Studies of Iron Sulfur Cluster Maturation and Transport DISSERTATION Presented in Partial Fulfillment of the Requirements for Th

Studies of Iron Sulfur Cluster Maturation and Transport

DISSERTATION

Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy in the Graduate School of The Ohio State University

Jingwei Li

Graduate Program in Chemistry

The Ohio State University

2015

Dissertation Committee:

Professor James A. Cowan, Advisor

Professor Ross E. Dalbey

Professor Claudia Turro

Jingwei Li

2015

Abstract

Cellular iron homeostasis is critically dependent on sensory and regulatory mechanisms that maintain a balance of intracellular iron concentrations. Divergence from a healthy iron concentration can result in common disease states such as anemia and ataxia. With the goal of understanding the molecular basis for such health problems, and advancing the knowledge based toward potential remedies, an understanding of the molecular details of cellular iron transport and the biological chemistry of iron species is an essential prerequisite. In that regard, iron-sulfur clusters are ubiquitous iron-containing centers in a variety of proteins and serve a multitude of roles that include electron transfer, catalysis of reactions, and sensors of cellular oxygen and iron levels.

Recently, a substantial body of evidence has suggested an essential role for cellular glutathione (a molecule normally implicated with eliminating reactive oxygen species from cells) in the regulation, stabilization and biosynthesis of cellular iron-sulfur clusters in humans and other complex organisms. We have demonstrated that glutathione can naturally bind to iron-sulfur cluster precursors and have isolated and characterized this species and shown it to be stable under physiological conditions. More importantly, we have demonstrated that the glutathione-bound iron-sulfur cluster can be transported by a

ii critical export protein from the cellular mitochondrion. The glutathione iron-sulfur cluster can also be bound by other critical cellular proteins and we hypothesize to be a new component of the so-called labile iron pool – an ill-defined collection of iron species prevalent inside a cell.

To develop a further understanding of the cellular chemistry of these species, a multidisciplinary approach is being taken to elaborate their structural and functional chemistry. Proteoliposome studies are underway to explore the transport chemistry of the glutathione-iron-sulfur cluster complex as a viable substrate candidate for the mitochondrial export protein. Protein engineering of the transport protein will yield vital information on the interactions of the glutathione iron sulfur cluster with the protein and help to elucidate the mechanistic origin of the dysfunctional mitochondrial iron transport systems associated with this protein, which also underlie two common hereditary diseases

(namely sideroblastic anemia and cerebellar ataxia). A variety of biophysical and molecular biology techniques, in addition to protein and lipid biochemistry are also being applied to the problem. For example, tiron chelation assay and flow cytometry assays are also applied to track and study the kinetic parameters of cluster transport across the membrane bilayer.

My studies also extend to understanding the role of protein mobility in promoting the functional chemistry of these complex systems. We have found the dynamics of structural change to be unusual for many of the proteins involved in the cellular

iii biosynthesis and transport of iron cofactors, and an understanding of these properties will help us to fully understand how they function in the broader context of the cell.

Dedication

This document is dedicated to my parents.

Acknowledgments

My graduate school experiences have taught me many lessons, and also helped me to grow as a scientist and as a person. I am heartily grateful to my adviser, Professor James

A. Cowan, for his encouragement and guidance during this time. The great support and inspiring research environment Dr. Cowan has provided made it possible for me to explore a vast field of scholarly ideas and experiments. Dr. Cowan’s valuable advices have also helped to shape my career goal as a physician scientist, and his encouragements and supports provided a solid foundation for my pursuit of a career in biomedical research.

I am also thankful to my committee members: Dr. Ross Dalbey and Dr. Claudia Turro, for the valuable insights they shared with me, and the encouragement they have provided.

It has been an honor and a pleasure for me to work with so many great colleagues. I would like to thank Wenbin, Jeff, Seth and Lalintip for the great help they gave me when

I joined the group. I have learned especially a great deal in biochemical theories and techniques from Wenbin, and his help and support have been an essential element to my achievements during graduate school. I have enjoyed great discussions with my

vi colleagues: James, Jessica, Insiya, Zhen, Steve, Christine, Andrew and Sam, and deeply appreciate their valuable comments and suggestions on my projects and manuscripts. I would like to thank Steve for his tireless efforts in collaborating with me on various projects we worked on during the last stage of my graduate school career.

I would like to thank Dr. Dennis Bong for his generosity by letting me using his instruments such as the Dynamic Light Scattering spectrometer and the ultracentrifuge for my proteoliposome studies. I would like to thank Drs. Chunhua Yuan and Tanya

Young for their valuable insights and enormous help in NMR experiments. I’d like to thank Drs. Cecilia Chain, Gustavo Pasquevich and Alberto Pasquevich for their collaborative effort in Mossbauer studies.

Most importantly, I am thankful to my mother and all of her encouragement, love and support during my academic career. My accomplishments and achievements would not have been possible without her help.

vii

Vita

2009...... B.S. Chemistry, The Ohio State University

2009 to present ...... Graduate Teaching and Research Associate, Department of

Chemistry, The Ohio State University

Publications

1. Jingwei Li and J.A. Cowan, “Glutathione-Coordinated [2Fe-2S] Cluster. A Viable

Physiological substrate for Mitochondrial ABCB7 Transport”, Chemical

Communications, 10.1039/C4CC09175B

2. Wenbin Qi, Jingwei Li and J.A. Cowan, “A structural model for glutathione-

complexed iron–sulfur cluster as a substrate for ABCB7-type transporters”

Chemical Communications, 2014, 50, 3795-3798

3. Jingwei Li, Shu Ding and J.A. Cowan, “Iron-Sulfur Cluster Biosynthesis:

Thermodynamic and Structural analysis of Human NFU Conformational

Chemistry” Biochemistry, 2013, 52, 4904-4913

4. Wenbin Qi, Jingwei Li, C. Y Chain, G.A. Pasquevich, A. F. Pasquevich, and J. A.

Cowan “Glutathione-Complexed Iron-Sulfur Clusters. Reaction Intermediates and

viii

Evidence for a Template Effect Promoting Assembly and Stability”, Chemical

Communications, 2013, 49, 6313-6315.

5. Wenbin Qi, Jingwei Li and J.A. Cowan, “Human Ferredoxin-2 displays a unique

conformational change” Dalton Transactions. 2013, 42, 3088-3091.

6 Wenbin Qi, Jingwei Li, C. Y Chain, G.A. Pasquevich, A. F. Pasquevich, and J. A.

Cowan Glutathione Complexed Fe-S Centers. J. Am. Chem. Soc. 2012 134,

10745-10748.

Fields of Study

Major Field: Chemistry

Table of Contents

Abstract ...... ii

Dedication ...... v

Acknowledgments...... vi

Vita ...... viii

Publications ...... viii

Table of Contents ...... x

List of Tables ...... xvii

List of Figures ...... xx

List of Schemes ...... xxvii

Chapter 1: Introductions – Iron and iron-sulfur clusters: their homeostasis and biological relevance...... 1

Background ...... 1

Mechanism of iron sulfur cluster formation ...... 12

Maturation of iron-sulfur cluster containing proteins ...... 15

Redox properties of iron-sulfur cluster proteins ...... 17

Mitochondrial export of iron-sulfur clusters ...... 19

Study of cluster transport using proteoliposome models ...... 23

References ...... 25

Chapter 2: Thermodynamic and Structural Analysis of Human NFU Conformational

Chemistry ...... 40

Introduction ...... 40

Materials and Methods ...... 43

Expression and purification of human NFU ...... 43

Expression of 15N labeled N-terminal, C-terminal and Full Length NFU ...... 44

Differential Scanning Calorimetry ...... 44

Nuclear Magnetic Resonance Spectroscopy...... 45

Circular Dichroism Spectroscopy ...... 46

Results ...... 48

DSC Studies of N-Terminal Human NFU...... 48

VTCD Studies of N-Terminal Human NFU...... 52

DSC Studies of C-Terminal Human NFU ...... 53

VTCD Studies of C-Terminal Human NFU ...... 54

DSC and VTCD Studies of Full Length Human NFU...... 57

DSC Studies of a Mixture of N- and C-Terminal Domains of NFU ...... 60

VTCD Studies of a Mixture of N- and C-Terminal Domains of NFU ...... 62

[15N-1H] HSQC Studies of a Mixture of N- and C-Terminal Domains of NFU ...... 62

Discussion ...... 68

Conclusion ...... 76

Additional Figures ...... 77

References ...... 81

Chapter 3: Temperature Induced Conformation Study of Human Ferredoxin 2 ...... 85

Introduction ...... 85

Materials and Methods ...... 89

Cloning and Expression of human ferredoxin2 ...... 89

Purification of proteins ...... 89

hFd1 and hFd2 temperature dependent absorbance change monitored by UV/Vis

spectroscopy ...... 89

hFd2 temperature dependent CD signal change ...... 90

Rate of ferredoxin reduction by adrenodoxin reductase and cluster transfers

monitored by the cytochrome c assay ...... 90

Nuclear magnetic resonance spectroscopy ...... 91

Results ...... 92

Variable Temperature UV/Vis Study ...... 92

xii

Variable Temperature CD study ...... 93

DSC Studies ...... 96

Cytochrome C Assays ...... 98

Discussion and Conclusion ...... 102

References ...... 104

Chapter 4: Synthesis, characterization, and studies on glutathione coordinated [2Fe-2S] clusters ...... 106

Introduction ...... 106

Results ...... 108

Mass Spectrometry Studies ...... 108

UV/Vis Absorption Studies ...... 117

Mossbauer Spectroscopy Studies ...... 120

Solution Ionic Strength and Cluster Stability ...... 125

Nuclear Magnetic Resonance Studies ...... 141

Experimental Methods ...... 148

Cluster synthesis and ESI-MS analysis ...... 148

Synthesis of modified glutathiones...... 148

UV/Vis Absorption Assays ...... 150

Mössbauer Spectroscopy ...... 151

xiii

Salt-Dependence of cluster stability and measurement of kinetics ...... 153

Nuclear Magnetic Resonance Experiments ...... 156

Conclusion ...... 157

References ...... 159

Chapter 5: Synthesis, Characterization and Biological Chemistry of a Glutathione

Complexed [2Fe-2Se] Cluster ...... 165

Introduction ...... 165

Materials and Methods ...... 168

Synthesis and charaterization ...... 168

Nuclear Magnetic Resonance Experiments ...... 169

Mössbauer Spectroscopy ...... 170

Protein Purifications and Reconstitution of Fe2Se2 clusters ...... 170

FeSe cluster transfer between glutathione complex and hISU D37A ...... 172

Cluster transfer from hISU to Fdx2 ...... 172

Results ...... 173

Synthesis and characterization ...... 173

Nuclear Magnetic Resonance Spectroscopy ...... 176

[2Fe-2Se] glutathione cluster transfer to apo-hISU ...... 182

[2Fe-2Se] cluster transfer from holo-ISU to apo-Fdx2 ...... 191

xiv

Electrochemistry ...... 197

Conclusion ...... 201

References ...... 203

Chapter 6: Glutathione-Coordinated [2Fe-2S] Cluster as a Viable Physiological Substrate for Mitochondrial ABCB7 Transport...... 208

Introduction ...... 208

Materials and Methods ...... 211

Molecular cloning ...... 211

ATPase Activity assay ...... 213

Protein Purification ...... 215

Synthesis of liposomes ...... 220

Synthesis of Atm1p proteoliposomes ...... 221

Computational Modeling ...... 224

Synthesis of fluorescein-labeled glutathione ...... 225

Synthesis of fluorescein-labeled cluster ...... 227

Flow cytometry analyses of proteoliposomes ...... 228

Tiron Assay...... 231

Titration of fluorescein-labeled cluster to Atm1p ...... 233

Titration of GSH to a solution of fluorescein ...... 234

Data fitting and Statistical analysis ...... 235

Results ...... 236

Atm1p ATPase Activity Assays ...... 236

Flow Cytometry Assays...... 242

Tiron Assays ...... 251

Conclusion ...... 255

References ...... 257

Bibliography ...... 263

xvi

List of Tables

Table 1: 15N-1H HSQC acquisition and processing parameters...... 45

Table 2: Melting temperatures for N-terminal NFU, C-terminal NFU, a mixture of N- and

C-terminal domains, and full-length NFU determined by DSC...... 50

Table 3: ΔHcal and ΔHv for N-terminal NFU, C-terminal NFU, a mixture of N- and C- terminal domains, and full-length NFU determined by DSC...... 50

Table 4: Circular dichroism analyses of N-terminal NFU, C-terminal NFU, a mixture of

N- and C-terminal domains, and full-length NFU...... 53

Table 5: Melting temperatures of N-terminal NFU, C-terminal NFU, a mixture of N- and

C-terminal domains, and full-length NFU determined by VTCD...... 56

Table 6: Fitting parameters for the change in Van’t Hoff enthalpy determined by VTCD.

...... 56

Table 7: A list new cross-peaks observed in the HSQC spectrum of the 15N-labeled C-

NFU domain following complex formation with unlabeled N-NFU...... 64

Table 8: A list of new cross-peaks observed in the HSQC spectrum of full-length NFU relative to the spectra for the individual N- and C-NFU domains...... 66

Table 9: Temperature dependent secondary structural prediction and analysis by K2D3.

...... 96

xvii

Table 10: Enzymatic parameters for ferredoxin electron exchange with adrenodoxin reductase...... 98

Table 11: Cluster transfer kobs by use of the cytochrome c assay...... 99

Table 12: Mossbauer parameters of solid-state cluster and assigned oxidation state, geometry and percent contribution...... 124

Table 13: A list of ions used and values of solution ionic radii, charge density, hydration enthalpy, KS and correction factor from data fit...... 135

Table 14: Relaxation times of protons of GSH, GSSG, GSH + Fe3+ and GSH coordinated cluster...... 145

Table 15: Mossbauer parameters of the Fe2Se2(GS)4 cluster...... 181

Table 16: A comparison of Mossbauer parameters of selenium substituted cluster with previously published results...... 181

Table 17: Observed rate constants for the cluster transfer from glutathione coordinated clusters to apo hISU native or D37A proteins...... 183

Table 18: Observed rate constants for the cluster transfer from the holo scaffold protein hISU (WT or D37A) to apo Fd2...... 192

Table 19: Redox potentials determine by cyclic voltammetry for sulfur and selenium reconstituted proteins. (Data collected in collaboration with Stephen Pearson) ...... 200

Table 20: Concentrations and volumes of components for the PCR mutagenesis of wildtype Atm1p to R284E...... 212

Table 21: Reaction times and temperatures for the PCR mutagenesis of wild-type Atm1p to R284E...... 212

xviii

Table 22: Parameters for [2Fe-2S](GS)4 and glutathione stimulation of transporter

ATPase activity...... 239

xix

List of Figures

Figure 1: A scheme of iron trafficking for eukaryotic cells...... 2

Figure 2: A scheme of endocytosis of iron by transferrin and transferrin receptor...... 4

Figure 3: Various types of Fe-S clusters...... 5

Figure 4: Substrate-bound active site of aconitase...... 7

Figure 5: isc operon and associated gene products...... 9

Figure 6: A proposed model for iron-sulfur cluster biogenesis by hISU, and the subsequent cluster transport to apo proteins...... 13

Figure 7: Differential scanning calorimetry profile for a 0.3 mM N-NFU solution in phosphate buffer...... 48

Figure 8: Rescan of the N-NFU domain ...... 51

Figure 9: VTCD analysis of 10 μM N-NFU in 40 mM phosphate 100 mM NaCl buffer. 52

Figure 10: DSC and VTCD analysis of C-NFU in 40 mM phosphate 100 mM NaCl buffer...... 55

Figure 11: DSC and VTCD analysis of NFU in 40 mM phosphate 100 mM NaCl buffer.

...... 58

Figure 12: DSC and VTCD analysis of a N-NFU / C-NFU mixture in 40 mM phosphate

100 mM NaCl buffer...... 61

Figure 13: 1H-15N HSQC spectra of (A) 15N-N-NFU, (B) [15N-N-NFU + 14N-C-NFU] and

Figure 14: 1H-15N HSQC spectra of (A) 15N-C-NFU, (B) [14N-N-NFU + 15N-C-NFU] and

Figure 15: [15N-1H] HSQC spectra indicate an alternative conformation for C-NFU following the interaction with N-NFU...... 70

Figure 16: [15N-1H] HSQC NMR spectrum of an equimolar mixture of 14N-N-NFU and

15N-C-NFU ...... 71

Figure 17: Thermal profile for N-NFU...... 72

Figure 18: Gibbs free energy plot illustrates a mixture of N-NFU and C-NFU domains to be most thermally stable near physiological temperature...... 73

Figure 19: DSC and VTCD analysis of N-NFU in 40 mM phosphate buffer...... 77

Figure 20: DSC and VTCD analysis of C-NFU in 40 mM phosphate buffer...... 78

Figure 21: DSC and VTCD analysis of NFU in 40 mM phosphate buffer...... 79

Figure 22: DSC and VTCD analysis of N+C-NFU in 40 mM phosphate buffer...... 80

Figure 23: Crystal structure of hFd1 ...... 86

Figure 24: Crystal structure of hFd2 ...... 86

Figure 25: Comparison of UV/vis absorption spectra for hFd1 and hFd2...... 87

Figure 26: Comparison of CD spectra for hFd1 and hFd2...... 88

Figure 27: Absorbance change during heating cycle for hFd1 and hFd2...... 92

Figure 28: CD signal from the cluster center when holo hFd2 is incubated up to 55 C. .. 94

Figure 29: Far UV absorption spectra obtained at various temperatures ...... 95

xxi

Figure 30: Differential scanning calorimetry experiments of hFd2 (top) and hFd1

(bottom)...... 97

Figure 31: Overlay of variable temperature [15N-1H] HSQC spectra for hFd1 (A) and hFd2 (B) at 28 C (black) and 55 C (red)...... 100

Figure 32: Comparison of chemical shift perturbations for hFd1 and hFd2 after heating to

55 C...... 101

Figure 33: Analysis of [2Fe-2S](GS)4 formation by ESI mass spectrometry...... 109

- 2+ + + Figure 34: Simulated mass spectrum of [(GS )4[Fe2S2] +2H +Na ]...... 110

Figure 35: The template effect of pre-assembled glutathione tetramer ...... 111

Figure 36: ESI-MS data showing solution aggregates of glutathione molecules...... 112

Figure 37: Oligomers were observed for glutathione molecules ...... 113

Figure 38: A two-dimensional representation of a glutathione-complexed cluster aggregate...... 115

Figure 39: Plot of the peak intensity of the mixed valence Fe3+/Fe3+ form at m/z=1425.3, and the Fe3+/Fe2+ form at m/z=1426.3, versus reaction time...... 117

Figure 40: The UV-Vis spectrum of a 50 uM solution of [Fe2S2](GS)4 ...... 118

Figure 41: Time-dependent cluster stability study recorded by UV-Vis spectrophotometer at 330 nm...... 119

Figure 42: Mössbauer spectra of isolated cluster in solution state...... 121

Figure 43: Mössbauer spectrum from a sample extracted from the reaction mixture 10 min after mixing...... 123

Figure 44: A plot of cluster initial degradation rate at various concentrations of salt .... 128

xxii

Figure 45: A concentration standard curve for glutathione iron-sulfur clusters dissolved in buffer ...... 129

Figure 46: The KS values were determined from derived equations for various salts. ... 131

Figure 47: A plot of charge density (z/r) vs. log (KS) for selected ions...... 134

Figure 48: A plot of ΔHm,hyd vs. log (KS) for selected ions...... 136

Figure 49: The log of k/k0 was fit against the square-root of ionic strengths...... 138

Figure 50: A slow cluster breakdown was observed when dissolved in solution without additional salt added...... 139

Figure 51: NMR spectra of glutathione cluster...... 141

Figure 52: Variable temperature NMR studies of 1H resonances at 27 ºC, 30 ºC, 40 ºC and 50 ºC...... 143

Figure 53: Proton Homonuclear Decoupling of Cluster at 4.70 ppm...... 144

Figure 54: Comparisons of relaxation times between free and [2Fe-2S] bound glutathione...... 145

Figure 55: Sodium cyanide titration to a solution of glutathione cluster...... 147

1 Figure 56: H NMR spectrum of N-acetylglutathione in D2O...... 149

1 Figure 57: H COSY NMR spectrum of N-acetylglutathione in D2O...... 150

Figure 58: [Fe2Se2](GS)4 cluster absorption spectrum ...... 174

Figure 59: ESI-MS spectrum of Fe2Se2GS4 cluster...... 175

Figure 60: Cluster stability under various conditions...... 176

1 Figure 61: H NMR spectra comparison of Fe2S2GS4 cluster and Fe2Se2GS4 cluster. ... 177

Figure 62: Mossbauer spectra of the Fe2Se2GSH4 cluster ...... 180

xxiii

Figure 63: CD scans of Fe-S reconstituted proteins and Fe2S2(GSH)4 cluster...... 184

Figure 64: CD scans of Fe-Se reconstituted proteins and Fe2Se2(GSH)4 cluster...... 185

Figure 65: CD scans of apo hISU and apo Fd2...... 186

Figure 66: Transfer of Fe2S2(GSH)4 in solution to apo WT hISU monitored by CD. .... 187

Figure 67: Transfer of Fe2S2(GSH)4 in solution to apo D37A hISU monitored by CD. 188

Figure 68: Transfer of Fe2Se2(GSH)4 in solution to apo WT hISU monitored by CD. .. 189

Figure 69: Transfer of Fe2Se2(GSH)4 in solution to apo D37A hISU monitored by CD.

...... 190

Figure 70: Transfer of Fe2S2 reconstituted WT hISU to apo Fd2 monitored by CD. .... 193

Figure 71: Transfer of Fe2S2 reconstituted D37A hISU to apo Fd2 monitored by CD. . 194

Figure 72: Transfer of Fe2Se2 reconstituted WT hISU to apo Fd2 monitored by CD. ... 195

Figure 73: Transfer of Fe2Se2 reconstituted D37A hISU to apo Fd2 monitored by CD. 196

Figure 74: Cyclic voltammetry experiments...... 198

Figure 75: Cluster transfer from glutathione cluster to D37A apo hISU monitored by UV/

Vis spectroscopy...... 201

Figure 76: An overview of the ATPase activity assay...... 213

Figure 77: Atm1p and Mg-ATP binding parameters...... 214

Figure 78: Purification of Atm1p...... 217

Figure 79: SDS-PAGE gel of Atm1p...... 218

Figure 80: SDS-PAGE of WT Atm1p and mutants...... 219

Figure 81: ATPase activity of purified Atm1p...... 220

Figure 82: Lipids were mixed at a molar ratio of 1:1:1 to synthesize lipid membrane. . 221

xxiv

Figure 83: Proteoliposome synthesis...... 223

Figure 84: ATPase activity of reconstituted proteoliposome...... 224

Figure 85: ESI-MS analysis of Fl-GSH...... 226

Figure 86: Raw flow cytometry data ...... 229

Figure 87: Control experiments to monitor the integrity of the proteoliposomes throughout the transport reaction...... 230

Figure 88: Synthesis of proteoliposome, ATP-driven cluster transport and ferric ion quantitation of product...... 232

Figure 89: Tiron coordinates to ferric ions, and absorb at 550 nm...... 233

Figure 90: Titration experiment showing the decrease in fluorescence signal intensity as labeled-cluster was added to Atm1p...... 234

Figure 91: Titration experiment showing GSH quenching the fluorescein fluorescence signal ...... 235

Figure 92: Glutathione stimulates the ATPase activity of Atm1p ...... 237

Figure 93: WT Atm1p ATPase activity is stimulated by cluster ...... 238

Figure 94: Atm1p ATPase activity is stimulated by [2Fe-2S](GS)4...... 239

Figure 95: Modelled structured of WT Atm1p and R284E mutant...... 241

Figure 96: Crystal structure of NaAtm1p ...... 242

Figure 97: Titration experiment showing cluster quenching the fluorescein fluorescence signal ...... 244

Figure 98: Atm1p-mediated cluster transport into fluorescein-loaded proteoliposome . 245

xxv

Figure 99: Initial velocity derived from the rate of fluorescein signal quenching following addition of cluster and Mg-ATP (exp)...... 246

Figure 100: Control experiments for cluster transport into fluorescein-loaded proteoliposome ...... 247

Figure 101: Fluorescence flow cytometry measurements of the proteoliposomes incubated with Mg-ATP (12 uM) and fluorescein labeled glutathione [2Fe-2S] cluster 248

Figure 102: Observed exchange rate of fluorescein labeled glutathione with solution glutathione...... 250

Figure 103: An area plot of fluorescein-GSH intensity observed on ESI-MS following degradation of Fluorescein labeled cluster...... 251

Figure 104: Concentration of iron inside the proteoliposome following 1 hr incubation with Mg-ATP and cluster...... 253

Figure 105: Proposed mechanism for mitochondrial iron-sulfur cluster transport by

ABCB7-type export proteins driven by ATP hydrolysis...... 255

xxvi

List of Schemes

Scheme 1: NFU sequence comparison ...... 41

Scheme 2: A proposed cluster degradation mechanism in the presence of salt ions based on time-dependent absorption spectroscopy results...... 130

Scheme 3: Synthesis of Fe2Se2GS4 cluster...... 169

Scheme 4: Cluster transfer between glutathione coordinated complex, hISU and Fd2 .. 183

xxvii

Chapter 1: Introductions – Iron and iron-sulfur clusters: their homeostasis and biological relevance.

Background

Iron is an essential element for many biological processes, and it is found in pathways in all eukaryotes. For example, in mammals, iron is used as a vital component in oxygen transport pathways in blood. The iron heme group found in hemoglobin serves as an essential cofactor to reversibly bind to oxygen molecules. Due to its relatively high nuclear stability, iron is one of the most abundant metals found on earth. Perhaps because of this and the vast chemical characteristics of iron, nature has shaped biological organisms to utilize iron in their cellular processes and functions. However, iron homeostasis and intracellular transport must be carefully handled by cells, as elevated levels of iron generates harmful oxygen radical species and lead to apoptosis. These tasks are meticulously regulated by complicated mechanisms, which over the past decades have been a popular facet of bioinorganic chemistry.

Figure 1: A scheme of iron trafficking for eukaryotic cells. Iron is transported from outside of cells via transferrin proteins, and endocytosis uptakes the iron into the cytoplasm and replenishes the labile iron pool.

Every day we ingest about 1-2 mg iron from food source, these iron are found in forms of both heme and nonheme motifs. The acidity and peptidase activities of our digestive systems aids the liberations of irons from their cofactors, and these irons are ultimately absorbed in the small intestines. Heme iron is relatively easier to be absorbed by the divalent metal transporters found in the brush border cells of the small intestines.

Nonheme irons, commonly found in the ferric state, must be reduced prior to absorption, and this redox reaction is carried out by the ferrireductase found in the duodenum. The

2 absorbed iron is transferred into muscle, bones and storage, at same time we lose about 1-

2mg of iron daily. Therefore, we have a constant hemeostatic flux of incoming and outgoing iron.

Iron are transferred throughout the body bound to the transferrin protein (Tf), which delivers two iron atoms to transferrin receptor (TfR) proteins embedded on the cellular surface (Figure 1). Iron is subsequently endocytosized into the cell in the form of a vesicle (Figure 2). Proton pumps lowers pH within the vesicle, causing iron to be release into the cytoplasm, as a part of the labile iron pool: a pool of chelatable and redox-active iron, which is transitory crossroad of cell iron metabolism and ready to be utilized by the cell for various functions.

Figure 2: A scheme of endocytosis of iron by transferrin and transferrin receptor.

To prevent over accumulation of iron and reactive oxygen species (ROS) in the cytoplasm, cells utilize highly compact iron storage method via the ferritin protein. This redox controlled iron storage pathway ensures the sequestration of excess iron ions, and provides a source of intracellularly stored iron when the need arises. The expression of

TfR, ferritin, and heme biosynthesis proteins are controlled by the iron-regulatory protein

1 (IRP1) via a trans-acting mechanism to stem-loop structures in cognate mRNAs, termed iron-responsive elements (IREs).

Other pathways for cells to utilize and avoid over accumulation of iron is via transport proteins such as ferroportin and mitoferrin, which export ferrous ions to the extracellular 4 space and mitochondria, respectively. Ferroportin is especially prevalent in hepatocytes and enterocytes, while mitoferrin is most commonly found in hematopoietic cells and is important in heme synthesis of hemoproteins and iron-sulfur cluster (ISC) assembly within the mitochondria. ISC are found in bacteria, plants and animals and are essential cofactors for many catalytic and redox proteins (Figure 3).

Figure 3: Various types of Fe-S clusters. From 1-Fe center such as Rubredoxin to more complicated proteins, such as 4Fe-4S cluster in High Potential Iron Sulfur Proteins (HiPIP).

ISC possess a vast range of chemical properties and consequently a plethora of enzymatic roles within the cell. An important role for E. Coli ISC is their regulation and response to reactive oxygen species and intracellular reduction potential via the soxRS system. Upon the oxidation of the ISC of SoxR by superoxide, and SoxS is upregulated which subsequently activates the transcription of the target antioxidation gene products, such as endonuclease IV, glucose 6-phosphate dehydrogenase and superoxide dismutase.1-4

Interestingly, another role for the iron-sulfur cluster cofactors includes aiding proteins in their structural integrity, such as the 4Fe-4S cluster found in E.Coli Endonuclease III. In cases such as this, the iron atoms of the cluster serves as Lewis acids for protein ligands,

5 and the holo structure is stabilized by both coordination bonds and the protein’s secondary and tertiary interactions.

