ROLES OF UREG AND UREF IN UREASE ACTIVATION
By
Jodi Lynn Boer

A DISSERTATION
Submitted to
Michigan State University
In partial fulfillment of the requirements
For the degree of
DOCTOR OF PHILOSOPHY
Biochemistry and Molecular Biology
2012

ABSTRACT
ROLES OF UREG AND UREF IN UREASE ACTIVATION
By
Jodi Lynn Boer
Urease hydrolyzes urea to ammonia and carbamic acid, which then spontaneously
decomposes into another ammonia molecule and carbonic acid. The active site contains two
nickel atoms bridged by a carbamylated lysine residue. In order to assemble the active site,
four accessory proteins, UreD, UreE, UreF, and UreG, are necessary. This dissertation focuses
on the roles UreG and UreF play in the activation process by investigating the individual
characteristics of these proteins as purified, as well as determining how they interact with the
other urease accessory proteins in the Klebsiella aerogenes system.
UreG is a GTPase required for the assembly of the urease active site; however, it has no
GTPase activity when purified alone. A Strep-tagged version, UreGStr, as well as several sitedirected variants were constructed and their effects on urease activation, metal-binding
properties, and protein: protein interactions with other urease-related proteins were assessed.
The Strep-tag had no effect on the ability of UreG to participate in urease activation, but the
K20A, D49A, C72A, H74A, D80A, and S111A variants essentially abolished enzyme activity.
UreGStr binds one nickel or zinc ion per monomer (Kd ~ 5 µM for each metal ion). The binding
site includes residue Cys72 as shown by the 12-fold increase in the Kd for nickel ions in this
variant, as well as a lack of a thiolate-to-nickel charge-transfer band in the UV-visible spectrum.
Based on homology to HypB, a hydrogenase maturation protein, His74 is also a likely metal

ligand.

Pull-down assays in cell-free extracts demonstrated that Asp80 is essential for

stabilizing the UreGStr interaction with UreABC—UreD—UreF.

In vitro pull-down assays

demonstrated that the interaction between UreGStr and UreE is metal-dependent. This result
suggests that UreE transfers its bound Ni to UreG in the UreABC—UreD—UreF—UreG complex,
where it can then be passed to the nascent active site, possibly via UreD.
UreF was proposed previously to be a GTPase activating protein (GAP) for the GTPase
UreG. Based on the UreF crystal structure from Helicobacter pylori, sixteen residues in K.
aerogenes UreF were chosen for mutagenesis to alanine. When produced in the context of the
urease gene cluster, cell-free extracts of nine site-directed mutants had less than 10% of the
wild-type activity. Using the UreE-F construct and its variants in the same context, UreE-F was
demonstrated to co-purify with urease apoprotein, UreD, and in some cases UreG from cellfree extracts. The variants that did not bind UreG correlated with low urease activity mutant
cells, and mapped to a distinct surface on the UreF structure, defining the UreG binding site. In
contrast to the GAP hypothesis for UreF, the UreABC—UreD—UreF(K165A)—UreGStr complex
had higher GTPase activity than the wild-type complex according to urease activation assays on
purified protein. Further studies showed that the urease activity and GTPase activity were
uncoupled in the K165A UreF-containing complex.

Additional experiments with these

complexes demonstrated that UreF gates the GTPase activity of UreG in order to enhance the
fidelity of urease activation and guard against incorrect metal insertion in the presence of Zn.

