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3 LIBRARY
3 0(‘5 Micnlagu State
University

 

 

This is to certify that the
dissertation entitled

PURIFICATION, KINETIC ANALYSIS, AND
SPECTROSCOPIC CHARACTERIZATION OF WILD-TYPE
Aspergillus nidulans XANTHINE HYDROXYLASE AND
SELECTED VARIANTS

presented by

Meng Li

has been accepted towards fulfillment
of the requirements for the

PhD degree in CHEMISTRY

 

\
.

 

Major Professor's Siénature

X/%7

Date

MSU is an affirmative-action, equal—opportunity employer

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PLACE IN RETURN BOX to remove this checkout from your record.
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DATE DUE

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6/07 p:/CIRC/DateDue.indd-p.1

PURIFICATION, KINETIC ANALYSIS, AND SPECTROSCOPIC
CHARACTERIZATION OF WILD-TYPE Aspergillus nidulans XANTHINE
HYDROXYLASE AND SELECTED VARIANTS
By

Meng Li

A DISSERTATION

Submitted to
Michigan State University
In partial fulfillment of the requirements
For the degree of

DOCTOR OF PHILOSOPHY
Department of Chemistry

2007

ABSTRACT
PURIFICATION, KINETIC ANALYSIS, AND SPECTROSCOPIC
CHARACTERIZATION OF WILD-TYPE Aspergillus nidulans XANTHINE
HYDROXYLASE AND SELECTED VARIANTS
By

Meng Li

Xanthine hydroxylase (XanA) is the first member in the Fe(II)/(x-ketoglutarate
dependent dioxygenase superfamily to catalyze the hydroxylation of a purine base.
Hiso-tagged XanA of Aspergillus nidulans was purified from both the fungal
mycelium and recombinant Escherichia coli cells. Comparison studies revealed very
different quaternary structures and posttranslational modifications; however, the
kinetic properties of XanA purified from both hosts are very similar. Extensive kinetic
studies were carried out for His6-tagged XanA isolated from bacterial host. The
enzyme exhibits no significant isotope effect when using 8-2H-xanthine whereas it
demonstrates a two-fold solvent isotope effect. XanA is Fe(II)-specific and has high
selectivity for substrates. A homology model of XanA was generated on the basis of
the structure of the related enzyme taurine/OLKG dioxygenase (TauD). The putative
active site residues were mutated to alanine, and the variants were purified and
kinetically characterized by using native substrate and chemical analogues. The three
mutants of the predicted metal ligands showed an increased Kd of Fe(II), H149A and
D151A are completely inactive and H340A exhibited only trace amount of activity.
Asn358 is crucial for catalysis. The Q356A and N358A variants had significantly

increased Kiapp over control protein for 6,8-DHP, suggesting that Gln356 and Asn358

 

 

 

 

 

 

hydrogen bond with the C-6 hydroxyl group of substrate. The E137A and D138A
variants demonstrated enhanced activity with 9-methylxanthine, consistent with
Glu137 and Asp138 being proximal to N-9 of substrate. Based on the extensive
kinetic studies and the homology model, a preliminary xanthine binding mode was
proposed and this is the first step to understand substrate binding for XanA when an
enzyme crystal structure is not available. Electronic spectroscopy and one- and
two-dimensional electron spin echo envelope modulation (ESEEM) experiments have
been used to study the coordination chemistry at the nitric oxide (NO)-bound
non-heme Fe(II) sites of XanA and TauD. Electron paramagnetic resonance (EPR)
spectra indicated the electronic environment perturbation of the {FeNO}7 center were
perturbed after addition of primary substrate for both enzymes. ESEEM and
HYSCORE (hyperfine sublevel correlation spectroscopy) detected hyperfine
couplings from lH and MN nuclei in different protein complexes, and XanA exhibited
patterns very similar to TauD. Interestingly, a new coupling coming from a
non-exchangeable IH approximately 3.2 A from the metal ion of TauD was detected
in the NO-bound ternary complex. A particular residue from the enzyme probably
contributes to this signal and one candidate, Trp248, was examined by analysis of the
W248F variant. This study represents the first biochemical characterization of
purified xanthine/orKG dioxygenase. It suggests the substrate binding mode through
mutagenesis studies, it provides the first spectroscopic information for this enzyme,

and it yields insights into the coordination properties of F e(II) in this protein.

To my dearest father, mother
&
Beloved husband

 

 

 

COU

Ion

222i
an]

ad

pr

 

ACKNOWLEDGMENTS

I would not have been able to complete a PhD without the aid and support of
countless people over the past five years. I must first express my deepest gratitude
towards my advisor, Prof. Robert Hausinger, for his guidance, encouragement and
continuous support through the course of my work. The extensive knowledge, vision,
and creative thinking of Dr. Hausinger have always been the source of inspiration for
me. Also, I would like to thank Prof. Joan Broderick who had been an excellent
advisor and great teacher to me for the first three years of my PhD.

I would like to thank many people for their help during my studies: Prof. Claudio
Scazzocchio and his graduate student Gabriela M. Montero-Moran initiated this
project and provided the xanA gene. Prof. John McCracken assisted with EPR and
ESEEM analyses and performed the HYSCORE simulations. Prof. Michael Feig and
Andrew W. Stumpff—Kane constructed the homology model. Prof. David J. Lowe and
Fabrice Jourdan synthesized the 8-hydroxypurine, 2,8-dihydroxypurine and
6,8-dihydroxypurine. Christine Drevet provided the alignment figure of XanA with
putative orthologous sequences from several fungi. Dr. Tina Miiller constructed the
XanA variants and carried out ferroxidase assays. thalia Kalliri and Dr. Piotr
Grzyska provided wild-type and W248F TauD samples. Prof. James Geiger provided
a model of NO-bound TauD. J. C. Price and Prof. Marty Bollinger provided

deuterium-labeled taurine. Prof. Nicholas Burzlaff provided NOG. Dr. Rajendra Bose

Muthukumaran initiated the HYSCORE studies. Prof. Dan Jones provided mass
spectrometry assistance.

I also thank the many graduate and undergraduate students I have worked with in
Prof. Hausinger’s group: Jana, Kim, Piotr, Scott, Soledad, Thalia, Tina, Aaron,
Andrea, Bruce, Kimberly, Melody and Rachel. The friendly and supportive
atmosphere inherent to the whole group contributed essentially to the final outcome of
my studies. I wish to thank Bruce in particular who has been a great help and assisted
me during the mutagenesis study. They each helped make my time in the PhD
program more fun and interesting.

Last but not least, I wish to thank my family who have always supported me, and
my husband, Jihua, who always believes in me. I could not finish this long journey

without you.

vi

TABLE OF CONTENTS

LIST OF TABLES .................................................................... xi
LIST OF FIGURES .................................................................. xii
ABBREVIATIONS ................................................................... xvii
CHAPTER 1
Introduction .............................................................................. 1
Fe(II)/a-ketoglutarate-dependent hydroxylases ................................. 2
Protein modification ....................................................... 4
Prolyl 4-hydroxylase .................................................. 4
Transcriptional regulation ................................................ 5
Prolyl hydroxylase domain containing enzymes &
Factor-inhibiting HIF ................................................. 5
Jumonji C (ijC)-domain-containing histone demethylases 6
Repair of alkylated DNA/RNA .......................................... 8
AlkB ..................................................................... 8
Biosynthesis of antibiotics and plant products ........................ 11
Clavaminate synthase ................................................. 11
F lavanone 3-hydroxylase ............................................. 12
Synthesis or decompose of small molecules ........................... 14
Taurine/aKG dioxygenase ........................................... l4
2,4-DichIorophenoxyacetate /aKG dioxygenase .................. 15
Xanthine hydroxylase ................................................. 16
Related enzymes ..................................................................... 18
Isopenicillin N synthase ................................................... l8
4-Hydroxyphenylpyruvate dioxygenase ................................ 20
Structural studies of Fe(II)/aKG-dependent hydroxylases .................... 21
Mechanistic studies of Fe(II)/dKG-dependent hydroxylases ................. 27
Thesis outline ........................................................................ 38
References ........................................................................... 41
CHAPTER 2
Purification and properties of Aspergillus nidulans xanthine hydroxylase ...... 58
Abstract .............................................................................. 59
Introduction .......................................................................... 60
Experimental procedures ........................................................... 64
Growth of E. coli cells overproducing A. nidulans XanA ............ 64
Purification of Hisé-tagged XanA from E. coli ......................... 64

vii

Enzyme assays ............................................................. 65

Metal analyses ............................................................ 66
Protein analytical methods ................................................ 66
Mass spectrometry ......................................................... 66
Structural homology modeling ........................................... 67
Results ................................................................................ 68
Properties of recombinant XanA in E. coli cell extracts .............. 68
Purification of XanA from E. coli ....................................... 68
Effects of pH on stability and activity of XanA from E. coli ......... 72
Kinetic analyses of XanA from E. coli .................................. 72
Differential protein properties of XanA purified from the two host
cells ......................................................................... 75
Discussion ........................................................................... 80
Purification ................................................................. 80
XanA enzyme stability ................................................... 8O
XanA enzyme activity .................................................... 81
Posttranslational modifications .......................................... 81
Insights from a homology model ....................................... 84
References .................................................................. . ........ 86
CHAPTER 3

Kinetic characterization, isotope effects, and effects of alternate metal ions,
salt, and substrate analogues on Aspergillus nidulans xanthine hydroxylase 91

Abstract .............................................................................. 92
Introduction .......................................................................... 93
Experimental procedures ........................................................... 97
Purification of bacterial-derived XanA ................................. 97
Enzyme assays ............................................................. 97
Sources and synthesis of chemical analogues of aKG and xanthine. 97
8-2H-Xanthine preparation ................................................ 98
2H20 solvent isotope effect ............................................... 98
Structural modeling ........................................................ 98
Results ................................................................................ 100
Isotope effects .............................................................. 100
Effect of other metal ions and salt ....................................... 100
aKG analogues ................................................................................. 104
Xanthine analogues .......................................................................... 104
Structural model of XanA ................................................. 104
Discussion ............................................................................ 109
Isotope effects ............................................................... 109
Metal ion and salt effects .................................................. 110
- Co—substrate and substrate specificity .................................... 110
Postulated substrate-binding mode 112
References ............................................................................ l 15

viii

 

 

 

 

 

 

CHAPTER 4
Characterization of active site variants of xanthine hydroxylase from Aspergillus

nidulans ................................................................................... 1 22
Abstract .............................................................................. 123
Introduction .......................................................................... 1 25
Experimental procedures ....... '. ................................................... 128

Site-directed mutagenesis, protein overproduction and purification..128
Fluorescence Quenching Analyses of Fe(II) and OLKG Binding 128

Enzyme assays ............................................................. 129
Kinetic analysis of 6,8-DHP inhibition ................................. 130
Results ................................................................................ 132
Fe(II) and orKG Binding to XanA and its Variants .................... 132
Kinetic comparison of XanA and its variants with xanthine ......... 134
Alternative substrates for XanA and its variants ....................... 141
6,8-DHP inhibition ......................................................... 143
Identification of a reactive thiol at the XanA active site .............. 155
Oxygen reactivity of XanA in the absence of primary substrate 159
Discussion ........................................................................... 164
Model of xanthine binding based on kinetic analyses of XanA
and its variants .............................................................. 164
Lack of binding by related compounds ................................. 169
Comparison to xanthine binding in other systems ..................... 171
References ............................................................................ l 73
CHAPTER 5

Coordination chemistry at the Fe(II) site of taurine/a-ketoglutarate dioxygenase
and Aspergillus nidulans xanthine hydroxylase by various spectroscopies 175

Abstract .............................................................................. 176
Introduction ......................................................................... 1 78
Experimental procedures .......................................................... 183
Materials .................................................................... 183
Enzyme purification ....................................................... 183
Analysis of Fe(II), orKG, and substrate binding to TauD and
XanA by using electronic spectroscopy ................................. 183
Electronic spectroscopy of NO-treated samples ........................ 184
Electron paramagnetic resonance analyses ............................. 184
HYSCORE spectra simulation ........................................... 186
Results ................................................................................ 187
Binding of (xKG and xanthine to Fe(II)-XanA monitored by
electronic spectroscopy ................................................... 187
Electronic spectroscopy of NO complexes of TauD and XanA 187
EPR of {FeNO}7 adducts of TauD and XanA ........................ I90

One-dimensional ESEEM of {FeNO}7 adducts of TauD and

ix

XanA ....................................................................... 1 94

Two-dimensional ESEEM of {FeNO}7 adducts of TauD .......... 201
Two-dimensional ESEEM of {FeNO} 7 adducts of XanA .......... 213
Discussion ........................................................................... 221
References ....................................................... . .................... 229
CHAPTER 6
Conclusions .......................................................................... 236
Future research ...................................................................... 240
Constructing double or triple mutants ................................. 240
Determination of dissociation constant (Kd) .......................... 240
Steady-state and transient kinetic studies by stopped-flow
UV-Vis spectroscopy ..................................................... 237
Resonance raman and Mossbauer spectroscopy ...................... 242
More HYSCORE spectroscopy studies ............................... 242
Problems ................................................................... 243
References ........................................................................... 245

LIST OF TABLES

CHAPTER 4

TABLE 4.1: Determination of Kd of Fe(II) and OLKG for XanA and. Selected
Variants. ................................................................................... 139

TABLE 4.2: Substrate, Cosubstrate, and Substrate Analog Kinetic Parameters
of XanA and Selected Variants. ........................................................ 140

TABLE 4.3: Inhibition of Wild-Type XanA and its Variants by 6,8-DHP. ...... 154

TABLE 4.4: Oxygen Reactivity of XanA and Selected Variants in the absence
of Primary Substrate. .................................................................... 161

xi

LIST OF FIGURES

(Images in this dissertation are presented in color.)

CHAPTER 1
FIGURE 1.1: General reaction of Fe(Il)/0tKG-dependent hydroxylases. ........ 1
FIGURE 1.2: General reaction of prolyl 4-hydroxylases (P4H). .................. '1

FIGURE 1.3: General reaction of HIF-or-specific asparaginyl hydroxylases
(FIH). ...................................................................................... 1

FIGURE 1.4: General reaction of ijC-domain-containing histone -
Demethylases (JHDM). ................................................................. 9

FIGURE 1.5: General reactions of AlkB. ............................................. 9

FIGURE 1.6: Reactions of Clavaminate synthase (CAS). .......................... 13
FIGURE 1.7: General reaction of Flavanone 3B—hydroxylase (F3H). ........... 13
FIGURE 1.8: General reaction of TauD. ............................................. 13
FIGURE 1.9: General reaction of deA. ............................................. 17

FIGURE 1.10: General reaction of Moco-containing xanthine oxidase (XO). .. 17

FIGURE 1.1]: General reaction of isopenicillin N synthase (IPNS). ............ 17

FIGURE 1.12: General reaction of 4-hydroxypheny1pyruvate dioxygenase

(HPPD). .................................................................................. 17
FIGURE 1.13: Double-stranded B-helix structure. ................................. 25
FIGURE 1.14: OLKG binding modes. ................................................. 25

FIGURE 1.15: Proposed mechanism for the hydroxylation reaction catalyzed
by Fe(II)/0tKG-dependent dioxygenase. ............................................ 30

FIGURE 1.16: Proposed mechanism for oxidative cyclization catalyzed by
CAS. ...................................................................................... 36

FIGURE 1.17: Proposed mechanism for a desaturation reaction. ............... 36

 

 

 

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FIGURE 1.18: General reaction of halogenase. ..................................... 36
CHAPTER 2

FIGURE 2.1: General mechanism of Fe(II)/0tKG-dependent xanthine
hydroxylases. ............................................................................ 6 1
FIGURE 2.2: Activity assay of XanA cell extracts. ............................... 69

FIGURE 2.3: SDS-PAGE analysis of the purified XanA from E. coli and
A. nidulans. ...................................................................... . ....... 71

FIGURE 2.4: pH dependence of XanA activity. .................................... 73

FIGURE 2.5: Substrate and co-substrate concentration dependencies of XanA.74

FIGURE 2.6: F e(II) concentration dependence of XanA. .......................... 76
FIGURE 2.7: Measurement of native size of XanA derived from E. coli. ...... 77
FIGURE 2.8: Mass spectrometric analysis of fungus-derived XanA. ............ 79

FIGURE 2.9: Location of potential sites of glycosylation and phosphorylation

in XanA. .................................................................................. 83
CHAPTER 3

FIGURE 3.1: General mechanism of Moco-containing xanthine oxidases. 94
FIGURE 3.2: Solvent deuterium isotope effect on XanA activity. ............... 101
FIGURE 3.3: Divalent cation inhibition of XanA. ................................. 102
FIGURE 3.4: NaCl inhibition of XanA. ............................................. 103
FIGURE 3.5: NOG inhibition of XanA. ............................................. 105
FIGURE 3.6: Homology model of XanA. ........................................... 107

FIGURE 3.7: Comparison of sequences from XanA orthologues in selected

fungi. ...................................................................................... 113

CHAPTER 4
FIGURE 4.1: Depiction of the putative active site pocket of XanA revealed

from a homology model. ............................................................... 126

xiii

i—M

>t‘

 

 

F1

 

 

 

 

FIGURE 4.2: Denaturing polyacrylamide gel electrophoretic analysis of Xan
variants purified from E. coli. ......................................................... 133

FIGURE 4.3: Fluorescence spectroscopic analysis of XanA interaction with
Fe(II). ..................................................................................... 135

FIGURE 4.4: Fluorescence spectroscopic analysis of XanA interaction with
OLKG .................................................................................... 137

FIGURE 4.5: UV-visible spectra of 1-methy1xanthine, 9-methy1xanthine, and

their products. ............................................................................ 142
FIGURE 4.6: Interaction of XanA with 6,8-DHP. .................................. 144
FIGURE 4.7: General mechanism for slow-binding inhibition. ................... 145
FIGURE 4.8: Interaction of Q101A with 6,8-DHP. ................................. 147
FIGURE 4.9: Interaction of K122A with 6,8-DHP. ................................. 148
FIGURE 4.10: Interaction of E1 37A with 6,8-DHP. ................................ 149
FIGURE 4.11: Interaction of D138A with 6,8-DHP. ................................ 150
FIGURE 4.12: Interaction of Q356A with 6,8-DHP. ................................ 151
FIGURE 4.13: Interaction of C357A with 6,8-DHP. ................................ 152
FIGURE 4.14: Interaction of N358A with 6,8-DHP. ................................ 153
FIGURE 4.15: Inactivation of XanA by DTNB. .................................... 156

FIGURE 4.16: Time-dependent loss of XanA activity for highly diluted
enzyme. .................................................................................... 158

FIGURE 4.17: Oxygen consumption studies of XanA and its variants. .......... 160

FIGURE 4.18: Hydrogen peroxide production by Q101A and wild-type XanA.163 2

FIGURE 4.19: Proposed xanthine binding on the active site. ..................... 166
CHAPTER 5 .
FIGURE 5.1: TauD active site. ......................................................... I80

xiv

FIGURE 5.2: General mechanism of TauD. ......................................... 181
FIGURE 5.3: UV-visible absorption spectra of XanA complexes. ............... 188

FIGURE 5.4: UV-visible absorption spectra of {FeNO}7 complexes of TauD
and XanA. ................................................................................ 189

FIGURE 5.5: Thermodynamics and kinetics of NO binding to TauD and
XanA. ..................................................................................... 191

FIGURE 5.6: EPR spectra of the {FeNO}7 complexes of TauD and XanA. 193

FIGURE 5.7: One-dimensional ESEEM spectra of different TauD complexes
prepared in H20 and 60% 2H20 buffer at 172.0 mT and 340.0 mT. .............. 195

FIGURE 5.8: One-dimensional ESEEM spectra of different XanA complexes
prepared in H20 and 60% szO buffer at 172.0 mT and 340.0 mT. .............. 198

FIGURE 5.9: HYSCORE spectra of different TauD complexes prepared in
H20 and 60% 21120 buffer at 172.0 mT. ............................................. 203

FIGURE 5.10: HYSCORE spectra of taurine-aKG-Fe(II)-TauD complex
prepared in H20 buffer at 172.0 mT (A), 340.0 mT (C) and 60% 2H20 buffer
at 172.0 mT (B), 340.0 mT (D). ....................................................... 204

FIGURE 5.11: HYSCORE spectra of different TauD complexes prepared in
H20 and 60% szO buffer at 340.0 mT. ............................................. 206

FIGURE 5.12: UV-Vis spectra and HYSCORE of ternary complex
taurine-NOG-Fe(II)-TauD. ............................................................ 208

FIGURE 5.13: HYSCORE spectra at 172.0 mT of taurine-zH-labeled-aKG
-Fe(II)-TauD prepared in 60% 2&0 buffer (A) and 90% szO buffer. ......... 209

FIGURE 5.14: HYSCORE spectra of 2H-Iabeled taurine-aKG-Fe(II)-TauD
complex prepared in H20 buffer at 172.0 mT (A) and 340.0 mT (B). ........... 210

FIGURE 5.15: HYSCORE spectra of taurine-aKG-Fe(II)-TauD complex
prepared in 60% 2H20 buffer at 172.0 mT (A), 182.0 mT (B) and 192.0 mT

(C). ........................................................................................ 214
FIGURE 5.16 Two depictions of TauD active site and the XanA active site. 215

FIGURE 5.17: HYSCORE spectra at 172.0 mT of taurine—aKG-Fe(I.I)-TauD

XV

 

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HI~
Hf

 

Flt»
Hg

llt .
pk"
at I

Fe
but

Cl‘
Flt
(ng

 

W248F prepared in H20 buffer (A) and 60% 2H20 buffer (B). .................. 216

FIGURE 5.18: HYSCORE spectra of different XanA complexes prepared in
H20 and 60% 2&0 buffer at 172.0 mT. ............................................. 217

FIGURE 5.19: HYSCORE spectra of different XanA complexes prepared in
H20 and 60% 21120 buffer at 340.0 mT. ............................................. 218

FIGURE 5.20: HYSCORE spectra of xanthine-aKG-Fe(II)-XanA complex
prepared in H20 buffer at 172.0 mT (A), 340.0 mT (C) and 60% 2H20 buffer
at 172.0 mT (B), 340.0 mT (D). ....................................................... 219

FIGURE 5.21: HYSCORE spectra at 172.0 mT of Fe(II)-TauD (A), (XKG-
Fe(II)-TauD (B), and taurine-aKG-Fe(II)-TauD complex prepared in H20
buffer. ..................................................................................... 226

CHAPTER 6

FIGURE 6.1: HYSCORE spectra of NO treated ternary complex TauD-Fe-
aKG-taurine in 50 mM Tris-H20, pH = 8.0 at (A) 3400 G and (B) 1720G 244

xvi

aKG

ABTS

CAS

2,4-D

DAOCS

DEA/NO

2,8-DHP
6,8-DHP
DSBH
DTNB
EPR
ESEEM
F3H

F IH

HIF
HPPD
HYSCORE
IPNS
JHDM

LMCT

ABBREVIATIONS

a-Ketoglutarate
2,2’-Azino-bis(3-ethylbenzthiazoline—6-sulfonic acid)
Clavaminate synthase

2,4-Dichlor0phenol

Deacetoxycephalosporin C synthase
Diethylammonium (Z)-1-(N,N-diethylamino)diazen-l-ium
-1,2-diolate

2,8-Dihydroxypurine

6,8-Dihydroxypurine

Double-stranded B-helix
5,5’-Dithiobis(2-nitrobenzoic acid)

Electron paramagnetic resonance

Electron spin-echo envelope modulation

Flavanone 3fl-hydroxylase

HIF-a-specific asparaginyl hydroxylases

Hypoxia —inducible factor

4-Hydroxyphenylpyruvate dioxygenase

Hyperfine sublevel correlation spectroscopy
Isopenicillin N synthase

ijC domain-containing histone demethylase

Li gand-to-metal charge-transfer

xvii

.\1l

M1

N1?

\l

01

 

n4 ll.|.1 ll: J!

P3

 

 

 

MLCT

Moco

NO

NOG

NTA

OPDA

P3H

P4H

PNGase F

SDS-PAGE

TauD

deA

XAS

XanA

XO

Metal-to-ligand charge-transfer
Molybdopterin cofactor

Nitric acid

N-oxalylglycine
Nitrilotriacetic acid
o-Phenylenediamine

Proline 3—hydroxylase

Prolyl 4-hydroxylases
N-glycosidase F

Sodium dodecyl sulfate-polyacrylamide gel electrophoresis
Taurine/OLKG dioxygenase
2.4-D dioxygenase

X-ray absorption spectroscopy
Xanthine/(XKG dioxygenase.

Xanthine oxidase

xviii

CHAPTER 1

INTRODUCTION

 

(if

d1

31

 

 

 

 

Fe(II)/a—KETOGLUTARATE-DEPENDENT HYDROXYLASES
Fe(II)/a-ketoglutarate (aKG)—dependent hydroxylases catalyze a wild range of
reactions including protein side-chain modification (some of which are involved in
transcriptional regulation), repair of alkylated DNA/RNA, biosynthesis of antibiotics
and plant products, lipid metabolism, and synthesis or decomposition of a wide
variety of small molecules (1-13). Structural studies reveal a conserved
double-stranded B-helix (DSBH) core of this superfamily, also called a B-strand
“jellyroll” structure. This conserved structure is responsible for binding the iron
through three amino acid side chains in a His'-X-Asp/Glu-X,,-His2 motif (where n can
range from 44 to over 150). In addition, the metal is bound by aKG using its C-l
carboxylate and C-2 keto groups. Binding of co-substrate and substrate triggers the
ligation of dioxygen to metal, stimulates the oxidative decomposition of aKG to
succinate and C02, and leads to the formation of a highly reactive Fe(IV)-oxo species
which is proposed to hydroxylate the primary substrate. The overall reaction is
illustrated in Figure 1.1. In this chapter, I briefly introduce several representatives of
this superfamily of enzymes, including selected Fe(II)/0tKG-dependent hydroxylases
as well as others that are structurally or mechanistically related. I discuss the general
structural and mechanistic features of these enzymes based on kinetic and
spectroscopic studies. Finally, I provide an outline of my thesis to highlight the

purpose of my studies.

FIGURE 1.1: General reaction of Fe(Il)/0tKG-dependent hydroxylases.

 

Fe(ID/aKG—dependent
R-H hydroxylases ; R-OH
0H '
0 H0
0 0
+ 002
H00 H00
aKG succinate

FIGURE 1.2: General reaction of prolyl 4-hydroxylases (P4H).

0
R §~R R\ §—R

N “ P4H N
O 7’"

02 + “KG CO2 + succinate
OH

Prolyl side chain 4-Hydroxyprolyl side chain

FIGURE 1.3: General reaction of HIF—a-specific asparaginyl hydroxylases (FIH).

02 + aKG C02 + succinattce3

Asparaginyl side chain [S-Hydroxy asparaginyl side chain

 

Protein Modification

Prolyl 4-hydr0xylase

Prolyl 4-hydroxylase (P4H), the first Fe(II)/aKG-dependent hydroxylase
identified (14), catalyzes the hydroxylation of proline residues to yield the
trans-4-hydroxyprolyl group as shown in Figure 1.2. In mammals, P4H is the key
enzyme in the biosynthesis of collagens (2), a family of extracellular matrix proteins,
as 4—hydroxyproline residues are essential for the folding of the newly synthesized
collagen polypeptide chains into triple-helical structure. The enzyme recognizes the
Gly-X-Pro motif and about 10% of the Pro position is modified. P4H has also been
isolated from plants (15, 16), where it hydroxylates proline-rich structural
glycoproteins of the cell walls. Genes encoding P4H have been cloned from human,
plant, insect, nematode, and other sources, including Paramecium bursaria Chlorella
virus-l (15-20). Generally, the protein is composed of two subunits, a catalytic
subunit on, which contains separate catalytic and peptide substrate-binding domains,
and a protein disulfide isomerase subunit, [3, except for algal and plant enzymes which
are identified as monomer (15, 16). His412, Asp414 and His483 were proposed to be
the metal ligands for human enzyme via a mutagenesis study (21, 22), and aKG is
thought to chelate the Fe(II) center in a bidentate fashion as described above with the
C-5 carboxylate forming a salt bridge with Lys493. The peptide substrate-binding
domain consisting of residues 144-244 of the human 0; subunit was crystallized and
diffracted to at least 3A (23), however, there is no complete structure reported since

the catalytic domain was not included in the crystal.

Transcriptional Regulation

Prolyl hydroxylase domain-containing enzymes & F actor-inhibiting HIF

Fe(II)/aKG-dependent hydroxylases were known for decades to catalyze protein
post-translational modifications (24-26), and included the prolyl, lysyl, and
aspartyl(asparaginyl) hydroxylases. Recently, some representatives of these enzymes
were shown to play a novel role involving regulation of the hypoxic response, one of
the most important ways in which animals respond to reduced levels of dioxygen (3, 4,
27-29). Hypoxia —inducible factor (HIF) is an (113 heterodimeric transcription factor
enabling the transcription of an array of genes that work to compensate for the effects
of low oxygen tension. Under low oxygen condition, the HIF-a subunit translocates
to the nucleus, dimerizes with HIF-B and together they recruit coactivators, such as
p300, to initiate a transcriptional response. Under normal dioxygen concentraions,
HIF-(x is targeted for degradation by two independent pathways. One path involves
hydroxylation of either Pro402 or Pr0564 by HIF-a-specific prolyl hydroxylase
domain-containing enzymes, using an oxygen, Fe(II), and OtKG-dependent reaction (3,
4, 30). The hydroxylated HIF-(x forms a complex with the von Hippel-Lindau tumor
suppressor protein, elongin B, and elongin C, resulting in polyubiquitinylation and
destruction of the transcription factor subunit. The other pathway involves
hydroxylation at the pro-S position of the B-carbon of Asn803 in HIF-ot through
factor-inhibiting HIF (FIH) (5, 31-33) as shown in Figure 1.3, The action of this
HIF-a-speciflc asparaginyl hydroxylase prevents interaction of HIF-oz with the p300

transcription coactivator, thus repressing HIF transcriptional activity. An X-ray

bf

311

3C1

pr:

CU]

of

8p:

an

the

CI.

 

 

crystallography study predicted that FIH is comprised of a B-strand jellyroll core with
both Fe(II) and the co-substrate OLKG bound in the active site (34). The metal ligands
are Hisl99, Asp201, and His279, whereas Tyr145, Thr196, and Ly5214 stabilize
binding of the (xKG C-5 carboxylate in a unique type of interaction. The crystal
structure of FIH, in complex with Fe(II), OLKG and the C-terminal transactivation
domain of HIP-0t, was also obtained. Asn803 of CAD is precisely orientated in the
active site to allow hydroxylation to occur at its B carbon; however, oxidation was
prevented by the anaerobic conditions used for crystallization. Similarly, anaerobic
conditions in the cell prevents HIF hydroxylation allowing it to function as a
transcription factor.

Jumonji C (ijC)- domain-containing histone demethylases

Covalent histone modifications have an important role in regulating a wide range
of processes including gene activity, chromatin structure, dosage compensation and
epigenetic memory (35). One such modification is methylation, occurring on arginine
and lysine residues, the extent of which is controlled by a balance between enzymes
that catalyze the addition and removal of this modification (36). It was long thought
that histone methylation was irreversible until a novel ijC domain-containing
protein, JHDMI (ijC domain-containing histone demethylase 1), that specifically
demethylates histone H3 at lysine 36 (H3K36) was discovered in 2005 (3 7). From
then on, more JHDM members have been identified and JHDMs have become the
third and largest class of demethylase enzymes. PADI4 (petidylarginine deiminase 4)

converts methyl-arginine to citrulline as opposed to directly reversing arginine

 

 

m:
ll}

16‘

111;
Ii};
11]

l'C‘.

yl

 

methylation, so it cannot strictly be considered a histone demethylase (38, 39). LSDl
(lysine specific demethylase 1), a representative of a second class of enzymes, directly
reverses histone H3K4 or H3K9 modifications by an oxidative demethylation reaction
in which flavin is cofactor (40, 41). It is worth noticing that the state of histone
methylation, in addition to the site of lysine modification, is important in determining
the functional outcome of this epigenetic modification. Unlike the other two classes,
which can only remove mono- and dimethyl lysyl modifications, the JHDMs can
remove all three histone lysine-methylation states. In this very active area of
investigation, JHDMs already have been shown to reverse H3K36 (JHDMI) (37),
H3K9 (JHDM2A) (42) and both H3K9 and H3K36 (JHDM3 and JMJDZA-D)
methylation (43-46). A set of 98 ijC-domain-containing proteins from human to
yeast has been reported based on the analysis of public protein-domain databases (47).
Curiously, FIH, functioning as an asparagine hydroxylase for the HIF-a transcription
factor, was included in this enzyme family. Even though FIH is only found in higher
eukaryotes such as mice and humans, it is still a useful template for study of other
ijC-domain-containing proteins.

The reaction catalyzed by JHDMs uses Fe(II) and OLKG as cofactors to
hydroxylate the methyl group of the modified lysyl side chain of histones, with the
resulting intermediate spontaneously decomposing as shown in Figure 1.4. In
agreement with the reaction, formaldehyde and succinate were detected during the
demethylation by JHDMlA (a human JHDMl homologue) (37). Based on the

recently solved structure (2.28 A) of the catalytic domain of JMJDZA (48), the ijC

 

dc

311

hr.

 

 

domain folds into eight conserved B-sheets forming the typical jellyroll-like structure.
The iron atom is chelated by three absolutely conserved residues: Hisl88, Glu190,
and Hi5276. Two water molecules binding to the metal center also were observed in
the native structure. In the presence of OLKG, the two water molecules are replaced by
two oxygen atoms from aKG which associate with Fe(II) via the C-1 carboxylate and
C-2 keto groups. aKG is further stabilized by three hydrogen bonds formed between
OLKG and the side chains of Tyrl32, Asn198 and Lys206. The potential substrate
binding site and binding mechanism were proposed, but a complex structure of
JMJD2A and its cognate substrate peptide is still required to verify this model.

Repair of Alkylated DNA/RNA

AlkB

Cellular DNA can be damaged by various intracellular and environmental
alkylating agents to produce alkylation base lesions which may cause genetic changes
that lead to diseases such as cancer (49-51). In E. coli, an adaptive responsive
pathway mediated by Ada protein is initiated by this treatment (52). Methyl transfer to
Cys69 of Ada converts it into an activator that increases expression of alkB and two
other genes, alkA and aidB. AlkA is a glycosylase that cleaves specific methylated
bases from DNA and performs the first step of the well-known base excision repair
pathway of base lesions. The precise function of AidB is still unknown. The activity
of AlkB was unknown for almost 20 years after its involvement in alkylation damage
was shown; however, based on phenotypic studies, AlkB was known to function as a

DNA repair enzyme that demethylates l-methyladenine and 3-methylcytosine of

FIGURE 1.4: General reaction of ijC—domain-containing histone demethylases

(J HDM).
.CHa CH20H
+H2N ‘ +H2N
Spontaneous
0Ez + u—fffm GCO2 + succinate HCHO
R s
N R
H O H o
Monomethylated lysyl side chain Lysyl side Chill“

FIGURE 1.5: General reactions of AlkB.

NH2 NH; NH;

4.
I” «II: °“’°H ”f“
/ /
/<N lN/J AlkB J Spontaneous /<N I ’J
DNA DNA/ TDNA

02 + GKG C0: + succinate HCHO
Nl-methyladenine

NH, NH2

grow N,CH20H lL/IIITN
‘WTJLO AlkB INT/LO Spontaneous *0
°7fir

OLA 0, + aKG C02 + succinate D'LA HCHO D'LA

Adenine

Ns—methylcytoslne Cytoslne

single-strand polynucleotides (53, 54). Sequence profile analysis revealed that AlkB
belongs to the Fe(II)/0tKG-dependent dioxygenase superfamily (55). In 2002, two
groups confirmed the biochemical activity of AlkB at the same time by detecting the
formation of formaldehyde and showing that the AlkB enzyme utilizes an oxidative
demethylation pathway to repair baselesions (1, 56). In this pathway, oxidation of the
methyl group gives a hydroxylated intermediate, which spontaneously decomposes to
release the repaired base and formaldehyde as illustrated in Figure 1.5. So far, in vitro
studies have shown that the sites repaired by AlkB include l-methyladenine and
3-methylcytosine in DNA or RNA, as well as adducts containing ethyl, hydroxyethyl,
propyl, and hydroxylpropyl groups (5 7, 58).

Homologs of AlkB have been discovered based on sequence and fold similarity,
and the alkB gene is conserved from bacteria to human (55, 59-61). At least eight
homologs exist in the human genome, and the functions of two of them, ABHZ and
ABH3, as well as two mouse homologs mAbh2 and mAbh3 have been characterized
(57, 62). In contrast, ABHI, which is 52% similar and 23% identical to E. coli AlkB,
along with ABH4, ABH6 and ABH7 did not show any activity on any of the known
substrates for AlkB. In 2006, crystal structures of substrate and product complexes of
E. coli AlkB were resolved from 1.8 A to 2.3 A (63). The catalytic core contains a
DSBH in which eight B-strands form a jellyroll-like structure matching that observed
in other superfamily members. Fe is bound through the Hisl31, Asp133 and Hisl87
facial triad and OLKG chelates to Fe(II) via its C-l carboxylate and C-2 keto groups

with Arg204 and Arg210 forming salt bridges to C-5 carboxylate. The alkylated

10

 

 

TIL.

 

 

 

l-methyladenine base is bound in a deep, predominantly hydrophobic cavity and the
nucleotide backbone is bound in a cooperative hydrogen bonding network.
Interestingly, there is a unique subdomain, 90 N-terminal residues, that holds the
methylated trinucleotide substrate into the active site through contact to the
polynucleotide backbone. Through amide hydrogen exchange studies and
crystallographic analyses (63), this substrate-binding “lid” was shown to be flexible
but its flexibility is reduced on binding the substrate. This suggested that AlkB has
evolved an elegant molecular design in which this flexible nucleotide-recognition lid
recognizes invariant nucleic acid backbone features to produce high-affinity docking
of chemically diverse modified bases into its conserved catalytic core. Analysis of
crystal structures of substrate and product complexes provided valuable information
about the mechanism of AlkB demethylation which will be elucidated in later sections
of this chapter.

Biosynthesis of Antibiotics and Plant Products

Clavaminate synthase

Soon after the introduction of ,B-lactam compounds (like penicillin) as antibiotics,
bacteria developed resistance mechanisms involving fl-lactamases, that hydrolyze the
antibiotic to give biologically inactive products (64). Clavulanic acid is a natural
compound which inhibits fl-lactamase mediated hydrolysis and Clavaminate synthase
(CAS) is a remarkable Fe(II)/0tKG-dependent dioxygenase that catalyzes three
separate oxidative reactions in the biosynthesis of Clavulanic acid (65), Figure 1.6. At

first, CAS catalyzes the hydroxylation of a monocylic fl-lactam to give a

11

 

 

911.1

3111'

CH

 

 

guanidine-containing alcohol which is hydrolyzed by proclavaminate .
amidinohydrolase to yield proclavaminic acid. CAS then catalyzes an oxidative
cyclization reaction to give dihydroclavaminic acid which is subsequently desaturated
by CAS to form Clavulanic acid. There are two functionally identical isozymes (CASl
and CASZ) with a sequence identity of 87% from Streptomyces clavuligerus and one
single isozyme from Streptomyces antibioticus mostly used to carry out the
trifunctional activities (66, 67). CAS] was used to obtain an X-ray crystal structure of
CAS-Fe(II)-aKG-substrate (6). It showed that iron is ligated by the side chains of
Hisl44, Glu146 and His279. Of interest, the presence of a glutamyl rather an aspartyl
iron ligand is pretty rare in this superfamily and this might reflect the need for
flexibility to perform its trifunctional role since glutamate is generally less rigid than
aspartate. (XKG binds Fe(II) via its C-1 carboxylate and C-2 keto groups and interacts
with Arg293 and Thrl72 though its C-5 carboxylate. N-OL-acetyl-L-arginine and
proclavaminic acid were used to study the enzyme-substrate binding, with subtle
differences observed in the structures that provide some insight into the mechanisms
of oxidative cyclization and desaturation.

Flavanone 3-hydr0xylase

Flavonoids, including flavones, isoflavones, flavonols, anthocyanins, and other
compounds, are widely distributed in plants where they fulfill many functions
including producing yellow or red/blue pigmentation in flowers and protecting them
from attack by microbes and insects (68). Flavanone 3,6-hydroxylase (F3H) catalyzes

one of the key steps of flavonoid biosynthesis by hydroxylating (ZS)- flavanones to

12

FIGURE 1.6: Reactions of Clavaminate synthase (CAS).

0 0

EN COOH Ef COOH

NHZ CAS 7 NH2 OH

Mg 7? m

0, + aKG CO2 + succinate H

Deoxyguanidinoproclavaminic acid Guanidinoproclavaminic acid
0 0 Ffo
N
CE .0... if 0
CAS 0 OOH CAS COOH

02 + aKG CO2 + succinate 01 + “KG C02 + succhateNH

HzN 2 .

H2

Prodavaminic acid Clavulanic acid

FIGURE 1.7: General reaction of Flavanone 3B-hydroxylase (F3H).

OH OH
H0 ' 0 “$0 H O O “NO

F3H

OH 0 O
02 + “KG CO2 + succinate 0H

Flavanone trans-Dilly droi'iavonol

FIGURE 1.8: General reaction of TauD.

SO ’ “ND 30 ‘ s ta
HZN’V 3.7T. HzN/Y 3.32232“. HzN/VO+ H2303

02 + aKG C02 + succhate OH

Taurine Aminoacetaldehyde 3mm

13

(2R,3R)-dihydroflavonols (69). F3H activity was first demonstrated in crude extracts
of Matthiola incana flower buds (70). The enzyme expressed in E. coli was purified
from Petunia hybrida in 1986 and for the first time, this recombinant F3H was shown
to be an F e(II)/0tKG-dependent dioxygenase (71), as shown in Figure 1.7. In 2000,

recombinant P. hybrida enzyme was purified by a more convenient two—step protocol
with higher yield (72). Mutagenesis studies of the recombinant P. hybrida gene
indicated that HisZ20, Asp222, and Hi3278 comprise the iron binding site, while

Arg288 and Ser290 assist in binding aKG (8, 73).

Synthesis or Decomposition of Small Molecules

T aurine/aK G dioxygenase

In the absence of sulfate, E. coli can utilize aliphatic sulfonates, such as
ethanesulfonate, butanesulfonate, L-cysteate, isethionate (2-hydroxylethanesulfonate),
and taurine (2-aminoethanesulfonate), as sulfur sources for growth (74, 75). The
tauABCD gene cluster on the E. coli chromosome is involved in the utilization of
taurine as sulfur source with tau/18C encoding an uptake system for taurine and the
tauD gene product showing similarity to Fe(II)/0LKG-dependent dioxygenases (I3,
76). Biochemical characterization confirmed that TauD belongs to this superfamily
(77) and extensive kinetic, structural and spectroscopic studies have been carried out
since then. Indeed, TauD is the most well—characterized member of this enzyme
family. This enzyme hydroxylates taurine to create an unstable intermediate that
decomposes to aminoacetaldehyde and sulfite (77), which is subsequently used as a

sulfur source. Substrate hydroxylation is coupled to the oxidative decarboxylation of

14

 

 

uk

311.:

iii“

It“

01..

C-

P1

511

 

 

aKG to succinate and C02 as shown in Figure 1.8. The X-ray crystal structure of
anaerobic TauD in the presence of aKG and taurine reveals a pentacoordinate ferrous
iron bound to the protein through a 2-His-1-carboxyiate facial triad made up of
residues His99, AsplOl and His255 (78, 79). The orKG is bound in a bidentate
manner with the C-1 carboxylate and the C-2 keto groups coordinated to Fe, while the
C-5 carboxylate forms a salt bridge with Arg266 and a hydrogen bond with Thr126.
Primary substrate binding induces protein conformational changes (79). More detailed
structural and mechanical studies will be discussed in the later sections of this chapter.

2, 4-Dich10r0phen0xyacetate /aK G dioxygenase

The 2,4-dichloropheoxyacetic acid (2,4-D) biodegradation pathway has been
studied as a model for microbial decomposition of chloroaromatic compounds. In
Alcaligenes eutrophus, there are five genes, tfdABCDE, responsible for this pathway
and the tfdA gene product (deA) catalyzes the hydroxylation of 2,4-D with
subsequent decomposition of the hydroxyl-intennediate to yield glyoxylate and.
2,4-dichlor0phenol, coupled with oxidative cleavage of aKG to CO2 plus succinate
(13, 76, 80), as shown in Figure 1.9. deA has a relatively broad substrate range,
including other phenoxyacetic acid derivatives, thiophenoxyacetic acid,
naphthoxyacetic acid, benzofuran-2-carboxylate, and various cinnamic acids (81). Of
interest, some orthologs from Alcaligenes denitrificans, Sphingomonas
herbicidovorans MG, and Delftia acidovorans MCI had shown stereospecficity to
degradation when examined with enantiomeric 2-phenoxypropionate analogues

(82-85). Biophysical methods had been applied to study the coordination chemistry of

15

Fe(II)- and Cu(II)-bound deA, including electron paramagnetic resonance (EPR),
electron spin-echo envelope modulation (ESEEM) and X-ray absorption spectroscopy
(XAS) (86-89). On the basis of observations by EPR and ESEEM, the presence of two
histidine ligands was proposed, with the ligand geometry undergoing changes upon
co-substrate and substrate binding (87). Mutagenesis studies indicated that Fe(Il) is
coordinated by Hisll4, Aspll6, and Hi5263, with Arg274 assisting in binding of
orKG (88, 89). A deA model was constructed using the TauD structure as a template,
since these two are closely related proteins (~30% sequence identity) (78). The model
is consistent with mutagenesis and spectroscopy studies.