In eukaryotic cells, ISC’s are known to regulate important cellular and biological functions, ranging from regulation of intracellular iron concentrations to tRNA modifications.5 In eukaryotes, cellular iron homeostasis is regulated by the iron regulon, which yields ~30 gene products that function in intracellular iron transport between mitochondria and vacuoles, such as MRS4, FET5 and FTH1. Interestingly, these genes are regulated based on the activity of the mitochondrial iron-sulfur cluster assembly and export machineries via Aft1 protein.6, 7 This indicates that for eukaryotic cells, the cellular iron concentration depends on the mitochondrial export product. Misregulated iron homeostasis, either during the cluster assembly process or the cytosolic transport process, results in transcriptional induction of the iron regulon, which causes the cell to over-accumulate iron in mitochondria6, 8-11 and results diseases such as x-linked sideroblastic anemia12, 13 or Friedreich’s ataxia.14 Other vital functions for cellular processes include examples such as NADH dehydrogenase, an essential protein located on the inner mitochondrial membrane for the aerobic cellular respiration.15, 16 In this case, the iron-sulfur prosthetic group is utilized as a redox center and provides a pathway for the electron transfer between NADH and coenzyme Q10. 16

The catalytic roles of iron-sulfur cluster cofactors are found in proteins such as aconitase and its conversion of citrate to isocitrate. In this case, irons within the cluster cofactor

6 serves as a Lewis acid during the rearrangement mechanism of the catalytic cycle. In these proteins, instead of protein ligand coordinating to the ISC, one of the coordination sites is bound to water molecules, and upon substrate binding, this site is utilized as a site for catalysis (Figure 4). 17

Figure 4: Substrate-bound active site of aconitase. Citrate is converted to isocitrate by aconitase, which utilizes [4Fe-4S]+ to catalyze the reaction. Iron expands it coordination number from 4 to 6 during the process, with two of the substrate’s oxygen atoms as additional ligands.17

Free ferric or ferrous ions are toxic to cellular component due to its participation in

Fenton reactions which produces reactive oxygen species. Sulfide are also toxic to biological systems, as it readily complexes with metals in key cellular respiratory enzymes. Therefore, the existence of enzymatic donors of iron and sulfur which interact with apo proteins to form iron-sulfur clusters. However, this is rather an inefficient process since each apo-protein must have their iron and sulfur donors which specifically interact with the recipient protein. The hypothesis and identification of scaffold proteins later solved this problem, and these iron sulfur cluster “factories” provide an efficient and 7 energy economical pathway for reconstitution of a wide range of iron sulfur cluster proteins with just a few scaffold motifs. It should be noted that this sophisticated iron and sulfur reconstitution system must be tightly regulated and stabilized, as loss of ferrous or sulfide during the transfer process would result in toxicity to cells.

Seminal studies on iron-sulfur cluster formation have revealed a consensus that a reducing solution condition was necessary for apo-proteins to uptake iron and sulfur. In more recent studies, auxiliary proteins have also been identified in vivo to aid the reconstitution process. These scaffold proteins were first identified in Azotobacter vinelandii nitrogenase system. 18-24

In bacteria, there are three known systems for iron-sulfur cluster biogenesis. In particular, the NIF system found in azototrophic bacteria serve as an example of ISC biosynthesis for the organism’s essential nitrogen fixation system. The other two systems are ISC and

SUF systems, which serves as iron-sulfur cluster maturation pathways, encoded and regulated by the isc and suf operons, respectively. The suf operon’s major role is to regulate the expression of iron-sulfur cluster chaperone proteins during oxidative stress,25-28 where as the isc operon controls a variety of proteins which functions to assemble iron sulfur clusters.29 ISC homologues have been identified in most sequenced genomes30, and thus likely represent a fundamental Fe-S cluster biosynthesis pathway.

The isc operon is regulates the gene products29 of a repressor protein IscR31, a sulfur donor protein IscS32, an iron sulfur cluster scaffold protein IscU33-35, and a redox protein

8 ferredoxin (Figure 5).36 Convincing experimental evidences31 have paved a way for the hypothesis that the gene products of the isc operon is regulated by the cellular concentration of iron-sulfur clusters, which is monitored and repressed when the concentration of intracellular holo IscR. When intracellular level of iron-sulfur clusters are low, the repressor is relieved which yields production of the aforementioned isc gene products and maintains an intracellular homeostatic concentrations of iron-sulfur clusters.

Figure 5: isc operon and associated gene products. HscA and Hsc B are co-chaperone proteins for IscU during the cluster synthesis process.29

Compared with the bacterial iron-sulfur cluster synthesis system, the eukaryotic system is relatively more complicated and involves with a wider range of proteins, and in some instances includes partial derivatives from the bacterial system. For example, the ISC biosynthesis pathway has been incorporated from the bacterial system, and in addition to this, new pathways have been uniquely identified as well, such as the cytosolic ISC assembly pathway (CIA). The ISC system is associated with the mitochondrial cluster synthesis pathway, whereas the cytosol relies on the CIA to construct and reconstitute iron-sulfur clusters to target proteins.37 Similar to the bacterial systems, iron-sulfur clusters in the eukaryotic system are synthesized by a several pathways. The central

9 biogenesis of iron-sulfur clusters are within the mitochondria, which contains an iron- sulfur cluster assembly machinery – ISC, which has been proposed to be inheritated from a similar system found in protomitochondria of α-proteobacteria. The other two systems are found in the cytosol and nucleus, each includes components involved in the maturation of Fe/S proteins within their respective compartments. However, it is known that these iron-sulfur protein assembly and maturation processes require the participation of components from the mitochondria, as well as an ABC transporter on the mitochondrial membrane. It is interesting to note that the ISC system found in eubacteria is known to synthesize all cellular iron-sulfur protein, and perhaps this explains the vast and vital functions of the mitochondrial ISC system, where cytoplasmic iron homeostasis is observed to be severely impacted with mutated ABC transporter that translocate important iron-sulfur cluster species from the mitochondria.

The eukaryotic mitochondrial cluster assembly and maturation processes require several systematic components and many chaperone proteins for iron and sulfur, as well as electrons. Isu1 (homolog Isu2 in Saccharomyces cerevisiae) is the scaffold protein involved in the Fe/S cluster assembly process. This molecular “factory” accepts iron from iron donor frataxin and sulfur from Nfs1/Isd11 and catalyzes the formation of [2Fe-2S] cluster, which is subsequently transferred to apo-proteins. It is important to note that the iron transport from frataxin requires the iron to be in its ferrous oxidation state. Similarly,

Nfs1/Isd11liberates the sulfur in its reduced (2-) state. Electrons used in reducing the persulfide bond and the subsequent sulfide formation in by the desulfurase comes from

Yah1, a redox enzyme with homologs which are also found in yeast and bacterial systems.

Relative to the bacterial system, iron and iron sulfur cluster concentration regulation in eukaryotic systems involves a more complicated pathway. Expression of a series of iron transport and storage proteins is regulated by Aft1p, the iron responsive transcription factor.7, 38-42 For example, the transcription of iron sulfur cluster scaffold protein ISU1(or

ISU2 in yeast) is induced when aft1 is activated.43 Under iron-replete conditions, glutaredoxins Grx3/4p is reconstituted with 2Fe-2S clusters and this holo-protein interacts with Aft1p.44 This process results in dissociation of Aft1p from its promoter target, and terminate the activated transcription of associated iron regulatory genes.44 The concentration of iron sulfur clusters is controlled by a post-transcriptional regulation mechanism via cytosolic iron regulatory protein (IRP1). 45 Under iron replete conditions, a [4Fe-4S] is found in the IRP1 and it exhibits aconitase activity. When the cytosolic concentration of iron sulfur cluster is low, the holo-protein loses its labile cluster, which results in a structural change and IRP1 gains the affinity to bind the iron-responsive element (IRE). These IREs are found on mRNAs with translational products involved in cellular iron storage and uptake. Depending the identity and role of these gene products, the apo IRP1 may bind to either the 5’ or the 3’ end of the mRNA, which results in either blocked or extended translation of the mRNA, respectively. 46-49

Mechanism of iron sulfur cluster formation

The precise mechanism of iron-sulfur cluster biogenesis is not well understood. However, a few general mechanisms that are consistent with published data have been proposed.

One of these mechanisms is described in Figure 6. In this pathway, iron is first donated to the scaffold protein ISU by frataxin.50-53 This provides a nucleation site for the full 2Fe-

2S cluster assembly on hISU. For both hISU and frataxin, conserved acidic residues are found near the surface of the proteins. For frataxin, this anionic surface is defined by a total of twelve acidic residues, as a part of α1 helix and β1 sheet.54 Five conserved residues, E100, E108, E11, D112 and D124, are conserved in all known frataxin homologs in other animals, plants, yeast and eubacteria. The conserved anion patch may served as an essential role in iron binding for frataxin, as they are also found on the aforementioned frataxin homologs.55 Interestingly, frataxin is known to possess more than one iron binding site, and calorimetry experiments indicated that up to seven ferrous ions may be found on frataxin.50 One explanation for the existence of many iron binding sites on frataxin may be due to the vast structure and properties of protein partners that frataxin interacts with. Frataxin’s binding affinity for iron increases when it is bound to protein partners.56, 57 For example, the binding affinity for iron increased from the uM range to nM range when bound to ferrochelatase.58 Another interesting observation is that frataxin’s binding affinity for partner proteins is significantly lower for apo-frataxin, suggesting that iron must be at the interfaces of the anion patches between frataxin and hISU.50 This first step of iron delivery from frataxin to hISU is further supported by the

12 electronic absorption experiments, which demonstrated that it is the rate-limiting step in the presence of either protein sulfur donor or when inorganic Na2S was used as the sulfide source.50

Figure 6: A proposed model for iron-sulfur cluster biogenesis by hISU, and the subsequent cluster transport to apo proteins.

In the human iron-sulfur cluster synthesis system, sulfur in the form of sulfide anion is provided by Nfs1, an evolutionarily conserved pyridoxyl-5'-phosphate (PLP) enzyme which functions as a desulfurase.59 Cysteine is converted to alanine by Nfs1 and partner protein Isd11, this results in the formation of a persulfide product on the active site

13 cysteine.22, 60, 61 The intermediate persulfide intermediate is then reduced back to cysteine and an inorganic sulfide ion, which is subsequently contributed to the formation of iron- sulfur cluster on the scaffold protein hISU. This reduction of persulfide process is not well understood, however, it has been proposed that human NFU is a potential enzyme that donates its thiol electrons and catalyzes the formation of inorganic sulfide by Nfs1.62

This hypothesis is based on the facts that human NFU possess a highly conserved thioredoxin-like CXXC motif and is a functionally competent enzyme for persulfide reduction.63-66 Human NFU is also known to form complexes with NifS-like proteins, and its function may mimic its homologous C-terminal domain of NifU, which provide further supports for its role as a reductase that mediates Nfs1 cysteinyl persulfide bond cleavage.67 Protein secondary and tertiary structures dictate protein functions, and misfolded proteins are often the molecular causes of a large number of diseases, including cystic fibrosis68-73, Creutzfeldt-Jakob disease (CJD)74, 75, and Alzheimer's76-80.

Understanding the difference in the folding process can bring insight into the causes of how proteins function at the molecular level. The techniques of circular dichroism (CD) and Differential Scanning Calorimetry (DSC) are particularly powerful tool for this application, and were used to study the structure and function of the N-terminal and C- terminal domains of human NFU. These results are presented in later chapters of this thesis.

The sulfide transfer from Nfs1 to hISU is followed by oxidation of [2Fe-2S]0 to [2Fe-

2S]2+, catalyzed by ferredoxin.81 In this way, iron-sulfur clusters are formed transiently

14 on hISU.82-86 Of the cluster, iron forms coordination bonds with conserved cysteine residues found on hISU. There are a total of three cysteines, all of which are found near the surface of hISU, at the end of the barrel-like structure. 87, 88 This results in a solvent exposed cluster, which provides a mechanism for the final cluster product to be transferred to apo proteins, such as ferredoxin. A conserved aspartic acid residue (D37) is known as a part of the cluster transfer process, as its mutation lead to a much slower observed rate constant for the transfer process to ferredoxin.33, 89 Variable temperature electronic absorption experiments concluded that the conserved aspartic acid residue forms hydrogen bonds with a water molecule, and influences the solvent accessibility of

ISU-bound cluster. The D37A mutant also demonstrates a higher stability for the holo protein, indicating that the general cluster break down mechanism is likely due to hydrolysis, and the alanine mutant’s relatively more hydrophobic cluster pocket may reduce the tendency of water molecules from interacting with the cluster core.90

Maturation of iron-sulfur cluster containing proteins

Many apo-proteins require iron-sulfur cluster, synthesized by ISU, to interact with their natural partners and fully function. Example of these apo-proteins include ferredoxin, a relatively small (13kDa -14kDa) [2Fe-2S] redox protein previously introduced in the mechanistic section.91 In human, two ferredoxins have been identified in mitochondria, termed Fdx1 and Fdx2. Both proteins have been demonstrated to obtain their iron-sulfur cluster from ISU.92 Despite their high sequence similarity, these ferredoxins have been

15 demonstrated to differ in structure, protein partners and assume highly specific roles in distinct biochemical pathways within the mitochondria.93 These two proteins are functionally non-interchangeable, and participate in different fundamental physiological processes.93 The first ferredoxin (Fdx1) was originally identified to possess redox roles in iron-sulfur cluster maturation, conversion of heme O to heme A, and other steroidogenesis pathways.94-96 In 2002, Seeber and coworkers identified a second ferredoxin isoform (Fdx2) with conserved C-terminal sequence motif to Fdx1 in eukaryotes.92 In 2010, Sheftel et al. reported the proposed functions for each of these redox proteins.93 According to the study done by RNAi depletion in human cells and yeast complementation experiments, Fdx2 is a functional orthologue of Yah1, yeast redox protein responsible for iron-sulfur cluster maturation.97 This interesting observation prompted a re-evaluation of the functional role for Fdx1, which had been assumed to its mitochondrial roles due its sequence similarity to ferredoxins found in bacteria and plants. Fdx1 was demonstrated to only be able to reduce mitochondrial cytochrome P450 enzymes in bile acid formation, vitamin D synthesis and steroidogenesis during conversion of cholesterol to cortisol, pregnenolone, and aldosterone.96 Other important roles which had been previously tentatively assigned to Fdx1 were demonstrated to be the function of Fdx2. Fdx2 deficiency had a severe on effect on iron-sulfur protein biogenesis and cellular iron homeostasis. RNAi experiments of Fdx2 resulted in cluster loss of IRP, which increases the amount of gene product for transferrin receptor. The loss of iron- sulfur cluster on IRP also influenced the concentration ferritin, and these effects resulted in an over-accumulation of iron within the cell.

Redox properties of iron-sulfur cluster proteins

Despite the protein structural differences in these examples of ferredoxins, the overall mechanism of how redox chemistry works for iron-sulfur cluster proteins may be understood by the general molecular and electronic structures of these prosthetic groups.

Important factors fine-tune the redox potential of iron-sulfur clusters. These include the primary and secondary shell ligands, partial charge and hydrophobicity of the cluster pocket, as well as the network of hydrogen bondings near the cluster site.98 Of these, one of the most influential factors is the ligating atom. Sulfurs, in the form of thiolates, are most commonly found as protein Lewis base and forms a coordination bond to iron within a given iron-sulfur cluster. Examples of non-thiolate ligands are also found in nature, such as Rieske proteins, in which imidazoles are found to be coordinated to the iron center. In these examples, the lone-pair of the imine nitrogen is used to form coordination bonds with iron. In general, the atom or ligand system of the coordinating residue directly influence the redox properties of the iron-sulfur cluster. For example, when less electron rich ligands such as alkoxides were used instead of thiolates, the reduction potential of the iron-sulfur clusters were observed to increase due to the destabilization of the oxidized cluster.99-102

The solution conditions, such as pH, salinity and temperature, may fine-tune the redox properties of the iron-sulfur cluster. Proteins found in different cellular compartment may behave differently due to the local concentration of protons. For example, ligand residues

17 such as histidines or aspartates near the cluster pocket may become protonated in the presence of a low pH environment. In general, this favors the reduction process by eliminating the local negative charge within the pocket.103-105 Similarly, the presence of charged residues in the cluster pocket also dictates the stability of the cluster oxidation state.101 The hydrophobicity of the pocket also selectively stabilize either the reduced or oxidized form of the cluster and plays a key role in the redox potential. Pockets buried deep within the hydrophobic portion of a protein typically have a higher reduction potential. For example, in order to minimize the charge-hydrophobic interaction within the pocket, the reduced state of the 4Fe-4S HiPIP centers is more favored.106-108 In this way, a general positive correlation between the reduction potential and the extent of hydration level has been observed for iron-sulfur cluster redox proteins.109 Similarly, backbone amides have been reported to be important in favoring reduced clusters due to the induced dipoles. Although not nearly as polar as a hydrogen bond formed with water molecules, the induced dipole moment found in some iron-sulfur cluster proteins have the influence to fine-tune the redox potential of the prosthetic center.108, 110, 111 In several cases, the presence of second shell hydrogen bondings is found to be the differentiating factor between different subclass of ferredoxins, and mutant proteins without these conserved residues resulted in decreased reduction potential.112, 113 High reduction potentials are observed for proteins with second coordination shell hydrogen bonds, such as HiPIP, and removal of conserved hydrogen bonds in several cases resulted in a decrease in the reduction potential.101, 114

The number of iron present, as well as reducible iron atoms, also influences the electronic structure of the overall iron-sulfur cluster.113, 115 For example, in some HiPIP and ferredoxin proteins, more than one electron transfer are observed, and often resulting in a partial charge shared by more than one iron atom within the cluster.113 The most common geometry of the coordination environment of each iron atom is distorted tetrahedral, with the geometry of the cluster almost a square or cube for [2Fe-2S] and [4Fe-4Fe] centers, respectively. Structurally, the torsion angle of the cysteinyl β-carbon bond also influences the reduction potential of the cluster.106, 116, 117 These slight difference in molecular geometries and electronic structures lead to a fine-tuned variation in redox properties in iron-sulfur cluster proteins.

Mitochondrial export of iron-sulfur clusters

After the cluster is synthesized by scaffold protein ISU, the final destination for these clusters is apo-proteins, within the mitochondria as well as other cellular compartments.8,

118 A conserved aspartate near the cluster pocket strongly influences the stability of the freshly synthesized cluster.90 Studies using human and yeast models suggested a general base role for the carboxylate group. This conclusion was also based on the observations that cluster stability is significantly lowered under acidic conditions due to competition of cysteine protonations.119 Therefore, during cluster transfer from holo-ISU to apo-target protein, the conserved carboxylate is proposed to interact with a Lewis base residue, which either facilitate to or directly coordinate to the cluster. The apparent pKa of 6.9

19 suggest that such residue could be a cysteine residue on the target protein.90 Compared to these iron-sulfur proteins, cluster stability of holo-ISU was documented to be relatively labile.120 The overall cluster transfer process is assisted by a chaperone system which consists of ATPase Ssq1, co-chaperone Jac1, and nucleotide exchange factor Mge1.121, 122

There is a conserved LPPVK motif on ISU, which interacts with Ssq1 followed by an

ATP hydrolysis-dependent interaction.123, 124 This interaction is thought to induce a conformation change to ISU, perhaps labilizing the iron-sulfur cluster through interaction with the conserved aspartate.89, 125-127 Regardless of its final destination, the assembled cluster must be stabilized after synthesis, either rapidly transferred to the target protein, or transiently stored in forms of coordinated complexes. Unlike cytosolic iron storage systems, such as ferritin, there are no known iron-sulfur cluster storage proteins. The intrinsic reactivity towards oxidation or hydrolysis of the 2Fe-2S clusters requires coordination environments which must prevent these reactions. Previous studies have provided convincing evidence of glutathione molecules to be involved in the maturation of cytosolic iron-sulfur clusters, perhaps due to its available thiolate ligands and a relative high mitochondrial concentration (up to 10 mM).128 Interestingly in the yeast system, glutathione was demonstrated only to effect the maturation of cytosolic iron-sulfur cluster protein, and not mitochondrial iron-sulfur cluster proteins.8, 45, 129 A defective cytosolic iron-sulfur cluster biogenesis system was observed in yeast cells with depleted GSH, demonstrating a similar phenotypic effect as the down-regulation of Atm1.45 This indicates that after cluster assembly within the mitochondria, there exists a mechanism where glutathione molecules, either directly or indirectly, interact with cluster product

20 and stabilizes its transport out of mitochondria.130-132 Interestingly, a glutathione coordinated iron-sulfur cluster had been documented decades ago.133 However, due to the limitations of characterization tools available at the time, biochemical and cellular experiments were not pursued. A stable glutathione iron-sulfur complex is a viable substrate candidate for mitochondrial cluster export protein ABCB7. The possible cellular presence of a glutathione cluster complex which transiently store iron-sulfur cluster and participate in cellular iron-sulfur cluster biogenesis machineries would underscore an essential physiological role for these types for coordination complexes.

The mitochondrial export protein responsible for cytosolic iron-sulfur cluster maturations is ABC transporter Atm1(ABCB7 in humans), located on the mitochondrial inner membrane.8, 134, 135 The substrate of these export proteins has not been identified.

However, it is known that the component exported by these proteins is vital for both cytosolic and nuclear iron-sulfur cluster maturation. The substrate for ABCB7 and Atm1p is also thought to interact with transcription factors Aft1/2, which ultimately regulate cellular iron homeostasis.41, 136, 137 There has been several proposed theories on the identity and properties of the transported substrate. For example, a recent report provides genetic and biochemical evidence that Atm1 specifically transports glutathione disulfide

(GSSG) and/or glutathione trisulfide (GS-S-SG).138 However, the contents of this recently published paper did not actually demonstrate specificity in the transport of GSH,

GSSG, and GSSSG, nor did it provide actual genetic evidence for GSSSG as a physiological substrate. Furthermore, the finding that Atm1 ATPase activity was

21 stimulated by GSSG, but not GSH, and contradicts the report by Lill and coworkers who found both GSSG and GSH to stimulate Atm1 ATPase activity.139 Moreover, there is no published evidence supporting any physiological role for GSSSG, nor has such a species been implicated in natural iron-sulfur cluster biosynthesis. It is unclear why “sulfide” would need to be transported, if this pathway was actually occurring, given the prevalence of recognized sulfur donor proteins in the cytosol.

An alternative theory for the identity of the transported substrate is the glutathione coordinated iron-sulfur cluster. This cluster is stable under physiological conditions, and glutathione is a natural ligand for proteins such as glutaredoxins, and it is found as a

7, 140-143 ligand for Fe2S2 cluster in complex formed by Aft1 and its partner proteins Fra1/2.

These clusters are readily synthesized from ferric chloride and sodium sulfide in a pH buffered glutathione solution. The resulting product is stable for months after lyophilization, and can be used as test substrates for the transporter experiments with

ABCB7/Atm1p. The binding of the transported substrate can be readily tested by studying the stimulation of adenosine triphosphatase (ATPase) activity by the ligand.

There are known examples of ligand induced structural changes which ultimately influence the ATPase activity for these types of enzymes.139, 144, 145 Glutathione coordinated iron-sulfur clusters are promising substrates for ABCB7 family transporters.

There are several reasons to pursue the enzymatic experiments for ABCB7/Atm1p with glutathione coordinated iron-sulfur clusters. First, mitochondrial iron-sulfur cluster export apparatus is essential for cytosolic iron-sulfur cluster proteins, and a defect in the

22 mitochondrial iron-sulfur cluster machinery lead to an impaired cytosolic iron homeostasis.8, 128, 129 For example, yeast strain carrying a deletion in the ATM1 gene is an auxotrophy for leucine. This is due to a lack of iron-sulfur cluster in the cytoplasm, which lead to defected formation of iron-sulfur cluster protein Leu1, an isopropylmalate isomerase. Atm1 depleted cells shows growth with defect in the maturation of iron-sulfur cluster within the mitochondria, and not cytoplasm.8, 146 Similar phenotypes are also observed in Atm1p depletion, suggesting its vital role in cluster mitochondrial iron-sulfur cluster export. Due to the size constraints, with less than 10 A of channel diameter found in ABCB7 ortholog from Novosphingobium aromaticivorans (NaAtm1), the identity of the substrate is unlikely to be a protein-bound iron-sulfur cluster. However, in order for the iron-sulfur cluster to be free from hydrolysis and oxidation, the substrate must be protected from a coordinating pocket. Glutathione coordinated iron-sulfur clusters are much smaller than a protein bound cluster, yet it is stable under physiological conditions, and therefore is a likely candidate as the substrate transported by ABCB7/Atm1.

Study of cluster transport using proteoliposome models

In order to mimic natural protein folding and functions, membrane enzymes have been traditionally studied under detergent or liposome conditions.147, 148 For transport proteins, proteoliposomes are often used as a model system to study their binding chemistry and protein functions. Detailed protocols have been established to reconstitute ABC family proteins into premade liposomes.149 These liposomes may consist of various types and

23 compositions of lipids that may match the properties of the native membrane bilayers, such as hydrophobic thickness, phase transition, curvature, and lateral pressure.148, 150-153

Dependence on the composition of liposome for specific membrane orientations for ABC family proteins have also been documented.154 With these reconstituted proteoliposomes, plethora types of enzymatic experiments have been performed using various types of spectroscopy methods. For example, prototypical substrate Hoechst 33342 has been used to study the ABC membrane protein HorA, which is known to transport its substrates in proteoliposome models. The translocation of Hoechst 33342 is commonly studied by fluorescence spectroscopy methods, where fluorescence of the substrate is observed to decrease upon incorporation into the liposome.154 Radioactively labeled substrates have also been studied for the ABC family transporters.155-157 For example, maltose transport by membrane protein MalK has been studied with 14C labeled substrate and reconstituted proteoliposomes.155, 158 Using chelators and electronic absorption spectroscopy techniques may also provide a valuable tool to study metal translocations in proteoliposome models. Using iron chelators such as Tiron (disodium 4,5-dihydroxy-1,3- benzenedisulfonate), EDTA (Ethylenediaminetetraacetic acid), and BPTD

(bathophenanthrolinedisulfonic acid), iron sulfur cluster transport may be qualitatively and quantitative studied. Standard iron quantitation methods may be performed with proteoliposomes loaded with transported substrates. These studies will complement the

ATPase stimulation studies with ABCB7/Atm1p, and the combined results would provide valuable mechanistic and functional insights on the transporter protein within the proteoliposome.

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Chapter 2: Thermodynamic and Structural Analysis of Human NFU Conformational Chemistry

Introduction

Human NFU is a multifunctional protein that has been demonstrated to interact with the histone cell cycle regulation homologue A (HIRA) as a transcriptional regulator due to its ability to influence the chromatin structure.1, 2 Human NFU has also been linked to

Lafora disease due to its interaction with laforin, a protein which is found to be mutated in the disease state.3 NFU family proteins have also been implicated in cellular iron- sulfur cluster biosyntheses, both in vivo and in vitro.4-11 For example, the functional site11 of human NFU is located in the C-terminal domain (C-NFU, 83 residues, Scheme 1) and shares significant sequence identity with Nfu in Synechocystis Sp.,6 Nfu1 in yeast,7, 8 and

NfuA in Azotobacter vinelandii,10 all of which have been reported to be involved in the iron-sulfur cluster assembly pathway within their respective organisms.

In some cases Fe-S cluster binding has been noted and a possible role as an intermediate carrier of 2Fe or 4Fe iron-sulfur clusters has been proposed. For example, the Nfu-type protein from Synechocystis Sp has been reported in a 2Fe-2S form6 while E. Coli NfuA has also been demonstrated as an atypical carrier for 4Fe-4S clusters.9, 10 The conserved 40

CXXC motif found in these proteins has also been identified in the C-terminal domain of

NifU, the iron-sulfur cluster scaffold protein in nitrogen fixation bacterial system.12

Overall, the factors that promote cluster binding versus alternative functional roles remain unclear. The subject of this study, human NFU, possesses a C-terminal domain

(C-NFU) that contains a pair of redox active cysteines that demonstrate thioredoxin-like activity.13, 14 This domain has been shown to bind and mediate persulfide bond cleavage of sulfur-loaded IscS, the sulfide donor protein in the final step of sulfide delivery for

[2Fe-2S] cluster assembly on ISU-type scaffold proteins.15-17 Alternative mechanisms of cysteinyl persulfide cleavage by NFU have also been proposed, including direct reduction via electrons derived from ferrous ions18 and human ferredoxin,19 as well as a possible role for oxidized Fd in removing electrons from the nascent reduced [2Fe-2S]+ cluster.19

Scheme 1: NFU sequence comparison showing the N-terminal domain in bold and the C- terminal domain underlined.

Building from an early report,20 protein structural flexibility has emerged as an important theme in iron-sulfur cluster biosynthesis, particularly in the chemistry of the scaffold protein that promotes cluster assembly from iron and sulfide.20-25 Previous studies have also revealed the C-terminal domain of human NFU, another protein involved in cluster maturation, to demonstrate molten-globule-type structural behavior that may be of functional significance.13, 26, 27 These studies included titration of full length and truncated constructs of NFU with 1-anilino-8-naphthalenesulfonic acid (ANS), the kinetics of trypsin digestion, and heteronuclear single-quantum coherence (HSQC) NMR spectroscopy. By contrast, the N-terminal domain (N-NFU) retains a well-defined structure.

Herein, we describe a series of studies to further advance the understanding of the structural properties and thermal stabilities of N-NFU, C-NFU, full-length NFU (NFU), as well as a mixture of N-NFU and C-NFU (N-NFU/C-NFU). These studies provide additional support for an emerging theme in the biochemistry of iron-sulfur cluster biosynthesis;13, 26, 27 namely, that key parts of the protein machinery underlying Fe-S cluster assembly must display structural flexibility in order to fully execute their functions in the context of a multi-step process that could involve a variety of multi- protein complexes.21-25, 28 To advance this investigation we have made use of differential scanning calorimetry (DSC), a thermal analytical technique that measures the heat capacity of a defined experimental sample, as well as variable temperature circular dichroism (VTCD) experiments in combination with high-field NMR spectroscopy.

These bioanalytical methods not only provide information concerning the thermodynamic stability of proteins of interest, but also more detailed information on the characteristics of intermediate states involved in melting and unfolding processes.29

Materials and Methods

Expression and purification of human NFU

Expression and purification of human NFU was performed from BL21 Lysozyme plus

(DE3) competent cells as previously described.13, 26, 27 In brief, 50 mL LB culture

(supplemented with 30 µg/ml kanamycin) was grown overnight followed by 1 L culture growth to an OD600 ~ 0.6 with subsequent addition of 1 mM IPTG for protein induction

(3 h). The harvested cells were resuspended in Tris-buffer (50 mM Tris-HCl, pH 7.5) followed by sonication. The cell lysate was centrifuged by use of a Sorvall® RC-5B

Refrigerated Superspeed Centrifuge (Du Pont Instruments) at 26,890 x g and 4 °C for 30 min and the resulting supernatant was loaded onto TALON® Metal Affinity Column

(Clontech) equilibrated with Tris-buffer and eluted with 20 mM imidazole in Tris-buffer.