To my parents, who always believed in me and taught me to do my best
To Tom, for encouraging me and loving me

iv

ACKNOWLEDGEMENTS

Many people have been essential to my success in graduate school. I would not have
been able to do any of this work without the guidance of Dr. Robert Hausinger, who has been a
wonderful person to work for. I have appreciated how he is always available to talk to. My
fellow graduate students in the Hausinger lab – Soledad, Jana, Eric, Nicholas, and Mark all
contributed to thought-provoking conversations. The post-docs who have worked here – “Big”
Scott, Tina, Piotr, Hajime, Lee, and William have all helped me out as well. I especially thank
Brittnie DeVries, who worked as “my” undergrad for two years and did much of the cloning,
protein purification, and pull-down assays described here. I thank Tejas and Chris who both
worked with me for a while, as well as the other undergrads who worked in the lab. I thank my
classmates who got me through the first year of classes. My committee members also have my
gratitude for their input.
I would not have gotten this far without the support of my family. My parents have
always believed in and pushed me to do my best. I thank Tom for all of his encouragement.
Though I didn’t know him when I started, he has been my rock through this push to the finish. I
thank my extended family for all their prayers and reminding me that there is life outside the
lab. I thank my friends, especially my Bible studies through the years for their prayer support.
Most of all, I am grateful to the Lord for all the blessings in my life.

v

TABLE OF CONTENTS

LIST OF TABLES ................................................................................................................................ ix
LIST OF FIGURES ............................................................................................................................... x
LIST OF ABBREVIATIONS ................................................................................................................ xii
CHAPTER 1
INTRODUCTION ............................................................................................................................... 1
Introduction to Ni-binding sites .................................................................................................. 2
Common features of Ni-binding sites...................................................................................... 2
Ni-binding sites in enzymes ..................................................................................................... 3
Ni-binding sites in non-catalytic proteins................................................................................ 8
Ni-substituted proteins ......................................................................................................... 12
Ni-binding to His-tagged proteins ......................................................................................... 14
Introduction to urease ............................................................................................................... 15
Urease structure .................................................................................................................... 16
Urease activation................................................................................................................... 18
Eukaryotic urease activation ................................................................................................. 24
Remaining questions.................................................................................................................. 27
References ................................................................................................................................. 29
CHAPTER 2
MUTAGENESIS OF KLEBSIELLA AEROGENES UREG TO PROBE NICKEL BINDING AND
INTERACTIONS WITH OTHER UREASE-RELATED PROTEINS .......................................................... 40
Abstract ...................................................................................................................................... 41
Introduction ............................................................................................................................... 42
Experimental Procedures........................................................................................................... 48
Vector construction, cell growth, and purification of Strep-tagged UreG ............................ 48
UreE purification ................................................................................................................... 51
Site-directed mutagenesis ..................................................................................................... 51
Circular dichroism ................................................................................................................. 52
Analytical gel filtration chromatography .............................................................................. 52
In vivo expression of UreGStr variants in the context of the urease operon ........................ 53
Urease activity assays ............................................................................................................ 53
Metal quantification .............................................................................................................. 54
Metal binding analysis ........................................................................................................... 54
UV/Visible spectroscopy........................................................................................................ 55
Pull-down assays ................................................................................................................... 55
Western blot .......................................................................................................................... 56

