 19 residue CTAD peptide corresponding to HIFα788-807, similar to previously described.11,16 NOG is a substrate analogue for αKG that binds in the same manner but inhibits chemistry,11 hence, it was used to avoid aberrant decarboxylation, which occurs ~ 6 fold faster in D201G compared to WT,16 yielding succinate and CO2.

The structure of (Zn+αKG)D201G/CTAD was previously reported,16 and revealed that the Zn center was 5 coordinate with a water molecule bound trans to the keto group of αKG at the carboxylate binding site (Figure 3.6A). Overall, our structure of

(Fe+NOG)D201G/CTAD was very similar to the structure of (Zn+αKG)D201G/CTAD with a rmsd = 0.184 Å when the backbones (residues 9 – 349 ) were aligned using

PyMoL, however, significant difference were observed at the active site. The structure of

(Fe+NOG)D201G/CTAD had unresolved density above the metal center and in the equatorial plane. The density was first modeled with two water molecules which resulted in significant positive density remaining post refinement, suggesting that water was not the correct model. We then tried one chloride at the carboxylate binding site and one water in the axial coordination position, as observed for the active site of SyrB2.15 The refinement indicated that chloride was an appropriate ligand but the axial site was still not

79 consistent with water as there was significant positive density observed. Next, chloride was placed in the axial position and water in the equatorial position. The refinement indicated that the model was consistent with chloride at the diffusible ligand site but water was insufficient to account for the electron density at the carboxylate binding site.

Figure 3.5: Crystal structure of (Fe+NOG+Cl)D201G. (A) The active site 2Fo – Fc map contoured to 1σ. (B) Active site of D201G showing the metal chloride bond lengths. (C) The 2Fo – Fc map contoured to 1σ showing the second coordination sphere residues that help position the target residue. (D) D201G active site second coordination sphere distances to N803

In the structure of (Fe+αKG)D201A, there was similar density above the metal center that was modelled with carbonate, assumed to result from oxidative decarboxylation during crystallization as D201A consumes αKG ~ 10 fold faster than

WT-FIH.16 Although we did not anticipate carbonate formation due to the use of NOG, we modeled the density with carbonate and observed that it refined well, but carbonate was bound asymmetrically in a bidentate fashion with unrealistic metal-ligand bond

80 lengths. The bond length was 1.80 Å for the O-atom near the axial position and 2.01 Å for the O-atom near the equatorial site, which does not compare well to the 2.4 Å (axial) and 2.6 Å (equatorial) Fe – O bond lengths for the carbonate ligand in the structure of

D201A.16

The density refined best with two chlorides (Figure 3.5A), one at the carboxylate binding site as seen in SyrB215 with a Fe – Cl bond length of 2.2 Å and another in the axial coordination position with a Fe – Cl bond length of 2.5 Å (Figure 3.5B). The Fe –

Cl bond length of 2.2 Å agrees well with the average Fe – Cl bond (2.2 Å – 2.3 Å) observed for 6-cordinate Fe2+ model complexes.50 The Fe – Cl bond length of 2.5 Å is slightly longer than the average Fe – Cl bond in model complexes but was similar to the

Fe – Cl bond (2.5 Å) in SyrB 2 (Figure 3.6D).15,50 An overlay of (Fe+αKG+Cl)SyrB2 and

(Fe+NOG+Cl)D201G/CTAD showed that the equatorial chloride ligands were in nearly the same position (Figure 3.6E). In SyrB2, there was a water molecule above the coordinated chloride which was within H-bonding distance to the coordinated axial aquo ligand (Figure 3.6D). This water molecule likely helps stabilize bound water in a similar fashion to the uncoordinated O-atom of the carboxylate residue in FIH and related enzymes. The axially ligated chloride in our structure was close to the position of the uncoordinated water molecule in SyrB2 (Figure 3.6E). Similarly, the structure of

(Zn+αKG)D201G/CTAD overlaid well with the structure of SyrB2 (Figure 3.6F) and showed that the equatorial aquo ligand in (Zn+αKG)D201G/CTAD was bound close to the Cl binding site in SyrB2 (Figure 3.6F). It appears that loss of the facial triad carboxylate opens a coordination site for other ligands to bind in the equatorial plane.

Figure 3.6: Structure of (Fe+NOG+Cl)D201G (green), (Zn+αKG)D201G (cyan), and (Fe+αKG+Cl)SyrB2 (white). (A) (Zn+αKG)D201G active site distances. (B) Comparison of (Zn+αKG)D201G (cyan) and (Fe+NOG+Cl)D201G (green). (C) Alternate view of (Zn+αKG)D201G (cyan) and (Fe+NOG+Cl)D201G (green) overlaid. (D) (Fe+αKG+Cl)SyrB2 active site distances. (E) Overlay of (Fe+αKG+Cl)SyrB2 (white) and (Fe+NOG+Cl)D201G (green). The Fe and Cl- for SyrB2 are colored grey and green respectively and the Fe and Cl- for D201G are colored magenta and yellow respectively. Waters are colored red (F) Overlay of (Fe+αKG+Cl)SyrB2 (white) and (Zn+αKG)D201G (cyan). The Fe and Cl- for SyrB2 are colored grey and green respectively and the Zn for D201G is colored light grey. Waters are colored red and the ones derived from SyrB2 are labelled.

A major difference that existed between the structure of (Zn+αKG)D201G/CTAD and our structure of (Fe+NOG+Cl)D201G/CTAD was the position of the target residue

(N803) and residues that help position the target residue (Q239 and R238) for chemistry

(Figure 3.6B). 51,52 Q239 formed a salt bridge with the target residue and R238 was H- bonded to the sidechain O-atom of N803 in the structure of (Zn+αKG)D201G/CTAD, as

82 seen for WT-FIH (Figure 3.6A).16,53 In the structure of (Fe+αKG+Cl)D201G, the target residue was shifted towards αKG, causing N803 to be out of optimal H-bonding distance with Q239 and R238 (Figure 3.5D). R238 and Q239 were best refined when split between two conformations, consistent with greater mobility as a result of the target residue being shifted (Figure 3.5C). With respect to the position of N803 in the

(Zn+αKG)D201G/CTAD structure, the Cα, Cβ, Cγ, and amine nitrogen were shifted towards NOG by 1.0 Å, 1.5 Å, 2.2 Å and 3.0 Å respectively in the

(Fe+NOG+Cl)D201G/CTAD structure, suggesting that chloride may play a role in positioning N803 for hydroxide rebound chemistry (Figure 3.6B).

- When overlaid, the positions of the Cl in the Fe-D201G structure and H2O in the

- Zn-D201G structure were ~ 4 Å from the Cα of G201, similar to the distance of the Cl ion in SyrB2 to the Cβ methyl group of A118 (~ 4 Å) (Figure 3.6C and D). Thus, it appears that in the absence of the carboxylate ligand, the active site of FIH provides enough room for exogenous ligands such as Cl- to bind. All other Fe – ligand bonds lengths were typical of Fe2+/αKG dependent enzymes and were all within 0.1 Å of each other for corresponding ligands in the (Fe+αKG+Cl)SyrB2, (Fe+NOG+Cl)D201G and

(Zn+αKG)D201G structures.

3.3.4 X-Ray absorption spectroscopy (XAS)

To further characterize the metal geometry, Fe K-edge XAS measurements were performed on a sample of (Fe+NOG+Cl)D201G in 50 mM Hepes pH 7.00 (100 mM

NaCl). Analysis of the XANES pre-edge features provides information on the geometry and coordination number of metal complexes. Although the transition is spin forbidden,

Fe2+ has vacancies in the 3d manifold, resulting in 1s → 3d electronic transitions that

83 give rise to the pre-edge features, which are sensitive to the geometric structure of the Fe center. The peak area of the pre-edge provides a measure of centrosymmetricity. Larger peak areas are associated with noncentrosymmetric geometries such as square pyramidal or distorted octahedral and small peak areas are observed for a centrosymmetric geometry such as octahedral.54–56

The XANES spectrum (Figure 3.7) exhibited an observable 1s → 3d pre-edge peak at ~ 7113 eV. The peak areas for octahedral geometries range from (3 – 7) x 10-2 eV.57 The peak area (0.12(4) eV) for (Fe+NOG+Cl)D201G was fairly large in comparison to the expected area for octahedral geometry, consistent with a noncentrosymmetric environment. This peak area was similar to the peak area (0.12(9) eV) of a square pyramidal ferrous model complex comprised of 4 N ligands and 1 Cl ligand.55 The large peak area can also be an indication of a 6 coordinate geometry that deviates from ideal octahedral geometry.55

Figure 3.7: Fe K-edge XANES spectrum of (Fe+NOG+Cl)D201G in 50mM Hepes pH

7.00.

The EXAFS region provides information on the coordination number (± 20%), ligand identities (Z ± 1), as well as the ligand distance from the metal center. The best

EXAFS fit (Table 3.2 and Figure 3.8) obtained consisted of three N/O donor ligands, one of which was an imidazole, one chloride, and one NOG fit as a unit to represent bidentate coordination through the O-atoms. The EXAFS analysis suggested a distorted octahedral geometry, which is consistent with the XANES analysis and the crystallographic data.

The three N/O donor bond lengths were 2.11(1) Å and the Cl- bond length was 2.51(3) Å, in agreement with the long Fe-Cl bond (2.5 Å) found in the crystal structure. NOG was modelled as a bidentate ligand and the Fe-O bond lengths were 1.89(1) Å and 1.99(1) Å, which deviate from the Fe-O bond lengths found for αKG (2.03 Å and 2.24 Å)

85 coordinated to Fe in PHD2,58 and the Fe-O bonds for NOG (2.06 Å and 2.24 Å) in our crystal structure of Fe-D201G. The Fe-Cl bond was also longer than observed for Fe-Cl bonds in model complexes (~2.3 Å),50 but the three N/O bond lengths were in agreement with typical bond lengths of N/O donors of αKG dependent enzymes.

The EXAFS region was difficult to fit owing to the heterogeneity in the sample.

Previous XANES analysis on PHD2 revealed that there was a mix of 5/6 coordinate Fe.58

Our best EXAFS fits indicated a distorted octahedral geometry, in agreement with the large pre-edge peak in the XANES spectrum and a noncentrosymmetric geometry. The best fits are the ones that yield lower R-factor and reduced χ2 values. Several fits were tried and based on R-factor and reduced χ2, the overall best fit suggested a model consisting of 3 N/O donors, 1.75 Cl, and NOG (Table 3.4 and Figure 3.9). This would result in a 7 coordinate Fe center which is unlikely, but suggests there is significant disorder at the metal center, and a second chloride may be near the Fe site. As all the best fits obtained involved chloride in the model (summarized in Table 3.5), it strongly suggests that chloride was bound to the Fe cofactor.

Table 3.2: EXAFS Analysis of (Fe+NOG+Cl)D201G

2 -3 2 2 2 N r (Å) σ (x10 Å ) ΔEo (eV) R factor χ Reduced χ

3N/O 2.11(1) 1(1)

(1 Imidazole)

1Cl 2.51(3) 7(3) 0(1) 0.0900 1976.10 126.35

1NOG-O1 1.89(1) 0(1)

1NOG-O2 1.99(1)

Figure 3.8: EXAFS analysis of (Fe+NOG+Cl)D201G in 50mM Hepes pH 7.00. Fourier transform (FT) XAS data with data in black and fit in red (see fit details in Table 3.2). Inset: Unfiltered k3-weighted EXAFS spectra for the fit (red) and data (black). Fits were performed in r-space (Δk=2 – 14Å-1; Δk=1 – 4.0Å) as described in materials and methods.

3.4 Discussion

The 2-His 1-carboxylate facial triad is a defining structural motif in the

Fe2+/αKG-dependent enzymes. The αKG-dependent halogenases coordinate Fe with only two protein derived His residues, and bind a chloride ligand in place of the facial triad carboxylate,14,15 suggesting that loss of the facial triad carboxylate permits binding of exogenous ligands to the Fe cofactor. In the consensus mechanism of αKG-dependent halogenases, ferryl formation leads to chloride rebound as opposed to hydroxide rebound, but the latter can occur as well. The partition between chloride rebound and hydroxide rebound is mainly dictated by substrate positioning, with substrates buried deeper in the active site favoring chlorination while more shallow substrates favor hydroxylation in

SyrB2.5,59 This suggests that with the proper coordination environment and substrate positioning, alternative rebound can be directed.

To gain insight into structural factors that influence ligand access, the facial triad carboxylate D201E and D201G constructs were expressed, purified, and studied using kinetics, spectroscopy and crystallography. Chloride was required for D201G activity but not for WT-FIH or D201E (Hangasky unpublished), suggesting that Cl- binds to the Fe cofactor in D201G to rescue activity. However, the change in the αKG-dependent MLCT in the presence of Cl- was within the noise of our instrument and could not be measured accurately. Hence structural studies were undertaken to investigate the presence of chloride on the metal environment.

While the D201E mutation is conservative, the D201G mutation creates an active site pocket that structurally mimics that of the αKG-dependent halogenases. In D201E, the longer sidechain may impose steric constraints that occlude water from the axial coordination site, resulting in a 5 coordinate geometry that reacts with oxygen and other ligands more readily than WT-FIH. The facial triad carboxylate has been implicated in being an H-bond partner for the axial water ligand and without the carboxylate, a weakened metal aquo bond results.19 With loss of the carboxylate ligand in the D201G variant, a 5-coordinate geometry would be favored, allowing O2 and other ligands access to the Fe cofactor. This line of reasoning is supported by the observation that D201E,

D201G, and D201A catalyze oxidative decarboxylation (~ 5 – 10 fold) significantly faster than WT FIH.16 Although they were able to catalyze oxidative decarboxylation, significant uncoupling was observed, suggesting that substrate positioning was not optimal for coupled rebound chemistry.