The purity of the eluted protein was checked by SDS-PAGE and identity was confirmed by ESI-Mass spectrometry.

Expression of 15N labeled N-terminal, C-terminal and Full Length NFU

15 1 15 For [ N- H] HSQC analyses, NH4Cl (99%, Cambridge Isotope Laboratory) supplemented M9 minimum medium (40 mM phosphate, 22 mM Glucose, 20 mM

14/15 NH4Cl,10 mM NaCl, 2 mM MgSO4, 0.1 mM CaCl2, 62 μM Kanamycin) was used to express 15N isotope labeled proteins. In brief, newly transformed BL21 Lysozyme plus

(DE3) competent cells were grown in 20 mL of LB medium to an OD600 of 1.0. After centrifugation, the cell pellet was resuspended in 0.5 L of unlabeled M9 medium and grown to OD600 of 0.8. The resulting cells were spun down and inoculated in 2 L of labeled M9 medium and grown to OD600 of 0.4, followed by IPTG induction (1 mM) for

3 h. Cell harvest and protein purification steps are described above.

Differential Scanning Calorimetry

All DSC samples (0.3 mM) were dialyzed against saline phosphate buffer (40 mM

Na2HPO4, 100 mM NaCl, pH 7.4) or phosphate buffer (40 mM Na2HPO4, pH 7.4) with

Spectra/Por® Dialysis Membrane MWCO: 10,000 (Spectrum Laboratories, Inc).

Resulting dialysis buffers were used as reference cell solvents for precision and repeatability. Prior to analyses, all sample and reference solutions were rigorously degassed with Microcal Thermovac2 (GE Healthcare). All DSC data acquisition were obtained on MicroCal VP-DSC (GE Healthcare) equipped with twin cells and operated on differential mode at a rate of 1.0 ºC min-1 from 15 ºC to 90 or 110 ºC. All data were processed with Origin 7 (Origin Labs) and fit according to a Two-state model or non- two-state model with respect to individual data sets.30

Nuclear Magnetic Resonance Spectroscopy.

[15N-1H] Heteroquantum Single Quantum Coherence (HSQC) spectra were recorded at

The Ohio State University Campus Chemical Instrument Center. Samples (0.45 mM) were exchanged to phosphate buffer (40 mM Na2HPO4, 100 mM NaCl, pH 7.4) in 10%

D2O by use of an Illustra MicroSpin™ G-25 Column (GE Healthcare). A standard water suppression [15N-1H] HSQC pulse sequence was used for data collection,31 followed by apodization, zero-filling, Fourier transformation and phase correction. Detailed acquisition and processing parameters are listed in Table 1.

Table 1: 15N-1H HSQC acquisition and processing parameters.

Acquisition Parameters F2 F1 Instrument Bruker Avance DRX 800 MHz Acquisition software Bruker XWINNMR v3.5 Probe 5mm CPTXI 1H-13C/15N Z-GRD Nucleus 1H 15N Pulse Sequence fhsqcf3gpph Size of FID 1024 128 Number of transients 64 Dummy Scans 16 TD0 1

Processing Parameters F2 F1 Processing program Bruker TopSpin 3.2 Size of real spectrum 1024 256 Spectrometer frequency 800.1300000 81.0764690 Spectral resolution (Hz) 5.868765 11.719922 Apodization(MHz) Gaussian Squared cosine Line Broadening (Hz) -3.00 0.30 bell 45

Circular Dichroism Spectroscopy

All circular dichroism (CD) samples (10 μM) were dialyzed to phosphate buffer (40 mM

Na2HPO4, pH 7.4) with Spectra/Por® Dialysis Membrane MWCO: 10,000 (Spectrum

Laboratories, Inc). Resulting dialysis buffers were used as reference cell solvents for precision and repeatability. Prior to analyses, all sample and reference solutions were rigorously degassed with Microcal Thermovac2 (GE Healthcare). All CD data acquisitions were obtained on Jasco J-815 CD Spectrometer (JASCO) equipped with

Quartz cells with 0.1cm path length. Secondary structure studies were collected at 8 averaging scans and 50 nm/min scan rate and monitored at 222 nm. The data were fit by

K2D3 program (European Molecular Biology Laboratory).32 Variable temperature studies were performed at a rate of 1.0 ºC min-1 from 20 ºC to 95 ºC. All data were processed with Origin 7 (Origin Labs). N-NFU and C-NFU VT data were fit to equation

1 while NC-NFU and full-length NFU VT data were fit to equation 2 for Tm and ΔHv determination:

Equation1

where R is the ideal gas constant in cal/mol, Tm is the melting temperature, ΔHv is the van’t Hoff enthalpy, Cp is the heat capacity, F and U are the mean residue ellipticity (θmr) of the folded and unfolded protein, respectively, 46

Equation 2

where the subscripts of Tm, ΔHv and Cp denote the transition parameters associated with the C-terminal and N-terminal domain, respectively.

Results

DSC Studies of N-Terminal Human NFU.

N-NFU DSC analyses were found to be in agreement with our previous NMR and CD studies27 inasmuch as N-NFU displays a rigid, well-behaved native structure at ambient temperature (Figure 7).

C) o 15

5 Cp (kcal/mole/ Cp 0

-5 10 20 30 40 50 60 70 80 90 100 110 120 Temperature (oC)

Figure 7: Differential scanning calorimetry profile for a 0.3 mM N-NFU solution in phosphate buffer. Origin was used to fit the data to a 2-peak, non-2-state model (MN2State). Values for Tm, ΔHcal and ΔvH were obtained from the fit and are listed in Table 2 and Table 3. The lower melting point was initialized at 79.4 ºC, which was obtained from a VTCD analysis.

The melting curve was analyzed and fit to a two peak non-two state model, yielding physical constants for melting temperatures (Tm) of 74.7 ± 0.6 and 78.4 ± 0.1 ºC (Table

2). The changes in molar enthalpy (ΔHcal) and van’t Hoff enthalpy (ΔHυ) from the fitting parameters were determined to be 134.6 ± 9.9 kcal/mol, and 141.4 ± 4.2 kcal/mol for the lower transition and 93.2 ± 9.8 kcal/mol, and 79.2 ± 1.6 kcal/mol for the higher transition, respectively (Table 3). The high Tm indicates the unfolding transitions for N-

NFU to stem from a well-folded state, while the reproducibility of melting for a sample following repetition of the heating cycle indicates that the melting process is reversible, but with partial protein degradation arising during each cycle (Figure 8). This evidence suggests that at high temperature the N-NFU domain exists in a stable conformation, and is able to fold back to the native state upon cooling.

Table 2: Melting temperatures for N-terminal NFU, C-terminal NFU, a mixture of N- and C-terminal domains, and full-length NFU determined by DSC.For a well-folded N- terminal domain, the change in melting temperatures for secondary and tertiary structures ter sec Δ(Tm -Tm ) is inversely related to the ionic strength. The majority of this observation is sec contributed by the decrease in Tm at lower ionic strength.

[NaCl] (mM) Tm1 (ºC) Tm2 (ºC) Tm2 -Tm1 (ºC) Tm3 (ºC) Tm4 (ºC) Tm4 -Tm3 (ºC)

100.0 74.7 ± 0.6 78.4 ± 0.1 3.7 - - - N-NFU 0.0 75.0 ± 0.6 80.0 ± 0.1 5.1 - - -

100.0 57-80 57-80 - - - - C-NFU 0.0 49-81 49-81 - - - -

N-NFU / 100.0 49.3 ± 0.3 58.1 ± 0.4 8.8 78.0 ± 1.5 80.6 ± 0.2 2.6 C-NFU 0.0 28.7 ± 0.7 41.4 ± 0.7 12.7 76.6 ± 0.5 80.9 ± 0.1 4.3

100.0 63.3 ± 3.4 67.0 ± 0.3 3.7 75.6 ± 7.9 77.1 ± 3.7 1.5 NFU 0.0 51.3 ± 0.6 58.1 ± 2.6 6.7 69.4 ± 4.9 76.1 ± 1.2 6.7

Table 3: ΔHcal and ΔHv for N-terminal NFU, C-terminal NFU, a mixture of N- and C- terminal domains, and full-length NFU determined by DSC.By comparison with melting temperatures in Table 2, the enthalpic contribution from loss of secondary structure is listed in bold. In the cases of the N-NFU / C-NFU mixture and full-length NFU protein, two sets of enthalpies are observed due to the melting steps for the two independent domains.

[NaCl] ΔH ΔH ΔH ΔH ΔH ΔH ΔH ΔH cal 1 v1 cal 2 v2 cal 3 v3 cal 4 v4 (mM) (kcal mol-1) (kcal mol-1) (kcal mol-1) (kcal mol-1) (kcal mol-1) (kcal mol-1) (kcal mol-1) (kcal mol-1) 100 134.6 ± 9.9 141.4 ± 4.2 93.2 ± 9.8 79.2 ± 1.6 - - - - N-NFU 0 59.1 ± 3.6 52.8 ± 1.4 66.2 ± 3.4 70.1 ± 4.1 - - - - 100 ------C-NFU 0 ------

N-NFU / 100 42.0 ± 0.4 71.8 ± 5.8 40.9 ± 0.4 60.4 ± 5.3 68.2 ± 1.5 71.9 ± 4.9 289.4 ± 1.5 128.4 ± 38.4 C-NFU 0 28.3 ± 1.2 48.9 ± 2.9 38.2 ± 1.2 42.9 ± 3.4 28.2 ± 0.5 28.3 ± 2.6 194.4 ± 0.6 105.6 ± 2.5 100 94.0 ± 7.1 41.8 ± 8.2 20.1 ± 7.6 36.9 ± 3.3 125.7 ± 7.9 69.0 ± 45.1 37.9 ± 13.3 40.4 ± 2.7 NFU 0 29.6 ± 9.3 32.9 ± 11.6 57.3 ± 8.9 49.1 ± 29.9 28.3 ± 4.9 32.2 ± 15.4 102.8 ± 4.4 67.9 ± 1.9

C) o

5 Cp (kcal/mole/ Cp 0

10 20 30 40 50 60 70 80 90 100 110 120 Temperature (oC)

Figure 8: Rescan of the N-NFU domain indicates reversible unfolding with 66% and 79% retention in ΔHcal and ΔHv, respectively. The fitting program used corresponded to MN2State, and the fitted results yielded Tm1 = 75.3 ºC, Tm2 = 77.6 ºC, ΔHcal 1 = 92.4 kcal/mol, ΔHcal 2 = 58.9 kcal/mol, ΔHv1 = 94.4 kcal/mol, and ΔHv2 = 80.8 kcal/mol.

VTCD Studies of N-Terminal Human NFU.

The secondary structure of N-NFU was analyzed by CD spectroscopy and yielded a composition of 31% α-helix, 21% β-sheet and 48% random coil (Table 4). The K2D3 program32 was used to obtain a more accurate prediction of secondary structure composition than was previously possible.13 Variable temperature studies yielded a structural transition at Tm = 79.5 ± 0.4 ºC with ΔHυ = 69.6 ± 6.1 kcal/mol (Figure 9, Table

5 and Table 6), which is consistent with DSC results for secondary structure loss at 3.7 ºC higher than the tertiary structure transition (Table 2).

-15

-16

-17

-18

-19 Ellipticity -20

-21

-22

290 300 310 320 330 340 350 360 370 Temp. (K)

Figure 9: VTCD analysis of 10 μM N-NFU in 40 mM phosphate 100 mM NaCl sec buffer.Data were fit to equation 1 to yield values of ΔHυ, Tm , and ΔCp of 69.56 kcal/mol, 79.44 ºC and 2.02 kcal/mol.K, respectively. Ellipticity data was directly used without converting to molar ellipticity units because the van’t Hoff enthalpies are sec independent of such a factor. Tm was used as an initial value for Tm in the DSC data fitting routine.

Table 4: Circular dichroism analyses of N-terminal NFU, C-terminal NFU, a mixture of N- and C-terminal domains, and full-length NFU.Data were obtained at 25 ºC with a path length of 0.1 cm and fit with the K2D3 program,32 which yields a higher accuracy in β- strand prediction compared with its predecessor K2D.13 Theoretical values for the N- NFU/C-NFU mixture were calculated from the average values of the two isolated domains.

N-NFU C-NFU N-NFU / C-NFU NFU

Experimental Theoretical

α-helix 31% 30% 32% 30 % 32%

β-sheet 21% 16% 18% 18% 20%

Random Coil 48% 54% 50% 52% 48%

DSC Studies of C-Terminal Human NFU

DSC analyses of the C-terminal domain of human NFU were consistent with a molten globule-like native structure27 as the melting curve was observed to be broad and asymmetric over a span of 43 ºC (Figure 10, top). Four transitions were calculated from the melting curve using the non-two-state model, with Tm’s ranging from 55.0 ºC to 72.9

ºC. The first transition curve was observed to be broad, with ΔHcal > ΔHυ in agreement with a molten globule-like native state structure. As the temperature increased, the calculated melting curves became sharper, resulting in ΔHcal < ΔHυ for the transition at

72.9 ºC. In conjunction with a negative peak observed at 74 ºC, the transitions at 70.0 ºC

53 and 72.9 ºC suggest protein aggregation at elevated temperature. As expected, due to the structural instability of C-NFU, the melting and refolding cycle was irreversible as opposed to its N-NFU counterpart, and denatured precipitate was observed after a single heating ramp.

VTCD Studies of C-Terminal Human NFU

The thermal melting profile of the C-NFU secondary structure was monitored over the entire scanning range, from 20 ºC to 95 ºC (Figure 10, bottom, and Table 5 and Table 6).

The overall CD signal change was determined to be 5.9 ± 0.2 mdeg. This is consistent with the DSC results that supported the low overall thermal stability of the C-NFU structure, which denatures over a broad range between 50 ºC ~ 80 ºC. Furthermore, the constant decrease of secondary structure (222 nm) over the entire heat ramp indicates multiple intermediates during melting, which is also consistent with DSC results using a four-state fitting model to define the structural loss. A single transition fit of the C-NFU melt resulted in a relatively low ΔHv (13.6 ± 3.2 kcal/mol), further indicating the absence of a distinct native-state secondary structure.

C) o 5

-5 Cp (kcal/mole/ Cp -10

-15 10 20 30 40 50 60 70 80 90 100 110 120 o Temperature ( C)

-11

-12

-13

-14

Ellipticity -15

-16

-17

290 300 310 320 330 340 350 360 370 Temp. (K)

Figure 10: DSC and VTCD analysis of C-NFU in 40 mM phosphate 100 mM NaCl buffer. (Top) Upscan of 0.3 mM C-NFU from 10 °C to 90 °C with rate of 1 K/min recorded by DSC. (Bottom) Upscan of 10 μM C-NFU with the same conditions monitored at 222 nm by CD. VTCD data were fit to equation 1 and results are listed in Table 5 and Table 6. Fitting results from DSC experiments are listed in Table 2 and Table 3.

Table 5: Melting temperatures of N-terminal NFU, C-terminal NFU, a mixture of N- and C-terminal domains, and full-length NFU determined by VTCD.The melting points were determined from molar ellipticity data fit with equations 1 or 2. As a result of the broad transitions observed for the C-NFU domain, Tm’s were estimated according to a single transition and indicated with asterisk marks. In rows containing data collected on a mixture of N-NFU / C-NFU and the full-length NFU, the Tm1 and the Tm2 parameters refer to melting temperatures for the C-terminal and N-terminal domains, respectively.

[NaCl] (mM) Tm1 (ºC) Tm2 (ºC) 100.0 79.5 ± 0.4 N-NFU 0.0 75.7 ± 0.1 100.0 59.4 ± 3.8 C-NFU 0.0 60.7 ± 0.5 100.0 59.2 ± 0.5 78.6 ± 0.2 N-NFU / C-NFU 0.0 41.9 ± 1.5 77.2 ± 0.3 100.0 67.8 ± 0.6 77.8 ± 0.9 NFU 0.0 50.2 ± 1.2 71.0 ± 0.4

Table 6: Fitting parameters for the change in Van’t Hoff enthalpy determined by VTCD.Increasing ionic strength resulted in an increase in the Van’t Hoff enthalpy, indicating a relatively more stable secondary structure for well-folded domains, such as N-NFU, and consistent with the trends observed in the melting temperatures defined in Table 5.

-1 -1 [NaCl] (mM) ΔHv1 (kcal mol ) ΔHv2 (kcal mol ) 100.0 69.6 ± 6.1 N-NFU 0.0 41.8 ± 4.5 100.0 13.6 ± 3.2 C-NFU 0.0 17.7 ± 0.9 100.0 43.3 ± 4.1 69.2 ± 1.1 N-NFU / C-NFU 0.0 44.1 ± 4.5 45.8 ± 0.7 100.0 23.9 ± 1.7 41.0 ± 2.7 NFU 0.0 31.8 ± 6.3 28.3 ± 3.3

DSC and VTCD Studies of Full Length Human NFU.

Two distinct secondary transitions were observed in VTCD profiles of NFU (Figure 11, bottom, and Table 5 and Table 6). The transition at 77.8 ± 0.9 ºC is tentatively assigned to the N-terminal domain due to the similarity in Tm in comparison to the N-NFU domain alone. The lower transition at 67.8 ± 0.8 ºC is likely due to the unfolding of the C- terminal domain, since it corresponds to the lowest thermally stable domain for NFU.

The van’t Hoff energies associated with the N-terminal and C-terminal domains were determined to be 41.0 ± 2.7 and 23.9 ± 1.7 kcal/mol, respectively. The distinct transition observed for the C-terminal domain at its Tm, together with the increased melting ΔHv relative to C-NFU alone, suggests that the secondary structure of this domain is stabilized in NFU.

C) o

5 Cp (kcal/mole/ Cp 0

10 20 30 40 50 60 70 80 90 100 110 120 Temperature (oC)

-14

-16

-18

-20

-22 Ellipticity -24

-26

-28 290 300 310 320 330 340 350 360 370 380 Temp. (K)

Figure 11: DSC and VTCD analysis of NFU in 40 mM phosphate 100 mM NaCl buffer.(Top) Upscan of 0.3 mM NFU from 10 °C to 110 °C at a rate of 1 K/min recorded by DSC. (Bottom) Upscan of 10 μM NFU under the same conditions, but monitored at 222 nm by CD. VTCD data were fit to equation 2 and results are listed in Table 5 and Table 6. Fitting results from DSC experiments are listed in Table 2 and Table 3.

The thermal melting curve of NFU (Figure 11, top) indicates a convoluted transition with

Tm near 72 ºC. Using the secondary structure unfolding ΔHv determined by the VTCD, the DSC plot was fit to four peaks that correspond to the loss of secondary and tertiary structure of the N- and C-domains of the NFU. In comparison with the DSC melting curves from the N-terminal domain (Figure 7) and the C-terminal domain (Figure 10, top), two of four peaks of the NFU melting curve correspond to the individual domains of the full length protein; the higher Tm of 77.1 ± 3.7 ºC corresponds to melting of the N-

NFU domain whereas the lower Tm of 67.0 ± 0.3 ºC corresponds to unfolding of the C-

NFU domain. The relatively modest difference between ΔHcal and ΔHυH for both transitions is characteristic of a two-state unfolding processes, suggesting the absence of melting intermediates or protein aggregation of the individual domains in the full length protein. Furthermore, the similarity of the two enthalpies in NFU compared with the individual domains, along with the reversibility of melting (Figure 8), suggests that in forming the full length protein, both domains have been structurally stabilized. These changes in thermodynamic and structural properties are more significant for the C-NFU domain, as forming the complete protein apparently alters its native state and results in a relatively more structurally-ordered macromolecule. The C-NFU domain in NFU is thermally well-behaved as the fitting yielded an overall melting ΔHcal of 114.1 ± 10.8 kcal/mol (Table 3). We speculate that in the full length protein, the C-terminal domain is stabilized by the N-terminal domain and therefore exhibits significant tertiary structure in comparison to the isolated domain.

DSC Studies of a Mixture of N- and C-Terminal Domains of NFU

The DSC results described thus far demonstrate unusual, but interesting interactions between the N- and C-terminal domains of NFU. To further elucidate the chemistry between the two segments we carried out DSC studies on solution mixtures of equimolar

N-NFU and C-NFU (Figure 12, top), which yielded three noteworthy observations. First, the transitions at Tm’s of 49.3 ± 0.3 and 58.1± 0.4 ºC correspond to unfolding of C-NFU in the solution mixture. The overall structural behavior of C-NFU in the presence of N-

NFU was observed to be more stable, with a symmetric melting profile and ΔHcal ≈ ΔHυ.

Furthermore, the secondary and tertiary structural Tm’s observed for C-NFU in solution with N-NFU are 8.93 ± 0.5 and 14.04 ± 3.4 ºC lower than that of the C-NFU domain of the full-length NFU protein, respectively, indicating that the degree of structural stabilization by N-NFU is stronger when the two domains are covalently attached. The interaction surface between the two domains is likely to be dominated by hydrophobic interactions, since decreasing buffer ionic strength was observed to weaken the degree of thermal stabilization of C-NFU. This is consistent with the results of isothermal titration calorimetry studies where no significant enthalpic response was observed by titrating the two domains.13 Rather, any binding would have to be promoted by entropy-driven interactions. Lastly, interaction between the two domains increases the overall structural stability of the N-terminal domain, which can be observed from the increased Tm and an overall increase of 129.7 ± 14.1 kcal/mol in molar enthalpy.

45 40 35

30 C)

o 25 20 15 10

5 Cp (kcal/mole/ Cp 0 -5 -10 10 20 30 40 50 60 70 80 90 100 110 Temperature (oC)

-10

-11

-12

-13

-14 Ellipticity -15

-16

-17 290 300 310 320 330 340 350 360 370 Temp. (K)

Figure 12: DSC and VTCD analysis of a N-NFU / C-NFU mixture in 40 mM phosphate 100 mM NaCl buffer.(Top) Upscan of a mixture of 0.3 mM N-NFU and C-NFU from 10 °C to 90 °C at rate of 1 K/min recorded by DSC. (Bottom) Upscan of a 10 μM mixture of 0.3 mM N-NFU and C-NFU under the same conditions, but monitored at 222 nm by CD.

VTCD Studies of a Mixture of N- and C-Terminal Domains of NFU

Similar to NFU, the VTCD melting curve of a mixture of N-NFU and C-NFU showed two distinct transitions (Figure 12, bottom, and Table 5). The lower transition was observed at 59.2 ± 0.5 ºC and corresponds to the C-NFU secondary structure melt. In comparison to the VTCD profile for C-NFU alone, the thermal transition was observed to be well behaved with ΔHv = 43.3 ± 4.1 kcal/mol in the presence of N-NFU (Table 6), which is more than three-fold larger than the C-NFU-only VTCD melting curve. The thermodynamic parameters evaluated for C-NFU are in agreement with its DSC profile and estimated melting enthalpies, and support the conformational change and structural stabilization of C-NFU following binding to N-NFU.

[15N-1H] HSQC Studies of a Mixture of N- and C-Terminal Domains of NFU

To further study the interaction between N-NFU and C-NFU, each of the two domains was prepared in an 15N-isotopically-enriched form. First, comparing the spectra from the isolated 15N-N-NFU domain, and either an equimolar mixture of 15N-N-NFU and 14N-

CNFU, or the 15N-labeled full length protein (Figure 13) revealed no shifted cross-peaks and no significant change in the conformation of the N-terminal domain. That is the structure appears conserved for both the isolated N-NFU, in complex with C-NFU, and as a separate domain in the full-length NFU protein. By contrast, experiments conducted with the C-terminal C-NFU domain under the same solution conditions as the DSC analyses demonstrated very distinct behavior. When equimolar mixtures of 14N-N-NFU and 15N-C-NFU were mixed and studied by [15N-1H] HSQC NMR experiments, a

62 comparison of the spectrum relative to that obtained for the isolated 15N-labeled C-NFU domain revealed 21 new 15N-1H cross-peaks (Figure 14 A and B, Table 7).

Figure 13: 1H-15N HSQC spectra of (A) 15N-N-NFU, (B) [15N-N-NFU + 14N-C-NFU] and (C) 15N-NFU.The N-terminal domain tertiary structure is not dependent on the presence of the C-terminal domain, and the N-terminal domain cross-peaks are observed to be unchanged upon addition of the C-terminal domain.

Table 7: A list new cross-peaks observed in the HSQC spectrum of the 15N-labeled C- NFU domain following complex formation with unlabeled N-NFU.No changes were observed in the observed cross-peaks for the complementary experiment with 15N-labeled N-NFU domain.

1H (ppm) 15N (ppm) 9.08 103.98 8.64 106.03 7.68 106.14 7.13 105.96 7.43 108.08 7.59 108.90 7.74 110.68 8.57 110.47 8.47 112.72 8.88 113.61 8.69 114.02 6.67 114.43 9.25 115.59 8.81 116.96 7.05 118.46 7.54 120.03 8.73 119.55 9.15 119.83 8.85 123.88 8.76 126.93 9.96 130.27

Figure 14: 1H-15N HSQC spectra of (A) 15N-C-NFU, (B) [14N-N-NFU + 15N-C-NFU] and (C) 15N-NFU.The C-terminal domain undergoes structural change in the presence of the N-terminal domain, as reflected by the peaks circled in red in (B). Cross peaks are also observed in spectrum (B) from the non-enriched N-NFU domain as a result of natural abundance 15N and provide an effective comparison of spectra for full-length NFU (C) relative to the sum of the individual domains (B). The cross-peaks circled in green in spectrum (C) highlight the chemical shifts of the full-length protein that are unique from the individual domains and in some cases overlaps with the peaks from the structure stabilized C-terminal domain. Chemical shifts for new cross-peaks identified in (B) and (C) are listed in Table 7 and Table 8, respectively.

Table 8: A list of new cross-peaks observed in the HSQC spectrum of full-length NFU relative to the spectra for the individual N- and C-NFU domains.No changes were observed in the observed cross-peaks for the N-NFU domain, and all changes derived from the C-NFU domain. Values in bold correspond to peaks observed to co-localize with cross-peaks from a solution mixture of 14N-N-NFU / 15N-C-NFU.

1H (ppm) 15N (ppm) 7.35 101.66 7.44 101.60 7.56 101.80 7.75 105.35 8.02 106.10 7.79 106.44 6.54 106.58 7.70 109.52 6.46 110.27 6.86 111.09 7.46 113.27 8.97 115.25 9.27 119.76 7.18 121.33 8.97 127.20 9.09 127.34 9.07 128.15 8.89 128.56 9.94 128.97 7.92 129.11 7.82 129.04 7.74 130.00 7.88 130.89

These peaks suggest an alternative tertiary conformation for C-NFU following interaction with the N-NFU domain. Furthermore, the observed new peaks exhibit distinct chemical shifts that span the chemical shift range from 6.67 to 9.96 ppm in the 1H domain, and from 103.98 to 130.27 ppm in the 15N domain, supporting a conformational change in C-

NFU with a greater level of tertiary structure and corresponding diversity in the chemical environment of its residues. When the spectrum for the 15N-labeled C-NFU domain was compared to that for the 15N-labeled full-length NFU (Figure 14 A and C) a total of 23 new cross-peaks were observed (Table 8). Significantly, only 4 of those new cross-peaks found for C-NFU in the full length NFU protein were co-localized with cross-peaks observed in the spectrum for the mixture of the 14N-N-NFU and 15N-C-NFU domains

(Table 8, bold). Apparently the structural transition induced by complex formation between the isolated N-NFU and C-NFU domains is incomplete, relative to the two domains in the full-length NFU protein.

Discussion

Comparison of the two isolated domains of the full-length NFU protein has shown the N- terminal fragment to display higher thermal stability relative to its C-NFU counterpart

(Figure 7and Figure 10, top, respectively). This is consistent with our previous studies demonstrating the N-terminus of NFU to possess a more rigid conformation than the C- terminal NFU.27 The C-terminal domain exhibits molten globule characteristics with lower thermostability (Figure 10, bottom) and does not display a well-folded tertiary structure according to both CD and NMR criteria.13, 26, 27 DSC studies of C-NFU reported herein displayed structural heterogeneity during the unfolding process (Figure

10, top), while the VTCD melt showed a consistent decrease in secondary structure throughout the temperature domain (Figure 10, bottom). The ability of N-NFU to promote conformational change and stabilize the structure of C-NFU was confirmed by both DSC and CD melting experiments (Figure 12) and 2D-NMR experiments (Figure

14). The degree of stabilization was observed to be greater for full-length NFU, relative to a mixture of the two domains (Figure 14 B vs. C). The secondary structures of the N-

NFU/C-NFU mixture contain 2% less random coil than the expected composition, which is consistent with the sharp CD melting curve due to the new C-NFU conformation

(Figure 12, bottom). Although N-NFU somewhat stabilizes the secondary and tertiary structure of C-NFU, there remains a significant amount of molten globular C-NFU within the mixture of the two proteins, with 50% random coil present in the secondary structure

(Table 4). Increasing ionic strength is known to stabilize the thermostability of molten globules.33 Accordingly, the difference in the degree of structural stabilization between

68 these two cases is also reflected in the results of DSC studies using a lower ionic strength buffer (0 mM NaCl), where a decrease of 18.6 ºC was observed in the Tm for C-NFU in the mixture of proteins, but only by 10.5 ºC in NFU (Table 2). Furthermore, by comparing the [15N-1H] HSQC NMR spectrum of full-length NFU relative to that from a mixture of the N-NFU and C-NFU domains, 23 new cross peaks were observed (Figure

14C and Figure 15), relative to the 21 new peaks observed for C-NFU following N-NFU binding in the co-complex of the two domains (Figure 14B and Figure 16), although only

4 of these new cross-peaks from each spectrum are found to co-localize (Table 7 and

Table 8). These residues underline the difference between the N/C-domain protein- protein interaction and covalently attached NFU. Nevertheless, the ability of N-NFU to structurally stabilize C-NFU was demonstrated in both cases through the appearance of new cross peaks in the NMR experiments (Figure 14) and is consistent with the progressive changes observed in the DSC plots for each isolated domain, relative to the mixture of domains and full-length protein (Figure 7, Figure 9 to Figure 12).