Xanthine hydroxylase

Most organisms that metabolize xanthine possess a molybdopterin cofactor
(Moco)-containing enzyme that hydroxylates the substrate to form uric acid while
transferring electrons to NAD (xanthine dehydrogenase) or oxygen (xanthine oxidase)
(90), Figure 1.10. These enzymes are conserved throughout living organisms,
including archaea, bacteria, fungi, plants, and metazoans. In 2005, a novel mechanism
for xanthine metabolism was discovered in certain fungi (91). All mutants of
Aspergillus nidulans defective in xanthine dehydrogenase (i.e., with mutations
affecting the structural gene th, the cnx genes for Moco synthesis, or th for
sulfuration of Moco) retained the ability to grow on xanthine as sole nitrogen source.
Genetic studies revealed that mm! was responsible for this capability. The XanA
sequence is 18% identical to TauD, suggesting that it is an Fe(II)/OLKG-dependent

dioxygenase and is the first member of this superfamily to hydroxylate a free purine

I6

FIGURE 1.9: General reaction of deA.

OO’ 00’

T. .1.
o 1;“. 'f :3“:

l
02 + aKG C0, + succinate

2.4—D 2.4-Dichlorophenol Glyoxylate

FIGURE 1.10: General reaction of Moco- containing xanthine oxidase (X0).

(it , ,fi ,___ £1}

Hypoxanthine Xanthine Uric acid

FIGURE 1.11: General reaction of isopenicillin N synthase (IPNS).

 

H H
,N SH ,N s
fiJ— R ‘m
0 N . IPNS 04"” _
COOH 7 3 COOH
0, 211,0

FIGURE 1.12: General reaction of 4-hydroxyphenylpyruvate dioxygenase (HPPD).
COOH

0 OH
0 0
OH 0’ CO” CH

17

 

 

11:1

51

C1“

0

 

base (91). This thesis will present the purification and extensive kinetic and
spectroscopic studies on this enzyme to understand its biochemical characteristics and

coordination chemistries at different stages of enzymatic reaction.

RELATED ENZYMES

The following section will focus on several enzymes that function independently
of aKG but are related to the Fe(II)/0tI(G-dependent hydroxylases by sequence or
mechanism.

Isopenicillin N synthase

Isopenicillin N synthase (IPNS) is a non-heme Fe(II)-dependent enzyme found in
fl-lactam antibiotic-producing microorganisms. It catalyzes the formation of
isopenicillin N from 5-(L-a-aminoadipoyl)-L—cysteinyl-D-valine (ACV) (92, 93), as
illustrated in Figure 1.1]. IPNS is structurally related to the Fe(II)/0tKG-dependent
hydroxylases, but it does not utilize orKG as a cosubstrate. Furthermore, unlike
Fe(II)/0tKG-dependent dioxygenases which incorporate the elements of dioxygen into
their substates, the two oxidative ring closures of ACV forming ,B-lactam and
thiazolidine rings catalyzed by IPNS result in the complete four-electron reduction of
1 equiv of dioxygen to 2 equiv of water. The monomeric enzyme has been purified
from a variety of fungi and bacteria (94-96). The first cloning, characterization and
expression in E. coli of the gene encoding the IPNS protein in Cephalosporium
acremom'um was reported in 1985 (97), and studies on the mechanism of the action of
IPNS have focused mainly on the structure of the ferrous iron active site and its role

in catalysis. Mdssbauer, EPR and UV-Vis spectra all show substrate perturbation of

the iron site, and nitric acid (NO) was used to complex with iron to create EPR and
UV-Vis detectable species (98, 99). UV-Vis absorption spectra suggested a possible
thiol coordination to iron of the IPNS-Fe(II)-ACV complex (98). High spin Fe(II) was
indicated by the Mossbauer spectrum of IPNS-Fe(II) and IPNS-Fe(II)-ACV (98).
ESEEM was used to study the active site structure of the IPNS from Cephalosporium
acremonium with Cu(II) as a spectroscopic probe (99). This study revealed two nearly
magnetically equivalent, equatorially coordinated His ligands, whereas lH nuclear
magnetic resonance spectra of Fe(II) and Co(II) derivatives indicated the likely
presence of three His ligands. Also, a comparison study of ESEEM spectra of Cu(II)
IPNS in D20 and H20 suggested that water is a ligand of Cu(II) and this is displaced
on the addition of substrate. In 1993, Fe K-edge X-ray absorption studies provided
more insight into the iron coordination environment with three His ligands and one
carboxylate associating with iron (100, 101). In the presence of substrate, an Fe-S.
interaction was observed indicating the coordination of substrate cysteine thiolate to
the metal center. The first crystal structure of recombinant Aspergillus nidulans IPNS
was obtained in complex with manganese and determined at a resolution of 2.5 A
(102). Eight of the ,B-strands fold to form a jelly-roll motif where the active site is
buried. The metal is attached by four protein ligands (His214, Asp216, His270, and
Gln330) in the absence of substrate. Of interest, Gln330 is not conserved in
Fe(II)/orKG-dependent dioxygenases and is only observed in IPNS. This might reflect
the differences in their mechanism and substrate specificity, in particular the fact that

IPNS does not use an a-keto acid as a co-substrate. Additional mutagenesis studies

19

have shown this residue is not important to catalysis (103), and a more recent crystal

structure has shown that this Gln residue is replaced by substrate binding (104). In

particular, a thiol-metal bond is formed in the IPNS-Fe(II)-ACV complex.
4-Hydr0xyphenylpyruvate diorvgenase

4-Hydroxyphenylpyruvate dioxygenase (HPPD) is a non-heme Fe(II) oxygenase
that catalyzes the conversion of 4-hydroxyphenylpyruvate to homogentisate (105) as
illustrated in Figure 1.12. This reaction involves decarboxylation, substituent
migration, and aromatic oxygenation. It shows close chemical parallels to
Fe(II)/orKG-dependent dioxygenases but is not related in sequence to this superfamily.
Rather, the motif, HX-30HX230E, is more similar to that of extradiol dioxygenases
HX265HX250E (105). (xKG is not used as a co-substrate and both atoms of dioxygen
are incorporated into the aromatic product. This conversion is the second step of a
catabolic pathway of tyrosine and has both agricultural and therapeutic significance
(106).

This enzyme has been purified from human as dimers and from bacteria as
tetramers (107-109). Aerobic purification of HPPD from Pseudomonas sp. strain P. J.
874 was blue (110). It is found that this 595 nm absorbance is linked to enzyme-bound
Fe(III) and the reduction of iron, which caused the disappearance of the color, restores
enzyme activity. This form of enzyme was investigated by resonance Raman
spectroscopy, indicating that this absorbance probably arises from coordination of an
active site Fe(III) atom by the hydroxyl oxygen atom of tyrosine in the phenolate form

(111). The crystal structure of Pseudomona fluorescens HPPD was solved at 2.4 A,

20

and the active site metal ion of HPPD was found to be surrounded by a bower of
B-sheet structure with iron liganded to the side chains of Hisl61, Hi3240 and Glu322
(112). Significantly, no tyrosine residue side chain is within contact distance of the
active site metal ion. This structure raises the possibility of self-hydroxylation of
nearby Phe residues to account for the spectroscopic results. Four fully conserved
phenylalanines lie close to the active site, and it was postulated that they could be
hydroxylated then subsequently ligate to the metal ion (113). The structure of
substrate binding to the active site is currently unknown, but a hypothetical mode of
binding was proposed by Serre et al.(112). Similar to Fe(II)/OLKG-dependent
dioxygenases, the or-keto acid moiety may bind the metal ion in a bidentate fashion,
which is consistent with results from the UV-Vis spectrum reported by Moran et al. in
2003 (114). A different binding mode was proposed based on the crystal structure of
HPPD doped with inhibitor NTBC (115, 116). In that proposal, the substrate (Jr-keto
acid moiety makes bidentate contact with the active metal ion; however, the phenol is

sandwiched between two conserved phenylalanine rings.

STRUCTURAL STUDIES OF F e(II)/aKG-DEPEN DENT HYDROXYLASES
Crystal structures have been determined for more than twenty
Fe(II)/OLKG-dependent dioxygenases and related enzymes as briefly introduced in the
previous sections. These structural studies provide valuable information to understand
the enzyme reaction mechanisms. Here, I will further discuss these structural studies,

highlight the similarities, point out the differences, and relate these analyses to

21

enzyme mechanism investigations.

All identified Fe(II)/(xKG-dependent dioxygenases appear to contain a DSBH (or
jellyroll) fold. The DSBH topology normally consiSts of eight B-strands that form a
B-sandwich structure comprised of two four-stranded antiparallel B-sheets, which
. twist in a right-handed fashion as shown in Figure 1.13. Sequentially adjacent strands
of the DSBH are in different sheets with the exception of the “hairpin” connected
fourth and fifth B-strands (117). In some members, such as deacetoxycephalosporin C
synthase (DAOCS), which catalyzes the conversion of penicillin N to
deacetoxycephalosporin C, this connection is a tight loop (118). But in the others,
such as TauD and CAS, this connection has a highly extended conformation which is
able to form part of the substrate binding pocket (6, 78). Crystallographic analyses
have revealed that the DSBH is stabilized by hydrophobic interactions and is
supported by conserved (Jr-helices, which pack alongside the major B-sheet. DSBH
enzymes can exist in various oligomeric forms. For examples, IPNS and CAS are
monomeric proteins (6, 102); DAOCS equilibrates between monomeric and trimeric
species in solution, but crystallizes as a trimeric protein (119); FIH and TauD are
dimeric proteins (5, 34, 78, 79); AtsK, catalyzing the oxygenolytic cleavage of a
variety of different alkyl sulfate esters to the corresponding aldehyde and sulfate, is
tetrameric (120) and CarC, which is involved in carbazoledegradation, is hexameric
(121).

The DSBH appears to act as a stable platform to anchor the Fe(II) and orKG The

metal ion is ligated by the three residues forming a conserved HXD/EXnH motif,

22

 

\l

 

 

which is located within the minor B-sheet (122). (The eight B-sheets can be
distinguished by two kinds (i) minor (II-strands 2, 7, 4, 5), referred to as minor due to
its shorter strand length (ii) major (B-strands 1, 8, 3, 6), which is often extended with
additional strands. See Figure 1.13. The HXD/E of the motif is located at the end of
the second strand of the DSBH and the distal histidine of the motif is found on the
seventh strand of the DSBH. This 2-His-1-carboxylate motif has been termed a facial
triad and also is found in other non-heme oxygenases, including the catechol
dioxygenases and the cyclooxygenases (123, 124). Exceptions to this arrangement are
known as in the enzyme that catalyzes hydroxylation of aspartate and asparagines
residues in epidermal growth factor domains where the distal His appears to be
missing (125).

Compared to the nearly identical arrangements of the three amino acid side
chains that bind to the metal, the (xKG binding mode is less rigid and resolved into
two distinct categories. In both cases, the C-1 carboxylate and C-2 keto groups chelate
the metal and the C-5 carboxylate is stabilized by a salt bridge to an Arg residue or by
ionic interaction with a Lys side chain. In all reported structures, the 2-keto group .
coordinates the iron in the position trans, or approximately trans, to the Asp or Glu
residue of the HXE/DXnH motif (122, 126). However, the coordination position of the
l-carboxylate of (xKG varies between being trans to the proximal histidine residues or
trans to the distal histidine of the triad. One category is represented by CAS, TauD
and FIH (6, 34, 78). In these enzymes, the 1-carboxylate of aKG is trans to the

proximal histidine, with the open coordination site either occupied by a water

23

molecule or vacant, as seen in Figure 1.14A. Primary substrate binding near the active
site could replace this water molecule so oxygen could bind in this position. In this
case, the open metal coordination site exists near the primary substrate, called the “in
line” binding mode. By contrast, some enzyme representatives have the open
coordination site oriented away from the substrate binding site, a situation known as
the “off line” binding mode, shown in Figure 1.148. In this category, the
l-carboxylate of OLKG is trans to the distal histidine and the vacant site is trans to the
distal histidine, represented by CarC and DAOCS (121, 127, 128). Sometimes, these
two modes are exchangeable. This aKG binding flexibility reflects that different
protein structures tend to stabilize one mode over the other. For example, the C-1
carboxylate of orKG flips from the “in line” to the “off line” mode when Fe(II) is
replaced by sodium in AtsK (120). For CAS, the addition of NO to
substrate-aKG-Fe-ACS complex causes the C-1 carboxylate of OLKG to flip from the
“in line” to the “off line” binding mode, as shown in Figurel.l4 (129). Significantly,
this discovery suggests that ligand migration involving an intermediate species may
be a general feature of Fe(II)/orKG-dependent dioxygenase reaction chemistry.
Structural studies indicate that orKG typically binds prior to the primary substrate,
but they also provide insight about the releaseof the products, whether C02 and
succinate dissociate during turnover, or they depart only after turnover is complete.
First, there are some contradictary Opinions on the existence of ferryl-oxo
intermediate with C02 bound. On one side, stoichiometric accumulation of CO2 was

detected before the completion of the catalytic cycle (130). The crystal structure of a

24

FIGURE 1.13: Double-stranded B-helix structure. A, Eight B-strands forming a
double-stranded B-helix (DSBH or jellyroll). B, cartoon of the DSBH motif better

illustrating the connections between the strands, modified from (131).

 

FIGURE 1.14: aKG binding modes. A , on-line, and 13, off line. The encircled S

represents the substrate binding site.

@ °
,0- o

   
  
  

 

A8P1o1 ”’11,,” x/ 9.
Fe'sl.‘ Ashes/0,,“ 5"
His” \‘ / Fe..\
O / ~
Hiszss “19133 H154“ 0
COO'
TauD coo- DAOCS
(9 69 °
.0' O
A5on1I/I , x" -
”1..., n. q
“x A391030 , 1
His,” \0/ " I'Fe'L‘
“'3279 ”[8101/ l 0
”15251 COO"
FIH coo- CarC
C9 °
.0" O
GIUj“ III/”h "'4’ ' .
'FQL~ Glu1“/,,,lh El"
[“51“ I \~ / ,Fe~~
”'8279 o HIS1“ H. I
I8219
COO‘
CAS coo. CAS-N0

A B

25

of a C-terminal truncated mutant of DAOCS complexed with Fe(II), succinate, and
unhydrated linear CO2 was determined (128), supporting the proposals for a
metal-bound CO2 intermediate during catalysis.This evidence suggests that Fe-C02
may be relatively stable and the CO2 is not always immediately released. Furthermore,
a self-inactivation reaction of TauD was affected by the addition of bicarbonate (132),
suggesting another mechanism of C02 binding to the metallocenter. 0n the other side,
computational studies suggest that the proposed ferryl-oxo intermediate can not form
with C02 bound. (133). Early steady state investigations indicated that CO2 is the first
product to dissociate, followed by succinate (108, 126, 130, 134, 135). In 2006,
co-crystallization of AlkB with succinate yielded a structure with monodentate
coordination of Fe by the product plus a second water molecule bound at the site
previously occupied by the C-1 carboxylate of orKG, supporting the proposal that the
release of CO2 from orKG occurs prior to release of succinate during turnover, and in
some cases its release is suggested to allow migration of the ferryl-oxo intermediate to
oxidize substrate (63). Succinate is believed to leave the active site after C02 departs,
and this usually occurs after primary substrate reactivity. An interesting exception
involves DAOCS, where the enzyme-substrate complex structure could only be
obtained in the absence of OLKG and succinate and the OLKG decomposition is thought
to occur temporily separated from substrate hydroxylation (128). Existence of a
product complex also is reported in the non-(xKG enzyme HPPD, and product release
is the rate-limiting step as observed for TauD (130, 136).

Another interesting structure feature for this enzyme family is the primary

26

substrate binding mode. There is more variation in the way the substrate binds than is
apparent in either Fe(II) or aKG binding. The substrates range from small molecules,
such as taurine, to large polymers, such as proteins or DNA/RNA. With the available
evidence, there is no clear link between the structural sub-family and the type of
reaction catalyzed. However, there are some common characteristics of substrate
binding. First, for some small molecule dioxygenase, such as DAOCS, the sequence
and length of the flexible C-terminus is important for substrate selectivity. For
example, mutation of the C-terminus of DAOCS, which has a polar side-chain,
enabled it to bind penicillin N, whereas, mutation to a hydrophobic side chain favored
selectivity toward penicillin G (137). Further mutation of the C-terminus resulted in
uncoupling of OLKG decomposition from substrate oxidation (119). Second, the
addition of primary substrate induces movement of the flexible elements of the
enzyme. For example, in IPNS Arg279 changes its direction from pointing towards
the exterior of the protein to one orientated in position to bind the C-terminal
carboxylate of the substrate (102, 104). Another example is TauD, which undergoes
dramatic changes of secondary structures upon adding primary substrate. In the
absence of taurine, several key hydrogen bonds in the active site are lost resulting in
the releases of a random coil region (79). Third, some large substrates can change

their conformations to fit into the active site better, such as FIH-(x (34).

MECHANISTIC STUDIES OF Fe(II)/aKG-DEPENDENT HYDROXYLASES

Prior to the detailed descriptions for each intermediate, I would like to briefly

27

summarize the reaction mechanism of Fe(II)/0tKG-dependent hydroxylases, highlight
some common features and point out uncertainties. Hydroxylation is the most
common reaction type catalyzed by this superfamily, so it is chosen to represent the
general mechanism.

In essence, substrate is oxidized by dioxygen coupled with oxidative
decarboxylation of orKG to generate succinate and C02, with one of the oxygen atoms
incorporated into succinate and the other into the product hydroxyl group. In all cases,
Fe(II) binds to the active site before the co-substrates and substrates, as shown by
Figure 1.15A. Usually, the addition of substrates begins with the binding of OLKG,
Figure 1.15B, with two H20 molecules being displaced (138-141). It was proposed
that subsequent addition of the primary substrate displaces another H20 and creates a
site for oxygen binding; this mechanism will limit the ability of the enzyme to reduce
dioxygen until such time that all organic substrates are appropriately localized and
oriented with respect to the active-site metal ion and thus avoids uncoupled or
abortive reactive intermediate pathways (142). The primary substrate isproposed to
coordinate in the vicinity of, not directly to, the metal ion, as shown in Figure 1.15C,
creating a five-coordinated Fe(Il) site (6, 78, 143). Subsequent binding of dioxygen
leads to an Fe(III)-superoxo or Fe(IV)-peroxo species, Figure 1.15D, the anion of
which attacks the 2-oxo group of aKG, which is activated by the Lewis acidity of the
iron (126). This reaction can lead to bicyclic peroxide or a persuccinate species Figure
1.15E (126). Notably, hybrid density-functional theory calculations have supported

the existence of such a persuccinate species (144). The ferryl-oxo species, Figure

28

1.15F, is formed after the collapse of the bicyclic peroxide or persuccinate and is
thought to oxidize the substrate by radical rebound-type chemistry, restoring the
ferrous species (Figure 1.15, F to A) (126).

Intermediate A is the resting state of enzyme, in which Fe(II) is six-coordinate
with the 2-His-1-Asp/Glu facial triad ligands and three H20 molecules that are clearly
revealed from the crystal structure of holoenzyme DAOCS (118). Until now, only two
structures of unliganded holoenzyme have been published: those for DAOCS and
proline 3-hydroxylase (P3H) (7, 118). Near-infrared magnetic circular dichroism
(MCD) spectroscopy of CAS detected two d—>d transitions belonging to the
six-coordinated distorted octahedral center; these investigations also showed that
(XKG binding is reduced in the absence of metal ion, confirming that the metal ion
binds prior to the co-substrate (138, 145, 146).This octahedral arrangement of the
Fe(II) ion was also indicated by the XAS studies with deA (86). Moreover, the
Mdssbauer spectrum of 57Fe-TauD shows a signal with an isomer shift 5: 1.27 10.05
mm/s and quadrupole splitting AEQ = 3.06 10.05 mm/s, consistent with high-spin
F e(II) in the active site (143). In summary, the holoenzyme species of this superfamily
contains a distorted six-coordinated high-spin Fe(II) ion center, surrounded by two
His, one Asp/Glu and three water molecules.

Two of the solvent molecules were replaced upon binding of aKG which
exhibits bidentate interactions with the iron atom via C-1 carboxylate and C-2 keto

groups, Figure 1.15B. Crystal structures showing this bidentate association of orKG

29

FIGURE 1.15: Proposed mechanism for the hydroxylation reaction catalyzed by

Fe(II)/aKG-dependent dioxygenase. See text for a description of the various

intermediates.

xi

”.1

o: A .0 o
9:38... s o. o 92>”? ma...

. \..mo....
0 o m :.a\ \ 0\1/
I.“ Inc
o 80.
80 ~ 1%

>0.E>mu.\..0 ...u ._ ..... l1~o
I.u\ ~0/ InO
....u

.../h...

:0.

QE~>GU ....... ...
I.m\m,o. .0 til
I.»

 

 

 

 

 

a... c». P: 0w 0. o 1
on 9:32.... ..u ...... o o 9:33. ...... no.2. o
o.
\ \
o z.u\\ 1.9.0.1 :.m\ :80
80. r 000. 80..
j
$10.0 . c at o
. O\
0:553. ...... .102 055$... ..._<./...01110 0 m
I-“\ ~lo I—uvm‘al DEV/P/
:5 .. 000. 15 000.

 

X

 

are available for many enzymes, such as FIH, CAS, DAOCS and AtsK (5, 6, 118,
120). The five-membered ring formed by OLKG binding to the metal is associated with
the metal-to—ligand charge-transfer (MLCT) absorption transitions with visible
maxima around 500 nm that can be observed in the absence of oxygen. These
transitions are now accepted as characteristic of bidentate (IKG binding to the metal
ion even though the intensities and maxima of these transitions differ slightly in each
enzyme, indicating differences in overlap between d orbitals of Fe and 1t. orbitals of
OtKG among different enzymes (142). The lilac-colored chromophore exhibits a 3......
at 530 nm for both (xKG-Fe(II)-TauD and orKG-Fe(II)-deA (140, 141), whereas the
feature is observed at 500 nm in orKG-Fe(II)-AlkB (I). Resonance Raman (RR)
spectroscopy detected two vibrational transitions with the complex orKG-Fe(II)-TauD
in H20: 470 cm"I indicative of metal-ligand stretching vibrations, and 1688 cm'1 due
to the C=0 vibration of orKG (147). There are two types of coordination chemistry
shown in Figure 1.15 for the OtKG bound state, intermediate B (in line) and B’(off
line). As described before, the C-1 carboxylate is trans to the proximal His ligand for
the in-line binding mode while it is trans to the distal His ligand for the off-line
binding mode. Under the off-line binding mode, the ferryl-oxo initially is oriented
away from the substrate binding site. In order to accomplish substrate oxidation,
migration is required of either the metal ligands or the ferry] intermediate in the later
step of the catalytic cycle.

The primary substrate is bound nearby, but not in contact with, the metal ion at

the active site, Figure 1.15C, however, substrate binding has a direct influence on the

31

metal’s coordination number. Crystal structures of substrate-(XKG-Fe(II)-enzyme
complex has been resolved for a few proteins, such as FIH, CAS, TauD and AlkB (6,
34, 78, 79). These structures indicate that substrate binding induces a conversion from
six-coordinate octahedral to five-coordinate square-pyramidal geometry by the release
of the last H20 molecule. CAS is an exception by only exhibiting an enlonged metal
to H20 bond distance (6). The departure of the H20 molecule is critical for enzyme
activity since 02 is supposed to bind in this open site to carry out the hydroxylation
reaction. It was shown that poor substrate binding sometimes retains six-coordination
while inclusion of native substrate almost invariably induces five-coordinate
rearrangements (6, 78, 121). One exception is AlkB, where the substrate-bound
enzyme retains a six-coordinate metal site (63, 148). Multiple types of spectroscopies
have been applied to monitor the effects of primary substrate binding on the active
site in the presence of orKG Upon substrate binding, the UV/Vis spectra often exhibit
a perturbation of the MLCT features including a slight increase in intensity, greater
resolution of the transitions, and a blue shifi (resulting in maxima at 520 nm for TauD
and 515 nm for deA) (140, 141). For TauD, RR also exhibited 10-cm’l shift in
features to 470 cm" and 1688 cm'I in the presence of taurine (1 4 7); this result was
attributed to a switch from six-coordination to five-coordination. Also, the Mbssbauer
spectra of taurine-aKG-Fe(II)-TauD complex gave a signal with an isomer shift 5 =
1.16 $0.05 mm/s and quadrupole splitting AEQ= 2.76 $0.05 mm/s, in accord with a
reduction in coordination number (143). For CAS, the d—>d transitions in the presence

of substrate identified by near-infrared MCD were in agreement with a .

32

five-coordination site (145, 14G. Substrate binding in the off-line mode is shown as
Figure 1150. It was reported that at least one third of the available crystal structures
of this superfamily indicate that the sixth site on the ferrous ion in the tertiary
complex is orthogonal to the substrate (142). Crystal structure studies on CAS and
AlkB provide good examples of enzymes that likely require rearrangements around
the active site (6, 63, 129).

From intermediate C to the end of catalytic cycle is the least understood part of
the mechanism. Only one intermediate, a ferryl-oxo species, has been directly
identified. This species was first detected in TauD, but has also been observed in
prolyl-4-hydroxylase (P4H) and the halogenase Cth3 (143, 149—151). This
intermediate from TauD has been investigated most thoroughly. The presumed
ferryl-oxo species exhibited an absorption near 318 nm by UV/Vis stopped-flow
spectroscopy (143). Using rapid freeze-quench techniques to trap the intermediate and
carrying out subsequent analysis by EPR and Mbssbauer spectroscopy, a species with
isomer shift 8: 0.31 10.03 mm/s and quadrupole splitting AEQ= 0.88 10.03 mm/s
was detected and assigned to an integer spin with S2 (143). Cryoreduction of this
intermediate formed a high-spin Fe(III) species; thus, the intermediate was identified
as some type of Fe(IV) species (143). Use of C-1 deuterated taurine decreased the rate
of decay of this intermediate by 37 fold, indicating that the Fe(IV) species participates
in hydrogen abstraction from the substrate (152). Furthermore, RR and cryogenic
continuous flow were applied to directly identify this intermediate as Fe(IV)=02’ by

detecting its isotope-sensitive vibrations (787 cm'1 for '80 and 821 cm" for 160) (149).

33

The Fe-O distance of 1.62 A was determined by EXAFS, consistent with literature
Fe(IV)-oxo models (153-155). The identification of Fe(IV)=02' is a big step for
understanding the mechanism of this enzyme superfamily, but there are still many
unsolved questions. None of the 02-dependent intermediates leading to or following
Fe(IV)=02'has been identified. Even though we knew that Fe(IV)=02' is responsible
for the hydrogen atom abstraction from TauD, whether or not this highly reactive
species has the capacity to facilitate all of the observed reactions of this superfamily is
unknown. Alternatively, different enzymes might use different activated oxygen
species to carry out their unique reactions. Nevertheless, this general mechanism
involving formation of Fe(IV)=02' is very appealing and could reasonably
accommodate the desaturation, ring-expansion, epimerization, halogenation and other
reactions of this enzyme superfamily. For instance, CAS catalyzes cyclization and
desaturation reactions from proclavaminic acid. The ferryl-oxo species could abstract
the C4, (S) hydrogen as illustrated in Figure 1.16, which leads to intermediate B, with
Fe(III)-OH and the C4’-centered radical (156). An attack of the C3-bound hydroxyl at
C4'coupled with hydrogen atom transfer to Fe(III)-OH leads to the product complex
C. Desaturation can be explained in a similar way. The Fe(IV)=02' species could first
hydroxylate the substrate, followed by a dehydration reaction to form the double bond.
Alternatively, the Fe(IV)=02’ could initiate two hydrogen atom transfers from the
substrate directly to the product, Figure 1.17. Regardless of the diversity of overall
reactions (that are observed, the initial processes of hydrogen abstraction could be

quite similar for each of the enzymes. The most recently discovered subgroup of

34

Fe(II)/orKG-dependent dioxygenase, the halogenases, replace the monodentate
carboxylate of the facial triad with a halide ion, Figure 1.18. The Fe(IV)=02' is
thought to abstract hydrogen atom from the substrate, then the newly formed radical
and chloride atom will combine together to give halogenated product (151, 157).

For the “in line” mode, the Fe(IV)=02' points to the intended position of the
primary substrate allowing the hydrogen abstraction and subsequent reactions to
proceed directly. In contrast, the Fe(IV)=02' points away from the substrate for the
“ off line” mode and migration of this ferryl-oxo species or shifting of the metal
ligands is necessary. One possibility is that orKG reorients from “off line” (Figure
1.15B,) to “in line” (Figure 1.15B) binding mode, so that the subsequent steps will
follow those of Figure 1.15 B to F. Another option is that orKG retains its off-line
orientation resulting in the formation of Fe(IV)=02‘ pointing away from the substrate,
as shown in Figure 1.15 B, to F’. Release of C02 will provide an open site for one
H2O molecule to bind, resulting in the formation of dihydroxylated intermediate,
Figure 1.15F", which can lose a molecule of water to complete the ferryl-oxo
migration, Figure 1.15 F'——» F" -—>F. Subsequently, Fe(IV)=02' is positioned near the
substrate and the following steps are equivalent to those in Figure 1.15 F to A.

Currently, the crystal structures of O2 adducts have only been determined for two
mononuclear non-heme iron enzymes, naphthalene dioxygenase and
homoprotocatechuate 2,3-dioxygenase (I58, 159). Both of these proteins contain

2-His-1-carboxylate facial triad, but their proposed reaction mechanisms are totally

different from that of the Fe(II)/01KG dependent dioxygenases and neither use OtKG as

35

co-substrate. For both enzymes, in the presence of primary substrate, 02 binds to the
iron center through a side-on rather than end-on mode which is favored by theoretical
calculations (160). Even though these enzymes provide valuable information about an
02 binding mode, the Fe(II)/OtKG dioxygenases are unlikely share this side-on 02
binding fashion due to their different protein structure and reaction mechanism.

It is well known that reactive oxygen species can cause severe damage to the
cell, and effective control of these reactive species is very important in viva (161-163).
As mentioned earlier, oxygen generally is the last substrate to bind to the metal ion so
that activation of oxygen will not occur until all the necessary cofactors or
co-substrates are in the proper position. This strategy guarantees a high coupling
efficiency, so the substrate covens into product concomitant with the oxidative
decarboxylation of (xKG. This is not always the situation in vitro, where the primary .
substrates may be absent or inhibitors or poor substrates may be present. Under these
conditions uncoupled reactions may take place and lead to enzyme inactivation and
self-modification, such as reported deA, AlkB, TauD, and HPPD (1, 11], 164-168).
Aerobic purification of HPPD resulted in a blue chromophore consistent with an
Fe(III) tyrosinate and the tyrosinate probably is formed from hydroxylation of one of
the phenyl rings near the active site (110). When deA reacted with dioxygen and
(xKG in the absence of primary substrate, a weak chromophore with 11m, around 580
nm was observed (164). RR and EPR were applied to identify this species as a Fe(III)
hydroxyindolate product arising from the hydroxylation of Trp112 adjacent to the

metal ion. A similar result was also observed using TauD, which first forms a

36

FIGURE 1.16: Proposed mechanism for oxidative cyclization catalyzed by CAS.

0 o 0
,Ef COOH J:Nl COOH 1:“
Fe(1v1=o2- Fe(III)—OH- Fe(Il10H20 COOH
HO ' HO _"
NH, NH, NH:
A B C

FIGURE 1.17: Proposed mechanism for a desaturation reaction.

R, R R R
R1’ 1‘4 R1“? 3*4 R2):::
H ) H ( R1
=8 H6
Glu/Asp ‘0 (2:0 Glu/Asp}: 0.} "(0‘0 Jew/Asp

Fe'," _. ‘0
His/ I His’Hi: ‘ ’ ‘ ’
Hi His
00 00'

FIGURE 1.18: General reaction of halogenase.

X
Halogenase

1
R-CH3 7"? R-CH,

0: + “KG C0, + succinate

37

tyrosyl radical then catalyzes the self-hydroxylation of Tyr73 to form a Catechol
which binds Fe(III) to develop a chromOphore with with 11m, near 550 nm, as
identified by RR as a Fe(lII)-catecholate species (166). These enzymes in their ferric
states are inactive, but some reductants, such as ascorbate and DTT, were reported to
be able to restore part of the enzyme activity (132). Completing reaction cycles in
uncoupled turnover reactions is commonly thought of as the reason for the ascorbate
requirement of many Fe(II)/01KG-dependent hydroxylases. Of interest, representatives
of these enzymes operate in organisms where ascorbate is not present (131), so they
could not be restored to a functional state by this reductant. It’s still unclear if the
self-hydroxylation observed in vitro is of physiological relevance. It’s been suggested
that the enzyme builds in a number of sacrificial amino acids that are susceptible to
oxygenation adjacent to the active site metal ion, so that in the absence of substrates
or when substrate is not positioned properly, the enzyme internally quenches the
reactive oxygen species rather than releasing it to solvent (13]). Given the facts that
the uncoupling reactions rates are much slower than the catalytic reactions involving
substrate (132, 166), and the relatively low cellular dioxygen concentration,

self-inactivation is unlikely to be physiologically relevant in viva.

THESIS OUTLINE
The following chapters describe my studies on the purification and
characterization of recombinant xanthine hydroxylase (XanA) from Aspergillus

nidulans, including the investigation of the substrate-binding mode by site-directed

38

mutagensis and examination of the metallocenter coordination chemistry by various
kinds of spectroscopies. In Chapter 2, I describe the purification of recombinant A.
nidulans XanA as a His-tag version, and I provide evidence that XanA belongs to the
family of Fe(II)/(XKG-dependent hydroxylases. I also examine the effects of pH,
different kinds of buffers, and salt concentration on the enzymatic activity to optimize
the assay conditions. In addition, I characterize the substrate and co-substrate
concentration dependence to characterize the kinetic properties of XanA. Of interest,
comparison of XanA expressed in different hosts revealed very different quaternary
structures and posttranslational modifications. In Chapter 3, I describe more extensive
kinetic characterizations, including effects of different divalent metals, co-substrate
analogs, and substrate analogs to understand their binding modes to the active site.
Finally, I use a homology model, created by a colleague using TauD as the template,
to help understand the active site structure. The studies described in Chapter 1 and 2
were published in Biochemistry 2007, 46, 5293-5304. In Chapter 4, I describe the
preparation and characterization of mutant proteins with putative active site residues
mutated to alanine based on the homology model constructed from Chapter 3.
Xanthine, OLKG alternative substrate analogs, inhibitors and chemical regents were
applied to characterize the kinetic properties of the seven active mutants. Kinetic
parameters from single mutants were compared with wild-type XanA to gain more
insight into the substrate binding mode. The study of oxygen consumption when
assayed without primary substrate provided more information on the potential of the

relevance of these residues to the active site. Combining the kinetic characterization

39

and oxygen consumption studies, several important substrate binding interactions are
proposed. In Chapter 5, I show that binding of orKG and xanthine to anaerobic
Fe(II)-XanA generated MLCT transitions typical of this enzyme family. In addition, I
show that nitric oxide (N0), 3 good oxygen surrogate, can be used to prepare various
{FeNO}7 complexes, which were extensively characterized by various kinds of
spectroscopies, including UV/Vis, EPR, and one- and two-dimensional ESEEM. I
used these different {FeNO}7 intermediates of XanA to investigate the coordination
chemistry at different reaction stages. For comparision, I used TauD, a well
characterized member of this superfamily, as a reference to better interpret the results
of my spectroscopic studies on XanA. Finally, I provide a concluding chapter that
summarizes the remaining questions and places my investigations in broader

perspective.

40

REFERENCES

(1)

(2)

(3)

(4)

(5)

(6)

(7)

(8)

(9)

(10)

Trewick, S. C., Henshaw, T. F., Hausinger, R. P., Lindahl, T., and Sedgwick, B.
(2002) Oxidative demethylation by Escherichia coli AlkB directly reverts
DNA base damage. Nature 419, 174-178.

Kivirikko, K. I., and Pihlajaniemi, T. (1998) Collagen hydroxylases and the
protein disulfide isomerase subunit of prolyl 4-hydroxylase. Adv. Enzymol. Rel.
Areas Mol. Biol. 72, 325-398.

Ivan, M., Kondo, K., Yang, H., Kim, W., Valiando, J., Ohh, M., Salic, A.,
Asara, J. M., Lane, W. S., and Kaelin, W. G, Jr. (2001) HIFor targeted for
VHL-mediated destruction by proline hydroxylation: implications for 02
sensing. Science 292, 464-467.

Jaakkola, P., Mole, D. R., Tian, Y.-M., Wilson, M. 1., Gielbert, J., Gaskell, S. J.,
von Kn'egsheim, A., Hebestreit, H. F., Mukherji, M., Schofield, C. J., Maxwell,
P. H., Pugh, C. W., and Ratcliffe, P. J. (2001) Targeting of HIF-or to the von
Hippel-Lindau ubiquitylation complex by 02-regulated prolyl hydroxylation.
Science 292, 468-472.

Dann, C. E., III, Bruick, R. K., and Deisenhofer, J. (2002) Structure of a
factor-inhibiting hypoxia-inducible factor 1: an essential asparaginyl
hydroxylase involved in the hypoxic response pathway. Proc. Natl. Acad. Sci.
USA 99, 15351-15356.

Zhang, Z., Ren, J., Stammers, D. K., Baldwin, J. E., Harlos, K., and Schofield,
C. J. (2000) Structural origins of the selectivity of the trifunctional oxygenase
clavaminic acid synthase. Nature Struct. Biol. 7, 127-133.

Clifton, I. J., Hsueh, L.-C., Baldwin, J. E., Harlos, K., and Schofield, C. J.
(2001) Structure of proline 3-hydroxylase. Evolution of the family of
2-oxoglutarate dependent dioxygenases. Eur: J. Biochem. 268, 6625-6636.

Lukacin, R., Groning, 1., Pieper, U., and Matem, U. (2000) Site-directed
mutagenesis of the active site serine290 in flavanone 3B-hydroxylase from

Petunia hybrida. Eur: J. Biochem. 267, 853-860.

Hedden, P., and Phillips, A. L. (2000) Gibberellin metabolism: new insights
revealed by the genes. Trends Plant Sci. 5, 523-528.

De Carolis, E., Chan, E, Balsevich, J., and De Luca, V. (1990) Isolation and

41

(11)

(12)

(13)

(14)

(15)

(16)

(17)

(18)

(19)

characterization of a 2-oxoglutarate dependent dioxygenase involved in the
second-to-last step in vindoline biosynthesis. Plant Physiol. 94, 1323-1329.

Hashimoto, T., and Yamada, Y. (1987) Purification and characterization of
hyoscyamine 6B-hydroxylase from root cultures of Hyoscyamus niger L.
Hydroxylase and epoxidase activities of enzyme preparations. Eur. J. Biochem.
164, 277-285. '

Jansen, G A., Mihalik, S. J., Watkins, P. A., Jakobs, C., Moser, H. W., and
Wanders, R. J. A. (1998) Characterization of phytanoyl-coenzyme A
hydroxylase in human liver and activity measurements in patients with
peroxisomal disorders. Clin. Chim. Acta 271, 203-211.

Fukumori, F., and Hausinger, R. P. (1993) Purification and characterization of

2,4-dichlorophenoxyacetate/or-ketoglutarate dioxygenase. J. Biol. Chem. 268,
24311-24317.

Hutton, J. J., Jr., Trappel, A. L., and Udenfriend, S. (1966) Requirement for
or-ketoglutarate, ferrous ion and ascorbate for collagen proline hydroxylase.
Biochem. Biophys. Res. Commun. 24, 179-184.

Hieta, R., and Myllyharju, J. (2002) Cloning and characterization of a low
molecular weight prolyl 4-hydroxylase from Arabidopsis thaliana. Effective
hydroxylation of proline-rich, collagen-like, and hypoxia-inducible
transcription factor a—like peptides. J. Biol. Chem. 277, 23965-23971.

Kaska, D. D., Myllyléi, R., Giinzler, V., Gibor, A., and Kivirikko, K. I. (1988)
Prolyl 4-hydroxy1ase from Volvox carteri. A low-Mr enzyme antigenically
related to the or subunit of the invertebrate enzyme. Biochem. J. 256, 257-263.

Helaakoski, T., Vuori, K., R., M., Kivirikko, K. I., and Pihlajaniemi, T. (1989)
Molecular cloning of the or-subunit of human prolyl 4-hydroxylase: the
complete cDNA-derived amino acid sequence and evidence for alternative
splicing of RNA transcripts. Proc. Natl. Acad. Sci. USA 86, 4392-4396.

Annunem, P., Koivunen, P., and Kivirikko, K. I. (1999) Cloning of the
Ot-subunit of prolyl 4-hydroxylase from Drosophila and expression and
characterization of the corresponding enzyme tetramer with some unique
properties. J. Biol. Chem. 274, 6790-6796.

Veijola, J., Koivunen, P., Annunem, P., Pihlajaniemi, T., and Kivirikko, K. I.
(1994) Cloning, Baculovirus expression, and characterization of the or subunit
of prolyl 4-hydroxylase from the nematode Caenorhaditis elegans. This or
subunit forms an active OLB dimer with the human protein disulfide

42

(20)

(21)

(22)

(23)

(24)

(25)

(26)

(27)

(28)

(29)

(30)

isomerase/B subunit. J. Biol. Chem. 269, 26746-26753.

Ericksson, M., Myllyharju, J., Tu, H., Hellman, M., and Kivirikko, K. I. (1999) .

Evidence for 4-hydroxyproline in viral prOteins. Characterization of a viral
prolyl 4-hydroxylase and its peptide substrates. J. Biol. Chem. 274,
22131-22134.

‘ Lamberg, A., Pihlajaniemi, T., and Kivirikko, K. I. (1995) Site-directed

mutagenesis of the or subunit of human prolyl 4-hydroxylase. Identification of
three histidine residues critical for catalytic activity. J. Biol. Chem. 270,
9926-9931.

Myllyharju, J., and Kivirikko, K. I. (1997) Characterization of the iron- and
2-oxoglutarate-binding sites of human prolyl 4-hydroxylase. The EMBO J. 16,
1173-1180.

Pekkala, M., Hieta, R., Kursula, P., Kivirikko, K. I., Wierenga, R. K., and
Myllyharju, J. (2003) Crystallization of the proline-rich-peptide binding
domain of human type I collagen prolyl 4-hydroxylase. Acta Crystallogr: D59,
940-942.

Myllyharju, J. (2003) Prolyl 4-hydroxylases, the key enzymes of collagen
biosynthesis. Matrix Biol. 22, 15-24.

Pirskanen, A., Kaimio, A. M., Myllyla, R., and Kivirikko, K. I. (1996)
Site-directed mutagenesis of human lysyl hydroxylase expressed in insect cells.
Identification of histidine residues and an aspartic acid residue critical for
catalytic activity. J. Biol. Chem. 271, 9398-402.

Walsh, G, and Jefferis, R. (2006) Post-translational modifications in the
context of therapeutic proteins. Nat. Biotechnol. 24, 1241-52.

Semenza, G L. (2000) HIF-1 and human disease: one highly involved factor.
Genes Dev. 14, 1983-91.

Schofield, C. J., and Ratcliffe, P. J. (2004) Oxygen sensing by HIF
hydroxylases. Nat. Rev. Mol. Cell Biol. 5, 343-54.

Hewitson,iK. S., McNeil], L. A., Elkins, J. M., and Schofield, C. J. (2003) The
role of iron and 2-oxoglutarate oxygenases in signalling. Biochem. Soc. Trans.

31,510-515.

Bruick, R. K., and McKnight, S. L. (2001) A conserved family of
prolyl-4-hydroxylases that modify HIF. Science 294, 1337-1340.

43

(31)

(32)

(33)

(34)

(35)

(36)

(37)

(38)

(39)

(40)

Hewitson, K. S., McNeil], L. A., Riordan, M. V., Tian, Y.-M., Bullock, A. N.,
Welford, R. W., Elkins, J. M., Oldham, N. J., Bhattacharya, S., Gleadle, J. M.,
Ratcliffe, P. J., Pugh, C. W., and Schofield, C. J. (2002) Hypoxia-inducible
factor (HIF) asparagine hydroxylase is identical to factor inhibiting HIF (FIH)
and is related to the cupin structural familly. J. Biol. Chem. 2 7 7, 26351-26355.

Lando, D., Peet, D. J., Whelan, D. A., Gonnan, J. J., and Whitelaw, M. L.
(2002) Asparagine hydroxylation of the HIF transactivation domain: a hypoxic
switch. Science 295, 858-861.

McNeill, L. A., Hewitson, K. S., Claridge, T. D., Seibel, J. F., Horsfall, L. E.,
and Schofield, C. J. (2002) Hypoxia-inducible factor asparginyl hydroxylase

(FIH-1) catalyses hydroxylation at the B-carbon of asparagine-803. Biochem. J.
367, 571-575.

Elkins, J. M., Hewitson, K. S., McNeill, L. A., Seibel, J. F., Schlemminger, 1.,
Pugh, C. W., Ratcliffe, P. J., and Schofield, C. J. (2003) Structure of
factor-inhibiting hypoxia-inducible factor (HIF) reveals mechanism of
oxidative modification of HIF-101. J. Biol. Chem. 278, 1802-1806.

Martin, C., and Zhang, Y. (2005) The diverse functions of histone lysine
methylation. Nat. Rev. Mol. Cell Biol. 6, 838-49.

Strahl, B. D., and Allis, C. D. (2000) The language of covalent histone
modifications. Nature 403, 41-5.