vi

Results ........................................................................................................................................ 57
Characterization of Strep-tagged UreG ................................................................................. 57
Targeting residues for mutagenesis ...................................................................................... 60
Effect of UreGStr variants on urease activity in cell extracts ................................................ 61
Metal binding to UreG, UreGStr, and UreGStr variants ......................................................... 63
Pull-down assays ................................................................................................................... 73
Discussion................................................................................................................................... 78
References ................................................................................................................................. 85
CHAPTER 3
KLEBSIELLA AEROGENES UREF: IDENTIFICATION OF THE UREG BINDING SITE AND ROLE IN
ENHANCING THE FIDELITY OF UREASE ACTIVATION .................................................................... 90
Abstract ..................................................................................................................................... 91
Introduction ............................................................................................................................... 92
Experimental Procedures........................................................................................................... 97
Plasmids, oligonucleotides, and site-directed mutagenesis ................................................. 97
Protein purification ............................................................................................................... 99
Sodium dodecyl sulfate (SDS)-polyacrylamide gel electrophoresis (PAGE) ....................... 101
Pull-down assays using cell-free extracts ............................................................................ 101
In vitro pull-down assays ..................................................................................................... 101
Urease activity and protein assays ..................................................................................... 102
Urease activation assays .................................................................................................... 103
GTPase activity assays ........................................................................................................ 103
Results ...................................................................................................................................... 105
Targets for site directed mutagenesis ................................................................................. 105
Mutants in ureF have variable effects on in vivo urease activity ........................................ 107
UreF variants primarily affect binding of UreG in cell-free extracts .................................. 109
UreF variants affect in vitro binding to (UreABC—UreD)3 and UreGStr ............................ 112
Examination of UreF as a GTPase activating protein .......................................................... 116
UreF functions to increase the fidelity of urease activation .............................................. 120
Discussion ................................................................................................................................ 124
References ............................................................................................................................... 130
CHAPTER 4
ADDITIONAL STUDIES, CONCLUSIONS, AND REMAINING QUESTIONS ...................................... 134
Additional Studies .................................................................................................................... 135
Metal analysis of (UreABC*)3 and Ure(AC)3 ....................................................................... 135
Crystallization Attempts ...................................................................................................... 138
XAS studies on UreG ............................................................................................................ 139
In vitro pull-down assays with UreE-F and MBP-UreD ........................................................ 140
Characterization of MBP-UreD—UreF—UreG .................................................................... 142
Characterization of UreABC—UreD—UreF—UreGStr ......................................................... 149

vii

Conclusions and Remaining Questions .................................................................................... 154
References ............................................................................................................................... 157
APPENDIX A ................................................................................................................................. 158
REFERENCES ................................................................................................................................ 161

viii

LIST OF TABLES

Table 2.1 Plasmids and E. coli strains used in this study ............................................................. 49
Table 2.2 Oligonucleotides used to generate UreG mutations ................................................... 52
Table 2.3 Effects of UreG mutations on the urease activity in soluble cell-free extracts ............ 63
Table 2.4 Thermodynamics of nickel ion binding to UreG, UreGStr, and its variants ................. 69
Table 3.1 Plasmid properties ....................................................................................................... 98
Table 3.2 Oligonucleotides used in this study ............................................................................. 99
Table 3.3 Effect of UreF variants on urease activity in cell-free extracts ................................... 109
Table 3.4 Binding of UreABC—UreD by control and variant UreE-F samples ............................ 114
Table 3.5 Binding of control and variant UreE-F by UreGStr ...................................................... 115
Table 4.1 Metal content of urease apoprotein samples subjected to activation conditions .... 138
Table 4.2 Samples sent for XAS analysis ..................................................................................... 140
Table 4.3 Binding of control and mutant UreE-F to MBP-UreD ................................................. 142
Table 4.4 GTPase activities of MBP-UreD—UreF—UreG samples ............................................. 145
Table 4.5 GTPase activities of MBP-UreD—UreF—UreG(T21A) samples................................... 146
Table 4.6 GTPase activity of His-MBP-UreD—UreF—UreG and variants ................................... 148
Table A.1 Plasmids used in this work .......................................................................................... 159