Figure 15: [15N-1H] HSQC spectra indicate an alternative conformation for C-NFU following the interaction with N-NFU.Overlap of the spectrum obtained from a 14N-N- NFU / 15N-C-NFU mixture (red) and NFU (black). At least 4 new cross peaks found in Figure 16 were observed to co-localize with the full-length protein and listed in Table 8. Black-only cross peaks indicate an alternate conformation for C-terminal NFU, in a solution with N-terminal NFU, relative to the C-terminal domain in the full-length protein.

Figure 16: [15N-1H] HSQC NMR spectrum of an equimolar mixture of 14N-N-NFU and 15N-C-NFU(red) in comparison to the sum of the two domains measured separately (black). Chemical shifts of novel cross-peaks are listed in Table 7. Red-only cross peaks indicate an alternate C-terminal conformation.

The Gibbs free-energy for the secondary structural transition was converted from CD melt data by using equation 3 and drawn to illustrate the unfolding profile for N-NFU

(Figure 17).

Equation 3

-3

-2

-1

G(kcal/mol)  2

3 275 300 325 350 375 T =348.88 K Temp. (K) m

Figure 17: Thermal profile for N-NFU.Data (dots) were converted to ΔG values by use of sec equation 3, and the Gibbs-Helmholtz equation, along with ΔHυ, Tm , and ΔCp, was used to form a plot of ΔG versus temperature (line). As expected, N-NFU is most thermally stable near human body temperature while melting of the protein is a favorable process at temperatures exceeding 79.4 ºC.

According to the plot, as temperature increases, the ΔG response is first observed to increase as the melting process is enthalpy driven. The free energy ΔG reaches the highest value at ~ 37 °C, since N-NFU is most likely thermostable at physiological 72 temperatures. The unfolding free energy decreases from 37 to 78 °C, because the unfolding process is entropy driven. Figure 18 shows the thermal profile of a mixture of

N- and C-NFU, and NFU, to be similar to the N-NFU thermal profile, where all thermal transitions were fit to yield a ΔGmax near 37 ºC.

-5 -4 -3 -2 -1 0 1

G(kcal/mol) 2  3 4 5 275 300 325 350 375 Temp. (K)

Figure 18: Gibbs free energy plot illustrates a mixture of N-NFU and C-NFU domains to be most thermally stable near physiological temperature.The solid line represents the calculated data for N-NFU, and the dashed line the calculated data for C-NFU. Squares show the experimental data for N-NFU, and triangles show the experimental data for C- NFU. The vertical dashed lines indicate the fitted Tm’s of C-NFU (315.0 K) and N- NFU (350.3 K).

The influence of N-NFU on C-NFU underlines the importance of the structural role of the

N-terminal domain on the native structure of NFU. In agreement with our previous studies, the N-terminal domain appears to maintain the overall structure of the full-length

73 protein.13, 26, 27 We conclude that hydrophobic interactions between the N- and C- terminal domains facilitate the folding of each, illustrated by the observation that the unfolding of C-NFU changes from a molten globule state to a relatively more thermally stable conformation. However, the C-terminal domain in the full-length protein is not fully structured as observed by NMR experiments.27 This is also supported by the increased thermostability of the C-terminal domain under high salt conditions. Structural disorder is generally considered to be a factor that decreases enzyme catalytic efficiency.

However, recent reports have shown molten globular enzymes adopting functional conformations upon binding to relevant binding partners or ligands, such as the case of chorismate mutase.34 Similarly, the molten globular C-terminal domain of NFU may adopt its functional conformation upon binding to sulfur donors and subsequently aid in the 2Fe2S cluster assembly with scaffolding proteins. As demonstrated by both ITC and kinetic studies, human NFU is known to bind to IscS in the process of reconstitution of human ISU scaffold protein.13 In order for the conserved CXXC thiolate groups to reduce the persulfide bond on IscS and subsequent relocation of sulfide ions into the ISU active site, it is likely that the C-NFU adopts a distinct and more structured conformation.

The absence of a thermally stable tertiary conformation for the C-terminal CXXC domain may provide the required structural flexibility to allow such a reaction. The decrease in flexibility of C-NFU in the presence of N-NFU was also observed in the secondary structure predictions, with an increase in ~2% α-helix content (5 residues) upon binding.

A recent bioinformatic study reported that the majority of protein-protein interaction are due to helix interactions (62%) with involvement commonly between 4 to 14 residues,35

74 such as the bacterial Bernase-Barstar system.36 The Zimm-Bragg theory describes that the rate determining step in formation of α-helix is the formation of the first loop, which requires ~20 kcal. However, the subsequent helix growth is exothermic and a thermodynamically favored process. Therefore, the formation of additional alpha helix content between the two terminals of NFU is likely to be entropy driven and forms spontaneously, leading to an alternative secondary and tertiary conformation for the C- terminal domain.

Conclusion

The term molten globule is ascribed to a diverse category of protein structural conformations that are broadly defined by a high degree of secondary structure, but lacking a rigid tertiary structure.37, 38 There are many examples for alternative conformers with distinct recognition or functional roles within a given protein molecule.39-41 Calorimetry results and our previous investigations13, 27 suggest the C- terminal domain of human NFU to exist in a molten globule state in both the truncated form and within the full-length protein, but with different conformations and physical properties. The C-terminal domain appears to adopt an alternative conformation following interaction with the N-terminal domain. This change in conformation is accompanied by an increase in the tertiary structure, as well as an increase in thermostability. This property of structural flexibility most likely underlies the required properties and functions of the protein, as also observed in the cases of molten globular clusterin and nucleosomes.42 Interestingly, the iron-sulfur scaffold protein T. maritima

IscU appears to equilibrate between structural conformers, and may be essential for its interactions with various protein partners.20 Structural isomerism within this family of scaffold proteins has also been emphasized by recent work from the Markley group.21-25

Due to the complexity of iron-sulfur cluster assembly systems, and numerous contributions from scaffold, partner and chaperone proteins, we propose that a dynamic tertiary structure is an important factor for the proper functioning of several key protein involved with Fe-S cluster formation.

Additional Figures

6 C) o 4

Cp (kcal/mole/ Cp 0

-2 0 10 20 30 40 50 60 70 80 90 100 110 120 Temperature (oC)

-7

-8

-9

-10

Ellipticity -11

-12

-13 290 300 310 320 330 340 350 360 370 Temp. (K)

Figure 19: DSC and VTCD analysis of N-NFU in 40 mM phosphate buffer.(Top) Upscan of 0.3 mM N-NFU from 10 °C to 90 °C with rate of 1 K/min recorded by DSC. (Bottom) Upscan of 10 μM N-NFU under the same conditions monitored at 222 nm by CD. VTCD data were fit to equation 1 and the results are listed in Tables 5 and 6. Fitting results from DSC experiments are listed in Tables 2 and 3.

C) o

5 Cp (kcal/mole/ Cp 0

-5 10 20 30 40 50 60 70 80 90 100 110 Temperature (oC)

-7

-8

-9 Ellipticity

-10

-11 290 300 310 320 330 340 350 360 370 Temp. (K)

Figure 20: DSC and VTCD analysis of C-NFU in 40 mM phosphate buffer.(Top) Upscan of 0.3 mM C-NFU from 10 °C to 90 °C with rate of 1 K/min recorded by DSC. (Bottom) Upscan of 10 μM C-NFU under the same conditions monitored at 222 nm by CD. VTCD data were fit to equation 1 and the results are listed in Tables 5 and 6. Fitting results from DSC experiments are listed in Tables 2 and 3.

15 C) o 10

Cp (kcal/mole/ Cp 0

-5 10 20 30 40 50 60 70 80 90 100 110 120 o Temperature ( C)

-10

-12

-14

-16 Ellipticity

-18

-20 290 300 310 320 330 340 350 360 370 Temp. (K)

Figure 21: DSC and VTCD analysis of NFU in 40 mM phosphate buffer.(Top) Upscan of 0.3 mM NFU from 10 °C to 90 °C with rate of 1 K/min recorded by DSC. (Bottom) Upscan of 10 μM NFU under the same conditions monitored at 222 nm by CD. VTCD data were fit to equation 1 and the results are listed in Tables 5 and 6. Fitting results from DSC experiments are listed in Tables 2 and 3.

C) 15 o

5 Cp (kcal/mole/ Cp

0 10 20 30 40 50 60 70 80 90 100 110 120 Temperature (oC)

-12

-14

-16

-18 Ellipticity -20

-22

290 300 310 320 330 340 350 360 370 Temp. (K)

Figure 22: DSC and VTCD analysis of N+C-NFU in 40 mM phosphate buffer.(Top) Upscan of a mixture of 0.3 mM N-NFU and C-NFU from 10 °C to 90 °C with rate of 1 K/min recorded by DSC. (Bottom) Upscan of 10 μM a mixture of 0.3 mM N-NFU and C-NFU under the same conditions monitored at 222 nm by CD. VTCD data were fit to equation 2 and the results are listed in Tables 5 and 6. Fitting results from DSC experiments are listed in Tables 2 and 3.

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Chapter 3: Temperature Induced Conformation Study of Human Ferredoxin 2

Introduction

Ferredoxins usually is found to contain either [2Fe-2S] or [4Fe-4S] clusters and function to mediate electron transfers in various biochemical pathways.1-3 Human ferredoxin 1

(hFd1) is involved in adrenal steroidogenesis, bile acid formation, and vitamin D synthesis. 4-6 In 2010, a second ferredoxin in human (hFd2) was identified. Despite the high similarity (43% identity and 69% similarity in protein sequence), it was recently reported that hFd1 and hFd2 play very specific roles on distinct pathways.7 The function of this second ferredoxin is found to be involved in Fe-S cluster and heme A biogenesis, similar to its yeast homologue.7 Protein structures for these two different forms of

Ferredoxin appear to be very similar. (Figure 23 and Figure 24) 8, 9 Furthermore, the electronic absorption profiles of the dark-red coloured proteins also appear to be similar.

(Figure 25 and Figure 26) In both cases, the strong absorption in the visible region is due to sulfur to iron charge transfer (LMCT) originated from the 2Fe-2S centers. The modest difference in the absorption spectra is likely due to the minute difference in coordination geometry of the iron-sulfur center.

Figure 23: Crystal structure of hFd1(PDB #: 3P1M) with [2Fe-2S] cluster shown in amber and yellow.

Figure 24: Crystal structure of hFd2(PDB#: 2Y5C) with [2Fe-2S] cluster shown in amber and yellow.

1.0 hFd1 hFd2 0.8

0.6

0.4 Absorbance 0.2

0.0 300 400 500 600 700 Wavelength (nm)

Figure 25: Comparison of UV/vis absorption spectra for hFd1 and hFd2.Both proteins are in their holo-state, and electronic transitions observed in this range are due to charge transfers between ligands and iron. (Note: spectra collected in collaboration with Wenbin Qi)

hFd1 hFd2

400

 200

0 Elipticity

-200

200 300 400 500 600 700 Wavelength (nm)

Figure 26: Comparison of CD spectra for hFd1 and hFd2. Both proteins are in their holo- state, and CD signals are due to charge transfers between ligands and iron, as well as the chiral environment of the cluster pocket. (Note: spectra collected in collaboration with Wenbin Qi)

Materials and Methods

Cloning and Expression of human ferredoxin2

Cloning of hFd2 into pET28-b(+) with N-terminal His-tag and transformation into E. coli BL21(DE3) was previously done by Wenbin Qi, and documented in his Doctoral

10 Thesis. For over-expression of the protein, cells were grown in LB at 37 C to an OD600 of ~ 0.6, and then induced with 100 mg/mL isopropyl-1-thio-α-D-galactopyranoside

(IPTG) for 3 hours at 37 C. Cells were harvested at 5,000 rpm for 10 min and stored as frozen pellets until purification.

Purification of proteins

Cell pellets were resuspended in 20 mM Tris, 500 mM NaCl, 5 mM imidazole, pH = 8.0 and lysed by sonication. Cell debris was removed by centrifugation at 4 C at 15,000 rpm for 30 min. The supernatant was applied through TALON (Cobalt Metal Affinity Column) column and eluted by 150 mM Imidazole in 50 mM Hepes, 100 mM NaCl, pH = 7.5. For hFd1, standard ion-exchange and size-exclusion column purification protocol established by Markley group was used.11 Eluted proteins were analyzed by SDS-PAGE.

hFd1 and hFd2 temperature dependent absorbance change monitored by UV/Vis spectroscopy

Temperature dependent absorbance experiments were performed by Wenbin Qi. In brief, a solution of protein (100 µL, 80 µM) heated at a rate of 1 C/minute from 25 C to 60 C

89 was recorded by Cary-50 UV/Vis Spectrometer (Varian). Absorbances at 414 nm were taken at 5 degree intervals (5 min intervals).

hFd2 temperature dependent CD signal change

A solution of hFd2 (80 uM) in hepes buffer (50 mM Hepes, 100 mM NaCl, pH 7.5) was heated from 25 C to 65 C at a rate of 1 C/min. CD spectra were taken at 10 C intervals using a JASCO J-815 Circular Dichroism (CD) Spectrometer. For far UV secondary structure analysis, data were obtained using Quartz cells with 0.1cm path length. Secondary structure studies were collected at 8 averaging scans and 50 nm/min scan rate. The data were fit by K2D3 program (European Molecular Biology Laboratory).12 A phosphate buffer (40 mM Na2HPO4, pH 7.4) was used for secondary structure determination.

Rate of ferredoxin reduction by adrenodoxin reductase and cluster transfers monitored by the cytochrome c assay

All solutions were purged by argon prior to use. For the measurement of enzymatic parameters, cytochrome c (80 M), adrenodoxin reductase (200 nM) and NADPH (400 M) were combined in 10 mM sodium phosphate buffer under anaerobic condition. Ferredoxin solutions at final concentrations of 0.1-8 M were added and the absorbance at 550 nm was monitored over 5 min to define the initial slope of the kinetic profiles. For measurement of the holo-ISU to apo ferredoxin cluster transfer rate, a solution of 20 L apo ferredoxin (200 M) was added to 20 L DTT (50 mM) and incubated at R.T. for 30 min. An aliquot of this solution (7 L) was subsequently added to 35 L of holo ISU 90

(200 M) and 6 L of this mixture was used in the cytochrome c assay. For the cytochrome c assay, the total reaction volume used was 100 L. This solution contained 80 M cytochrome c, 200 M adrenodoxin reductase, 400 M NADPH, and 10 mM sodium phosphate at pH 7.4. The absorbance was measured at 550 nm for the first minute and the resulting initial slope was used for subsequent calculations. For details see previously published work from this laboratory.16

Nuclear magnetic resonance spectroscopy

All NMR samples (0.45 mM) were exchanged to 90% phosphate buffer (40 mM

Na2HPO4, 100 mM NaCl, pH 7.4)/ 10% D2O with Illustra MicroSpin™ G-25 Column (GE Healthcare) and data were acquired in 5 mm NMR tubes (Wilmad-Labglass). [15N- 1H] Heteroquantum Single Quantum Coherence (HSQC) spectra for all samples were recorded on a Bruker DMX 600 MHz spectrometer (carrier frequency 600.13MHz) equipped with a 5 mm TXI (13C/15N) probe with x,y,z-axes gradients to a dimension of [1024 x 512] and processed with XWIN-NMR v1.1. Contour level was defined with a multiplier of 1.4 and a total of 8 positive layers. All NMR experiments were recorded at 301.1 K except for the variable temperature experiments, which were achieved with the spectrometer’s built-in calibrated temperature controller.

Results

Variable Temperature UV/Vis Study

Interestingly, an unusual transition to a higher extinction coefficient species was observed for hFd2 when solution was warmed to physiological temperature, and up to 55 C. This structural change is likely to influence the structure and function of hFd2. Similar change was not observed for hFd1, and the thermal stability of hFd2 was found to be greater than hFd1. Absorption at 414 nm (cluster LMCT) was observed to decrease for holo-hFd1 when temperature was increased above r.t., in contrast to holo-hFd2 displaying stability above physiological temperature (Figure 27).

0.2

0.1 hFd2

0.0

414 Abs

 -0.1 hFd1 -0.2

-0.3 20 30 40 50 60 70 Temperature (oC)

Figure 27: Absorbance change during heating cycle for hFd1 and hFd2.(Note: spectra collected in collaboration with Wenbin Qi)

Variable Temperature CD study

In contrast to the change in absorbance response, the CD signal from the cluster does not show a significant change when incubated up to 55 C (Figure 28), aside from slight degradation at 55 C after 30 min, suggesting that warming the sample, and the conformational change that subsequently results does not induce a significant change in the chiral environment of the cluster. The absorption data measured by UV/Vis and CD suggest the structural change of hFd2 invokes a change near the cluster coordination sphere, but the change does influence the chirality of the ligands coordinated to the iron sulfur cluster.

0 Ellipticity -2

-4

300 400 500 600

Wavelength (nm)

Figure 28: CD signal from the cluster center when holo hFd2 is incubated up to 55 C. To further investigate the origin of temperature dependent absorbance change, far UV

CD experiments was used to study the secondary structures of hFd2 at various temperatures. At each temperature, raw CD signals were converted to molar ellipticity and used as input data for K2D3 program for secondary structures prediction of hFd2.

Similar to the CD signals in the visible region, no significant change in secondary structure was observed during the conformational transition (Figure 29 and Table 9).

Figure 29: Far UV absorption spectra obtained at various temperatures: 35 ºC (black), 55 ºC (red), 75 ºC (blue) and 95 ºC (teal). Secondary structural analyses by K2D3 are summarized in Table 9.

Table 9: Temperature dependent secondary structural prediction and analysis by K2D3.

T (ºC) 25 35 45 55 65 75 85 95

α-helix (%) 17.5 16.4 19.2 17.3 18.2 17.4 9.7 9.7

β-strand (%) 21.2 22.6 21.6 22.6 22.3 22.3 26.8 26.9

DSC Studies

The temperature induced structural change of hFd2 was further studied by Differential

Scanning Calorimetry. hFd2 showed an increased heat intake between 35 C to 70 C. This transition agrees with temperature dependent absorption experiments done by Wenbin Qi

(Figure 27). In both studies, changes in protein behavior were clearly observed when solution temperature reached above 35 C. In contrast, studies with hFd1 showed poor stability of the holo cluster, with the loss of protein structure above 30 C. This was observed by both DSC and absorption methods. A decrease in absorbance at 414 nm for hFd1 is consistent with a loss of the iron-sulfur cluster. For the multi-state transition observed by DSC experiment, it is consistent with a loss of structure above 30 C, with protein aggregation near 80 C and resulted in a negative molar heat capacity.

4 C) o 3

1 Cp (kcal/mole/ Cp

-1 10 20 30 40 50 60 70 80 90 100 110 Temperature (oC)

C) o

0 Cp (kcal/mole/ Cp

-5 10 20 30 40 50 60 70 80 90 100 Temperature (oC)

Figure 30: Differential scanning calorimetry experiments of hFd2 (top) and hFd1 (bottom).

Cytochrome C Assays

To evaluate the functional implications for such a distinct conformational state, a standard cytochrome c assay13, 14 was used to determine the affinity of hFd1 and hFd2 to their physiological partner adrenodoxin reductase. Enzymatic parameters for hFd1 and hFd2, both at ambient temperature (25 C) as well as elevated temperature were determined and are listed in Table 10. At ambient temperature a relatively tighter binding to adrenodoxin reductase was observed for hFd1 with a Km of 2.1 μM, while hFd2 shows a Km value of 4.4 μM. Interestingly, at higher temperature the Km for hFd2 decreased to values closer to hFd1 (~1.8 μM), while the Km for hFd1 remained similar. Similar data are obtained for hFd2 for 37 and 55 C, supporting the view that the conformational transition has already occurred following incubation at physiological temperature.

Table 10: Enzymatic parameters for ferredoxin electron exchange with adrenodoxin reductase.

value ± σ hFd1 25 C hFd2 25 C hFd1 37 C hFd2 37 C hFd2 55 C

Km (μM) 2.1 ± 0.1 4.4 ± 1.5 2.0 ± 0.4 1.9 ± 0.3 1.8 ± 0.1

-1 kcat (s ) 0.30 ± 0.01 0.37 ± 0.06 0.21 ± 0.01 0.20 ± 0.02 0.20 ± 0.01

-1 -1 kcat/Km (μM s ) 0.14 ± 0.01 0.09 ± 0.02 0.10 ± 0.04 0.11 ± 0.01 0.11 ± 0.01

Vmax (μM/min) 1.8 ± 0.1 2.2 ± 0.3 1.2 ± 0.1 1.2 ± 0.1 1.2 ± 0.1

Table 11: Cluster transfer kobs by use of the cytochrome c assay.

-1 -1 -1 value ± σ kobs (min ) k2 (M min )

apo hFd2 (21.6 ± 3.9) x10-3 129 ± 23

apo hFd1 (9.3 ± 1.7) x10-3 56 ± 10

The reconstitution of apo ferredoxins by the Fe/S scaffold protein holo ISU was also monitored by use of the cytochrome c assay. Both hFd2 and hFd1 were capable of receiving the [2Fe-2S] cluster from the scaffold protein, although a faster rate was

-1 observed for hFd2 with observed reconstitution rate constants (kobs) of 21.6 min versus

-1 9.3 min (Table 11) for hFd2 and hFd1, respectively. The kobs determined for cluster transfer between AdR and hFd1 was consistent with our previously reported values.16

A distinct conformational change for hFd2 at elevated temperature, relative to hFd1, was also demonstrated more directly by following the change in chemical shift (Δδ) in the 2-

D [15N-1H] HSQC NMR spectra (Figure 31). Most of the cross peaks for hFd1 were observed to show a minor and uniform shift as a result of the rise of temperature, while the cross peaks observed for hFd2 were observed to move more dramatically and in different directions, in a manner consistent with a structural change. The weighted averages of the chemical shift change, Δδavg, were calculated from Eq.4 for hFd1 and hFd2 and plotted as shown in Figure 32. The overall average chemical shift change upon varying the temperature for hFd1 and hFd2 was calculated to be 0.017 ppm and 0.090 ppm, respectively.

(4)

It is important to note that along with the greater change in chemical shifts at elevated temperature, the [15N-1H] HSQC experiments for hFd2 also show the shifted signals to be spread over a wider range. These observations provide further support for a distinct change in tertiary conformation for hFd2 at higher temperatures, from physiological up to

55 C. A similar temperature dependent change of chemical shift was not observed for hFd1, as a convergence of the chemical shifts at higher temperatures indicated protein denaturation.

Figure 31: Overlay of variable temperature [15N-1H] HSQC spectra for hFd1 (A) and hFd2 (B) at 28 C (black) and 55 C (red).

100

Figure 32: Comparison of chemical shift perturbations for hFd1 and hFd2 after heating to 55 C. Chemical shift changes of amide proton and nitrogen signals, and Δδavg are shown for all comparable residues. The x-axes have been systematically numbered clockwise, as observed from the 2-D spectra, since the hFd2 residues have not been assigned.

101

Discussion and Conclusion

These results suggest that there is an alternative structural conformation for hFd2 that is stable at higher temperature which is not exhibited by hFd1. For hFd2, the native structure is likely to exist as the alternative conformation at higher temperatures (> r.t.), since the structural transition is observed at just over 30 C as observed by DSC and

UV/Vis experiments. Interestingly, the conformational transition for the reverse process was not observed for hFd2 upon cooling back to r.t. When a sample of hFd2 was heated to 60 C followed by cooling to 25 C, the absorbance at 414 nm was observed to increase during the warming process, and stayed unchanged during the cooling process (Figure

27). The higher absorbance conformation indicates that the high temperature conformation not an intermediate state formed during cluster degradation. Rather, a distinct and stable state reflected by a perturbed coordination environment around the

2Fe-2S cluster that results in an increase of cluster absorbance. In contrast, hFd1 displayed a decrease in absorbance at 414 nm upon heating to 60 C. Since the absorbance at this region is due to the sulfur to iron charge transfer of the iron-sulfur cluster, a decrease in absorbance is indicative for cluster loss. Cluster stability within holo-proteins may depend on the solution temperature, pH, ionic strength and the intrinsic dissociation properties of the prosthetic group. In this study, holo-hFd1 was found to be unstable at above physiological temperature. Loss of the 2Fe-2S center was also confirmed by the temperature dependent absorbance at 414 nm when it was cooled back to r.t. The absorbance was observed to be unchanged between 60 C and 25 C, indicating that the initial signal decrease during the warming step was due to loss of iron-sulfur cluster.

102

The tighter binding of hFd2 to adrenodoxin reductase under more physiological temperatures underlines the enhancement of protein-protein interactions arising from the structural change and is also consistent with the prediction that this second stable conformation is the actual functional conformation under physiological condition. Such unique thermally-induced changes in both structure and binding profiles were not observed for hFd1. Examples of heat-induced structural changes in proteins are known, including heat-shock proteins and other proteins such as the apo-α-lactalbumin,15-17 but such an effect has not previously been documented for any iron-sulfur cluster protein.

Accordingly, the ambient temperature structure determined for hFd2 may not fully represent the physiologically active state, and a greater difference in structure may contribute to the selection of either hFd1 or hFd2 for distinct cellular roles. It will also be of future interest to evaluate the prevalence and functional roles of such structural transitions across the broad family of Fe/S proteins.

NOTE: Figures 25, 26 and 27 were adapted from UV/Vis and CD data collected in collaboration with Wenbin Qi.

103

References

[1] Vickery, L. E. (1997) Molecular recognition and electron transfer in mitochondrial steroid hydroxylase systems, Steroids 62, 124-127.

[2] Grinberg, A. V., Hannemann, F., Schiffler, B., Muller, J., Heinemann, U., and Bernhardt, R. (2000) Adrenodoxin: structure, stability, and electron transfer properties, Proteins 40, 590-612.

[3] Miller, W. L. (2005) Minireview: regulation of steroidogenesis by electron transfer, Endocrinology 146, 2544-2550.

[4] Lange, H., Kaut, A., Kispal, G., and Lill, R. (2000) A mitochondrial ferredoxin is essential for biogenesis of cellular iron-sulfur proteins, Proc Natl Acad Sci U S A 97, 1050-1055.

[5] Rouault, T. A., and Tong, W. H. (2005) Iron-sulphur cluster biogenesis and mitochondrial iron homeostasis, Nat Rev Mol Cell Biol 6, 345-351.

[6] Barros, M. H., Nobrega, F. G., and Tzagoloff, A. (2002) Mitochondrial ferredoxin is required for heme A synthesis in Saccharomyces cerevisiae, J Biol Chem 277, 9997-10002.

[7] Sheftel, A. D., Stehling, O., Pierik, A. J., Elsasser, H. P., Muhlenhoff, U., Webert, H., Hobler, A., Hannemann, F., Bernhardt, R., and Lill, R. (2010) Humans possess two mitochondrial ferredoxins, Fdx1 and Fdx2, with distinct roles in steroidogenesis, heme, and Fe/S cluster biosynthesis, Proc Natl Acad Sci U S A 107, 11775-11780.

[8] Muller, A., Muller, J. J., Muller, Y. A., Uhlmann, H., Bernhardt, R., and Heinemann, U. (1998) New aspects of electron transfer revealed by the crystal structure of a truncated bovine adrenodoxin, Adx(4-108), Structure 6, 269-280.

[9] Chaikuad, A., Johansson, C., Krojer, T., Yue, W.W., Phillips, C., Bray, J.E., Pike, A.C.W., Muniz, J.R.C., Vollmar, M., Weigelt, J., Arrowsmith, C.H., Edwards, A.M., Bountra, C., Kavanagh, K., Oppermann, U. (2010) Crystal structure of human ferredoxin-1 (FDX1) in complex with iron-sulfur cluster, PDB ID: 3P1M.

[10] Qi, W. (2011) Studies of Iron-Sulfur Cluster Biogenesis and Trafficking, In Biochemistry Program, p 172, The Ohio State University, Columbus, Ohio.

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[11] Xia, B., Cheng, H., Bandarian, V., Reed, G. H., and Markley, J. L. (1996) Human Ferredoxin: Overproduction in Escherichia coli, Reconstitution in Vitro, and Spectroscopic Studies of Iron-Sulfur Cluster Ligand Cysteine-to-Serine Mutants, Biochemistry 35, 9488-9495.

[12] Louis-Jeune, C., Andrade-Navarro, M. A., and Perez-Iratxeta, C. (2011) Prediction of protein secondary structure from circular dichroism using theoretically derived spectra, Proteins 80, 374-381.

[13] Ziegler, G. A., Vonrhein, C., Hanukoglu, I., and Schulz, G. E. (1999) The structure of adrenodoxin reductase of mitochondrial P450 systems: electron transfer for steroid biosynthesis, J Mol Biol 289, 981-990.

[14] Wu, S. P., Wu, G., Surerus, K. K., and Cowan, J. A. (2002) Iron-sulfur cluster biosynthesis. Kinetic analysis of [2Fe-2S] cluster transfer from holo ISU to apo Fd: role of redox chemistry and a conserved aspartate, Biochemistry 41, 8876- 8885.

[15] Aprodu, I., Stanciuc, N., Banu, I., and Bahrim, G. (2012) Probing thermal behaviour of microbial transglutaminase with fluorescence and in silico methods, J Sci Food Agric.

[16] Lee, J. S., Satoh, T., Shinoda, H., Samejima, T., Wu, S. H., and Chiou, S. H. (1997) Effect of heat-induced structural perturbation of secondary and tertiary structures on the chaperone activity of alpha-crystallin, Biochem Biophys Res Commun 237, 277-282.

[17] Reiner, E., Davis, C. S., Schwab, B. W., Schopfer, L. M., and Richardson, R. J. (1987) Kinetics of heat inactivation of phenyl valerate hydrolases from hen and rat brain, Biochem Pharmacol 36, 3181-3185.