Tsukada, Y.-I., Fang, J., Erdjument-Bromage, H., Warren, M. E., Borchers, C.
H., Tempst, P., and Zhang, Y. (2006) Histone demethylation by a family of
ijC domain-containing proteins. Nature 439, 811-816.

Wang, Y., Wysocka, J., Sayegh, J., Lee, Y.-H., Perlin, J. R., Leonelli, L.,
Sonbuchner, L. S., McDonald, C. H., Cook, R. G, Dou, Y., Roeder, R. G,
Clarke, 8., Stallcup, M. R., Allis, C. D., and Coonrod, S. A. (2004) Human
PAD4 regulates histone arginine methylation levels via demethylimination.
Science 306, 279-283.

Cuthbert, G L., Daujat, S., Snowden, A. W., Erdjument-Bromage, H.,
Hagiwara, T., Yamada, M., Schneider, R., Gregory, P. D., Tempst, P., Bannister,
A. J., and Kouzarides, T. (2004) Histone deimination antagonizes arginine
methylation. Cell 118, 545-53.

Shi, Y., Lan, F., Matson, C., Mulligan, P., Whetstine, J. R., Cole, P. A., Casero,

R. A., and Shi, Y. (2004) Histone demethylation mediated by the nuclear
amine oxidase homolog LSDl. Cell 119, 941-53.

44

(41)

(42)

(43)

(44)

(45)

(46)

(47)

(48)

(49)

(50)

Metzger, E., Wissmann, M., Yin, N., Muller, J. M., Schneider, R., Peters, A. H.,
Gunther, T., Buettner, R., and Schule, R. (2005) LSDl demethylates repressive
histone marks to promote androgen-receptor-dependent transcription. Nature
43 7, 436-9.

Yamane, K., Toumazou, C., Tsukada, Y.-I., Erdjument-Bromage, H., Tempst,
P., Wong, J., and Zhang, Y. (2006) JHDM2A, a ijC-containing H3K9
demethylase, facilitates transcription activation by androgen receptor. Cell 125,
1-13.

Cloos, P. A. C., Christensen, J., Agger, K., Maiolica, A., Rappsilber, J., Antal,
T., Hansen, K. H., and Helin, K. (2006) The putative oncogene GASC]
demethylates tri- and dimethylated lysine 9 on histone H3. Nature 442,
307-311.

Whetstine, J. R., Nottke, A., Lan, F., Huart, M., Smolikov, S., Chen, Z.,
Spooner, E., Li, B, Zhang, G, Colaiacovo, M., and Shi, Y. (2006) Reversal of
histone lysine trimethylation by the JMJD2 family of histone demethylases.
Cell 125, 467-481.

Klose, R. J ., Yamane, K., Bae, Y., Zhang, D., Erdjument-Bromage, H., Tempst,
P., Wong, J., and Zhang, Y. (2006) The transcriptional repressor JHDM3A
demethylates trimethyl histone H3 lysine 9 and lysine 36. Nature 442,
312-316.

Fodor, B. D., Kubicek, S., Yonezawa, M., O'Sullivan, R. J., Sengupta, R.,
Perez-Burgos, L., Opravil, S., Mechtler, K., Schotta, G, and Jenuwein, T.
(2006) ijd2b antagonizes H3K9 trimethylation at pericentric
heterochromatin in mammalian cells. Genes Dev 20, 1557-62.

Klose, R. J., Kallin, E. M., and Zhang, Y. (2006) ijC-domain-containing
proteins and histone demethylation. Nat. Rev. Genet. 7, 715-27.

Chen, Z., Zang, J., Whetstine, J., Hong, X., Davrazou, F., Kutateladze, T. G,
Simpson, M., Mao, 0., Pan, C. H., Dai, S., Hagman, J., Hansen, K., Shi, Y.,
and Zhang, G (2006) Structural insights into histone demethylation by JMJD2
family members. Cell 125, 691-702.

Sedgwick, B., and Lindahl, T. (2002) Recent progress on the Ada response for
inducible repair of DNA alkylation damage. Oncogene 21, 8886-8894.

Sedgwick, B. (2004) Repairing DNA-methylation damage. Nat. Rev. Molec.
Cell Biol. 5, 148-157.

45

(51)

(52)

(53)

(54)

(55)

(56)

(57)

(58)

(59)

(60)

(61)

Rydberg, B., and Lindahl, T. (1982) Nonenzymatic methylation of DNA by the
intracellular methyl group donor S-adenosyl-L-methionine is a potentially
mutagenic reaction. EMBO J. 1, 211-6.

Teo, 1., Sedgwick, B., Kilpatrick, M. W., McCarthy, T. V., and Lindahl, T.
(1986) The intracellular signal for induction of resistance to alkylating agents
in E. coli. Cell 45, 315-24.

Kataoka, H., Yamamoto, Y., and Sekiguchi, M. (1983) A new gene (alkB) of
Escherichia coli that controls sensitivity to methyl methane sulfonate. J.
Bacterial. 153, 1301-1307.

Chen, B. J., Carroll, P., and Samson, L. (1994) The Escherichia coli AlkB
protein protects human cells against alkylation-induced toxicity. J. Bacteriol.
176, 6255-6261.

Aravind, L., and Koonin, E. V. (2001) The DNA-repair protein AlkB, EGL-9,
and leprecan define new families of 2-oxoglutarate-dependent- and
iron-dependent dioxygenases. Genome Biology 2, 7.1-7.8.

Falnes, P. 0., Johansen, R. F., and Seeberg, E. (2002) AlkB-mediated oxidative
demethylation reverses DNA damage in Escherichia coli. Nature 419,
178-182.

Duncan, T., Trewick, S. C., Koivisto, P., Bates, P. A., Lindahl, T., and
Sedgwick, B. (2002) Reversal of DNA alkylation damage by two human
dioxygenases. Proc. Natl. Acad. Sci. USA 99, 16660-16665.

Koivisto, P., Duncan, T., Lindahl, T., and Sedgwick, B. (2003) Minimal
methylated substrate and extended substrate range of Escherichia coli AlkB
protein, a l-methyladenine-DNA dioxygenase. J. Biol. Chem. 278,
44348-44354.

Wei, Y.-F., Carter, K. .C., Wang, R.-P., and Shell, B. K. (1996) Molecular
cloning and fitnctional analysis of a human cDNA encoding an Escherichia
coli AlkB homolog, a protein involved in DNA alkylation damage repair. Nucl.
Acids Res. 24, 931-937.

Kurowski, M. A., Bhagwat, A. S., Papaj, G, and Bujnicki, J. M. (2003)
Phylogenomic identification of five new human homologues of the DNA

repair enzyme AlkB. BMC Genomics 4, 48.

Colombi, D., and Gomes, S. L. (1997) An alkB gene homolog is differentially
transcribed during the Caulobacter crescentus cell cycle. J. Bacteriol. 179,

46

(62)

(63)

(64)

(65)

(66)

(67)

(68)

(69)

(70)

(71)

(72)

3139-45.

Aas, P. A., Otterlei, M., Falnes, P. 0., Vagbe, C. B., Skorpen, F., Akbari, M.,
Sundheim, 0., Bjoras, M., Slupphaug, G, Seeberg, E., and Krokan, H. E.
(2003) Human and bacterial oxidative demethylases repair alkylation damage
in both RNA and DNA. Nature 421, 859-863.

Yu, B., Edstrom, W. C., Hamuro, Y., Weber, P. C., Gibney, B. R., and Hunt, J.
F. (2006) Crystal structures of catalytic complexes of the oxidative DNA/RNA
repair enzyme AlkB. Nature 439, 879-884.

Matagne, A., Dubus, A., Galleni, M., and Frere, J. M. (1999) The
beta-lactamase cycle: a tale of selective pressure and bacterial ingenuity. Nat
Prod Rep 16, 1-19.

Lloyd, M. D., Mem't, K. D., Lee, V., Sewell, T. J., Wha-Son, B., Baldwin, J. E.,

Schofield, C. J., Elson, S. W., Baggaley, K. H., and Nicholson, N. H. (1999)
Product-substrate engineering by bacteria: studies on Clavaminate synthase, a
trifunctional dioxygenase. Tetrahedron 55, 10201-10220.

Busby, R. W., Chang, M. D.-T., Busby, R. C., Wimp, J., and Townsend, C. A.
(1995) Expression and purification of two isozymes of Clavaminate synthase
and initial characterization of the iron binding site. General error analysis in
polymerase chain reaction amplification. J. Biol. Chem. 270, 4262-4269.

Marsh, E. N., Chang, M. D.-T., and Townsend, C. A. (1992) Two isozymes of
Clavaminate synthase central to Clavulanic acid formation: cloning and
sequencing of both genes from Streptomyces clavuligerus. Biochemistry 31,
12648-12657.

Bohm, B. A. (1998) Introduction to Flavonoids, Harwood Academic
Publishers, Amsterdam, The Netherlands.

Prescott, A. G (1993) A dilemma of dioxygenases (or where biochemistry and
molecular biology fail to meet). J. Expert Bat. 44, 849-861.

Heller, W., and Forkmann, G. (1993) The flavonaids, advances in research
since 1986, 399-425.

Britsch, L., and Grisebach, H. (1986) Purification and characterization of
(ZS)-flavanone 3-hydroxylase from Petunia hybrida. Eur: J. Biochem. 156,
569-577.

Lukacin, R., Groning, 1., Schiltz, E., Britsch, L., and Matem, U. (2000)

47

(73)

(74)

(75)

(76)

(77)

(78)

(79)

(80)

(81)

Purification of recombinant flavanone 3B-hydroxylase from Petunia hybrida
and assignment of the primary site of proteolytic degradation. Arch. Biochem.
Biophys. 375, 364-370.

Lukacin, R., and Britsch, L. (1997) Identification of strictly conserved
histidine and arginine residues as part of the active site in Petunia hybrida
flavanone 3B-hydroxylase. Eur: J. Biochem. 249, 748-757.

Uria-Nickelsen, M. R., Leadbetter, E. R., and Godchaux, W., 111. (1993)
Sulfonate utilization by enteric bacteria. J. Gen. Microbial. 139, 203-208.

Van Der Ploeg, J. R., Weiss, M. A., Saller, E., Nashimoto, H., Saito, N.,
Kertesz, M. A., and Leisinger, T. (1996) Identification of sulfate
starvation-regulated genes in Escherichia coli: 3 gene cluster involved in the
utilization of taurine as a sulfur source. J. Bacterial. I 78, 5438-5446.

Fukumori, E, and Hausinger, R. P. (1993) Alcaligenes eutraphus JMP134
"2,4-dichlorophenoxyacetate monooxygenase" is an
or-ketoglutarate-dependent dioxygenase. J. Bacteriol. 175, 2083-2086.

Eichhom, E., van der Ploeg, J. R., Kertesz, M. A., and Leisinger, T. (1997)

Characterization of a-ketoglutarate-dependent taurine dioxygenase from
Escherichia coli. J. Biol. Chem. 272, 23031—23036.

Elkins, J. M., Ryle, M. J., Clifton, I. J., Dunning Hotopp, J. 0, Lloyd, J. S.,
Burzlaff, N. 1., Baldwin, J. E., Hausinger, R. P., and Roach, P. L. (2002) X-ray‘
crystal structure of Escherichia coli taurine/a-ketoglutarate dioxygenase
complexed to ferrous iron and substrates. Biochemistry 41, 5185-5192.

O'Brien, J. R., Schuller, D. J., Yang, V. S., Dillard, B. D., and Lanzilotta, W. N.
(2003) Substrate-induced conformational changes in Escherichia coli

taurine/a-ketoglutarate dioxygenase and insight into the oligomeric structure.
Biochemistry 42, 5547-5554.

Hausinger, R. P., Fukumori, F., Hogan, D. A., Sassanella, T. M., Kamagata, Y.,
Takami, H., and Saari, R. E. (1997) Biochemistry of 2,4-D degradation:
evolutionary implications, in Microbial Diversity and Genetics of
Biodegradatian (Horikoshi, K., Fukuda, M., and Kudo, T., Eds.) pp 35—51,

Japan Scientific Press, Tokyo.
Dunning Hotopp, J. C., and Hausinger, R. P. (2001) Alternative substrates for

2,4-dichlorophenoxyacetate/or-ketoglutarate dioxygenase. J. Molec. Catalysis
B 15,155-162.

48

(82)

(83)

(84)

(85)

(86)

(87)

(88)

(89)

(90)

(91)

Tett, V. A., Willetts, A. J., and Lappin-Scott, H. M. (1997) Biodegradation of .
the chlorophenoxy herbicide (R)-(+)-mecoprop by Alcaligenes denitrificans.
Biodegradation 8, 43-52.

Nickel, K., Suter, M. J.-F., and Kohler, H.-P. E. (1997) Involvement of two
or-ketoglutarate-dependent dioxygenases in enantioselective degradation of
(R)- and (S)-mecoprop by ‘Sphingomanas herbicidavarans MH. J. Bacteriol.
179, 6674-6679.

Westendorf, A., Miiller, A., and Babel, W. (2003) Purification and
characterization of the enantiospecific dioxygenases from Delftia acidovorans
MCl initiating the degradation of phenoxypropionate and phenoxyacetate
herbicides. Acta Biotechnol 23, 3-17.

Miiller, T. A., Zavodszky, M. I., Feig, M., Kuhn, L. A., and Hausinger, R. P.
(2006) Structural basis for the enantiospecificities of R- and

S-phenoxypropionate/or-ketoglutarate dioxygenases. Prat. Science 15,
1356-1368.

Cosper, N. J., Stalhandske, C. M. V., Saari, R. E., Hausinger, R. P., and Scott,
R. A. (1999) X-ray absorption spectroscopic analysis of Fe(II) and Cu(II)

forms of an herbicide-degrading Ot-ketoglutarate dioxygenase. J. Biol. Inorg.
Chem. 4, 122-129.

Whiting, A. K., Que, L., Jr., Saari, R. E., Hausinger, R. P., Fredrick, M. A., and
McCracken, J. (1997) Metal coordination environment of a Cu(II)-substituted '

or-keto acid-dependent dioxygenase that degrades the herbicide 12,4-D. J. Am.
Chem. Soc. 119, 3413-3414.

Dunning Hotopp, J. C., and Hausinger, R. P. (2002) Probing the
2,4-dichlorophenoxyacetate/or-ketoglutarate dioxygenase substrate binding site
by site-directed mutagenesis and mechanism-based inactivation. Biochemistry

41, 9787-9794.

Hogan, D. A., Smith, S. R., Saari, E. A., McCracken, J., and Hausinger, R. P.
(2000) Site-directed mutagenesis of 2,4-dichlorophenoxyacetic
acid/Ot-ketoglutarate dioxygenase. Identification of residues involved in
metallocenter formation and substrate binding. .1. Biol. Chem. 275,
12400-12409.

Hille, R. (2005) Molybdenum-containing hydroxylases. Arch. Biochem.
Biophys. 433, 107-116.

Cultrone, A., Scazzocchio, C., Rochet, M., Montero-Moran, G, Drevet, C.,

49

(92)

(93)

(94)

(95)

(96)

(97)

(98)

(99)

(100)

and Femandez-Martin, R. (2005) Convergent evolutiOn of hydroxylation
mechanisms in the fungal kingdoms: molybdenum cofactor-independent

hydroxylation of xanthine via or-ketoglutarate dependent dioxygenase. Molec.
Microbiol. 57, 276-290.

Kreisberg-Zakarin, R., Borovok, 1., Yanko, M., Aharonowitz, Y., and Cohen, G
(1999) Recent advances in the structure and function of isopenicillin N
synthase. Antonie van Leeuwenhoek 75, 33-39.

Schofield, C. J., Baldwin, J. E., Byford, M. F., Clifton, I. J., Hensgens, C., and
Roach, P. L. (1997) Proteins of the penicillin biosynthesis pathway. Curr:
Opinion Struct. Biol. 7, 857-864.

Pang, C. P., Chakravarti, B., Adlington, R. M., Ting, H. H., White, R. L.,
Jayatilake, G S., Baldwin, J. E., and Abraham, E. P.’(l984) Purification of
isopenicillin N synthetase. Biochem J 222, 789-95.

Baldwin, J. E., and Abraham, E. (1988) The biosynthesis of penicillins and
cephalosporins. Nat. Prod. Rep. 5, 129-45.

Castro, J. M., Liras, P., Laiz, L., Cortes, J., and Martin, J. F. (1988)
Purification and characterization of the isopenicillin N synthase of
Streptomyces lactamdurans. J. Gen. Microbiol. 134, 133-41.

Samson, S. M., Belagaje, R., Blankenship, D. T., Chapman, J. L., Perry, D.,
Skatrud, P. L., VanFrank, R. M., Abraham, E. P., Baldwin, J. E., Queener, S.
W., and et al. (1985) Isolation, sequence determination and expression in
Escherichia coli of the isopenicillin N synthetase gene from Cephalasporium
acremanium. Nature 318, 191-4.

Chen, V. J., Orville, A. M., Harpel, M. R., Frolik, C. A., Surerus, K. K., Munck,
E., and Lipscomb, J. D. (1989) Spectroscopic studies of isopenicillin N
synthase. A mononuclear nonheme Fez” oxidase with metal coordination sites
for small molecules and substrate. .1. Biol. Chem. 264, 21677-21681.

Jiang, F., Peisach, J., Ming, L.-J., Que, L., Jr., and Chen, V. J. (1991) Electron
spin echo envelope modulation studies of the Cu(I)-substituted derivative of
isopenicillin N synthase: a structural and spectroscopic model. Biochemistry
30, 11437-11445.

Randall, C. R., Zang, Y., True, A. E., Que, L., Jr., Chamock, J. M., Garner, C.
D.; Fujishima, Y., Schofield, C. J ., and Baldwin, J. E. (1993) X-ray absorption
studies of the ferrous active site of isopenicillin N synthase and related model
complexes. Biochemistry 32, 6664-6673. ‘

50

(101)

(102)

(103)

(104)

(105)

(106)

(107)

(108)

(109)

(110)

(111)

Scott, R. A., Wang, S., Eidsness, M. K., Kriauciunas, A., Frolik, C. A., and
Chen, V. J. (1992) X-ray absorption spectroscopic studies of the high-spin
iron(II) active site of isopenicillin N synthase: evidence for Fe-S interaction in
the enzyme-substrate complex. Biochemistry 31, 4596-4601.

Roach, P. L., Clifton, I. J., Ffilbp, V., Harlos, K., Barton, G J., Hajdu, J.,
Andersson, K., Schofield, C. J., and Baldwin, J. E. (1995) Crystal structure of
isopenicillin N synthase is the first from a new structural family of enzymes.
Nature (London) 375, 700-704.

Landman, 0., Borovok, 1., Aharonowitz, Y., and Cohen, G (1997) The
glutamine ligand in the ferrous iron active site of isopenicillin N synthase of
Streptomyces jumonjinensis is not essential for catalysis. FEBS Lett. 405,
172-174.

Roach, P. L., Clifton, I. J., Hensgens, C. M. H., Shibata, N., Schofield, C. J.,
Hajdu, J., and Baldwin, J. E. (1997) Structure of isopenicillin N synthase
complexed with substrate and the mechanism of penicillin formation. Nature
(London) 387, 827-830.

Moran, G R. (2005) 4-Hydroxyphenylpyruvate dioxygenase. Arch. Biochem.
Biophys. 433, 117-128.

Gissen, P., Preece, M. A., Willshaw, H. A., and McKiernan, P. J. (2003)
Ophthalmic follow-up of patients with tyrosinaemia type I on NTBC. J.
Inherit. Metab. Dis. 26, 13-6.

Wada, G H., Fellman, J. H., Fujita, T. S., and Roth, E. S. (1975) Purification
and properties of avian liver p-hydroxyphenylpyruvate hydroxylase. J. Biol.
Chem. 250, 6720-6.

Lindblad, B., Lindstedt, G, Lindstedt, S., and Rundgren, M. (1977)
Purification and some properties of human 4-hydroxyphenylpyruvate
dioxygenase (I). J. Biol. Chem. 252, 5073-84.

Buckthal, D. J ., Roche, P. A., Moorehead, T. J ., Forbes, B. J., and Hamilton, G
A. (1987) 4-Hydroxyphenylpyruvate dioxygenase from pig liver. Methalds
Enzymol. 142, 132-8.

Lindstedt, S., and Rundgren, M. (1982) Blue color, metal content, and
substrate binding in 4-hydroxyphenylpyruvate dioxygenase from
Pseudomonas sp. strain P. J. 874. J. Biol. Chem. 257, 11922-31.

Bradley, F. C., Lindstedt, S., Lipscomb, J. D., Que, L., Jr., Roe, A. L., and

51

(112)

(113)

(114)

(115)

(116)

(117)

(118)

(119)

(120)

Rundgren, M. (1986) 4-Hydroxyphenylpyruvate dioxygenase is an
iron-tyrosinate protein. J. Biol. Chem. 261, 11693-11696.

Serre, L., Sailland, A., Sy, D., Boudec, P., Rolland, A., Pebay-Peyroula, E., and
Cohen-Addad, C. (1999) Crystal structure of Pseudomonas fluorescens
4-hydroxyphenylpyruvate dioxygenase: an enzyme involved in the tyrosine
degradation pathway. Structure 7, 977-988.

Ryle, M. J., and Hausinger, R. P. (2002) Non-heme iron oxygenases. Curr.
Opinion Chem. Biol. 6, 193-201.

Johnson-Winters, K., Purpero, V. M., Kavana, M., Nelson, T., and Moran, G R.

(2003) (4-Hydroxyphenyl)pyruvate dioxygenase from Streptomyces
avermitilis: the basis for ordered substrate addition. Biochemistry 42,
2072-2080.

Gunsior, M., Ravel, J ., Challis, G L., and Townsend, C. A. (2004) Engineering
p-hydroxyphenylpyruvate dioxygenase to a p-hydroxymandelate synthase and
evidence for the proposed benzene oxide intermediate in homogentisate
formation. Biochemistry 43, 663-674.

Brownlee, J. M., Johnson-Winters, K., Harrison, D. H. T., and Moran, G R.
(2004) Structure of the ferrous form of (4-hydroxyphenyl)pyruvate

dioxygenase from Streptomyces avermitilis in complex with the therapeutic
herbicide, NTBC. Biochemistry 43, 6370-6377.

Stirk, H. J., Woolfson, D. N., Hutchinson, E. G, and Thornton, J. M. (1992)
Depicting topology and handedness in jellyroll structures. FEBS Lett 308, 1-3.

Valegard, K., Terwisscha van Scheltinga, A. 0, Lloyd, M. D., Hara, T.,
Ramaswamy, S., Perrakis, A., Thompson, A., Lee, W.-J., Baldwin, J. E.,
Schofield, C. J., Hajdu, J., and Andersson, I. (1998) Structure of a
cephalosporin synthase. Nature (London) 394, 805-809.

Lloyd, M. D., Lee, H.-J., Harlos, K., Zhang, Z.-H., Baldwin, J. E., Schofield,
C. J., Chamock, J. M., Gamer, C. D., Hara, T., Terwisscha Van Scheltinga, A.
C., Valegard, K., Viklund, J. A. C., Hajdu, J., Andersson, I., Danielsson, A.,
and Bhikhabhai, R. (1999) Studies on the active site of
deacetoxycephalosporin C synthase. J. Mol. Biol. 287, 943-960.

Miiller, I., Kahnert, A., Pape, T., Sheldrick, G M., Meyer-Klaucke, W., Dierks,
T., Kertesz, M., and Uson, I. (2004) Crystal structure of the alkylsulfatase
AtsK: insights into the catalytic mechanism of the Fe(II)
tx-ketoglutarate-dependent dioxygenase superfamily. Biochemistry 43,

52

(121)

(122)

(123)

(124)

(125)

(126)

(127)

(128)

(129)

(130)

(131)

3075-3088.

Clifton, I. J., Doan, L. X., Sleeman, M. C., Topf, M., Suzuki, H., Wilmouth, R.
C., and Schofield, C. J. (2003) Crystal structure of carbapenem synthase
(CarC). J. Biol. Chem. 278, 20843-20850.

Clifton, I. J., McDonough, ‘M. A., Ehrismann, D., Kershaw, N. J., Granatino,
N., and Schofield, C. J. (2006) Structural studies on 2-oxoglutarate

oxygenases and related double-stranded B-helix fold protein. J. Inorg.
Biochem. 100, 644-669.

Broderick, J. B. (1999) Catechol dioxygenases. Essays Biochem. 34, 173-189.

Costas, M., Mehn, M. P., Jensen, M. P., and Que, L., Jr. (2004) Dioxygen
activation at mononuclear nonheme iron active sites: enzymes, models, and
intermediates. Chem. Rev. 104, 939-986.

Lancaster, D. E., McDonough, M. A., and Schofield, C. J. (2004) Factor
inhibiting hypoxia-inducible factor (FIH) and other asparaginyl hydroxylases.
Biochem. Soc. Trans. 32, 943-5.

Hausinger, R. P. (2004) Fe(II)/or-ketoglutarate-dependent hydroxylases and
related enzymes. Crit. Rev. Biochem. Mal. Biol. 39, 21-68.

Valegard, K., Terwisscha van Scheltinga, A. C., Dubus, A., Ranghino, G,
Oster, L. M., Hajdu, J., and Andersson, I. (2004) The structural basis of
cephalosporin formation in a mononuclear ferrous enzyme. Nat. Struct. Mol.
Biol. 11, 95-101.

Lee, H. J., Lloyde, M. D., Clifton, I. J., Baldwin, J. E., and Schofield, C. J.
(2001) Kinetic and crystallographic studies on deacetoxycephalosporin C
synthase (DAOCS). J. Mol. Biol. 308, 937-948.

Zhang, Z., Ren, J.-s., Harlos, K., McKinnon, C. H., Clifton, I. J., and Schofield,
C. J. (2002) Crystal structure of a Clavaminate
synthase-Fe(II)-2-oxog1utarate-substrate-NO complex: evidence for metal
centred rearrangements. FEBS Lett. 517, 7-12.

Price, J. C., Barr, E. W., Hoffart, L. M., Krebs, C., and Bollinger, J. M., Jr.
(2005) Kinetic dissection of the catalytic mechanism of
taurinezor-ketoglutarate dioxygenase (TauD) from Escherichia coli.

Biochemistry 44, 8138-8147.

Hewitson, K. S., Granatino, N., Welford, R. W., McDonough, M. A., and

53

(132)

(133)

(134)

(135)

(I36)

(137)

(138)

(139)

(140)

Schofield, C. J. (2005) Oxidation by 2-oxoglutarate oxygenases: non-haem
iron systems in catalysis and signalling. Philos. Transact. A Math. Phys. Eng.
Sci. 363, 807-28; discussion 1035-40.

Ryle, M. J., Koehntop, K. D., Liu, A., Que, L., Jr., and Hausinger, R. P. (2003)
Interconversion of two oxidized forms of TauD, a non-heme iron hydroxylase:
evidence for bicarbonate binding. Proc. Natl. Acad. Sci. USA 100, 3790-3795.

Neidig, M. L., Decker, A., Choroba, O. W., Huang, F., Kavana, M., Moran, G.
R., Spencer, J. B., and Solomon, E. I. (2006) Spectroscopic and electronic
structure studies of aromatic electrophilic attack and hydrogen-atom
abstraction by non-heme iron enzymes. Proc Natl Acad Sci U S A 103,
12966-73.

Holme, E. (1975) A kinetic study of thymine 7-hydroxylase from Neurospara
crassa. Biochemistry 14, 4999-5003.

Tuderman, L., R., M., and Kivirikko, K. I. (1977) Mechanism of prolyl
hydroxylase reaction. 1. Role of co-substrates. Eur: J. Biochem. 80, 314-348.

Johnson-Winters, K., Purpero, V. M., Kavana, M., and Moran, G R. (2005)
Accumulation of multiple intermediates in the catalytic cycle of
(4-hydroxyphenyl)pyruvate dioxygenase from Streptomyces avermitilis.
Biochemistry 44, 7189-7199.

Lee, H.-J., Lloyd, M. D., Clifton, I. J., Harlos, K., Dubus, A., Baldwin, J., E.,
Frere, J.-M., and Schofield, C. J. (2001) Alteration of the co—substrate
selectivity of deacetoxycephalosporin C synthase. The role of arginine 258. J.
Biol. Chem. 276, 18290-18295.

Pavel, E. G, Zhou, J., Busby, R. W., Gunsior, M., Townsend, C. A., and
Solomon, E. I. (1998) Circular dichroism and magnetic circular dichroism
spectroscopic studies of the non-heme ferrous active site in Clavaminate

synthase and its interaction with or-ketoglutarate cosubstrate. J. Am. Chem. Soc.
120, 743-753.

Neidig, M. L., Kavana, M., Moran, G R., and Solomon, E. I. (2004) CD and
MCD studies of the non-heme ferrous active site in
(4-hydroxyphenyl)pyruvate dioxygenase: correlation between oxygen

activation in the extradiol and alpha-KG-dependent dioxygenases. J. Am.
Chem. Soc. 126, 4486-7.

Ryle, M. J., Padmakumar, R., and Hausinger, R. P. (1999) Stopped-flow
kinetic analysis of Escherichia coli taurine/a—ketoglutarate dioxygenase:

(141)

(142)

(143)

(144)

(145)

(146)

(147)

interactions with a-ketoglutarate, taurine, and oxygen. Biochemistry 38,
15278-15286.

Hegg, E. L., Whiting, A. K., Saari, R. E., McCracken, J ., Hausinger, R. P., and
Que, L., Jr. (1999) Herbicide-degrading or-keto acid-dependent enzyme deA:
metal coordination environment and mechanistic insights. Biochemistry 38,
16714-16726.

Purpero, V., and Moran, G R. (2007) The diverse and pervasive chemistries of
the Ot-keto acid dependent enzymes. J. Biol. Inorg. Chem. 12, 587-601.

Price, J. C., Barr, E. W., Tirupati, B., Bollinger, J. M., Jr., and Krebs, C. (2003)
The first direct characterization of a high-valent iron intermediate in the
reaction of an or-ketoglutarate-dependent dioxygenase: a high-spin Fe(IV)

complex in taurine/(x-ketoglutarate dioxygenase (TauD) from Escherichia coli.
J. Biol. Chem. 42, 7497-7508.

Borowski, T., Bassan, A., and Siegbahn, P. E. M. (2004) Mechanism of
dioxygen activation in 2-oxoglutarate-dependent enzymes: 3 hybrid DFT study.
Chem. Eur: J. 10,1031-1041.

Zhou, J., Gunsior, M., Bachmann, B. 0., Townsend, C. A., and Solomon, E. I.
(1998) Substrate binding to the a—ketoglutarate-dependent non-heme iron

enzyme Clavaminate synthase 2: coupling mechanism of oxidative
decarboxylation and hydroxylation. J. Am. Chem. Soc. 120, 13539-13540.

Zhou, J., Kelly, W. L., Bachmann, B. 0., Gunsior, M., Townsend, C. A., and
Solomon, E. I. (2001) Spectroscopic studies of substrate interactions with
Clavaminate synthase 2, a multifunctional or-KG-dependent non-heme iron
enzyme: correlation with mechanisms and reactivities. J. Am. Chem. Soc. 123,

7388-7398.

Ho, R. Y. N., Mehn, M. P., Hegg, E. L., Liu, A., Ryle, M. A., Hausinger, R. P.,
and Que, L., Jr. (2001) Resonance Raman studies of the iron(II)-or-keto acid
chromophore in model and enzyme complexes. J. Am. Chem. Soc. 123,
5022-5029.

(148) Bleijlevens, B., Shivarattan, T., Sedgwick, B., Rigby, S. E., and Matthews, S. J.

(149)

(2007) Replacement of non-heme Fe(II) with Cu(II) in the a-ketoglutarate
dependent DNA repair enzyme AlkB: spectroscopic characterization of the
active site. J. Inorg. Biochem., ASAP.

Proshlyakov, D. A., Henshaw, T. F., Monterosso, G R., Ryle, M. J., and
Hausinger, R. P. (2004) Direct detection of oxygen intermediates in the

55

(150)

(151)

(152)

(153)

(154)

(155)

(156)

(157)

non-heme Fe enzyme taurine/a-ketoglutarate dioxygenase. J. Am. Chem. Soc.
126, 1022-1023.

Hoffart, L. M., Barr, E. W., Guyer, R. B., Bollinger, J. M., Jr., and Krebs, C.
(2006) Direct spectroscopic detection of a C-H-cleaving high-spin Fe(IV)
complex in a prolyl-4-hydroxylase. Proc. Natl. Acad. Sci. USA 103,
14738-14743. ‘

Galonic, D. P., Barr, E. W., Walsh, C. T., Bollinger, J. M., Jr., and Krebs, C.
(2007) Two interconverting Fe(IV) intermediates in aliphatic chlorination by
the halogenase Cth3. Nat Chem Biol 3, 113-6.

Price, J. C., Barr, E. W., Glass, T. E., Krebs, C., and Bollinger, J. M., Jr. (2003)
Evidence for hydrogen abstraction from C1 of taurine by the high-spin Fe(IV)

intermediate detected during oxygen activation by taurineza-ketOglutarate
dioxygenase (TauD). J. Am. Chem. Soc. 125, 13008-13009.

Wolter, T., Meyer-Klaucke, W., Muther, M., Mandon, D., Winkler, H.,
Trautwein, A. X., and Weiss, R. (2000) Generation of oxoiron (IV)
tetramesitylporphyrin pi-cation radical complexes by m—CPBA oxidation of
ferric tetramesitylporphyrin derivatives in butyronitrile at - 78 degrees C.
Evidence for the formation of six-coordinate oxoiron (IV)
tetramesitylporphyrin pi-cation radical complexes FeIV = 0(tmp*)X (X = Cl-,
Br-), by Mossbauer and X-ray absorption spectroscopy. J. Inorg. Biochem. 78,
117-22.

Penner—Hahn, J. E., McMurry, T. 1., Renner, M., Latos-Grazynsky, L., Eble, K.
S., Davis, I. M., Balch, A. L., Groves, J. T., Dawson, J. H., and Hodgson, K. 0.
(1983) X-ray absorption spectroscopic studies of high valent iron porphyrins.
Horseradish peroxidase compounds I and II and synthetic models. J. Biol.
Chem. 258, 12761-4.

Riggs-Gelasco, P. J., Price, J. C., Guyer, R. B., Brehm, J. H., Barr, E. W.,
Bollinger, J. M., Jr., and Krebs, C. (2004) EXAFS spectroscopic evidence for
an Fe=0 unit in the Fe(IV) intermediate observed during oxygen activation by
taurinezalpha-ketoglutarate dioxygenase. J. Am. Chem. Soc. 126, 8108-9.

Borowski, T., de Marothy, S., Broclawik, E., Schofield, C. J., and Siegbahn, P.
E. (2007) Mechanism for cyclization reaction by clavaminic acid synthase.
Insights from modeling studies. Biochemistry 46, 3682-91.

Blasiak, L. C., Vaillancourt, F. H., Walsh, C. T., and Drennan, C. L. (2006)

Crystal structure of the non-haem iron halogenase SyrB2 in syringomycin
biosynthesis. Nature 440, 368-71.

56

(158)

(159)

(160)

(161)

(162)

(163)

(164)

(165)

(166)

(167)

(168)

Karlsson, A., Parales, J. V., Pareles, R. 15., Gibson, D. T., Eklund, H., and
Ramaswamy, S. (2003) Crystal structure of naphthalene dioxygenase: side- -on
binding of droxygen to 1ron Science 299, 1039- 1042.

Kovaleva, E. G, and Lipscomb, J. D. (2007) Crystal structures of Fe2+
dioxygenase superoxo, alkylperoxo, and bound product intermediates. Science
316, 453-7.

Goddard, W. A., 3rd, and Olafson, B. D. (1975) Ozone model for bonding of
an O2 to heme in oxyhemoglobin. Proc Natl Acad Sci U S A 72, 2335-9.

Valko, M., Rhodes, C. J., Moncol, J., Izakovic, M., and Mazur, M. (2006) Free
radicals, metals and antioxidants in oxidative stress-induced cancer. Chem.
Biol. Interact. 160, 1-40.

Weiss, S. J. (1982) Neutrophil-mediated methemoglobin formation in the
erythrocyte. The role of superoxide and hydrogen peroxide. J. Biol. Chem. 25 7,
2947-53.

Simon, R. H., Scoggin, C. H., and Patterson, D. (1981) Hydrogen peroxide
causes the fatal injury to human fibroblasts exposed to oxygen radicals. J. Biol.
Chem. 256, 7181-6.

Saari, R. E., and Hausinger, R. P. (1998) Ascorbic acid-dependent turnover
and reactivation of 2,4-dichlorophenoxyacetic acid/a-ketoglutarate
dioxygenase using thiophenoxyacetic acid. Biochemistry 3 7, 3035-3042.

Welford, R. W. D., Schlemminger, I., McNeill, L. A., Hewitson, K. S., and
Schofield, C. J. (2003) The selectivity and inhibition of AlkB. J. Biol. Chem.
278, 10157-10161.

Ryle, M. J., Liu, A., Muthukumaran, R. B., Ho, R. Y. N., Koehntop, K. D.,
McCracken, J., Que, L., Jr., and Hausinger, R. P. (2003) 02- and
or-ketoglutarate-dependent tyrosyl radical formation in TauD, an Ot-keto
acid-dependent non-heme iron dioxygenase. Biochemistry 42, 1854-1862.

Lindstedt, S., and Rundgren, M. (1982) Blue color, metal content, and

substrate binding in 4-hydroxyphenylpyruvate dioxygenase from
Pseudomonas Sp. strain P. J. 874. J. Biol. Chem. 25 7, 11922-11931.

Liu, A., Ho, R. Y. N., Que, L., Jr., Ryle, M. J., Phinney, B. S., and Hausinger,

R. P. (2001) Alternative reactivity of an a—ketoglutarate-dependent iron(II)
oxygenase: enzyme self-hydroxylation. J. Am. Chem. Soc. 123, 5126-5127.

57

CHAPTER 2

PURIFICATION AND PROPERTIES OF ASPERGILL US N UDULANS

XANTHINE HYDROXYLASE

The work described in this chapter was combined with additional studies and
published: Montero-Moran*, G. M.; Li*, M.; Rendon-Huerita, E.; Jourdan, F.; Lowe, D.
J.; Stumpf‘f-Kane, A. W; Feig, M.; Scazzocchio, C.; and Hausinger, R. P. “Purification
and characterization of the Fe(II)- and oi-Ketoglutarate-Dependent Aspergillus nidulans
Xanthine Hydroxylase from Aspergillus nidulans” Biochemistry, 2007, 46 (18),
5293-5304 (‘ Co-first author). The studies briefly mentioned here regarding purification,
Mr estimation, kinetic characterzaion, glycosylation analysis and phosphorylation testing
of protein from the A. nudulans host were carried out by other authors. One figure
included here (the subunit size comparison from both hosts examined by SDS-PAGE)

was provided by Dr. Montero-Moran.

58

ABSTRACT

Hisfi-tagged xanthine/a-ketoglutarate (OLKG) dioxygenase (XanA) of Aspergillus
nidulans was purified from both the fungal mycelium and recombinant Escherichia coli
cells, and the properties of the two forms of the protein were compared. The kinetic
parameters were similar for XanA from the two sources (ka 30-70 3", Km of OLKG 31-50
“M, Km of xanthine ~45 11M, and pH optima at 7.0-7.4); however, the protein properties
were markedly distinct. Evidence was obtained for both N- and O-linked glycosylation on
the fungus-derived XanA, which aggregated into an apparent dodecamer, while
bacterial-derived XanA was free of glycosylation and behaved as a monomer.
Furthermore, the phosphorylation status differed for the two enzyme forms and the
fungus-derived sample was shown to undergo extensive truncation at its amino terminus.
The sites of posttranslational modification on the two forms of the enzyme are discussed
in terms of a homology model of XanA. These studies represent the first biochemical

characterization of purified xanthine/aKG dioxygenase.

59

INTRODUCTION

Most organisms that metabolize xanthine possess a molybdopterin cofactor
(Moco)-containing enzyme that hydroxylates the substrate to form uric acid while
transferring electrons to NAD (xanthine dehydrogenase) or oxygen (xanthine oxidase) (I).
These enzymes, referred to here as xanthine hydroxylases, are conserved throughout
living organisms, including archaea, bacteria, fungi, plants, and metazoans.

In 2005, a novel mechanism for xanthine metabolism was discovered in certain
fungi (2). This finding arose out of the observation that all mutants of Aspergillus
nidulans defective in xanthine dehydrogenase (i.e., with mutations affecting the structural
gene th, the cnx genes for Moco synthesis, or th for sulfuration of Moco) retained the
ability to grow on xanthine as sole nitrogen source (2, 3). A mutation affecting this
alternative process was identified, and the cognate gene xanA was localized to
chromosome VIII (4). Subsequently, the xanA gene and its homologues from
Schizosaccharomyces pombe and Neurospora crassa were cloned and further
homologues were identified in several other fungi (but not outside the fungal kingdom).
The XanA sequence shows some similarity with the TauD group of Fe(II)- and
a—ketoglutarate (orKG)-dependent dioxygenases (2), including a clear conservation of the
Fe(II)- and aKG-binding sites. This homology suggested that the alternative xanthine
oxidation mechanism present in some fungi might utilize an Fe(II)-dependent
xanthine/OLKG dioxygenase. Such activity, depicted in Figure 2.1, was demonstrated in

both crude and partially purified extracts of fungal mycelia of strains that expressed the

60

FIGURE 2.]: General mechanism of F e(11)/0tKG-dependent xanthine hydroxylases.

 

H
J; 6 5| :8\> Fe(II), XanA ; 1 I N>=o
N 0 H

O N H
H OH
o HO
O
02 + O + 002
H00 "°°

aKG succinate

61

xanA gene (2).

The wide range of Fe(II)/0tKG hydroxylases utilize a diverse array of primary
substrates (reviewed in (5)); however, XanA is the first described enzyme of this group to
hydroxylate a free purine base. In the fungal kingdom, this enzyme coexists with the
classical xanthine hydroxylase; i.e., some fungi possess both xanthine hydroxylase and
xanthine/OLKG dioxygenase, while others possess only one or the other. Notably, yeasts as
evolutionarily distant as S. pombe and Kluveromyces lactis are able to metabolize
xanthine through the activity of a XanA homologue (2). They lack a classical xanthine
dehydrogenase, and they are incapable of synthesizing Moco, which is universally
present in the classical xanthine hydroxylases. The discovery of the novel
Fe(II)/0LKG-dependent XanA enzyme poses both evolutionary and mechanistic problems.
Is the xanthine-binding site of the newly identified enzyme at all similar to that of the
classical xanthine hydroxylases (6, 7) or to the recently described xanthine transporters (8,
9) Is the mechanism of hydroxylation similar to that described for TauD (10, 11) What are
the evolutionary advantages and disadvantages of possessing the Moco-containing and
Fe(II)/0LKG-dependent enzymes.

As a first step towards answering the above questions, I purified the Hisb-tagged
versions of XanA from Escherichia coli and compared it to the corresponding protein
purified from fungal mycelium. I confirm that the enzyme is an Fe(II)/0tKG dioxygenase
and determine the pH dependence and kinetic parameters associated with the Fe(II), ocKG,

and xanthine concentration dependencies. Comparison studies with mycelium-derived

62

enzyme show similar kinetic properties but the proteins differ in quaternary structure and
identity of posttranslational modifications. In addition, fungus-derived protein is
truncated at its amino terminus. Finally, 1 use a homology model of XanA to provide
insights into the sites of posttranslational modification. These studies present the first
detailed biochemical analysis of a purified Fe(II)-dependent xanthine/OLKG dioxygenase.
More detailed kinetic inhibition and homology modeling studies are described in Chapter

3.

63

EXPERIMENTAL PROCEDURES

The plasmid pxanA-Hiso was provided by Gabriela M. Montero-Moran, Institut de
Ge'ne'tique et de Microbiologie, Universite’ Paris-Sud, Batiment 409, UMR 8621 CNRS, 91405
Orsay Cedex, France.

Growth of E. coli Cells Overproducing A. nidulans XanA. Plasmid pxanA-Hi56 was
transformed into XLlBlue E. coli cells (Stratagene) as described by Hanahan (12). A
single colony of E. coli XLlBlue (pxanA-Hisé) was used to inoculate 50 mL of Luria
Base Broth (Difco) containing 100 11g mL'l ampicillin. The cells were grown to
saturation at 37 °C for 16 h and used to inoculate l L of LB media. The culture was
grown at 37 ° C with vigorous shaking until reaching OD600 0.5-0.7, adjusted to contain
0.4 mM isopropyl-B-D-thiogalactopyranoside, transferred to 25 °C, and vigorously
shaken for 16 h. Alternatively, E. coli XLlBlue (pxanA-Hisé) was grown aerobically in
200 mL LB-ampicillin medium at 37 °C to an OD600 of 0.6, induced with 0.75 mM IPTG,
and incubated at 25 °C for 14 h with constant shaking (140 rpm). In either case, cultures
were harvested by centrifugation at 8,000-9,000 rpm for 10 min at 4 ° C.

Purification of His6-tagged XanA from E. coli. Approximately 5 g of E. coli XLBlue
(pxanA-Hisé) cell paste was suspended in 10-15 mL of lysis buffer containing 100 mM
Tris, pH 8.0, 300 mM NaCl, 25 mM imidazole, and trace amounts of lysozyme, leupeptin,
DNase I and RNase A. This suspension was incubated at room temperature for 30 min,
transferred to an ice bath for 30 min, disrupted by using a French pressure cell at ~500 psi

at 4 ° C, and spun for 45 min at 100,000 g. The soluble cell extracts (30 mL) were loaded

64

onto a 10 mL Ni-nitrilotriacetic acid (NTA) column using 100 mM Tris buffer, pH 8.0,
containing 300 mM NaCl and 25 mM imidazole, followed by elution with 100 mM Tris
buffer, pH 8.0, with 300 mM NaCl, 250 mM imidazole and 15 % glycerol. The fractions
containing XanA were collected and incubated with 1 mM EDTA at 4 °C for 5 h, then
concentrated to 5-10 mg mL'I by using a Centriprep (Amicon Corp.) with a YM-lO
membrane.