ix

LIST OF FIGURES

Figure 1.1 Active sites of Ni-containing enzymes .......................................................................... 7
Figure 1.2 Ni-binding sites in Ni-sensor or Ni-delivery proteins .................................................. 11
Figure 1.3 Ni-substituted protein metallocenters ........................................................................ 13
Figure 1.4 His-tag used for affinity purification ........................................................................... 14
Figure 1.5 Genetic structures of the urease gene clusters from K. aerogenes and H. pylori ....... 17
Figure 1.6 Urease structures can form different supramolecular structures .............................. 17
Figure 1.7 Interactions among urease-related proteins ............................................................... 19
Figure 1.8 Crystal structure (3FS5) of (UreH—UreF)2 from H. pylori ........................................... 21
Figure 1.9 Crystal structure (1GMW) of truncated UreE from K. aerogenes ............................... 24
Figure 2.1 Simplified scheme of the urease activation process .................................................. 43
Figure 2.2 Multiple sequence alignment of the K. aerogenes UreG, B. pasteurii UreG, and M.
jannaschii HypB ........................................................................................................... 46
Figure 2.3 Purification of UreGStr and size exclusion chromatography native size analysis ........ 59
Figure 2.4 Equilibrium dialysis analysis of metal binding to wild type and Strep-tagged UreG ... 65
Figure 2.5 Equilibrium dialysis analyses to assess the competition of nickel and zinc ions ......... 67
Figure 2.6 UV-visible spectral titrations of UreGStr and its C28A variant with NiCl2 ................... 71
Figure 2.7 Difference in UV-visible absorbance spectra according to Ni concentration.............. 72
Figure 2.8 Interactions of UreGStr with urease proteins using cell-free extracts and in vitro
interactions between UreGStr and UreE ................................................................... 75
Figure 2.9 Chromatography of the UreGStr:UreE complex .......................................................... 76
Figure 2.10 Model of UreG:UreE interaction in the urease activation process .......................... 84
Figure 3.1 Simplified scheme of the urease activation process .................................................. 93

x

Figure 3.2 Alignment of UreF sequences from selected microorganisms.................................. 106
Figure 3.3 Mutagenesis of K. aerogenes UreF ............................................................................ 107
Figure 3.4 Effect of UreF variants on urease activity in cell-free extracts.................................. 108
Figure 3.5 UreE-F pull-down analyses of cell-free extracts ....................................................... 111
Figure 3.6 UreF mutations affecting UreG binding .................................................................... 112
Figure 3.7 In vitro pull-down assay examples ............................................................................ 116
Figure 3.8 Effect of K165A UreF on GTP-dependent activation and phosphate release by
UreABC—UreD—UreF—UreGStr ............................................................................ 119
Figure 3.9 Comparison of activation time course for UreABC—UreD—UreF—UreGStr complex
and UreABC—UreD—UreF(K165A)—UreGStr ........................................................ 121
Figure 3.10 Zn inhibition of UreABC—UreD—UreF—UreGStr activation for complexes containing
wild type and K165A UreF ........................................................................................ 123
Figure 4.1 UreB of K. aerogenes urease blocks access to the active site ................................... 136
Figure 4.2 Fractions of MBP-UreD—UreF—UreG tested for GTPase activity ............................ 145
Figure 4.3 Fractions of MBP-UreD—UreF—UreG(T21A) tested for GTPase activity.................. 146
Figure 4.4 Fractions of His-MBP-UreD—UreF—UreG, His-MBP-UreD—UreF—UreG(T21A), and
His-MBP-UreD—UreF(K165A)—UreG that were tested for GTPase activity .......... 147
Figure 4.5 Effect of adding GTP to MBP-UreD—UreF—UreG .................................................... 149
Figure 4.6 Gel filtration profile of UreABC—UreD—UreF—UreGStr .......................................... 150
Figure 4.7 Native gel of UreABC—UreD—UreF—UreGStr and UreABC—UreD—UreF—UreG .. 151
Figure 4.8 Thermal stability of UreABC—UreD—UreF—UreGStr ............................................... 153