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Chapter 4: Synthesis, characterization, and studies on glutathione coordinated [2Fe-2S] clusters

Introduction

Glutathione (γ-glutamyl-cysteinyl-glycine, GSH) is an important cellular redox buffering agent as a result of its high cellular concentration (10 mM in mitochondria) and low reduction potential (-0.16 V vs. NHE).1-4 Glutathione molecules possess various essential cellular functions including redox homeostasis, cell signalling pathways, regulations of ion channel activity and protein folding.5-9 GSH serves as a backup to thioredoxin for maintenance of cytosolic reduction potential.10 Glutathione also protect cells towards reactive oxygen species and serve as a cofactor for glutathione peroxidase.11

Interestingly, GSH has been shown to participate in cytosolic iron-sulfur cluster biosynthesis12, 13 with a recent study suggesting an essential role in cellular iron-sulfur cluster assembly. Glutaredoxins are redox enzymes that utilizes iron-sulfur cluster centers to oxidized glutathione molecules. Glutaredoxin have also been proposed to be involved in iron-sulfur cluster biogenesis.13-15 Glutaredoxins form an iron-sulfur cluster bridged dimer by incorporating two molecules of glutathione, where they provides two thiolate ligands.16-20 Previously, contributions from our group have documented glutaredoxin to undergo cluster exchange with the scaffold protein ISU, and this prompted us to investigate the potential regulatory role of glutathione in cellular iron-sulfur cluster 106 assembly.13 These natural coordination compounds formed by glutathione and iron-sulfur clusters may also serve as important evidences for the potential involvement of glutathione in cellular iron-sulfur cluster assembly pathways.18 Over the past few decades, chemists have synthesized many iron-sulfur clusters using small, non-protein coordinated ligands.21, 22 These iron-sulfur cluster compounds are typically soluble and stable only in non-nucleophilic solvents.23 In 1972, Sugiura et al. synthesized an iron- sulfur cluster coordinated by glutathione molecules. Due to technological limilations, physicochemical properties of these novel clusters were not fully studied. Since glutathione molecules are highly abundant in cells and soluble in water, their coordination product with iron-sulfur clusters are potentially biologically relevant, and deserve careful physicochemical and biochemical investigation on their stability and interactions with cellular components.

107

Results

Mass Spectrometry Studies

The glutathione iron-sulfur cluster complex was synthesized by mixing reduced glutathione, ferric ion and sulfide in water at pH 8.6.24 Formation of this cluster complex was evidenced by electronic absorption, NMR, Mössbauer and electrochemical experiments.24 Mass spectrometry (Electrospray Ionization Mass Spectrometry, ESI-MS) was also demonstrated to be a valuable aid in cluster characterization. Although several iron-sulfur cluster proteins have been studied by ESI-MS the general instability of non- protein-bound iron-sulfur cluster compounds has rendered the characterization of these small complexes by mass spectrometry to be challenging and very rare.25 Nevertheless, when the reaction mixture was analyzed by ESI-MS, an exact mass peak at m/z=1425.3

(Figure 33) was documented and consistent with a cluster carrying two ferric ions,

- 2+ + + + [(GS )4[2Fe-2S] +2H +Na ] . Simulated ESI-MS spectrum based on the natural abundance of isotopes is graphed in Figure 34.

108

Figure 33: Analysis of [2Fe-2S](GS)4 formation by ESI mass spectrometry.ESI-MS of reaction mixture of GSH, ferric chloride and sodium sulfide at 0 min (left), 10 min (middle), and 40 min (right). The exact mass peak at m/z=1425.3 is consistent with - 2+ + + + - [(GS )4[2Fe-2S] + 2H + Na ] where GS is the thiolate form of glutathione, and the - + + + + peak at m/z=1426.3 is consistent with both [(GS )4[2Fe-2S] + 3H + Na ] and an - 2+ + + + isotopic peak of [(GS )4[2Fe-2S] + 2H + Na ] . Exact mass peaks at both m/z=1425.3 and m/z=1426.3 are greater, showing the formation of the cluster in both bis-ferric Fe3+/Fe3+ and mixed valence Fe3+/Fe2+ forms. Note: Spectra collected in collaboration with Wenbin Qi.

109

- 2+ + + Figure 34: Simulated mass spectrum of [(GS )4[Fe2S2] +2H +Na ].

Mass spectrometry results also indicate that the glutathione iron-sulfur cluster self- assembly process is due to the fact that in solution, glutathiones form tetrameric pre- assembled templates. These pseudo-macrocyclic ligands mimic a dynamic combinatorial library, a process where the forming clusters select salt-bridge stabilized glutathione tetramers. The peak observed at 1427.3 in Figure 33 correspond to aggregates of

2- + + 2- glutathione in the reaction mix ([(GS )4 + 9Na ] , with thiolate and carboxylate (GS )).

Exchange of Na+ with K+ resulted in the expected mass shift (Figure 35) observed for the tetramer aggregate. This tetrameric form of glutathione molecules may provide the explanation of the stability of a complex that should not be stable, based on prior literature precedent for ligand complexes of iron-sulfur centers. 110

Figure 35: The template effect of pre-assembled glutathione tetramerwas also observed by ESI-MS with K+ as counter ions. The base peak at 1533.0 corresponds to + [C40H61K8N12O24S4] . Additional peaks are due to the natural distribution of potassium isotopes.

111

Figure 36: ESI-MS data showing solution aggregates of glutathione molecules.

Previous crystallographic studies have documented a hydrogen bond network in crystal structures of glutathione.26 The results presented here indicate that the glutathione tetramers also form in solution. Interestingly, larger multimers were observed, such as pentamers and hexamers. Up to 19-mer were observed (1947.0 at 3+ charged state, see

Figure 36). However, the tetrameric state demonstrated to be the most prominent species, with highest signal relative to other multimeric peaks. It is important to note that the peak intensity often does not necessarily reflect the concentration of the species due to other factors such as ionization efficiency. Aggregation is essentially eliminated by carboxyl ester formation or amine acetylation, respectively (Figure 37). Without these charged groups, the modified glutathione molecules were unable to form iron-sulfur cluster, as observed by formation of insoluble iron sulphides. This supports the notion that the

112 charged functional groups are important in forming intermolecular salt-bridges or hydrogen bonds.

Figure 37: Oligomers were observed for glutathione molecules(top). When charges were eliminated from the functional groups, the oligomers were observed to decrease in intensity. For example, No oligomers were observed in the solution of N- acetylglutathione, and Minor oligomerization was observed in the solution of glutathione ethyl ester, most likely reflecting hydrogen bonding between the ester and protonated amine.

113

Figure 38 illustrates a likely intermolecular salt bridge and/or hydrogen bonding network for glutathione tetramer. The cluster pocket appears to be of the correct size to serve as a preassembled iron-sulfur cluster chelate and ready to accept free iron and sulfide to form the stable cluster complex. This pocket mimics a protein binding site by both providing a pre-assembled ligand set. This pocket may also protect the cluster core from solvent access in the folded state. Interestingly, other glutathione aggregates were not observed in the presence of cluster. These results suggest that there is a synergic interaction with the tetramer species selected by the cluster core, which in turn is stabilized by the glutathione tetramer.

114

Figure 38: A two-dimensional representation of a glutathione-complexed cluster aggregate. Salt bridge formation between carboxylates and protonated amines appear to favor aggregation of glutathione.

115

The intensities of the peaks at 1425.2 and 1426.3 (Figure 39) were observed to increase as the reaction continued. In the theoretical isotopic distribution profile, the pure oxidized complex has the ratio 0.48. Thus, the exact mass peak at m/z=1426.3 represents both an isotopic peak from the oxidized complex as well as the mixed valence complex

- + + + + [(GS )4[2Fe-2S] + 3H + Na ] . Based on the theoretical peak intensity ratio, we were able to deconvolute the intensities of peaks at 1425 and 1426 and calculate the actual intensity of bis-ferric Fe3+/Fe3+ and mixed valence Fe3+/Fe2+ species, respectively. By plotting calculated peak intensity versus time and fitting to first order exponential kinetics, the observed reaction rate constants for the formation of each of the Fe3+/Fe3+ and Fe3+/Fe2+ species are 0.16 ± 0.03 min-1 and 0.34 ± 0.10 min-1, respectively (Figure

39).

116

bisferric mixed valent 6000

4000

2000 Peak Intensity Peak 0

0 10 20 30 40 Time (minutes)

Figure 39: Plot of the peak intensity of the mixed valence Fe3+/Fe3+ form at m/z=1425.3, and the Fe3+/Fe2+ form at m/z=1426.3, versus reaction time. The peak at 1426.3 was deconvoluted based on the natural isotopic distributions (48%). Note: Spectra collected in collaboration with Wenbin Qi.

UV/Vis Absorption Studies

After purification, the glutathione iron-sulfur clusters are observed to be black powder- like solids, and stable under moisture-free desiccators. The powder appears to be hygroscopic and since the cluster core is vulnerable toward hydrolysis without appropriate buffer, the cluster must be placed under dry environments for prolonged storage. The dried cluster is highly soluble in water, up to 10 mM stock cluster solutions have been made without encountering solubility issues. Dissolved cluster appears to be in dark brown/black, with extinction coefficient at 330 nm determined to be ~7600 M-1cm-1.

The cluster solution is unstable in water, with cluster degradation observed within the 117 first 30 minutes (t1/2 = 19 min). This is possibly due to the irreversible loss of a glutathione ligand and the cluster core is subsequently hydrolyzed by water molecules.

However, under a physiological concentration of glutathione (10 mM), the cluster is observed to be slowed significantly. Furthermore, the complex is stable for days under anaerobic atmosphere, indicating that the cluster hydrolysis process must involve an oxidation mechanism which is prevented in the absence of dioxygen.

0.6

0.4

0.2 Absorbance

0.0 300 400 500 600 700 Wavelength (nm)

Figure 40: The UV-Vis spectrum of a 50 uM solution of [Fe2S2](GS)4shows characteristic iron-sulfur cluster transitions at 330 nm and 415 nm due to sulfur to iron charge transfers.

118

Anaerobic 1.8 Aerobic, 10 mM GSH Aerobic 1.6

1.4

1.2

1.0

0.8

Absorbance 330 nm 330 Absorbance 0.6

0.4 0 10 20 30 40 50 60 Time (minutes)

Figure 41: Time-dependent cluster stability study recorded by UV-Vis spectrophotometer at 330 nm.Dissolved cluster was observed to be labile toward degradation under aqueous conditions. However, with excess glutathione in solution, or in the absence of molecular oxygen, the dissolved cluster solution was observed to be stable.

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Mossbauer Spectroscopy Studies

Mossbauer spectroscopy studies were conducted in collaboration with C. Y. Chain, G. A.

Pasquevich, and A. F. Pasquevich at Departamento de Física, Facultad de Ciencias

Exactas, Universidad Nacional de La Plata, Argentina. All samples were synthesized by

Jingwei Li in the Cowan Laboratory. Data acquisition and analysis were done by C.Y.

Chain in the Pasquevich Laboratory.

When the cluster was dissolved in solution, a doublet was observed, and the fitted interactions correspond to the oxidized cluster (Figure 42).The fully oxidized cluster showed parameters of δ = 0.39 mm/s and ΔEQ = 0.68 mm/s, which falls within the range of ferric ions in tetrahedral geometry, coordinated by softer ligands such as sulfur.27-30

When the cluster was dissolved in a solution of glutathione, a mixed valent species of the cluster was observed by Mossbauer spectroscopy. In this case, three interactions were observed, with two additional interactions due to the mixed valent iron atoms.

120

Figure 42: Mössbauer spectra of isolated cluster in solution state.The solution was frozen and spectrum was taken at 212 K. The solution contained 9.3 mM cluster, 10 mM GSH at pH 8.6. The solid line corresponds to a quadrupolar interaction characterized by δ = 0.39 mm/s and ΔEQ = 0.68 mm/s. (Data collected in collaboration with C.Y. Chain)

When compared with ESI-MS results, variations in the relative ratio of the oxidized and mixed valent species are due to the different reaction quenching time points. This difference has also be previously documented when clusters were analyzed by ESI-MS techniques.31 In comparison with solution state clusters, Mossbauer spectroscopy studies of the lyophilized sample showed both oxidized (Fe3+/Fe3+) and mixed valent (Fe2+/Fe3+)

121 species of the cluster (Figure 43). Mossbauer parameters were determined from the overall spectra by deconvoluting the spectrum, and these values are listed in Table 12.

122

Figure 43: Mössbauer spectrum from a sample extracted from the reaction mixture 10 min after mixing. Inside curves correspond to the three interactions contributing to the overall absorption spectrum and fit to deconvoluted Voigtian doublets indicated by square brackets. The data was fit to three Lorentzian doublets (indicated by square brackets) as described in the materials and methods section. (Data collected in collaboration with C.Y. Chain)

123

Table 12: Mossbauer parameters of solid-state cluster and assigned oxidation state, geometry and percent contribution.(Data collected in collaboration with C.Y. Chain)

Species Contribution δ (mm/s) ΔEQ (mm/s) Oxidation state Geometry

I 56 % 0.21 0.55 III 4-coord, Td II 23 % 1.10 2.54 II 5- or 6-coord. III 21 % 0.33 0.79 III 4-coord, Td

These data suggest that after an initial phase, the solution reaches an equilibrium in which there is a steady-state ratio of fully oxidized cluster (species I) and mixed valence cluster (species II and III). The ethanol precipitated product showed both the mixed valence Fe3+/Fe2+ and fully oxidized Fe3+/Fe3+ cluster species (Figure 43). Isomeric and quadrupolar shifts of III agree with literature values of ferric iron in the reduced 2Fe-2S cluster.32-35 The fitted spectra also indicated an equimolar ratio of species II and III, and support the identity of the one-electron reduced diiron cluster. Mössbauer experiments show that when the freshly synthesized cluster complex is isolated by ethanol precipitation and re-dissolved with excess glutathione then only fully oxidized cluster is

2- present, exhibiting the NMR and other physical characteristics of the [2Fe-2S](GS)4 cluster.24 These results indicate that while the fully oxidized cluster is the most stable solution state for the cluster, the conditions of the initial reaction mixture (with excess iron and sulfide and thiols) maintains a thermodynamic fraction of reduced cluster.

The major difference between the solid-state and solution spectra is species II (Table 12) which shows a feature with δ ~ 1.10 mm/s that is assigned to either a five or six coordinate Fe2+ center in the mixed-valent cluster based on similarity to published data,

124 allowing for differences in sample temperature.36-39 Moreover, the Gaussian standard deviation sigma of the interaction corresponding to Fe2+ is larger than the other peaks, showing that the atomic arrangement around Fe2+ is less defined. Expanded coordination reflects both the larger high-spin ferrous ion and the presence of an intramolecular chelate effect through carboxylate ligation from one or two carboxylates on two glutamate residues (Figure 37), and is consistent with prior reports of ferrous centers in binuclear iron36-39 as well as proton relaxation studies that indicated the glutamate side chains to lie in close proximity to the cluster center.40

Solution Ionic Strength and Cluster Stability

Note: cluster stability and salt-dependence studies were conducted in collaboration with

Stephen Pearson. All sample synthesis and data acquisition were performed by Jingwei

Li and Stephen Pearson.

Results presented thus far have demonstrated GSH coordinated [2Fe-2S] cluster to be stabilized by intramolecular salt-bridges. In the absence of excess glutathione, the cluster is readily and irreversibly hydrolyzed, as its optical spectrum shows a decrease in absorbance due to charge transfer bands. Previously we demonstrated that the glutathione tetramer in solution may facilitate the formation of the cluster complex by Nature’s equivalent of dynamic combinatorial chemistry, by formation of an apo template (an intermediate) that provides a scaffold for assembly of the [2Fe-2S] cluster by coordination of four GSH molecules. Intramolecular glutathione-glutathione interactions 125 through salt-bridge formation were proposed at the time based on results from functional group modifications and ESI-MS analyses. Tetrameric states of GSH were not observed when carboxyl ester or amine acetylation modifications were made 41. When considered with previously determined GSH 1H relaxation rates, these experimental results suggested that GSH aggregates are held together by intramolecular interactions 40. The resulting “macrocyclic” tetramer provides a coordination pocket for the [2Fe2S] cluster and stabilizes it toward hydrolysis reactions.

To further elucidate the effects of the inter-glutathione interactions, the kinetics of cluster breakdown was studied in the presence of varying solution concentrations of halide salts of various alkali and alkaline earth metals. These kinetic profiles suggested a sequential reaction mechanism, where the binding of salt ions lead to cluster degradation; presumably because loss of salt bridge contact results in loss of the macrocyclic effect, and also expose the cluster more externally to solvent. Based on initial rates of degradation, the binding affinities between salt ions and cluster were modeled, and the physical properties of these cations and anions (including charge, ionic radius and hydration enthalpy) were plotted against the equilibrium constant (KS). The identity of the ions dictated their binding affinities to the negatively-charged [2Fe-2S](GS)4 cluster and their ability to compete with carboxylate-amine salt-bridges. Furthermore, solution ionic strengths and the cluster breakdown rates were fit to Debye-Hückel equations and observed to be positively correlated. This relationship was observed for all salts used and provides direct evidence for the existence of salt-bridges within the glutathione

126 coordinated iron-sulfur cluster. Such recognition of cluster binding model lends further support to a theme where it reflects Nature’s equivalent of dynamic combinatorial chemistry 42.

The overall absorbance of the cluster solutions was observed to decrease over time, and the extinction coefficient was used to convert the raw absorbance values to the concentration of cluster remaining in solution. For example, over a period of 30 minutes,

18 uM of [2Fe-2S](GS)4 cluster was observed to be hydrolyzed in a solution of 0.62 M

NaCl (calculated from Figure 44).

127

Figure 44: A plot of cluster initial degradation rate at various concentrations of salt (NaCl) suggest a pre-equilibrium binding kinetic profile between salt ions and cluster. The initial rates were determined from the first 15 min of cluster degradation profiles measured by absorption spectroscopy. The signals were then converted to concentrations of clusters, and rates were determined from the slope of the time vs. concentration plot. Absorbance of the cluster at 330 nm was used to calculate the concentration. Data obtained from the 0 mM NaCl experiment was used to subtract out minor hydrolytic breakdown. Standard curve for the cluster concentration was calculated based on 0.2 mL sample solution in 96-wellplate sample well and illustrated in Figure 45.

128

1.0

0.8

0.6

0.4

0.2 Absorbance(330 nm)

0.0 0 50 100 150 200 [Fe S (GSH) ] (uM) 2 2 4

Figure 45: A concentration standard curve for glutathione iron-sulfur clusters dissolved in buffer A. 96-well plates were used for data collection. Solution volumes were set at 200 uL. Extinction coefficient was measured to be 3024 M-1cm-1 with these conditions. The overall equation for conversion between cluster concentration and absorbance at 330 nm is: Abs = 3024*[Fe2S2(GSH)4] + 0.217.

129

The hydrolysis product is observed to be light-yellow colored following cluster degradation and loss of absorbances at the charge-transfer bands. The cations in solution measurably increased the Fe-S cluster degradation rate (for example NaCl in Figure 44).

This general trend was also observed for all alkali and alkaline earth salts of halides.

Equation 4 was derived from a mechanism (Scheme 2) of cluster degradation, and used to quantitatively determine the binding affinity of different ions to the negatively charged

[2Fe-2S](GS)4 cluster (Figure 46). The solution salt concentration range of 0 – 0.62 M was sufficient for fitting the rates to their binding affinities toward cluster in terms of KS values corresponding to a consecutive kinetic profile. The range of KS values spread from

0.8 ± 0.1 to 74.0 ± 7.1 M-1, with values significantly higher for the alkali and alkaline earth metal salts than halide salts.

Scheme 2: A proposed cluster degradation mechanism in the presence of salt ions based on time-dependent absorption spectroscopy results.See Experimental Methods section for detailed mechanistic explanations.

130

102

] ] 1

-1 10

S K

100

2 2 2 2 LiCl KCl NaI NaCl CsCl MgCl CaCl SrCl BaCl NaFNaClNaBr

MgCl Figure 46: The KS values were determined from derived equations for various salts.The ionic strength used for each salts are identical and listed in the Experimental Methods section. The initial rates were determined using the absorbances from the first 15 minutes of the plate reader data. Triplicate sets were averaged for each salt and the error bars represent the standard error determined from individual data fits.

131

The positive relationship between cluster breakdown rate and solution ionic strength most likely reflects the counter-ion screening effect, which reduces the cluster intramolecular salt-bridge stability. The salt dependence of the rate of this reaction can be modeled using the primary kinetic salt effect 43, 44. Two ions, P+ and N-, react via Coulombic attraction and form a stable and ionic interaction. Upon addition of charged ions, the stability of the salt-bridge is weakened, which leads to an overall decrease in cluster stability. The first step of the reaction is binding of salt ions to the cluster complex. This is primarily due to electrostatic attractions and it is a reversible process. The binding constants between ions and cluster were measured for individual salts used from equation 4 (Experimental

Methods Section). Our data suggest that the binding constant between different ions used depended on the physical characteristics of the ions such as the charge density. The second step of the reaction is the interaction between the bound salt ions with the functional groups responsible for the intramolecular salt-bridges. Our previous studies on these charged functional groups suggested that they are essential for cluster formation, and thus a cluster with disrupted salt-bridges is likely to rapidly hydrolyzed and degraded. In summary, the first half of the reaction may be considered as a binding step between the solution salt ions and cluster. This results in an activated cluster-ion complex with which the intra-molecular salt bridges may be significantly altered due to the bound salt ions. This intermediate species is subsequently hydrolyzed, which is an irreversible reaction.

132

There is also a positive relationship between log(KS) and charge density (Figure 47).

These slopes are more prominent for the halides than for alkali and alkaline earth metals.

Similarly, the ion charge density has a higher effect on binding of alkali metals than alkaline earth metals. These observations may be explained by the charge of the ions, where the 2+ alkaline earth metals are attracted to the overall negatively-charged cluster molecule with less dependence on the charge density. For negatively charged halides, our data suggest that the charge density strongly influences the binding of the anions to the cluster. Although the cluster is overall negatively-charged, cluster break down rates depended on the identity and concentration of anions. These observations suggest that the anions are interacting with positively-charged functional groups within the cluster and consequently weaken intramolecular salt-bridges.

133

- - Figure 47: A plot of charge density (z/r) vs. log (KS) for selected ions.These are: Cl , Br , I- for halides, Li+, Na+, K+ for alkali metals, and Mg2+, Ca2+, Sr2+ for alkaline earth metals. Linear fits are drawn in to qualitatively demonstrate the relationship between the axial variables. Error bars represent signal-to-noise ratio calculated from KS in Table 13.

134

Table 13: A list of ions used and values of solution ionic radii, charge density, hydration enthalpy, KS and correction factor from data fit.Mean ionic-radii were used for equation 7 for fitting of CF. Ionic charge densities were calculated by taking the ratio of absolute charge and volume. Literature values were obtained from the following sources: a – Reference 45, b – Reference 46, c – Reference 47 and d – Reference 48.

Furthermore, hydration enthalpy may also have an effect on KS. Lower charge density leads to decreased hydration, increasing the electrostatic interaction between hydrated ions and cluster, which in turn decreases cluster stability. Figure 48 is a plot of previously determined hydration enthalpies (ΔHm,hyd) for selected ions and their binding affinities toward the cluster. Ions such as halides were observed to have stronger dependence between the ΔHm,hyd and KS to the negatively charged cluster. The ΔHm,hyd has relatively less effect on cluster binding affinities for highly charged alkaline earth metals with high

ΔHm,hyd, as observed with a lower slope on the plot of ln(KS) vs. ΔHm,hyd. When hydration enthalpy is low, the binding affinity of ions to the cluster is dictated by ΔHm,hyd . For

135 example, the trend of halides (black traces in Figure 48) showed a high dependence of KS on ΔHm,hyd. In contrast with the halides, alkaline earth metals with high magnitudes of

ΔHm,hyd showed less effect binding to cluster.

- - - Figure 48: A plot of ΔHm,hyd vs. log (KS) for selected ions.These are: Cl , Br , I for halides, Li+, Na+, K+ for alkali metals, and Mg2+, Ca2+, Sr2+ for alkaline earth metals. Linear fits are drawn in to qualitatively demonstrate the relationship between the axial variables. Error bars represent signal-to-noise ratio calculated from KS in Table 13.

136

A modified Debye-Hückel equation was used to further study the effect of solution ion concentration on cluster stability. This fit for NaCl solution is illustrated in Figure 49.

The first term of equation 7 is derived from electrostatic screening, and this term is dependent on the charge and the concentration of the ionic species in solution. The second term describes the binding of water molecules to ions at high ionic strengths. This term is responsible for the concave down feature at high salt concentrations. The base

-1 rate was measured to be k0 = 0.00571 min (Figure 50). The quality of fit between experimental results and counterion screening theory was measured by the correction factor, CF, for each salt used. For the 11 different salts used, CF values were observed to be close to 1 (0.453 – 1.49), with the averaged CF of 1.02. These values suggest that ions in solution destabilize the cluster intramolecular interactions.

137

Figure 49: The log of k/k0 was fit against the square-root of ionic strengths.The modified Debye-Hückel equation (equation 7) was used to fit the cluster degradation data obtained by varying the ionic strength using NaCl. The CF values were measured from the fit and reported in Table 13.

138

Figure 50: A slow cluster breakdown was observed when dissolved in solution without additional salt added. Kinetic rate constant (k0) was determined from this plot, and used for calculations for CF values in Equation 7. A minor absorbance increase was observed due to precipitate formation, and this contribution does not influence the calculated rates of cluster break down as each measurement was baseline-corrected with 0 mM salt solutions prior to fit.

Previous results from protein models such as the PYP system showed dependence on ion concentration in solution, as its activity peaked near [KCl] ~ 720 mM 49. The rational was that at low ionic strength, the activity coefficient decreased due to counterion screening, whereas at higher ionic strength binding of water by ions lead to an increase in the activity coefficient. Our data indicate that the salt induced cluster breakdown rate peaked

139 near 520 mM with NaCl solution and agrees with the ion hydration model predicted by the modified Debye-Hückel equation. It is also important to note that the peak activity not only depends on the hydration of ions, but also the interaction of the ions with salt- bridges being studied. Therefore, the approximated concentrations corresponding to peak activities between different systems should only be used for the purpose of qualitative comparison.

Results from this report further support our previous observations where modifications to

GSH lead to the loss of its ability to form the tetrameric scaffold 41. Solution chemistry also suggests that physiological pH is important in protonation of GSH functional groups.

These polar groups not only are important in salt-bridge formation and stabilizing the cluster from hydrolysis, but may also interact with physiological protein partners, such as the iron-sulfur cluster scaffold protein hISU, during biosynthesis and transport of iron- sulfur clusters. Future studies on covalently linked GSH multimers may provide further insights on the importance of inter-GSH salt-bridges and cellular GSH solution dynamics.

140

Nuclear Magnetic Resonance Studies

The chemical property of glutathione coordinated [2Fe-2S] was further investigated by

NMR technique. 1H NMR scan of the cluster dissolved in solution indicated chemical shifts of the cysteine side-chain protons have moved down field upon coordination to the cluster (Figure 51).

Figure 51: NMR spectra of glutathione cluster.A solution of 1 mM glutathione complexed [2Fe-2S] cluster showing cysteine α and β protons (bottom) shifting downfield upon coordinating to iron-sulfur cluster (middle). The two cysteine β protons are observed to shift from 2.96 ppm to 3.32 ppm, and from 2.89 ppm to 2.99 ppm. The cysteine α-proton shift from 3.72 ppm into the 1HO2D at 4.70 ppm.

141

Unpaired electrons on iron usually cause hyperfine shift and thus resonance broadening when studied by NMR techniques. For example, when a 0.5 mM ferric chloride was added to the glutathione solution, the signal to noise level of the glutathione proton peaks were observed to be significantly rendered (Figure 51 top vs. bottom). This is due to the coordination of glutathione to ferric ions, and since the ferric center is paramagnetic, it caused the relaxation rate of nearby protons to increase. Previous NMR studies on iron- sulfur cluster proteins demonstrated that these prosthetic groups also cause peak broadening of nearby resonances to various extents, depending on the identity, structure and chemical environment of the cluster pocket.49-59 Interestingly, glutathione protons of the [[Fe2S2](GSH)4] cluster remained within the diamagnetic window (Figure 51, middle). The cysteine α proton disappeared from the spectrum, however, various temperature and proton homonuclear decoupling NMR studies showed that this proton had been shifted into the 1HO2D peak at 4.70 ppm (Figure 52 and Figure 53, respectively).

142

Figure 52: Variable temperature NMR studies of 1H resonances at 27 ºC, 30 ºC, 40 ºC and 50 ºC.All peaks were observed to shift downfield with increasing temperature. The cysteine α proton was observed to be shifted into the water peak, but with higher temperature, it moved out of the water peak and observed at 50 ºC.

143

Figure 53: Proton Homonuclear Decoupling of Cluster at 4.70 ppm.A doublet of quartet was reduced to a doublet of doublet when the cysteine α-proton was decoupled from the β-protons, indicating that the α-proton was in the 1HO2D peak at 4.70 ppm.

Proton NMR results indicate that there is strong antiferromagnetic coupling between the pair of ferric centers in the relatively symmetric coordination environment and negligible paramagnetic influence. Consistent with this, variable temperature studies show the protons on cluster-bound glutathiones to display a negligible temperature dependence, relative to free glutathione.

In order to further study the effects of iron-sulfur cluster coordination on glutathione protons, spin-lattice (T1) and spin-spin (T2) relaxation time studies were performed

144

(Figure 54). These data were converted to R1 and R2, respectively, and tabulated in Table

14.

Table 14: Relaxation times of protons of GSH, GSSG, GSH + Fe3+ and GSH coordinated cluster.These data were used to calculate the cluster induced relaxation ratio for each proton (R1M and R2M).