Our initial screen identified NO as an inhibitor of WT-FIH, D201E, and D201G,

- - N3 as an inhibitor of D201E and D201G, and OCN as an activator of D201G. The dose

88 response assays revealed that NO inhibited D201E (IC50 = 4 µM) the most, followed by

D201G (IC50 = 22 µM) and then WT (IC50 = 65 µM). This data suggested that mutation of the facial triad carboxylate allowed exogenous ligands better access to the active site of FIH, an expected outcome if the Fe centers in D201G and D201E favored a 5 coordinate state. Recently, NO was demonstrated to inhibit the HIF hydroxylase PHD2 by binding to the active site metal.43 NO was also recently demonstrated to inhibit FIH activity in cells via an unknown mechanism.44 As binding of NO resulted in electronic absorption transitions typical of NO binding to Fe2+ in non heme enzymes,39–41 we interpreted this to mean that NO bound to the Fe cofactor to inhibit chemistry.

Interestingly, the intensity of the peak at 440 nm due to NO binding was positively correlated to the degree of inhibition and followed the same trend, D201E >

D201G > WT-FIH, consistent with better access to the active site in the D201E and

D201G variants compared to WT-FIH. As the intensity depends on the amount of ligand character in the overlapping orbitals,41 it suggests that removal of the native carboxylate residue facilitates formation of stronger bonds with exogenous ligands. With the longer sidechain in D201E, the steric constraints may dictate the Fe ligand interaction, resulting in increased orbital overlap that gives rise to stronger Fe-ligand bonds compared to WT-

FIH. The lack of steric constraints in D201G would provide greater degrees of freedom for Fe-ligand interactions, which could result in the ligand adopting the most favorable conformation to enhance bonding interactions compared to WT.

- N3 inhibited D201G the most, and then D201E, but failed to inhibit WT. The

- larger size of N3 compared to NO probably contributed to this preference. The more open active site of D201G confers less steric barriers while D201E, even with a vacant

89 coordination site, would be expected to impose more steric barriers to ligand binding due

- to the longer sidechain. The electronic absorption spectra were consistent with N3

- - - - binding to D201E, and both N3 and OCN binding to D201G. In SyrB2, N3 and OCN binding caused red shifts in the MLCT centered at ~ 500 nm to 545 and 540

6 6 respectively. The binding affinity for azide (KD = 120 µM) in SyrB2 is comparable to

- the IC50 of inhibition in D201G and D201E. OCN enhanced D201G activity ~ 3 fold, and the MLCT shifted from 485 nm to 500 nm, as seen for WT FIH. This suggests a chemical rescue and that the active site environment of D201G with OCN- is similar to

WT with OCN- replacing the carboxylate ligand in D201G. We attribute the red shift to stabilization of the Fe t2g orbitals, likely a result of greater π symmetry overlap due to

OCN- binding in the equatorial position trans to the αKG keto group. Partial chemical rescue was previously observed for the D101A variant in TauD as 100 mM formate resulted in a ~4 fold increase in activity.48

The role of the active site carboxylate has previously been defined as an H-bond acceptor for the bound aquo ligand based on DFT and VTVH MCD and CD experiments on the chlorinating enzyme CytC3 and the hydroxylase TauD.19 Due to the anionic nature of the carboxylate, the O-H bond of bound H2O becomes polarized, leading to a shorter

Fe-O bond as more OH character is gained.19 The subsequent rearrangement of the carboxylate ligand to form new H-bond contacts with CTAD upon substrate binding

11 weakens the Fe-H2O bond, stimulating aquo release and allowing O2 binding, as suggested by computations and spectroscopy on FIH.52

In both the CytC3 and TauD, the MCD and CD data revealed that the active site

Fe was comprised of a distorted octahedral geometry, consistent with a weakly bound

2+ H2O ligand, and switched to a mixture of 6/5-coordinate Fe upon substrate binding with

αKG coordinated in a bidentate fashion throughout.19 Our XAS analysis and crystallography are consistent with a distorted octahedral site for D201G, which structurally resembles CytC3 at the active site. The XANES region analysis suggested a mixture of 5/6 coordinate Fe centers, consistent with the existence of 2 distinct ferrous species seen in the MCD features of CytC3 and TauD.19 The EXAFS analysis best fit involved the coordination of Fe by 3 N/O donors, one of which was an imidazole, 1 Cl-, and NOG. The active site of (Zn+αKG)D201G/CTAD was reported to be 5-coordinate with the Fe coordinating ligands being 2 His residues, αKG in a bidentate fashion, and

- one H2O molecule. In our structure of (Fe+NOG+Cl )D201G, the active site is similar, but instead of an aquo ligand, there were two Cl- ligands bound to the Fe center. Perhaps the use of the native metal provides the proper electronic environment for Cl- binding, or it’s a result of the crystallization conditions.

Although carbonate was a good fit during refinement, we excluded carbonate binding based on the fact that (i) the presence of 2 mM carbonate in our steady state assays did not inhibit D201G activity, (ii) the refinement of carbonate leads to asymmetric binding with unusual Fe-O bond lengths, (iii) the use of NOG prevents oxidative decarboxylation, limiting CO3 formation, and (vi) the EXAFS analysis was consistent with the presence of chloride in the Fe coordination sphere. Hence, we assigned the unresolved density tentatively to two chlorides as the refinement suggested this was the best model. We cannot rule out the possibility that some other unknown ligand generated during crystallization was bound. The discrepancy in the structure and

EXAS analysis is attributed to more heterogeneity in the XAS sample compared to the

91 well-ordered crystal used for crystallography. In solution, a mix of different states (5/6- coordinate geometries) are averaged in the XAS analysis, resulting in the data fitting best with a single Cl- model. The heterogeneity also contributes to the different bond lengths observed crystallographically and by XAS. Interestingly, the r-factor and reduced χ2 were lowered when the model was expanded to included ~ 1 extra Cl-, however this would yield a 7-coordinate geometry which is unlikely.

According to the consensus mechanism, an open coordination site must exist for

2 - O2 binding in order for oxidative decarboxylation to occur. Hence, one Cl ligand is expected to dislodge and open a coordination site for O2. The long Fe-Cl bond found in the axial position, suggests that this Cl- ligand is loosely associated with Fe and can be displaced by O2. The net result would be a 6C geometry in which O2 is bound in the expected position for ferryl formation to lead to hydroxylation. It is also possible that the

- Cl bound trans to the keto group of NOG vacates to yield an O2 binding site, however, this would lead to O2 being incorrectly positioned for decarboxylation chemistry and ferryl formation would occur at a position cis to the relative position of the target residue.

In clavaminate synthase, the O2 mimic NO was bound to a site cis to the diffusible ligand site,60 and it cannot be ruled out as a possibility in D201G.

It appears that Cl- binding leads to changes in the positioning of the target residue, hinting at a more dominant role in sterics as opposed to tuning the electronic environment of the Fe cofactor, consistent with the lack of observable spectral changes in the presence of chloride. The position of N803 is shifted towards αKG when compared to the position in WT-FIH,11 and to the position in the (Zn+αKG)D201G structure.16 The B-factor for

N803 was high (~ 100 Å2) compared to the average B-factor (~ 67 Å2), consistent with

92 increased mobility of N803. We also observed alternate conformations for the substrate stabilizing residues, suggesting increased flexibility in the N803 positioning. We propose that Cl- rescues activity by shifting the target residue to an optimal position for hydroxide rebound chemistry. Rescue of D201G activity was not observed in the presence of NaF or

NaBr (Hangasky unpublished), suggesting F- and Br- fail to provide the same steric environment as Cl-. It’s been suggested that substrate positioning is the dominant model to explain the preference for OH- and Cl- rebound.5,59 Thus, rebound chemistry in FIH may be tunable with the proper geometry and active site environment. Unfortunately, we

- - did not detect the transfer of Cl , N3 , or any other non-native ligand. In SyrB2, nitration chemistry was enhanced ~ 2.5 fold and azidation was enhanced ~ 13 fold in the A118G variant in the absence of Cl-,6 implying that alternative rebound chemistry can be achieved and improved.

3.5 Conclusion

Taken together with previous studies on TauD and CytC3, which lack a carboxylate ligand that coordinates to the axial H2O molecule, our data implicates the facial triad carboxylate in controlling ligand access to the Fe cofactor in the Fe2+/αKG dependent oxygenases. In order for halogenation chemistry to occur, the active site must incorporate a halide ion to the Fe cofactor, generate the ferryl intermediate, perform HAT on the substrate, and transfer the bound halide to the substrate radical. Our kinetic, spectroscopic and crystallographic data are consistent with the binding of a halide (Cl) and other non-native ligands to the Fe cofactor in D201G. D201G also satisfies the prerequisite for ferryl formation, and HAT, but halide transfer is impeded. The lack of alternative rebound observed in D201G suggests that substrate positioning is not

93 optimized for alternative rebound, which perhaps may be achieved with further mutagenesis of the substrate binding residues (Q239 and R238). FIH provides an ideal model system for “biorthogonal” conjugation chemistry as it has been shown to hydroxylate other residues in non-native substrates, including His, Asp, Ile, Leu, and

Ser.13,61,62

3.6 Appendix

3.6.1 Supplemental

Table 3.3: Data collection and refinement statistics for D201G. Values in parentheses represent the highest resolution shell Data set (Fe2++NOG+Cl)D201G λ (Å) 1.08 Space group P41212 Unit cell (Å, °) a = b = 86.42, c = 148.06 α = β = γ = 90 Resolution (Å) 74 – 2.6 (2.64 – 2.6) Rmerge (%) 7.7 Mean I/σ (I) 28.3 (2.4) Completeness 99.27 (99.92) Multiplicity Refinement Resolution range (Å) 74 – 2.6 Number of Reflections 15948 Average B factor (Å2) 66.5 Rwork/Rfree (%) 19.04/25.22 R.m.s deviations Bond lengths (Å) 0.014 Bond angles (Å) 1.565 Ramachandran plot, residues in (%) Preferred region 93.02 Allowed region 6.69 Disallowed region 0.29

Table 3.4: EXAFS Analysis of (Fe+NOG+Cl)D201G with 1.75 Cl

2 -3 2 2 2 N r (Å) σ (x10 Å ) ΔEo (eV) R factor χ Reduced χ

3N/O 2.11(1) 1(1)

(1Imidazole)

1.75Cl 2.53(3) 13(4) 0(1) 0.0840 1786.23 114.21

1NOG-O1 1.89(1) 0(1)

1NOG-O2 1.99(1)

Figure 3.9: EXAFS analysis of (Fe+NOG+Cl)D201G for 1 and 1.75 Cl in 50mM Hepes pH 7.00. Fourier transform (FT) window of 2 – 14 Å-1 with data in black and fits in red (1Cl) and blue (1.75 Cl). Fit details for 1 Cl are in Table 3.2 and details for 1.75 Cl are in Table 3.4). Inset: Unfiltered k3-weighted EXAFS spectra for the fits and data. Fits were performed in r-space (Δk=2 – 14Å-1; Δk=1 – 4.0Å) as described in materials and methods.

Table 3.5: Table of EXAFS fits (continued onto next few pages)

Reduced 2 -3 2 2 2 N r (Å) σ (x10 Å ) ΔEo (eV) R factor χ χ #Var 2N/O 2.11(3) 1(2) 2(4) 0.2717 3413.21 416.09 3 3N/O 2.10(3) 4(2) 1(2) 0.1744 3089.66 376.64 3 4N/O 2.09(2) 7(2) 0(2) 0.1333 2159.28 263.23 3 5N/O 2.08(3) 11(2) -1(2) 0.1237 1960.34 238.98 3 6N/O 2.08(3) 14(3) -2(2) 0.1321 2145.33 261.53 3 7N/O 2.07(4) 18(3) -2(2) 0.1494 2506.26 305.53 3

Fe-S(2.2) Reduced 2 -3 2 2 2 N r (Å) σ (x10 Å ) ΔEo (eV) R factor χ χ #Var 2S 2.20(4) 6(3) -15(6) 0.3543 7216.30 879.70 3 3S 2.21(4) 10(3) -15(5) 0.2736 5347.59 651.90 3 4S 2.21(4) 13(3) -15(4) 0.2262 4260.11 519.33 3 5S 2.22(4) 16(3) -15(4) 0.1980 3611.67 440.28 3 6S 2.22(4) 19(3) -15(4) 0.1816 3231.16 393.89 3 7S 2.23(4) 21(3) -15(3) 0.1730 3021.43 368.33 3

Fe-Cl(2.3) Reduced 2 -3 2 2 2 N r (Å) σ (x10 Å ) ΔEo (eV) R factor χ χ #Var 2Cl 2.18(4) 7(3) -14(6) 0.3312 6666.47 812.68 3 3Cl 2.18(4) 12(3) -15(5) 0.2498 4812.27 586.64 3 4Cl 2.19(4) 14(3) -15(4) 0.2050 3800.75 463.33 3 5Cl 2.19(4) 17(3) -15(4) 0.1808 3247.88 395.93 3 6Cl 2.20(4) 20(3) -15(3) 0.1687 2960.08 360.85 3 7Cl 2.20(4) 23(3) -15(3) 0.1639 2831.02 345.11 3

Fe -N(2.0) Fe- N(2.2) Reduced 2 -3 2 2 2 N r (Å) σ (x10 Å ) ΔEo (eV) R factor χ χ #Var 2N/O 2.01(3) 1(3) 1N/O 2.13(2) -4(2) -2(2) 0.1013 1980.93 319.34 5

2N/O 2.11(2) -1(1) 2N/O 1.97(2) 2(2) -3(2) 0.0649 1227.08 197.82 5

3N/O 2.01(2) 5(3) 1N/O 2.13(2) -3(1) -3(2) 0.0692 1302.00 209.89 5

4N/O 2.01(3) 10(3) 1N/O 2.12(2) -2(2) -3(2) 0.0709 1326.12 213.78 5

3N/O 1.98(3) 6(3) 2N/O 2.12(2) 0(2) -4(2) 0.0764 1448.24 233.47 5

5N/O 2.10(2) 10(2) 1N/O 2.38(3) -1(3) 1(2) 0.0991 1690.20 272.50 5

4N/O 2.09(3) 8(4) 2N/O 2.32(22) 37(75) 1(4) 0.1237 1957.01 315.49 5

3N/O 2.27(32) 42(67) 3N/O 2.09(3) 6(3) 1(6) 0.1302 2090.02 336.93 5

6N/O 2.09(3) 13(3) 1N/O 2.38(3) -1(3) 0(2) 0.1031 1831.88 295.32 5

5N/O 2.09(3) 10(3) 2N/O 2.42(12) 26(39) 1(3) 0.1226 1952.41 314.75 5

4N/O 2.09(3) 8(3) 3N/O 2.36(17) 46(69) 2(4) 0.1240 1975.94 318.54 5

Fe-S(2.2) Fe- S(2.3) Reduced 2 -3 2 2 2 N r (Å) σ (x10 Å ) ΔEo (eV) R factor χ χ #Var 2S 2.22(4) 4(3) 1S 2.40(7) 3(6) -10(5) 0.2493 4980.06 802.83 5