xi

ABBREVIATIONS

CD

circular dichroism

DTT

dithiothreitol

EDTA

Ethylenediaminetetraacetic acid

GAP

GTPase activating protein

HEPES

4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid

ICP-AES

inductively coupled plasma-atomic emission spectroscopy

IPTG

isopropyl β-D-1-thiogalactopyranoside

MBP

maltose binding protein

MWCO

molecular weight cut-off

NTA

nitrilotriacetic acid

NMR

nuclear magnetic resonance

PAGE

polyacrylamide gel electrophoresis

PCR

polymerase chain reaction

PAR

4-(2-pyridylazo)-resorcinol

SAXS

small-angle x-ray scattering

SDS-PAGE

sodium dodecyl sulfate-polyacrylamide gel electrophoresis

TCEP

tris(2-carboxyethyl)phosphine

Tris

2-amino-2-(hydroxymethyl)-1,3-propanediol

(UreABC)3

urease apoprotein

xii

(UreABC*)3

complex formed by incubating purified (UreAC)3 with UreB

UreE-F

translational fusion of UreE and UreF

UreGStr

UreG tagged with Strep II

XAS

X-ray absorption spectroscopy

xiii

Chapter 1

Introduction

Portions of this chapter were adapted from
Boer, J. L., and Hausinger, R. P. (2012) Ni-binding sites in proteins, in Encyclopedia of
Metalloproteins (Vladimir Uversky, R. H. K., Eugene A. Permyakov, Ed.) (in press)
and
Carter, E. L., Flugga, N., Boer, J. L., Mulrooney, S. M., and Hausinger, R. P. (2009) Interplay of
metal ions and urease, Metallomics 1, 207-221

1

This thesis describes the characterization of two proteins, UreF and UreG, required for
activation of urease, a Ni-containing enzyme. To place my work in context, this introductory
chapter (i) describes well-defined structures of Ni-binding sites in proteins, (ii) provides an
overview of what is known about urease and its maturation, and (iii) highlights remaining
questions related to urease and describes where my work has filled in some gaps.
INTRODUCTION TO NI-BINDING SITES
Ni is essential to many life forms, but excess levels are toxic and carcinogenic; thus, cells
have devised mechanisms to sense Ni concentrations and maintain homeostatic control of this
metal ion (1, 2). Ni-binding sites occur in Ni-dependent enzymes and a wide variety of proteins
that transport or deliver Ni, regulate transcription of genes involved in Ni metabolism, or
adventitiously bind the metal ion. Understanding the features of Ni-binding sites in proteins
with well-defined structures gives a wide view of how Ni is used in nature, and also allows for a
better understanding of how Ni-binding proteins that do not have defined structures may
chelate the metal. Understanding of Ni-binding sites is applicable to the urease maturation
system, where we only have partial knowledge of some of the Ni-binding sites, and do not
know the specifics of how the Ni is transferred into the active site.
Common features of Ni-binding sites
2+

The most common oxidation state for Ni in biology is Ni , although redox active sites
+1

are present in some Ni enzymes leading to Ni

+3

or Ni

states. The metal ion is most often

found in six-coordinate octahedral configuration; however, planar or tetrahedral fourcoordinate geometries and various five-coordinate ligand environments are known.

2

The

imidazole group of His side chains is the most common amino acid ligand of Ni, with other
amino acids coordinating the metal via the sulfur atoms of Cys or Met and the carboxylate
groups of Glu and Asp residues (2). Backbone amides and amino-terminal amine groups bind
the metal ion in some proteins, and Ni can be part of a larger metallocluster or incorporated
into a tetrapyrrole bound to some proteins.
Ni-binding sites in enzymes
Enzymes utilize Ni to perform a wide variety of reactions, from hydrolysis to redox
chemistry. A key aspect of Ni binding at enzyme active sites is that the metal ion must retain at
least one open coordination site in order to bind the substrate. This open site may be occupied
by water in the resting enzyme. Ni ligands in these enzymes vary widely, typically involving
amino acid side chains but also sometimes utilizing backbone atoms and non-protein cofactors
(1-5). Figure 1.1 illustrates the active sites with known structures of Ni-containing enzymes. All
are crystal structures except for that of acireductone dioxygenase, which is derived from NMR
studies.
Glyoxalase I binds the hemithioacetal derived from addition of glutathione and
methylglyoxal, then uses Ni to catalyze its conversion to S-D-lactoylglutathione (6).