C C 1 1 800 C 800 C

(ms) 2 2 1 700 (ms) 700 E 2 E   600 600 E E   500 500

400 400

300 300

200 200

100 100

0 0 Spin-Spin relaxation time T time relaxation Spin-Spin

Spin-Lattice relaxation time T time relaxation Spin-Lattice GSH Cluster GSH Cluster

Figure 54: Comparisons of relaxation times between free and [2Fe-2S] bound glutathione.Spin-Lattice (T1) relaxation times (left) and Spin-Spin (T2) relaxation times (right) show significant increases in longitudinal and transverse magnetization recovery rates, respectively.

Without paramagnetic species in solution, relaxation times for GSH and GSSG were observed to be in the ranges of hundreds of milliseconds. However, when ferric chloride

145 was added to a solution of GSH, the relaxation times, especially T2, decreased drastically.

This is consistent with the idea that the unpaired electrons increases the relaxation rates of the protons on GSH. These T2 times were different from those found in [2Fe-2S] clusters. This agrees with the idea that the two ferric irons are antiferromagnetically coupled, and thus the overall paramagnetism is lower than that of ferric ions. The ratio of

GSH T1 and cluster T1 yields a ratio of cluster induced relaxation rates (R1M and R2M).

The ratios of R1M/R2M indicate contact relaxation is relatively more dominant over dipolar relaxation for Cβ1 and Cβ2 protons than Eβ and Eγ protons. These observations, along with the increased relaxation rates for protons is consistent with cluster ligation by thiolate ligands. These results also agrees with optical and Mossbauer absorption data.

The increased rates for glutamate protons agrees with the proposed model that GSH molecules forms a pocket via salt-bridges or hydrogen bonds, which protects the cluster core from hydrolysis.

When the cluster was titrated with iron chelating ligands such as cyanide ions, free glutathione molecules were observed as cluster was hydrolyzed (Figure 55). Free glutathione, as observed by the cysteine α-proton, appeared on the proton NMR spectrum with 10 equivalence of sodium cyanide was added to the solution. Furthermore, the S/N ratio of the peaks significantly increased as more sodium cyanide was added. This is consistent with the idea that the paramagnetic effect of ferric ions on glutathione molecules were reduced as cyanide coordinates to iron.

146

Figure 55: Sodium cyanide titration to a solution of glutathione cluster.As the concentration of cyanide ion was increased, cysteine α-protons appeared near 4.25 ppm, indicating glutathione dissociated from the iron-sulfur cluster.

147

Experimental Methods

Cluster synthesis and ESI-MS analysis

The glutathione coordinated 2Fe-2S clusters were synthesized by mixing ferric chloride

(20 mM) and sodium sulfide (20 mM) to glutathione solution (50 mM, pH 8.6). The total volume of the solution is 5 mL. To isolate the cluster, 900 uL ethanol was added to 100 uL of the mixed solution, and vortexed briefly. The resulting mixture was centrifuged at

13,000 rpm for 10 min, washed twice with ethanol and dried under vacuum. The cluster is usually dried for at least 4 hours, and the resulting pellet should appear free of moisture. For ESI-MS analysis, the reaction mixture was applied to Bruker Micro-TOF

(ESI) spectrometer and data was analyzed by use of DataAnalysis software (Bruker). For the potassium adduct assay, KOH was used instead of NaOH for GSH solution pH adjustment.

Synthesis of modified glutathiones

N-acetylglutathione was prepared according to previously published literature by

Anderson et al.60 In brief, glutathione (GSH, 1.00 g, 3.25 mmol) was dissolved in formic acid (3 mL, 3.66 g) and stirred at room temperature. Acetic anhydride (1.5 mL, 1.62 g) was added dropwise to the stirred solution. The resulting mixture was allowed to react at room temperature for 2 h. Diethyl ether (45 mL) was added to the reaction mixture and the resulting precipitate was separated by gravity filtration. The precipitate was washed with petroleum ether (3 x 10 mL) and dried under vacuum. The crude product was

148 recrystallized from a 1:1 mixture of ethyl acetate and dimethyl formamide to yield N- acetylglutathione as a white solid (0.90 g, 2.57 mmol, 79%). Characterization of N- acetylglutathione was confirmed by 1H NMR experiments (Figure 56 and Figure 57).

Glutathione ethyl ester was purchased from Sigma-Aldrich and used without further purification.

1 Figure 56: H NMR spectrum of N-acetylglutathione in D2O.

149

1 Figure 57: H COSY NMR spectrum of N-acetylglutathione in D2O.

UV/Vis Absorption Assays

All UV/Vis measurements were obtained on Varian Cary 50 UV-Vis

Spectrophotometers. For cluster degradation studies, a solid sample of glutathione-iron- sulfur cluster is weighed and dissolved to 1 mM in solution containing the appropriate 150 concentration of salt of interest. This solution is then diluted ten-fold using the same salt solution, and a UV-Vis spectrophotometer was used to measure the cluster degradation during incubation. The characteristic cluster absorbance peaks at 330 nm and 415 nm were used to quantitatively determine the relative cluster concentration in the reaction mixture. The cluster break-down leads to formation of a brown precipitate. To remove this suspended precipitate in the solution, the reaction mixtures were centrifuged at

13,000 rpm in an Eppendorf centrifuge tube for 1 min prior to each absorbance measurement.

Mössbauer Spectroscopy

Note: Mossbauer spectroscopy studies were conducted in collaboration with C. Y. Chain,

G. A. Pasquevich, and A. F. Pasquevich at Departamento de Física, Facultad de Ciencias

Exactas, Universidad Nacional de La Plata, Argentina. All samples were synthesized by

Jingwei Li in the Cowan Laboratory. Data acquisition and analysis were done by C.Y.

Chain in the Pasquevich Laboratory.

A 10 mg sample of 57Fe metal was dissolved in 250 uL of a 1:1 mixture of concentrated

HCl and HNO3. The suspension was stirred for ~ 10 min until all solids had dissolved and gas evolution ceased. A solution of 5M NaOH was then slowly added in aliquots of

20 uL and the pH of the solution checked after each addition. Additions continued until a pH ~ 7.4 was obtained; typically ~ 300-340 uL 5M NaOH total. The color of the mixture

151 turned light yellow, and then dark orange. This 57Fe stock salt solution was subsequently used to synthesize 57Fe-labeled clusters. The solution was centrifuged to discard any precipitate, and then 0.077g GSH in 4 mL H2O, pH 7.4 was added. A 500 uL solution of

200mM Na2S was added and reaction continued for 10 min prior to precipitation with a ten-fold volume excess of ethanol with stirring. The resulting solution was centrifuged at

14,000 rpm and the supernatant removed by decanting. The solid was resuspended two additional times in ethanol with stirring and it was collected by centrifugation prior to final drying in a speedvac for a period of up to 4 h to obtain the 57Fe-S glutathione cluster complex. The Mössbauer absorber was prepared by distributing uniformly 82 mg of this sample in a 2 cm-diameter holder.

Mossbauer experiments were conducted with a constant acceleration at r.t. A Rh matrix and nominal 50 mCi 57Co source was used. For the process modules, ORTEC (142 pc,

572 A, and 551-TSCA) was used with a LND-4045 counter coupled with CMTE MA-

250 transducer. Velocity reference wave generations and spectra acquisition were obtained by instrumentation in recently developed by the Pasquevich Laboratory.61 As a reference, isomer shifts for α-Fe were taken at room temperature. Results were fit to

Lorentzian doublets with the Levenberg-Marquardt algorithm.62 Sample thickness were accounted for by considering the Mössbauer transmission integral63 , and relevant parameters such as isomer shifts (δ) and quadrupolar splittings (ΔEQ) were extracted from spectra fits.

152

Salt-Dependence of cluster stability and measurement of kinetics

Note: cluster stability and salt-dependence studies were conducted in collaboration with

Stephen Pearson. All sample synthesis and data acquisition were performed by Jingwei

Li and Stephen Pearson.

All ionic salts were dissolved in the reaction buffer (10 mM Tris-HCl/Trizma base, pH

8.6, buffer A) to the appropriate concentrations. Prior to the reactions, the buffer was flash frozen, thawed under vacuum, and purged with argon for 3 minutes. The buffer was kept under anaerobic atmosphere until use. A 96-well plate (Immulon ® 4HBX) was used to measure the kinetics of cluster breakdown on a SpectraMax M5 (Molecular Devices).

The solution conditions (final concentrations) are: 0, 50, 112.5, 200, 312.5, 450, and

612.5 mM for monovalent salts and 0, 16.7, 37.5, 66.7, 104.2, 150, and 204.2 mM for divalent salts. The final concentration of cluster was 100 uM. The reaction was monitored at 25.0ºC and 330 nm for 30 minutes with 30 second intervals. All data measurements were collected in triplicate sets.

Data fitting and derivations

Note: cluster stability and salt-dependence studies were conducted in collaboration with

Stephen Pearson. All sample synthesis and data acquisition were performed by Jingwei

Li and Stephen Pearson.

153

All ionic salts were dissolved in the reaction buffer (10 mM Tris-HCl/Trizma base, pH

The solution conditions (final concentrations) are: 0, 50, 112.5, 200, 312.5, 450, and

For the KS measurements, raw absorbances of each triplicate were averaged and converted to the concentration of cluster using the equation: absorbance = 3024 *

[cluster] + 0.217 derived from a calibration plot (Figure 45). The concentration of 0 mM salt trial was subtracted from each data set to remove minor absorbance shifts over the duration of the experiment. Observed rate constants were determined from the initial slopes of 15 minutes in unit of μM/min, and plotted against the salt concentrations and fit to equation 4. A mechanistic explanation of the salt-dependent cluster degradation is presented in Scheme 2. In short, salt ions bind to the glutathione coordinated cluster with an equilibrium constant of KS = k1/k-1. The bound salt ion then subsequently interact and disrupt the intramolecular salt-bridges (or hydrogen bondings), which results in a less stable, activated cluster complexǂ. The final step of the cluster breakdown is a fast,

154 irreversible hydrolysis reaction. The determined KS constants for various salts are presented in Figure 46.

+ constant Equation 4

For data fitting to the modified Debye-Hückel equation 49, concentrations of cluster determined from the plate-reader absorbances were used to determine the rate constants.

Equation 5

23 -1 Using SI values of Avogadro constant (NA) = 6.02 x 10 mol , Boltzmann’s constant (k)

-23 -19 = 1.38 x 10 J/K, proton charge (e) = 1.60 x 10 C, permittivity of vacuum (εo) = 8.85

-12 3 x 10 F/m, solvent dielectric constant (εr) = 78.38, solvent density (ρ) = 997.05 kg/m ,

Temperature (T) = 298 K and Equation 5, A was calculated to be 0.510, and B =

0.328*diameter of ions 65. These values were combined with the modified Debye-Hückel equation derived from previous published literature (equation 6)43, 44, 49 and rearranged to yield equation 7.

Equation 6

Equation 7

Where CF, a correction factor, is introduced to measure the quality of data fit. ZAZB was set to -1 and averaged ion diameters were used for d45. Constant C accounts for the hydration of ions at high concentrations, which may decrease the rate of hydrolysis

155 reactions. Values of log(k/k0) were plotted against to determine CF, and these values are presented in Table 13.

Nuclear Magnetic Resonance Experiments

For NMR experiments, 5 mm tubes were obtained from Wilmad-Labglass. All cluster experiments were done in D2O, which was purchased from Sigma-Aldrich. Cluster samples synthesized as previously described, and solution 1H NMR spectra were acquired at a concentration of 1 mM. For the glutathione and ferric chloride control, concentrations of 4 mM glutathione were used. Except for the variable temperature NMR experiments, all spectra were collected at 300.1 K. Proton spectra were recorded on a

Bruker DRX 500 MHz spectrometer equipped with a 5 mm TXI cryo-probe. The T1 and

T2 relaxation data were recorded on a Bruker DMX 600 MHz spectrometer equipped with a 5 mm triple-resonance probe. Variable temperature studies were recorded on a

Bruker DPX 400 MHz equipped with a 5 mm BBI probe. Standard pulse programs by

Bruker were used for advanced NMR experiments such as proton homonuclear decoupling, Spin-Lattice and Spin-Spin relaxation, and 1H-1H COSY experiments.

156

Conclusion

Several recent reports have demonstrated glutaredoxins (Grx) to form [Fe2S2] cluster- bridged dimmers. In these cases, glutathione provides two thiolate ligands, and serve as examples where glutathione coordinations to iron-sulfur clusters are found in nature.

Results presented in chapter demonstrate that glutathione molecules coordinate to iron- sulfur clusters under physiological glutathione concentration and other solution conditions. Results from optical, redox, ESI-MS, Mössbauer and NMR analyses suggest

2+ the cluster is in its oxidized form ([Fe2S2] ), with four glutathione molecules coordinated

2+ - 2- around it. NMR characteristics suggest that the iron centers in [[Fe2S2] (GS )4] are antiferromagneticly coupled. Using techniques such as mass spectrometry and

Mössbauer spectroscopy, the reduced intermediate cluster was also observed and documented. Furthermore, detection of the glutathione-complexed Fe-S cluster and reaction intermediates were also observed and documented. These were accomplished by using a combined application of mass spectrometric and Mössbauer techniques. The resulting cluster complex is stable at physiological pH and a potential component of the cellular labile iron pool.40, 65 Formation of hydrolytically stable small-molecule ligated iron-sulfur clusters in aqueous conditions has not been document prior to this study. The stability of the glutathione coordinated iron-sulfur centers are likely due to the salt- bridges and hydrogen bondings on glutathione molecules. These interactions are also observed for free glutathione molecules in solution, suggesting a pre-formed pseudo-

157 macrocyclic molecule and plausible role for this species as an active physiological component of cellular iron chemistry and iron-sulfur cluster biosynthesis.

158

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164

Chapter 5: Synthesis, Characterization and Biological Chemistry of a Glutathione Complexed [2Fe-2Se] Cluster

Introduction

Iron sulfur clusters are prosthetic groups that find wide utility in biological chemistry, including cellular iron regulation, redox chemistry and structural stabilization of proteins.1, 2 Cluster biogenesis pathways are widely conserved in all three kingdoms of life.3-6 The basic clusters vary in size from as small as one iron center, in the case of rubredoxin, to up to 4Fe-4S units in ferredoxins.7, 8 More complex clusters that incorporate other metal centers and non-stoichiometirc Fe-S ratios have also been identified, such as the 3Fe-4S and Ni-Fe clusters found in membrane bound NiFe hydrogenases9.

Until recently, water stable coordination complexes of Fe-S centers have not been well characterized as a result of hydrolytic instability. We have reported the synthesis and characterization of a glutathione-coordinated cluster possessing biologically relevant activities, as well as stability under physiological solution conditions. The proposed role for this cluster is stabilization and transient storage of 2Fe-2S centers following synthesis at the scaffold protein hISU, and subsequent cluster transport and reconstitution into iron- 165 sulfur cluster proteins. This is observed by both glutathione cluster extraction from holo protein and reconstitution of the apo protein.10 This model is consistent with the observed unstable holo form of the scaffold protein hISU after cluster synthesis, presumably due to solvent accessibility of the cluster.11

Selenium substituted iron-sulfur cluster proteins have similar yet altered enzymatic parameters, such as higher catalytic efficiency observed in the iron-selenium cluster found in C. pasteurianum ferredoxin dehydrogenase.12 Selenium substitution in iron- sulfur cluster proteins has also been used as a probe of electronic structure and reactivity in a number of Fe-S proteins, such as putidaredoxin13, adrenodoxin14 and nitrogenase15, which have all been reconstituted with selenide in place of bridging sulfide.16 Cluster transport of native Fe-S clusters between ISU-type scaffold protein and relevant protein partners has also been extensively studied.17-19 Selenium substituted iron sulfur clusters yield distinct and interesting enzymatic and spectroscopic properties compared to the

20-25 native sulfur derivatives. To date, Fe2Se2 cluster transport to apo-proteins has yet to be extensively studied.

In this report, the synthetic glutathione complexed iron-selenide cluster was demonstrated to be stable in aqueous conditions under anaerobic conditions. Furthermore, this glutathione iron-selenium cluster is a good candidate for the study of cluster uptake and transport between iron-sulfur cluster proteins. The large change in absorption spectra

166 when cluster is transferred from the glutathione complex to the protein provides a useful spectroscopic probe.

Herein, we report the synthesis and characterization of the selenium substituted iron- sulfur glutathione cluster and exchange chemistry with iron-sulfur scaffold proteins and target apo ferredoxins. The optical spectrum showed characteristic absorption peaks in the iron-selenium reconstituted proteins. ESI-MS, Mossbauer and NMR spectroscopy were used to confirm and compare the selenium cluster with the native sulfide derivative reported previously. Also, the ability of iron-sulfur cluster assembly protein hISU to be reconstituted with glutathione 2Fe-2Se cluster, and subsequently transport the cluster to the apo Ferredoxin2 (Fd2) protein is described. The kinetics of selenium cluster transfer steps were studied by electronic absorption techniques. In comparison to the sulfur derivatives, transfer rates of selenium substituted clusters were observed to vary depending on the clusters’ local environment in the native hISU and its mutant (D37A, which has a less hydrophilic binding pocket).

167

Materials and Methods

Synthesis and charaterization

A solution of dithiothreitol (DTT, 25 mM) was used to reduce sodium selenite (2.5 mM) in situ to generate sodium selenide (Scheme 3). Ferric chloride (2.5 mM) and sodium selenide (2.5 mM) were added to a glutathione solution (0.077 g in 4.0 mL H2O at pH

8.6). The resulting mixture was vortexed and precipitated by addition of ethanol (45 mL). The resulting product was isolated via centrifugation at 13,000 rpm for 10 min, and washed with additional ethanol (2 x 50 mL) and dried under vacuum. The crude product was ground to fine powder and triturated with ethanol (10 mL) to remove residual impurities. The resulting [Fe2Se2](GS)4 complex was dissolved in water for subsequent

ESI, UV-Vis, CD and cyclic voltammetric experiments. A Bruker MicroTOF spectrometer was used to acquire ESI-MS results and DataAnalysis (Bruker) software was used to process and plot the results. A Varian Cary 50 UV-Vis spectrophotometer was used for all UV-Vis measurements. Potentiostat/Galvonostat Model 263 (EG&G

Princeton Applied Research) was used for electrochemical measurements. Pyrolytic

Graphite Working Electrodes (Pine Research Instrumentations) and standard Ag/AgCl electrode were used to measure the reduction potential of the cluster and reconstituted proteins. For preparation of protein films on the PGE electrode, a 50 mM Tris-HCl buffer at pH 8 was used. All CD data acquisitions were obtained on Jasco J-815 CD

Spectrometer (JASCO) equipped with Quartz or glass cells with 1.0 cm path length.

Triplicates were done for each reaction and the results were averaged. Data were

168 smoothed by the Jasco instrument data analysis program over 25 data points. Observed rate constants were obtained by Origin 7 data fitting software.

Scheme 3: Synthesis of Fe2Se2GS4 cluster.Previously published protocols for Fe2S2GS4clusters yields GSH contaminated clusters. Therefore, additional trituration steps were needed to further purify the product.

Nuclear Magnetic Resonance Experiments

NMR samples were dissolved in 100% D2O in 5 mm NMR tubes (Wilmad-Labglass).

Solution samples of glutathione and the [2Fe-2Se] glutathione cluster complex were freshly prepared at concentrations of 4 mM (pH 8.6) and 1 mM, respectively. All NMR experiments were recorded at 300.1 K. Proton (1H) spectra for all samples were recorded on a Bruker DRX 500 MHz spectrometer equipped with a 5 mm TXI cryo- probe and processed with XWIN-NMR v3.5 software.

169

Mössbauer Spectroscopy

Note: Mossbauer spectroscopy studies were conducted in collaboration with C. Y. Chain,

G. A. Pasquevich, and A. F. Pasquevich at Departamento de Física, Facultad de Ciencias

Exactas, Universidad Nacional de La Plata, Argentina. All samples were synthesized by

Jingwei Li in the Cowan Laboratory. Data acquisition and analysis were done by C.Y.

Chain in the Pasquevich Laboratory.

Preparation of an 57Fe3+ solution for Mössbauer sample preparation was performed as described previously.26 The resulting solution (25 mM, 62.5 uL) was added to 9.6 mg

GSH in 500 uL H2O at pH 8.6. A 62.5 uL solution of 25 mM sodium selenide was added and reaction continued for 10 min prior to precipitation and subsequent isolation following the general cluster protocol described earlier. The Mössbauer data acquisition and data processing were done in collaboration with C.Y. Chain (Pasquevich Laboratory) at Departamento de Física, Facultad de Ciencias Exactas, Universidad Nacional de La

Plata, Argentina. Same experimental methods were used to acquire the data, and these parameters were described in Chapter 4.

Protein Purifications and Reconstitution of Fe2Se2 clusters

Note: Iron-selenium protein reconstitution and cluster transfer experiments were conducted in collaboration with Stephen Pearson. All sample synthesis and data acquisition were performed by Jingwei Li and Stephen Pearson.

170

Both hISU (native and D37A) and Fdx2 expression and purification were performed as previously reported.11, 27 The D37A mutant was incubated in 500 mM EDTA pH 8.0 to convert it to its apo form. The resulting apo-protein was dialyzed against a Tris-HCl buffer (50 mM Tris-HCl, 100 mM NaCl, pH 8.6) prior to reconstitution. To be consistent with native hISU, the D37A mutant was reconstituted in the same manner. Cluster reconstitution experiments were performed under argon atmosphere to prevent irreversible cluster oxidation by molecular oxygen. The resulting solution (2 mL of 200 uM) was argon purged following addition of DTT (12.5 mM). Final concentrations of ferric chloride (2.5 mM) and sodium selenide (2.5 mM) were anaerobically added from concentrated stock solutions. The resulting mixture was stirred under anaerobic atmosphere at room temperature for 30 min and the reconstituted protein was loaded on to a gel-filtration PD-10 column purged with argon. The holo protein was eluted with 50 mM Tris-HCl, 100 mM NaCl, pH 8.6. The sulfur derivatives of native and D37A hISU were reconstituted in the same manner, with the only change being the use of sodium sulfide in place of sodium selenide.

To convert Fd2 from its holo to apo form, 2 mL of Fd2 (300 μM) was added to 10 mL of

10 M Urea, 1M EDTA pH 7.4 and incubated at 50oC until the solution became colorless

(about 15 minutes). The protein was then dialyzed against 50 mM Tris-HCl, 100 mM

NaCl, pH 8.6. The apo Fd2 was then reconstituted by incubating apo Fd2 (250 μM) with

DTT (12.5 mM), ferric chloride (7.5 mM), and sodium selenide (7.5 mM) while stirring

171 anaerobically for 30 minutes. The reconstituted protein was then applied to a PD-10 column as stated above. The eluted product was collected in an anaerobically purged glass vial, flash frozen, and stored at -80oC until use when they were thawed under vacuum.

FeSe cluster transfer between glutathione complex and hISU D37A

A 90 µM solution of apo hISU in 500 µL 50 mM Tris, 100 mM NaCl, pH = 8.6 was mixed with 500 μL of 200 μM Fe2Se2 GS4 cluster and the UV/Vis and CD spectrum recorded at 5 min intervals for 120 min. The measured absorbance at were plotted against time. For sulfur derivatives, 330 nm and 433 nm were used for the native and

D37A mutant, respectively. For selenium derivative, 364 nm and 381 nm were used for the native and D37A mutant, respectively. These appropriate wavelengths were chosen based on the peak signal difference between reactants and products. For comparative purposes, CD spectra of sulfur or selenium reconstituted proteins, as well as apo hISU

(both native and D37A) and Fd2 were also recorded, these spectra are listed in Figure 63-

Figure 65.

Cluster transfer from hISU to Fdx2

All protein and reagent solutions were degassed and prepared under an argon atmosphere.

Solutions of reconstituted native (4.4 uM for sulfur derivative, 4.9 uM for selenium derivative) or D37A (6.4 uM for sulfur derivative, 2.6 uM for selenium derivative) hISU in Tris-HCl (50 mM,100 mM NaCl, pH 8.6, 210 μL) was added to apo Fd2 (final

172 concentration of 4.5 uM, 8 mM DTT) and 2.25 mL Tris-HCl 50 mM,100 mM NaCl, pH

8.6. UV/Vis and CD absorption spectra were collected at 5 min intervals over 120 minutes.

Results

Synthesis and characterization

We have recently synthesized a glutathione coordinated iron-sulfur cluster and demonstrated its physiological relevance by monitoring cluster exchange with iron-sulfur cluster assembly protein hISU.28 Since selenium substituted clusters have characteristics similar to the native iron-sulfur derivative,12, 21, 23, 29, 30 we have investigated the synthesis of the glutathione coordinated iron-selenium clusters and its ability to transfer between iron-sulfur cluster proteins. The previously published protocol for synthesis of the iron- sulfur glutathione derivative was used to synthesize and purify the selenium substituted cluster, but it yielded crude products with contamination by free glutathione (Scheme 3).

This contamination was removed by further triturating the pulverized crude product with ethanol. The final product appeared to be a dark brown powder and readily dissolved in water. The absorption profile showed electronic transitions at 360 nm, 415 nm, and 480 nm (Figure 58, left), which were similarly observed with other selenium substituted centers, such as the [2Fe-2Se] reconstituted spinach ferredoxin31, parsley ferredoxin32 and

Chromatium vinosum HiPIP21.

173

Figure 58: [Fe2Se2](GS)4 cluster absorption spectrum (black) in comparison with [Fe2Se2](GS)4 (red). The overall extinction coefficient between the two chalcogenide derivatives were observed to be similar, with a bathochromic shift of transitions was observed for the selenium derivative. Right: kinetic of absorption profile of [Fe2Se2](GS)4 under aerobic conditions. The observed rate constant was 0.071 ± 0.006 min-1. The increased absorbance at 360 nm is due to the hydrolyzed cluster selenium product.

ESI-MS confirmed the identity of the cluster with a base peak m/z value of 1499.1

(Figure 59). Similar to the iron-sulfur glutathione cluster, the selenium substituted cluster was stable under argon atmosphere but aerobically unstable and degraded within 30 min after dissolving in water (Figure 58, right). The mechanism of cluster degradation is likely to be initiated by glutathione oxidation followed by hydrolysis, as cluster degradation was not observed under anaerobic conditions (Figure 60). Previously we have demonstrated that excess glutathione in solution can stabilize the Fe2S2GS4 cluster from degradation, presumably due to the poised reduction potential and solution equilibrium.28 However, since selenide is more susceptible to oxidization than sulfide, the

174 cluster proved unstable in an aerobic environment, even with excess glutathione in solution and therefore all reactions were carried out under strict anaerobic conditions.

Figure 59: ESI-MS spectrum of Fe2Se2GS4 cluster.The base peak is at 1499.1 and + corresponds to C40H67Fe2N12O24S4Se2 , with iron and selenium natural abundant isotopes contributing to a distribution of alternative masses and additional peaks.

175

Figure 60: Cluster stability under various conditions.Spectra were taken after incubation for 1h under various solution conditions. The cluster is observed to be stable only under an argon atmosphere.

Nuclear Magnetic Resonance Spectroscopy

1 The H NMR spectrum of the [Fe2Se2](GS)4 cluster showed similar chemical shifts when compared with the [Fe2S2](GS)4 cluster, where the cysteinyl protons were shifted downfield following coordination to the cluster (Figure 61). As expected, the protons are shifted slightly upfield for the selenium substituted clusters due to the softer character of selenium when compared with sulfur.30, 34 Interestingly, the upfield shift is more pronounced for the glutamate β and γ protons (30 and 50 ppb, respectively) than the cysteine β protons (10 and 20 ppb), presumably due to through-space dipole-dipole interaction contributed by the weak paramagnetism of the antiferromagnetically coupled

176 irons. Nonetheless, the effect of cluster coordination is observed to be significant and differs from free glutathione. These differences have also been observed and reported in works by Holm’s group, where chalcogenide substitutions lead to minor ligand proton chemical shifts. For example, chemical shifts of o-H and p-H of the synthetic cluster

2- [Fe2X2(SPh)] (X = S/Se) were observed to shift upfield in the selenium substituted center.30

1 Figure 61: H NMR spectra comparison of Fe2S2GS4 cluster and Fe2Se2GS4 cluster.The protons in proximity to the cluster center showed upfield shifts.

177

Mössbauer spectroscopy

Mössbauer spectroscopy was used to characterize the coordination environment of iron within the selenium substituted cluster (Table 15, Figure 62). The spectrum exhibits two quadrupole doublets: I (56 %, δ = 0.34 mm/s, ΔEQ = 0.93 mm/s) and II (44 %, δ = 0.25

35 mm/s, ΔEQ = 0.45 mm/s). Species II is within the range of tetrahedral ferric ions and

2+ assigned as oxidized [Fe2Se2] cluster coordinated by glutathione. Similar hyperfine shifts have been observed for selenium substituted iron-sulfur centers in protein, such as

2Fe-2Se cluster in Putidaredoxin.36 By comparing the Mössbauer fit with the FeSe precipitate control experiment, another possible identity of the impurity interaction could be due to iron-selenide precipitate. Species I was also found in a control experiment where GSH was not used (Interaction V, Table 15). This indicates that this interaction is due to a precipitate formed from iron and selenide. Since it is not soluble in water or ethanol, it remained in the sample and consequently observed on the Mössbauer spectrum. This is also observed as non-zero absorption at 800 nm on the UV-Vis absorption profile. Control experiments without selenide resulted in two interactions that were not observed in the cluster spectrum (interactions III and IV, Table 15). These hyperfine parameters are in agreement with octahedral ferric and ferrous coordination environments or sulfur ligands,35 and are likely to be a coordination product by mixing

Fe3+ and GSH. In the presence of DTT, a higher percentage of the product was found to be in the reduced ferrous-GSH product (21% with DTT vs. 18% without DTT). This observation supports a model where the mixture of Fe3+ and GSH, in the absence of Se2-,

178 resulted in a coordination product, which was found to be either oxidized or reduced depending on the solution potential.

In comparison with Mössbauer profiles of iron-sulfur clusters, the iron-selenium glutathione cluster does not exhibit mixed-valent multi-iron centers.36-40 The selenium derivatized clusters yielded characteristics only of oxidized homovalent ferric centers.