Enzyme Assays. Xanthine/aKG dioxygenase activity was measured at 25 °C by
using the following typical assay conditions (total volume of 1 mL): 1 mM orKG, 40 11M
Fe(NH4)2(SO4)2, and 200 uM xanthine in 50 mM MOPS buffer, pH 7.4. Variations of
these conditions included use of alternate buffers, different pH values, and varied
concentrations of substrates. The absorbance at 294 nm was monitored to determine the
uric acid production (8294 12,200 M’I cm") with a correction for loss of the xanthine
absorbance at this wavelength (measured 8294 2,000 M’l cm") for an overall change in
3294 of 10,200 M" cm". Units of activity (U) were defined as umol min’1 of uric acid
produced and the specific activity (U mg") was measured as umol min‘l (mg of purified
XanA)".

In addition to the above spectroscopic assay, xanthine/aKG dioxygenase activity was
measured by two alternative methods. Oxygen consumption measurements were carried
out in air-saturated MOPS medium (pH 7) at 25 °C by using a Clark-type oxygen electrode.
Quantification of aKG consumed during the reaction was assessed by HPLC. Aliquots

(300 uL) of the reaction mixtures were quenched by addition of 5 uL 6 M H2SO4, the

65

samples were centrifuged for 5 min at 20,000 g, and the supernatant was
chromatographed on an Aminex HPX-87H column (Bio-Rad Laboratories) in 0.013 M
H2804 with detection by using a differential refractometer (Waters, Model R401).

Metal Analyses. The iron concentration was measured by utilizing the KMnO4
oxidation, ascorbate reduction, and ferrozine chelation protocol of Bienert (13).

Protein Analytical Methods. Routine determinations of protein concentration were
carried out by the method of Bradford (14) with bovine serum albumin used as the
standard. Qualitative measurement of protein overexpression and assessment of protein
purity made use of sodium dodecyl sulfate polyacrylamide gel electrophoresis
(SDS-PAGE) (15), with stacking and running gels containing 5 % and 12 % acrylamide.
Standard proteins used for comparison included phosphorylase b, M, 97,400; bovine
serum albumin, M, 66,200; ovalbumin, M, 45,000; carbonic anhydrase, M, 31,000;
trypsin inhibitor, M, 21,500; and lysozyme, M, 14,400 (Bio-Rad Laboratories). The native
size of XanA isolated from E. coli was estimated by gel filtration chromatography using a
Protein-pak Diol(OH) 10 um column (Waters, 0.5 or 1 min mL", room temperature in
100 mM Tris buffer, pH 7.5, containing 300 mM NaCl), a SuperoseTM 6 HR 10/30 GL
column (Pharrnacia, 1 mL min'1 in 50 mM MOPS, pH 6.8, containing 0.15 M NaCl). The
calibration proteins were thyroglobulin, M, 670,000; y-globulin, M, 158,000; ovalbumin,
M, 44,000; myoglobin, M, 17,000; and vitamin B12, M, 1,350 (Bio-Rad).

Mass Spectrometry. (Assisted by Dan Jones, Department of Biochemistry &

Chemistry, MSU). Mass spectrometry analyses were performed by using a Waters

66

(Milford, MA) LCT Premier mass spectrometer coupled to a Shimadzu (Columbia, MD)
LC-20AD HPLC and SIL-5000 autosampler. Samples were analyzed using electrospray
ionization in positive ion mode. On-line desalting and separation from detergents was
performed using a Thermo Hypersil-Keystone BetaBasic cyano column (1.0 x 10 mm)
coupled to the electrospray ionization probe.Aliquots were injected onto the column
using a flow rate of 0.1 mL/min of 95% solvent A (0.15% aqueous formic acid)/5%
solvent B (acetonitrile). Gradient elution was performed by using the following
parameters: (0-1 min: 95%A/5%B; linear gradient to 30%A/70%B at 6 min; hold at
30%A/70%B until 9 min). Electrospray spectra were processed using MassLynx software
(Waters, Milford, MA), and zero-charge state mass spectra were obtained by
deconvolution using the MaxEntl algorithm.

MALDI mass spectra were generated on a Voyager-DE STR mass spectrometer
(Applied Biosystems, Foster City, CA) in positive ion linear mode using sinapinic acid as
matrix. Samples were processed using strong cation exchange ZipTips (ZipTipSCX,
Millipore, Billerica, MA) to remove detergent and reversed phase C18 ZipTips for
desalting, following the manufacturer's recommended protocols, before spotting the
MALDI target.

Structural Homology Modeling. (Assisted by Michael Feig and Andrew W.
Stumpff-Kane, Department of Biochemistry and Molecular Biology, MSU.) Details
related to the creation of the homology model using TauD (PDB code IOS7, chain A) as

a template structure (16, 1 7) are described in Chapter 3 (18).

67

RESULTS

Properties of Recombinant XanA in E. coli Cell Extracts. When assayed by using
standard conditions (1 mM aKG, 40 11M Fe(NH4)2(SO4)2, and 200 uM xanthine in 50
mM MOPS buffer, pH 7.4, 25‘ °C), fresh extracts of E. coli cells overproducing
Hisé-tagged XanA exhibited xanthine/orKG dioxygenase specific activity of
approximately 19 U (mg protein)’1 (Figure 2.2A). As expected, the activity was strictly
dependent on the presence of aKG in the assay as previously described for the crude
enzyme extracted from A. nidulans (Figure 2.28) (2). Unlike the earlier findings,
however, significant activity was detected in the absence of added Fe(II) (16.9 U mg")
(Figure 2.2C) and some activity was retained when 5 mM EDTA was included in the
assay buffer (2.01 U mg"), Figure 2.2D. These data indicate that endogenous Fe(II) is
tightly bound to the bacterial-produced enzyme sample. The activity of these extracts was
irreversibly lost over 3 weeks when stored at 4 °C using a protein concentration of 15-17
mg mL", or after 1 week in the added presence of 1 mM EDTA. While long-term
incubation of cell extracts with EDTA was undesirable, studies with enriched enzyme
samples showed that EDTA treatment provided stable apoprotein (when maintained at
high protein concentrations) that could be activated by addition of ferrous ions.

Purification of XanA from the E. coli. XanA produced in E. coli XLlBlue (pxan-His6)
was purified to homogeneity from cell extracts (Figure 2.3A) by Ni-NTA chromatography.
Inclusion of 15 % glycerol in the chromatography buffers, which were maintained on ice,

helped to minimize protein precipitation during purification. About 5 % of the soluble

68

FIGURE 2.2: Activity assay of XanA cell extracts. (A) Complete assay: 15 pg XanA
crude extract, 40 uM F e(ll), 1 mM aKG, 100 uM xanthine in 50 mM MOPS, pH=7.4, 25
°C. (B) without aKG. (C) without Fe(II). (D) with 5 mM EDTA. UV-Vis spectra were

recorded from 240 to 320 nm every 15 seconds after initiating the enzymatic reaction by

adding primary substrate.

 

 

 

 

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69

activity was located in the flow-though fractions, perhaps indicating that some XanA
interacts tightly with other proteins that fail to bind the resin. As measured by the
standard assay protocol, the Ni-NTA column fractions containing purified XanA
accounted for 33 % of the activity that had been observed in cell extracts. When this pool
was treated with 1 mM EDTA at 4 ° C for 5 h and then concentrated to 5-12 mg mL", the
activity increased such that 60 % of the activity of cell extracts was recovered and
yielded a final specific activity for the purified enzyme of 70-80 U (mg protein)" at 25 °C
which corresponds to a km, of 49-56 s'l (assuming M, = 42 kDa per subunit). The enzyme
recovered from the Ni-NTA column contained 0.26 to 0.5 moles of Fe per mole of
subunit according to the colorimetric assay, while that incubated with EDTA lacked
detectable Fe, and a sample incubated with exogenous Fe(II) and then chromatographed
on a Sephadex G-25 gel filtration column contained 1.3 moles of Fe per mole of subunit.
Concentrated XanA derived from E. coli was stable for at least one month at 4 °C when
stored in 100 mM Tris buffer, pH 8.0, containing 300 mM NaCl, 250 mM imidazole, 1
mM EDTA and 15 % glycerol, or at least two months if frozen at -80 ° C.

For comparison, the specific activity of the isolated protein from the firngal host was
measured as 22-40 U (mg protein)" at 30 °C equivalent to a km, of 15.4-30 5" (again
assuming M, = 42 kDa per subunit), comparable with those from bacterial host. (The
activity of XanA isolated from fungal host was measured by Gabriela M. Montero-Moran,

Institut de Génétique et de Microbiologie, Université Paris-Sud, Batiment.)

70

FIGURE 2.3: SDS-PAGE analysis of the purified XanA from E. coli and A. nidulans. (A)
Analysis of fractions during purification of XanA from E. coli. Lane 1, molecular weight
standards; lanes 2 and 3, suspension of cultures before and after induction by IPTG; lane
4, cell extracts afier lysis; lane 5, cell pellet after lysis; lane 6, flow-through fraction from
Ni-NTA chromatography; lane 7-13, fractions of purified XanA obtained after Ni-NTA
chromatography. (B) Comparison of the purified XanA protein derived from A. nidulans
and E. coli. Lane M, markers; lane 1, sample purified from the fungus; lane 2, protein
isolated from the bacterium (7 pg each). Stacking and running gels contain 5 % and 12 %

acrylamide.

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Effects of pH on Stability and Activity of XanA from E. coli. The effect of pH on the
stability of XanA was examined. After incubating the XanA samples in various pH
buffers at 4 °C for 3 h, the activity remaining was examined by using the standard assay
procedure. The results (data not shown) indicate that each purified enzyme is stable over
a wide pH range (7.0-11.0).

The effect of pH on the activity of the enzyme was also examined. XanA isolated
from E. coli was assayed by using a series of different buffers as depicted in Figure 2.4.
The enzyme activity exhibited a narrow pH optimum of 7.0-8.0 with pH 7.4 yielding
optimal activity for most buffers (Tris, MOPS, MES, imidazole, and HEPES).

Kinetic Analyses of XanA from E. coli. For the E. coli-derived enzyme, the results of
studies using varied orKG concentrations provided a Km of 3 1.1 i 1.6 uM and km of 66.5
s" at 25 °C (Figure 2.5A), while those for varied xanthine concentrations provided a Km
of 45.2 :t 3.6 uM and kca, of 71.4 s" (Figure 2.5B). Fe(lI) is required for xanthine/aKG
dioxygenase activity, with half-maximal activity at ~7 11M when using the apoprotein
isolated from the bacterium (Figure 2.6).

The kinetic parameters were very similar for XanA isolated from the two sources.
When the protein isolated from A. nidulans enzyme was assayed at 30 °C with varied
aKG or xanthine concentrations the measured Km values were 50 11M 2t 6 and 46 d: 4 pM,
respectively, but with a smaller Item that ranged from 15 s" to 30 5" depending on the

preparation provided by Gabriela M. Montero-Moran.

72

FIGURE 2.4: pH dependence of XanA activity. The activity of XanA (0.51 pg ml")
derived from E. coli was assayed in the following buffers (50 mM) and pH values at 25
°C: imidazole, pH 6.0-8.2 (O);MES, pH 666.8 (+); MOPS, pH 6.6-7.8 (X); HEPES, pH
7.0-8.2 (V); Tris, pH 7.4-9.0 (o);CHES, pH 9.4-9.8 (A); CAPS, pH 9.6-10.0 (0). Assay

solutions also contained 1 mM aKG, 40 uM Fe(NH4)2(SO4)2, and 200 uM xanthine.

 

 

73

FIGURE 2.5: Substrate and co-substrate concentration dependencies of XanA. The
effects of varying the concentrations of (A) aKG and (B) xanthine on xanthine/aKG
dioxygenase activity were examined for the E. coli-derived protein at 25 °C. Except for
the compound being varied, the assay solutions contained 40 pM Fe(II), 1 mM aKG, and
200 uM xanthine in 50 mM MOPS buffer, pH 7.4. The data were fit to the

Michaelis-Menten equation.

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The stoichiometry of the enzymatic reaction was examined for XanA by UV-Vis
spectroscopy and 02 electrodes. The degradation of 100 pM xanthine was accompanied
by the production of 92 uM uric acid as measured by UV-Vis spectrosocopy, and this
coincided with the consumption Of 110 uM oxygen determined by O; electorde (data not
shown).

Differential Protein Properties of XanA Purified from the Two Host Cells. Gel
filtration chromatography provided an estimated M, of 39-42 kDa for the native enzyme
isolated from the bacterial host (consistent with a monomeric structure) (Figure 2.7);
however, similar analysis of the protein from the fungal host showed that it was
oligomeric, with an approximate M, of 500 kDa that was consistent with about twelve
subunits per native molecule (data from Gabriela M. Montero-Moran.) (18).The SDS-PAGE
results highlight a key difference between the XanA proteins isolated from the two
sources; i.e., the apparent M, of the E. coli-derived protein is larger than that of the
protein derived from the fungus (Figure 2.3B) (Provided by Montero-Morén). This
finding led us to investigate the possibility of unique posttranslational modifications in
the proteins produced in the bacterial and eukaryotic hosts. G M. Montero-Moran and
co-workers have shown the presence of N-glycosylation, O-glycosylation and Thr
phosphorylation in the protein purified from A. nidulans, but both Thr and Ser
phosphorylation in the one isolated from bacterial host (18).

Mass spectrometric methods were used to further characterize the two enzyme forms.

Electrospray ionization mass spectrometry of bacteria-derived XanA indicated a single

75

FIGURE 2.6: Fe(ll) concentration dependence of XanA. The effects of varying the
concentration of Fe(II) on xanthine/aKG dioxygenase activity were examined by using

the E. coli-derived protein at 25 °C in solutions containing 1 mM aKG and 200 1.1M

xanthine in 50 mM MOPS buffer, pH 7.4.

 

 

 

76

FIGURE 2.7: Measurement of native size of XanA derived from E. coli. (I) represents

the standards and (1:1) represents the XanA isolated form E. coli.

 

 

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15 '
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3 I I l .

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elution volume (mL)

77

species with a molecular mass of 41,992 Da (data not shown), which matches very well
to the theoretical mass (41,996.50 Da; using the ExPASY Compute pI/Mw tool at
ca.expasy.org) for the His6-tagged protein missing its amino-terminal Met residue. This
sample provided no evidence Of phosphorylation (differing from the immunological
detection of phosphoserine and phosphothreonine by our collaborators) or glycosylation.
In contrast to this single species, the fungus-derived protein sample exhibited a complex
electrospray ionization mass spectrum centered near 36,000 Da (Figure 2.8). The
spectrum of Figure 2.8 includes features separated by 162 mass units, consistent with
glycosylation involving hexose sugars, as well as features separated by 80 mass units,
indicating phosphorylation. The smallest component of the spectrum exhibits a mass of
35,171 Da, indicating that the non-glycosylated and non-phosphorylated fungal protein is
severely truncated compared to the theoretical mass of full-length protein of 42,127.69.
This truncation must occur at the N-terminus, since the C-terrninal Hisé-tag was used for

enzyme purification, and consists of approximately 60 residues.

78

FIGURE 2.8: Mass spectrometric analysis of fungus-derived XanA. The XanA protein

isolated from A. nidulans was analyzed by electrospray ionization mass spectrometry.

The figure depicts a series of peaks separated by 162 mass units and 80 mass units,

consistent with glycosylation and phosphorylation, respectively.

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79

DISCUSSION

In this study I describe the isolation and general properties of xanthine/aKG
dioxygenase, a novel enzyme found exclusively in the fungal kingdom (2).

Purification. Immobilized metal ion chromatography is very effective for purifying
recombinant XanA from the bacterial host cells. His-tagged versions of several other
representative Fe(ll)/0LKG dioxygenases have been studied, including AlkB, deA, SdpA,
phytanoyl-CoA hydroxylase, and the oxygen-sensing prolyl 4- and asparaginyl
hydroxylases (19-23). The His-rich sequences generally have modest if any effect on
activity.

XanA Enzyme Stability. Purified XanA is unstable at room temperature (denaturing
within an hour even in buffers containing 15 % glycerol), when agitated (e.g., during
stirring in an Amicon concentrator), or when incubated at pH values below 6.5 (the pI
estimated for XanA is 5.82 according to the ExPASy ProtParam tool at ca.expasy.org).
EDTA treatment enhances the activity of the highly concentrated bacterial-derived
enzyme and this compound is maintained in the storage buffer to ensure maximal lifetime
of the activity. I attribute the enhancement effect of EDTA to its ability to remove
inhibitory Ni(II) (co-eluted with the enzyme from the NTA resin) and Fe(III) (the
oxidized, inactive state of the metal) from the enzyme so that the apoprotein can bind
Fe(lI) in the assay buffer. Several other Fe(II)/0LKG dioxygenases are purified as their
apoprotein forms by inclusion of chelators (24, 25) to prevent Fe(II) oxidation and to

eliminate Ni(II), if purified using an NTA column. In contrast, some representatives of

80

these enzymes have been purified anaerobically to assure that the metal remains in its
reduced form (26, 27).

XanA Enzyme Activity. The kinetic properties of XanA as purified from E. coli (~70
U mg ", Km of 31 pM for orKG, and Km of 45 uM for xanthine at pH 7.4) compare well
with those of XanA isolated from A. nidulans (30 U mg", and Km values of 50 uM and 46
uM at pH 7.0), and these results are compatible with those reported earlier for enriched
sample from the fungus (40 U mg", 50 uM, and 23 uM) (2). The reaction requires Fe(II)
(half-maximal activity at 7 uM for standard conditions), consistent with the results of .
related family members.

Posttranslational Modifications. My comparison of recombinant XanA purified from
E. coli with that of A. nidulans provided by my collaborators reveals very different
quaternary structures and posttranslational modifications. Whereas the protein derived
from the bacterial source chromatographs on a size exclusion column as a monomer, that
isolated from the fungus is much larger, with apparent M, of 500 kDa. The sample from
the fungus was also glycosylated and phosphorylated while the one from bacterial was
suggested to contain both phosphoserine and phosphothreonine by our collaborators.
Treatment of the fungal protein with PNGase F was found to result in a dramatic shift in
electrophoretic mobility, but not all glycosylation was removed by this process indicating
the presence of both N— and O-glycoconjugates. Comparing the size of the subunit from
both sources along with the mass spectrometry results suggested an extensive truncation

takes place at the N-terminus of the fungal enzyme. Despite the extensive differences in

81

post-translational modifications between the two forms of the enzyme the kinetic
parameters are nearly identical, consistent with the modifications not affecting catalytic
activity. Analogous biochemical comparisons between the enzyme form isolated from its
native eukaryotic host and the form isolated from E. coli have not been reported for other
F e(II)/aKG dioxygenase family members.

An extant question is the role of the extensive N-terminal processing of enzyme
isolated from its natural fungal host. I speculate that this feature may relate to cellular
targeting of the XanA enzyme. It is not unreasonable to propose that an oxidoreductase
such as XanA could be peroxisomal. In fact, the immediate downstream enzyme, urate
oxidase, has been shown to be peroxisomal in every organism where its localization has
been investigated, including amoebas, mammals and plants (28-31). A urate oxidase-green
fluorescent protein fusion shows a particulate intracellular distribution in Aspergillus
nidulans, fully consistent with a peroxisomal localization (G. Langousis and G. Diallinas,
unpublished data). XanA does not possess a C-terminal peroxisomal targeting signal
(variations of an SKL tripeptide, denoted PTSI); however, it contains a RSALYTHL
sequence (residues 40-47) that resembles the PTSZ N-terminal import sequence (variations
of RLXsHL) found in a minority of peroxisomal proteins throughout the eukaryotes
(32-34). In some instances, including mammalian (35), yeast (36), and plant (37)
peroxisomal proteins, it has been shown that the PST2 is contained in a pre-sequence that is
cleaved upon peroxisomal entry. The function of XanA glycosylation could be understood

in this context as a mechanism to protect the protein from further proteolysis.

82

FIGURE 2.9: Location of potential sites of glycosylation and phosphorylation in XanA.
The positions of selected residues are depicted in a homology model, described in the
chapter 3, with the potential N-glycosylation site (Asn118) in blue, Thr and Ser residues
that could be either glycosylated or phosphorylated (T hr5 and Ser37) in orange and
magenta, potential 0-glycosylation sites (Thr10, Thr195, and Ser316) in red, and other
possible phosphothreonines (Thr38, Thr58, Thr66, Thr120, and Thr274) or
phosphoserines (Ser15, Serl42, Serl99, Ser208, Ser217, Ser2l8, and Ser34l) illustrated
in yellow and cyan. The amino terminus of the fungus-derived protein is truncated
through the first approximately 60 residues. The brown sphere shows the postulated
position of Fe(II). Additional glycosylation (Ser231) and phosphorylation (Thrl77 and
Ser184) are not shown because they are located on non-modeled protein loops that

represent insertions compared to the template structure (TauD, PDB code 1087, chain A).

stems
Ser142 56'2"

  

83

Insights from a Homology Model. A XanA homology model (Figure 2.9), whose
generation is described in detail in Chapter 3 (18), provides critical insights into the likely
sites of posttranslational modification in the protein. This model is based on the
full-length sequence of the XanA protein, but we assume that the overall fold is
maintained even in the truncated version derived from the fungal host because this region
of the protein is external to the DSBH core of the protein. Consistent with the PNGase F
results that demonstrate the presence of N-glycosylation sites in the fungus-derived
protein, the NetNGlyc 1.0 server (R. Gupta, E. Jung, and S. Brunak, unpublished)
predicts glycosylation of Asnl.18, and the homology model places this residue on the
protein surface. The PNGase F-treated fungal-derived protein still stains as a glycoprotein,
consistent with O-Iinked glycoconjugates in the protein. Thr5 and ThrIO are predicted to
be glycosylated by the NetOGlyc 3.1 server (38) while the YingOYang 1.2 server (R.
Gupta, J. Hansen, and S. Brunak, unpublished) predicts Thr5, Ser37, Thr195, Ser23 1 , and
Ser3l6 as possible sites of glycosylation. The XanA homology model reveals that all of
these positions except for Thr195 are surface exposed or, for Ser23], on surface loops
that were not modeled. On the basis of my mass spectrometry results showing truncation
of approximately 60 residues from the N-terminus, I suggest Ser23] or Ser3l6 as the
most likely sites of O-glycosylation.

A wide range of potential Ser, Thr, or Tyr phosphorylation sites are identified in
XanA by the NetPhos 2.0 server (39) and by Prosite (www.cxpasvorg). Phosphoproteins

are common in eukaryotes and are well known in E. coli (40). Given the immunological

84

evidence for phosphothreonine in XanA from both sources, the Thr phosphorylation sites
are of special interest and are predicted by at least one program to include Thr5, Thr38,
Thr58, Thr66, Thr120, Thrl77, and Thr274. The first three residues are likely to be
missing in the fungus-derived protein based on the mass spectrometry results. The
homology model predicts that Thr66 and Thr120 are buried (although their disposition is
less clear for the truncated protein), whereas the other residues are exposed to the surface
or, for Thrl77, on a loop that was not modeled. The bacterial-derived protein also reacts
with antibodies directed against phosphoserine, whereas XanA derived, from the fungus
reacted only weakly with these antibodies. We attribute this difference in reactivity
between the two proteins to the phosphorylation site being inaccessible in the
fungal-derived sample, either due to nearby glycosylation or due to being located at a
subunit interface in the multimeric protein. Sites of Ser phosphorylation predicted by at
least one program include Ser15, Ser37, Serl42, Ser184, Serl99, Ser208, Ser217, Ser218,
and Ser34l. The first two residues are likely to be absent in the fungus-derived protein.
Residues Ser199 and Ser341 are predicted to be somewhat buried in the model, Ser184
occurs on a loop that could not be modeled, and the other residues are predicted to be
surface exposed. Of note, Ser341 would be made more inaccessible by glycosylation of
Thr195; thus, potentially explaining the source specificity behavior for Ser

phosphorylation.

85

REFERENCES

(1)

(2)

(3)

(4)

(5)

(6)

(7)

(8)

(9)

Hille, R. (2005) Molybdenum-containing hydroxylases. Arch. Biochem. Biophys.
433, 107-116.

Cultrone, A., Scazzocchio, C., Rochet, M., Montero-Moran, G, Drevet, C., and
F emandez-Martin, R. (2005) Convergent evolution of hydroxylation mechanisms
in the fungal kingdoms: molybdenum cofactor-independent hydroxylation of

xanthine via or-ketoglutarate dependent dioxygenase. Molec. Microbiol. 5 7,
276-290.

Darlington, A. J ., and Scazzocchio, C. (1968) Evidence for an alternative pathway
of xanthine oxidation in Aspergillus nidulans. Biochim. Biophys. Acta 166,
557-568.

SeaIy-Lewis, H. M., Scazzocchio, C., and Lee, S. (1978) A mutation defective in
xanthine alternative pathway of Aspergillus nidulans: its use to investigate the
specificity of uaY mediated induction. Molec. Gen. Genet. 164, 303-308.

Hausinger, R. P. (2004) Fe(lI)/or-ketoglutarate-dependent hydroxylases and
related enzymes. Crit. Rev. Biochem. Mol. Biol. 39, 21-68.

Glatigny, A., Hof, P., Romao, M. J., Huber, R., and Scazzocchio, C. (1998)
Altered specificity mutations define residues essential for substrate positioning in
xanthine dehydrogenase. J. Mol. Biol. 278, 431-438.

Truglio, J. J ., Theis, K., Leimkt'rhler, S., Rappa, R., Rajagopalan, K. V., and Kisker,
C. (2002) Crystal structure of the active and alloxanthine-inhibited forms of
xanthine dehydrogenase from Rhodobacter capsulatus. Structure 10, 115-125.

Goudela, S., Karatza, P., Koukaki, M., Frilingos, S., and Diallinas, G (2005)
Comparative substrate recognition by bacterial and fungal purine transporters of
the NAT/NCS2 family. Molec. Membrane Biol. 22, 263-275.

Koukaki, M., Vlanti, A., Goudela, S., Pantazopoulou, A., Gioule, H., Toumaviti,
S., and Diallinas, G (2005) The nucleobase-ascorbate transporter (NAT) signature
motif in UapA defines the function of the purine translocation pathway. J. Mol.
Biol. 350, 499-513.

86

(10)

(11)

(12)

(13)

(14)

(15)

(16)

(17)

(18)

(19)

Eichhom, E., van der Ploeg, J. R., Kertesz, M. A., and Leisinger, T. (1997)
Characterization of or-ketoglutarate-dependent taurine dioxygenase from
Escherichia coli. J. Biol. Chem. 272, 23031-23036.

Grzyska, P. K., Ryle, M. J., Monterosso, G R., Liu, J., Ballou, D. P., and
Hausinger, R. P. (2005) Steady-state and transient kinetic analyses of
taurine/a-ketoglutarate dioxygenase: effects of oxygen concentration, alternative
sulfonates, and active site variants on the Fe(IV) intermediate. Biochemistry 44,

3845-3855.

Hanahan, D. (1983) Studies on transformation of Escherichia coli with plasmids.
J. Mol. Biol. 166, 557-580.

Beinert, H. (1978) Micro methods for the quantitative determination of iron and
copper in biological material. Meth. Enzymol. 54, 435-445.

Bradford, M. M. (1976) A rapid and sensitive method for the quantitation of
microgram quantities of protein utilizing the principle of protein-dye binding.
Anal. Biochem. 72, 248-254.

Laemmli, U. K. (1970) Cleavage of structural proteins during the assembly of the
head of bacteriophage T4. Nature (London) 22 7, 680-685.

Elkins, J. M., Ryle, M. J., Clifton, I. J., Dunning Hotopp, J. C., Lloyd, J. S.,
Burzlaff, N. 1., Baldwin, J. E., Hausinger, R. P., and Roach, P. L. (2002) X-ray
crystal structure of Escherichia coli taurine/a-ketoglutarate dioxygenase
complexed to ferrous iron and substrates. Biochemistry 41, 5185-5192.

O'Brien, J. R., Schuller, D. J., Yang, V. S., Dillard, B. D., and Lanzilotta, W. N.
(2003) Substrate-induced conformational changes in Escherichia coli

taurine/a-ketoglutarate dioxygenase and insight into the oligomeric structure.
Biochemistry 42, 5547-5554.

Montero-Moran, G M., Li, M., Rendon-Huerta, E., Jourdan, F., Lowe, D. J.,
Stumpff-Kane, A. W., Feig, M., Scazzocchio, C., and Hausinger, R. P. (2007)
Purification and characterization of the Fe(II)- and or-ketoglutarate-dependent
xanthine hydroxylase from Aspergillus nidulans. Biochemistry 46, 5293-5304.

Trewick, S. C., Henshaw, T. F., Hausinger, R. P., Lindahl, T., and Sedgwick, B.
(2002) Oxidative demethylation by Escherichia coli AlkB directly reverts DNA
base damage. Nature 419, 174-178.

87

(20)

(21)

(22)

(23)

(24)

(25)

(26)

(27)

(28)

(29)

Muller, T. A., Zavodszky, M. I., F eig, M., Kuhn, L. A., and Hausinger, R. P. (2006)
Structural basis for the enantiospecificities of R- and
S-phenoxypropionate/or-ketoglutarate dioxygenases. Prot. Science 15, 1356-1368.

Searls, T., Butler, D., Chien, W., Mukherji, M., Lloyd, M. D., and Schofield, C. J.
(2005) Studies on the specificity of pro- and mature-forms of phytanoyl-CoA
2-hydroxylase and mutation of the iron binding ligands. J. Lipid Res. 46,
1660-1670.

McDonough, M. A., Li, V., Flashman, E., Chowdhury, R., Mohr, C., Liénard, B.
M. R., Zondlo, J., Oldham, N. J., Clifton, I. J., Lewis, J., McNeil], L. A., Kurzeja,
R. J. M., Hewitson, K. S., Yang, E., Jordan, S., Syed, R. S., and Schofield, C. J.
(2006) Cellular oxygen sensing: crystal structure of hypoxia-inducible factor
prolyl hydroxylase (PHD2). Proc. Natl. Acad. Sci. USA 103, 9814-9819.

Dann, C. E., III, Bruick, R. K., and Deisenhofer, J. (2002) Structure of a
factor-inhibiting hypoxia-inducible factor 1: an essential asparaginyl hydroxylase
involved in the hypoxic response pathway. Proc. Natl. Acad. Sci. USA 99,
15351-15356.

Ryle, M. J., Padmakumar, R., and Hausinger, R. P. (1999) Stopped-flow kinetic-
analysis of Escherichia coli taurine/or-ketoglutarate dioxygenase: interactions
with or-ketoglutarate, taurine, and oxygen. Biochemistry 38, 15278-15286.

Fukumori, F., and Hausinger, R. P. (1993) Purification and characterization of
2,4-dichlorophenoxyacetate/a-ketoglutarate dioxygenase. J. Biol. Chem. 268,
24311-24317.

Mishina, Y., Chen, L. X., and He, C. (2004) Preparation and characterization of
the native iron(III)-containing DNA repair AlkB protein directly from Escherichia
coli. J. Am. Chem. Soc. 126, 16930-16936.

Vaillancourt, F. H., Yin, J., and Walsh, C. T. (2005) SyrBZ in syringomycin E
biosynthesis is a nonheme Fell a-ketoglutarate- and Oz-dependent halogenase.

Proc. Natl. Acad. Sci. USA 102, 10111-10116.

Muller, M., and Moller, K. M. (1969) Urate oxidase and its association with
peroxisomes in Acanthamoeba sp. Eur: J. Biochem. 9, 424-430.

Leighton, F., Poole, B., Lazarow, P. B., and de Duve, C. (1969) The synthesis and

88

(30)

(31)

(32)

(33)

(34)

(35)

(36)

(37)

(38)

(39)

turnover of rat liver peroxisomes. I. Fractionation of peroxisome proteins. J. Cell.
Biochem. 41, 521-535.

Nguyen, T., Zelechowska, M., Foster, V., Bergmann, H., and Verma, D. P. S.
(1985) Primary structure of the soybean nodulin-35 gene encoding uricase II
localized in the peroxisomes of uninfected cells of nodules. Proc. Natl. Acad. Sci.
USA 82, 5040-5044.

Breidenbach, R. W., Kahn, A., and Beevers, H. (1968) Characterization of
glyoxysomes from castor bean endosperm. Plant Physiol. 43, 705-713.

de Hoop, M. J., and Ab, G (1992) Import of proteins into peroxisomes and other
microbodies. Biochem. J. 286, 657-669.

Petriv, O. 1., Tang, L., Titorenko, V. 1., and Rachubinski, R. A. (2004) A new
definition for the consensus sequence of the peroxisome targeting signal type 2. J.
Mol. Biol. 341, 119-134.

Reumann, S. (2004) Specification of the peroxisome targeting signals type 1 and
type 2 of plant peroxisomes by bioinformatics analyses. Plant Physiol. 135,
783-800.

Swinkels, B. W., Gould, S. J., Bodnar, A. G, Rachubinski, R. A., and
Subramaniam, S. (1991) A novel, cleavable peroxisomal targeting signal at the
amino-terminus of rat 3-ketoacyl-C0A thiolase. EMBO J. 10, 3255-3262.

Lee, J. G, Lee, Y. J., Lee, C. H., and Maeng, P. J. (2006) Mutational and
functional analysis of the cryptic N-terminal targeting signal for both
mitochondria and peroxisomes in yeast peroxisomal citrate synthase. Biochem

(Tokyo) 140, 121-133.

Kato, A., Hayashi, M., Kondo, M., and Nishimura, M. (2000) Transport of
peroxisomal proteins synthesized as large precursors in plants. Cell Biochem.
Biophys. 32, 269-275.

Julenius, K., Molgaard, A., Gupta, R., and Brunak, S. (2005) Prediction,
conservation analysis and structural characterization of mammalian mucin-type
O-glycosylation sites. Glycobiology 15, 153-164.

Blom, N., Gammeltoft, S., and Brunak, S. (1999) Sequence- and structure-based

prediction of eukaryotic protein phosphorylation sites. J. Mol. Biol. 295,

89

1351-1362.

(40) Cortay, J.-C., Rieul, C., Duclos, B., and Cozzone, A. J. (1986) Characterization of
the phosphoproteins of Escherichia coli cells by electrophoretic analysis. Eur. J.

Biochem. 159, 227-237.

90

CHAPTER 3

KINETIC CHARACTERIZATION, ISOTOPE EFFECTS, AND EFFECTS OF
ALTERNATE METAL IONS, SALTS, AND SUBSTRATE ANALOGUES ON

ASPERGILL US NIDULANS XANTHINE HYDROXYLASE

The studies described in this chapter were combined with additional investigations
and published: Montero-Moran*, G. M.; Li*, M.; Rendon-Huerita, E.; Jourdan, F.; Lowe,
D. J .; Stumpff-Kane, A. W; Feig, M.; Scazzocchio, C.; and Hausinger, R. P. “Purification
and characterization of the Fe(II)- and a-Ketoglutarate-Dependent Aspergillus nidulans
Xanthine Hydroxylase from Aspergillus nidulans” Biochemistry, 2007, 46 (18),
5293-5304 (. Co-first author). Michael Feig and Andrew W. Stumpff-Kane constructed
the homology model, David J. Lowe and Fabrice Jourdan synthesized the
8-hydroxypurine, 2,8-dihydroxypurine and 6,8-dihydroxypurine, and Christine Drevet
provided the alignment figure of XanA with putative orthologous sequences from several

fungi.

91

ABSTRACT

Xanthine/or-ketoglutarate (OtKG) dioxygenase (XanA) of Aspergillus nidulans was
previously purified from fungal mycelium and recombinant Escherichia coli cells, and
the two forms of the enzyme were shown to exhibit distinct posttranslational
modifications while retaining similar kinetic parameters as described in Chpater 2. Here,
I characterize several additional kinetic properties of the XanA isolated form E. coli. The
enzyme exhibits no significant isotope effect when using 8-2H-xanthine; however, it
demonstrates a two-fold solvent deuterium isotope effect. Cu(II) and Zn(II) potently
inhibit the Fe(II)-specific enzyme, presumably by binding to the Fe(II) site, whereas
Co(Il), Mn(ll), and Ni(II) are weaker inhibitors. NaCI inhibits the enzyme, resulting in
decreases in km, and increases in Km of both ocKG and xanthine. The 01KG cosubstrate can
be substituted by a—ketoadipate (resulting in a 9-fold decrease in kca, and a 5-fold increase
in the Km compared to the normal (x-keto acid), while the OtKG analogue N-oxalyl glycine
is an effective competitive inhibitor (K, 0.12 uM). No alternative purines are able to
effectively substitute for xanthine as a substrate, and only one purine analogue
(6,8-dihydroxypurine, DHP) results in significant inhibition. A homology model of XanA
was generated on the basis of the structure of the related enzyme TauD (PDB code lOS7)
and used to provide insights into the mode of substrate and inhibitor binding. These
studies present the first analysis of xanthine/OLKG dioxygenase isotope effects,

interactions With alternative metal ions and NaCl, and behavior with substrate analogues.

92

INTRODUCTION

In Chapter 2, I described the purification and basic properties of the
Fe(II)-dependent xanthine/a-ketoglutarate (aKG) dioxygenase (XanA) of Aspergillus
nidulans. This enzyme, exclusively associated with the fungal kingdom, catalyzes the
oxidative decarboxylation of OLKG, forming succinate and carbon dioxide, concomitant
with xanthine hydroxylation to uric acid, as shown'in Figure 2.1. I showed that
fungus-derived XanA differed from the recombinant form isolated from Escherichia coli
both in quaternary structure and identity of posttranslational modifications, but the two
enzyme forms exhibited very similar kinetic parameters. In contrast to the
phylogenetically restricted XanA enzyme, a molybdopterin cofactor (Moco)-containing
enzyme xanthine oxidoreductase or xanthine hydroxylase (1) is present in all kingdoms
of life, including fungi, and carries out an analogous reaction, the key step of which is
depicted in Figure 3.1. A glutamic acid general base activates the hydroxyl group
coordinated to Mo(Vl) (chelated by an enedithiolate moiety of the pterin) and the
resulting nucleophile attacks the C—8 position of xanthine with concomitant hydride
transfer to Mo=S. The product subsequently dissociates and the resulting Mo(IV)-SH
intermediate reoxidizes by sequential electron transfer steps through two [2Fe-2S]
clusters and an enzyme bound flavin adenine dinucleotide. The reduced flavin passes on
the electrons to NAD (xanthine dehydrogenase) or in some cases to oxygen (xanthine
oxidase). Whereas the Moco-containing xanthine hydroxylases have been extensively

characterized, little is known of the Fe(II)/0tKG-dependent XanA.

93

FIGURE 3.1: General mechanism of Moco-containing xanthine oxidases

// r) AIL/Em. f“
/:/\:V\H o\o u/KO /M\°§o

\O_</NN/

 

'o _ HO
H%O >¢o
Glu Glu

94

IZ

NH

The Fe(II)/0tKG hydroxylases encompass a wide range of enzymes with diverse
primary substrates (reviewed in (2)). The prolyl, lysyl, and aspartyl(asparaginyl)
hydroxylases catalyze posttranslational modifications of proteins, in some cases
associated with hypoxic signaling (3). J ij domain-containing proteins have been shown
to catalyze methylated-histone demethylation reactions (4). AlkB repairs l-methyl-A or
3-methyl-C lesions in DNA (or RNA) by using an analogous oxidative dealkylation
reaction (5, 6). The lipid-metabolizing enzyme phytanoyl-CoA hydroxylase participates
in the metabolism of phytanic acid, and deficiency of this enzyme leads to Refsum
disease (7). Finally, a wide variety of small molecules are synthesized or decomposed by
members of this enzyme family. For example, plants synthesize gibberellins, flavonoids,
and some alkaloids by action of these enzymes (8), thymine 7-hydroxylase sequentially
hydroxylates the methyl group of free thymine (9), and deA, deA, and SdpA
decompose specific phenoxyalkanoic acid herbicides by hydroxylation of the side chain
(10, 11). The best-characterized representative of this protein family is taurine/OtKG
dioxygenase (TauD), responsible for the decomposition of taurine
(2-aminoethanesulfonate) to aminoacetaldehyde and sulfite (12). In particular, a series of
transient kinetic studies have demonstrated the interrnediacy of an Fe(IV)-0x0 species in
the TauD catalytic mechanism (13-19). In addition, the TauD crystal structure has been
solved (20, 21) and aberrant chemistry leading to side chain hydroxylation reactions has
been extensiVely defined (22-24). The XanA sequence shows some similarity with the

TauD group of dioxygenases (25), including a clear conservation of the Fe(II)- and

95

aKG-binding sites.

The studies reported here continue our characterization of the bacteria-derived forms
of A. nidulans XanA. I describe its solvent and substrate kinetic isotope effects, define the
effects of different divalent metal ions and several OLKG and xanthine analogues, and I
identify alternate substrates and inhibitors. Finally, a homology model of XanA was
generated and I use it to provide insights into the mode of substrate/inhibitor binding.
These studies present the first analysis of isotope effects, interactions with alternative

metal ions and NaCl, and behavior of substrate analogues for xanthine/OLKG dioxygenase.

96

EXPERIMENTAL PROCEDURES

Purification of Bacterial-Derived XanA. XanA was purified from recombinant E.
coli cells as previously described in Chapter 2 (26).

Enzyme Assays. Xanthine/OtKG dioxygenase activity was measured at 25 °C by using
the following typical assay conditions (total volume of 1 mL): 1 mM OtKG, 40 uM
Fe(NH4)2(SO4)2, and 200 uM xanthine in 50 mM MOPS buffer, pH 7.4. Variations of
these conditions included use of varied concentrations of these and other additives. The
absorbance at 294 nm was monitored to determine the uric acid production (8294 12,200
M" cm") with a correction for loss of the xanthine absorbance at this wavelength
(measured 8294 2,000 M" cm") for an overall change in 8294 of 10,200 M" cm". Initial
rates were calculated by using linear data collected over a time period where 10% or less
of the substrate was consumed. Units of activity (U) were defined as umol min" of uric
acid produced and the specific activity (U mg") was measured as umol min" (mg of
purified XanA)".

Sources and Synthesis of Chemical Analogues of aKG and Xanthine. OLKG,
a—ketoadipate, a—ketobutyric acid, pyruvate, phenylpyruvate, 4-hydroxyphenylpyruvate,
purine, 6-methylpurine, 2-hydroxypurine, 2-hydroxy-6-methylpurine, hypoxanthine,
xanthine, l-methylxanthine, 3-methylxanthine, 7-methylxanthine, 9-methylxanthine,
allopurinol, and allantoin were from Sigma-Aldrich. N-oxalylglycine (NOG) was a gift
from Dr. Nicolai Burzlaff.

8-Hydroxypurine and 6,8-dihydroxypurine (6,8-DHP) were prepared from the

97

commercially available (Sigma Aldrich) 4,5-diaminopyrimidine and
4,5-diamino-6-hydroxypyrimidine, respectively, following a modified method described
previously (27). 2,8-Dihydroxypurine (2,8-DHP) was prepared following a published
method (28). Each of these compounds was provided by David J. Lowe and Fabrice
Jourdan.

8-2H-Xanthine Preparation. 8-2H-xanthine was prepared by incubating a 1 % (w/v)
xanthine solution in szO (99.9 %, Sigma & Aldrich), with 0.3 M NaOzH (> 99 % 2H,
Sigma-Aldrich) in a serum vial sealed with a butyl rubber stopper for 20 h in a 100 °C
oven. The proton-deuterium exchange was monitored by using NMR spectroscopy to
integrate the resonance at 6.9 ppm due to the proton on C-8. The final solution was
diluted and neutralized to pH 7.0. The 8-2H-xanthine precipitated from the solution and
was dried under vacuum for at least 3 h. Yields were approximately 90 %. The purity of
8-2H-xanthine was monitored by electrospray ionization mass spectrometry, and the
conversion was shown to be 98.4 % as a molar ratio.

21120 Solvent Isotope Eflect. In 1 mL 50 mM MOPS, p211 8.0 (p2H values were
determined by adding 0.4 to the pH 7.6 electrode reading), 0.5] pg of purified XanA was
assayed in 40 uM Fe(NH4)2(SOo)2, 1 mM aKG and 0-200 uM xanthine. All buffers were
prepared in szO. These data were compared to assays carried out in the same conditions
at pH 7.6. Of note, the E. coli-derived enzyme exhibits optimal activity over a range from
pH 7.0 to 8.0.

Structural Modeling. (Assisted by Andrew W. Stumpff-Kane and Michael Feig,

98

Department of Biochemistry and Molecular Biology, MSU.) A homology model of XanA
was generated on the basis of the structure of the related enzyme TauD (PDB code lOTJ,
chain A, and PDB code IOS7, chain A) (21). XanA is 18 % identical to TauD over 280
residues, and has three significant additional regions of 15, 22, and 18 amino acids.
Multiple templates for the XanA structure were obtained with PSI-BLAST (29, 30)
starting both from the amino acid sequence of XanA and from that of several closely
related proteins for which no structure has yet been reported, and with 3D-Jury (31) using
the Bioinfo Meta Server (http://bioinfo.pl/Meta). Additional suboptimal alignments for.
each template were generated using probA (32) to produce a large pool of possible
models. A structural model was constructed from each alignment, with side chains
reconstructed using the MMTSB Tool Set (33, 34), and the best model from the entire set
of models was selected according to combined energy scores from DFIRE (35),
MMGB/SA (36), and RAPDF (3 7), using a correlation-based approach (38) in
combination with clustering. The orKG and iron were placed into the active site according

to the TauD structure.