2S 2.20(4) 5(3) 2S 2.38(6) 9(7) -10(4) 0.2006 3804.79 613.37 5

3S NO FIT 1S 5

4S NO FIT 1S 5

3S 2.21(3) 7(2) 2S 2.41(5) 7(5) -10(3) 0.1581 2902.37 467.89 5

5S NO FIT 1S 5

4S 2.23(3) 10(2) 2S 2.44(4) 7(4) -11(3) 0.1339 2378.41 383.42 5

3S 2.20(3) 8(3) 3S 2.41(5) 12(6) -10(3) 0.1435 2515.15 405.47 5

6S NO FIT 1S 5

5S 2.23(3) 12(2) 2S 2.45(4) 7(4) -11(3) 0.1224 2119.77 341.73 5

4S 2.21(3) 10(2) 3S 2.43(5) 12(5) -11(3) 0.1298 2209.60 356.22 5

Fe -N(2.0) Fe- S(2.2) Reduced 2 -3 2 2 2 N r (Å) σ (x10 Å ) ΔEo (eV) R factor χ χ #Var 2N/O 1.95(6) 4(5) 1S 2.23(4) 2(3) -11(5) 0.2237 4514.23 727.73 5

1N/O 1.89(4) 0(4) 2S 2.22(4) 6(3) -14(4) 0.2065 4238.71 683.32 5

2N/O 1.95(7) 6(6) 2S 2.25(5) 8(4) -11(5) 0.2001 3670.53 591.72 5

3N/O 2.09(3) 4(2) 1S 2.60(23) 26(39) 1(3) 0.1547 2605.88 420.09 5

1N/O NO FIT 3S 5

1N/O NO FIT 4S 5

4N/O 2.09(3) 7(2) 1S 2.69(16) 23(31) 1(2) 0.1222 1915.89 308.86 5

2N/O 1.98(8) 8(7) 3S 2.29(6) 15(7) -9(5) 0.1838 3164.87 510.21 5

3N/O 2.09(3) 4(2) 2S 2.61(18) 35(31) 1(3) 0.1451 2383.32 384.21 5

5N/O 2.08(5) 17(10) 1S 1.82(6) 17(12) 1(3) 1(3) 1730.69 279.00 5

1N/O NO FIT 5S

4N/O 2.09(3) 7(2) 2S 2.70(14) 34(29) 1(2) 0.1190 1851.80 298.53 5

2N/O NO FIT 4S 5

3N/O 2.09(3) 5(2) 3S 2.62(17) 43(28) 1(3) 0.1392 2253.89 363.35 5

6N/O 2.06(2) 2(1) 1S 2.02(2) 0(1) -3(1) 0.0630 1220.85 196.81 5

1N/O NO FIT 6S 5

5N/O 2.09(3) 11(3) 2S 2.75(12) 31(27) 0(2) 0.1175 1851.59 298.49 5

2N/O 2.00(7) 8(9) 5S 2.30(8) 25(8) -9(5) 0.1639 2728.01 439.78 5

4N/O 2.09(3) 7(3) 3S 2.71(14) 43(31) 1(2) 0.1179 1832.09 295.35 5

3N/O NO FIT 4S 5

Fe-N(2.0) Fe- Cl(2.3) Reduced 2 -3 2 2 2 N r (Å) σ (x10 Å ) ΔEo (eV) R factor χ χ #Var 2N/O 1.94(5) 4(4) 1Cl 2.21(3) 2(2) -11(5) 0.1964 3999.91 644.82 5

1N/O 1.89(4) 0(3) 2Cl 2.19(3) 6)2) -14(4) 0.1770 3640.83 586.94 5

2N/O 1.92(6) 6(5) 2Cl 2.22(4) 8(3) -12(4) 0.1769 3313.50 534.17 5

3N/O 2.09(3) 4(3) 1Cl 2.57(22) 28(39) 1(3) 0.1557 2625.13 423.19 5

1N/O 1.88(4) 1(3) 3Cl 2.20(3) 10(3) -14(3) 0.1555 3018.69 486.64 5

1N/O 1.88(5) 2(4) 4Cl 2.21(4) 15(3) -14(3) 0.1551 2862.09 461.40 5

4N/O 2.09(3) 7(2) 1Cl 2.66(17) 25(33) 1(2) 0.1220 1908.59 307.68 5

2N/O 1.95(9) 8(8) 3Cl 2.25(6) 15(6) -10(5) 0.1740 3029.63 488.40 5

3N/O 2.09(3) 5(3) 2Cl 2.58(18) 38(31) 1(3) 0.1466 2411.81 388.81 5

5N/O 2.09(3) 11(3) 1Cl 2.71(12) 20(23) 0(2) 0.1162 1815.24 292.63 5

1N/O 1.90(8) 5(8) 5Cl 2.23(5) 19(4) -13(4) 0.1639 2866.47 462.10 5

4N/O 2.09(3) 7(2) 2Cl 2.66(14) 37(30) 1(2) 0.1186 1839.57 296.56 5

2N/O 1.98(9) 9(10) 4Cl 2.26(7) 21(8) -10(5) 0.1677 2844.58 458.57 5

3N/O 2.09(3) 5(3) 3Cl 2.58(18) 45(29) 1(3) 0.1411 2282.61 367.98 5

6N/O 2.06(5) 20(11) 1Cl 1.79(7) 15(9) 0(3) 0.1176 1807.51 291.39 5

1N/O 1.68(26) 28(82) 6Cl 2.18(5) 19(4) -17(6) 0.1621 2818.61 454.39 5

5N/O 2.09(3) 11(3) 2Cl 2.72(13) 34(29) 0(2) 0.1170 1837.87 296.28 5

2N/O 2.0097) 9(10) 5Cl 2.27(8) 26(9) -9(5) 0.1633 2732.65 440.53 5

4N/O 2.09(3) 7(3) 3Cl 2.67(14) 45(31) 1(2) 0.1172 1813.00 292.27 5

3N/O 2.09(3) 5(3) 4Cl 2.59(18) 51(27) 1(3) 0.1371 2196.50 354.10 5

Best 1st shell (R @ 1 - 4.0)

Fe -N(2.0) Reduced 2 -3 2 2 2 N r (Å) σ (x10 Å ) ΔEo (eV) R factor χ χ #Var 2N/O 2.11(2) 1(2) 3(3) 0.3336 7481.57 380.94 3 3N/O 2.10(2) 4(2) 1(2) 0.2434 5122.66 260.83 3 4N/O 2.09(2) 7(2) 0(2) 0.2075 4191.65 213.43 3 5N/O 2.08(2) 11(2) -1(2) 0.2011 4030.41 205.22 3 6N/O 2.08(3) 14(3) -1(2) 0.2109 4268.92 217.36 3 7N/O 2.08(3) 18(3) -2(2) 0.2285 4688.34 238.72 3

100

Fe-S(2.2) Reduced 2 -3 2 2 2 N r (Å) σ (x10 Å ) ΔEo (eV) R factor χ χ #Var 2S 2.21(3) 6(2) -14(5) 0.4230 9714.92 494.66 3 3S 2.21(3) 10(2) -15(4) 0.3491 7787.69 396.53 3 4S 2.22(3) 13(2) -15(3) 0.3055 6658.88 339.05 3 5S 2.22(3) 16(2) -14(3) 0.2794 5980.22 304.50 3 6S 2.23(3) 19(2) -14(3) 0.2640 5578.25 284.03 3 7S 2.23(3) 21(3) -14(3) 0.2556 5354.10 272.62 3

Fe-Cl(2.3) Reduced 2 -3 2 2 2 N r (Å) σ (x10 Å ) ΔEo (eV) R factor χ χ #Var 2Cl 2.18(3) 7(2) -14(4) 0.4029 9172.66 467.05 3 3Cl 2.19(30 11(2) -15(4) 0.3282 7250.07 369.15 3 4Cl 2.19(3) 14(2) -15(3) 0.2867 6189.14 315.13 3 5Cl 2.19(3) 17(2) -15(3) 0.2637 5598.90 285.08 3 6Cl 2.20(3) 20(2) -15(3) 0.2517 5282.66 268.98 3 7Cl 2.20(3) 23(3) -15(3) 0.2464 5132.10 261.31 3

Fe -N(2.0) Fe- N(2.2) Reduced 2 -3 2 2 2 N r (Å) σ (x10 Å ) ΔEo (eV) R factor χ χ #Var 2N/O 2.01(2) 1(3) 1N/O 2.13(2) -4(1) -1(2) 0.1923 4193.62 237.74 5

2N/O 1.97(2) 2(2) 2N/O 2.12(2) -1(1) -2(1) 0.1619 3471.66 196.81 5

3N/O 2.02(2) 6(3) 1N/O 2.13(2) -3(1) -2(2) 0.1629 3481.66 197.38 5

4N/O 2.02(3) 10(3) 1N/O 2.12(2) -2(2) -3(2) 0.1648 3519.50 199.52 5

3N/O 1.98(3) 7(3) 2N/O 2.12(2) 0(2) -3(2) 0.1735 3721.70 210.99 5

5N/O 2.10(2) 1092) 1N/O 2.38(2) -1(2) 2(1) 0.1750 3637.89 206.23 5

4N/O 2.09(2) 7(3) 2N/O 2.35(11) 28(36) 2(2) 0.1996 4000.42 226.79 5

3N/O 2.09(2) 5(2) 3N/O 2.27(21) 40(37) 2(4) 0.2027 4081.25 231.37 5

101

6N/O 2.09(2) 13(2) 1N/O 2.38(2) -1(2) 1(1) 0.1827 3863.80 219.04 5

5N/O 2.10(3) 10(3) 2N/O 2.43(9) 22(25) 2(2) 0.2010 4044.92 229.31 5

4N/O 2.10(3) 8(3) 3N/O 2.38(11) 41(42) 2(3) 0.2015 4058.62 230.09 5

Fe-S(2.2) Fe- S(2.3) Reduced 2 -3 2 2 2 N r (Å) σ (x10 Å ) ΔEo (eV) R factor χ χ #Var 2S 2.22(3) 4(2) 1S 2.40(4) 3(4) -10(3) 0.3216 7273.49 412.34 5

2S 2.21(2) 4(2) 2S 2.39(4) 8(5) -10(3) 0.2748 6024.33 341.52 5

3S NO FIT 1S 5

4S NO FIT 1S 5

3S 2.22(2) 7(2) 2S 2.42(3) 7(3) -10(2) 0.2353 5056.08 286.63 5

5S NO FIT 1S 5

4S 2.23(2) 9(2) 2S 2.44(3) 7(3) -10(2) 0.2133 4508.13 255.57 5

3S 2.41(4) 11(4) 3S 2.21(2) 7(2) -10(2) 0.2291 4826.72 273.63 5

6S NO FIT 1S 5

5S NO FIT 2S 5

4S 2.22(2) 10(2) 3S 2.44(4) 11(4) -10(2) 0.2087 4323.29 245.09 5

Fe -N(2.0) Fe- S(2.2) Reduced 2 -3 2 2 2 N r (Å) σ (x10 Å ) ΔEo (eV) R factor χ χ #Var

102

2N/O 1.95(4) 5(4) 1S 2.24(3) 2(2) -10(4) 0.3079 6946.53 393.80 5

1N/O 1.89(3) 1(3) 2S 2.22(3) 6(2) -14(30 0.2949 6706.86 380.21 5

2N/O 1.98(6) 7(5) 2S 2.28(4) 10(4) -9(4) 0.2853 6026.54 341.65 5

3N/O 2.10(2) 4(2) 1S 2.66(13) 23(25) 2(2) 0.2201 4515.90 256.01 5

1N/O NO FIT 3S 5

1N/O NO FIT 4S 5

4N/O 2.10(2) 7(2) 1S 2.75(6) 12(10) 2(1) 0.1897 3779.28 214.25 5

2N/O 1.99(5) 7(5) 3S 2.30(5) 17(6) -8(3) 0.2625 5421.11 307.33 5

3N/O 2.10(2) 4(2) 2S 2.66(10) 31(19) 2(2) 0.2089 4230.68 239.84 5

5N/O 2.08(4) 18(9) 1S 1.81(5) 15(8) 2(2) 0.1896 3732.57 211.60 5

1N/O NO FIT 5S

4N/O 2.10(2) 7(2) 2S 2.72(8) 27(16) 2(2) 0.1871 3698.66 209.68 5

2N/O 2.61(3) 1(3) 4S 2.19(3) 13(2) -17(3) 0.2390 5196.69 294.60 5

3N/O 2.10(2) 4(2) 3S 2.66(10) 38(18) 2(2) 0.2031 4082.57 231.44 5

6N/O 2.17(3) 1395) 1S 2.22(3) 4(3) 2(2) 0.1956 4008.08 227.22 5

1N/O NO FIT 6S 5

103

5N/O 2.10(2) 11(2) 2S 2.74(7) 24(14) 1(2) 0.1901 3787.62 214.72 5

2N/O 2.01(5) 8(6) 5S 2.32(6) 26(7) -8(3) 0.2413 4907.27 278.20 5

4N/O 2.09(2) 7(2) 3S 2.71(9) 36(17) 1(2) 0.1873 3703.65 209.96 5

3N/O NO FIT 4S 5

Fe-N(2.0) Fe- Cl(2.3) Reduced 2 -3 2 2 2 N r (Å) σ (x10 Å ) ΔEo (eV) R factor χ χ #Var 2N/O 1.94(4) 4(3) 1Cl 2.22(2) 2(2) -11(3) 0.2857 6457.88 366.10 5