The hyperfine parameters for these interactions were observed to be similar for the sulfur and selenide derivatives, both at room temperature in solid form and in solution form at lower temperatures (Table 16). This finding of similar zero-field hyperfine parameters for selenium substituted iron-sulfur clusters agrees with previous reports of protein clusters, such as aconitases41, as well as synthetic “cubane type”clusters.42, 43 These Mössbauer results agree with the 1H NMR results, where the substitution of sulfur with selenium did not significantly change the interactions and bonding characteristic between the ferric iron and its ligands.

179

Figure 62: Mossbauer spectra of the Fe2Se2GSH4 cluster(right) and control experiment of FeSe precipitate (left). These spectra are fit to Voigt line shapes as described in the Materials and Methods section, and the resulting parameters are listed in Table 15. (Data collected in collaboration with C.Y. Chain)

180

Table 15: Mossbauer parameters of the Fe2Se2(GS)4 cluster.and comparison with control experiments without GSH, Se2- or DTT. The experimental procedures for the selenium cluster are documented in the Materials and Methods section. For the control experiments, the concentrations for each reagent were identical to the cluster synthesis conditions, except for the omitted chemical as indicated under each sample description. The corresponding precipitates formed in each case were isolated by centrifugation at 13,000 rpm on table-top microtube centrifuge. These controls were further washed with ethanol (same method as the cluster), and dried under vacuum.

Table 16: A comparison of Mossbauer parameters of selenium substituted cluster with previously published results.

181

[2Fe-2Se] glutathione cluster transfer to apo-hISU

We investigated the ability of this glutathione coordinated iron-selenium cluster to reconstitute the scaffold protein hISU (Scheme 4). The observed rate constants are reported in Table 17 (for raw data, see Figure 66 - Figure 69). In contrast with the native form, the D37A mutant yielded a higher transfer rate from glutathione coordinated cluster to the apo protein. (0.0775 min-1 for D37A vs. 0.0273 min-1 for native hISU) The faster observed rate constant for the selenium derivatives are possibly due to the hydrophobic coordination pocket found in the D37A mutant, which facilitates the cluster transfer of the larger and less electron dense selenide clusters. For comparative purposes, the transfer rates of sulfur derivatives were also studied. For the sulfur derivative, the native form had a faster cluster transfer rate. This can be explained by the role of conserved aspartate residue in the cluster pocket facilitating the cluster transfer as observed before.43 More specifically, Wu et al reported that the conserved aspartate residue to play a key role in the stabilizing the iron-sulfur clusters in hISU protein, and its substitution to alanine resulted in a decrease of cluster transfer to ferrodoxin-1 protein by an order of magnitude.43 Results from the current study with ferredoxin-2 agrees with these previously published observations for hISU reconstituted with iron-sulfur clusters.

182

Table 17: Observed rate constants for the cluster transfer from glutathione coordinated clusters to apo hISU native or D37A proteins.Final concentrations are: 100 uM cluster, 45 uM apo protein and 100 uM DTT. (Data collected in collaboration with Stephen Pearson)

Scheme 4: Cluster transfer between glutathione coordinated complex, hISU and Fd2were studied for Fe2X2GS4 clusters, where X = S or Se. These studied were conducted by measuring the ellipticity on Circular Dichroism Spectroscopy. Rate constants for these transfer reactions are listed in Table 17 and Table 18. Data used to yield these parameters are Figure 63 - Figure 73.

183

-1

-2

-3

-4

300 400 500 600 700 Baseline corrected CD signal (mdeg) signal CD corrected Baseline Wavelength (nm)

Figure 63: CD scans of Fe-S reconstituted proteins and Fe2S2(GSH)4 cluster.Black: 100 uM Fe2S2GSH4; Red: 4.5 uM Fe-S reconstituted native hISU; Blue: 4.5 uM Fe-S reconstituted D37A mutant hISU; Green: 4.5 uM Fe-S reconstituted Fd2. All spectra were recorded in a Tris-HCl buffer (50 mM Tris-HCl, 100 mM NaCl, pH 8.6).

184

2 1 0 -1 -2 -3 -4 -5

300 400 500 600 700 Baseline corrected CD signal (mdeg) signal CD corrected Baseline Wavelength (nm)

Figure 64: CD scans of Fe-Se reconstituted proteins and Fe2Se2(GSH)4 cluster.Black: 100 uM Fe2Se2GSH4; Red: 4.5 uM Fe-Se reconstituted native hISU; Blue: 4.5 uM Fe-Se reconstituted D37A mutant hISU; Green: 4.5 uM Fe-Se reconstituted Fd2. All spectra were recorded in a Tris-HCl buffer (50 mM Tris-HCl, 100 mM NaCl, pH 8.6).

185

2 1 0 -1 -2 -3 -4 -5

300 400 500 600 700 Baseline corrected CD signal (mdeg) signal CD corrected Baseline Wavelength (nm)

Figure 65: CD scans of apo hISU and apo Fd2.Concentrations are: 45 uM apo hISU (native and D37A are red and blue traces, respectively) and 4.5 uM apo Fd2 in green trace. All spectra were recorded in a Tris-HCl buffer (50 mM Tris-HCl, 100 mM NaCl, pH 8.6).

186

Fe S (GSH) to apo WT hISU 0.2 2 2 4

0.0

-0.2

-0.4

-0.6

RawCD signal change (330nm) 0 20 40 60 80 100 120 Time (min)

Figure 66: Transfer of Fe2S2(GSH)4 in solution to apo WT hISU monitored by CD.The signal change at 330 nm was monitored over 120 minutes with scans taken every 5 minutes.

187

2.0 Fe S (GSH) to apo D37A hISU 2 2 4 1.5

1.0

0.5

0.0

-0.5

-1.0

-1.5

CDRaw signal change (433nm) -2.0 0 20 40 60 80 100 120 Time (min)

Figure 67: Transfer of Fe2S2(GSH)4 in solution to apo D37A hISU monitored by CD.The signal change at 433 nm was monitored over 120 minutes with scans taken every 5 minutes.

188

1.00 Fe Se (GSH) to apo WT hISU 2 2 4

0.75

0.50

0.25

0.00

-0.25

RawCD signal change (364nm) 0 20 40 60 80 100 120 Time (min)

Figure 68: Transfer of Fe2Se2(GSH)4 in solution to apo WT hISU monitored by CD.The signal change at 364 nm was monitored over 120 minutes with scans taken every 5 minutes.

189

Fe Se (GSH) to apo D37A hISU 2 2 4

1.5

1.0

0.5

0.0

-0.5 0 20 40 60 80 100 120 RawCD signal change (381nm) Time (min)

Figure 69: Transfer of Fe2Se2(GSH)4 in solution to apo D37A hISU monitored by CD. The signal change at 381 nm was monitored over 120 minutes with scans taken every 5 minutes.

190

[2Fe-2Se] cluster transfer from holo-ISU to apo-Fdx2

The kinetic profile of Fe-Se cluster transfer from holo-hISU to apo-Fd2 was also studied by Circular Dichroism Spectroscopy. The concentration corrected observed rate constant for this step (k2,obs’) are listed in Table 18 (for raw data, see Figure 70 - Figure 73). The resulting k2,obs is similar to cluster transfer rates for iron-sulfur clusters in E.Coli

-1 18, 19 ferredoxin and glutaredoxin systems, typically with kobs < 0.1 min . The observed rate constants for the selenium derivatives are faster than the sulfur derivatives in the case of the D37A hISU, which is similar to the cluster transfer reaction between glutathione cluster and apo hISU. In contrast with the sulfur derivatives, the mutation of the conserved aspartate to alanine had an opposite effect on observed cluster exchange rate for the selenium derivatives. The mutation of the conserved aspartate residue caused the cluster transfer reaction rate to decrease by over 25-fold (see Table 18, sulfur derivatives). A 10-fold rate increase was observed with the alanine derivative. These observations may be explained by similar arguments presented above, where a relatively more hydrophobic cluster environment in D37A hISU may be the reason for the kinetic differences between the sulfur and selenium cluster derivatives.

191

Table 18: Observed rate constants for the cluster transfer from the holo scaffold protein hISU (WT or D37A) to apo Fd2.k2,obs’ was calculated by dividing the observed rate by the initial concentration of holo protein. Concentrations of holo proteins are listed in Materials and Methods. Initial concentration of apo Fd2 was 4.5 μM. (Data collected in collaboration with Stephen Pearson)

-1 Transfer Reaction k2,obs’ (min ) x100

holo [Fe2S2] WT ISU to apo Fd2 2.38 ± 0.48

holo [Fe2S2] D37A ISU to apo Fd2 0.0865 ± 0.0070

holo [Fe2Se2] WT ISU to apo Fd2 0.397 ± 0.015

holo [Fe2Se2] D37A ISU to apo Fd2 3.53 ± 0.76

192

Fe S holo WT hISU to apo Fd2 2.0 2 2

1.5

1.0

0.5

0.0

RawCD signal change (435nm) 0 10 20 30 40 50 60 Time (min)

Figure 70: Transfer of Fe2S2 reconstituted WT hISU to apo Fd2 monitored by CD.The signal change at 435 nm was monitored over 120 minutes with scans taken every 5 minutes.

193

Fe S holo hISU D37A to apo Fd2 0.7 2 2 0.6 0.5 0.4 0.3 0.2 0.1 0.0 -0.1

RawCD signal change (420nm) 0 20 40 60 80 100 120 Time (min)

Figure 71: Transfer of Fe2S2 reconstituted D37A hISU to apo Fd2 monitored by CD.The signal change at 420 nm was monitored over 120 minutes with scans taken every 5 minutes.

194

0.5 Fe Se holo WT hISU to apo Fd2 2 2

0.0

-0.5

-1.0

-1.5

-2.0 0 20 40 60 80 100 120 RawCD signal change (590nm) Time (min)

Figure 72: Transfer of Fe2Se2 reconstituted WT hISU to apo Fd2 monitored by CD.The signal change at 590 nm was monitored over 120 minutes with scans taken every 5 minutes.

195

Fe Se holo hISU D37A to apo Fd2 2 2 1.2

1.0

0.8

0.6

0.4

0.2

0.0

0 20 40 60 80 100 120 RawCD signal change (473nm) Time (min)

Figure 73: Transfer of Fe2Se2 reconstituted D37A hISU to apo Fd2 monitored by CD.The signal change at 473 nm was monitored over 120 minutes with scans taken every 5 minutes.

196

Electrochemistry

Cyclic voltammetry experiments were carried out to study the redox properties of the iron center in the selenium substituted cluster. The results show that there were two distinct irreversible reduction processes. These reduction potentials (Ep,c) were observed at -190 mV and -604 mV vs. NHE. The glutathione clusters substituted with selenium in this study resulted in a higher first electron reduction for the cluster. The second reduction was observed at -604 mV. Due to the window of the measurable reduction potential in aqueous solution, the second reduction event for the iron-sulfur cluster was not observed.

To study the reversibility of the reduction processes for the selenium substituted clusters,

Emin was varied between -478 mV to -778 mV. These studies revealed that unlike the iron-sulfur glutathione cluster, the first reduction process was reversible. The second reduction process was not reversible (Figure 74, right), as observed by change of returning wave currents in cycles with Emin of -678 mV. The breakdown of cluster is especially evident in the Emin = -778 mV scan, with more of the cluster observed to precipitate out of solution.

197

A)  5 0

0 E = -778 mV

min Current ( Current -50

-5

-800 -600 -400 -200 0 200 400 -100 Potential (mV vs. NHE)

 E = -678 mV min -150

Current( -200 15

E = -578 mV

10 min A)

 -250

Current ( Current 0 -300

E = -478 mV -5 min -350 -800 -600 -400 -200 0 200 400 -1000 -500 0 500 1000 1500 Potential (mV vs. NHE) Potential (mV vs. NHE)

Figure 74: Cyclic voltammetry experimentsshow that the selenium cluster (bottom left) displays two distinct irreversible reduction process were observed in the Fe/Se cluster. One of the reduction occurs at a more negative reduction potential (-604 mV) than the sulfur derivative (top left, -305 mV), while the other reduction potential (-190 mV) is lower than the sulfur derivative. Emin was varied to determine the reversibility of the reduction process (right).

198

Cyclic voltammetry experiments of sulfur and selenium reconstituted hISU (native and

D37A) and Fd2 were conducted and results are listed in Table 19. Overall, general trends of more negative reduction potentials were observed for the selenium derivatives. These trends were also observed for the ferredoxin-1 protein (-274 mV and -288 mV for sulfur and selenium derivatives, respectively),44 as well as other selenium substituted 2Fe-2S proteins such as putidaredoxin from P. putida.45 These observations are consistent with the general relationship between electronegativity and reduction potentials.46, 47

Furthermore, these electrochemical data agree with the Mossbauer spectroscopy results in that the Fe electron densities were found to be higher in the selenium derivatives. For example, the reduction potentials for iron-selenium centers were found to be more negative than sulfur derivatives. The isomeric shift (δ), another spectroscopic measurement of electron density of the iron atom, was also found to be higher for the selenium derivative than the sulfur derivative. These results collectively indicate a higher electron density in the [Fe2Se2] core, which may result in a longer Fe-S(cys) distance relative to the sulfur centers. In this way, the [Fe2Se2] cluster coordinated by glutathiones may be intrinsically less thermodynamically stable, resulting in a faster cluster transfer to apo-hISU. Interestingly, previous studies have indicated that the activation barrier for cluster transfer reactions is dictated by both enthalpy and entropy contributions.44, 49, 50

Therefore, the selenium substituted glutathione cluster with higher iron electron density and longer Fe-S(cys) bond therefore lower bonding enthalpy, may explain the reactivity of cluster transfer with apo-hISU proteins.

199

Table 19: Redox potentials determine by cyclic voltammetry for sulfur and selenium reconstituted proteins. (Data collected in collaboration with Stephen Pearson)

o E 1/2 (mV) Fe2X2 Fd2 Fe2X2 WT ISU Fe2X2 D37A hISU X = S -103.5 -17.7 -34.7 X = Se -133.3 -27.6 -50.4

200

Conclusion

A selenium substituted glutathione iron sulfur cluster has been synthesized and characterized. The cluster behaved similarly to the native sulfur derivative when incubated with physiologically relevant proteins such as the cluster scaffold protein hISU.

Interestingly, Fe2Se2 clusters were also observed to be transferred between holo-hISU to apo-Fd2. These transfers were monitored by Circular Dichroism Spectroscopy. When

[Fe2Se2](GS)4 cluster is incubated with apo-hISU D37A derivative, the absorbance at 360 nm was also observed to increase and the spectrum was fit to yield a time constant of

39.5 ± 1.8 min (Figure 75).

1.10 )

1.05 470

1.00

0.95

0.90

0.85

0.80

0.75

Normalized Intensity (Abs Intensity Normalized 0 20 40 60 80 100 120 Time (min)

Figure 75: Cluster transfer from glutathione cluster to D37A apo hISU monitored by UV/ Vis spectroscopy. The extinction coefficient for the holo protein is lower than that of the cluster, and the absorbance change was observed to decrease over time as the cluster is transferred. The apparent rate constant is 0.010 ± 0.005 min-1M-1, which is slower than the iron sulfur cluster transfer rate constant observed previously.43 201

The additional kinetics observed by time dependent UV-Vis spectroscopy experiments agrees very well with results presented in Table 17. Iron-selenium cluster formation by the iron-sulfur cluster synthesis machinery has been previously documented by

Hallenbeck et al. in the Azotobacter vinelandii system using selenocysteine.22 However, iron-selenium cluster transport between iron-sulfur cluster proteins has not been previously demonstrated. The rate of cluster transport is similar to holo-hISU/apoferredoxin in E. coli systems. These findings suggest that similar to the iron-sulfur clusters, the selenium substituted clusters interacts with glutathione molecules and is able to be transported between cluster proteins. Stadtman et al. reported natural selenium/molybdenum clusters are the catalytic center in Clostridium barkeri nicotinic acid hydroxylase.48 Furthermore, there have been several reports which hypothesized that selenium content in laboratory diet of rats may be incorporated as selenide derivatized proteins and protect against oxidative destruction in vivo by vitamin E.49, 50 Together with theoretical and computational modeling results,51 it is logical to hypothesize the possible existence of natural occurring iron selenium cluster proteins, which may possess important physiological roles in biological systems.

These results demonstrate physiologically relevant exchange reactions between iron- chalcogenide cores and glutathione versus protein ligands. Assembly of [2Fe-2X] (X =

S/Se) clusters via scaffold protein ISU provide a pathway to a cellular pool of glutathione-complexed cluster, as well as provides a mechanism for biosynthesis and transfer of [2Fe-2Se] cores to Se-dependent proteins enzymes.

202

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[45] Mukai, K., Huang, J. J., and Kimura, T. (1974) Studies on Adrenal Steroid Hydroxylases - Chemical and Enzymatic Properties of Selenium Derivatives of Adrenal Iron-Sulfur Protein, Biochim Biophys Acta 336, 427-436.

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[46] Wilson, G. S., Tsibris, J. C. M., and Gunsalus, I. C. (1973) Electrochemical Studies of Putidaredoxin and Its Selenium Analog, J Biol Chem 248, 6059-6061.

[47] Song, I. K., and Barteau, M. A. (2004) Redox properties of Keggin-type heteropolyacid (HPA) catalysts: effect of counter-cation, heteroatom, and polyatom substitution, J Mol Catal a-Chem 212, 229-236.

[48] Vanysek, P. (2000) Electrochemical Series, 95th ed. ed.

[49] Gladyshev, V. N. K., S.V.Stadtman, T.C. (1994) Nicotinic acid hydroxylase from Clostridium barkeri: electron paramagnetic resonance studies show that selenium is coordinated with molybdenum in the catalytically active selenium-dependent enzyme, Proc. Natl. Acad. Sci. USA 91, 232-236.

[50] Giasuddin, A. S. M., Caygill, C. P. J., Diplock, A. T., and Jeffery, E. H. (1975) Dependence on Vitamin-E and Selenium of Drug Demethylation in Rat-Liver Microsomal Fractions, Biochem J 146, 339-350.

[51] Diplock, A. T., and Lucy, J. A. (1973) Biochemical Modes of Action of Vitamin-E and Selenium - Hypothesis, Febs Lett 29, 205-210.

[52] Turker, L. E., S. (2003) A theoretical study on certain iron-sulfur and iron-selenium clusters, J. Mol. Struct. 623, 17-21.

207

Chapter 6: Glutathione-Coordinated [2Fe-2S] Cluster as a Viable Physiological Substrate for Mitochondrial ABCB7 Transport

Introduction

Eukaryotic cluster assembly involves a pathway based on proteins in the bacterial ISC operon, and both cytosolic and nuclear iron-sulfur clusters are dependent on mitochondrial iron sulfur cluster assembly.1, 2 The exact mechanism of how the mitochondrial and cytosolic iron-sulfur cluster assembly pathways are associated remains unclear.3-6 Atm1p is a yeast homolog of the ABC7-type ATP-dependent mitochondrial membrane transporter that is linked both to mitochondrial iron homeostasis,7-10 and the regulation of cytosolic iron concentration.11-14 Deletion of the human homolog leads to severe iron-sulfur cluster deficiency in the cytoplasm, but not in mitochondria.6, 15-19 In humans, natural mutants of the transporter have been identified in patients affected with

X-linked sideroblastic anaemia and cerebellar ataxia.20 Key roles in cytoplasmic iron- sulfur cluster assembly have also been reported in A. thaliana for other Atm1 family proteins, indicating their importance in the plant kingdom,21 and a role suggested in heavy metal detoxification by a bacterial homolog of Atm1p.22 While the identity of the transporter substrate remains uncertain, glutathione (GSH) has been implicated.18, 19

Previously, contributions from the Lill group have shown that glutathione stimulate the 208

ATPase activity of Atm1p/ABC7.23 The involvement of glutathione in mitochondrial cluster export is further supported by its role in the maturation of cytosolic iron-sulfur cluster proteins, but not mitochondrial cluster proteins. These observations are also consistent with a close genetic relationship between ATM1 and GSH1.19 Therefore, glutathione is likely a key component in the mitochondrial iron-sulfur cluster export mechanism.

In this chapter, glutathione complexed [2Fe-2S] clusters are used as substrates for biochemical studies with Atm1p to test its candidacy as a natural substrate for the transporter. Given the evidence implicating glutathione in cluster export and neither a bare cluster core, nor a protein-bound cluster, are likely substrate candidates for the exporter, glutathione coordinated iron-sulfur cluster complexes are viable substrate candidate for the ABC7-type transporter. A wide range of techniques were used for this purpose, including substrate stimulated ATPase activity assay, protein structure computational modelling (In collaboration with Wenbin Qi), flow cytometry, and tiron chelation assays for proteoliposome models. In particular, evidence of the ability of

2- Atm1p to transport [2Fe-2S](GS)4 by use of independent flow-cytometric methods and tiron absorbance assays in a model proteoliposome complex are presented. These results also allowed quantification of cluster transport activity and determination of kinetic rate constants. These experiments provide support for the physiological relevance of glutathione-complexed [2Fe-2S] clusters, demonstrate the viability of such species as

209 natural transporter substrates, and present a quantitative methodology for study of metal translocation proteins and their proteoliposome products.

In previous chapters, glutathione Fe/S cluster complex was demonstrated to be stable in the presence of physiological glutathione concentration. Studies done by Wenbin Qi also showed that this complex is able to exchange cluster with iron-sulfur cluster scaffold protein.24, 25 These results suggest that this complex is a potential cluster carrier in a cellular environment. Results from this chapter provides evidence in support of [2Fe-

2S](GS)4 as a likely iron-sulfur cluster substrate for the Atm1p/ABC7 transporter in both solution and proteoliposome-bound forms, and identify a likely substrate binding site on the transporter.

210

Materials and Methods

Molecular cloning

Molecular cloning of the recombinant yeast Atm1p to pASK-IBA2 was done by Wenbin

Qi, and detailed procedures are documented in his dissertation thesis.26 Atm1p R284E mutant was obtained by using the standard site directed mutagenesis protocol. In brief, mutation primers were obtained from IDT DNA Inc. and are:

TGGAGGACACATTTTGAAAGGGATGCTAACAAGGC, and TTGTTAGCATCCCT

TTCAAAATGTGTCCTCCA. Experimental conditions such as concentration, volume and PCR time segments are listed in Table 20 and Table 21, respectively. Transformation of the resulting mutant plasmid and the subsequent protein purification were performed with the same protocols as described for the wild-type protein.

211

Table 20: Concentrations and volumes of components for the PCR mutagenesis of wildtype Atm1p to R284E.

Table 21: Reaction times and temperatures for the PCR mutagenesis of wild-type Atm1p to R284E.

212

ATPase Activity assay

The ATPase activity of purified Atm1p was monitored by using EnzChek Phosphate assay kit (Invitrogen). In brief, protein was mixed with 200 µM MESG substrate, 1 unit purine nucleoside phosphorylase, 1 mM ATP, 1 mM Mg2+ in 1x reaction buffer.

Formation of product (ribose 1-phosphate and 2-amino-6-mercapto-7-methylpurine) was monitored at 360 nm (Figure 76). Initial rates of reaction were also measured with increasing concentrations of ATP, and a Michaelis-Menten plot of Atm1p ATPase activity of was constructed (Figure 77).

Figure 76: An overview of the ATPase activity assay.Inorganic phosphate reacts with MESG in the presence of PNP, and yield a product which absorbs at 360 nm.

213

Data Fit 2.5x10-6

2.0x10-6

) -1 1.5x10-6

V = 2.32 +/- 0.03 uM/min MAX -6 K = 54.6 +/- 0.4 uM 1.0x10 m

5.0x10-7 Initial Velocity (M*min Velocity Initial

0.0

0.0 0.5 1.0 1.5 2.0 [Mg-ATP] (mM)

Figure 77: Atm1p and Mg-ATP binding parameters.Data were fit to standard Michaelis- Menten equation to yield Vmax and Km values. The solution conditions are: [protein] = 1.2 uM, [MESG] = 200 uM, 1 unit of purine nucleoside phosphorylase and in 1x reaction buffer (60 uL).

214

Protein Purification

For the over-expression of protein, cells were grown in LB media (100 mg/L ampicillin) to an OD550 of ~ 0.6 prior to induction with 20 µg/L anhydrotetracycline over night. After the cells were harvested, Atm1p purification was done either following the Strep-tag column method established by Wenbin Qi,26 or by column chromatography via MonoS then Superose-12 FPLC method described herein.

For the Strep-tag method, cells were suspened in 50 mM Trish-HCl pH 8.0 buffer with 75 mM NaCl. The cells were then incubated on ice with 10 mM EDTA and 1 mg/mL for 30 minutes. Then, cells were lysed for 10 seconds every minute for 10 minutes, and cell debris was removed by centrifugation for 30 min at 5000 rpm. Ultracentrifuge was used to pellet the membrane fraction of protein at 86,000 rpm. The pellet was dissolved in 20 mM MOPS-KOH at pH 6.5 with 200 mM NaCl and 20% Sorbitol. A stock solution of n- dodecylmatoside was used to introduce a final concentration of 0.5 %. The resulting solution was incubated on ice for 30 min, then centrifuged at 86,000 rpm to remove insoluble impurities. The final supernatant containing Atm1p was diluted equal volume with 100 mM Tris-HCl at pH 8.0 with 150 mM NaCl, 1 mM EDTA, 10 % Sorbitol and

0.025% n-dodecylmaltoside. A StrepTactin (IBA-LifeSciences) column was then used to selectively bind and elute Atm1p.

For the MonoS/Superose-12 purification method, cells were incubated on ice with 50 mM HEPES buffer at pH 7.5 with 10 mM EDTA, 1 mg/mL lysozyme for 30 minutes.

215

Cells were lysed and the supernatant was obtained with the same method as described above. The supernatant was then applied to FPLC equipped with an ion-exchange MonoS column. Protein was eluted with a 50 mM HEPES buffer at 7.5 and linear NaCl gradient

(0 – 400 mM). Fractions showed ATPase activity were further purified with a Superose-

12 gel-filtration column to isolate Atm1p. The purity of Atm1p was confirmed by SDS-

PAGE method (Figure 78).

216

Fraction 1 0.040 Fraction 2 Fraction 3 Fraction 4

) 0.035 -1

0.030

0.025

0.020

0.015

0.010

ATPase activity (initial slope min slope (initial activity ATPase 0.005

0.000

Figure 78: Purification of Atm1p.(top) FPLC elute fractions from MonoS column. (bottom left) FPLC elute fractions from Superose-12 column, with silver-staining to show Atm1p in protein fraction #1. (bottom right) ATPase activity assays indicate Atm1p is in protein fraction #1 of the elute.

217

In agreement with previous findings, Atm1p (and mutants) appears as a single band near

66 kDa. This is consistent with reported observations for Atm1p, although the MW of the protein is 76 kDa.23, 26 The yield of Atm1p was observed to significantly increase when purified at 4 ºC (with FPLC system in the coldbox). With 12 g of harvested cells, 2.6 mg

Atm1p was obtained when the protein was purified at 4 ºC (Figure 79 and Figure 80).

Figure 79: SDS-PAGE gel of Atm1p.Protein fraction were collected at 1 mL/min and 8 min/tube. Atm1p was observed in fraction 15. With purification done in 4 ºC, 0.2 mg protein was obtained per gram of cell pellet, and the resulting protein fractions were visible on the SDS-PAGE gel stained with Coomassie Brilliant Blue stain.

218

Figure 80: SDS-PAGE of WT Atm1p and mutants.Gel was stained by Coomassie Brilliant Blue.

The stability of purified Atm1p is low, with precipitation observed after days stored in the 4 ºC coldbox. According to the Strep-tag purification protocol established by Wenbin

Qi, the final step included addition of a solution detergent which may prolong the stability of Atm1p when stored in the coldbox. Therefore, a final concentration of 0.025% n-dodecylmaltoside was added to the eluted protein after purification. With detergent in solution with Atm1p, its stability significantly increased, and precipitation of protein was not observed. Furthermore, the ATPase activity of Atm1p was not influence by 0.025%

219 n-dodecylmaltoside, as shown in Figure 81. The ATPase activity of Atm1p was also found to be depended on its concentration.

Figure 81: ATPase activity of purified Atm1p.Added n-dodecylmaltoside did not influence the activity, but it was observed to stabilize the purified protein, and prevent it from crashing out of solution. The ATPase activity also depends on the concentration of Atm1p (magenta vs. brown).

Synthesis of liposomes

DOPG (Dioleoylphosphatidyl-Glycerol), DOPC (Dioleoylphosphatidyl-Choline), and

DOPE (Dioleoylphosphatidyl-Ethanolamine) were purchased from Avanti Polar Lipids,

Inc. and dissolved to 15 mM stock concentrations. An equimolar mixture of DOPG, 220

DOPC and DOPE (133 μL, 131 μL and 124 μL, respectively) were vortexed and dried over Argon gas. The resulting residue was suspended in 1 mL 50 mM HEPES, 100 mM

NaCl buffered at pH 7.5 and extruded through a 400 nm membrane 21 times. The quality and size of the lipid was checked by Dynamic Light Scattering Spectrometry (Malvern

Instruments) and kept at 4 ºC until used. The same liposome synthesis method was followed for fluorescein-encapsulated liposomes, except that the re-suspension HEPES buffer contained 1 mM fluorescein.

Figure 82: Lipids were mixed at a molar ratio of 1:1:1 to synthesize lipid membrane.

Synthesis of Atm1p proteoliposomes

The following procedure for proteoliposome synthesis is based on a modified protocol originally established by Poolman et al.27, 28 The previously synthesized liposome was

221 diluted 2-fold to a total volume of 2 mL. A stock solution of 10% Triton X was titrated into the liposome solution at 2 μL increments until the OD550 reached a maximum, and an additional 16 μL was added to reach the “loose” state (Figure 83). It proved to be important to maintain the liposome on ice during the Triton X addition. Subsequently,

Atm1p protein (80 μL, 1.2 μM) that had been purified by use of a protocol established by

Qi et al.,29 was added to the loosened liposome and incubated on ice for 15 min. Biobeads were used to remove the Triton X detergent and restore the rigidity of the proteoliposome. Aliquots of 40 mg Biobeads were added at 0, 30, 60 min and overnight.