99

RESULTS

Isotope Eflects. To test for substrate or solvent isotope effects on the overall reaction
rate, activity assays were carried out by using 8—2H-xanthine or in 2H20 and compared to
control studies with unlabeled xanthine in H20. No significant isotope effect was
observed when the reaction was carried out with 8—2H-xanthine (98.4 % enriched with 2H)
and compared to non-labeled substrate (data not shown). In contrast, a significant solvent
deuterium isotope effect was observed (Figure 3.2), with Vmax reduced by 50 % (72.1 U
mg" dropping to 34.0 U mg") while Km was nearly unaffected (45.2 and 48.9 mM,
respectively).

Eflect of Other Metal Ions and Salt. Metal ions other than Fe(II) were tested and
found to be unable to stimulate activity when added to the apoprotein. When various
metal ions were added to the assay buffer in concentrations equivalent to that of Fe(II),
both Cu(II) and Zn(ll) completely inhibited the xanthine/aKG dioxygenase activity, with
partial inhibition observed with Co(Il), Mn(ll), and (much less pronounced) Ni(II)
(Figure 3.3). The inactive metal ions are presumed to compete for the Fe(II)-binding site.
The activity of XanA was shown to decrease in buffers containing NaCl (Figure 3.4A).
Kinetic analyses revealed that 0.5 M NaCl salt increased the Km of ocKG to 0.74 i 0.08
mM (Figure 3.4B), increased the Km of xanthine to 105 i 5.8 uM (Figure 3.4C), and
decreased the kca, to 35.4 s'1 and 43.2 s", respectively in the two studies. These findings
indicate that the Km of OLKG is significantly affected by ionic strength, while the Km of

xanthine and kca, are less affected, and demonstrate that the salt content of fractions

100

FIGURE 3.2: Solvent deuterium isotope effect on XanA activity. The effects of varying
the concentration of xanthine on xanthine/orKG dioxygenase activity were examined at
25 °C in 50 mM MOPS buffer, pZH 8.0 (I) (p2H values were determined by adding 0.4 to
the pH electrode reading) or pH 7.6 (A) containing 40 uM Fe(ll), 1 mM OLKG, and 0-200

11M xanthine.

 

 

 

o ' 5'0 160 130 260

Xanthine (pM)

101

FIGURE 3.3: Divalent cation inhibition of XanA. The effects of several divalent cations
(M", at 40 uM concentration) on xanthine/(XKG dioxygenase activity were examined by
using the E. coli-derived protein at 25 °C in solutions containing 40 uM Fe(II), 1 mM
aKG and 200 uM xanthine in 50 mM MOPS buffer, pH 7.4. Samples included (I) Fe(II)

only and Fe(Il) plus (2) Mg(ll), (3) Mn(ll), (4) Co(II), (5) Ni(II), (6) Zn(II), (7) Cu(II).

80

60

 

 

1 2 3 4 5 6 7
Fe(II)+M(II)

102

FIGURE 3.4: NaCl inhibition of XanA. (A) The effects of varying NaCl concentrations
on xanthine/aKG dioxygenase activity were examined by monitoring the absorbance at
294 nm over time for the E. coli-derived protein at 25 °C in solutions containing 40 11M
Fe(II), 1 mM aKG and 200 11M xanthine in 50 mM. MOPS buffer, pH 7.4. The
concentrations of NaCl examined were: 0 mM (6); 200 mM (I); 400 mM (A); 600 mM
(V); 800 mM (C). Using 0.5 M NaCl, the effects of varying the concentrations of (B)
(XKG and (C) xanthine were examined. Data in panels B and C represent initial rates and

were fit to the Michaelis-Menten equation.

     

 

 

 

 

0.71 45‘ I
0.3!
E 0.5: g 30"
a 0.4‘ 5
.3 ‘ 15 B
< 0.3:
0.2 ‘
v v v v I *7 T V ‘l 0 jjjjjjjjjj
o 40 30 120 150 o 1 2 3 4 5
Time (3) alpha-KG (W)
45"
3o,

Ulrng

.51 c

 

 

0 V v f 1 v I V T T u .
o 40 00 120 160 200
xanthine (9")

103

recovered during protein isolation must be considered when assaying the enzyme.

aK G Analogues. In addition to (xKG, a-ketoadipic acid is a cosubstrate of XanA and
results in an activity of 9.2 U (mg protein)" or about 1/10 of that observed with (XKG
Kinetic analyses revealed a kca, of 7.6 s" and 3 Km of 0.16 mM for this alternative
co-substrate compared to a kca, of 61 s" and Km of 31 11M for OLKG (26). In contrast,
pyruvate, (x-ketobutyric acid, phenylpyruvate, and 4-hydroxyphenylpyruvate were not
used as co-substrates. NOG, a known inhibitor of several Fe(II)/01KG-dependent
dioxygenase family members (39-43), was shown to compete with OLKG and provided a
K, of 0.12 11M for inhibition of the enzyme (Figure 3.5).

Xanthine Analogues. XanA was shown to be highly specific for xanthine. On the
basis of the spectroscopic assay using standard conditions with 12 nM enzyme, no
activity was detected when the enzyme was assayed with 80, 100, or 200 11M
hypoxanthine, l-methylxanthine, 3-methylxanthine, 7-methylxanthine, 9-methylxanthine,
purine, 6-methy1purine, 2-hydroxypurine, 8-hydroxypurine, 2,8-dihydroxypurine,
2-hydroxy-6-methylpurine, allopurinol, allantoin, or adenosine diphosphate. Similarly,
significant inhibitory effects were not observed with 100 or 200 11M of any of these
compounds (although very modest inhibition was noted with 2,8-DHP). The one
xanthine-like compound that does inhibit the enzyme, but does not serve as a substrate, is
6,8-DHP. The kinetic inhibition mechanism of 6,8-DHP will be discussed in more detail
in Chapter 4.

Structural Model of XanA . XanA was aligned with TauD and a homology model was

104

FIGURE 3.5: NOG inhibition of XanA. (A) The effects of varying concentrations of
NOG (0, 0.2, 0.6, 1.2, and 2 uM) on xanthine/ocKG dioxygenase activity were examined
by using the E. coli-derived protein at 25 °C in solutions containing 40 1.1M Fe(II), 1 mM
aKG and 200 11M xanthine in 50 mM MOPS buffer, pH 7.4. Each set of initial rate data
was fit to the Michaelis-Menten equation. (B) A replot of the values of apparent Km
divided by apparent Vmax as a function of inhibitor concentration.

1001

no.1

 

20 / A

’ ,

 

 

 

0 vvvvv . vi ~fi - . .
o 100 200 300 400 500 600
alpha-Kama)
10 '
8 l-
K
36 ‘
E4 - B
2 I-
oI + L l
0 1 2 3
~061le

105

created by my collaborators using the TauD structure (20, 21) as the template. The overall
sequence identity is 18%, however the sequence identity of the most relevant structural
regions near the active site is 33%. Moreover, given recent advances in structure
prediction, sequence identities as low as 18% identity are commonly sufficient to support
comparative modeling with a good match of predicted secondary structure elements (44,
45) as in the case of XanA. Consequently, it is reasonable to assume that our model of the
XanA structure captures the overall features correctly, while structural details near the
active site are likely represented more accurately. Nevertheless, any predicted model
without further experimental validation remains speculative and is subject to some level
of uncertainty.

As illustrated in Figure 3.6A, the XanA protein is predicted to contain the DSBH
- fold comprised of eight B-strands with connecting loops, as is typical of this enzyme
family (2, 46). Three loops in the sequence, comprising residues 72-88, 173-190, and
219-231, had no cotmterparts in TauD and were not modeled (indicated by boxes at the
appropriate positions in the figure), but these are all distant from the putative active site
region. The homology model contains the Fe(II)-binding site (Hisl49, Asp151, and
His340) expected from prior sequence alignments (25). The co-substrate (shown in
yellow) was positioned into the model so as to chelate F e(II) in a similar fashion as OLKG '
occurs in TauD. The aKG C-5 carboxylate is predicted to form a salt bridge with Arg352
(depicted in red), while Ly5122 (in green) is well positioned to stabilize the C-1

carboxylate of the co-substrate.

106

FIGURE 3.6: Homology model of XanA. A, Ribbon diagram depicting the XanA
homology model that was predicted using TauD (PDB code 1087) as a template. The
polypeptide chain is depicted in blue to red coloration (N-terminus to C-terminus) with
gaps indicated by boxes for the non-modeled loops involving residues 72-88, 173-190,
and 219-231. The postulated metal ligands (Hisl49, Asp151, and His340) are illustrated
in green, yellow, and red, respectively, to the right of the magenta Fe(II) sphere. Bound
aKG (yellow) chelates the metals and is suggested to be stabilized by a salt bridge to the
C-5 carboxylate involving Arg352 (red) and a hydrogen bond to the C-1 carboxylate via
Lysl22 (green). B, Putative active site pocket derived from the XanA homology model.
A closeup view (20 A slab) of the XanA homology model depicting possible positions of
residues at the active site pocket, shown in the same orientation as illustrated in Figure 9.
OtKG in stick form is shown chelating the metal with its carbon atoms colored yellow.
The three side chains that bind the metal are shown in stick form using green (Hisl49),
yellow (Asp151), and red (His340) coloration, as in Figure 9. Residues predicted to line
the active site pocket are shown in stick form with their carbon atoms in white, while

other residues are shown as lines with green carbon atoms.

107

FIGURE 3.6:

 

108

DISCUSSION

In this study I describe several properties of xanthine/OLKG dioxygenase, a novel
xanthine-metabolizing enzyme found exclusively in the fungal kingdom (25). Below, I
place my findings on isotope effects, interactions with various metal ions, salts, and
substrate analogues, into the larger context of other Fe(II)/01KG dioxygenases, and I
relate key findings to our homology model of the protein.

Isotope Eflects. Although the xanthine C-H bond is broken at 08 during turnover,
substitution of the proton at this position by 2H did not lead to a substrate isotope effect.
This result demonstrates that C-H cleavage is not the rate-detennining step in the reaction.
The deuterated substrate might be useful in future experiments to examine individual
steps in the reaction by using transient kinetic approaches, as was elegantly demonstrated
with deuterated substrate and stOpped-flow techniques for TauD (15) and, more recently,
prolyl 4-hydroxylase (4 7). In contrast to the situation with labeled xanthine, a solvent
isotope effect was observed upon substituting H20 with D20 (Figure 3.2). This
substitution had. little effect on the Km of xanthine while decreasing Vmax by 40 %
compared to the assay in H20. This result suggests that a chemical group possessing an
exchangeable proton is important in the rate-detennining step of the overall reaction.
Options for the protonatable group include a general base or general acid protein side
chain or a metallocenter species such as Fe(III)-OOH or Fe(III)-OH. The finding of a
solvent deuterium isotope effect contrasts with the case of TauD, where product release is

the slow step in catalysis and no solvent isotope effect is observed (13, 19).

109

Metal Ion and Salt Eflects. Zn(II) and Cu(II) are potent inhibitors of XanA, and
several other metals also inhibit the enzyme (Figure 3.3). This situation resembles that
known for related enzymes such as TauD where Co(ll) and Ni(II) inhibition has been
studied (39), Clavaminate synthase where Co(lI) inhibition was characterized (48), or
deA where the Cu(II)-inhibited enzyme was analyzed (49). The inhibitory metal ions are
likely to substitute for Fe(II) and utilize the same set of amino acid side chain ligands.

The inhibitory effects of NaCl on XanA (Figure 3.4) represent, to my knowledge, the
only systematic characterization of any salt effect on an Fe(II)/01KG dioxygenase. The
presence of salt leads to a large increase in Km of (XKG, a small increase in Km of xanthine,
and a reduction in km. I attribute the Km effect to the ability of salt to interfere with salt
bridge formation and other stabilizing interactions between OLKG or xanthine and the
protein. Similar salt effects are likely to apply to a wide range of other family members;
thus, one must exercise caution in the choice of ionic strength when doing enzyme
assays.

Co-Substrate and Substrate Specificity. The co-substrate specificity of XanA is
somewhat more relaxed than that for the primary substrate, with a—ketoadipic acid (with
one extra carbon compared to aKG) also yielding activity. The increase in Km and
decrease in km, for the incorrectly-sized analogue is easily rationalized in terms of the
XanA homology model where both the C-1 and C—5 carboxylates of aKG are predicted to
interact with the protein (with Ly5122 and Arg352, respectively) while also chelating the

active site metal ion. Alternative OL-ketoacids are known to support (xKG-dependent

110

activities of several other enzyme family members including TauD, deA, deA, SdpA,
and an alkyl sulfatase (10, 12, 50, 51). The aKG homologue NOG is a competitive
inhibitor of XanA (Figure 3.5), consistent with its known inhibition of several other
representatives of the Fe(Il)/(xKG-dependent hydroxylases. Of interest, the measured K,
of NOG for XanA (0.12 11M) is well below the Km of OLKG in this enzyme and much
below the reported 290 11M K, for inhibition of TauD (39). More generally, the K, of NOG
can vary widely among family members (e.g., it is reported to be l.9-7.0 11M for collagen
prolyl 4-hydroxylase (40) and 1.2 mM for an oxygen-sensing asparaginyl hydroxylase
(43)), presumably due to distinct interactions with the active site protein side chains in
the target enzymes.

XanA proved to be exquisitely specific to its primary substrate. For example,
allopurinol (a known substrate and inhibitor of xanthine oxidase (52)), l-methylxanthine
and 2-hydroxy-6-methylpurine (alternative substrates of the Moco-containing enzyme (53,
54), and several other purine-type compounds were neither substrates nor inhibitors of
XanA. Of the compounds tested, only 6,8-DHP bound tightly to the enzyme (more date .
and discussion of this inhibitor are provided in Chapter 4). For comparison, other
members of the Fe(II)/01KG-dependent dioxygenases range widely in their substrate
specificities. The prolyl hydroxylases involved in the hypoxic response appear to be
highly specific for recognizing a single prolyl residue in the HlFlor protein (55, 56). By
contrast, deA utilizes a wide range of phenoxyacetic acids (10) and a yeast

orKG/sulfonate dioxygenase metabolizes a diverse array of sulfonates (5 7).

ll]

Postulated Substrate-Binding Mode. Additional structural or mutagenesis studies are
required to characterize the mode of substrate binding to XanA; however, our homology
model (Figure 3.6) allows me to identify potential key active site residues (Figure 3.6B).
the model depicts a pocket adjacent to the metallocenter and lined by a series of putative
active site residues (Gln99, ProlOO, GlnlOl, IlellO, Thr120, Lysl22, Glul37, Ala152,
leu154, Gln356, and Asn358) that are generally well conserved in sequences of XanA
orthologues (Figure 3.7). Pr0100, GlnlOl, Gln356, and Asn358 are universally conserved
in the XanA sequences. Gln99 counterparts also are present often, but Val occupies this
position in some representatives. Lysl22, which is suggested to stabilize the OLKG C-l
carboxylate and bind substrate, either is retained or conservatively replaced by Arg or Thr.
In some fungi, various residues (Lys, Gln, Arg, Asn, and Glu) replace Thr120, but all of
these are ble to function in hydrogen bonding. Similarly, Asp, Gln, and Ser, all capable of
similar hydrogen bond interactions, replace Glul37 in other XanA homologues. Among
the hydrophobic residues predicted to surround the active site, Ile110 is replaced by Val
or Phe, Ala152 is retained or replaced by Ser, Leu154 is strictly conserved, and Leu251
(not shown) is retained or replaced by other large side chains in other homologues.
Aromatic groups, often involved in tt-tt stacking interactions with nucleic acids and
known to occur in xanthine hydroxylase (58, 59) and uric acid oxidase (60), do not
appear to be important for binding xanthine in xanthine/orKG dioxygenase. These
predictions set the stage for future chemical modification, mutagenesis, and structural

efforts to test these interactions.

112

FIGURE 3.7: Comparison of sequences from XanA orthologues in selected fungi. Black
shading represents identical sequences and gray shading indicates conservative

replacements.

chryaosporfum
cinereual
neotorman
graminearum
crassa
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maydie

po 9
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U
a
2
H
B
Q

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cinereusl 7 ' ' KKSILHPDLKTI PO' VQ

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maydls

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cinereusz

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chrysosporium s 'K I-hlnnGTGDh=
cinereuel 17 OTYDHCTGDFIFI
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113

FIGURE 3.7':

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.neotorman
.graminearum
.crassa
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.maydls

.pombe
.albtcans
.ctnereusz

.chrysosporlum
.ctnereusl

neotorman
graminearum

. crassa
.ntdulans

mayors

albtcans
c1nereus2

.chryaosporlum
.clnoreuel
.neotorman
.gramrnearum
.craaaa
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.maydls

.albrcans
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432

380

387

413

371

 

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ll4

REFERENCES

(1)

(2)

(3)

(4)

(5)

(6)

(7)

(8)

(9)

(10)

Hille, R. (2005) Molybdenum-containing hydroxylases. Arch. Biochem. Biophys.
433, 107-116.

Hausinger, R. P. (2004) Fe(II)/01-ketoglutarate-dependent hydroxylases and
related enzymes. Crit. Rev. Biochem. Mol. Biol. 39, 21-68.

Schofield, C. J., and Ratcliffe, P. J. (2004) Oxygen sensing by hydroxylases. Nat.
Rev. Molec. Cell Biol. 5, 343-354.

Tsukada, Y.-l., Fang, J., Erdjument-Bromage, H., Warren, M. E., Borchers, C. H.,
Tempst, P., and Zhang, Y. (2006) Histone demethylation by a family of ijC
domain-containing proteins. Nature 439, 811-816.

Trewick, S. C., Henshaw, T. F., Hausinger, R. P., Lindahl, T., and Sedgwick, B.
(2002) Oxidative demethylation by Escherichia coli AlkB directly reverts DNA
base damage. Nature 419, 174-178.

Aas, P. A., Otterlei, M., Falnes, P. O., Vagbe, C. B., Skorpen, F., Akbari, M.,
Sundheim, O., Bjoras, M., Slupphaug, G, Seeberg, E., and Krokan, H. E. (2003)
Human and bacterial oxidative demethylases repair alkylation damage in both
RNA and DNA. Nature 421, 859-863.

Mukherji, M., Chien, W., Kershaw, N. J., Clifton, I. J., Schofield, C. J.,
Wierzbicki, A. S., and Lloyd, M. D. (2001) Structure-function analysis of

phytanoyl-CoA 2-hydroxylase mutations causing Refsum's disease. Human Molec.
Genet. 10, 1971-1982.

Prescott, A. G, and John, P. (1996) Dioxygenases: molecular structure and role in
plant metabolism. Annu. Rev. Plant Physiol. Plant Molec. Bio]. 47, 245-271.

Thomburg, L. D., Lai, M.-T., Wishnok, J. S., and Stubbe, J. (1993) A non-heme

iron protein with heme tendencies: an investigation of the substrate specificity of
thymine hydroxylase. Biochemistry 32, 14023-14033.

Fukumori, F., and Hausinger, R. P. (1993) Purification and characterization of

2,4-dichlorophenoxyacetate/or-ketoglutarate dioxygenase. J. Biol. Chem. 268,
24311-24317.

115

(11)

(12)

(13)

(14)

(15)

(16)

(17)

(18)

(19)

Miiller, T. A., Zavodszky, M. 1., Feig, M., Kuhn, L. A., and Hausinger, R. P. (2006)
Structural basis for the enantiospecificities of R- and
S-phenoxypropionate/(x-ketoglutarate dioxygenases. Prot. Science 15, 1356-1368.

Eichhom, E., van der Ploeg, J. R., Kertesz, M. A., and Leisinger, T. (1997)
Characterization of a—ketoglutarate-dependent taurine dioxygenase from
Escherichia coli. J. Biol. Chem. 2 72, 23031-23036.

Ryle, M. J., Padmakumar, R., and Hausinger, R. P. (1999) Stopped-flow kinetic
analysis of Escherichia coli taurine/Ot-ketoglutarate dioxygenase: interactions
with a—ketoglutarate, taurine, and oxygen. Biochemistry 38, 15278-15286.

Price, J. C., Barr, E. W., Tirupati, B., Bollinger, J. M., Jr., and Krebs, C. (2003)
The first direct characterization of a high-valent iron intermediate in the reaction
of an a—ketoglutarate-dependent dioxygenase: a high-spin Fe(IV) complex in

taurine/Ot-ketoglutarate dioxygenase (TauD) from Escherichia coli. J. Biol. Chem.
42, 7497-7508.

Price, J. C., Barr, E. W., Glass, T. E., Krebs, C., and Bollinger, J. M., Jr. (2003)
Evidence for hydrogen abstraction from C1 of taurine by the high-spin Fe(IV)

intermediate detected during oxygen activation by taurineza-ketoglutarate
dioxygenase (TauD). J. Am. Chem. Soc. 125, 13008-13009.

Proshlyakov, D. A., Henshaw, T. F., Monterosso, G R., Ryle, M. J., and Hausinger,
R. P. (2004) Direct detection of oxygen intermediates in the non-heme Fe enzyme
taurine/Ot-ketoglutarate dioxygenase. J. Am. Chem. Soc. 126, 1022-1023.

Riggs-Gelasco, P. J., Price, J. C., Guyer, R. B., Brehm, J. H., Barr, E. W.,
Bollinger, J. M., Jr., and Krebs, C. (2004) EXAF S spectroscopic evidence for an
Fe=O unit in the Fe(IV) intermediate observed during oxygen activation by
taurineza-ketoglutarate dioxygenase. J. Am. Chem. Soc. 126, 8108-8109.

Price, J. C., Barr, E. W., Hoffart, L. M., Krebs, C., and Bollinger, J. M., Jr. (2005)
Kinetic dissection of the catalytic mechanism of taurineza-ketoglutarate
dioxygenase (TauD) from Escherichia coli. Biochemistry 44, 8138-8147.

Grzyska, P. K., Ryle, M. J., Monterosso, G R., Liu, J., Ballou, D. P., and
Hausinger, R. P. (2005) Steady-state and transient kinetic analyses of
taurine/or-ketoglutarate dioxygenase: effects of oxygen concentration, alternative
sulfonates, and active site variants on the Fe(IV) intermediate. Biochemistry 44,

116

(20)

(21)

(22)

(23)

(24)

(25)

(26)

(27)

3845-3855.

Elkins, J. M., Ryle, M. J., Clifton, I. J., Dunning Hotopp, J. C., Lloyd, J. S.,
Burzlaff, N. 1., Baldwin, J. E., Hausinger, R. P., and Roach, P. L. (2002) X-ray
crystal structure of Escherichia coli taurine/Ot-ketoglutarate dioxygenase
complexed to ferrous iron and substrates. Biochemistry 41, 5185-5192.

O'Brien, J. R., Schuller, D. J., Yang, V. S., Dillard, B. D., and Lanzilotta, W. N.
(2003) Substrate-induced conformational changes in Escherichia coli

taurine/Ot-ketoglutarate dioxygenase and insight into the oligomeric structure.
Biochemistry 42, 5547-5554.

Ryle, M. J., Liu, A., Muthukumaran, R. B., Ho, R. Y. N., Koehntop, K. D.,
McCracken, J., Que, L., Jr., and Hausinger, R; P. (2003) 02- and
or-ketoglutarate-dependent tyrosyl radical formation in TauD, an a—keto
acid-dependent non-heme iron dioxygenase. Biochemistry 42, 1854-1862.

Ryle, M. J., Koehntop, K. D., Liu, A., Que, L., Jr., and Hausinger, R. P. (2003)
Interconversion of two oxidized forms of TauD, a non-heme iron hydroxylase:
evidence for bicarbonate binding. Proc. Natl. Acad. Sci. USA 100, 3790-3795.

Koehntop, K. D., Marimanikkuppam, S., Ryle, M. J., Hausinger, R. P., and Que,
L., Jr. (2006) Self-hydroxylation of taurine/a—ketoglutarate dioxygenase.
Evidence for more than one oxygen activation mechanism. J. Biol. Inorg. Chem.
11, 63-72.

Cultrone, A., Scazzocchio, C., Rochet, M., Montero-Moran, G, Drevet, C., and
Fernandez-Martin, R. (2005) Convergent evolution of hydroxylation mechanisms
in the fungal kingdoms: molybdenum cofactor-independent hydroxylation of

xanthine via or-ketoglutarate dependent dioxygenase. Molec. Microbiol. 5 7,
276-290.

Montero-Moran, G M., Li, M., Rendon-Huerta, E., Jourdan, F., Lowe, D. J.,
Stumpff—Kane, A. W., Feig, M., Scazzocchio, C., and Hausinger, R. P. (2007)
Purification and characterization of the Fe(II)- and on-ketoglutarate-dependent
xanthine hydroxylase from Aspergillus nidulans. Biochemistry 46, 5293-5 304.

Albert, A., and Brown, D. J. (1954) Pruine studies. Part 1. Stability to acid and

alkali. Solubility. Ionization. Comparison with pteridines. J. Chem. Soc,
2060-2071.

117

(28)

(29)

(30)

(31)

(32)

(33)

(34)

(35)

(36)

(37)

(38)

Dornow, a., and Hinz, E. (1958) Syntheses of nitrogen-containing heterocycles.
XVIII. ortho-condensations of heterocyclic o-aminocarboxylic acid derivatives.
Chem. Ber. 91, 1834-1840.

Altschul, S. F., Madden, T. L., Schaffer, A. A., Zhang, J. H., Zhang, Z., Miller, W.,
and Lipman, D. J. (1997) Gapped BLAST and PSI-BLAST: A new generation of
protein database search programs. Nucl. Acids Res. 25, 3389-3402.

Schaffer, A. A., Aravind, L., Madden, T. L., Shavirin, S., Sponge, J. L., Wolf, Y. I.,
Koonin, E. V., and Altschul, S. F. (2001) Improving the accuracy of PSI-BLAST
protein database searches with composition-based statistics and other refinements.
Nucleic Acids Research 29, 2994-3005.

Ginalski, K., Elofsson, A., Fisher, D., and Rychlewski, L. (2003) 3D-Jury: a
simple approach to improve protein structure predictions. Bioinformatics 19,
1015-1018.

Muckstein, U., Holfacker, I. L., and Stadler, P. F. (2002) Stochastic pairwise
alignments. Bioinformatics 18 (Suppl. 2), 8153-8160.

Feig, M., Rotkiewicz, P., Kolinski, A., Skolnick, J., and Brooks, C. L., 111. (2000)
Accurate reconstruction of all-atom protein representations from side-chain-based
low-resolution models. Proteins 41, 86-97.

Feig, M., Karanicolas, J., and Brooks, C. L., 111. (2004) MMTSB tool set:
enhanced sampling and multiscale modeling methods for applications in structural
biology. J. Molec. Graphics Modeling 22, 377-395.

Zhang, C., Liu, S., Zhu, Q. Q., and Zhou, Y. Q. (2005) A knowledge-based energy
function for protein-ligand, protein-protein, and protein-DNA complexes. J. Med.
Chem. 48, 2325-2335.

Feig, M., and Brooks, C. L., 111. (2002) Evaluating CASP4 predictions with
physical energy functions. Proteins: Struct. Funct. Genet. 49, 232-245.

Samudrala, R., and Moult, J. (1998) An all-atom distance-dependent conditional
probability discriminatory function for protein structure prediction. J. Mo]. Bio].
275, 895—916.

Stumpff-Kane, A. W., and Feig, M. (2006) A correlation-based method for the

enhancement of scoring functions on funnel-shaped energy landscapes. Proteins:

ll8

(39)

(40)

(41)

(42)

(43)

(44)

(45)

(46)

(47)

(48)

Struct., F unct. Bioinform. 63, 155-164.

Kalliri, E., Grzyska, P. K., and Hausinger, R. P. (2005) Kinetic and spectroscopic
investigation of Co”, Ni”, and N-oxalylglycine inhibition of the
Fe(II)/(x-ketoglutarate dioxygenase, TauD. Biochem. Biophys. Res. Commun. 338,
191-197.

Baader, E., Tschank, G, Baringhaus, K. H., Burghard, H., and Gunzler, V. (1994)
Inhibition of prolyl 4-hydroxylase by oxalyl amino acid derivatives in vitro, in
isolated microsomes and in embryonic chicken tissues. Biochem. J. 300, 525-530.

Cunliffe, C. J., Franklin, T. J., Hales, N. J ., and Hill, G B. (1992) Novel inhibitors
of prolyl 4—hydroxylase. 3. Inhibition by the substrate analogue N-oxaloglycine
and its derivatives. J. Med. Chem. 35, 2652-2658.

Chan, D. A., Sutphin, P. D., Denko, N. C., and Giaccia, A. J. (2002) Role of prolyl
hydroxylation in oncogenically stabilized hypoxia-inducible factor-101. J. Biol.
Chem. 277, 40112-40117.

McDonough, M. A., McNeill, L. A., Tilliet, M., Papmichael, C. A., Chen, Q.-Y.,
Banerji, B., Hewitson, K. S., and Schofield, C. J. (2005) Selective inhibition of
factor inhibiting hypoxia-inducible factor. J. Am. Chem. Soc. 127, 7680-7681.

Tramontano, A., and Morea, V. (2003) Assessment of homology-based predictions
in CASP5. Proteins: Struct. Funct. Genet. 53, 352-368.

Kryshtafovych, A., Venclovas, C., Fidelis, K., and Moult, J. (2005) Progress over
the first decade of CASP experiments. Proteins: Struct. Funct. Genet. S7,
225-236.

Clifion, I. J., McDonough, M. A., Ehrismann, D., Kershaw, N. J., Granatino, N.,
and Schofield, C. J. (2006) Structural studies on 2-oxoglutarate oxygenases and
related double-stranded B-helix fold protein. .1. Inorg. Biochem. 100, 644-669.

Hoffart, L. M., Barr, E. W., Guyer, R. B., Bollinger, J. M., Jr., and Krebs, C. (2006)
Direct spectroscopic detection of a C-H-cleaving high-spin Fe(IV) complex in a
prolyl-4-hydroxylase. Proc. Nat]. Acad. Sci. USA 103, 14738-14743.

Busby, R. W., Chang, M. D.-T., Busby, R. C., Wimp, J., and Townsend, C. A.

(1995) Expression and purification of two isozymes of Clavaminate synthase and
initial characterization of the iron binding site. General error analysis in

119

(49)

(50)

(51)

(52)

(53)

(54)

(55)

(56)

(57)

polymerase chain reaction amplification. J. Biol. Chem. 270, 4262-4269.

Hegg, E. L., Whiting, A. K., Saari, R. E., McCracken, J., Hausinger, R. P., and
Que, L., Jr. (1999) Herbicide-degrading or-keto acid-dependent enzyme deA:
metal coordination environment and mechanistic insights. Biochemistry 38,
16714-16726.

Miiller, T. A., Fleischmann, T., van der Meer, J. R., and Kohler, H.-P. E. (2006)
Purification and characterization of two enantioselective

(x-ketoglutarate-dependent dioxygenases, deA and SdpA, from Sphingomonas
herbicidovorans MH. Applied and Environmental Microbiology 72, 4853-4861.

Kahnert, A., and Kertesz, M. A. (2000) Characterization of a sulfur-regulated
oxygenative alkylsulfatase from Pseudomonas putida S-3l3. J. Biol. Chem. 275,
31661-31667.

Elion, G B., Kovensky, A., and Hitchings, G. H. (1966) Metabolic studies of
allopurinol, an inhibitor of xanthine oxidase. Biochem. Pharmacol. 15, 863-880.

Edmondson, D., Ballou, D. P., Vn Heuvelen, A., Palmer, G, and Massey, V. (1973)

Kinetic studies on the substrate reduction of xanthine oxidase. J. Biol. Chem. 248,
6135-6144.

McWhirter, R. B., and Hille, R. (1991) The reductive half-reaction of xanthine
oxidase. Identification of spectral intermediates in the hydroxylation of
2-hydroxy-6-methylpurine. J. Biol. Chem. 266, 23724-23731.

Ivan, M., Kondo, K., Yang, H., Kim, W., Valiando, J., Ohh, M., Salic, A., Asara, J.
M., Lane, W. S., and Kaelin, W. G, Jr. (2001) HIFa targeted for VHL-mediated
destruction by proline hydroxylation: implications for 02 sensing. Science 292,
464-467.

Jaakkola, P., Mole, D. R., Tian, Y.-M., Wilson, M. I., Gielbert, J., Gaskell, S. J.,
von Kriegsheim, A., Hebestreit, H. F., Mukherji, M., Schofield, C. J ., Maxwell, P.
H., Pugh, C. W., and Ratcliffe, P. J. (2001) Targeting of HIF-01 to the von
Hippel-Lindau ubiquitylation complex by Oz—regulated prolyl hydroxylation.
Science 292, 468-472.

Hogan, D. A., Auchtung, T. A., and Hausinger, R. P. (1999) Cloning and

characterization of a sulfonate/(x-ketoglutarate dioxygenase from Saccharomyces
cerevisiae. J. Bacteriol. 181, 5876-5879. .

120

(58)

Okamoto, K., Matsumoto, K., Hille, R., Eger, B. T., Pai, E. F., and Nishino, T.
(2004) The crystal structure of xanthine oxidoreductase during catalysis:
implications for reaction mechanism and enzyme inhibition. Proc. Natl. Acad. Sci.

USA 101, 7931-7936.

(59) Truglio, J. J ., Theis, K., Leimkiihler, S., Rappa, R., Rajagopalan, K. V., and Kisker,

(60)

C. (2002) Crystal structure of the active and alloxanthine-inhibited forms of
xanthine dehydrogenase from Rhodobacter capsulatus. Structure 10, 115-125.

Retailleau, P., Colloc'h, N., Vivarés, D., Bonneté, F., Castro, E., El Hajji, M.,
Momon, J.-P., Monard, G, and Prangé, T. (2004) Complexed and ligand-free
high-resolution structures of urate oxidase (on) from Aspergillus flavus: a
reassignment of the active-site binding mode. Acta Crystallogr. D60, 453-462.

121

CHAPTER 4

Characterization of Active Site Variants of Xanthine Hydroxylase from

Aspergillus nidulans

Tina Muller constructed the XanA variants and carried out oxygen consumption

studies.

122

ABSTRACT

Xanthine/a-ketoglutarate (OtKG) dioxygenase (XanA) is a non-heme mono-
nuclear Fe(ll) enzyme that decarboxylates orKG to succinate and C02 while
catalyzing the hydroxylation of Xanthine to generate uric acid. In the absence of a
XanA crystal structure, a homology model was used to target several putative active
site residues for mutagenesis. Wild-type XanA and ten enzyme variants were purified
from recombinant Escherichia coli cells and extensively characterized. From analysis
of the quenching of the endogenous fluorescence of XanA, the H149A, D151A, and
H340A mutants displayed significant increases in K, of Fe(II) and the K122A variant
exhibited a large increase in Kd of OLKG, consistent with the proposed roles of the
corresponding residues in Fe(II)- and OtKG-binding. The H149A and DlSlA variants
were inactive whereas the H340A variant exhibited 0.13 U mg" (0.17 % of the
wild-type enzyme). The N358A variant exhibited the next largest change in
xanthine-related kinetics with a 12-fold larger Km and 2-fold decrease in km, compared
to wild-type XanA, pointing to a key role of Asn358 in catalysis. The E137A and
D138A variants demonstrated enhanced activity with 9-methy1xanthine, a poor
substrate of the enzyme, consistent with Glul37 and Aspl 38 being proximal to N-9 of
substrate. The Q356A and N358A variants had significantly increased K?” over
control protein for 6,8-dihydroxypurine, identified as a slow-binding competitive
inhibitor of XanA, suggesting that Gln356 and Asn358 hydrogen bond with the C-6
hydroxyl group of substrate. In contrast, the K?” decreased for the E137A and

D138A proteins, consistent with repulsion between these carboxylates and the

123

deprotonated C-8 hydroxyl group suspected to bind Fe(II). Support for Cys357
residing at the active site was obtained by using thiol-specific reagents that inactivated
wild-type enzyme with partial protection by substrate, whereas the C357A variant was
resistant to these reagents. In the absence of substrates, the Q101A, Q356A, and
C357A variants showed elevated ferroxidase activity, indicating increased oxygen
reactivity of their metallocenters and/or enhanced Fe(II) access to those sites. These
results were combined into a model depicting Fe(II) and substrate interaction with the
XanA active site and provide insight into the specificity of the enzyme and selected

aspects of its reactivity.

124

INTRODUCTION

Xanthine dehydrogenases and xanthine oxidases are molybdopterin cofactor
(Moco)-containing enzymes that transform xanthine into uric acid (1). These enzymes
are conserved throughout living Organisms, including archaea, bacteria, fungi, plants,
and metazoans. Surprisingly, mutants of Aspergillus nidulans with known defects in
xanthine dehydrogenase activity (i.e., with mutations affecting the structural gene th,
the cnx genes for Moco synthesis, or hrB for sulfuration of Moco) were found to
retain the ability to grow on xanthine as sole nitrogen source (2). This capacity arises
from an alternative xanthine-degrading activity encoded by the xanA gene found only
in selected fungi (3). Recombinant Hisé-tagged XanA protein was purified from both
A. nidulans mycelia and Escherichia coli cells, extensively characterized, and shown
to be an Fe(II)- and a-ketoglutarate (OLKG)-dependent hydroxylase that catalyzes the
reaction shown in Figure 2.1 (4). On the basis of its sequence similarity to
taurine/aKG dioxygenase or TauD (5), a well-studied and crystallographically
characterized member of the Fe(II)/0tKG dioxygenase family (6, 7, 8), a homology
model was constructed for XanA (4). The overall sequence identity of XanA and
TauD is only 18 % over 280 residues and three loops in XanA could not be modeled;
however, the relevant structural regions near the active sites are 33 % identical and it
is reasonable to assume that the model captures the overall features correctly. The
XanA model (Figure 4.1) predicts that (i) Fe(II) binds to the Hisl49, Asp151, and
His340‘ residues of the protein, (ii) (xKG chelates the metal by using its C-l

carboxylate and C-2 keto group, with its C-l carboxylate further stabilized by

125

FIGURE 4.1: Depiction of the putative active site pocket of XanA revealed from a
homology model. Hisl49, Asp151, and His340 side chains (blue) and aKG (yellow)
coordinate the Fe(H)(orange sphere). Seven residues lining the putative active site and
capable of participating in hydrogen-bonding interactions are colored green (Gln99,
Ly5122, Glul37, Asp138, Gln356, Cys357, Asn358), with another shown in purple

(GlnlOl) at the active site entrance.

:1
(3513357 .. Asn358
,“ \
C

‘\ . ,
Asp151 l’ ‘
~ --- Gln99 \
Hrs34o Gln356
Lys122 A
W "b , .9
H1949 :. p *1 4’
1 :3 F Gln101 -.
arKG .

l .
\\~_ 1 Glu137

Asp138

126

interaction with Ly3122 and its C-5 carboxylate forming a salt bridge with Arg352,
and (iii) xanthine binds in an active site pocket lined with potential hydrogen bond
donors or acceptors (Gln99, GlnlOl, Glul37, Asp138, Ly3122, Gln356, Cys357, and
Asn358) whereas aromatic residues are not important for binding substrate.

In this study, the three residues thought to bind Fe(II) and each of the potential
hydrogen-bondin g residues at the XanA active site was replaced by Ala, a residue that
is incapable of hydrogen bonding. Fluorescence quenching methods were used to
assess the K, values of Fe and OtKG for the mutant enzymes. The kinetic properties of
the variants were analyzed with xanthine, the altemate substrate 9-methylxanthine,
and 6,8-dihydroxypurine (6,8-DHP), which was shown to be a slow-binding
competitive inhibitor of XanA. Chemical modification studies were carried out with
thiol-specific reagents for the wild-type enzyme and the C357A variant. Finally, the
reactivity of each metallocenter with oxygen was examined in the absence of substrate.
On the basis of these results, we identify critical residues that participate in binding of
Fe(II), orKG, and the primary substrate, and we obtain insight into the enzyme

reactivity.

127

EXPERIMENTAL PROCEDURES

Site-Directed Mutagenesis, Protein Overproduction, and Enzyme Purification.
The Q99A, Q101A, K122A, E137A, D138A, H149A, D151A, H340A, Q356A,
C357A, and N358A variants of XanA were created by mutagenesis of xanA (encoding
a Hisb-tagged version of XanA, termed wild-type XanA for convenience) in a
modified version of vector pThioHisC (4) using the Quickchange II site-directed
mutagenesis kit (Stratagene). Each mutation was confirmed by sequence analysis
(Davis Sequencing, Davis, CA). Plasmids were transformed into XLlBlue E. coli
cells (Stratagene) and the variant proteins were overproduced during cell growth in
Luria Base Broth (Difco) and induced by using isopropyl-B-D-thiogalactopyranoside
as described previously (4). Cultures were harvested by centrifugation at 8,000 g for
10 min at 4 °C, the cells were disrupted by use of a French pressure cell, membranes
and insoluble materials were removed by ultracentrifugation (45 min at 100,000 g),
and the wild-type and variant forms of XanA were purified by using Ni-loaded
nitrilotriacetic acid (NTA)-bound resin, as previously reported (4).

Fluorescence Quenching Analyses of Fe(II) and 6K0 Binding. Fluorescence
measurements were made on a luminescence spectrometer, model LS-50B (Perkin
Elmer Limited, UK). The temperature of the cells was maintained at 25 °C.
Fluorescence measurements were carried out at an excitation wavelength of 280 nm (10
nm band-width) with emission monitored from 300 to 400 nm (5 nm bandwidth).
Samples were prepared in 50 mM Tris buffer, pH 8.0, and Fe(II) or aKG was added

to 35 11M or 350 11M, respectively. The data were fit to equation 1, where [L] is the

128

ligand (Fe(II) or aKG) concentration.
Afluorescence = Afluorescencemax [L]/(Kd + [L]) (1)

Enzyme Assays. Enzymatic assays of XanA and its variants were carried out at
25 °C by using the following typical assay conditions (total volume of 1 mL): 1 mM
aKG, 40 11M Fe(NH4)2(SO4)2, and 200 11M xanthine in 50 mM MOPS buffer, pH 7.4
(4). For kinetic analyses, the concentrations of orKG or xanthine were varied while
holding the concentrations of the other components constant. The absorbance at 294
nm was monitored to determine the uric acid production (overall change in 8294 of
10,200 M" cm"). Units of activity (U) were defined as umol min'l of uric acid
produced and the specific activity (U mg") was measured as umol min" (mg of
purified XanA)" as described before (4). Analogous studies were carried out with
9-methylxanthine (Sigma-Aldrich), but in this case the absorbance at 291 nm was
used to calculate the production of 9-methyluric acid (overall change in 829, of 6,800
M'l cm"). Trace activity was detected with l-methylxanthine (Sigma-Aldrich), but
complete kinetic studies could not be performed.

In addition to the spectrophotometric measure of xanthine conversion to uric
acid (or the use of substituted xanthines to form substituted uric acids), the various
forms of the enzyme were analyzed for ferroxidase activity (reduction of oxygen by
using excess ferrous ions as reductant) by use of a Clark-type oxygen electrode. These
assays were carried out in air-saturated MOPS medium (pH 7.4) at 25 °C. Less than
12% of the initial levels of 02 were consumed in these assays, and the data were fit to

equation 2. [02], is the 02 concentration at time t, [02],, is the initial 02 concentration, vb

129

is the background rate of 02 reduction under the given conditions, v0 is the ferroxidase
initial velocity, and kapp is the apparent first-order rate constant for the transition from V0
to vb.

[02]: = [0216 — vb! —(vo - vb)[1-6XP(-kappt)l/kapp (2)

As a complement to the electrode assay for 02 reduction, the production of
hydrogen peroxide was quantified by using a spectrophotometric assay. Timed aliquots
of reaction mixtures were added to assay solutions containing 100 mM potassium
phosphate (pH 5.0), 8.7 mM 2,2’-azino-bis(3-ethylbenzthiazoline-6-sulfonic acid)
(ABTS), and approximately 0.003 units of horseradish peroxidase (Sigma-Aldrich).
The oxidation of ABTS was monitored at 420 nm and compared to a standard curve
generated with hydrogen peroxide.

A final method to assay the activity of the enzyme focused on quantification of the
OtKG consumed during the reaction. Aliquots (250 11L) of reaction mixtures were
. incubated for selected time periods, quenched by addition of 1 mL 0.5 mg/ml
o-phenylenediamine (OPDA, Sigma-Aldrich) stock solution (dissolved in 1 M
phosphoric acid, pH 2, containing 0.25% (v/v) B-mercaptoethanol), and the samples
were heated for 3 min at 100 °C. The absorbance at 333 nm was monitored to
determine aKG consumption by comparison to standard curves.

Kinetic Analysis of 6,8-DHP Inhibition. Spectrophotometric progress curves
(containing 90 or more data points, typically at 15 5 intervals) were initiated by
adding XanA to solutions containing several fixed concentrations of substrate and

selected concentrations of 6,8-DHP (graciously provided by D. J. Lowe and F.

130

Jourdan). The data were analyzed according to equation 3 by using a nonlinear
regression program to give the individual parameters for each progress curve: A
(absorbance at 294 nm), A0 (for baseline correction), v, (initial velocity), v,
(steady-state velocity), and kobs (apparent first-order rate constant for the transition
from v, to v,).