1N/O 1.89(3) 0(2) 2Cl 2.20(2) 6(2) -14(3) 0.2694 6109.05 346.32 5

2N/O 1.93(5) 6(4) 2Cl 2.23(3) 8(3) -11(3) 0.2654 5719.62 324.25 5

3N/O 2.10(2) 4(2) 1Cl 2.62(14) 25(26) 2(2) 0.2208 4528.37 256.72 5

1N/O 1.88(3) 0(3) 3Cl 2.21(3) 10(2) -14(3) 0.2471 5430.00 307.83 5

1N/O 1.88(4) 2(3) 4Cl 2.22(3) 15(3) -13(3) 0.2437 5222.85 296.09 5

4N/O 2.10(2) 7(2) 1Cl 2.68(10) 20(19) 1(2) 0.1912 3787.75 214.73 5

2N/O 1.97(6) 8(6) 3Cl 2.26(5) 16(5) -9(4) 0.2565 5326.20 301.94 5

3N/O 2.10(2) 4(2) 2Cl 2.62(11) 34(21) 2(2) 0.2106 4262.60 241.65 5

5N/O 2.10(2) 11(2) 1Cl 2.73(7) 14(12) 1(1) 0.1887 3742.77 212.18 5

1N/O 1.92(8) 6(8) 5Cl 2.23(4) 2(4) -12(3) 0.2480 5168.47 293.00 5

104

4N/O 2.09(2) 7(2) 2Cl 2.67(9) 31(18) 1(2) 0.1874 3695.66 209.51 5

2N/O 1.99(6) 8(7) 4Cl 2.28(6) 22(6) -9(4) 0.2473 5075.15 287.71 5

3N/O 2.09(2) 4(2) 3Cl 2.62(11) 42(19) 1(2) 0.2053 4119.70 233.55 5

6N/O 2.06(4) 22(10) 1Cl 1.79(5) 14(6) 0(2) 0.1961 3878.72 219.89 5

1N/O 1.68(24) 31(80) 6Cl 2.19(4) 19(3) -16(5) 0.2463 5150.91 292.01 5

5N/O 2.09(2) 11(2) 2Cl 2.71(8) 28(16) 1(2) 0.1906 3793.99 215.08 5

2N/O 2.01(5) 8(7) 5Cl 2.29(6) 28(7) -8(3) 0.2415 4919.72 278.90 5

4N/O 2.09(2) 7(2) 3Cl 2.67(10) 41(19) 1(2) 0.1873 3691.53 209.27 5

3N/O 2.09(2) 5(2) 4Cl 2.61(11) 49(19) 1(2) 0.2022 4034.03 228.69 5

Best 1st shell + imid (R @ 1 - 4.0)

Fe -Im(1Q5Y) - NOG(4V2W) - 4 Reduced 2 -3 2 2 2 N r (Å) σ (x10 Å ) ΔEo (eV) R factor χ χ #Var 4N/O 2.10(2) 6(2) (1Im0) 4(2) 0.2437 5167.57 263.12 3

3N/O 2.04(2) 6(3) 1Im0 2.12(2) -1(1) -1(1) 0.1332 2828.56 160.35 5

4N/O 2.08(2) 6(2) (2Im0) 2(2) 0.2771 6000.68 305.54 3

2N/O 2.00(3) 4(3) 2Im0 2.11(2) 1(1) -1(1) 0.1744 3817.51 216.42 5

3N/O 2.00(2) 4(2) (1Im0) 1Im0 2.12(1) -2(1) -1(2) 0.1758 3929.48 222.76 5

105

2N/O 2.03(3) 5(5) 1Im0 2.11(2) -2(1) 1Im0 1.98(5) 3(5) -2(1) 0.1375 3008.11 192.34 7

3N/O 2.00(2) 4(2) (1Im0) 1Im7 2.12(1) -3(1) -1(2) 0.1707 3762.48 213.30 5

2N/O 2.01(3) 5(4) 1Im0 2.01(5) 3(7) 1Im7 2.12(2) -3(1) -2(1) 0.1400 3022.98 193.29 7

4N/O 2.07(2) 6(2) (2Im7) 0(2) 0.2755 5873.01 299.04 3

2N/O 1.99(2) 3(3) 2Im7 2.11(2) 0(1) -1(1) 0.1762 3822.12 216.68 5

3N/O 2.00(2) 4(2) (1Im7) 1Im7 2.11(1) -3(1) -1(2) 0.1740 3854.21 218.50 5

2N/O 2.04(4) 6(6) 1Im7 2.11(2) -3(1) 1Im7 1.97(5) 1(4) -2(1) 0.1382 2991.25 191.26 7

2N/O 2.12(2) 0(2) 1NOG-O1 1.92(3) 1NOG-O2 2.01(3) 2(3) -1(2) 0.2085 4728.08 268.04 5

2N/O 2.11(2) 0(1) (1Im0) 1NOG-O1 1.92(2) 1NOG-O2 2.02(2) 2(3) 0(2) 0.1928 4516.26 256.03 5

1N/O 1.90(2) -2(1) 1Im0 2.07(2) 0(2) 1NOG-O1 1.98(2) 1NOG-O2 2.08(2) 0(2) -4(2) 0.1422 3258.72 208.36 7

2N/O 2.11(2) 0(1) (1Im7) 1NOG-O1 1.92(2) 1NOG-O2 2.02(2) 1(2) 0(2) 0.2010 4660.57 264.21 5

1N/O 2.14(3) -2(2) 1NOG-O1 1.91(3) 1NOG-O2 2.01(3) 1(5) -1(2) 0.1702 3904.46 249.65 7

106

1Im7

3N/O 1.96(4) 7(3) (1Im0) 1Cl 2.23(3) 3(2) -8(3) 0.2708 5895.08 334.19 5

2N/O 2.00(2) 3(2) 1Cl 2.38(6) 13(8) 1Im0 2.10(2) -2(1) -3(2) 0.1480 3148.39 201.31 7

3N/O 1.96(4) 7(3) (1Im7) 1Cl 2.23(3) 3(2) -8(3) 0.2715 5957.39 337.73 5

2N/O 1.99(2) 2(2) 1Cl 2.37(5) 11(7) 1Im7 2.10(2) -3(1) -3(2) 0.1487 3155.21 201.74 7

3N/O 1.98(4) 8(3) (2Im0) 1Cl 2.24(3) 4(2) -7(3) 0.2651 5686.02 322.34 5

1N/O 1.93(2) -1(2) 1Cl 2.32(5) 11(7) 2Im0 2.07(2) 0(1) -4(2) 0.1911 4158.21 265.88 7

2N/O 1.95(2) 1(2) (1Im0) 1Im0 2.09(2) -3(1) 1Cl 2.31(4) 9(5) -4(2) 0.1577 3450.59 220.63 7

1N/O 1.98(5) 3(6) 1Cl 2.32(4) 10(6) 1Im0 1.95(4) 0(4) 1Im0 2.09(2) -3(1) -4(2) 0.1388 3007.96 220.53 9

3N/O 1.98(4) 7(4) (2Im7) 1Cl 2.24(3) 4(3) -7(3) 0.2620 5607.41 317.89 5

1N/O 1.92(2) -2(1) 1Cl 2.33(5) 10(6) 2Im7 2.06(2) 0(1) -4(1) 0.1689 3682.88 235.48 7

2N/O NO FIT (1Im7) 1Im7 1Cl

107

1N/O 1.97(5) 2(7) 1Cl 2.33(1) 35(43) 1Im7 1.96(2) -4(1) 1Im7 2.09(4) -1(4) -4(2) 0.1323 2861.62 209.80 9

2N/O 1.96(2) 1(2) (1Im0) 1Im7 2.09(2) -4(1) 1Cl 2.33(4) 8(5) -4(2) 0.1531 3296.78 210.80 7

1N/O 2.02(4) 0(5) 1Cl 2.32(5) 10(7) 1Im0 2.09(1) -2(1) 1Im7 1.92(3) -1(2) -4(2) 0.1412 3079.45 225.77 9

1Cl 2.19(3) 2(2) 1Im0 1.71(9) 10(12) 1NOG-O1 1.85(4) 1NOG-O2 1.95(4) 1(2) -15(3) 0.2230 5066.99 323.99 7

1Cl 2.22(3) 4(3) 1NOG-O1 1.92(6) 1NOG-O2 2.02(6) 3(4) 1Im7 1.86(6) 3(5) -10(4) 0.2376 5280.73 337.65 7

Fe-Im(1Q5Y) - NOG(4V2W) - 5 Reduced 2 -3 2 2 2 N r (Å) σ (x10 Å ) ΔEo (eV) R factor χ χ #Var 5N/O 2.09(2) 8(2) (1Im0) 3(2) 0.2158 4416.64 224.88 3

4N/O 2.04(2) 10(2) 1Im0 2.11(1) -1(1) -1(1) 0.1112 2313.84 131.17 5

5N/O 2.08(2) 8(2) (2Im0) 1(2) 0.2314 4822.40 245.54 3

3N/O 2.01(2) 8(3) 2Im0 2.11(2) 2(1) -2(1) 0.1325 2806.18 159.08 5

4N/O 2.01(2) 7(2) (1Im0) 1Im0 2.12(1) -2(1) -1(1) 0.1318 2862.90 162.30 5

3N/O 2.03(3) 9(4) 1Im0 1.98(5) 5(6) -2(1) 0.1049 2232.75 142.76 7

108

1Im0 2.11(2) -1(1)

4N/O 2.01(2) 7(2) (1Im0) 1Im7 2.12(1) -2(1) -1(1) 0.1342 2873.71 162.91 5

3N/O 2.05(3) 9(4) 1Im0 2.11(2) -1(1) 1Im7 1.95(5) 3(4) -2(1) 0.1069 2290.43 146.45 7

3N/O 2.09(3) 5(3) 1NOG-O1 2.09(18) 1NOG-O2 2.18(18) 27(24) 1(3) 0.2185 4468.49 253.32 5

3N/O 2.12(2) 2(1) (1Im0) 1NOG-O1 1.91(2) 1NOG-O2 2.00(2) 2(2) 0(2) 0.1718 3923.04 222.40 5

3N/O (2Im0) 1NOG-O1 1NOG-O2

2N/O 1.98(5) 4(3) 1Im0 2.10(2) -2(1) 1NOG-O1 2.03(5) 1NOG-O2 2.12(5) 9(10) -2(2) 0.1252 2675.61 171.08 7

1N/O 1.89(2) -2(1) 2Im0 2.07(2) 3(2) 1NOG-O1 1.98(2) 1NOG-O2 2.07(2) 1(2) -3(1) 0.1121 2540.80 162.46 7

2N/O 1.95(4) 4(3) (1Im0) 1Im0 2.10(2) -2(1) 1NOG-O1 1.98(4) 1NOG-O2 2.07(4) 6(6) -3(2) 0.1304 2911.34 186.15 7

1N/O 1.89(3) 0(3) 1Im0 1.98(6) 3(9) 1Im0 2.10(2) -1(3) 1NOG-O1 1.98(2) 1NOG-O2 2.09(2) 2(3) -4(1) 0.1036 2343.33 171.80 9

2N/O 1.97(4) 4(4) (1Im0) -3(2). 0.1371 3020.27 193.12 7

109

1Im7 2.10(2) -3(1) 1NOG-O1 1.97(4) 1NOG-O2 2.06(4) 9(8)

1N/O 1.90(3) -1(2) 1Im0 2.09(4) 0(3) 1Im7 2.00(12) 6(19) 1NOG-O1 1.99(2) 1NOG-O2 2.08(2) 1(3) -4(2) 0.1112 2517.42 184.57 9

3N/O 2.11(2) 2(1) (2Im7) 1NOG-O1 1.91(2) 1NOG-O2 2.00(2) 2(3) 0(2) 0.1801 4060.93 230.21 5

1N/O 2.12(2) -2(1) 2Im7 2.05(6) 8(6) 1NOG-O1 1.91(3) 1NOG-O2 2.01(3) 5(4) -2(2) 0.1496 3339.41 213.52 7

4N/O 2.11(2) 5(2) (1Im0) 1Cl 2.79(6) 10(9) 5(2) 0.2150 4530.30 256.82 5

4N/O (2Im0) 1Cl

3N/O 2.01(2) 6(2) 1Cl 2.43(7) 18(10) 1Im0 2.11(1) -1(1) -2(1) 0.1073 2210.68 141.35 7

4N/O 2.05(3) 7(2) (1Im7) 1Cl 2.39(11) 18(16) 0(3) 0.2509 5149.33 291.92 5

2N/O 1.97(2) 3(2) 1Cl 2.38(6) 15(9) 2Im0 2.10(2) 1(1) -3(1) 0.1340 2828.19 180.83 7

3N/O 1.97(2) 4(2) (1Im0) 1Im0 2.10(1) -2(1) 1Cl 2.34(5) 11(6) -3(1) 0.1176 2508.19 160.37 7

2N/O 2.00(3) 6(5) 1Cl 2.36(5) 13(7) 1Im0 1.97(5) 3(5) -4(1) 0.0975 2056.04 150.74 9

110

1Im0 2.10(2) -2(1)

4N/O 2.03(3) 8(3) (2Im7) 1Cl 2.31(6) 12(8) -3(2) 0.2376 4828.91 273.75 5

2N/O 1.96(2) 2(2) 1Cl 2.38(5) 13(8) 2Im7 2.09(1) 0(1) -3(1) 0.1338 2836.94 181.39 7

3N/O 1.97(2) 4(2) (1Im0) 1Im7 2.10(1) -3(1) 1Cl 2.36(4) 11(6) -3(1) 0.1160 2452.51 156.81 7

2N/O 1.97(3) 4(3) 1Cl 2.38(5) 12(6) 1Im0 2.01(4) 2(5) 1Im7 2.11(2) -3(1) -3(1) 0.0962 2016.45 147.84 9

2N/O 1.89(6) 6(6) (1Im0) 1Cl 2.22(3) 4(3) 1NOG-O1 1.96(7) 1NOG-O2 2.05(7) 4(6) 10(4) 0.2276 4953.87 316.75 7

1N/O 1.91(2) -2(2) 1Cl 2.55(9) 19(16) 1Im0 2.08(2) 0(2) 1NOG-O1 1.98(2) 1NOG-O2 2.07(2) 0(2) -3(2) 0.1123 2493.00 182.78 9