The Biobeads were removed by gentle centrifugation at 3,000 rpm for 1 min and separated from the decantate. The reconstituted proteoliposome was removed from the buffer by ultracentrifugation at 80,000 rpm for 20 min and re-dissolved in 1 mL 50

HEPES, 100 mM NaCl buffered at pH 7.5. The resulting proteoliposome was kept on ice and typically used within 24 hours of synthesis. ATPase activity assay was performed with the reconstituted proteoliposome , shown in Figure 84. The solution conditions are: total volume = 100 uL, 1 mM Mg-ATP, 0.1 unit PNP, 1x reaction buffer, 200 uM MESG, and 72 uL proteoliposome.

222

Figure 83: Proteoliposome synthesis.(top) incorporation of Atm1p into liposome using Triton-X100. Addition of detergent increases the fluidity of the membrane bilayer. Protein is added when the absorbance is less than the initial value with no detergent added (circled). (bottom) For incorporation of fluorescein in the proteoliposome, a fluorescein solution was used to ensure the concentration of fluorescein stayed constant during treatment with Biobeads.

223

Figure 84: ATPase activity of reconstituted proteoliposome.A control with a delayed addition of Mg-ATP was performed to show that the first few minutes of the activity may be due to impurities in solution, as observed by the initial increase followed by a slower rate exhibited by Atm1p.

Computational Modeling

Computational modeling was performed in collaboration with Wenbin Qi. Model structures for human ABCB7 and yeast Atm1p were generated by SWISS-Model,30 and

PQR files generated by PDB2PQR from the modeled PDB file by use of the PDB2PQR server (http://nbcr-222.ucsd.edu/pdb2pqr_1.8/).31 The electrostatic potential maps were then calculated using the generated PQR file and APBS software.4,32 The modeled

Atm1p were visualized by Chimera.33

224

Synthesis of fluorescein-labeled glutathione

Oxidized glutathione (GSSG, 25.88 mg, 42 μmol) was dissolved in 0.8 mL 100 mM sodium bicarbonate buffered at pH 8.0. A solution of 5-(and 6-) carboxyfluorescein succinimidyl ester (NHS-Fluorescein, 10 mg, 21 μmol), dissolved in 0.1 mL DMSO, was added drop-wise to the buffered GSSG. The solution was allowed to stir at r.t. for 2 hours in the dark. After reaction, the disulfide bond was reduced by addition of 0.1 mL TCEP

(26.4 mg, 0.106 mol) dissolved in bicarbonate buffer, and stirred at r.t. for 10 min. The pure product (Fl-GSH) was purified from the crude mixture by HPLC (Agilent 1100) equipped with a reversed-phase C-18 column (Gemini 100 x 21.20 mm 5 micron) and eluted with buffers A: nano-pure H2O, 0.1% TFA, and B: Acetonitrile, 0.1% TFA. The retention time of the product was 37 min with a flow rate of 5 mL/min and gradient of

+1% B/min. The resulting fractions were dried over vacuum and stored as a powder until used. The yield of the final fluorescein-GSH conjugate (Fl-GSH) after reduction by

TCEP was >90% and characterization by ESI-MS demonstrated a product peak with m/z

+ of 666.1 that corresponded to Fl-GSH (C31H28N3O12S ).

225

Figure 85: ESI-MS analysis of Fl-GSH.

226

Synthesis of fluorescein-labeled cluster

Fl-GSH was used to make fluorescein-labeled cluster by combining labeled and unlabeled glutathione in a ratio of Fl-GSH : GSH of 1:20 in order to limit the number of

Fl-GSH molecules incorporated per equivalent of cluster to one labeled glutathione molecule. The singly-labeled product appears to be the limiting form produced, most likely reflecting the fact that modification of two GSH yields an unstable cluster, as a result of disruption of stabilizing salt bridges,34 and so multiply-modified derivatives are not observed.

A solution of glutathione (19.3 mg, 62.9 μmol) was dissolved in 1 mL nano-pure H2O and purged under argon by 3x freeze-thaw cycles. The pH of this solution was adjusted by addition of stock NaOH solution (5 M, 16.5 μL) to a final pH of 8.6. The mixture of

GSH and Fl-GSH was made by mixing 20 μL of the GSH solution with 80 μL of purified

Fl-GSH dissolved in H2O. Solutions of ferric chloride (0.0217 g in 3 mL H2O) and sodium sulfide (0.0321 g in 3 mL H2O) were made fresh prior to synthesis and added to the GSH solution (12.5 μL each). The resulting mixture was stirred and purified with the

2- same protocol for [2Fe-2S](GS)4 cluster as previously described. The yield of the final fluorescein-GSH conjugate (Fl-GSH) after reduction by TCEP was >90% and characterization by ESI-MS demonstrated a product peak with m/z of 1868.6 that

- corresponded to a cluster with one fluorescein label (C61H70Fe2N12Na5O30S6 ).

227

Flow cytometry analyses of proteoliposomes

Liposomal flow cytometry experiments were conducted on a FACS Calibur Flow

Cytometer (BD Biosciences) equipped with 488 nm / 633 nm lasers and dichroic filters at the University Cell Analysis and Sorting Core (UCAS, OSU). A detection filter setting of FL2 (585/42 nm) was used for detection of rhodamine-labeled liposomes and FL1

(530/30 nm) was used to detect fluorescein labeled iron-sulfur cluster. For detection of liposomes, Forward (FSC) and Side (SSC) scattering detector voltages were set at 8.68 V and 5.08 V, respectively. Compensation of the dichroic filters was set at FL1 – 10 % FL2 to remove false positive fluorescence signals. Samples were injected into the instrument at low speed setting and recorded until a final count of 10,000 events. The resulting data was processed on Cell Quest Pro (BD Biosciences) and plots were made as FSC vs. SSC and FL1/2 vs. event count without post-acquisition smoothing.

The fluorescence response from the labeled proteoliposomes was observed to change over the incubation period due to cluster transport into the vesicle (Figure 86). A solution of freshly made proteoliposome (1.0 mL in 50 mM HEPES and 100 mM NaCl at pH 7.5) was mixed with Mg-ATP (200 μL, 75 μM) and cluster (13 mM) in GSH solution (10 μL,

150 mM, pH 8.6) and incubated at 25 ºC for 60 min. To determine the kinetic profile for cluster transport, reactions were started at 10 min intervals up to 50 min. Normalized intensities (average + standard deviation) for experiments with cluster and relevant controls were plotted and shown in Figure 98 and Figure 100. Initial slopes were fit to the first 30 min of data for most samples (except for the more rapid cluster transport, which

228 was fit to the initial 10 min) and initial velocities (average + standard deviations) were plotted in Figure 99. The integrity of the proteoliposome during the experimental conditions described in the main text was checked by use of dynamic light scattering spectroscopy. The measured Z-average and count/s were plotted (Figure 87), and the proteoliposomes were observed to be stable.

Figure 86: Raw flow cytometry datashowing proteoliposomes fluorescent signal changes over time for the experiment and control without Atm1p. FlowJo was used to quantitatively obtain the geometric mean for each experiment.

229

Liposome + MgATP + Cluster/GSH 200 Proteoliposome + MgATP + Cluster/GSH

180

160

Z-average (nm) Z-average 140

120 0 10 20 30 40 50 60 Time (min)

Liposome + MgATP + Cluster/GSH 1.2x108 Proteoliposome + MgATP + Cluster/GSH

1.0x108

8.0x107

6.0x107

counts/s 4.0x107

2.0x107

0.0 0 10 20 30 40 50 60 Time (min)

Figure 87: Control experiments to monitor the integrity of the proteoliposomes throughout the transport reaction.Both the size (top) and number (bottom) of the liposomes remained relatively unchanged over the course of the reaction, indicating that the fluorescence change observed during active transport reflects perturbation of the inner content of the proteoliposome. Synthesized proteoliposomes were dissolved in 50 mM HEPES and 100 mM NaCl buffered at pH 7.5. Data were collected on DLS (Malvern Instruments) at room temperature.

230

For studies of fluorescein-labeled cluster transport, a solution of freshly made proteoliposome (1.0 mL in 50 mM HEPES and 100 mM NaCl at pH 7.5) was mixed with

Mg-ATP (200 μL, 75 μM) and fluorescein-cluster (13 mM) in GSH solution (100 μL,

150 mM, pH 8.6) and incubated at 25 ºC for 60 min. Data points were gated with both

FSC and SSC > 20 to remove background noise and yield the overall count for the fluorescent proteoliposomes (Figure 101).

Tiron Assay

A solution of freshly made proteoliposome (0.2 mL in 50 mM HEPES and 100 mM NaCl at pH 7.5) was mixed with Mg-ATP (40 μL, 75 μM) and cluster (13 mM) in GSH solution (20 μL, 150 mM, pH 8.6) and incubated at 25 ºC for 60 min, and the product mixture was centrifuged at 6,000 rpm for 5 min to remove precipitate formed from cluster hydrolysis. The resulting decantate was removed and centrifuged at 80,000 rpm for 20 min to isolate the proteoliposome. The pellet was resuspended with 100 μL 50 mM

HEPES and 100 mM NaCl buffered at pH 7.5. The proteoliposome was denatured by adding concentrated HCl (30 μL, 12 M) to liberate the encapsulated iron. Addition of

MES buffer (10 mM, pH 6.5) and NaOH solution (5 M, 692 μL) was used to neutralize the reaction. A stock tiron solution (100 mM, 100 μL in MES buffer) was added to chelate the released ferric ions and incubated at room temperature for 10 min. The absorbance at 550 nm was measured by use of a UV/Vis spectrophotometer (Varian T-

50) and plotted against a standard curve to calculate the iron concentration inside the proteoliposome. The overall scheme for this assay is summarized in Figure 88 and

231

Figure 89. Control experiments with GSH, GSSG and iron were carried out using similar concentrations of these reagents and the results are shown in Figure 104.

Figure 88: Synthesis of proteoliposome, ATP-driven cluster transport and ferric ion quantitation of product.

232

Figure 89: Tiron coordinates to ferric ions, and absorb at 550 nm.The concentration of -1 -1 iron is then calculated with ε550 = 32,000 M cm .

Titration of fluorescein-labeled cluster to Atm1p

Labeled cluster from a stock solution (1 mM) was added to a solution of Atm1p (1.2 uM) in HEPES buffer (50 mM HEPES, 2 mM GSH, pH 7.5). The equivalent control experiment without Atm1p was used to calculate the net signal change, and these results are graphed in Figure 90, with a fitted KD = 118 ± 11 μM.

233

-50

-100

-150

Fluorescence Signal Fluorescence  -200 0 50 100 150 200

[Fl-cluster] (uM)

Figure 90: Titration experiment showing the decrease in fluorescence signal intensity as labeled-cluster was added to Atm1p.Labeled cluster from a stock solution (1 mM) was added to a solution of Atm1p (1.2 uM) in HEPES buffer (50 mM HEPES, 2 mM GSH, pH 7.5). The equivalent control experiment lacking Atm1p was used to calculate the net signal change, and these results are graphed and fitted yield a KD = 118 ± 11 μM.

Titration of GSH to a solution of fluorescein

Addition of GSH quenches the fluorescence response from fluorescein observed in a cuvette measurement. GSH from a stock solution (10 mM) was added to a solution of

Fluorescein (10 uM, 1 mL) in HEPES buffer (50 mM HEPES, 100 mM NaCl, pH 7.5).

These results are graphed in Figure 91.

234

0.00

-0.01

-0.02

-0.03

Normalized Fluorescence Unit Fluorescence Normalized 0.0 0.5 1.0 1.5 2.0  [GSH] (mM) f

Figure 91: Titration experiment showing GSH quenching the fluorescein fluorescence signal, most likely due to a heavy atom effect. A solution of fluorescein (10 uM , 1 mL) was titrated with a stock solution of glutathione (10 mM) in Hepes buffer (50 mM Hepes, 100 mM NaCl, pH 7.5).

Data fitting and Statistical analysis

For the control data shown in Figure 99, the first thirty minutes of the flow cytometry signal was fit to linear functions to obtain the observed initial rate of decrease. Standard p-test was performed for GSH control against experiments with clusters, which yielded p

= 0.0070 and z = 2.70, corresponding to a 99.3% level of confidence.

235

Results

Atm1p ATPase Activity Assays

Atm1p was purified as described in the Materials and methods section. Its identity was confirmed by both SDS-PAGE analysis and ATPase activity assay (Figure 78, Figure 79,

Figure 80 and Figure 81). When the concentration of Mg-ATP was varied, a Michaelis-

Menten type kinetic was observed for Atm1p ATPase activity. The Michaelis-Menten parameters determined from the plot were KM = 54.6 ± 0.4 uM and Vmax = 2.32 ± 0.03 uM/min. This is similar to previously determined values for Atm1p and other ABC transporters.26, 35 Substrated binding often causes ABC proteins to undergo conformation change, and consequently alters its ATPase activity.36, 37

With glutathione, but no cluster complex present, the rate of phosphate formation increases (Figure 92), which is consistent with previous studies that have shown that glutathione can stimulate the ATPase activity of Atm1p.23

236

Data Fit 2.2x10-6

2.2x10-6

) -6 -1 2.1x10

2.1x10-6

2.0x10-6

2.0x10-6 Initial Velocity (M*min Velocity Initial

2.0x10-6

1.9x10-6 0 1 2 3 4 5 [GSH] (mM)

Figure 92: Glutathione stimulates the ATPase activity of Atm1p, consistent with previously reported assays.

With additional concentrations of glutathione iron-sulfur cluster, the ATPase activity of

Atm1p was observed to increase. The dependence of activity on cluster concentration

(Figure 93) was fit to a nonessential activation model (equation 8),38

[S] [A][S] max β max KM αKDKM = [S] [A] [A][S] - c A Equation 8 1 KM KD αKDKM

where Vmax is the maximum initial ATPase activity of Atm1p in the absence of cluster,

[S] is the concentration of substrate Mg-ATP, [A] is the concentration of cluster stimulant, KD is the binding constant of the cluster to Atm1p, KM is the binding constant

237 of Mg-ATP to Atm1p, α accounts for the modification of KM by cluster, and β accounts for Vmax stimulation by cluster.

Figure 93: WT Atm1p ATPase activity is stimulated by cluster(left). The negative slope at higher cluster concentrations reflects contributions from cluster degradation and is also observed at 360 nm. Data is fit to eqn.8 and fitted parameters are listed in Table 22. R284E mutant data is shown in the right plot. The mutant retained its ATPase activity, but not stimulated by cluster.

The data illustrated in Table 22 demonstrates the glutathione cluster complex to serve as a modifier that increases the velocity of Atm1p-catalyzed phosphate formation by 1.9- fold and decreases the KD 0.6-fold, which supports the hypothesis that the glutathione iron-sulfur cluster is a likely substrate for this transporter in a manner consistent with previous genetic interaction and knock-out studies.19 The cluster complex shows saturation binding to the transporter with a measured KD of 68 uM.

238

Table 22: Parameters for [2Fe-2S](GS)4 and glutathione stimulation of transporter ATPase activity.Parameter definitions: Vmax, ATPase activity in the absence of cluster; S, [Mg-ATP]; KM: Michaelis-Menten constant for Mg-ATP; β, an activity multiplier reflecting the stimulation of Vmax by cluster; α, a modifier of KM reflecting the impact of cluster on Mg-ATP binding; KD, dissociation constant for cluster; c, rate of cluster degradation.

Stimulant cluster GSH R284E

Vmax (uM/min) 2.19 ± 0.04 2.07 ± 0.02 2.50 ± 0.08 S (mM) 1.00 ± 0.01 1.00 ± 0.01 1.00 ± 0.01 KS (uM) 54.6 ± 5.3 54.2 ± 8.9 55.0 ± 16.5 β 1.85± 0.05 1.14 ± 0.01 0.86 ± 0.02 α 0.55 ± 0.06 0.87 ± 0.14 0.68 ± 0.05 KD (uM) 68 ± 2 689 ± 215 1820 ± 660 c (hr-1) 0.020 ± 0.004 - 0.004 ± 0.001

3.8 3.6 3.4

M/min) 3.2  3.0 2.8 2.6 2.4 2.2

Initial Velocity ( Velocity Initial 2.0 0 20 40 60 80 100 [Cluster] (M)

Figure 94: Atm1p ATPase activity is stimulated by [2Fe-2S](GS)4.The data are corrected for magnesium-induced cluster degradation and fit to eqn.7 to yield the fitted parameters listed in Table 22. Solid squares (■), native Atm1p; vacant squares (□), R284E Atm1p.

239

The relative KD’s for [2Fe-2S](GS)4 and glutathione indicate a much higher affinity for the cluster complex (68 uM versus > 689 uM, respectively). Prior observation of very modest levels of stimulation of Atm1p/ABC7 ATPase activity by glutathione are consistent with a glutathione cluster as a natural transporter substrate, with the more modest levels of stimulation reflecting weaker intrinsic binding to the transporter (Table

22), as a result of partial occupation of some of the contact sites on the transport protein occupied by the full tetrameric glutathione complex cluster. No stimulation of ATPase activity by cluster was observed when R284 was substituted with Glutamate, although full ATPase activity was retained. To further study and understand the role of this important residue, computational modelling was performed to investigate its possible role based on structure.

In collaboration with Wenbin Qi, yeast Atm1p structure was generated by using the recently solved ABCB10 transporter structure (~ 30% identity and 50% sequence similarity to the ABCB7 transporter) and SWISS-MODEL (Figure 95). Electrostatic surface map showed two positively-charged pockets at the bottom of the transmembrane segment.26 The R284 residue is located in one of the positively charged pockets.

240

Figure 95: Modelled structured of WT Atm1p and R284E mutant.Zoomed box indicate the location of R284 residue, which showed an importance in ATPase stimulation by cluster. This positive patch is also a potential binding site for the negatively-charged 2- {[2Fe-2S](GS)4} complex.

Recently, the crystal structure of Novosphingobium aromaticivorans Atm1p was solved, and it shares ~45% sequence identity with yeast and human Atm1p.22 R210 (equivalent to yeast R284) is found to be a conserved residue in 12 different organisms. In the NaAtm1p structure, R210 was found to coordination site for oxidized glutathione (Figure 96). The

NaAtm1p was also found to be in positively charged patches.

241

Figure 96: Crystal structure of NaAtm1p(PDB: 4MRS) complexed with oxidized glutathione. R210 is the conserved residue equivalent to R284 in yeast Atm1p.

Of these two positively charge patches, one lies between the two transmembrane helix bundles, and facing inward to the channel in the dimeric transporter. It’s highly probable that in its native dimeric state, these two sites of Atm1p may function to create a positive binding pocket for [2Fe-2S](GS)4 cluster. Substitution of Arg284 within the arginine-rich region with Glu appears to eliminate cluster substrate binding and ATPase stimulation.

Flow Cytometry Assays

Glutathione-coordinated iron-sulfur cluster is stable under physiological matrix conditions, where excess cellular glutathione prevents cluster hydrolysis. ESI mass 242 spectrometry, combined with studies of functional group modification suggest salt- bridges to be important in glutathione tetramer formation, which creates a macrocyclic ligand that accepts the [2Fe-2S]2+ cluster core from the scaffold protein ISU.24, 34

Functional studies of Atm1p were conducted with protein-embedded proteoliposomes and transport monitored by a novel application of both flow cytometry and tiron-ligated absorption assays. The flow cytometer was able to both detect proteoliposomes ~400 nm in diameter and quantitate the fluorescence signals of the content inside. In the case of fluorescein-loaded liposomes, a decrease in fluorescence signal was observed throughout the cluster transport reaction, because of the inner-filter effects of iron-sulfur cluster transport into the liposome (Figure 97). The advantage of particle-specific signal measurements allow events during proteoliposome detection to be correlated with the fluorescence signal associated with the proteoliposome rather than the bulk of the solution. Moreover, the cuvette pathlength is 56-fold thinner than the conventional 10 mm cuvette found in standard fluorimeters, and the thinner cuvette design eliminates

98% of the solution noise that is not associated with the proteoliposome.

243

700

600

500

400

300

200

100 Fluorescence Signal Fluorescence 0

0 200 400 600 800 1000 [cluster] (M)

Figure 97: Titration experiment showing cluster quenching the fluorescein fluorescence signal, most likely due to heavy atom and inner filter effects. A solution of glutathione iron-sulfur cluster solution (10 mM) was added in 1uL increments to 100uL 10uM fluorescein solution.

The addition of cofactor Mg-ATP provides the energy source for Atm1p to transport substrate into the proteoliposome. Flow cytometric studies of the reaction mixture containing Atm1p proteoliposome, cluster and Mg-ATP, yielded a time-dependent variation of the overall geometric mean of the event count vs. fluorescence, which decreased by ~30% over the course of an hour (Figure 98). The geometric mean was plotted against incubation time and fit to a linear function prior to plotting the slope

2- (Figure 99). Quantitative measurement of [2Fe-2S](GS)4 cluster transport into Atm1p- embedded proteoliposomes yielded an observed rate constant of 0.06 ± 0.01 min-1, in good agreement with the transport kinetics determined for other ABC transporter, such as 244

OpuA.27 The corresponding control experiments without ATP or Atm1p showed a relatively unchanged geometric mean, indicating that the overall fluorescence of the proteoliposome only decreases in the presence of the transporter protein Atm1p and the cofactor Mg-ATP.

Figure 98: Atm1p-mediated cluster transport into fluorescein-loaded proteoliposomeis demonstrated by incubation in the presence of Mg-ATP. The fluorescent response is measured by flow cytometry at 530 nm, and the geometric mean of the signal indicates a decreasing kinetic profile for fluorescein within the proteoliposome as a result of the inner filter effect from [2Fe-2S] cluster, which partially absorbs the fluorescein signal within the proteoliposome. Cluster transport, along with control experiments conducted in the absence of Atm1p and Mg-ATP are denoted in black, red, and green, respectively. Controls with GSH only, GSH + Fe3+, and GSH + S2- are shown in Figure 100.

245

16 14 12

10 * 1000 *

) 8 -1

min 6 ( 4 2 0

Initial Velocity Velocity Initial -2 -4 exp -Atm1p -ATP GSH GSH+Fe3+ GSH+S2-

Figure 99: Initial velocity derived from the rate of fluorescein signal quenching following addition of cluster and Mg-ATP (exp).The Atm1p proteoliposome fluorescence signal was measured by flow cytometry at time intervals of 10 min over a period of 50 min. Control experiments in the absence of Atm1p or Mg-ATP demonstrate that cluster transport is absent. A cluster concentration of 13 mM was used, compared to 38 mM of GSH in control experiments.

246

Figure 100: Control experiments for cluster transport into fluorescein-loaded proteoliposomewith GSH only (blue), GSH + Fe3+ (cyan) and GSH + S2- (magenta). The dashed line corresponds to the observed signal fit from experiments with clusters.

247

Figure 101: Fluorescence flow cytometry measurements of the proteoliposomes incubated with Mg-ATP (12 uM) and fluorescein labeled glutathione [2Fe-2S] cluster (1 mM) after 1 hr at r.t. and pH 7.5, with Atm1p (left), and without Atm1p (right).

During proteoliposome synthesis, the phospholipids were resuspended in the presence of

1 mM fluorescein, which encapsulates the fluorophore within the liposome. Subsequent insertion of Atm1p is also performed with excess fluorescein present in the solution to prevent diffusion during the addition of detergent. The final product contains the fluorescent probes and therefore can be readily quantified by flow cytometry. An interesting phenomenon was observed for the GSH only, and GSH/S2- control experiments, where the rate constants are non-zero, but significantly slower than that of the cluster experiment. This agrees with prior observations from the ATPase stimulation assay our laboratory29 as well as results reported by the Lill Laboratory.23 These results suggest that glutathione alone may stimulate Atm1p ATPase activity, and may be 248 transported weakly. Additional ferric and sulfide ions did not provide any significant increase in fluorescence signal change (Figure 99 and Figure 100).

Complementary studies were also carried out by monitoring the transport of fluorescein- labeled glutathione cluster into a non-fluorescent proteoliposome. Synthesis and characterization of the fluorescein-labeled glutathione Fe-S cluster are detailed in the

Materials and Methods section. Binding of the labeled cluster to Atm1p was assessed by titration of fluorescein-labeled cluster to Atm1p (Figure 90) to yield an observed KD =

118 ± 11 μM, which compares favorably with the 68 ± 2 uM determined for the unlabeled cluster by inhibition of ATPase activity,29 and demonstrating that the labeled cluster can orient in the substrate pocket without significant steric interference. The resulting fluorescein-labeled cluster was used to assay cluster transport into the proteoliposome, and the final proteoliposome product (containing labeled cluster) was detected by flow cytometry methods (Figure 101). Over the incubation period of 1 h, sufficient numbers of fluorescein-labeled clusters were taken up to yield the kinetic profile for proteoliposome transport by Atm1p.

Due to an associative glutathione ligand exchange mechanism (Figure 102), the actual rate of fluorescein labeled cluster transfer is likely to be higher than observed. As Fl- cluster was incubated with proteoliposome, some Fl-GSH may have been replaced by glutathione in solution, and therefore the fluorescence signal observed in Figure 101 is likely lower than the intrinsic rate of fluorescein labeled cluster transport. This issue is

249 difficult to be address, because at low concentrations of solution glutathione, fluorescein labeled cluster is easily hydrolyzed and oxidized (Figure 103). However, at high concentration of solution glutathione, an increase in exchange rate was observed, indicating an associative exchange mechanism of glutathione ligands.

Figure 102: Observed exchange rate of fluorescein labeled glutathione with solution glutathione.At concentrations greater than 50 mM, solution glutathione begin to exchange with cluster ligated fluorescein labeled glutathione. At concentrations lower than 5 mM, solution glutathione is too dilute to stabilize the [2Fe-2S] cluster, and cluster break down leads to an increase in fluorescence signal (increased observed rate).

250

Figure 103: An area plot of fluorescein-GSH intensity observed on ESI-MS following degradation of Fluorescein labeled cluster.As the cluster break down (hydrolyzed by water), the intensity of fluorescein-GSH (666.1 m/z) increased over one hour.

Tiron Assays

Flow cytometry experiments were further complemented by use of tiron-iron complex formation in a colorimetric assay. Tiron yields a higher extinction coefficient in the visible range relative to other iron chelators such as BPTD and EDTA.39 Following cluster transport across the membrane bilayer, the concentration of iron was readily and independently quantitated following release by acidification of the proteoliposome product with concentrated HCl. The product was then neutralized by addition of NaOH 251 and buffered with MES for tiron chelation detection, a complexing agent selective for ferric ions.40 The overall method is summarized in Figure 88.

A limiting factor for cluster uptake is the finite size and volume of the liposome.27

However, after 1 h incubation, significantly higher ferric ion concentrations were detected inside the proteoliposome, relative to control experiments carried out in the absence of protein and Mg-ATP (Figure 104), and the time-dependent change in absorbance yielded an observed rate constant of 0.07 ± 0.02 min-1 for cluster transfer.

The raw measured concentration of 3.6 ± 0.3 μM reflects an estimated liposomal concentration of ~ 0.3 mM after correction for sample dilutions.27

252

Figure 104: Concentration of iron inside the proteoliposome following 1 hr incubation with Mg-ATP and cluster.The concentration of iron was quantitated by tiron coordination and comparison of absorbance values to a standard calibration curve. The final concentrations represent the results from diluted samples following work up of the liposome samples. Data, such as that shown in this plot is intended to compare relative concentrations, rather than absolute values. In fact, the absolute concentrations will be higher, but can only be estimated based on the inner proteoliposome volume. According to Geertsma et al.,27 the total internal volume is estimated to be ~ 5 uL, and the corresponding concentration of the glutathione Fe-S cluster inside the liposome after the transport experiment is therefore estimated to be ~ 0.3 mM.

A low level of iron was also observed in the control experiments for the tiron assay and this background noise was not removed by additional washing of the proteoliposomes. A rational interpretation of this observation is that a small component of the cluster may become interstitially trapped in the membrane bilayer, and is then solubilized along with the transported cluster and contributes to the overall iron concentration. Background 253 signal was also observed in cluster transport experiments that were analyzed by OES-MS, where an elevated level of iron was detected in the proteoliposome product in the absence of the transporter protein.26 This explanation is also supported by the control experiments from the flow cytometry data, where no significant amount of signal change was observed without the essential components of the reaction. It is significant that essentially no transported iron was detected in controls that used free iron ion rather than cluster, and so neither transported nor background-associated iron appear to stem from any product of cluster degradation in the reaction solution (and in fact no degradation would be expected under the experimental conditions used).

254

Conclusion

In summary, the results support a model (Figure 105) for ABCB7-type transporters that uses glutathione-coordinated iron sulfur cluster as the physiological substrate for

2- mitochondrial cluster export to the cytosol. The [2Fe-2S](GS)4 cluster demonstrates a two-fold greater stimulation of ATPase activity at a concentrations three orders of magnitude lower than that required for stimulation by glutathione alone,29 and most likely the association and stimulation by free glutathione represents partial population of the site

2- occupied by [2Fe-2S](GS)4 . This also provides a rational for the likely transport of glutathione-coordinated heavy metals,22 which most likely mimic the [2Fe-2S] core.

Figure 105: Proposed mechanism for mitochondrial iron-sulfur cluster transport by ABCB7-type export proteins driven by ATP hydrolysis.The transporter shown in the figure is adapted from the crystal structure of Novosphingobium aromaticivorans Atm1p. 22

255

Results from electronic absorption studies support the independent findings from flow cytometry experiments, and quantitative measurement of substrate transport rates into

2- Atm1p-embedded proteoliposomes yielded an observed rate constant for [2Fe-2S](GS)4 cluster transport from the independent colorimetric assays of 0.07 ± 0.02 min-1, in good agreement with the 0.06 ± 0.01 min-1 obtained from flow cytometry. The results of this chapter provide evidences and support the proposed model of glutathione-complexed

[2Fe-2S] cluster as a natural substrate for ABCB7-type mitochondrial transporters.

Complementary analytical techniques used in the current study also demonstrate a novel quantitative methodology for study of metal translocation proteins and their proteoliposome products. These methods will be invaluable in ongoing quantitative studies of metal cofactor transport for iron-sulfur clusters and serve as a foundation to study other metal cofactor species.

256

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