A = A0 + Vst + (Vi' Vs)[]' exp('kObst)]/ kobs (3)

131

RESULTS

Site-directed variants of recombinant A. nidulans XanA, an Fe(II)- and
OtKG-dependent hydroxylase of xanthine, were investigated to further characterize the
protein features important to catalysis. In the absence of a crystal structure for XanA,
we relied on apreviously reported homology model of XanA (4) that identifies the
likely triad of metal-binding residues and predicts the active site pocket is lined by
eight residues capable of hydrogen bonding interactions with substrate. To test this
hypothetical‘model,‘ Ala was used to replace each of these residues and the enzymatic
properties of the resulting variant proteins were characterized. In particular, the
binding of Fe(lI) and OtKG to the XanA variants was assessed by fluorescence
quenching methods and the kinetic properties were studied using xanthine, the
alternative substrate '9-methylxanthine, and 6,8-DHP, a known XanA inhibitor (4). In
addition, the behavior of wild-type enzyme and C357A variant were compared using
thiol-specific chemical modification reagents. Finally, the ferroxidase activities of the
active mutant proteins Were examined in the absence of primary substrate.

Fe(II) and otKG Binding to XanA and its Variants. Eleven XanA variants (Q99A,
Q101A, K122A, E137A, D138A, H149A, D151A, H340A, Q356A, C357A, and
N358A) were produced in E. coli XLlBlue cells as Hisé-tagged fusion proteins, along
with the wild-type protein. Seven variant proteins were purified to homogeneity from
cell extracts by Ni-NTA chromatography whereas three (H149A, D151A, and H340A)
were enriched by this chromatographic step (Figure 4.2), and Q99A XanA failed to

bind to Ni-loaded NTA resin. We interpret the Q99A XanA results to indicate protein

132

FIGURE 4.2: Denaturing polyacrylamide gel electrophoretic analysis of XanA
variants purified from E. coli. Lane I, markers; lane 2, Q101A (1.5 pg) ; lane 3,
K122A (4.2 pg); lane 4, E137A (1.0 pg); lane 5, D138A (1.2 pg); lane 6, Q356A (0.5
pg); lane 7, C357A (0.3 pg); lane 8, N358A (6.4 pg); lane 9, H149A (0.17 pg); lane
10, D151 (0.8 pg); lane 11, H340A (0.13 pg) variant protein. Stacking and running

gels contain 5 % and 12 % acrylamide, respectively.

kDa l 2 3 4 5 6 7 8
97.4
66.2

45.0

    

31.0

21.5

 

l 4.4

133

misfolding during overexpression, and this variant was not further investigated.

The binding of OLKG and xanthine to XanA was previously shown to result in
quenching of the endogenous fluorescence of the protein (4), thus allowing estimation
of the K., of the substrates. The apparent quenching of fluorescence by xanthine is
complicated by the significant absorption of the excitation wavelength by the
substrate, precluding the confident estimation of the K, of xanthine. In contrast, the
fluorescence quenching approach was extended to study the K, of both OLKG and Fe(II)
(neither of which absorbs at 280 nm) for the mutant proteins (Figure 4.3 and 4.4, and
Table 4.1). The three variants involving the putative ligands to the metallocenter
displayed significantly increases in the Kd of Fe(II), consistent with their metal
binding assignments. Similarly, the K122A variant exhibited an increased Kd of (xKG,
consistent with its suggested function in stabilizing the binding of the cosubstrate. Of
interest, the fluorescence of the H149A variant was not quenched by addition of OLKG
perhaps suggesting that it may an important role in protein folding.

Kinetic Comparison of XanA and its Variants with Xanthine. Although all
variants appeared to be equally overproduced, the Q99A, H149A, D151A, and
H340A variants displayed no activity in cell extracts. When purified, H340A XanA
exhibited trace activity (0.13 U mg") corresponding to 0.17 % of that associated with
wild-type enzyme. These four variants were not further analyzed kinetically. The
kinetic parameters for XanA and the seven active mutant proteins are listed in Table
4.2. All of these variants exhibit perturbations in their Km (xanthine), K.,, (OtKG), and

kca, compared to the control enzyme, consistent with the mutant proteins having

134

FIGURE 4.3: Fluorescence spectroscopic analysis of XanA interaction with Fe(II).
The changes in the endogenous fluorescence (excitation at 280 nm and emission at
335 nm) due to quenching by Fe(II) were examined in 50 mM Tris buffer, pH. 8.0,
solutions containing (A) 0.12 pM XanA, (B) 0.0] pM H149A, (C) 0.03 pM D151A,
(D) 0.04 pM H340A, (E) 0.05 pM Q101A, (F) 0.12 pM K122A, (G) 0.05 pM E137A,
(H) 0.07 pM D138A, (I) 0.02 pM Q356A, (J) 0.02 pM C357A, and (K) 0.07 pM

N358A at 25 °C. Fe(II) concentrations ranged from 0-35 pM.

 

 

 

 

 

 

 

 

 

 

 

 

 

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D151A, (C) 0.04 pM H340A, (D) 0.50 pM Q101A, (E) 0.12 pM K122A, (F) 0.13 pM

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TABLE 4.]: Determination of Kd of Fe(II) and OLKG for XanA and Selected Variantsa

 

 

Mutant Kd of Fe(II) (uM) Kd of aKG (11M)
XanA 85:03 115 :5
Q101A 21.5:12 108:12
K122A 9.3 : 0.6 236 : 29
E137A 11.7: 0.6 146: 13
D138A 12.3 :1.0 136: 14
H149A 36.8 :1.3 NDb
D151A 28.2: 1.4 169: 16
H340A 30.7 :14 175 :16
Q356A 15.9 : 0.6 108 : 6
C357A 13.5:09 124: 14
N358A 10.1 : 1.0 168 :9

 

a Estimated on the basis of quenching of the endogenous fluorescence with excitation
at 280 nm and emission at 335 nm at 25 °C. The protein concentrations: 0.12 M for
XanA, 0.01 M for H149A, 0.03 M for D151A, 0.04 uM for H340A, 0.05 uM for
Q101A in the Fe(II) assay and 0.5 uM in the aKG assay, 0.12 11M for K122A, 0.05
uM for E137A with Fe(II) and 0.13 uM in the (xKG assay, 0.07 11M for D138A in the
Fe(II) assay and 0.18 11M in the (XKG assay, 0.02 M for Q356A in the Fe(II) assay
and 0.05 11M in the OtKG assay, 0.02 M for C357A in the Fe(II) assay and 0.10 M
in the aKG. assay, and 0.07 uM for N358A in the Fe(II) assay and 0.1] uM in the

aKG assay. b ND, fluorescence quenching was not detected.

139

TABLE 4.2: Substrate, Cosubstrate, and Substrate Analog Kinetic Parameters of

XanA and Selected Variantsa

 

 

XanA Xanthineb (JtKGc 9-Methylxanthined
sample
Km kcat kcat/ Km Km kcat kcat/ Km Km kcat kcat/ Km
01M) 6") (.844 s") (11M)(s")01M" s") (mM) (s") (mM' s")
Wild-Type 45 :4 72:2 1.6 31 :2 67:1 2.16 0.40: 3.8: 9.5
0.07 0.3
0101A 122 : 9 36 :1 0.30 59: 6 21 :1 0.36 0.78: 0.66: 0.85
0.07 0.03
K122A 88:5 60:] 0.68 32:2 42:1 1.31 0.88: 1.9: 2.16
0.07 0.1
E137A 71:5 40:1 0.56 41:8 31:1 0.76 1.09: 5.9: 5.41
0.17 0.5
D138A 69 : 6 34 : 2 0.49 53 : 7 25 :1 0.47 0.58 : 4.25 : 7.33
0.09 0.24
Q356A 57 : 5 35 : 1 0.61 42 : 3 28 : 1 0.67 0.68 : 1.4 : 2.05
0.07 0.1
C357A 58 : 5 32 :1 0.55 47 : 4 19 :1 0.40 0.55 : 4.5 : 8.18
0.05 0.2
N358A 554:24 38:2 0.07 48:8 17:] 0.34 1.0: 1.26: 1.26
0.4 0.35

 

a Except for the compound being analyzed, the assay solutions contained 40 11M
Fe(II), 1 mM aKG, and 200 11M xanthine in 50 mM MOPS buffer, pH 7.4, and were
maintained at 25 °C. b Xanthine concentrations ranged from 0-250 11M, except for the
N358A variant, which ranged from 0-400 pM. c aKG concentrations ranged from

0-500 pM. d 9-methy1xanthine concentrations ranged from 0-1.0 mM.

140

modifications involving active site residues.

Alternative Substrates for XanA and its Variants. In agreement with prior results
(4), XanA was shown to be incapable of using 3-methylxanthine, 6-methylpurine, or
7-methylxanthine as the primary substrate. In contrast, trace levels of activity were
detected for l-methylxanthine (about 10'4 of that observed for xanthine) when using
highly concentrated enzyme solutions, and low levels of activity were observed with
9-methylxanthine. The absorbance spectra of the products arising from
l-methylxanthine and 9-methylxanthine exhibited features at 289 nm and 291 nm
(Figure 4.5), consistent with the formation of l-methyluric acid and 9-methy1uric acid
(9), respectively. Generation of the product from 9-methylxanthine was coupled to 02
consumption, as shown by use of an oxygen electrode, and to the decomposition of
0tKG, as analyzed by the OPDA assay (data not shown).

The kinetic parameters associated with XanA utilization of 9-methy1xanthine for
wild-type enzyme and the seven variants are shown in Table 2. For wild-type XanA,
the Km of 9-methy1xanthine is 8.9-fold larger than that for xanthine while the kw is
approximately 5 % of that for the true substrate. Comparison of the data obtained for
the control enzyme with those for the mutant proteins provides insight into potential
protein interactions involving the 9-methyl group. For example the E137A variant
exhibited a 1.6-fold increase in km for 9—methylxanthine versus the wild-type enzyme.
Although not statistically significant, 3 small increase (1.1-fold) in km also was noted
for the D138A variant. These results suggest that substituting Ala for the carboxylate

at position 137, and perhaps also at position 138, partially compensates for the

141

FIGURE 4.5: UV-visible spectra of l-methylxanthine, 9-methylxanthine, and their
products. (A) Spectrum of 9-methylxanthine in assay solution (solid) and the sample
after 30 min of reaction (dotted) at 25 0C. The assay solution included 40 11M Fe(II), 1
mM aKG, 3.6 uM XanA, and 200 uM 9-methylxanthine in 50 mM MOPS, pH 7.4. (B)
Spectrum of l-methylxanthine in assay solution (solid) and the sample after 100 min
of reaction at 25 °C (dotted), using conditions as above. (C) The spectrum of the
product arising from the enzymatic transformation of 1-methy1xanthine was
calculated from B (dotted line) by measuring the absorption at 290 nm (0.747
absorbance units), determining £290 for l-methylxanthine (1,497 M'I cm") and
l-methylun'c acid (10,273 M'l cm'l), using these results to estimate that the
concentrations of l-methylxanthine (149 0M) and l-methylun'c acid (51 11M) which
totaled to 200 11M, and subtracting the spectrum of 149 1.1M l-methylxanthine from
the spectrum of the mixture. This was compared to the spectrum of authentic

1-methyluric acid (solid line)

 

 

 

 

 

 

.3 om
‘

 

 

 

142

presence of a methyl group at N-9 of xanthine by allowing for more productive active
site binding.

6,8-DHP Inhibition. 6,8-DHP was previously shown to inhibit wild-type XanA
(4), but its mechanism of inhibition Was not examined. By contrast only very modest
inhibition was noted with 2,8-DHP, and 8-hydroxypurine did not inhibit the enzyme
(4). These results indicated that XanA interactions with the C-6 oxygen were most
important for inhibitor binding.

To examine the inhibitory effect of 6,8-DHP on XanA, progress curves of uric
acid production were obtained under different inhibitor and substrate concentrations.
The curves obtained for a constant substrate concentration with varied levels of
inhibitor reveal a time-dependent inhibition of XanA by 6,8-DHP (Figure 4.6A)
whereby the initial rate constant, vi, decreases to a steady-state rate constant, vs, with
an apparent. first-order rate constant, kobs. The values of V., vs, and [robs were
characterized according to equation 2 by using a variety of assay conditions. For
varied inhibitor concentrations, kub, exhibited a hyperbolic dependence on the
concentration of 6,8-DHP (Figure 4.6B), consistent with a slow-binding inhibitory
mechanism involving the reversible binding of inhibitor followed by a conformational
change to create a tightly bound state (10). To distinguish the type of slow-binding
inhibition mechanism, progress curves were obtained for a constant concentration of
6,8-DHP in the presence of varied substrate concentration (Figure 4.6C). A replot of
kobs versus substrate concentration reveals a negative slope (Figure 4.6‘D) and

confirms the mechanism as being competitive (11), as depicted in Figure 4.7. Thus,

143

FIGURE 4.6: Interaction of XanA with 6,8-DHP. (A) Progress curves for XanA
inhibition by varied concentrations of 6,8-DHP. 2.4 nM XanA was added to 50 mM
MOPS buffer, pH 7.4, at 25 °C containing 40 11M Fe(II), 1 mM aKG, 200 11M
xanthine, and 0 mM (n), 0.02 mM (.), 0.04 mM (A), 0.06 mM (V), 0.1 mM (0) or
0.12 mM (4) 6,8-DHP. The production of uric acid was monitored at 294 nm and the
progress curves were fit to equation 1. (B) Dependence of kobs on the concentration of
6,8-DHP. The curve was fit to equation 2. (C) XanA progress curves were obtained as
above, but in the presence of 0.1 mM 6,8-DHP and 50 11M (4), 100 1.1M (V), 150
11M (A), 200 11M (0) or 250 11M (I) xanthine. The progress curves were fit to

equation I. (D) Dependence of kobs on the concentration of xanthine.

-" 0.00251 0

O
b
1.
x
1
'1
1
‘1
I
1
I
I

A

A 1

:1" ,x z ‘7 0.00201
‘ t

0

v

§ 0.0015{ B
.3 0.0010'

400 600 30° 100° 120° o'msno002004000000 030 03?
Time (sec) 5 B-DHP (mM)

0 e e e
d
4 - 1 A n - 1 -

Abs 294 nm

  

 

 

 

 

2.01110"-
2.71110“-
..2‘ 2.01110": .
3 2.5x10"«
0 .31
1: 2.41110 1 D
0 4
2 2.31110 .
2.2x1o"- '
“Twang,“ “°° 5'0 ' 160 ' 150 ' 200 ' 250 '
Xanthine (0M)

IOD‘A

 

 

 

 

144

FIGURE 4.7: General mechanism for slow-binding inhibition.

 

 

 

 

145

6,8-DHP competes with substrate to form an initial enzyme inhibitor complex (E-I)
that slowly transforms to a more stable complex (E-I‘). For such a mechanism, kobs
varies with the inhibitor concentration [I] according to equation 4 (10) , where K, =
k4/k3. This equation simplifies to equation 5 when one defines K?” = Ki(1 + [S]/Km).

kobs = k6 + k5[I]K./(1 + [SI/Km + [ll/K1) (4)
kobs = k6+ kill/(1’9“pp + [1]) (5)
Using the data from Figure 4.68 for wild-type XanA, k5 = 2.0 X 10'3 8", k6 = 8.1 x

104's", and Kiapp = 12.6 11M. k5 is larger than k6 suggesting the rate of formation of
B]. complex is much faster than its dissociation, which is in agreement with
slow-binding inhibition mechanism.

The slow-binding inhibition kinetic properties of the active variant forms of
XanA with 6,8-DHP were analyzed in a similar manner (see Figures 4.8-4.14) and the
k5, k6 and K-f'pp are compared to the control enzyme in Table 4.3. Compared with the
K?” of 12.6 11M calculated for wild-type XanA, the values for the E137A and D138A
proteins were smaller, ranging from 3.0-3.6 11M. This decrease in K?” is consistent
with elimination of negative interactions between 6,8-DHP (possibly involving the
deprotonated C-8 hydroxyl group that likely binds to Fe(II)) and the Glul37 or
Aspl38 carboxylates; i.e., the charge repulsion is eliminated in the two mutant
proteins containing the small and hydrophobic Ala side chains. Kf’pp increased
significantly for both the Q356A and N358A proteins, suggesting that stabilizing
interactions provided by the Gln356 and Asn358 side chains are abolished in these
mutants. In. particular, this result is consistent with hydrogen bonding between the C-6

hydroxyl group (or the corresponding keto tautomer) of 6,8-DHP and Gln356 and

146

FIGURE 4.8: Interaction of Q101A with 6,8-DHP. (A) Progress curves for Q101A
XanA inhibition by varied concentrations of 6,8-DHP. 1.9 uM Q101A was added to
50 mM MOPS buffer, pH 7.4, at 25 0C containing 40 11M Fe(II), 1 mM aKG 200 11M
xanthine, and 0 mM (I), 0.01 mM (0), 0.02 mM (A), 0.04 mM (V), 0.06 mM (4) or
0.08 mM (b) 6,8-DHP. The production of uric acid was monitored at 294 nm and the
progress curves were fit to equation 1. (B) Dependence of kobs on the concentration of
6,8-DHP. The curve was fit to equation 2. (C) XanA progress curves were obtained as
above, but in the presence of 0.1 mM 6,8-DHP and 50 1.1M (I), 100 11M (0), 150 0M
(A), 200 11M (V) or 250 11M (4) xanthine. The progress curves were fit to equation

1. (D) Dependence of kobs on the concentration of xanthine.

 

 

 

 

 

  

 

   

 

 

 

 

 

 

 

 

 

 

 

 

 

10« 0.004-
. ‘ ‘ -
0.04 0.0031
5 ~ .
v 0.0. ‘7
a ‘ 3 0002‘
g 0.40 g 1
‘ -K 0.W1<
< 0.2. A 1 B
00.-.-.-r. 0.000..-.-.r--
o 400 000 1200 0.00 0.02 0.04 0.00 0.00
“”9 (39°) 0,11-an (mM)
0.0« .
. 0.0000-
05 .
0.0000-
5 .- .
3 . 4‘ 0.0000.
3 . I a 1
N 30.“:
n 1
2 3 0.0040. .
0.0030. D
‘12'00' 00'100‘100'200'200'
Time (sec) Xanthine (pM)

147

FIGURE 4.9: Interaction of K122A with 6,8-DHP. (A) 6.1 uM K122A was added to
50 mM MOPS buffer, pH 7.4, at 25 °C containing 40 11M Fe(II), 1 mM OLKG, 200 11M
xanthine, and 0 mM (I), 0.01 mM (0), 0.02 mM (A), 0.04 mM (V), 0.06 mM (4),

0.08 mM (D) or 0.1 mM (0) 6,8-DHP. Panels B-D are the same as for Figure 83.

1.0-
0.31

1
0,51

 

0,41

Abs 294nm

0.2‘ 7 A

 

 

 

 

'0'400‘060'12100v '0.00'0.04'0.00'0.'12
W" 189°) 6,8-DHP (mM)

 

  

 

 

Abs 294 nm

C 1 D
o '400'000'12'00' 50*150'150'2fifi250'
Time (sec) Xanthine (pM)

   

 

 

 

148

FIGURE 4.10: Interaction of E137A with 6,8-DHP. (A) 0.27 uM E137A was added to
50 mM MOPS buffer, pH 7.4, at 25 °C containing 40 1.1M Fe(II), 1 mM aKG, 200 11M
xanthine, and 0 mM (I), 0.005 mM (0), 0.01 mM (A), 0.02 mM (V), 0.04 mM (4),
0.06 mM (b) or 0.08 mM (0) 6,8-DHP. 0.54 11M E137A was used for plot (C) and

(D). Panels B-D are the same as for Figure S3.

 

 

 

  

 

   

 

 

 

 

 

 

 

 

 

 

6.31
0.51
E 0.4.
3 0.3:
N .
3 0.2.
< 0.1:
0.0 fi.vA.-.r 0.000.:.-.-r~ffi
0 400 000 1200 0.00 0.02 004 0.00 0.00
Time (sec) 6,8-DHP (mM)
. 0.020-
00. J '
0.010-
2 ~ 1
F I
' '10 0.0124
g V
a 30000
.n .3 . .
< 0.0044 D ' -
0.0.-.-.fi.- .-.-.f.fi..
0 400 000 1200 00 100 150 200 200
Time (sec) Xanthine (an)

149

FIGURE 4.11: Interaction of D138A with 6,8-DHP. (A) 0.83 11M D138A was added
to 50 mM MOPS buffer, pH 7.4, at 25 °C containing 40 11M Fe(II), 1 mM OtKG, 200
11M xanthine, and 0 mM (I), 0.005 mM (0), 0.01 mM (A), 0.02 mM (V), 0.04 mM

(4), 0.06 mM (D) or 0.08 mM (0') 6,8-DHP. 1.6 11M D138A was used for plot (C)

and (D). Panels B-D are the same as for Figure S3.

 

 

 

 

 

   

 

 

 

 

 

 

 

 

 

 

0.0- :_.
0.003-
E 0.0<
1: . ,7: .
3 0,4. g 0.002:
N 10
1
.3 02 '3
< . .1: 0.001
0.0 . - A - -
f ' ' OlomI ' I r U V V v V ‘
° ‘°° '°° 12°° 0.00 0.02 0.04 0.00 0.00
71"“ ““1 0.0-011p (mM)
0.00301
0.0. '
S E 0.00324
0.4- c
3 3 1
N ‘ N
00 0.0020
.n 0.21 3
< , C < 1 D ' I
0.0: va v - w - 0.0024 1a.r.-.-+.
0 400 000 1200 so 100 150 200 250
Time (sec) Xanthine (till)

150

FIGURE 4.12: Interaction of Q356A with 6,8-DHP. (A) 53.5 nM Q356A was added to
50 mM MOPS buffer, pH 7.4, at 25 °C containing 40 1.1M Fe(II), 1 mM aKG, 200 1.1M
xanthine, and 0 mM (I), 0.1 mM (0), 0.2 mM (A), 0.3 mM (V), 0.4 mM (4) or 0.5

mM (b) 6,8-DHP. Panels B-D are the same as for Figure S3.

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

  

 

 

   

 

 

0.0-
A
g ‘7
3’, 0.41 3
N ‘ 3
3 0.21 g
“ A
0.0 v v v 1 . I - l T I V I j I V I fl I v
0 400 000 1200 0.0 0.1 0.2 0.3 0.4 0.5
Time (sec) 6,8-DHP (mM)
0-3‘ 0.0070.
0.00054 '
0.0 .
E p 0.00004 .
C In 1
v 0.4« v 0.0055-
G a .
" ‘5 0.0050- - _
3 0.2 4‘ .
< C 0.0045: D .
0.0..fifi...- 0.0040.-.-.-.-,-
0 400 000 1200 50 100 150 200 250
Xanthine (all) Xanthine (0'4)

151

FIGURE 4.13: Interaction of C357A with 6,8—DHP. (A) 2.3 11M C357A was added to
50 mM MOPS buffer, pH 7.4, at 25 °C containing 40 11M Fe(II), 1 mM OtKG, 200 11M

xanthine, and 0 mM (I), 0.02 mM (0), 0.06 mM (A), 0.10 mM (V), 0.14 mM (4) or

0.18 mM (b) 6,8-DHP. Panels B-D are the same as for Figure S3.

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

E
c
v
a:
N
(
B
_ .-.-.. °-°°°‘.-.-.-.+.-.
0 400 000 1200 0.00 0.04 0.00 0.12 0.10 0.20
Time (sec) 3.8-0"? ("1")
0.0040
E0.0 0.0030:
c ,1"
v 0.44 'g, 0.0032 '
0 v
N a '
£03 30.0020 '
.1:
. D .
0.0fifi—f.rfiv 0.0024 W.
‘2‘” 50 100 150 200 250
Time (39°) Xanthine (01111)

152

FIGURE 4.14: Interaction of N358A with 6,8-DHP. (A) 1.4 uM N358A was added to
50 mM MOPS buffer, pH 7.4, at 25 °C containing 40 uM Fe(II), 1 mM (XKG, 200 uM
xanthine, and 0 mM (I), 0.02 mM (0), 0.04 mM (A), 0.10 mM (V), 0.15 mM (4),
0.20 mM (D) or 0.30 mM (0) 6,8-DHP. 2.3 uM N358A was used for plot (C) and (D).

Panels B-D are the same as for Figure S3.

  
 
   

 

 

 

 

 

 

 

 

  

 

 

 

  

 

 

0.51 0.00324
0.4- 0.0030.
E 1 A J
c '7 00020
V0.31 . u ‘
OD “—- v 4
N
“03 3 0.0020‘
n o 4
<01 4‘ 0.0024.
°'° ' ' - T - . - 0.0022 - . - i f 1
° ‘°° °°° 12°° 0.0 0.1 0.2 0.3
Time (sec) 6,8-DHP (mM)
0.000
1.2- i I
‘ 0.m7- .
1,0. ‘
. A ‘
E 0.0. ,7 0.006
‘ ‘ am;
3 0.0: a . 4
°‘ .0
0.4. 00.0042 ,
1 * I D
‘M‘ C 0.0034 ' .
0.0-,-,+1_ 1-.er-....
0 400 000 1200 50 100 150 200 250
Time (sec) Xanthine 0,1”)

153

TABLE 4.3: Inhibition of Wild-Type XanA and its Variants by 6,8-DHPa

 

XanA sample

1‘5 (5")

hi§0

Kiapp (HM)

 

“manme

QHHA

K122A

E137A

D138A

Q3 56A

C357A

N358A

(ZOiQDx103
(l7iODx103
(17:0nxnfi
(18i00x103
(sciomx103
(58ionx103
(lliQDx103

(94irmxio‘

(&li0&x104
(54irnxlo‘
(20:0nxio‘
(25irnx104
(lliLDx104
(L2i0nx103
(Isionxiot

(23ionx103

12.6 i 2.4

9.8:t 1.7

lO.4:tl.l

3.] i0.4

3.6 :t 0.4

lOZiIZ

16.6 d: 2.0

81i39

 

a Kinetic analyses were carried out as described in the text, with the kinetic

parameters defined according to equation 5.

154

Asn358. For most of the mutants, k5 is larger than k6; however, N358A exhibited the
opposite result (k5 = 9.4 x 10'4 3", k6 = 2.3 x 10'3 s"). This finding is consistent with
the N358A enzyme lacking an important stabilizing hydrogen bond(s) to the C-6
hydroxyl group of 6,8-DHP, perhaps involved in the k5 transition and formation of the
E-I.complex (Figure 4.5).

Identification of a Reactive T hi0! at the XanA Active Site. The homology model
of XanA depicts Cys357 at the active site (Figure 4.1), a prediction that was directly
tested by chemical modification studies. As illustrated in Figure 4.15A, the incubation
of XanA with various concentrations of 5,5’-dithiobis (2-nitrobenzoic acid) (DTNB or
Ellman’s reagent, specific for reacting with thiol groups) led to concentration-
dependent, first-order losses of activity. Significantly, the added presence of xanthine
provided some protection to the enzyme (A symbols in Figure 4.15A), consistent with
a reaction between DTNB and a Cys residue at the active site. In contrast, OLKG had
much less of a protective effect when examined alone ([3 symbols) or with xanthine
(data not shown). Inactivation studies also were carried out with iodoacetamide
(Figure 4.15B), a less specific reagent to thiol groups than DTNB. Higher
concentrations of iodoacetamide were required to obtain XanA inactivation rates
compared to DTNB. Either xanthine or (xKG provided some protection against
enzyme inactivation by this reagent, and the combined presence of xanthine and aKG
provided increased protection against iodoacetamide. These results obtained with
iodoacetamide agree well with those using DTNB and are consistent with the

presence of a reactive Cys residue near the xanthine-binding site. When the DTNB

155

FIGURE 4.15: Inactivation of XanA by DTNB. (A) XanA (51 pg ml") was incubated
on ice with O (O), 2.5 uM (I), 125 uM (A), 750 11M (0) or 1.25 mM (0) DTNB in 50
mM MOPS buffer, pH 7.4, for the indicated times, then diluted lOO-fold into buffer
containing 40 11M Fe(II)and 1 mM aKG After blanking the spectrophotometer,
xanthine was added to 200 pM. Additional samples containing 750 uM DTNB were
examined with added 0.2 mM xanthine (--A--) or 1 mM aKG (--D--). The
combination of added xanthine plus OLKG was equivalent to added xanthine alone. (B)
XanA inactivation by iodoacetamide. The enzyme (51 pg ml") was incubated on ice
with O (O), 20 mM (A), 50 mM (I), 60 mM (0) and 100 mM (x) iodoacetamide in 50
mM MOPS buffer, pH 7.4, for the indicated times, then diluted lOO-fold into buffer
containing 40 uM Fe(II)and 1 mM aKG After blanking the spectrophotometer,
xanthine was added to 200 pM. Additional samples containing 60 mM iodoacetamide
were examined with added 0.2 mM xanthine (-—+--) or 0.2 mM xanthine plus 1 mM
OLKG (--D--). The inclusion of OtKG alone was equivalent to the case of added
xanthine. (C) C357A XanA (51 ug ml") was incubated on ice with 0 (O), 15 uM (I),
0.75 mM (A) and 1.25 uM (0) DTNB in 50 mM MOPS buffer, pH 7.4, for the
indicated times, and diluted and assayed as above. (D) Time—dependent inactivation of

wild-type XanA (I) and the C357A variant (O).

Time (min) Time (min)
0 5 10 15 20 25 0 5 10 15 20 25

 

     

 

 

 

 

 

 

 

 

. M {R . '
I E
43.02 ,, A O E 0.0121
g’ o I A " ‘ '
5 .0“ _ . . g 0.000- _
32 I: .
3: '0-03 f C 0 12 0.00M .
- .000 - g ‘ 2 . o
5 0.000.
.01 - 03‘032'03'03003113274

 

156

inactivation studies were repeated with the C357A variant enzyme, the reagent
yielded very little inactivation compared to the control enzyme (Figure 4.15C and D).
This result confirms the positioning of Cys357 at the XanA active site. During the
course of the above studies, XanA was found to be labile when examined at dilute
concentrations (e.g., 12 nM or 0.51 pg mL"), such as those used in routine enzyme
assays, whereas the concentrated XanA samples were quite stable during storage. For
example when highly diluted XanA was incubated at 25 °C in 50 mM MOPS buffer
(pH 7.4), the activity decreased by ~70 % in about 6 min (Figure 4.16). Inclusion of
40 pM Fe(NH4)3(SO4)2 during this incubation stabilized the protein such that ~55 %
activity was retained over this time period. Addition of both 40 pM Fe(NH4)2(SO4)2
and 1 mM aKG further stabilized the protein, with ~80 % activity remaining after 6
min. HPLC measurements showed no detectable change of OLKG concentrations
during these incubations. The approximately first-order loss of activity in the presence
of 40 pM Fe(NH4)2(SO4)2 and 1 mM aKG was not affected by inclusion of 80 pM
ascorbate (data not shown); however, the protein was found to be more stable in the
absence of oxygen. The oxygen-dependent instability of XanA apoprotein hinted at
the reactivity of a cysteine thiol in the protein. Consistent with this hypothesis, the
activity of the Cys357 protein was quite stable when examined at dilute (12 nM)
concentrations (Figure 4.16). This result confirms that metal-independent oxidation of
Cys357 is associated with XanA inactivation; although the mechanism of this

inactivation process remains unclear.

157

FIGURE 4.16: Time-dependent loss of XanA activity for highly diluted enzyme.
Wild-type XanA was diluted to 0.5] pg ml'1 in aerobic buffer, 50 mM MOPS (pH 7.4),
without additives (V), with 40 pM Fe(II) (A), and with 40 pM Fe(II) plus 1 mM
aKG (0), or was diluted into anaerobic buffer (I). For comparison, the C357A variant
(0.51 pg ml") was examined in aerobic buffer (0). At the indicated time points, the
missing components were added and assays (25 °C) were initiated by adding 200 pM

xanthine.

 

 

i 4 6 ii i 1‘?
Time (min)

158

Oxygen Reactivity ananA in the Absence of Primary Substrate. The addition of
XanA to a buffered solution containing 40 pM Fe(II) led to a slow rate of 0;
consumption, similar to the background rate of consumption that occurred in the
absence of enzyme (data not shoWn). For the Q101A, Q356A, and C357A mutant
proteins, and to a lesser extent the E137A variant, a distinct pattern was observed with
15-20 pM 02 being consumed with a much greater initial rate constant, denoted v0 in
Table 4.4. This rapid phase of oxygen consumption also was observed for enzyme
samples added to solutions containing Fe(II) plus (XKG (Figure 4.17). The presence of
cosubstrate often led to an increase in V.,, which is especially noticeable for the K122A
and N358A variants (Table 4.4). Control experiments carried out with Q101A and
E137A forms of XanA confirmed that the amount of oxygen consumed in this rapid
process correlated with the concentration of added Fe(II), so that approximately twice
the amount of O; was consumed when using 100 pM Fe(II). The use of higher
concentrations of the metal ion in the assay allowed for a more reliable calculation of
v0 for the wild-type enzyme (Table 4.4) even though the background rate of oxygen
consumption also was increased.

We attribute the rapid phase of oxygen consumption in the above studies to
enzyme-catalyzed ferroxidase activity in which the exogenous metal ions are used to
provide electrons for the reduction of O; to hydrogen peroxide. Support for this
activity was obtained by using the ABTS assay for quantifying peroxide production.
To illustrate, the Q101A variant stimulated the production of hydrogen peroxide

significantly faster than the wild-type protein, which generated H203 more rapidly

159

FIGURE 4.17: Oxygen consumption studies of XanA and its variants. An oxygen
electrode was used to monitor 0; consumption at 25 °C in 50 mM MOPS buffer, pH 7.4,
containing 1 mM aKG and 40 pM Fe(II) (circles) or 100 pM Fe(II) (triangle), and 1 pM
enzyme (wild-type, dark blue; Q101A, cyan; D138A, purple; C357A, green). Enzyme-

free control samples (black symbols, no lines) were analyzed for comparison.

 

Oz-concentratlon (nM)

 

 

 

220

 

time (sec)

160

TABLE 4.4: Oxygen Reactivity of XanA and Selected Variants in the Absence of

Primary Substratea

 

 

Mutant Fe(II) only Fe(II) + OLKG
v0 (11M s") vo (11M 8“)
. 0.030 d: 0.002
W - . i .
ild type 0 023 0 0001 (0.136 i 0.002)
0.46 i 0.01
.3 :1: .
0101A 0 8 0 008 (0.240 d: 0.002)
K122A 0.05] i 0.001 0.143 i 0.001
0.192 :1: 0.018
3 . i .
El 7A 0 17 O 03 (0.264 i 0.004)
D138A 0.061 i 0.003 0.097 3: 0.002
Q356A 0.39 at 0.01 0.48 :t 0.03
C357A 0.70 i 0.03 0.73 :t 0.01
N358A 0.064 i 0.001 0.159 i 0.002

 

8 These representative data were obtained by using an oxygen electrode to measure
the effects of adding enzyme samples (final concentrations of 1 pM) to solutions
containing 50 mM MOPS buffer, pH 7.4, and 40 pM Fe (or 100 pM Fe for the values
shown in parentheses), or the same plus 1 mM OLKG. The data shown for each sample
are derived from replicate measurements obtained on the same day. The immediate
response (V.,) is attributed to ferroxidase activity with electrons from excess Fe(II)

used to reduce oxygen.

16]

than the non-protein controls (Figure 4.18) when assayed in the presence of 120 pM
Fe(II) and 1 mM aKG,. The rates of H202 production correlated with the rates of
oxygen consumption for the mutant examined. The H2O2 produced in these reactions
accounted for 48-63 % of the expected levels of this product (with the low
stoichiometry likely due to the known Fe(III) inhibition of peroxidase). The increased
levels of ferroxidase activity observed in several of the mutants compared to the
wild-type enzyme could reasonably relate to increased accessibility of exogenous
Fe(II) to the enzyme metallocenter or to altered binding of (XKG resulting in enhanced

metal center reactivity.

162

FIGURE 4.18: Hydrogen peroxide production by Q101A and wild—type XanA. The
wild-type (O) and mutant (I) XanA samples (1 pM) were incubated with 120 pM
Fe(II) and 1 mM (xKG in 50 mM MOPS buffer, pH 7.4, at room temperature. At
selected time intervals, aliquots Were mixed with ABTS assay solution (pH 5.0)
containing horseradish peroxidase and the absorbance was monitored at 420 nm. A

non-protein control sample (A) was examined for comparison.

2.5-
1 . I I I
2.0- . °
. O
A ‘ O A
2 1.5. ‘
:3. A
U ‘ .
N 10.
O
N ' A
I 0.5-
0.0 - - 1 - . - 1 - .

 

 

O
OH}
..I
O
..5
0|
N
O
N
0'

Time (min)

163

DISCUSSION

To identify active site residues and better define the reactivity of XanA, a
recently identified Fe(II)/aKG-dependent xanthine hydroxylase (4), we extensively
characterized a suite of enzyme Variants (Q99A, Q101A, K122A, E137A, D138A,
H 149A, D151A, H340A, Q3 56A, C357A, and N358A variants). The residues targeted
for mutagenesis, suspected metal ligands or potentially capable of hydrogen-bonding
interactions with substrate, were selected on the basis of a homology model of the
enzyme (4) that depicts them binding Fe(II) lining the active site pocket (Figure 4.1).
Significantly, these residues are well conserved in sequences of XanA orthologues (4).
For example, Gln101, Hisl49, Asp151, His340, Gln356, Cys357, and Asn358 are
universally conserved in the available XanA sequences. Gln99 often is present, but
Val occupies this position in some representatives. Lysl22, which was suggested to
stabilize interactions with the C-1 carboxylate of OLKG, either is- retained or
conservatively replaced by Arg or Thr. Although Asp, Gln, and Ser replace Glul37 in
other XanA orthologues, these replacement residues are capable of similar hydrogen
bond interactions. More flexibility exists for Asp138 which is replaced by Glu, Gln,
His, and Lys in other XanA sequences. Aromatic residues are not predicted to lie near
the active site, reducing the possibility of 1H: stacking interactions with the base,
though other hydrophobic residues are suggested to occur in this region. In addition to
these various side chains, the substrate might also interact with unidentified backbone
amide and carbonyl groups.

Model ofXanthine Binding Based on Kinetic Analyses of XanA and its Variants.

I64

The results from our extensive kinetic analyses of wild type and variant XanA
enzymes with substrates and inhibitors were combined along with determination of
the Kd of Fe(II) and OLKG along with the ferroxidase activities into a model for
xanthine binding to XanA, as depicted in Figure 4.19 (with substrate entering the
active site from the right and some hydrogen bonds likely to arise from interactions
with backbone amide groups). In this figure, the Fe(II) is shown bound by the Hisl49,
Asp151, and His340 side chains, as is almost certain from sequence comparisons to
other related enzymes (4). Mutations involving these residues abolished or greatly
diminished activity and led to an increase in the Kd of Fe(II). Furthermore, OLKG is
illustrated as chelating the metal, in agreement with other members of this enzyme
family, with stabilization provided by the appropriately positioned Arg352 (supported
by sequence comparisons) and Ly5122. Consistent with this role for Ly5122, the
K122A variant displayed an increase in the Kd of OLKG The primary substrate is
shown binding to the enzyme active site via a constellation of hydrogen bonding
interactions (some of which cannot be identified from our analyses), so that the
disruption of any one hydrogen bond will lead to only modest effects on the kinetics.
The Km and kw parameters measured by steady-state kinetics for xanthine utilization
by the wild type and variant enzymes cannot be used to directly infer which residues
bind the substrate; however, this baseline information allows for such inferences to be
made when combined with the data obtained from studies using 9-methylxanthine and
6,8-DHP. In addition, the ferroxidase measurements of the variants provide added

insight about putative active site residues. The suspected positioning and role of each

165

FIGURE 4.19: Proposed xanthine binding on the active site.

CYS357
, . . H
ASH358‘ ' \
O\~ 9 ASP151 i. Gln356
\ 0 7
. |‘ ~ ‘ ‘ ‘ N ' a ’ '
H6340: Ffi LYS122 </ I \ N
-0 His149 /N N O. ‘ "101
' ‘ 7
: a J.
Afgész’ ' Asp138 ',
G'U137 ?

I66

residue is described below.

The N358A variant displayed the largest reduction in kcm/Km of all mutants
retaining significant activity, consistent with this side chain having a critical role in
catalysis. This variant also gave riSe to one of the largest effects when assessing K?”
of 6,8-DHP inhibition, a result that can be interpreted as arising from the loss of a
critical interaction involving the C-6 hydroxyl group of the inhibitor. Together, these
results lead us to the depiction of Asn358 interacting with both the N-7 and the C-6
enol oxygen of xanthine (Figure 4.19). This mode of interaction with the substrate is
compatible with the homology model (Figure 4.1) that predicts Asn358 is positioned
deep in the active site near the metallocenter.

Gln356 also is shown interacting with the C—6 hydroxyl group of the substrate
(Figure 4.19). Support for this interaction derives primarily from the significant
increase in K?” of 6,8-DHP for the Q3 56A variant. The homology model predicts that
this residue is positioned near Asn358 in the active site (Figure 4.1). Of interest, the
Q356A variant exhibits enhanced ferroxidase activity that could be explained by
increased access of exogenous Fe(II) to the metallocenter when this side chain is
absent, in agreement with the model and Figure 4.19.

Cys357, located between Gln356 and Asn358, is present at the XanA active site
as shown by the results of chemical modification studies-especially the finding that
binding of substrate protects this residue from the thiol-specific reagents. The C357A
variant was not susceptible to inactivation by these reagents. Further support for this

residue being located near the XanA active site is derived from the increased

167

ferroxidase activity of the C357A variant, attributable to enhanced access to the bound
metallocenter by exogenous Fe(II) when this thiol group is removed or to perturbation
of the reactivity of the metal site. No specific interactions between Cys357 and
substrate were identified experimentally (Figure 4.19), but removal of this side chain
resulted in an increase in km for 9-methylxanthine compared to the wild-type enzyme
indicating a possible improvement in substrate orientation during catalysis. The
homology model (Figure 4.1) predicts the thiol group does not point into the active
site where xanthine binds, but small changes in torsion angles could reposition this
side chain to allow for interaction with substrate. Significantly, changes in the kinetic
properties of the C357A variant may be partially attributed to positional alterations of
the adjacent Gln356 andAsn358 side chains.

Both Glu] 37 and Asp138 are proposed to accept hydrogen bonds from the proton
at N-9 of xanthine .(Figure 4.19). This suggestion is based primarily on the
comparison of the kinetic properties for the wild-type protein versus the E137A and
D138A variants when using xanthine and 9-methylxanthine. Notably, these mutant
proteins exhibit greater kca. than the control protein with this alternate substrate as if
the absence of these \side chains provides more optimized interactions with the
substrate. Likely related to this finding, the E137A and D138A variants exhibit
decreased Kiapp for 6,8-DHP compared to the wild-type protein. We attribute these
results to a decreased repulsion of the deprotonated C-8 hydroxy group of the
inhibitor (the form that is most likely to coordinate the metallocenter) by the variants,

which lack the carboxylate groups. The homology model predicts that Asp138 lies

168

very close to the bound Fe(II), with Glul37 a short distance away (Figure 4.1). These
residues are located on the opposite wall of the active site compared to the Gln356,
Cys357, and Asn358 residues, consistent with the depiction in Figure 4.19.

The substitution of Gln101 by Ala leads to small effects on the substrate and
inhibitor kinetic parameters of XanA, but a significant increase in ferroxidase activity.
While we cannot suggest a specific site(s) of interaction between Gln101 and xanthine,
we speculate (in Figure 4.19) that this residue restricts access to the active site by
exogenous Fe(II). Compatible with this proposal, Gln101 is positioned near the active
site entrance in the homology model (Figure 4.1).

Although the Kd of OLKG is increased in the K122A variant, consistent with the
earlier prediction that Ly5122 interacts with the C-1 carboxylate of (XKG (4), the
substitution of this side chain by Ala had little effect on the kinetic parameters of the
enzyme. Nevertheless, when compared to the very small ferroxidase activity of
control enzyme the K122A variant exhibits a pronounced increase in ferroxidase
activity in the presence of Fe(II) plus (xKG over that of metal alone, suggesting that
Ly5122 is located near the OLKG binding site and serves to minimize this aberrant
activity.

Lack of Binding by Related Compounds. The proposed binding interactions for
xanthine illustrated in Figure 4.19 are consistent with the exquisite specificity of
XanA. Xanthine derivatives methylated at N-l, N-3, or the C-6 hydroxyl group are
very poor or not substrates (4) because of steric clashes or the interruption of critical

interactions. Although hypoxanthine (lacking the C-2 oxygen compared to xanthine)

I69

might appear to closely mimic the normal substrate, this mono-hydroxylated
compound will have too little resonance energy to overcome the Iactam intrinsic
stability so hypoxanthine will predominate as the C-6 keto tautomer (4), thus
disrupting several features important to binding. Xanthine derivatives hydroxylated at
C-8 (such as 6,8-DHP) cannot serve as substrates for the enzyme, but the hydroxyl
group at C-8 could reasonably function as a metal ligand and contribute to inhibition.
It is clear, however, that 8-hydroxypurine lacks many interactions proposed to be
critical for binding xanthine, explaining why it is not an inhibitor of the enzyme (4).
2,8-DHP is a somewhat better inhibitor, but it still lacks several hydrogen bond
interactions likely to be present with xanthine and exhibits only weak binding. By
contrast, the model correctly predicts that 6,8-DHP should be a potent inhibitor due to
the interactions involving its N-l, C—6 hydroxyl group, N-7, and N-9 proton, along
with metal coordination by the C-8 hydroxyl group.