The

substrate likely displaces one or both water ligand(s) when coordinating to the Ni, and the
2+

oxidation state remains Ni

throughout the reaction. This protein illustrates how multiple

peptide chains, in this case two, can cooperate to form a single Ni-binding site.
Acireductone dioxygenase uses the Ni at its active site to catalyze the cleavage of 1,2dihydroxy-3-keto-5-methylthiopentane (acireductone) by reaction with oxygen, producing
methylthiopropionate, formic acid, and carbon monoxide (7). Ni likely remains in the 2+

3

2+

oxidation state during the reaction, acting as a Lewis acid. This enzyme alternatively binds Fe ,
resulting in the distinct products formic acid plus the α-keto acid precursor of Met, which is
used as part of a salvage pathway (8).
Ni-superoxide dismutase catalyzes the disproportionation of two molecules of
superoxide to form oxygen and hydrogen peroxide, identical in chemistry to the Cu/Zn-, Mn-,
and Fe-superoxide dismutases, which are unrelated in sequence (9, 10). In this case, Ni cycles
2+

between Ni

in a square planar configuration and Ni

3+

in a square pyramidal coordination.

Unique to the Ni superoxide dismutase, the protein backbone forms part of the coordination
sphere of the metal, using both the N-terminal amine and amide nitrogen ligands. His, a fifth
ligand in the oxidized form of the enzyme, swings away from the metal ion in the reduced form.
Methyl coenzyme M reductase catalyzes the final step in methane formation in
methanogenic

archaea.

The

enzyme

reacts

methyl-coenzyme

M

(methyl-S-

thioethanesulfonate) with coenzyme B (N-7-mercaptoheptanoylthreonine phosphate),
producing methane and disulfide-linked CoM-CoB. The active site contains an F430 cofactor,
which is a Ni-containing tetrapyrrole related to, but extensively modified from, sirohemes and
corrinoids (11). A Gln side chain coordinates at an axial ligand position, and substrates react at
the other axial site. A very similar Ni-tetrapyrrole is found in an enzyme related in sequence to
methyl coenzyme M reductase, but in that case it participates in anaerobic methane oxidation
+1

(12). The Ni cofactors in these enzymes undergo redox chemistry that includes Ni
Ni

+2

and/or Ni

+3

along with

states.

Urease (discussed in detail below) catalyzes the hydrolysis of urea to ammonia and

4

carbamate, which then dissociates into another molecule of ammonia and bicarbonate. The
urease active site contains a dinuclear Ni center, with the two metals bridged by a Lys
carbamate (13, 14). The metallocluster is thought to both bind the urea and activate a water
molecule for nucleophilic attack.
[NiFe] hydrogenases catalyze the reversible reduction of protons to H2. The active site
contains both Ni and Fe, with the Ni coordinated by four Cys residues (or in some cases three
Cys and one Se-Cys) while Fe is coordinated by two of the same Cys residues and several
diatomic molecules identified as cyanide or carbon monoxide (15, 16). There is a bridging
ligand as well, with its identity depending on the state of the enzyme. Several other iron-sulfur
clusters typically are present to serve as a conduit for electrons to electron carrier proteins.
The mechanism is still being investigated, although spectroscopic evidence shows the Ni cycles
between +2 and +3 oxidation states.
Carbon monoxide dehydrogenase (CODH) catalyzes the reversible oxidation of CO to
CO2.

The protein contains a catalytic C-cluster with variations on a [Ni-4Fe-5S] cluster,

depending on the source organism (17, 18). Ni is the likely site of CO binding, and one of the Fe
atoms activates a hydroxide to attack the CO to form the product. The two electrons generated
by this oxidative reaction are transferred through other iron sulfur clusters in the protein to
reduce partner electron carrier proteins.
Acetyl-coenzyme A (CoA) synthase (ACS) often is found in a large complex that includes
CODH. In this protein complex, CO2 is reduced by the metallocenter of CODH to form CO and at
a separate active site this toxic intermediate becomes linked to a methyl group, provided by a