2N/O 1.89(5) 4(5) (1Im7) 1Cl 2.22(3) 4(3) 1NOG-O1 1.97(5) 1NOG-O2 2.06(5) 4(7) -9(3) 0.2268 4958.22 317.03 7

1N/O 1.91(3) -2(2) 1Cl 2.54(10) 21(20) 1NOG-O1 1.98(3) 1NOG-O2 2.08(3) 1(3) 1Im7 2.08(3) -2(2) -2(2) 0.1186 2613.30 191.60 9

1Cl 2.23(3) 5(3) 2Im0 1.92(9) 12(9) 1NOG-O1 1.94(7) 1NOG-O2 2.03(7) 6(7) -9(4) 0.2246 4794.12 306.54 7

111

1Cl 2.22(3) 5(3) 1NOG-O1 1.96(7) 1NOG-O2 2.06(7) 7(8) 2Im7 1.93(8) 9(8) -8(4) 0.2256 4819.36 308.15 7

1Cl 2.47(6) 11(8) 1Im0 2.02(5) 1(5) 1NOG-O1 1.90(2) 1NOG-O2 2.00(2) 3(3) 1Im7 2.12(2) -3(2) -2(2) 0.1155 2553.12 187.18 9

3N/O 1.98(4) 8(4) (1Im0) 2Cl 2.28(4) 12(5) -7(3) 0.2497 5115.51 290.00 5

3N/O 2.00(4) 9(3) (2Im0) 2Cl 2.28(4) 13(5) -6(2) 0.2353 4778.36 270.89 5

2N/O 1.98(2) 392) 2Cl 2.38(5) 21(7) 1Im0 2.10(1) -2(1) -3(2) 0.1283 2669.92 170.71 7

3N/O 1.98(4) 7(4) (1Im7) 2Cl 2.28(4) 12(5) -7(3) 0.2559 5275.88 299.09 5

1N/O 1.92(2) -1(2) 2Cl 2.32(4) 17(6) 2Im0 2.06(2) 1(1) -5(1) 0.1530 3241.27 207.25 7

2N/O 1.95(2) 2(2) (1Im0) 1Im0 2.09(1) -2(1) 2Cl 2.31(3) 16(5) -5(2) 0.1234 2637.65 168.65 7

1N/O 1.96(5) 3(6) 2Cl 2.32(4) 17(5) 1Im0 1.95(4) 1(4) 1Im0 2.09(2) -3(1) -5(1) 0.1107 2351.07 172.37 9

3N/O 2.00(4) 8(3) (2Im7) 2Cl 2.29(4) 14(5) -6(4) 0.2376 4828.73 273.74 5

1N/O 1.91(20 -2(1) 2Cl 2.33(4) 17(5) -4(1) 0.1381 2954.40 188.90 7

112

2Im7 2.06(2) 0(1)

2N/O 1.95(2) 1(2) (1Im0) 1Im7 2.08(2) -3(1) 2Cl 2.32(4) 16(5) -5(2) 0.1261 2663.78 170.32 7

1N/O 1.94(3) 0(2) 2Cl 2.34(4) 17(5) 1Im0 1.99(3) 1(4) 1Im7 2.09(2) -4(1) -5(1) 0.1076 2268.29 166.30 9

1Im0 1.87(4) 2(3) 2Cl 2.21(4) 12(5) 1NOG-O1 1.96(5) 1NOG-O2 2.05(5) 4(6) -10(3) 0.2404 5187.57 331.69 7

1Im7 1.87(4) 1(3) 2Cl 2.21(4) 12(5) 1NOG-O1 1.96(5) 1NOG-O2 2.06(5) 4(6) -10(3) 0.2393 5164.55 330.22 7

Fe-Im(1Q5Y) - NOG(4V2W) - 6 Reduced 2 -3 2 2 2 N r (Å) σ (x10 Å ) ΔEo (eV) R factor χ χ #Var 6N/O 2.08(2) 11(2) (1Im0) 2(2) 0.2043 4119.31 209.74 3

5N/O 2.05(2) 15(3) 1Im0 2.11(1) 0(1) -2(1) 0.1139 2377.51 134.78 5

6N/O 2.07(2) 10(2) (2Im0) 0(2) 0.2087 4251.66 216.48 3

4N/O 2.02(2) 12(3) 2Im0 2.11(2) 3(2) -2(1) 0.1290 2703.10 153.24 5

5N/O 2.01(2) 10(2) (1Im0) 1Im0 2.12(1) -1(1) -1(1) 0.1126 2406.41 136.42 5

4N/O 2.04(3) 15(4) 1Im0 1.98(4) 4(6) 1Im0 2.11(4) -1(1) -3(1) 0.1006 2145.54 137.19 7

5N/O 2.01(2) 9(2) (1Im0) -1(1) 0.1209 2553.20 144.74 5

113

1Im7 2.12(1) -1(1)

4N/O 2.04(3) 10(3) 1Im0 2.11(2) -1(1) 1Im7 2.19(42) 32(94) -1(3) 0.1095 1708.36 109.23 7

4N/O 2.10(2) 6(2) 1NOG-O1 2.28(6) 1NOG-O2 2.38(6) 11(11) 3(2) 0.1941 3893.28 220.71 5

4N/O 2.12(2) 4(2) (1Im0) 1NOG-O1 1.90(3) 1NOG-O2 1.99(3) 3(3) 1(2) 0.1737 3819.84 216.55 5

3N/O 2.01(3) 6(3) 1Im0 2.11(1) -1(1) 1NOG-O1 2.10(7) 1NOG-O2 2.19(7) 17(14) -1(2) 0.1051 2170.31 138.77 7

4N/O 2.10(2) 4(2) (2Im0) 1NOG-O1 1.89(2) 1NOG-O2 1.99(2) 3(3) 0(2) 0.1675 3717.23 210.73 5

2N/O 1.93(3) 3(2) 2Im0 2.09(2) 2(1) 1NOG-O1 2.02(3) 1NOG-O2 2.12(3) 6(6) -3(1) 0.1171 2510.72 160.54 7

3N/O 1.98(4) 7(3) (1Im0) 1Im0 2.11(2) -2(1) 1NOG-O1 1.99(5) 1NOG-O2 2.09(5) 12(10) -2(2) 0.1173 2560.80 163.74 7

2N/O 1.96(6) 7(7) 1Im0 2.11(2) -2(2) 1Im0 1.99(6) 4(9) 1NOG-O1 2.03(5) 1NOG-O2 2.12(5) 10(12) -3(2) 0.1002 2148.20 157.50 9

3N/O 1.99(3) 6(3) (1Im0) 1Im7 2.11(2) -2(1) 1NOG-O1 2.00(6) 1NOG-O2 2.10(6) 15(12) -2(2) 0.1244 2681.42 171.45 7

114

2N/O 1.95(3) 4(3) 1Im0 2.03(4) 1(5) 1Im7 2.11(2) -3(2) 1NOG-O1 2.05(5) 1NOG-O2 2.14(5) 12(11) -3(1) 0.1001 2131.11 156.24 9

4N/O 2.11(2) 4(2) (2Im7) 1NOG-O1 1.90(2) 1NOG-O2 1.99(2) 4(2) 0(2) 0.1762 3854.24 218.50 5

2N/O 1.94(3) 2(2) 2Im7 2.09(2) 0(1) 1NOG-O1 2.04(4) 1NOG-O2 2.14(4) 10(9) -2(1) 0.1244 2656.91 169.88 7

5N/O 2.10(2) 8(2) (1Im0) 1Cl 2.79(5) 10(8) 4(2) 0.1903 3858.21 218.72 5

4N/O 2.02(4) 9(3) (1Im0) 2Cl 2.33(7) 20(9) -3(3) 0.2284 4603.20 260.96 5

4N/O 2.33(2) 10(3) 1Im0 2.11(2) -1(1) 1Cl 2.53(14) 28(22) -2(2) 0.1020 2098.91 134.20 7

3N/O 2.01(2) 7(2) 1Im0 2.11(1) -1(1) 2Cl 2.46(8) 32(13) -2(1) 0.1020 2081.35 133.08 7

5N/O 2.10(2) 8(2) (1Im7) 1Cl 2.78(5) 8(6) 4(2) 0.1950 3949.91 223.92 5

4N/O 2.02(3) 9(3) (1Im7) 2Cl 2.31(5) 19(8) -4(2) 0.2174 4362.74 247.33 5

4N/O 2.03(2) 10(3) 1Im7 2.11(2) -1(1) 1Cl 2.54(17) 30(26) -2(2) 0.1200 2468.65 157.85 7

5N/O 2.04(3) 10(2) (2Im0) 1Cl 2.33(7) 15(12) -3(2) 0.2118 4258.97 241.44 5

115

4N/O 2.02(3) 10(3) (2Im0) 2Cl 2.30(5) 17(7) -5(2) 0.2125 4250.20 240.95 5

3N/O 1.99(2) 7(3) 2Im0 2.11(2) 2(1) 1Cl 2.44(9) 21(15) -2(1) 0.1174 2468.46 157.83 7

2N/O 1.96(2) 3(2) 2Im0 2.09(1) 1(1) 2Cl 2.38(5) 24(9) -3(1) 0.1225 2562.01 163.81 7

4N/O 1.99(1) 7(2) (1Im0) 1Im0 2.11(1) -2(1) 1Cl 2.37(6) 14(8) -2(1) 0.1017 2150.53 137.50 7

3N/O 1.97(2) 5(2) (1Im0) 1Im0 2.10(2) -2(1) 2Cl 2.34(4) 21(6) -4(1) 0.1040 2193.33 140.24 7

3N/O 2.01(3) 11(5) 1Im0 1.98(5) 4(7) 1Im0 2.11(2) -2(1) 1Cl 2.39(9) 20(14) -3(1) 0.0920 1948.92 142.89 9

2N/O 1.99(4) 7(6) 1Im0 2.10(2) -2(1) 1Im0 1.97(5) 3(6) 2Cl 2.36(6) 24(9) -4(1) 0.0909 1910.02 140.03 9

5N/O 2.04(3) 10(2) (2Im7) 1Cl 2.33(8) 16(13) -2(2) 0.2152 4327.40 245.32 5

4N/O 2.02(30 9(3) (2Im7) 2Cl 2.31(5) 19(8) -4(2) 0.2174 4363.74 247.33 5

3N/O 1.99(2) 7(3) 2Im7 2.11(2) 1(1) 1Cl 2.44(10) 21(16) -2(1) 0.1328 2792.60 178.56 7

2N/O 1.95(2) 2(2) 2Im7 2.09(2) 0(1) 2Cl 2.39(5) 24(9) -3(1) 0.1269 2681.15 171.43 7

116

4N/O 1.99(2) 7(2) (1Im0) 1Im7 2.11(1) -2(1) 1Cl 2.38(6) 14(8) -3(1) 0.1068 2252.03 144.00 7

3N/O 1.97(2) 4(2) (1Im0) 1Im7 2.10(2) -3(1) 2Cl 2.35(5) 21(7) -4(1) 0.1084 2275.08 145.47 7

3N/O 2.03(4) 10(6) 1Im0 2.11(2) -1(1) 1Im7 1.96(5) 4(6) 1Cl 2.40(14) 26(22) -3(1) 0.0989 2104.56 154.30 9

2N/O 2.01(4) 7(6) 1Im0 2.10(2) -2(1) 1Im7 1.95(5) 3(4) 2Cl 2.36(7) 27(11) -4(1) 0.0989 2093.78 153.51 9

3N/O 2.11(1) 1(1) (1Im0) 1Cl 2.51(3) 7(3) 1NOG-O1 1.89(1) 1NOG-O2 1.99(1) 0(1) 0(1) 0.0900 1976.10 126.35 7

2N/O 1.90(6) 6(6) (1Im0) 2Cl 2.22(4) 11(5) 1NOG-O1 2.00(7) 1NOG-O2 2.09(7) 7(12) -9(3) 0.2359 4967.94 317.65 7

2N/O 2.00(14) 6(8) 1Cl 2.58(20) 19(22) 1Im0 2.11(2) -1(1) 1NOG-O1 1.99(13) 1NOG-O2 2.09(13) 9(26) -1(2) 0.1042 2169.84 159.08 9

1N/O 1.91(2) -2(2) 2Cl 2.55(6) 25(11) 1Im0 2.09(2) -1(1) 1NOG-O1 1.98(2) 1NOG-O2 2.07(2) 0(2) -2(1) 0.0945 2053.02 150.52 9

3N/O 1.93(5) 7(5) (1Im7) 1Cl 2.22(3) 3(3) 1NOG-O1 2.02(8) 8(13) -8(4) 0.2249 4848.85 310.04 7

117

1NOG-O2 2.12(8)

2N/O 1.90(5) 5(5) (1Im7) 2Cl 2.21(3) 10(5) 1NOG-O1 2.01(7) 1NOG-O2 2.10(7) 6(12) -9(3) 0.2344 4981.55 218.52 7

2N/O 1.98(7) 4(5) 1NOG-O1 2.02(2) 1NOG-O2 2.11(2) 8(20) 1Im7 2.11(2) -2(1) 1Cl 2.60(18) 21(26) -1(2) 0.1152 2400.39 175.99 9

1N/O 1.91(3) -1(2) 2Cl 2.55(7) 26(13) 1Im7 2.09(2) -2(2) 1NOG-O1 1.98(2) 1NOG-O2 2.07(2) 1(3) -2(1) 0.1017 2193.70 160.83 9

3N/O 2.11(1) 2(1) (2Im0) 1Cl 2.50(3) 7(4) 1NOG-O1 1.89(2) 1NOG-O2 1.98(2) 1(2) -1(1) 0.1046 2329.11 148.92 7

2Im0 2.10(2) 1(1) 2Cl 2.50(5) 19(7) 1NOG-O1 1.90(2) 1NOG-O2 2.00(2) 2(2) -1(1) 0.1288 2787.59 178.24 7

1N/O 1.89(1) -2(1) 1Cl 2.57(6) 15(11) 2Im0 2.09(2) 3(2) 1NOG-O1 1.97(2) 1NOG-O2 2.07(2) 0(2) -2(1) 0.0868 1952.27 143.13 9