The product of the xanthine/aKG dioxygenase reaction, uric acid, does not
significantly inhibit the enzyme despite its close structural resemblance to the
substrate. A critical difference between xanthine and uric acid is the distinction in pK,|
of the compounds. Whereas the pKa of xanthine is 7.4 (12), that of uric acid is 5.8
with the singly ionized species identified as the deprotonated N-3 and the dianion
shown to be the N-3 plus N-9 deprotonated species (13). Thus, in solution at neutral
pH, uric acid is predominantly a species in which the C-2 enol is deprotonated while
C-6 and OS are keto groups. The binding mode of substrate shown in Figure 4.19 is

inconsistent with these interactions, perhaps leading to ejection of the product from

l70

the active site.

To better understand the weak interaction of uric acid with XanA it is also
instructive to examine the structure of a protein that specifically accepts uric acid as a
substrate. Uric acid oxidase has been crystallized in the presence of several substrate
analogues, including 9-methyluric acid (14). In that case, the compound binds as the
2,6,8-tri-keto species where both N-3 and N-7 are deprotonated. Specifically, a Gln
OEl hydrogen bonds to the N-l proton, the C-2 keto oxygen hydrogen bonds to a
backbone amide as well as to an Arg residue, the same Arg is bound by the
deprotonated N-3, the C-6 keto oxygen binds a Gln EN2, the deprotonated N-7 binds
to a backbone amide, and the C-8 keto group binds to another backbone amide. If the
uric acid formed in XanA were to adopt this structure, many of the postulated
hydrogen bond interactions suggested for xanthine would be disrupted. In addition,
the C-8 keto oxygen would likely exhibit repulsion by the nearby AspISI. Thus, one
can readily rationalize the dissociation of the product from XanA.

Comparison to Xanthine Binding in Other Systems. Although the structural
details differ greatly, the general features of the proposed mode of xanthine binding in
XanA resemble the suggested interactions of xanthine with xanthine hydroxylase (15,
16) and the fungal xanthine/uric acid transporter (17, 18). Similar to the proposed
Gln356 and Asn358 hydrogen bonds to the xanthine C-6 enol proton, a Glu binds to
the xanthine C-6 hydroxyl group in xanthine hydroxylase. In addition, the
uncharacterized stabilization of the xanthine C-2 keto group in XanA is reminiscent of

the C-2 keto group binding to an Arg residue in xanthine hydroxylase. Furthermore,

17]

the potentially bridging interactions of XanA Asn358, between the N-7 and C-6
hydroxyl group of xanthine resemble the xanthine hydroxylase Glu interaction with
both the N-3 and N-9 protons. No structure is available for any xanthine/uric acid
transporter, but mutagenesis studies (17, 18) suggest that a particular Gln residue
binds the deprotonated xanthine N-9 nitrogen, much like the model generated here.
Final determination of XanA interactions with substrate must await elucidation of the

crystal structure in the presence of substrate or inhibitor.

I72

REFERENCES

(I)

(2)

(3)

(4)

(5)

(6)

(7)

(8)

(9)

(10)

Hille, R. (2005) Molybdenum-containing hydroxylases. Arch. Biochem.
Biophys. 433, 107-116.

Darlington, A. J., and Scazzocchio, C. (1968) Evidence for an alternative
pathway of xanthine oxidation in Aspergillus nidulans. Biochim. Biophys. Acta
166, 557—568.

Cultrone, A., Scazzocchio, C., Rochet, M., Montero—Moran, G, Drevet, C.,
and Femandez-Martin, R. (2005) Convergent evolution of hydroxylation
mechanisms in the fungal kingdoms: molybdenum cofactor-independent

hydroxylation of xanthine via a-ketoglutarate dependent dioxygenase. Molec.
Microbiol. 57, 276-290.

Montero-Morén, G M., Li, M., Rendon-Huerta, E., Jourdan, F., Lowe, D. J.,
Stumpff—Kane, A. W., Feig, M., Scazzocchio, C., and Hausinger, R. P. (2007)
Purification and characterization of the Fe(II)- and a-ketoglutarate-dependent
xanthine hydroxylase from Aspergillus nidulans. Biochemistry 46, 5293-5304.

Eichhom, E., van der Ploeg, J. R., Kertesz, M. A., and Leisinger, T. (1997)
Characterization of a-ketoglutarate-dependent taurine dioxygenase from
Escherichia coli. J. Biol. Chem. 272, 23031-23036.

Hausinger, R. P. (2004) Fe(ll)/0t-ketoglutarate-dependent hydroxylases and
related enzymes. Crit. Rev. Biochem. Mol. Biol. 39, 21-68.

Clifion, I. J., McDonough, M. A., Ehrismann, D., Kershaw, N. J., Granatino,
N., and Schofield, C. J. (2006) Structural studies on 2-oxoglutarate

oxygenases and related double-stranded B-helix fold protein. J. Inorg.
Biochem. 100, 644—669.

Purpero, V., and Moran, G R. (2007) The diverse and pervasive chemistries of
the OL-keto acid dependent enzymes. J. Biol. Inorg. Chem. (On-line).

H.S.Forrest, D. H., and J.M. Lagowski. (1961) Uric acid riboside. Part 1.
Isolation and reinvestigation of the sturcture. J. Chem. Soc, 963-968.

Morrison, J. F., and Walsh, C. T. (1988) The behavior and significance of

slow-binding enzyme inhibitors. Adv. Enzymol. Rel. Areas Mol. Biol. 61,
201-301.

I73

(11)

(12)

(13)

(14)

(15)

(16)

(17)

(18)

Ha, T. J., and Kubo, I. (2007) Slow-binding inhibition of soybean
lipoxygenase—l by dodecyl gallate. J. Agric. Food. Chem. 55, 446-51.

Dawson, R. M. C., Elliott, D. C., Elliott, W. H., and Jones, K. M. (1986) Data
for Biochemical Research, Third ed., Clarendon Press, Oxford.

Kahn, K., Serfozo, P., and Tipton, P. A. (1997) Identification of the true
product of the urate oxidase reaction. J. Am. Chem. Soc. 119, 5435-5442.

Retailleau, P., Colloc'h, N., Vivarés, D., Bonneté, F., Castro, E., El Hajji, M.,
Momon, J.-P., Monard, G, and Prangé, T. (2004) Complexed and ligand-free
high-resolution structures of urate oxidase (on) from Aspergillus flavus: a
reassignment of the active-site binding mode. Acta Crystallogr D60, 453-462.

Okamoto, K., Matsumoto, K., Hille, R., Eger, B. T., Pai, E. F., and Nishino, T.
(2004) The crystal structure of xanthine oxidoreductase during catalysis:

implications for reaction mechanism and enzyme inhibition. Proc. Natl. Acad.
Sci. USA 101, 7931-7936.

Truglio, J. J., Theis, K., Leimkiihler, S., Rappa, R., Rajagopalan, K. V., and
Kisker, C. (2002) Crystal structure of the active and alloxanthine-inhibited
forms of xanthine dehydrogenase from Rhodobacter capsulatus. Structure 10,
115-125.

Goudela, S., Karatza, P., Koukaki, M., Frilingos, S., and Diallinas, G (2005)
Comparative substrate recognition by bacterial and fungal purine transporters
of the NAT/NCSZ family. Molec. Membrane Biol. 22, 263-275.

Koukaki, M., Vlanti, A., Goudela, S., Pantazopoulou, A., Gioule, H.,
Toumaviti, S., and Diallinas, G (2005) The nucleobase-ascorbate transporter
(NAT) signature motif in UapA defines the function of the purine translocation
pathway. J. Mol. Biol. 350, 499-513.

I74

 

CHAPTER 5

COORDINATION CHEMISTRY AT THE FE(II) SITE OF
TAURINE/a-KETOGLUTARATE DIOXYGENASE AND ASPERGILL US

NIDULANS XANTHIN E HYDROXYLASE BY VARIOUS SPECTROSCOPIES

Dr. John McCracken assisted with EPR and ESEEM analyses and performed the
HYSCORE simulations; Efthalia Kalliri and Piotr Grzyska provided TauD and variant -

W248F; Dr. James Geiger simulated the structure of the NO-bound TauD active site.

175

 

ABSTRACT

Electronic spectroscopy and one- and two-dimensional electron spin echo
envelope modulation (ESEEM) experiments have been used to study the coordination
chemistry at the nitric oxide (NO)-bound non-heme Fe(II) sites of xanthine
hydroxylase (XanA) and taurine/a-ketoglutarate (OLKG) dioxygenase (TauD). NO,
which serves as a surrogate for molecular oxygen, binds to the metallocenter and
generates electronic transitions that were used to establish binding constants. The
diatomic gas spin-couples to the integer spin Fe(II) to yield an S = 3/2 paramagnetic
center with an axial EPR spectrum characterized by g2=4.00 and g“ = 2.00.
One-dimensional ESEEM spectra were taken across the EPR lineshape and show
modulations from 14N and 1H. At g = 4 the contributions from these coupled nuclei
overlap, making it necessary to use the two dimensional, 4-pulse hyperfine sublevel
correlation spectroscopy (HYSCORE) method to resolve contributions from bound
histidine nitrogens, coordinated H20, and ambient H2O. For the extensively
characterized enzyme TauD, HYSCORE spectra collected for samples in aqueous
buffer and 60% 2H20-buffer show changes in H20 and histidyl coordination as 0tKG
and taurine are added to the enzyme. Prior to co-substrate addition, HYSCORE
spectra show a substantial distribution of exchangeable, IH hyperfine couplings.
When OLKG is added, the lH HYSCORE is considerably altered with the dominant
hyperfine coupling arising from an exchangeable, strongly coupled proton of rhombic
symmetry. Subsequent addition of substrate taurine to yield the ternary complex

(Fe(II)-NO/aKG/taurine) at the active site showed a new lH hyperfine interaction that

I76

is not exchangeable in 2H2O. The HYSCORE cross-peaks from this lH show a
hyperfine tensor of axial symmetry characterized by a dipole-dipole distance of 3.2 A ,
and an isotropic contribution of 0.75 ea 0.3 MHz. X-ray crystal structure of TauD
shows C7 of Trp248 lies around 14.5 A to Fe(II), however, HYSCORE spectrum of
W248F did not exhibit substantial difference than that of TauD. Analogous studies
were carried out with the less well characterized enzyme XanA, providing the first
spectroscopic information for this enzyme and yielding insights into the coordination

properties of Fe(II) in this protein.

177

 

INTRODUCTION

The Fe(II)— and a-ketoglutarate (aKG)—dependent dioxygenases encompass a
wide range of enzymes with diverse primary substrates (reviewed in (1-3)). To
illustrate the versatility of this enzyme family, the prolyl, lysyl, and
aspartyl(asparaginyl) hydroxylases catalyze posttranslational modifications of
proteins (4), in some cases associated with hypoxic signaling (5). ijC
domain-containing proteins catalyze methylated-histone demethylation reactions (6).
AlkB repairs l-methyl-A or 3-methyl-C lesions in DNA (or RNA) by using an
analogous oxidative dealkylation reaction (7, 8). The lipid-metabolizing enzyme
phytanoyl-CoA hydroxylase participates in the metabolism of phytanic acid, and
deficiency of this enzyme leads to Refsum disease (9). Finally, a wide variety of small
molecules are synthesized or decomposed by members of this enzyme family. For
example, plants synthesize gibberellins, flavonoids, and some alkaloids by action of
these enzymes (10), thymine 7-hydroxylase sequentially hydroxylates the methyl
group of free thymine (11), and deA, deA, and SdpA decompose specific
phenoxyalkanoic acid herbicides by hydroxylation of their side chains (12, 13). The
studies described here focus on two representative hydroxylases: TauD and XanA.

TauD or taurine/(xKG dioxygenase, which catalyzes the reaction depicted in
Figure 1.8., is the best-characterized member of this group of enzymes. The
Escherichia coli enzyme is induced under conditions of sulfur starvation (14) and
allows the cells to use taurine, a widely available sulfonate (15), as a sulfur source.

The crystal structure of TauD has been reported (16, 17) and reveals a typical

I78

2-His-l-carboxylate Fe(II)-binding motif of protein side chains (18), chelation of the
metal by OLKG, and the binding of primary substrate near to, but not directly
coordinating, the metallocenter (Figure 5.1). The general mechanism of this enzyme
(Figure 5.2) follows that proposed for another enzyme in this class approximately 25
years ago by Hanauske-Abel and Giinzler (19). In the absence of the co-substrates (A),
three side chain ligands and three water molecules bind the hexacoordinate Fe(II).
The chelation of OLKG to Fe(II) through the C-1 carboxylate and C-2 carbonyl
moieties (B) displaces two of the waters, but leaves the final water bound resulting in
low reactivity with oxygen (20). The addition of taurine (C) triggers release of the
remaining water and creates a site for O2 binding (D) that stimulates further oxidative
chemistry. A critical Fe(IV)-oxo intermediate (E) was identified in this enzyme by a
combination of stopped-flow ultraviolet—visible, continuous-flow resonance Raman,
freeze-quench Mossbauer and extended x-ray absorbance fine structure approaches
(21-2 7). This intermediate abstracts a hydrogen atom from substrate (F) and hydroxyl
rebound leads to the product. Nitric oxide (NO) can be used as a surrogate for O2, and
the NO-bound form of this archetype enzyme has served as the paradigm for applying
electron spin-echo envelop modulation (ESEEM) spectroscopy to the examination of
substrate binding by use of deuterium labeled substrate (28).

XanA or xanthine/aKG dioxygenase is a recently identified member of the
Fe(II)/OLKG-dependent hydroxylases (29). This enzyme carries out the reaction shown
in Figure 2.1, in which OLKG decomposition is coupled to xanthine hydroxylation to

form uric acid. XanA is found only in selected fungi (29) and its chemistry contrasts

I79

FIGURE 5.1: TauD active site. The Fe(II) is bound to the His99, Asp101, and Hi5255
side chains and is chelated by OLKG. The substrate taurine binds near, but does not

directly coordinate, the metal center.

E, taurine

 

I80

FIGURE 5.2: General mechanism of TauD.

on2

G'UIA‘ph,_ F‘I", LT‘ Gln/ASPI,,‘_F
'0 ° Burs/H: /"‘ His
0

coo 2 H20

H20

. M- O. T
‘\ ..uIH O OG'UIASPI:,,F 9‘2 ,0. o
A Glu/Asp:-.Fell- 2 Glu/Aspl“, "F ' i v'
His’ | \Hzo ’FG/f‘o’ D
HIS Hil‘ HISO His
000'
°JKLCWR W,
C02
R-H

O
Glu/Asp”. ,,,,, 'e'v O

GIU’ASDMMH "I
HI:’ /F/ 0 His” FI 0 E
HIS ' HIS .

COO“ coo-

 

 

 

I81

with the well-characterized metabolism of xanthine by the molybdopterin
cofactor-containing enzymes that hydroxylate this substrate while transferring
electrons to NAD (xanthine dehydrogenase) or oxygen (xanthine oxidase) (30). The
recombinant Hisb-tagged form of XanA from Aspergillus nidulans was purified from
both its natural host and E. coli and extensively characterized (31, 32). The general
protein properties differ for samples from the two sources, persumably due to the
suspected lysosomal localization of the fungus-derived enzyme. Nevertheless, the
enzymatic properties of the two forms of XanA are very similar and the general
mechanism of this enzyme is suggested to parallel that of TauD (Figure 5.2). XanA is
related in sequence to TauD (18% identity) and the latter structure was used to create
a homology model of XanA (31); however, no crystal structure or spectroscopic
studies of XanA have been reported.

Here, the metal ion coordination properties of the NO-bound forms of TauD
and XanA are examined by electronic and one- and two-dimensional ESEEM
spectroscopies. The spectroscopic results of the TauD model system are analyzed in
terms of the known structure of this protein, and spectroscopic studies of a
site-directed variant provide additional insights in the Fe(II) coordination environment.
This spectroscopic approach is then applied to the XanA system to provide new

structural insights into the active site of this poorly characterized system.

182

EXPERIMENTAL PROCEDURES

Materials. 2H2O was purchased from Cambridge Isotope Laboratories, Inc.
(Andover, MA). Diethylammonium(Z)-l -(N,N-diethylamino)diazen- l -ium-l ,2-diolate
(DEA/NO) was purchased from Cayman Chemical (Ann Arbor, MI). An ammonium
salt of taurine deuterated at C] (the carbon adjacent to the sulfonate group) was a gift
from J. C. Price and J. M. Bollinger, and prepared as described (22). N-oxalylglycine
(NOG) was a kind gift from N. Burzlaff. Other general chemicals were purchased
from Sigma-Aldrich. 2H-Iabeled (xKG was prepared by 24h incubation of OLKG in
2H2O at room temperature.

Enzyme Purification. TauD, XanA, and the W248F variant of TauD were purified
as their His6-tagged forms, as previously described (26). Concentration of the samples
made use of Centriprep (Amicon Corp.) units with YM-lO membranes.

Analysis of Fe(II), (KC, and Substrate Binding to TauD and XanA by Using
Electronic Spectroscopy. All stock solutions for UV—visible studies were prepared
inside serum vials sealed with butyl rubber stoppers. Stock solutions of aKG (30
mM), taurine (100 mM), and xanthine (100 mM) were prepared in 50 mM Tris buffer
(pH 8.0), and made anaerobic by 10 rounds of vacuum degassing and flushing with
argon by using a vacuum manifold. Ferrous ammonium sulfate stock solutions (6 mM)
were prepared by 10 rounds of degassing and flushing with argon inside sealed serum
vials. Following the procedures reported for TauD (20), XanA (0.45 mM, in 50 mM
Tris, pH 8.0) was made anaerobic by 10 rounds of degassing and flushing with argon

in a 1 cm path length, 300-pl quartz cuvette fitted with a rubber stopper. After

183

blanking against the protein solution, spectra were recorded for samples to which
anaerobic aliquots of F e(II), OtKG, or xanthine had been added.

Electronic Spectroscopy of NO-treated Samples. 10 mg DEA/NO was dissolved
in 2 ml, 10 mM NaOH in tightly capped brown glass vials degassed as described
above. The DEA/NO stock solutions were made anaerobic by 5 rounds of degassing
and flushing with argon and stored at -80 °C. The concentrations were determined
from product information (8250 nm= 6,500 M"°cm") provided by Cayman chemical.
Using a 50 pl syringe, the indicated concentrations of DEA/NO were added by to
anaerobic protein samples in 300 pl quartz cuvettes fitted with rubber stoppers, and
electronic spectra (300 nm to 800 nm) were recorded on a Shimadzu UV-2401PC
spectrometer at 2 min intervals at 25 °C. Equation 1 was used for calculation of the
dissociation constant, Kd, where the concentration of free NO is equal to the
concentration of total NO added minus the concentration of enzyme—NO complex.

Fraction of maximal NO-bound TauD complex = [NO]/(Kd + [NO]) (1)

Electron Paramagnetic Resonance (EPR) Analyses. The samples from the quartz
cuvettes described above were transferred to degassed EPR tubes by using a 1 ml
syringe after incubation for approximately 35 min in the case of TauD or 50 min for
XanA. The EPR samples were frozen and stored under liquid N2. Additional protein
samples were prepared by diluting the concentrated proteins into the desired volumes
of 50 mM Tris, pH 8.0, prepared in 60% 2H2O.

Continuous wave-X band EPR spectra were recorded at 5 K on a Bruker

ESP300E spectrometer equipped with an Oxford liquid He cryostat by using 100 KHz

I84

 

modulation, 1.99 mW microwave power, and 9.58 GHz microwave frequency. One-
and two-dimensional ESEEM spectra were recorded on a Bruker E-680X
spectrometer operating at X—band and equipped with a model ER4118—MD-X-5-Wl
probe that employs a 5 mm dielectric resonator. The sample temperature was
maintained at 4.2 K using an Oxford Instruments liquid helium flow system equipped
with a CF-935 cryostat and an ITC-503 temperature controller. ESEEM data were
collected using a three-pulse, stimulated echo sequence (90°-t-90°-T-90°) with ‘t = 120
ns; 90° microwave pulse widths of 16 ns (full width half maximum) and peak powers
of 200 W. A four-step phase cycling sequence, (+x, +x, +x), (-x, +x, +x), (+x, -x, +x),
and (-x, -x, +x), together with the appropriate addition and subtraction of the
integrated spin-echo intensities served to actively remove the contributions of
two-pulse echoes and baseline offsets from the data (33). An integration window of 48
ns was used to acquire spin-echo amplitudes, and data set lengths were 512 Points.
Two dimensional ESEEM, also called hyperfine sublevel correlation spectroscopy
(HYSCORE), data were collected using a four-pulse, stimulated echo sequence
(90°-T-90°-t.-180 °-t2 -90°) with t' = 120 ns, 90° microwave pulse widths of 16 ns and
200 W peak power; 180° microwave pulse widths of 28 ns and 200 W peak power;
starting t1 and t2, 40 ns; and time increment, 28 ns. One- and two- dimensional
ESEEM spectral processing involves removal of the ESEEM decay by subtraction of
an exponential or polynomial decay feature, created with a Hamming window and
Fourier. Transformation. The absolute value, or square root of the resulting power

spectrum, is displayed.

I85

H I’SCORE spectral simulations. HYSCORE spectral simulations were done in
the time domain using the analytical expressions for S = 1/2, I = 1/2 HYSCORE
modulation function developed by Gemperle, et al. together with the orientation
averaging scheme developed by Hoffman and coworkers (33, 34). The output of these
simulations was processed using the same scheme outlined above for the Bruker

spectrometer via a program written for MATLAB.

186

 

RESULTS

Binding of (ZKG and Xanthine to Fe(II)-XanA Monitored by Electronic
Spectroscopy. Anaerobic Fe(II)-bound TauD is known to form lilac-colored
chromophores when incubated With OLKG (Max 530 nm, 8530 140-250 M'l-cm") or
OLKG plus taurine (Max 520 nm, 8520 254 M'l'cm") (20, 26, 35), and these features
have been attributed to metal-to-ligand charge—transfer transitions of six-coordinate
and five-coordinate sites, respectively (36—38). For comparison, anaerobic solutions of
Fe(II)-XanA were examined in a similar manner (Figure 5.3). The addition of OLKG to
Fe(II)-XanA yielded a very broad absorption spectrum centered at 506 nm with 8506 of
145 M'l-cm". Subsequent binding of xanthine to OLKG-Fe(lI)-XanA yielded no
significant shift in the absorption maximum, but led to enhanced definition of features
at 470 and 580 nm while diminishing the overall signal intensity (8506 125 M" cm").
Unlike the case of TauD, where the addition of Fe(II) in the absence of co-substrates
yielded a weak absorbance feature at ~650 nm due to metal interaction with low
concentrations of an endogenous catechol arising from enzyme self-hydroxylation of
Tyr73 (35), no analogous chromphore was observed in this region for Fe(II)-XanA.

Electronic Spectroscopy of N0 complexes of T auD and XanA. The effects of
added NO on the UV-visible spectra of various complexes of Fe(II)-bound TauD and
XanA were examined, as shown for the substrate-OLKG-Fe(II)-protein spectra in
Figure 5.4. For each sample, NO binds to the Fe(II) site to form a yellow
chromophoric species denoted {FeNO}7 in the nomenclature of Feltham and Enemark

(39). Only small differences were noted among the spectra of NO-treated Fe(II)-,

187

 

FIGURE 5.3: UV-visible absorption spectra of XanA complexes. (A) Spectra of
anaerobic XanA (black), Fe(II)-XanA (red), aKG-Fe(H)-XanA (blue) and xanthine-aKG-
Fe(II)-XanA (cyan) obtained in 50 mM Tris buffer, pH 8.0. The XanA subunit (0.45 mM)
was mixed with ferrous ammonium sulfate (slightly <0.45 mM) and adjusted to contain 1

mM aKG and 1 mM xanthine. (B) Difference spectra of Fe(II)-XanA in the presence of

aKG (solid) and OLKG plus xanthine (dashed).

I
\
1500- ‘
‘I
.

1000-

8 (MA, cm'1)

 

 

 

500 . - . . . --
400 500 000 700 000

 

 

 

188

FIGURE 5.4: UV—visible absorption spectra of {FeNO}7 complexes of TauD and
XanA. The spectra of substrate-aKG-Fe(II)-protein complexes (0.45 mM protein
subunit) were obtained prior to (solid lines) and afler (dashed lines) incubation with
0.9 mM DEA/NO at 25 °C for (A) TauD (incubation time 35 min) and (B) XanA
(incubation time 50 min). (C) Difference spectrum of xanthine-aKG—Fe(II)-XanA in
the presence and absence of NO. (D) Difference spectrum of

taurine-aKG-Fe(II)-TauD in the presence and absence of NO.

 

 

 

'400'500 060'760'000
run
1000-

800

. ID
600 ‘
400 -

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30'560T060 700'000
nm

 

189

ocKG-Fe(ll)-, and substrate-aKG-Feal)-enzymes. In all cases, NO addition resulted in
an intense transition near 450 nm (A8443 423 M'l-cm" for TauD and A3435 518
M"°cm'l for XanA) and a second less intense feature near 650 nm (A8630 180 M'l-cm"
for TauD and A5650 32 M"-cm" for XanA). These results are in close agreement with
results previously reported for the herbicide-degrading Fe(II)/aKG hydroxylase deA
(40).

To examine the binding affinity of NO for Fe(II)-TauD, aKG-Fe(lI)-TauD, and
taurine-aKG-Fe(II)-TauD, a series of binding curves were generated by monitoring
the absorbance changes at 443 nm with varied NO concentrations (Figure 5.5A). The
data were used to calculate Kd values of 0.178 mM for taurine-aKG-Fe(II)-TauD,
0.341 mM for OLKG-Fe(lI)-TauD, and 0.279 mM and for Fe(II)-TauD. The smallest
Kd was noted for sample containing both the primary substrate and OLKG, in
agreement with the reported situation for deA (40).

Whereas the absorbance changes associated with the {FeNO}7 species of TauD
were typically complete within 35 min, in the case of the XanA complexes the
{FeNO}7 species were slower to develop and did not reach completion after 50 min
(Figure 5.5B). These kinetics precluded calculation of Kd values for the XanA
complexes, but it was shown that for identical concentrations of DEA/NO the
{FeNO}7 species was generated fastest in sample containing both substrate and aKG
(Figure 5.58).

EPR of {FeN0}7 Adducts of TauD and XanA. X-band CW—EPR spectra for

NO-treated Fe(II)-TauD, aKG-Fe(II)-TauD, taurine-aKG-Fe(II)-TauD, Fe(II)-XanA,

I90

FIGURE 5.5: Thermodynamics and kinetics of NO binding to TauD and XanA. (A)
NO binding curves were generated for the interaction of varied concentrations of
DEA/NO with Fe(II)-TauD (dashed, A), (xKG-Fe(II)-TauD (dotted, e) and
taurine-aKG-Fe(II)-TauD (solid, I). The ordinate represents the fraction of
NO—bound protein complex as monitored by the absorbance change at 443 nm after
35 min at 25 °C. The abscissa represents the free (unbound) NO concentration. (B)
Kinetics of the absorbance changes at 435 nm for Fe(II)-XanA (0),0tKG-Fe(II)-XanA
(A), xanthine-Fe(II)-XanA (V) and xanthine-aKG-Fe(II)-XanA (I). DEA/NO (0.9
mM) was added to XanA (0.45 mM) containing ferrous ammonium sulfate (slightly <
0.45 mM), and aKG (1 mM), xanthine (1 mM), or both OLKG and xanthine (1 mM

each) were added to the sample that was incubated at 25 °C.

 

 

 

 

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191

aKG-Fe(II)-XanA, and xanthine-aKG~Fe(II)-XanA were obtained to monitor the
changes in the electronic environment of the enzyme metallocenters upon binding
OLKG and substrate (Figure 5.6). All the NO-bound samples possessed a nearly axial
signal ofS = 3/2 with g2 = 4.00 and g” = 2.00. This observation is typical for {FeNO}7
complexes and arises from the Mg = :t1/2 Kramers doublet of the S = 3/2 coupled spin
system. The lineshape reflects an axially symmetric zero field splitting interaction
with its principal axis directed along g” which approximates the Fe-NO bond (41). The
region near g 2 was used to investigate structural changes since the g” = 2.00 signal
was relatively weak and confounded by the signal arising from free NO. The EPR
lineshapes of complexes Fe(II)-TauD and aKG-Fe(II)-TauD were almost identical, as
shown in Figure 5.6A and B, and the calculated E/D value is 0.015 for both indicating
a slight rhombic character. The terms D and E refer to axial and rhombic zero-field
splitting constants, respectively, and the ED ratio reflects the degree of rhombic
distortion of the {FeNO}7 center (with a purely axial geometry displaying E/D z 0
_ while a highly rhombic feature exhibits an E/D 2 0.33). Addition of primary substrate,
taurine, generated a more rhombic specieswith features at g = 4.27, 4.07 and 3.89. It
is composed of two distinct populations: an axial species (E/D= 0.009, 68%) and a
more-rhombic one (E/D= 0.03, 32%). Similar results were observed for XanA
complexes. Both Fe(II)-XanA and aKG-Fe(II)-XanA exhibited axial signals (E/D r“:
0), whereas a more rhombic EPR lineshape with features at g = 4.16, 4.03 and 3.78
was observed after addition of xanthine. The more axial species (E/D = 0.01) accounts

for 77% of the signal, whereas the remaining 23% comes from a more rhombic

I92

FIGURE 5.6: EPR spectra of the {FeNO}7 complexes of TauD and XanA. Samples

(0.45 mM protein and slightly less than 0.45 mM ferrous ammonium sulfate) of (A)

TauD or (D) XanA were examined alone or in the presence I mM aKG (B) and (E),

or with 1 mM aKG plus I mMisubstrate after incubation (35 min for TauD (C), 50

min for XanA (F)) with 0.9 mM DEA/NO at 25 °C. The solid line represents

experimental data and dashed line represents the simulation.

1.2::108 ‘

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species (E/D = 0.045). A feature at g = 4.3 existed in all the spectra and was assigned
to free Fe(III) ion.

One-Dimensional ESEEM of {FeNO}7 adducts of T auD and XanA. Across the
EPR lineshape, two magnetic fields, 172.0 mT and 340.0 mT (around g 2 =, 4.00 and g”
= 2.00), were selected to collect ESEEM spectra.

Three—pulse ESEEM spectra collected at 172.0 mT for Fe(II)-TauD,
OLKG-Fe(II)-TauD, and taurine-aKG-Fe(II)-TauD in Tris buffer prepared in H2O are
shown in Figure 5.7 A, C and E, and corresponding spectra collected in 60% 2H2O are
shown in Figure 5.7 B, D and F. The sharp peak at a frequency around 7.4 MHz could
be assigned to the matrix proton in the protein. This feature exists in all TauD
complexes prepared in H2O, but no in the 2H20 samples. The broad peaks from 9.5
MHz to 16 MHz could derive from one or more populations of strongly-coupled, and
these couplings seem stronger in the presence of OLKG or both OLKG and taurine. The
signal from 0 to 5 MHz is more complicated; it shows contributions from the histidyl
ligands and more than one sets of strongly coupled protons. Compared to the spectra
taken in H2O, the signals from 2H2O prepared samples are cleaner. The significant 2H
modulation dominates the spectra at a frequency around 1.1 MHz as shown in Figure
5.7 B, D and F. However, the strong proton couplings in taurine-aKG-Fe(II)-TauD
still exist and become more defined when comparing Figure 5.7 E and F.

The spectra that result from repeating the measurements for TauD samples at
340.0 mT are shown in Figure 5.7 G, I and K (H20) and Figure 5.7 H, J, and L (for

samples prepared in 60% 2H2O). The feature at frequency 14.6 MHz is consistent with

194

FIGURE 5.7: One-dimensional ESEEM spectra of different TauD complexes
prepared in H20 and 60% an0 buffer at 172.0 mT and 340.0 mT. Fe(II)-TauD
complex examined at 172.0 mT in H2O buffer (A) and 60% 2H2O buffer (B),
OLKG-Fe(II)—TauD complex examined at 172.0 mT in H2O buffer (C) and 60% 2H2O
buffer (D), taurine-aKG-Fe(II)-TauD complex examined at 172.0 mT in H2O buffer
(E) and 60% ano buffer (F). Fe(II)-TauD complex examined at 340.0 mT in H2O
buffer (G) and 60% 2H2O buffer (H), aKG-Fe(II)-TauD complex examined at 340.0
mT in H2O buffer (1) and 60% 2H2O buffer (J), taurine-OLKG-Fe(II)-TauD complex
examined at 340.0 mT in H2O buffer (K) and 60% 2H20 buffer (L). Protein
concentration, 2 mM; ferrous ammonium sulfate, slightly less than 2 mM; OLKG and
taurine, 4 mM. ESEEM data were collected under the conditions as described in

experimental procedures.

I95

FIGURE 5.7:

N U
I

intensity (x 10.0)

intensity ( x 10.0)

 

L

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0 4 8 12 15 20 24 28 I 1'6 20 2'4 2'8

1.2 _
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197

 

MHz

FIGURE 5.8: One-dimensional ESEEM spectra of different XanA complexes
prepared in H20 and 60% 2H2O buffer at 172.0 mT and 340.0 mT. Fe(II)-XanA
complex examined at 172.0 mT in H2O buffer (A) and 60% 2H2O buffer (B),
aKG-Fe(II)-XanA complex examined at 172.0 mT in H2O buffer (C) and 60% 2H2O
buffer (D), xanthine-aKG-Fe(II)-XanA complex examined at 172.0 mT in H2O buffer
(E) and 60% 2H2O buffer (F). Fe(II)-XanA complex examined at 340.0 mT in H2O
buffer (G) and 60% 2H2O buffer (H), aKG-Fe(II)-XanA complex examined at 340.0
mT in H2O buffer (1) and 60% 2H2O buffer (J), xanthine-aKG-Fe(II)-XanA complex
examined at 340.0 mT in H2O buffer (K) and 60% 2H2O buffer (L). Protein
concentration, 0.45 mM; ferrous ammonium sulfate, slightly less than 0.45 mM; aKG
and taurine, 1 mM. ESEEM data were collected under the conditions as described in

experimental procedures.

I98

 

FIGURE 5.8:

A 2.5

 

 

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200

the proton Larmor frequency at g = 2. The low frequency region that contains 14N and
1H couplings are more complicated than was observed at g = 4 since bound NO might
also contribute to this feature. A striking difference is that the broad peaks contributed
from the strong proton couplings observed at g = 4 disappeared at g = 2 spectra.

The corresponding spectra obtained with XanA at the two fields (Figure 5.8)
were very similar to those observed for TauD. For both enzymes, when probed at g =
4, the contributions from the coupled nuclei were overlapped making it necessary to
use the two dimensional, 4-pulse HYSCORE method to resolve contributions from
bound histidine nitrogens, coordinated H20, and ambient H20.

Two-Dimensional ESEEM of {FeN0}7 adducts of T auD. TauD was used as the
model system to develop HYSCORE spectroscopy as a tool for analysis of
Fe(II)/(1K6 dioxygenases. As described in the following paragraphs, the HYSCORE
analysis of TauD included the use of H20 and 2H-labeled buffers, studies at several
magnetic field strengths, examination of 2H-labeled substrates, investigation of the
effect of an inhibitor, and the study of W248F variant enzyme.

Comparison of HYSCORE spectra collected at 172.0 mT (perpendicular to the
Fe-NO axis) for TauD samples prepared in aqueous buffer versus buffer containing
60% 2H20 reveal features associated with exchangeable protons and perturbations
induced by the binding of co-substrates (Figure 5.9). In particular, the HYSCORE
spectrum of the enzyme with Fe(II) alone shows two major cross-correlations in the
(+,+) quadrant, Figure 5.9A. The stronger correlations lying in low frequencies (0.94,

1.86 MHz) and (1.77, 0.88 MHZ), which are shown more clearly in Figure 5.10A, are

20]

likely due to 14N couplings. The HYSCORE spectrum also shows at least three pairs
of high frequency correlations, (5.32, 9.14 MHZ) (9.01, 5.76 MHZ), (5.79, 9.57 MHz)
(9.50, 5.89 MHZ) and (4.1, 11.2 MHZ) (11.3, 3.9 MHZ). The correlations with higher
frequencies exhibit a weak, disordered system of overlapping IH arches which most
likely reflects a high degree of disorder at the site prior to co-substrate addition. When
aKG was added, Figure 5.9C , the low frequency feature remains the same, whereas,
the higher frequency correlations, (3.94, 11.58 MHZ) (11.58, 3.78 MHZ), (4.85, 11.64
MHz) (11.86, 4.33 MHZ) and (5.37, 9.70 MHZ) (9.70, 5.55 MHZ), become better
defined and take on the appearance of a wedge-shape that could be indicative of a 1H
hyperfine coupling of rhombic symmetry. Subsequent addition of substrate taurine to
yield the NO-bound ternary taurine-aKG-Fe(II)-TauD complex at the active site
yielded a new IH hyperfine coupling, at frequency correlations (4.48, 11.80 MHZ) and
(11.61, 4.73 MHZ), with smoother contours and better resolution as shown in Figure
5.9E. In addition, there is a less resolved coupling at (2.1, 12.0 MHZ) and (12.3, 2.2
MHZ) lying right beside the major lH interactions. Figure 5.9B, D and F show the
HYSCORE spectra at g = 4 of Fe(II)-TauD, OLKG-Fe(II)-TauD and taurine-aKG-Feal)
-TauD complexes prepared in 60% 2H20 buffer. Compared to signals in H20 buffer,
the low frequency couplings slightly changed in shape, Figure 5.10B, and in their
correlation peaks (1.03, 2.04 MHZ) and (2.07, 1.00 MHZ). For complexes Fe(II)-TauD
and aKG—Fe(II)-TauD, the cross peaks at high frequency disappeared from the
spectrum when the samples are exchanged against 2H20 buffer, however, in the

presence of taurine, this proton correlation is not exchangeable.

202

 

FIGURE 5.9: HYSCORE spectra of different TauD complexes prepared in H20 and 60%

2H20 buffer at 172.0 mT. Fe(II)-TauD complex examined in H20 buffer (A) and 60%

2H20 buffer (B), aKG-Fe(H)-TauD complex examined in H20 buffer (C) and 60% 2H20

buffer (D), taurine-aKG-Fe(II)-TauD complex examined in H20 buffer (E) and 60%

2H20 buffer (F). buffer. Protein concentration, 2 mM; ferrous ammonium sulfate, slightly

< 2 mM; aKG and taurine, 4 mM.
1H A 15J
E 10< E 10.
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203

FIGURE 5.10: HYSCORE spectra of taurine-aKG-FeaD-TauD complex prepared in

H20 buffer at 172.0 mT (A), 340.0 mT (C) and 60% 2H20 buffer at 172.0 mT (B), 340.0

mT (D). All the spectra were examined at a threshold high enough to show l4N couplings.

Protein concentration, 2 mM; ferrous ammonium sulfate, slightly < 2 mM; aKG and

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

taurine, 4 mM.
15
1s. A B
101
.. 1o- 1;
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204

Figure 5.11 exhibits the HYSCORE spectra that result from repeating the
measurement and processing procedure described for Figure 5.9 at 340.0 mT. An
intact contour centered at the diagonal of (+,+) quadrant (14.5, 14.5 MHZ) was
observed for all three Complexes, Fe(II)-TauD, (xKG-Fe(II)-TauD and
taurine-aKG-Fe(II)—TauD, whether they were prepared in H20 or 60% 2H20 bufi'er.
However, the feature at low frequency changed its appearance as substrate was added.
Two weak I4N couplings were shown for complexes Fe(II)-TauD and
(xKG-Fe(II)-TauD at correlations (2.26, 2.94 MHz) (2.94, 2.26 MHZ) and (1.34, 3.34
MHZ) (3.33, 1.35 MHZ), Figure 5.11A and C. In the presence of taurine, only one
coupling remained at correlations (2.15, 3.04 MHz) and (3.04, 2.16 MHZ), Figure
5.11E. The low frequency couplings are very similar for all three complexes when
they were prepared in 2H20 buffer, as illustrated in Figure 5.11B, D, F and 5.11D.
This result is probably due to the dominate feature of 2H signals. One significant
change for the spectra taken at 340.0 mT compared to those taken at 172.0 mM is that
the 1H cross peaks at high frequency are no longer exist.

Three possibilities were considered to account for the origin of the
non-exchangeable protons in NO-treated taurine-aKG-Fe(II)-TauD. First, this lH
could come from C-3 of aKG. To test this option, NO-treated
taurine-NOG-Fe(II)-TauD in H20 was examined by HYSCORE spectroscopy. NOG
is a known inhibitor of several Fe(II)/OLKG-dependent dioxygenase family members
(42-45), including XanA and TauD (31, 46), and is known to compete with OLKG for

binding to the metallocenter. The addition of NOG to Fe(II)-TauD yields a broad

205

FIGURE 5.11: HYSCORE spectra of different TauD complexes prepared in H20 and
60% 2P120 buffer at 340.0 mT. Fe(II)-TauD complex examined at in H20 buffer (A) and
60% szO buffer (B), OLKG-Fe(II)-TauD complex examined in H20 buffer (C) and 60%
2H20 buffer (D), taurine-aKG-FeaD-TauD complex examined in H20 buffer (B) and

60% 2H20 buffer (F). Protein concentration, 2 mM; ferrous ammonium sulfate, slightly <

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

2 mM; OtKG and taurine, 4 mM.
20
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206

absorbance near 400 nm rather than the 530 nm MLCT feature observed for (XKG (46).
After adding taurine and NO to this sample, the protein complex exhibited an
electronic spectrum with a well-defined peak at 408 nm and a feature between 550 nm
and 800 nm, Figure 5.12A. Very similar 111 cross peaks were observed in the
HYSCORE spectrum of this sample in H30 at 172.0 mT compared to that generated
by adding NO to the taurine-aKG-Fe(II)-TauD complex (Figure 5.128). This result
suggested that the non-exchangeable proton likely does not arise from the C-3
position of OLKG Further confirmation of this point was obtained by using aKG that
was incubated for 24 hr in szO at 25 °C to exchange the protons at C-3. This sample
was confirmed to be predominantly intact by the demonstration that its use generated
70% of the activity (km) for freshly prepared aKG When the 172.0 mT HYSCORE
spectra of NO-treated tauine-zH-labeled-(xKG-Fe(II)-TauD (2 mM protein in 60%
2H20 or 0.45 mM TauD in 90% 2H20) were compared to the unlabeled samples, no
significant differences were noted (Figure 5.13), again emphasizing that C-3 OLKG
protons are unlikely to be the source of non-exchangeable protons observed in the
sample.

A second possible source of the non-exchangeable protons is from the primary
substrate. To examine this option, HYSCORE spectra were obtained in H20 for
NO-treated taurine-aKG-Fe(II)-TauD where the taurine was deuterated at C1. The
non-exchangeable IH was still present in the HYSCORE spectra at 172.0 mT,
although a slight change in the low frequency region was observed at 340.0 mT

(Figure 5.14D), a new correlation appeared at (2.21, 2.21 MHZ) compared to

207

FIGURE 5.12: UV-Vis spectra and HYSCORE of ternary complex taurine-NOG-Fe-(ID-
TauD. (A) Left panel: UV-Vis Spectra of anaerobic TauD (black), Fe(II)-TauD (red),
NOG-Fe(m-TauD (blue) and taurine-NOG-Fe-(II)-TauD (cyan) obtained in 50 mM Tris
buffer, pH 8.0. The TauD subunit (2 mM) was mixed with ferrous ammonium sulfate
(slightly < 2 mM) and adjusted to contain 4 mM orKG and 4 mM taurine. Right panel,
difference spectra of Fe(II)-TauD in the presence of NOG (solid) and NOG plus taurine
(dashed). HYSCORE spectra of taurine-NOG-Fe(II)-TauD complex prepared in H20

buffer at 172.0 mT (B) and 340.0 mT (C).

 

 

 

 

 

 

 

 

 

 

 

 

 

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f2 (MHZ) f2 (MHZ)

208

FIGURE 5.13: HYSCORE spectra at 172.0 mT of taurine-ZH—labeled-0tKG-Fe(II)-TauD
prepared in 60% 2H20 buffer (A) and 90% 2H20 buffer. Protein concentration for (A) is 2
mM and (B) is 0.45 mM; ferrous ammonium sulfate for (A) is slightly < 2 mM and (B) is

< 0.45 mM; OLKG and taurine, 4 mM for (A) and 1 mM for (B).

 

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209

FIGURE 5.14: HYSCORE spectra of 2H-labeled taurine-orKG-FeflD-TauD complex
prepared in H20 buffer at 172.0 mT (A) and 340.0 mT (B). Protein concentration, 2 mM;

ferrous ammonium sulfate, < 2 mM; (xKG and taurine, 4 mM.

 

 

 

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210

non-deuterated spectrum, Figure 5.10C.

The third possibility to account for the observed non-exchangeable 1H is that it
derives from a side chain of TauD. To attempt to identify this proton, the distance
between the lH and Fe was estimated by analyzing the cross-peak lineshape of the
HYSCORE spectrum for NO-treated taurine-aKG-Fe(II)-TauD in H20 at 172.0 mT.
These cross peaks are indicative of an axial hyperfine coupling and can be analyzed
for their dipole-dipole and isotropic hyperfine contributions, |T| and lAisol, respectively,
using a graphical analysis developed by Dikanov and Bowman (4 7). In this analysis,
an arc is constructed down the long-axis of the contour and correlated frequency pairs
(vmvB) are read from points along the arc. One then constructs a plot of V.,? vs. v52 and
extracts the slope, Qa, and y-intercept, Ga, from a linear least squares fit to the points.
A150 and |T| are then determined from knowing the Larmor frequency of the nucleus,
v), and the system of equations given below. v. , the 1H Larmor frequency at g = 4, as
7.323 MHz, [3 (Bohr constant) is 9.27408 x 10'24 JT", 6,, (Bohr magneton constant) is
5.05082x 10'27 JT", g. (electron g factor) is 4.00, gn (nuclear g factor) is 5.586, and h

(Planck’s constant) is 6.62618x 10'34 Is.