2Cl 2.50(5) 19(7) 2Im0 2.10(2) 1(1) 1NOG-O1 1.90(2) 1NOG-O2 2.00(2) 2(2) -1(1) 0.1288 2787.59 178.24 7

3N/O 1.95(6) 9(6) (2Im7) 1Cl 2.23(3) 4(3) 1NOG-O1 2.02(8) 1NOG-O2 2.11(8) 11(13) -7(3) 0.2157 4554.70 291.23 7

118

2Cl 2.52(4) 17(6) 1NOG-O1 1.89(2) 1NOG-O2 1.98(2) 1(2) 2Im7 2.11(1) 2(1) 0(1) 0.1071 2300.57 147.10 7

2N/O 2.07(3) -1(3) (1Im0) 1Im7 2.16(4) -3(3) 1NOG-O1 1.88(2) 1NOG-O2 1.98(2) 1(2) 1Cl 2.49(4) 6(4) -1(1) 0.0925 2044.88 149.92 9

1N/O 2.12(3) -1(3) 1Cl 2.52(5) 9(6) 1Im0 2.10(4) 1(4) 1Im7 2.12(29) 17(32) 1NOG-O1 1.90(2) 1NOG-O2 1.99(2) 1(2) -1(3) 0.0898 1964.42 168.77 11

2Cl 2.48(5) 19(7) 1Im0 2.03(4) 1(5) 1Im7 2.12(2) -3(2) 1NOG-O1 1.90(2) 1NOG-O2 1.99(2) 2(2) -2(1) 0.0965 2081.55 152.61 9

Fe-Im(1Q5Y) - NOG(4V2W) - Best+Frac

Reduced 2 -3 2 2 2 N r (Å) σ (x10 Å ) ΔEo (eV) R factor χ χ #Var 3N/O 2.11(1) 1(1) (1Im0) 1Cl 2.51(3) 7(3) 1NOG-O1 1.89(1) 1NOG-O2 1.99(1) 0(1) 0(1) 0.0900 1976.10 126.35 7

3N/O 2.11(1) 1(1) (1Im0) 1.5Cl 2.52(3) 11(4) 1NOG-O1 1.89(1) 1NOG-O2 1.99(1) 0(1) 0(1) 0.0849 1820.56 116.41 7

3N/O 2.11(1) 1(1) (1Im0) 1.75Cl 2.53(3) 13(4) 1NOG-O1 1.89(1) 1NOG-O2 1.99(1) 0(1) 0(1) 0.0840 1786.23 114.21 7

119

3N/O 1.95(6) 10(7) (1Im0) 2Cl 2.23(5) 11(6) 1NOG-O1 2.05(12) 1NOG-O2 2.15(12) 14(26) -8(4) 0.2332 4832.71 309.00 7

2.5N/O 1.92(6) 8(6) (1Im0) 1.5Cl 2.22(3) 7(4) 1NOG-O1 2.00(9) 1NOG-O2 2.10(9) 8(13) -9(4) 0.2332 4935.01 315.54 7

Reduced 2 -3 2 2 2 N r (Å) σ (x10 Å ) ΔEo (eV) R factor χ χ #Var 3N/O 2.11(1) 2(1) (2Im0) 1Cl 2.50(3) 7(4) 1NOG-O1 1.89(2) 1NOG-O2 1.98(2) 1(2) -1(1) 0.1046 2329.11 148.92 7

3N/O 2.11(1) 2(1) (2Im0) 1.5Cl 2.51(4) 12(5) 1NOG-O1 1.89(2) 1NOG-O2 1.99(2) 1(2) -1(1) 0.1008 2205.54 141.02 7

3N/O 2.11(1) 2(1) (2Im0) 1.75Cl 2.51(4) 14(5) 1NOG-O1 1.89(2) 1NOG-O2 1.98(2) 1(2) -1(1) 0.1000 2176.84 139.19 7

3N/O 2.11(1) 2(1) (2Im0) 2Cl 2.51(4) 17(5) 1NOG-O1 1.89(2) 1NOG-O2 1.98(2) 1(2) -1(1) 0.0996 2160.29 138.13 7

3N/O 2.11(1) 2(1) (2Im0) 2.5Cl 2.52(4) 22(6) 1NOG-O1 1.89(2) 1NOG-O2 1.98(2) 1(2) -1(1) 0.0996 2150.98 137.53 7

2.5N/O 2.10(1) 2(1) (2Im0) -1(1) 0.1090 2362.14 151.04 7

120

2Cl 2.51(4) 18(6) 1NOG-O1 1.89(2) 1NOG-O2 1.99(2) 1(2)

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(59) Kulik, H. J.; Drennan, C. L. Substrate Placement Influences Reactivity in Non- Heme Fe(II) Halogenases and Hydroxylases. J. Biol. Chem. 2013, 288, 11233– 11241.

(60) Zhang, Z.; Ren, J. S.; Harlos, K.; McKinnon, C. H.; Clifton, I. J.; Schofield, C. J. Crystal Structure of a Clavaminate Synthase-Fe(II)-2-Oxoglutarate-Substrate-NO Complex: Evidence for Metal Centred Rearrangements. FEBS Lett. 2002, 517, 7– 12.

(61) Yang, M.; Chowdhury, R.; Ge, W.; Hamed, R. B.; McDonough, M. a.; Claridge, T. D. W.; Kessler, B. M.; Cockman, M. E.; Ratcliffe, P. J.; Schofield, C. J. Factor- Inhibiting Hypoxia-Inducible Factor (FIH) Catalyses the Post-Translational Hydroxylation of Histidinyl Residues within Ankyrin Repeat Domains. FEBS J. 2011, 278, 1086–1097.

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CHAPTER 4

HYDROGEN SULFIDE INHIBITION AND RESCUE OF THE HUMAN HYPOXIA SENSING ENZYME FIH

4.1 Introduction

Hydrogen sulfide (H2S) is emerging as an effector of numerous physiological

1–5 processes related to metabolism and the cardiovascular system. H2S has also displayed pro angiogenic/wound healing and anti-cancer effects, further adding to its potential as a

5–10 therapeutic agent for a wide array of human diseases. Currently, H2S releasing agents are being developed as therapeutics for protection against myocardial reperfusion injury,

4,5 lethal hypoxia, and inflammation. Many of the physiological responses to H2S treatment involve genes regulated by the transcription activator, Hypoxia Inducible

Factor (HIF), suggesting that some of the physiological effects of H2S may arise from effects on the HIF pathway. Despite the recent surge in reports of the therapeutic potential, the molecular mechanisms governing cellular responses to H2S remain poorly understood.6,7

HIF is a heterodimeric protein consisting of a nuclear localized β-subunit and a cytosolic α-subunit, which is regulated by the non-heme Fe2+/αKG dependent oxygenase,

Factor Inhibiting HIF (FIH).11,12 Under hypoxic conditions, HIF-1α translocates to the nucleus to dimerize with HIFβ, then recruits the transcriptional co-activator p300 to initiate gene expression.13 FIH hydroxylates HIF-1α-Asn803 in the c-terminal transactivation domain (CTAD), preventing HIF mediated gene expression.13 HIF regulated genes are associated with angiogenesis and vascularization, maintaining basal metabolism, erythropoiesis, and lung development.14,15 It’s been recently demonstrated that H2S treatment leads to an increase in HIF-1α mRNA and upregulation of HIF

128 dependent target genes.16,17 As the HIF hydroxylases Prolyl Hydroxylase Domain 2

(PHD2) and FIH control HIF-1α abundance and transcriptional ability respectively, the activity of the HIF hydroxylases and physiological responses to H2S treatment may be mechanistically linked.

Scheme 4.1: Proposed consensus mechanism of FIH and αKG-dependent

oxygenases

The active site of FIH contains a 2His 1Asp facial triad coordinated to the Fe2+ center, a hallmark of the Fe2+/αKG dependent enzyme superfamily.18–20 The chemistry of

FIH proceeds through a sequential mechanism in which binding of HIF-CTAD stimulates aquo release from the diffusible ligand site, followed by O2 binding and the subsequent oxidative decarboxylation of αKG to form a reactivity ferryl intermediate (Scheme 4.1)21–

23 which has been trapped and identified as the hydroxylating species in related enzymes

24,25 26 taurine dioxygenase (TauD) and prolyl 4 hydroxlase (PH4) . The ferryl hydroxylates the β carbon of CTAD-Asn803. CTAD binding appears to alter second coordination

129 sphere contacts that lead to aquo release. Asp201 H-bonds to the axial aquo ligand but rearranges upon CTAD binding to form new H-bond contacts with the backbone amide of CTAD and other second sphere residues.21,23 The subsequent rearrangement of the active site may serve to modulate the access of O2 and related molecules to this diffusible ligand site.

Given the correlation between H2S and HIF regulated pathways, a potential mechanism of action for H2S would be to directly bind to the open coordination site of

FIH. Under conditions of low CTAD, FIH can also autohydroxylate to form an inactive

3+ Fe complex. As H2S is a known reducing agent, another possible mode of action would be to quench the ferryl or reduce Fe3+ in autohydroxylated FIH to Fe2+. Interestingly, the

− active site appears to be well suited for polar species such as H2S/HS as there would be a stabilizing effect from hydrogen bonding to the Asp201 sidechain.

There have not been any in vivo demonstrations of H2S directly inhibiting the HIF hydroxylases that we are aware of, but studies have shown that exposure of C. elegans to

17 H2S leads to HIF-1α nuclear translocation and expression of HIF-1α target genes, suggesting inhibition of the HIF hydroxylases. Here, we provide the first in vitro evidence that H2S directly interacts with one of the HIF hydroxylases, FIH. This provides a rationale for many of the physiological effects of H2S, and may provide intriguing directions for therapeutics related to anemia, cancer, and ischemic diseases. In the present work, we demonstrate through kinetics and spectroscopy that the anion, HS− inhibits FIH activity by ligating to the metal cofactor. We also show that HS− can reduce the ferric center of inactive FIH to the active ferrous state, restoring activity. This suggests a dual role for H2S in modulating FIH activity in cells.

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4.2 Methods

4.2.1 Protein Purification

All reagents were purchased and used as received from commercial vendors unless stated otherwise. Expression and purification of FIH was carried out as previously

27 described in the literature. The N-terminal His6 tagged protein was purified using a Ni-

NTA column with subsequent removal of the tag via thrombin digestion. EDTA treatment was used to remove exogenous metals followed by further purified using size- exclusion chromatography to afford the active dimer. Protein concentrations were

-1 -1 27 determined by absorption spectroscopy measurements (ɛ280 = 48800 M cm ).

4.2.2 CTAD Purification

A 39 amino acid peptide substrate corresponding to the c- terminal transactivation domain (CTAD) of HIF-1α788-826 containing a Cys800 → Ala substitution with sequence

DESGLPQLTSYDAEVNAPIQGSRNLLQGEELLRALDQVN was ordered from

EZBiolabs (Asn803 bolded) as a desalted synthetic peptide with free N and C termini. The peptide was dissolved in 25% acetonitrile and purified by reverse-phase HPLC as previously reported.27,28 Two rounds of purification yielded CTAD >95% pure as determined using matrix assisted laser desorption ionization mass spectrometry (MALDI-

MS). The concentration was determined by measuring the UV-absorption at 293 nm in

-1 -1 28 0.1 M NaOH (ε293 = 2400 M cm ).

4.2.3 Dose Response Assays

Dose response assays were carried out according to previous reports from the

29 literature. Reaction components, Asc (2 mM), FeSO4 (25 µM), αKG (25 µM), CTAD

131

(100 µM), and sulfide (0 – 0.52 mM) were incubated in 50 mM Hepes pH 6.00 – 8.00 at

37°C for 1 minute and the reaction initiated by adding 5 μL of apo FIH to bring the final volume to 50 μL and a final concentration of FIH = 0.5 µM. Reactions were carried out

30 under ambient O2 conditions (~ 217 µM]). Aliquots of the reaction (5 μL) were quenched in 20μL of 75% acetonitrile with 0.2% trifluoroacetic acid saturated with 3,5

Dimethoxy-4-hydroxycinnamic acid at designated time points. Initial rates were determined by calculating the mole fraction of peptide converted to product based on the relative intensities of the un-hydroxylated peptide (m/z = 4255) and hydroxylated peptide

(m/z = 4271) using MALDI-MS.27,29

4.2.4 EPR Spectroscopy A 500 µL sample containing FIH (100 µM), Fe2+ (100 µM), and αKG (500 µM) in air saturated Hepes pH 7.40 was prepared and incubated at 23°C aerobically for 45 minutes to allow auto-oxidation of FIH to occur, forming Fe3+ at the active site.31

Aliquots of freshly prepared Na2S (50 mM) in 50 mM air saturated Hepes pH 7.40 were added to aliquots of the samples such that the final volumes were 100 µL and the final concentrations of sulfide were 0.5 mM and 1 mM. The samples were allowed to incubate for an additional 5 minutes at 23°C, at which point they were transferred to EPR tubes and flash frozen in liquid nitrogen. EPR samples were run using a finger dewar on a

Bruker Elexsys E-500 EPR equipped with a DM4116 cavity at 9.609 GHz frequency, 6.0 mW power, 10G modulation amplitude, 100 GHz modulation frequency, 327 ms time constant, 77K.

4.2.5 Electronic Absorption Spectroscopy

An aerobic sample containing FIH (1 mM), FeSO4 (1 mM), αKG (2 mM), and sulfide (2 mM) was prepared and the spectrum collected from 300 – 1000 nm on an HP

132

845x UV-visible spectrophotometer. We noticed that the auto-hydroxylated ferric chromophore28 dominated the electronic transitions, so additional aerobic UV-Vis experiments were conducted in the presence of N-oxalylglycine (NOG), an αKG

21 analogue that inhibits chemistry. A sample containing FIH (0.467 mM), FeSO4 (0.467 mM), NOG (2 mM), and H2S (0.467 mM) in 50 mM Hepes pH 7.40 was prepared and purified by size exclusion chromatography using buffer containing 50 mM Tris 50 mM

NaCl pH 8.00. The FIH containing fractions were concentrated and buffer exchanged into

50mM Hepes pH 7.40. The protein concentration was verified (0.424 mM) and the electronic absorption spectrum was measured from 300 – 1000 nm.

Anaerobic UV-visible measurements were recorded using an Avantes AvaSpec-

128 Ultrafast Fiber Optic Spectrometer set up in a glovebox. All reagents were degassed using argon and brought into the glovebox with the exception of FeSO4 and Na2S, which were brought in as solids and prepared using degassed water and Hepes pH 7.40 respectively. The sample contained FIH (1 mM), FeSO4 (1 mM), αKG (2 mM), and sulfide (2 mM) in a total volume of 100 µL.

4.2.6 Rescue Assays

A 1 mL sample containing FIH (50 µM), FeSO4 (50 µM), and αKG (1 mM) in

Hepes buffer pH 7.40 was prepared and incubated at 23°C overnight (~16 hours) to allow auto-hydroxylation and inactivation of FIH to occur. Auto-hydroxylated FIH was treated with a 10 mM stock of varied reducing agents, such that the working concentrations were

49 µM FIH and 200 µM reductant, then incubated at 37°C for 15 minutes. Reactions were initiated by adding 5 µL of the treated samples to fresh tubes containing 45 µL of

100 µM CTAD pre-incubated at 37°C in 50 mM Hepes pH 7.40 for 2 minutes, bringing

133 the total volume to 50 µL and the final concentration of FIH = 4.9 µM. Aliquots (5 µL) were quenched after 5 minutes as for the dose response assays, then analyzed using

MALDI-MS.

For dose response measurements, aliquots of the auto-hydroxylated FIH samples were incubated with varying concentrations of sulfide at 37°C for 2 minutes and the reaction initiated by adding CTAD. Reactions were quenched as described above after 5 minutes and the concentration of hydroxylated CTAD determined. The final concentrations were FIH (4.9 µM), FeSO4 (4.9 µM), αKG (185 µM), sulfide (0 – 1.5 mM), and CTAD (100 µM). The data was fit to a simple dose response model (equation

1) in which A1 is the top asymptote, A2 is the bottom asymptote, and p is the hill coefficient.

퐴2−퐴1 푦 = 퐴1 + (1) 1+10(퐿푂퐺푥−푥)푝

4.2.7 Dynamic Light Scattering (DLS)

DLS samples were prepared by mixing FIH (50 µM), FeSO4 (49 µM), αKG (250

µM), and H2S (49 µM) in 50 mM Hepes pH 7.40. Measurements were taken with a

MALVERN Zetasizer Nano ZS instrument and analyzed as previously described.32

Briefly, samples were incubated for 2 minutes at 25°C and 4 scans (12 measurements per scan) were collected using incident excitation laser of 633 nm. The data from the 4 scans were averaged and analyzed using the Stokes-Einstein equation as previously reported.32

4.3 Results

As H2S is garnering increased attention as a potential therapeutic agent for various diseases, identifying molecular targets of H2S could aid in therapeutic development.

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Recent studies have implicated the HIF hydroxylases as potential targets of H2S in

16,17 cells. To test the hypothesis that FIH is a target of H2S, we performed inhibition assays under ambient [O2]. We measured a dose response curve at pH 7.00 and observed

3 moderate inhibition of FIH (IC50 = 64 μM). As the pKa of H2S is 6.9, we measured the

− inhibition assays over the pH range 6.00 – 8.00 in order to test for the H2S/HS speciation. We observed that the IC50 values were pH dependent and decreased as pH was increased from 6.00 to 8.00 (Figure 4.1), pointing to HS− as the relevant inhibitor, since FIH activity is pH-independent in this range.33 As sulfide is ~ 80% HS− and ~ 20%

34 − H2S at physiological pH (pH 7.40), our use of the term H2S or HS refers to total

− + sulfide, which is distributed in a pH dependent equilibrium H2S + H2O ⇋ HS + H3O .

Figure 4.1: pH dependent inhibition of FIH (0.5 µM) in 50 mM Hepes pH 6.00 – 8.00 at 37°C; αKG (25 μM), FeSO4 (25 μM), CTAD (100 μM), sulfide (0 – 520 μM), ascorbate (2 mM). The inset is an example of a dose response curve at pH 7.40. To determine a Ki(app) for sulfide inhibition, we performed steady state kinetic assays with varied concentrations of CTAD in the presence of constant sulfide concentrations (Figure 2). To obtain Ki(app), the data was fit to an uncompetitive inhibition

135

-1 27 27 binding model (equation 2) in which Vmax (fixed at 31 min ) and KM (fixed at 77 µM) have their traditional meanings, [S] denotes the substrate concentration and [I] denotes the inhibitor ([sulfide]tot) concentration. A global fit performed in OriginPro version 9.1 yielded Ki(app) = 60 ± 9 µM.

v 푘cat ×[CTAD] = [Sulfide] (2) E 퐾 +[S](1+ ) M(CTAD) Ki(푎푝푝)

In enzymatic reactions, kcat/KM for a given substrate reports on all steps involving diffusional encounter with the substrate up through and including the first irreversible step, which is oxidative decarboxylation in the case of FIH (Scheme 4.2). According to the consensus mechanism, substrate binding precedes O2 binding, which leads to the first

30 35 irreversible step. Under conditions of ambient [O2] = 217 µM (KM(O2) = 90 µM), both kcat and kcat/KM (CTAD) report on O2 reaction as this is the first irreversible step. A regression analysis plot (Figure 4.2) generated from the kinetic data revealed a decreasing slope, suggesting that sulfide was an uncompetitive inhibitor for CTAD, and a step after

CTAD binding is perturbed, likely O2 binding or O2 reaction.

Scheme 4.2: Kinetic mechanism of FIH

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Figure 4.2: (Left) Steady state kinetics of FIH in the presence of sulfide (Right) Regression analysis of steady state kinetics data. Assays were performed at 37°C with FIH (0.5 µM), ascorbate (2 mM), αKG (500 µM), FeSO4 (25 µM), and CTAD (24 – 288 µM) in 50 mM Hepes pH 7.40.

As HS− inhibition could arise from many potential mechanisms, we measured the electronic environment of FIH in the presence of sulfide to test for binding. The use of electronic absorption spectroscopy for monitoring ligand access to non heme Fe centers has been demonstrated to be an effective method.36,37 Binding of αKG in Fe2+/αKG- dependent enzymes leads to electronic transitions at ~ 500 nm attributed to metal to ligand charge transfer (MLCT) transitions from Fe2+ to the bidentate coordinated αKG.38–

40 Anaerobic addition of αKG to (Fe)FIH resulted in a broad absorption centered at 500 nm (Figure 4.3), indicating that αKG bound to Fe2+ in the FIH active site. Adding sulfide to the sample of (Fe+αKG)FIH caused the solution to turn a light yellow color with a new electronic transitions centered near 400 nm, and two weak transitions near 500 and 600 nm (Figure 3),suggesting binding of sulfide to the Fe cofactor in FIH. Binding of HS− to

Fe2+ in the αKG-dependent oxygenase SyrB2 resulted in a change of the MLCT centered at 500 nm to two transitions centered at 540 nm and 600 nm.36

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Figure 4.3: Electronic absorption spectra of aerobic and anaerobic (Fe+αKG)FIH treated with sulfide. (black) Anaerobic (red) Aerobic. FIH (1 mM), FeSO4 (1 mM), αKG (2 mM), and sulfide (2 mM).

To ensure that the electronic transitions observed were due to binding of sulfide at the active site, we decided to purify sulfide treated FIH by size exclusion chromatography

(SEC). An aerobic sample was prepared and upon addition of αKG we noticed that the

αKG dependent MLCT was shifted to ~583 nm, consistent with the formation of Fe3+ as a result of auto-hydroxylation of FIH (Figure 4).28,31 When sulfide was added, the transition at 583 nm disappeared and new transitions centered at 440 nm, 515 nm, and 605 nm

(Figure 4.3) appeared. This suggested that sulfide reduced the ferric iron to the ferrous

− 2+ 3+ form. Studies have shown that H2S, mainly the anion (HS ), binds to Fe or Fe at heme or binuclear enzyme active sites. 37,39–41 Binding of HS− to Fe3+ in superoxide dismutase and cytochrome c oxidase (CcO) reduced Fe3+ → Fe2+, concomitant with a red shift in the soret band in CcO and a disappearance of the broad absorption near 350 nm in SOD,37,41 suggesting reduction of Fe3+FIH → Fe2+FIH

138

To prevent auto-hydroxylation of FIH, we substituted αKG with N-oxalylglycine, an αKG mimic that binds in a similar manner but inhibits chemistry.21,28,42 Binding of sulfide to (Fe+NOG)FIH produced broad transitions at λmax ~ 440 nm, and 600 nm

(Figure 4.4). The sample was purified by SEC and protein containing fractions were pooled and then concentrated with 50 mM Hepes pH 7.4 using a 30 kDa molecular weight cutoff filter. The resultant electronic absorption spectra displayed transitions with

λmax = 440 nm, 515 nm and 605 nm, similar to the transitions observed in the presence of

αKG after sulfide treatment. This suggested that sulfide was bound relatively tightly to

Fe2+ in FIH (Figure 4). Inductively coupled plasma atomic emission spectroscopy measurements post purification by SEC demonstrated that the FIH to Fe ratio was 1:0.6, consistent with an intact cofactor site in FIH. Our data suggests that sulfide binds to the active site metal in FIH as the electronic transitions and extinction coefficients observed are compatible with those observed for model complexes (ɛ ~ 700 – 6100 M-1 cm-1) of thiolate ligated non-heme Fe centers.43 Anaerobic and aerobic addition of HS- to Fe and

αKG or NOG resulted in blackish solutions that failed to exhibit the well defined peaks observed in the presence of FIH (Figure 4.10).

Figure 4.4: Electronic absorption spectra of (Fe+NOG+Sulfide)FIH in 50 mM Hepes pH 7.40. (left) Spectra in the absence and presence of sulfide before and after purification with SEC; FIH (467 µM), Fe2+ (467 µM), NOG (2 mM), sulfide (0, 467 µM). After SEC, [FIH] = 424 µM. (Right) Difference spectra calculated by subtracting the spectrum of (Fe+NOG)FIH.

139

Dynamic light scattering (DLS) experiments were conducted to test for metal sequestration by sulfide as this technique is a sensitive reporter of trace amounts of particulate formation in solutions. The hydrodynamic diameter (DH) of (Fe+αKG)FIH was 8.2 ± 2.3 nm (Figure 4.5), consistent with the reported crystallographic dimensions.21

Adding sulfide to the preformed (Fe+αKG)FIH sample produced a DH of 7.8 ± 2.4 nm

(Figure 5), within uncertainty to the hydrodynamic diameter in the absence of sulfide, indicating that Fe-S colloids were not formed. Formation of Fe-S colloids yields a hydrodynamic diameter of 168 ± 42 nm (Figure 4.5).

Figure 4.5: DLS of (Fe+αKG)FIH and (Fe+αKG+Sulfide)FIH. FIH (50 μM), FeSO4 (49 μM), αKG (250 μM), Sulfide (50 µM) in 50 mM Hepes pH 7.40.

FIH and other αKG dependent oxygenases catalyze auto-hydroxylation of nearby active site residues in the absence of prime substrate, rendering the enzymes inactive.31,44

Auto-hydroxylated FIH (FIH-OH) was demonstrated to be inactive due to formation of an Fe3+-O-Trp296 adduct, resulting in an indigo colored protein solution with an electronic

28 absorption transition centered at 583 nm. H2S is known to reduce metalloprotein active sites,37,41,45 raising the possibility that sulfide could potentially restore inactive FIH

140

3+ 3+ activity by reducing Fe . H2S rapidly converts ferric (Fe ) heme centers to the ferrous

(Fe2+) states.46–48

A sample of FIH was incubated with FeSO4 and αKG at 23°C overnight for auto- hydroxylation. Following incubation, the sample turned a light indigo color and displayed a transition centered at 583 nm. Aliquots of this sample were treated with 200 µM of a variety of reducing agents: ascorbate (Asc), dithiotrietol (DTT), glutathione (GSH), N- acetylcysteine (NAC), and sulfide (HS−) for 15 minutes at 37°C to test for the reduction of auto-hydroxylated FIH. Post treatment, the samples were assessed for activity by mixing with fresh buffer containing CTAD, then analyzed for hydroxylated peptide using

MALDI MS. Auto-hydroxylated FIH was inactive as previously reported,28 and rescue of activity was observed for the HS− treated sample (Figure 6). The HS− treated sample had significantly hydroxylated CTAD (~ 38 µM CTAD-OH) while other reductants yielded < 5 µM CTAD hydroxylated (Figure 4.6). This suggests that sulfide was able to access the active site of FIH and reduce the inactive Fe3+ cofactor to the active Fe2+ form.

Figure 4.6: Rescue of auto-hydroxylated FIH activity by reductants. FIH (4.9 μM), Fe2+ (4.9 μM), αKG (185 μM), CTAD (100 µM), reductant (20 µM) in 50 mM Hepes pH 7.40.

141

Fe2+ in non-heme enzymes are generally EPR silent due to the Fe having a low spin S = 2 spin configuration. Oxidation of Fe2+ yields a high spin S = 5/2 Fe3+ center that can be observed using X-band EPR spectroscopy. Samples of auto-hydroxylated FIH were treated with sulfide then monitored by EPR to test for reduction of Fe3+. The EPR spectrum of the auto-hydroxylated FIH exhibited geff = 4.30, 4.25 (Figure 7), corresponding to high spin Fe3+ (S = 5/2). This oxidation was FIH dependent as the sample of Fe and αKG in buffer failed to produce an EPR signal. Addition of sulfide resulted in loss of the high spin Fe3+ signal (Figure 7), indicating that Fe3+ was reduced to

Fe2+ in the HS− treated sample.