Fa = [i4vi(Qa + 1)]/(Qa - l) (2)
m = (2/3)-{a[oa(r., a 4v.)]/2v. — 4v.2 + ref/4} ”1’ (3)
A.,, = (Fa - T)/2 (4)
T = gcgnflcfln/hr3 (5)

In practice, the assignment of the labels or and B to the frequency pairs is

arbitrary and one just has to pick a convention and stick to it. The signs for the first

211

 

term in the square root argument for calculating [Tl are chosen to yield a real root. The
result of using this system of equations is two unique pairs of values for Also and T.
One set features Am and T parameters of the same sign, while for the other, Aiso and T
have opposite signs. For the higher are of Figure 5913, the v0.2 vs. V32 plot yielded Q,
= -0.6364 and GCl = 108.47 MHZZ. Plugging these numbers into equation 2-5 gave T =
5.0 MHZ, Aiso = 1.1 MHZ and r, the distance between the transition metal ion and
nuclei of interest, approximate 3.2 A.

A field dependent experiment was carried out for the taurine-aKG-Fe(II)TauD
sample in 60% 2HZO at 172.0 mT, 182-.0 mT, and 192.0 mT to further study the
gradual hyperfine coupling changes and determine the Aim and T values, Figure 5.15.
After simulation, the non-exchangeable lH cross-peaks are characterized by Aiso =
0.75 i 0.3 MHZ, T = 5.0 MHZ, and fire = 90° ('H is perpendicular to the Fe-NO bond
axis). This result is consistent with the graphical analysis described above.

Examination of the TauD crystal structure reveals six residues that could
position one or more protons at approximately this distance: the three metal ligands
(His99, Asp101, and His255) and Asn95, Trp248, and Arg270, Figure 5.16A. Since
the crystal structure was obtained under anaerobic conditions, the NO ligand was
modeled into the crystal structure using bond lengths and angles from model
compounds studies and placing the Fe-NO bond opposite to the axial histidine ligand
(41). In particular, proton of C-7 in the benzene ring of Trp248 provided the shortest
distance of 3.7 A to the iron center, Figure 5.16B. To test the importance of the

Trp248 C-7 proton relative to the HYSCORE spectrum, we made use of the W248F

212

 

variant that had previously been characterized and shown to retain 37% of the activity
(km) of wild-type TauD (26). In addition, this mutant protein exhibits UV—visible
spectra similar to the control enzyme when incubated anaerobically with aKG and
taurine. The taurine—aKG-Fe(II)-TauD W248F complex was treated with NO and
examined by HYSCORE at 172.0 mT (Figure 5.17). When using 2 mM W248F, the
lH hyperfine coupling was still clearly observed and remained non-exchangeable in
60% 2&0 buffer. Furthermore, the simulation result from taurine-aKG-Fe(II)-TauD
resembled spectrum of taurine-(xKG-Fe(II)-TauD W248F complex as illustrated in
Figure 5.17C. Analogous experiments were not carried out with Arg270 mutants
because this residue is well conserved in TauD sequences and we have found that the
R270K variant is inactive. In the case of Asn95, the available variants (N95A and
N95D) exhibit very large increases in Km (taurine) and would not be suitable for these
studies.

Two-Dimensional ESEEM of {FeN0}7 adducts of XanA. The HYSCORE
spectra of XanA at 172.0 mT (Figure 5.18) and 340.0 mT (Figure 5.19) exhibited
patterns very similar to TauD. A weak, exchangeable IH hyperfine coupling at (5.73,
9.23 MHZ) and (9.37, 5.48 MHZ) was observed before addition of (xKG and xanthine;
stronger, exchangeable IH couplings appeared when orKG was added; and further
interaction of xanthine caused a new, non-exchangeable lH coupling at (4.25, 11.71
MHZ) and (11.69, 3.78 MHz). The low frequency l4N couplings are similar to TauD
spectra in 2H20 prepared buffer, comparing Figure 5.20A to Figure 5.11A. When the

172.0 mT spectra were examined at a less sensitive contour or the spectra were

213

 

FIGURE 5.15: HYSCORE spectra of taurine-aKG-Fe(II)-TauD complex prepared in
60% 2H20 buffer at 172.0 mT (A), 182.0 mT (B) and 192.0 mT (C). Red dots are

simulations. Protein concentration, 2 mM; ferrous ammonium sulfate, slightly < 2 mM;

 

aKG and taurine, 4 mM.
14. i 7 . 3172023.
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a 5
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'6 is 1'0
r2 (MHz)

214

FIGURE 5.16: Two depictions of the TauD active site (A, B) and the XanA active site
(C). aKG in stick form with its carbon atoms colored yellow is shown chelating the metal
(red sphere). The three side chains that bind the metal are shown in stick form with
carbon in yellow, oxygen in red and hydrogen in white. Residues located close to the
metal center and potentially capable of accounting for the non-exchangeable hyperfine
coupling with the paramagnetic center are shown in stick form with their carbon atoms in
blue and nitrogen in dark blue. Panel B highlights the distance and geometry of Trp248
versus the NO-bound metal center of TauD. Panel C highlights the distance and geometry

of Asp138 and ILe150 to metal center of XanA.

rm.
Taurlm
270
Arg Asn95

His”

HI8255

 

215

FIGURE 5.17: HYSCORE spectra at 172.0 mT of taurine-aKG-Fe(H)—TauD W248F

prepared in H20 buffer (A) and 60% 2H20 buffer (B). Protein concentration is 2 mM;

ferrous ammonium sulfate, slightyly < 2 mM; aKG and taurine, 4 mM. Spectrum (C)

shows the spectrum (A) overlapping with simulations in red.

 

 

 

 

 

 

 

 

 

 

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f2 (MHz)

216

FIGURE 5.18: HYSCORE spectra of different XanA complexes prepared in H20 and
60% 21120 buffer at 172.0 mT. Fe(II)-XanA complex examined at in H20 buffer (A) and
60% 2H20 buffer (B), aKG-Fe(II)-XanA complex examined in H20 buffer (C) and 60%
2H20 buffer (D), xanthine-aKG—FeaD-XanA complex examined in H20 buffer (B) and
60% 2H20 buffer (F). Protein concentration, 0.45 mM; ferrous ammonium sulfate,

slightly < 0.45 mM; ocKG and xanthine, 1 mM.

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

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217

 

FIGURE 5.19: HYSCORE spectra of different XanA complexes prepared in H20 and
60% 2HgO buffer at 340.0 mT. Fe(ID-XanA complex examined at in H20 buffer (A) and
60% 2H20 buffer (B), aKG-Fe(II)—XanA complex examined in H20 buffer (C) and 60%
2H20 buffer (D), xanthine-aKG-FeflD-XanA complex examined in H20 buffer (E) and
60% 2H20 buffer (F). Protein concentration, 0.45 mM; ferrous ammonium sulfate,

slightly < 0.45 mM; OLKG and xanthine, 1 mM.

 

 

 

 

 

 

 

 

 

 

 

 

 

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218

 

 

 

FIGURE 5.20: HYSCORE spectra of xanthine-aKG-Fe(H)—XanA complex prepared in
H20 buffer at 172.0 mT (A), 340.0 mT (C) and 60% 2H20 buffer at 172.0 mT (B), 340.0
mT (D). All the spectra were cut threshold high enough to show l4N couplings. Protein

concentration, 0.45 mM; ferrous ammonium sulfate, slightly < 0.45 mM; ocKG and

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

xanthine, 1 mM.
16¢ A 16‘ B
4;: 10~ 7: 10<
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219

studied using 340.0 mT field, non-exchangeable protons were not convincingly
observed and the features associated with nitrogen atoms were much less obvious than
in the case of TauD. The potential donor of this non-exchangeable proton coule be
Asp138, as shown in Figure 5.16C, or IlelSO, which occupies the similar position of

Trp248 in TauD.

220

DISCUSSION

Characterization of the coordination environment of most Fe(II) enzymes is
hampered by the difficulty of obtaining high—quality crystal structures of these
proteins under anaerobic conditions. One approach to overcome this hurdle is to make
use of various spectroscopic methods to examine the active site environments.
Whereas Fe(II) sites generally are uninformative when examined by electronic and
EPR spectroscopies, prior studies with TauD (20, 28) and other Fe(II)/aKG
dioxygenase family members (40, 48) have shown that the (xKG-bound Fe(II) species
exhibit a visible chromophore attributed to a .MLCT transition (36) and that their
NO-bound centers yield visible species that are paramagnetic and capable of study by
EPR methods. Here, we use the highly soluble, easily obtained, and
well-characterized TauD enzyme as a model system to extend these methods to
include one- and two-dimensional ESEEM, and then we apply these methods to the
study of XanA, a recently identified member of this group of enzymes.

As expected from TauD and other family members, (xKG-Fe(II)-XanA
exhibited characteristic MLCT transitions that were more defined in the presence of
xanthine. Three partially resolved bands, 58] nm, 506 nm and 465 nm, could be
assigned to electron transfer from Fe(II) dyz, d xiy’, d 2’ to the 1!. orbital of aKG
respectively (36). In the presence of NO, XanA developed a distinct UV-visible
spectrum as seen in many Fe(II) proteins. The two main features, were previously
assigned by Solomon and coworkers to NO' to Fe(III) charge transfer transitions, in

which the most intense transition at 443 nm for TauD and 435 nm for XanA are due

22]

 

to the interaction of the out-of-plane (the plane created by the bend Fe-N=O bond)
NO" 21:. to Fe(III) dyz orbitals, and the broad transitions around 650 nm for both
enzymes could be due to the overlap of two transitions, the in-plane NO' 21:. to Fe(III)
d,z and Fe(IIl) d,y to Fe(III) dxif orbitals (41).

NO-treated TauD and XanA complexes in the absence of substrate exhibited a
nearly axial signals of S = 3/2 resulting from the strong antiferromagnetic coupling of
the high-spin Fe(III) (S = 5/2) with NO' (S = 1) (41). In the presence of primary
substrate, both enzymes exhibited a mixed signal at g = 4 composed of a major axial
species (~70%) and a more rhombic one (~30%). Increasing the co-substrate or
substrate ratio to protein from 2.0 to 5.0 did not alter the overall EPR lineshape and
the percentages of each component. Moreover, the diagnostic MLCT transition related
to aKG bidentate binding to Fe(II) did not increase when the ratios of added (XKG or
taurine to protein increased. These results indicated that the two different species were
not caused by slow equilibrium between the protein and substrates, but more likely
caused by slightly different ligand environments of the ternary complexes
taurine-aKG-Fe(II)-TauD or xanthine-aKG-Fe(II)-XanA. Different environments
could be associated with distinct Fe-NO angles arising from different ligand
geometries of the equatorial histidine and aspartate. Both of the observed species
could be catalytic relevant, or only one might lead to turnover. Neither UV-Vis nor
HYSCORE spectroscopies are sensitive enough to discriminatethe basis of the two
species.

One-dimensional ESEEM spectra of {FeNO}7 adducts of TauD gave mixed

222

signals attributed to l4N (bound histidine and NO) and 'H (coordinated H20, ambient
H20, and protons from the protein). Even though 1 could not clearly identify the
source of each feature by l-D ESEEM, the individual signals were useful for later
studies. The general features of the spectra include the following, (i) Two distinct
regions were present in the spectra including a low frequency region, from 0-5 MHz,
and a high frequencydomain, usually above proton Larmor frequency. The high
frequency signals are more resolved, especially, for the case of a non-exchangeable
proton/protons that appears in the presence of substrate. (ii) The low frequency
features are more complicated, probably due to a mixture of different 14N and 1H
couplings. (iii) ESEEM spectra were dominated by 2H modulations when protein
samples were prepared in 60% 2H20 buffer.

The important advantage of HYSCORE techniques lies in the creation of cross
peaks whose coordinates are nuclear frequencies from opposite-electron-spin
manifolds. By using HY SCORE, our ability to identity couplings from different
nuclei is greatly increased, as well as the capability to discriminate noise from signals.
When TauD is incubated with Fe(II) alone, two water molecules should be
coordinated to the Fe center, consistent with HY SCORE spectra at g = 4 which
showed a substantial distribution of exchangeable lH hyperfine couplings attributed to
accessible water molecules. There should be no bound water molecules when (xKG is
added, however, better defined proton couplings appeared in the spectrum. My
explanation is that even though there is no metal bound water, some ambient water

which binds near the {FeNO}7 site are still close enough to modulate the electronic

223

center. Such protons may be attributed to the C-1 protonated carboxylate of (XKG, or
those interacting with the oxygen of the Fe-NO adduct. These proton cross-peaks
disappear from the spectrum when the samples are exchanged against 2&0 buffer
since they come from ambient water. The most interesting result is the appearance of
the non-exchangeable proton in the ternary complex. This contribution is not derived
from (xKG or taurine, as shown from comparison studies of perdeuterated taurine and
the aKG surrogate, NOG. Both graphical analysis and simulation of field dependent
spectra for the taurine-aKG-Fe(II)-TauD sample gave consistent results: Aiso z 1.0
MHz, T z 5.0 MHz, r z 3.2 A and this proton is perpendicular to Fe-NO bond axis, in
agreement with the disappearance of cross-peaks at g = 2. Interestingly,
xanthine-aKG-Fe(II)-XanA exhibited a very similar non-exchangeable lH signals
with much lower intensity, while only exchangeable 1H couplings were observed for
complexes without primary substrate. From these observations, it appears that
whatever residue gives rise to this strong signal, it is coupled to the {FeNO}7 center in
this unique fashion only when primary substrate is added. Perhaps this residue plays a
role in positioning substrate or in guiding the chemistry of the center once the high
valent iron-0x0 intermediate is prepared. Based on TauD the crystal structure, this
proton could be contributed from any of six residues, including the three metal ligands
or Trp248, ,Asn95, and Arg270. W248F TauD has been used to test the importance of
Trp248 and its HYSCORE spectrum showed no substantial difference with wild-type
TauD. These results do not completely rule out Trp248 as a candidate for this coupling

since the phenylalanine residue partially resembles tryptOphan structurally, and it is

224

known that W248F retains 37% of the activity of wild-type TauD. The three metal
ligands are less likely as a source of this proton since no non-exchangeable proton
cross-peaks were observed for Fe(II)-TauD and aKG—Fe(II)-TauD. However, the
nitrogen couplings were greatly affected by adding primary substrate, thus implying
that the positions of metal ligands may shift upon addition of taurine to create an
optimized site for 02 coordination, compatible with the finding that NO has the
greatest affinity to the enzyme in the presence of both co-substrate and substrate.
Further mutagenesis studies will help to locate this proton; however, the mutants need
to be active to confirm proper folding of the protein, so the residue could not simply
be mutated to alanine. The lack of crystal structure of XanA makes it even more
difficult to locate this proton donor. The HYSCORE spectra of XanA exhibited
patterns very similar to TauD, however, the poor signal-to-noise ratio forces one to set
the contour plot threshold high. As a result, only the most intense spots instead of
complete contour were observed for XanA complexes and an accurate comparison
with TauD results is difficult. To overcome this problem, I attempted to set the
threshold of TauD HYSCORE spectra high enough so that only the most intense
couplings can be seen, as shown in Figure 5.2]. The comparison of XanA with this
plot reveals that the correlations of proton hyperfine couplings are not exactly the
same for XanA and TauD in the Fe(II) bound only and OLKG-Fe(II) bound complexes,
as shown in Figure 5.21 and Figure 5.18A, C and E. All the XanA samples exhibited a
less sensitive contour and much less obvious signals than in the case of TauD,

however, this loss of useful information is not just from the decreased sample

225

FIGURE 5.21: HYSCORE spectra at 172.0 mT of Fe(II)-TauD (A), aKG-Fe(II)-TauD
(B), and taurine-aKG-Fe(II)-TauD complex prepared in H20 buffer. The threshold of the
spectra was adjusted to show ("‘1’ the most intense spots. Protein concentration, 2 mM;

ferrous ammonium sulfate, slightly < 2 mM; aKG and taurine, 4 mM.

 

 

 

 

 

 

 

 

 

 

 

 

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o .‘ r ' 1 ' I
o 5 1o 15
rzwuz)
15- C
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o 5 1o 15
f2 (MHZ)

226

concentration of XanA, as compared with TauD at lower concentration (data not
shown). These facts imply that structural differences exist between the two proteins.
The differences could be associated with changes in the coordination geometry from
metal ligands or aKG, or the active site may be affected by surrounding residues to
different extents. Nevertheless, it is not surprising to see these differences based on
discrepencies observed before (32), including selectivity for primary substrate,
spacings between the conserved Asp to the distal His, solvent isotope effect, MLCT
transitions of Fe(II)-orKG-protein complex, electronic rhombicity for the ternary
complex, also the reactivity of the replaced proton (H at C-1 of taurine, H at C-8 of
xanthine).

In summary, I applied different spectroscopies, including UV-Vis, EPR, and one-
and two- dimensional ESEEM, to gain insights into XanA, a recently described
Fe(II)/(1K6 hydroxylase family member that is the first representative to catalyze the
oxidation of purine base. A well-characterized member, TauD, was used as a model
system. (xKG-Fe(II)-XanA exhibited characteristic MLCT transitions that were altered
by the presence of xanthine verifying the (XKG bidentate binding to Fe(II). NO treated
XanA complexes developed similar spectra of UV-Vis, EPR, one- and two-
dimensional ESEEM compared with those of TauD. This study provides the first
spectroscopic information for XanA and yields insights into the coordination
properties of Fe(II) in this protein. An non-exchangeable proton coupling signal was
observed for the ternary complex, this new feature only exists in the presence of the

primary substrate and may play an important role of positioning the substrate in the

227

binding pocket.

228

 

REFERENCES

(1)

(2)

(3)

(4)

(5)

(6)

(7)

(8)

(9)

(10)

Hausinger, R. P. (2004) Fe(II)/or-ketoglutarate-dependent hydroxylases and
related enzymes. Critical Reviews in Biochemistry and Molecular Biology 39,
21-68.

Clifton, I. J., McDonough, M. A., Ehrismann, D., Kershaw, N. J., Granatino,
N., and Schofield, C. J. (2006) Structural studies on 2-oxoglutarate

oxygenases and related double-stranded B-helix fold protein. Journal of
Inorganic Biochemistry 100, 644-669.

Purpero, V., and Moran, G. R. (2007) The diverse and pervasive chemistries
of the a-keto acid dependent enzymes. J. Biol. Inorg. Chem. 12, 587-601.

Kivirikko, K. I., and Pihlajaniemi, T. (1998) Collagen hydroxylases and the
protein disulfide isomerase subunit of prolyl 4-hydroxylase. Advances in
Enzymology and Related Areas of Molecular Biology 72, 325-398.

Schofield, C. J., and Ratcliffe, P. J. (2004) Oxygen sensing by hydroxylases.
Nature Reviews in Molecular Cell Biology 5, 343-354.

Tsukada, Y.-I., Fang, J., Erdjument-Bromage, H., Warren, M. E., Borchers, C.
H., Tempst, P., and Zhang, Y. (2006) Histone demethylation by a family of
ijC domain-containing proteins. Nature 439, 811-816.

Trewick, S. C., Henshaw, T. F ., Hausinger, R. P., Lindahl, T., and Sedgwick,
B. (2002) Oxidative demethylation by Escherichia coli AlkB directly reverts
DNA base damage. Nature 419, I74-178.

Aas, P. A., Otterlei, M., Falnes, P. O., Vagbe, C. B., Skorpen, F., Akbari, M.,
Sundheim, 0., Bjoras, M., Slupphaug, G., Seeberg, E., and Krokan, H. E.
(2003) Human and bacterial oxidative demethylases repair alkylation damage
in both RNA and DNA. Nature 421, 859-863.

Mukherji, M., Chien, W., Kershaw, N. J., Clifton, I. J., Schofield, C. J.,
Wierzbicki, A. S., and Lloyd, M. D. (2001) Structure-function analysis of
phytanoyl-CoA 2-hydroxylase mutations causing Refsum's disease. Human
Molecular Genetics 10, 1971-1982.

Prescott, A. G., and John, P. (1996) Dioxygenases: molecular structure and

role in plant metabolism. Annual Review of Plant Physiology and Plant
Molecular Biology 4 7, 245-271.

229

(11)

(12)

(13)

(14)

(15)

(16)

(17)

(18)

(19)

(20)

Thomburg, L. D., Lai, M.-T., Wishnok, J. S., and Stubbe, J. (1993) A
non-heme iron protein with heme tendencies: an investigation of the substrate
specificity of thymine hydroxylase. Biochemistry 32, 14023-14033.

Fukumori, F., and Hausinger, R. P. (1993) Purification and characterization of

2,4-dichlorophenoxyacetate/a-ketoglutarate dioxygenase. Journal of
Biological Chemistry 268, 24311-24317.

Miiller, T. A., Zavodszky, M. I., Feig, M., Kuhn, L. A., and Hausinger, R. P.
(2006) Structural basis for the enantiospecificities of R- and

S—phenoxypropionate/a—ketoglutarate dioxygenases. Protein Science 15,
1356-1368.

Eichhom, E., van der Ploeg, J. R., Kertesz, M. A., and Leisinger, T. (1997)

Characterization of or-ketoglutarate-dependent taurine dioxygenase from
Escherichia coli. Journal of Biological Chemistry 272, 23031-23036.

Kertesz, M. A. (1999) Riding the sulfur cycle - metabolism of sulfonates and
sulfate esters in Gram-negative bacteria. FEMS Microbiology Reviews 24,
135-175.

Elkins, J. M., Ryle, M. J., Clifton, l. J., Dunning Hotopp, J. C., Lloyd, J. S.,
Burzlaff, N. 1., Baldwin, J. E., Hausinger, R. P., and Roach, P. L. (2002) X-ray
crystal structure of Escherichia coli taurine/a-ketoglutarate dioxygenase
complexed to ferrous iron and substrates. Biochemistry 41, 5185-5192.

O'Brien, J. R., Schuller, D. J., Yang, V. S., Dillard, B. D., and Lanzilotta, W.
N. (2003) Substrate-induced conformational changes in Escherichia coli

taurine/a-ketoglutarate dioxygenase and insight into the oligomeric structure.
Biochemistry 42, 5547-5554.

Koehntop, K. D., Emerson, J. P., and Que, L., Jr. (2005) The
2-His-l-carboxylate facial triad: a versatile platform for dioxygen activation

by mononuclear non-heme iron(II) enzymes. Journal of Biological Inorganic
Chemistry 10, 87-93.

Hanauske-Abel, H. M., and Gunzler, V. (1982) A stereochemical concept for
the catalytic mechanism of prolylhydroxylase. Applicability to classification
and design of inhibitors. Journal of Theoretical Biology 94, 421-455.

Ryle, M. J., Padmakumar, R., and Hausinger, R. P. (1999) Stopped-flow
kinetic analysis of Escherichia coli taurine/a-ketoglutarate dioxygenase:
interactions with a-ketoglutarate, taurine, and oxygen. Biochemistry 38,

230

15278-15286.

 

(21) Price, J. C., Barr, E. W., Tirupati, B., Bollinger, J. M., Jr., and Krebs, C. (2003)
The first direct characterization of a high-valent iron intermediate in the
reaction of an a-ketoglutarate-dependent dioxygenase: a high-spin Fe(IV)

complex in taurine/a-ketoglutarate dioxygenase (TauD) from Escherichia coli.
Journal of Biological Chemistry 42, 7497-7508.

(22) Price, J. C., Barr, E. W., Glass, T. E., Krebs, C., and Bollinger, J. M., Jr. (2003)
Evidence for hydrogen abstraction from C1 of taurine by the high-spin Fe(IV)
intermediate detected during oxygen activation by taurine:0t-ketoglutarate
dioxygenase (TauD). Journal of the American Chemical Society 125,
13008-13009.

(23) Proshlyakov, D. A., Henshaw, T. F., Monterosso, G. R., Ryle, M. J., and
Hausinger, R. P. (2004) Direct detection of oxygen intermediates in the

non-heme Fe enzyme taurine/a-ketoglutarate dioxygenase. Journal of the
American Chemical Society 126, 1022-1023.

(24) Riggs-Gelasco, P. J., Price, J. C., Guyer, R. B., Brehm, J. H., Barr, E. W.,
Bollinger, J. M., Jr., and Krebs, C. (2004) EXAFS spectroscopic evidence for
an Fe=O unit in the Fe(IV) intermediate observed during oxygen activation by

taurineza-ketoglutarate dioxygenase. Journal of the American Chemical
Society 126, 8108-8109.

(25) Price, J. C., Barr, E. W., Hoffart, L. M., Krebs, C., and Bollinger, J. M., Jr.
(2005) Kinetic dissection of the catalytic mechanism of

taurineza-ketoglutarate dioxygenase (TauD) from Escherichia coli.
Biochemistry 44, 8138-8147.

(26) Grzyska, P. K., Ryle, M. J., Monterosso, G. R., Liu, J., Ballou, D. P., and
Hausinger, R. P. (2005) Steady-state and transient kinetic analyses of
taurine/a-ketoglutarate dioxygenase: effects of oxygen concentration,
alternative sulfonates, and active site variants on the Fe(IV) intermediate.
Biochemistry 44, 3845-3855.

(27) Sinnecker, S., Svensen, N., Barr, E. W., Ye, S., Bollinger, J. M., Jr., Neese, F.,
and Krebs, C. (2007) Spectroscopic and computational evaluation of the
structure of the high-spin Fe(IV)-0x0 intermediates in taurine:
alpha-ketoglutarate dioxygenase from Escherichia coli and its His99Ala ligand
variant. J Am Chem Soc 129, 6168-79.

(28) Muthakumaran, R. B., Grzyska, P. K., Hausinger, R. P., and McCracken, J.
(2007) Probing the Fe-substrate orientation for taurine/a-ketoglutarate

231

 

(29)

(30)

(31)

(32)

(33)

(34)

(35)

(36)

(37)

dioxygenase using deuterium ESEEM spectroscopy. Biochemistry in press.

Cultrone, A., Scazzocchio, C., Rochet, M., Montero-Moran, G., Drevet, C.,
and Feméndez-Martin, R. (2005) Convergent evolution of hydroxylation
mechanisms in the fungal kingdoms: molybdenum cofactor-independent

hydroxylation of xanthine via a-ketoglutarate dependent dioxygenase.
Molecular Microbiology 5 7, 276-290.

Hille, R. (2005) Molybdenum—containing hydroxylases. Archives of
Biochemistry and Biophysics 433,107-116

Montero-Moran, G. M., Li, M., Rendém-Huerta, E., Jourdan, F ., Lowe, D. J.,
Stumpff-Kane, A. W., Feig, M., Scazzocchio, C., and Hausinger, R. P. (2007)
Purification and characterization of the FeIl- and a-ketoglutarate-dependent
xanthine hydroxylase from Aspergillus nidulans. Biochemistry 46, 5293-5304.

Montero-Moran, G. M., Li, M., Rendon-Huerta, E., Jourdan, F., Lowe, D. J.,
Stumpff-Kane, A. W., Feig, M., Scazzocchio, C., and Hausinger, R. P. (2007)
Purification and characterization of the Fe(II)- and a-ketoglutarate—dependent
xanthine hydroxylase from Aspergillus nidulans. Biochemistry 46, 5293-5304.

Gemperle, C., Aebli, G., Schweiger, A., and Ernst, R. R. (1990) PHase cycling
in pulse EPR. J. Magn. Reson. 88, 241-256.

Hoffman, B. M., Venters, R. A., and Martinsen, J. (1985) General theory of
polycrystalline ENDOR patterns. Effects of finite EPR and ENDOR
component linewidths. J. Magn. Reson. 62, 537-542.

Grzyska, P. K., Miiller, T. A., Campbell, M. G., and Hausinger, R. P. (2007)
Metal ligand substitution and evidence for quinone formation in

taurine/a-ketoglutarate dioxygenase. Journal of Inorganic Biochemistry 10],
797-808.

Pavel, E. 6., Zhou, J., Busby, R. W., Gunsior, M., Townsend, C. A., and
Solomon, E. I. (1998) Circular dichroism and magnetic circular dichroism
spectroscopic studies of the non-heme ferrous active site in Clavaminate

synthase and its interaction with a-ketoglutarate cosubstrate. Journal of the
American Chemical Society 120, 743-753.

Zhou, J., Kelly, W. L., Bachmann, B. 0., Gunsior, M., Townsend, C. A., and
Solomon, E. I. (2001) Spectroscopic studies of substrate interactions with
Clavaminate synthase 2, a multifunctional or-KG-dependent non-heme iron
enzyme: correlation with mechanisms and reactivities. Journal of the

American Chemical Society 123, 7388-7398.

232

(38)

(39)

(40)

(41)

(42)

(43)

(44)

(45)

(46)

Ho, R. Y. N., Mehn, M. P., Hegg, E. L., Liu, A., Ryle, M. A., Hausinger, R. P.,
and Que, L., Jr. (200]) Resonance Raman studies of the iron(lI)-0t-keto acid
chromophore in model and enzyme complexes. Journal of the American
Chemical Society 123, 5022-5029.

Enemark, J. H., and Fleltham, R. D. (1974) Principles of structure, bonding,
and reactivity for metal nitrosyl complexes. Coordination Chemistry Reviews
13, 339.

Hegg, E. L., Whiting, A. K., Saari, R. E., McCracken, J., Hausinger, R. P., and
Que, L., Jr. (1999) Herbicide-degrading (Jr-keto acid-dependent enzyme deA:
metal coordination environment and mechanistic insights. Biochemistry 38,
16714-16726.

Brown, C. A., Pavlovsky, M. A., Westre, T. E., Zhang, Y., Hedman, B.,
Hodgson, K. 0., and Solomon, E. I. (1995) Spectroscopic and theoretical
description of the electronic structure of S = 3/2 iron-nitrosyl complexes and

their relation to Oz activation by non-heme iron enzyme active sites. Journal
of the American Chemical Society 11 7, 715-732.

Baader, E., Tschank, G., Baringhaus, K. H., Burghard, H., and Gunzler, V.
(1994) Inhibition of prolyl 4-hydroxylase by oxalyl amino acid derivatives in
vitro, in isolated microsomes and in embryonic chicken tissues. Biochemical

Journal 300, 525-530.

Cunliffe, C. J., Franklin, T. J., Hales, N. J., and Hill, G. B. (1992) Novel
inhibitors of prolyl 4-hydroxylase. 3. Inhibition by the substrate analogue
N-oxaloglycine and its derivatives. Journal of Medicinal Chemistry 35,

2652-2658.

Chan, D. A., Sutphin, P. D., Denko, N. C., and Giaccia, A. J. (2002) Role of
prolyl hydroxylation in oncogenically stabilized hypoxia-inducible factor-10L.
Journal of Biological Chemistry 277, 40112-40117.

McDonough, M. A., McNeill, L. A., Tilliet, M., Papmichael, C. A., Chen,
Q.-Y., Banerji, B., Hewitson, K. S., and Schofield, C. J. (2005) Selective
inhibition of factor inhibiting hypoxia-inducible factor. Journal of the
American Chemical Society 12 7, 7680-7681.

Kalliri, E., Grzyska, P. K., and Hausinger, R. P. (2005) Kinetic and
spectroscopic investigation of Co", Ni", and N-oxalylglycine inhibition of the
Fen/a-ketoglutarate dioxygenase, TauD. Biochemical and Biophysical
Research Communications 338, 191-197.

233

(47)

(48)

Dikanov, S. A., and Bowman, M., K. (1995) Cross-peak lineshape of
two-dimensional ESEEM spectra in disordered S=1/2, l=1/2 spin systems.
Journal of Magnetic Resonance A 116, 125-128.

Hogan, D. A., Smith, S. R., Saari, E. A., McCracken, J., and Hausinger, R. P.
(2000) Site-directed mutagenesis of 2,4-dichlorophenoxyacetic
acid/a-ketoglutarate dioxygenase. Identification of residues involved in
metallocenter formation and substrate binding. Journal of Biological
Chemistry 275, 12400-12409.

234

 

CHAPTER 6

CONCLUSIONS AND FUTURE RESEARCH

235

CONCLUSIONS

Moco-independent Aspergillus nidulans xanthine hydroxylase (XanA) was
purified as the His6-tagged recombinant protein from both the fungus and a
heterologous bacterial host ’ and characterized as a novel member of
Fe(II)/orKG-dependent dioxygenase superfamily (l). Ferrous XanA catalyzes xanthine
hydroxylation to uric acid, concomitant with the oxidative decarboxylation of aKG,
forming succinate and carbon dioxide. Comparison of XanA isolated from A. nidulans
and E. coli revealed very different quaternary structures and posttranslational
modifications; however, the kinetic properties of XanA purified from both hosts are
very similar (E. coli : ~70 U mg 4, Km of 3] uM for OLKG, and Km of 45 uM for
xanthine at pH 7.4; A. nidulans : 30 U mg '1, and K.,, values of 50 uM and 46 uM at
pH 7.0). Fe(II) is irreplaceable for enzyme activity and some divalent metals, such as
Ni(II), Zn(II) and Cu(II), are competitive inhibitors of Fe(II). A solvent isotope effect
was observed upon substituting H20 with 2H20. This result suggests that a chemical
group possessing an exchangeable proton such as Fe(III)-OOH or Fe(III)-OH is
important in the rate-determining step of the overall reaction. Although the xanthine
C-H bond is broken at C-8 during turnover, substitution of the proton at this position
by 2H did not lead to a substrate isotope effect. This result demonstrates that C-H
cleavage is not the rate-determining step in the reaction. The co-substrate can be
substituted by a-ketoadipate, but not by other (it-keto acids tested. NOG, a known
inhibitor of several other representatives of this superfamily, is a competitive (with

aKG) inhibitor of XanA with a K, of 0.12 pM. XanA displays high specificity

236

towards its primary substrate. 9-methylxanthine and l-methylxanthine act as
alternative substrates with significant activity loss, respectively around 20 and 104
times less than xanthine. 6,8-DHP is a competitive, slow-binding inhibitor with Kiapp
of 12.6 11M, while 2,8-DHP and 8-HP have almost no inhibitory effect, indicating the
potentially important role of the C-6 carbonyl or enol for primary substrate binding.
The above studies represent the first biochemical characterization of purified
xanthine/aKG dioxygenase, and provide valuable baseline information to carry out
further mechanistic and spectroscopic studies.

Fe(II)/aKG-dependent dioxygenases catalyze a wild range of reactions and
utilize substrates ranging from small molecules, such as taurine, to large polymers,
such as proteins or DNA/RNA (2). It is well known that the highly conserved DSBH
structure acts as a stable platform to anchor the Fe(II) and aKG (2). The metal ion is
ligated by the three residues forming a conserved HXD/EXnH motif, while aKG
binds Fe(II) in a bidentate fashion through the C-1 carboxylate and C-2 keto groups.
The most interesting question is how to bind different substrates within this scaffold
to carry out the unique reactions for each enzyme. A homology model of XanA was
generated on the basis of the structure of the related enzyme TauD (I). The XanA
protein is predicted to contain the DSBH comprised of eight B-strands with the Fe(II)
binding ligands (Hisl49, Asp151, and His340), and the co-substrate positioned to
chelate Fe(II) in a bidentate fashion. The aKG C-5 carboxylate is predicted to form a
salt bridge with Arg352, while Ly5122 is well positioned to stabilize the C-1

carboxylate of the co-substrate. Xanthine binds in an active site pocket lined with

237

potential hydrogen bond donors or acceptors (Gln99,-Gln101, Glul37, Asp138,
Ly3122, Gln356, Cys357, and Asn358). Ala was chosen to replace each of the
potential hydrogen-bonding residues at the XanA active site and the eight single
mutants were constructed and purified. Afier analyzing the kinetic properties of the
variants with xanthine, the alternate substrate 9-methylxanthine, 6,8-DHP and
thiol-specific reagents, critical residues that participate in binding of the primary
substrate were identified and a model for xanthine binding to XanA was proposed,
Figure 4.17. Xanthine is shown binding to the enzyme active site via a constellation of
hydrogen bonding interactions which explains the high selectivity towards the
primary substrate. The xanthine binding model derived from my extensive kinetic
comparisons will need to be verified by crystal structure determination, but crystals of
XanA are not available currently.

After carrying of biochemical characterization and mutational/kinetic studies
(focusing on the primary substrate binding mode) of XanA, my ultimate goal and the
most challenging part of this project was to further uncover the mechanism of this
enzymatic reaction by using of spectroscopic methods. Knowing the structures of
each intermediate is the key to understand the mechanism. I started with investigating
the coordination chemistry of different enzyme complexes: XanA-Fe,
XanA-Fe(II)-aKG and XanA-Fe(II)-0tKG-xanthine. I confirmed the bidentate binding
of orKG to Fe(II) by observing the diagnostic MLCT features around 500 nm by
UV-Vis spectroscopy of the XanA-Fe(II)-OLKG and XanA-Fe(II)-0tKG-xanthine

complexes. Consistent with a shift from 6-coordinate to 5-coordinate geometry, I

238

observed enhanced definition of the spectroscopic features after adding substrate.
Since high spin Fe(II) is EPR silent, I used NO, a good 02 surrogate, to bind Fe(II) to
form an {FeNO}7 complex which is EPR active. I found that adding primary substrate
disturbs the electronic environment of Fe(II) and this is supported by the more
rhombic EPR line shape for XanA-Fe(II)-aKG-xanthine compared with XanA-Fe(II)
or XanA-Fe(II)-aKG. Electron nuclear hyperfine coupling between Fe(II) and
surrounding ligands was investigated by using one- and two-dimensional ESEEM
spectroscopies. In this part, I used TauD as a reference and examined even more
extensively than XanA. One-dimensional ESEEM spectra for both XanA and TauD
show modulations from ”N and 1H. At g = 4, the contributions from these coupled
nuclei are overlapped making it necessary to use the two dimensional, 4-pulse
HYSCORE method to resolve contributions from bound histidine nitrogens,
coordinated H20, and ambient H20. For TauD-Fe(II), HYSCORE spectra show a
substantial distribution of exchangeable, lH hyperfine couplings. When co-substrate
orKG is added, the 1H HYSCORE is considerably altered with the dominant hyperfine
coupling arising from an exchangeable, strongly-coupled proton of rhombic symmetry.
Subsequent addition of substrate taurine, to yield the ternary complex at the active site,
showed a new, IH hyperfine interaction that was not exchangeable in 2H20. The
HYSCORE cross-peaks from this lH show a hyperfine tensor of axial symmetry
characterized by a dipole-dipole distance of 3.2 A and an isotropic contribution of 1.1
MHz. Comparison of these data with the X-ray crystal structure of TauD and the

results of parallel studies of TauD variants suggests that this 1H is likely from W248.

239

However, the W248F variant did not show any substantial difference in its spectral
features compared with those of wild-type TauD. Interestingly,
XanA—Fe(II)-0tKG-xanthine exhibited very similar non-exchangeable lH signals with
much lower intensity. Exchangeable IH hyperfine couplings were also observed for
XanA-Fe(II) and the XanA-Fe(II)-aKG complex. This comparison study suggested
that the coordination chemistry of TauD and XanA are very similar, even though the
origin of this strong, non-exchangeable 1H is still unclear, its appearance in both
enzyme complexes means this structural change induced by adding primary substrate
could be quite common for this superfamily and important for understanding the

mechanism.

FUTURE RESEARCH

Constructing Double or Triple Mutants. Eight single mutants (Q99A, Q101A,
E137A, D138A, K122A, Q356A, C357A, and N358A) were constructed and, except
Q99A, all of them exhibited substantial activities. This finding suggested that
modifying only one residue will not cause significant change of primary substrate
binding. Double or even triple mutants might affect xanthine binding more efficiently.
N358A has shown the most significant Km change, so it could be considered first;
Q99A can be excluded; C357A did not show any direct interaction with substrate
based on the homology model, so it could be considered the last possibility. So
(Q101A, N358A), (K122A, N358A), (E137A, N358A), (D138A, N358A) and

(Q3 56A, N358A) could be the first set of double mutants to try out.

240

Determination of Dissociation Constant (K.,). Dissociation constant (Kd) is a more
accurate parameter than Km to examine substrate binding. Fluorescence spectroscopy
was applied to try to determine the Kd. Fluorescence measurements were carried out at
an excitation wavelength of 280 nm with emission monitored from 300 to 400 nm. The
binding of Fe, orKG and xanthine each quenched the endogenous fluorescence of the
proteins. However, the quenching by xanthine is artificial since xanthine absorbs at
268 nm which is too close to the excitation wavelength 280 nm. So the quenching
observed upon adding xanthine is actually caused by fewer photons being absorbed by
the protein. So finding a better to determine the dissociation constant (Kd) of xanthine

would be very helpful to understand primary substrate binding mode for XanA.

Steady-State and Transient Kinetic Studies by Stopped-Flow UV-Vis spectroscopy.
Stopped-flow UV-Vis spectroscopy is a powerful technique to detect intermediates
forming within ms to s. The chromophores associated with XanA-Fe(II)-0tKG and
XanA-Fe(II)-0tKG-xanthine could be examined to see if their formation is fast enough
to be catalytically relevant. Next, the oxygen reactivity of XanA-Fe(II)-0tKG and
XanA-Fe(II)-aKG-xanthine could be examined. The Fe(IV)=Oz' intermediate has
been observed by stopped-flow methods applied to three family members, and in each
case exhibited an absorption near 318 nm by UV/Vis. We could assess whether a
similar signal is formed in XanA. To enhance the chances of observing this species,
one can attempt to slow down the decay reaction. Using 2H20 instead of H20 may be

a useful approach since XanA is known to exhibit a significant solvent isotope effect.

241

 

Alternatively, the intermediate may be longer-lived in the presence of a poor
substrates, such as using 9-methylxanthine instead of xanthine. Probably the best
approach is to use 8-2H-xanthine instead of xanthine. Even though I did not observed
any isotope effect during steady-state kinetic studies, the deuterated substrate is likely
to reduce the rate of the hydrogen abstraction in the reaction and should facilitate the
transient kinetic approaches. If transient absorption changes are observed with
wild-type XanA, single or double mutants which are potentially important for

substrate binding or enzymatic reactions also could be examined by this technique.

Resonance Raman and Mossbauer Spectroscopy. The UV-Vis spectrum provides
information on electron transitions, but in order to further define the properties of the
lntennediate species, more sophisticated methods need to be applied, such as
resonance Roman and Mossbauer spectroscopy. For RR, typical vibrational modes of
each intermediate, combining the information from modeling studies and other
members of this superfamily, could help us assign each species. Mossbauer
spectroscopy is very sensitive to Fe redox state and coordination, and it detects the Fe
center no matter whether it is paramagnetic or diamagnetic, so it is a very powerful
technique to investigate Fe-containing proteins. So, freeze-quench Mossbauer

analyses may be a useful tool for further studies.

More HYSCORE Spectroscopy studies. In order to assign the non-exchangeable 1H

in the TauD-Fe(II)—aKG-taurine complex to certain residue, more single mutants

242

 

could be prepared, such as substitutions involving Asn95 and Arg270, which also lie
within 5 A around Fe(II) of TauD. The metal binding ligands are also possible targets
to examine if incorporation of deuterated histidine and aspartate could be
accomplished. The same strategy is applicable for wild-type XanA and mutants such
as D138A (this residue lies within 5A around Fe(II) of XanA), and might be the first
mutant to examine. Another interesting study is to assign the strong couplings we
observed at g“ = 2.00 (not observed at g f. 4.00) for both TauD and XanA. It could

come from NO or axial histidine ligand as shown in Figure 6.1.

Problems Application of many techniques mentioned above, such as, HYSCORE,
RR, Mossbauer, even UV—Vis, require substantial amounts of concentrated (at least
0.5 mM), active and stable protein. XanA isolated from E. coli barely meets this
requirement and only freshly prepared protein can be used. So, the further studies of
the XanA enzyme mechanism will benefit from additional investigation designed to
improve the protein yield and stability.

From my HYSCORE study, I found the non-exchangeable IH only appearing
when the primary substrate is present. This suggests that a protein structural change is
induced by adding substrate. Mutation of targeted residues could help to locate this
proton; however, the mutants need to be active to confirm proper folding of the
protein. That is why we could not simply mutate the metal ligands to alanine. Further

mutagenesis studies will need to take this problem into account.

243

FIGURE 6.1: HYSCORE spectra of NO treated ternary complex TauD-Fe-OLKG

-taurine in 50 mM Tris-H20, pH = 8.0 at (A) 3400 G and (B) 17200

 

 

 

 

 

 

 

 

 

20

15 \Q
A \It.
N
I 101 a
E
v t _
E 54 I

o.
o ‘—l ' I ' I 1 I 1 I 1—I fiT
-2o -15 -1o -6 o I 10 15 20
f2 (MHz)
A

151
7.710- A \1 1
I ., 1 ,
E 1‘5 ».
V
a: 51 :§\

1 1‘

o I ' I ' I ' I t I ‘7I ‘ I

-15 -1o -5 o 5 1o 15
12 (MHz)
B

244

REFERENCES

(1)

(2)

Montero-Moran, G. M., Li, M., Rendon-Huerta, E., Jourdan, F., Lowe, D. J.,
Stumpff-Kane, A. W., Feig, M., Scazzocchio, C., and Hausinger, R. P. (2007)
Purification and characterization of the Fe(II)- and a-ketoglutarate-dependent
xanthine hydroxylase from Aspergillus nidulans. Biochemistry 46, 5293-5304.

Hausinger, R. P. (2004) Fe(II)/0t-ketoglutarate-dependent hydroxylases and
related enzymes. Crit. Rev. Biochem. Mol. Biol. 39, 21-68.

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