Figure 4.7: X-Band EPR of auto hydroxylated FIH samples. FIH (100 μM), FeSO4 (100 μM), αKG (500 μM), Sulfide (0 – 1 mM) in aerobic 50 mM Hepes pH 7.40. 9.609 GHz, 6.0 mW power, 10 G modulation amplitude, 100 GHz, 77 K.

The concentration dependence of sulfide rescue of auto-hydroxylated FIH was tested by incubating auto-hydroxylated FIH with sulfide for 2 minutes and then initiating turnover by adding CTAD. Reactions were quenched after five minutes and assessed for

142 hydroxylation via MALDI-MS. We observed that sulfide rescue of FIH activity was dose dependent, exhibiting an EC50 = 333 ± 32 µM (Figure 4.8). Interestingly, at doses > 600

µM, there was inhibition that increased with increasing [sulfide] (Figure 4.10), consistent

− with higher concentrations of HS inhibiting activity. H2S is known to exhibit different effects based on its concentration. At high concentrations, H2S inhibits CcO activity but

41 acts as a reductant at low concentrations. Thus, low concentrations of H2S can preserve mitochondrial function by decreasing the production of reactive oxygen species.49 As a nueroprotectant, low concentrations of H2S mediate oxidative stress whereas high concentrations have been shown to enhance cerebral damage.49

Figure 4.8: Concentration dependence for sulfide rescue of auto-hydroxylated FIH activity. FIH (5 μM), FeSO4 (5 μM), αKG (185 μM), sulfide (0 - 600 µM) in 50 mM Hepes pH 7.40. EC50 = 333 ± 32 µM

4.4 Discussion

There is an overwhelming amount of evidence that links the effects of H2S treatment to the HIF pathway,16,17,50 suggesting that sulfide may be a physiological regulator of HIF activity. Exposure of C. elegans to 50 ppm H2S ( ~ 2 µM at STP.) led to

143

HIF-1α nuclear localization and increased HIF target gene expression, which was

17 positively correlated to survival in otherwise lethal concentrations of H2S. In fact, HIF-

17 1α was required for survival to lethal doses of H2S, suggesting that HIF may aid in relieving the stress caused by H2S inhibition of CcO. Treatment of vascular smooth muscle cells with 300 µM H2S resulted in stabilization of HIF-1α mRNA and HIF controlled gene mRNA such as VEGF, and stimulated binding to the HIF response element (HRE),51 an outcome that could be explained by the inhibition of the HIF hydroxylases in cells. Additionally, H2S has also exhibited pro-angiogenic activity in cultured endothelial cells, an effect that is enhanced under hypoxic conditions and

52,53 thought to proceed through a pathway involving upregulation of HIF-1α. Na2S at a dose of 25.6 µmol/kg per day significantly augmented ischemic tissue VEGF levels and

53 restored ischemic hind-limb blood flow in mice. If H2S bound to the iron in the HIF hydroxylases, the effects of sulfide treatment would be more potent under hypoxic conditions, suggesting a mechanistic linkage between H2S treatment and the HIF hydroxylases.

Our data demonstrated that FIH was inhibited in a dose and pH dependent manner by sulfide with maximum inhibition near physiological pH (IC50 ≈ 30 µM at pH 7.40), pinpointing the anion HS− as the inhibiting species as ~ 80% of the total available sulfide exists as HS− at pH 7.40.34,54 We also showed that sulfide rescued FIH activity in a dose dependent manner with EC50 = 333 µM. In studies where H2S was reported to be an inhibitor or reducing agent, it’s been suggested that the anionic form (HS−) was the relevant species causing the observed effects.37,45,55

144

To date, there have been limited accounts of H2S directly inhibiting enzymes,

− 55 including carbonic anyhdrase and its various isozymes (HS , Ki = 0.0006 - 70 mM), and

56 CcO, (H2S, Ki = 0.2 μM). Due to CcO inhibition, H2S is a toxic substance to aerobic organisms. However, H2S treatment at a modest concentration (80 ppm or 3.4 µM in air at 15°C) induced a reversible suspended-animation state in mice, marked by low demand

4 for O2 and reduced metabolic rate. This state resembled hibernation, suggesting a global metabolic response rather than simple CcO inhibition. The recent discovery that HIF plays a role in up-regulating the transcription of certain homeostasis networks and

16 − proteins in response to H2S treatment, coupled with our observation that HS inhibits

FIH, implies that direct action on FIH and other hydroxylases that regulate HIF, such as the prolyl hydroxylase, may account for some of the broader metabolic effects of H2S.

In humans, the concentration of H2S that leads to acute poisoning is reported to be

57,58 1000 – 2000 ppm (~ 41 – 82 µM at STP.) However, the toxicity of H2S is mitigated by its rapid oxidation in the mitochondria, and the presence of > 30 µM HS− was shown to not disturb oxidative phosphorylation.59 Intoxicating mice with NaSH elevated endogenous H2S levels by 57%, 18% and 64% in brain, liver, and kidney respectively, suggesting that the concentration range must be steep for the switch over from beneficial to toxic effects.59 As such, no cytotoxic effects were observed in HCT116 human colon

60 cancer cells upon treatment with 125 µM – 1 mM H2S or SH-SY5Y human

61 neuroblastoma cells in the presence of 25 – 250 µM H2S.

Current estimates of endogenous H2S levels in living tissue are still unclear as different observations have been reported based on the analytical method with estimates

62–65 ranging from virtually undetectable to low micromolar. Recently, the use of H2S

145 specific electrodes have led to more reliable detection of sulfide levels. Reports of H2S levels in serum using H2S electrodes in healthy individuals and patients with chronic obstructive pulmonary disease are estimated to range from 30 – 100 µM.66,67

Concentrations of H2S in mammalian brain tissue have been reported to be as high as 50

– 160 µM,59 suggesting that the concentrations at which FIH is affected in our assays

(IC50 = 34 µM, EC50 = 333 µM) are physiologically relevant in certain tissues, and are within range of the concentrations reported to elicit physiological changes. For example,

59 H2S was reported to act as a vasorelaxant (IC50 = 125 µM) of rat aorta and portal veins

68 and demonstrated to inhibit NO synthase isoforms (IC50 = 130 – 210 µM). H2S levels in plasma have been demonstrated to rise to as high as 41 µM in various disease models such as pancreatitis, haemorrhagic, endotoxic, and septic shock, along with upregulation

6 of the mRNA of CBS and CSE, the H2S generating enzymes.

As the therapeutic potential for H2S in ischemic and hypoxia related disorders is now being realized, there is an urgent interest in elucidating molecular mechanisms of

− − action for H2S. Our data implicates FIH as a potential target of H2S/HS in cells (HS

− being more relevant). As HS is likely competing with O2 for binding, the concentration needed to attenuate FIH activity in vivo may be less due to the fact that [O2] in cells is

14,69 typically about 5 % of ambient O2.

We established that HS− accessed the (Fe+αKG)FIH cofactor to inhibit and rescue activity by three methods. First, HS− reacted with Fe resulting in electronic absorption bands with λmax = 440 nm, 515 nm and 605 nm. These electronic transitions persisted following purification of HS− treated FIH by SEC, suggesting that the active site was intact and contained HS− bound to Fe. As the transitions arise from an EPR silent Fe

146 center and the DLS data suggested no Fe-S colloid formation, we interpret the transition to be S → Fe2+ ligand to metal charge transfers transition. The anaerobic (Fe+αKG)FIH sample exhibited the characteristic MLCT at 500 nm indicative of αKG binding to Fe2+ in

FIH. Upon addition of H2S, the transition at 440 nm was prevalent but the peaks at 515 nm and 605 nm were weak, likely due to slight protein precipitation. HS− binding to Fe2+ in related enzyme SyrB2 resulted in transitions centered at 540 nm and 600 nm.36 The lack of a carboxylate ligand in SyrB2 likely results in different orbital overlap with bound

HS−, resulting in the observed difference in the transitions of SyrB2 compared to FIH.

A comparison of binding affinities shows that Fe2+ binds more tightly to the FIH active site than in solution with HS−. HS− binds Fe2+ with moderate affinity (log β for

HS−/Fe2+ is 5.2).70 However, (Fe+αKG)FIH is estimated to bind Fe2+ with high affinity

+ + 27 2+ based on the binding of CO2 (log β for (CO2 +αKG)FIH is 6.9); Fe binding to the related enzyme taurine dioxygenase (apo-TauD) was similarly strong (log β = 7.0).71

Based on thermodynamic considerations, Fe2+ is bound more tightly by FIH than by HS−.

This is consistent with the electronic absorption spectra being unchanged after purification of (Fe+NOG+HS−)FIH by SEC and the hydrodynamic diameter being consistent with the presence of soluble Fe bound FIH following the addition of HS− to

(Fe+αKG)FIH .

Second, HS− reduced air-oxidized (Fe3++αKG)FIH back to the Fe2+ state. Air

3+ oxidized (Fe +αKG)FIH was light indigo, and displayed an EPR signal (geff = 4.30,

4.25) indicative of Fe3+. As the g-values were not widely dispersed from g near 4.3, this suggested a nearly pure rhombic Fe3+ center that possessed close to ideal octahedral geometry.37 Upon HS− addition, the light indigo color and EPR signal disappeared,

147 indicating reduction to (Fe2++αKG)FIH. The rescue of FIH activity upon treatment with

HS− is consistent with reduction of Fe3+ back to Fe2+, which was also observed by the change in the characteristic (Fe3++αKG)FIH UV-Vis transition centered at 583 nm. We interpret this to indicate that HS− accessed the active site in order to reduce the Fe cofactor.

− 3+ The antioxidant properties of H2S are well documented. Addition of HS to Fe superoxide dismutase (Fe3+SOD) led to a colorless sample coincident with the disappearance of the absorption transition at 350 nm, suggesting reduction of Fe3+ to

2+ 37 Fe . EPR experiments demonstrated that low concentrations of H2S fully reduced the

3+ 41 Fe form of oxidized CcO in quantitative yield. H2S has also been demonstrated to act as an antioxidant that provides protection against oxidative stress in numerous cell lines.72–74 The conversion of HS− → S0 + H+ + 2e- has a redox potential of +0.17 V, making it a weaker reductant than other well established reducing agents such as glutathione (-0.25 V), ascorbate (-0.066 V), and dithiotrietol (-0.33 V), which are much more abundant in cells.34,75 Despite being a weaker reductant, reduction of Fe3+ in FIH seemed to be HS− specific under our conditions, perhaps speaking to the fact that FIH may be a physiological target for HS−. The smaller size of HS− may afford it better access to metalloprotein active sites compared to the more abundant intracellular thiols such as

GSH and cysteine.

Third, initial rates of turnover were unaffected by the order of reagent addition.

When HS− was added to pre-formed (Fe+αKG)FIH and the reaction initiated with CTAD, the rate was the same as when HS− was pre-incubated with Fe2+ and αKG and turnover initiated with FIH. As rates are unchanged when Fe is pre-loaded in FIH or allowed to

148 react with HS− in solution, this excludes kinetic formation of a solution-based HS−/Fe2+ adduct as the mechanism of inhibition. Importantly, the addition of excess Fe (6 equivalent with respect to HS−) failed to increase FIH activity, suggesting that inhibition is due to HS− binding to Fe2+ as opposed to Fe sequestration or ferryl quenching.

− Our kinetic data suggests that HS likely competes with O2 for binding as it is an uncompetitive inhibitor of CTAD and the IC50 is unchanged in the presence of high

[αKG] (Supplemental Figure 1). As the regression analysis for kcat/KM(CTAD), which reports on O2 binding, showed a decreasing slope, it suggested that diffusional encounter

2+ with O2 was perturbed. The relatively tight binding of sulfide to Fe in FIH supports this model. UV-Vis and electrocatalytic data indicated that H2S bound to the reduced 5-

2+ 41 coordinate Fe site of CcO and served as a competitive inhibitor of O2.

Scheme 4.3: Proposed model for sulfide interaction with FIH.

In sum, our data indicates that HS− directly interacts with the Fe2+ cofactor in

(Fe+αKG)FIH. As FIH is part of the control mechanism to regulate HIF-controlled gene

149 expression in response to hypoxia, this suggests that FIH may serve as a pathway for regulating HIF activity in response to H2S in vivo. The IC50 (~30 µM) of inhibition and

EC50 (~330 µM) of rescue are reasonable as these are within the range of some estimates

58,66 of H2S levels in humans cells and serum. Based on the evidence presented here and in literature,13-15 we propose a model (Scheme 4.3) in which HS− directly binds to the exposed coordination site of (Fe+αKG)FIH, inhibiting its activity and thereby promoting

HIF transcriptional activity. However, in the absence of prime substrate, auto- hydroxylated (inactive) FIH is rescued by HS− binding to Fe3+ and reducing it to Fe2+.

Coincidentally, maximum inhibition is observed at approximately physiological pH, lending credence to the hypothesis that the HIF hydroxylases are physiological targets of

H2S. The data presented here offer insight into possible physiological targets as well as a mechanism of action for H2S function in cells, and suggests that other αKG dependent oxygenases may also be potential targets.

4.5 Supplemental Figures

The IC50 is unchanged in the presence of high αKG, suggesting that sulfide is an uncompetitive inhibitor of αKG.

150

Figure 4.9: IC50 measurement at high αKG levels FIH (0.5 μM) in 50 mM Hepes pH 7.00 at 37°C; αKG (500 μM), FeSO4 (25 μM), CTAD (100 μM), Sulfide (0 – 120 μM), ascorbate (2 mM).

Figure 4.10: Concentration dependence for sulfide rescue of auto-hydroxylated FIH activity. Rescue is followed by inhibition. FIH (5 μM), FeSO4 (5 μM), αKG (185 μM), Sulfide (0 – 1.5 mM) in 50 mM Hepes pH 7.40.

Table 4.1: Summary of IC50 values pH IC (µM) 50 6.00 307 ± 19

6.25 244 ± 15

6.50 152 ± 3

6.75 137 ± 3

7.00 64 ± 3

7.50 34 ± 2

8.00 45 ± 4

151

Figure 4.10: Anaerobic (black) and aerobic (red) UV-vis samples of Fe (400 µM), NOG/αKG (2 mM), and HS- (400 µM) in 50 mM Hepes pH 7.40.

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