5

corrinoid/iron-sulfur protein, and to CoA, thus forming acetyl-CoA (18, 19). The CO-producing
and CO-consuming sites are linked by an ~70 Å long molecular tunnel through the protein
complex. In addition to the Ni cluster in the CODH, the acetyl-CoA synthase active site contains
two Ni atoms as well as a [4Fe-4S] cluster. One relatively stable Ni is bound via two backbone
amides and two Cys residues. The second Ni, easily replaced by Cu or Zn, is coordinated by the
same two Cys along with a third Cys that also serves as a ligand to the iron sulfur cluster. An
analogous protein complex found in acetate-degrading methanogens carries out nearly the
reverse reaction (20). It splits acetyl-CoA into a CO, CoA, and methyl group bound to CoM; in
this case, oxidation of CO to CO2 is used to provide electrons for conversion of methyl-CoM into
methane.

Several mechanisms have been proposed for the C-C and C-S bond

formation/cleavage events, with variations in the order of substrate binding and the oxidation
states of the Ni sites.

6

Figure 1.1: Active sites of Ni-containing enzymes. (A) Ni-Glyoxalase I (1F9Z, Escherichia coli).
The Ni (green sphere) bridges two subunits (carbons colored by subunit) and is bound by two
His and two Glu, with two coordinated waters (red spheres). (B) Acireductone dioxygenase
(1ZRR, NMR structure, Klebsiella oxytoca). Ni is coordinated by three His and one Glu, leaving
two open coordination sites filled by water. (C) Ni-superoxide dismutase (1T6U, Streptomyces
coelicolor). In the oxidized form (shown), the Ni is coordinated by two Cys, the amino-terminal
amine, a backbone amide, and an axial His. In the reduced form, the His swings away, leaving a
square-planar geometry (not shown). (D) Methyl coenzyme M reductase (1MRO,
Methanothermobacter margburgensis). The structure contains a Ni-tetrapyrrole, termed
coenzyme F430, with axial coordination by Gln and coenzyme M. (E) Urease (1FWJ, Klebsiella
aerogenes). Each Ni is coordinated by two His, terminal waters, and, in one case, by Asp, and
the metals are bridged by a lysine carbamate and a hydroxyl group. (F) [NiFe]-Hydrogenase
(1YRQ, Desulfovibrio fructosovorans). The Ni is coordinated by four Cys residues, two of which
also coordinate Fe (that has a carbon monoxide and two cyanide ligands), plus a bridging ligand
whose identity and presence depend on the state of the enzyme. (G) Carbon monoxide
dehydrogenase (CODH, 1SU7, Carboxydothermus hydrogenoformans). The Ni is part of a [Ni4Fe-5S] metallocluster, where the composition varies for proteins from different sources. (H)
Acetyl-CoA synthase (ACS, 2Z8Y, Morella thermocetica). One Ni is bound in square-planar
geometry to two backbone amides and two Cys while a second Ni (shown here replaced by Cu
in blue) is bound to the same two Cys as well as another that is linked to a [4Fe-4S] cluster. A
third Ni (not shown) at a distant site in this protein is in a cluster that functions like and closely
resembles that in CODH. For interpretation the references to color in this and all other figures,
the reader is referred to the electronic version of this dissertation.

7

Ni-binding sites in non-catalytic proteins
Non-catalytic Ni-binding proteins contain metallocenters (Figure 1.2) that function in Ni
sensing, regulation, transport, and delivery (3, 21). These proteins use the same amino acids as
already noted for the Ni-enzymes, but an open coordination site is not required. As with some
enzymes, non-protein ligands may be used.
Ni sensors and regulators
Ni sensors and regulators play important roles in Ni homeostasis by allowing organisms
to maintain constant cellular Ni concentrations through modulation of the transcription of
genes encoding Ni uptake or efflux pumps. From a structural perspective, the best studied
example of such a protein is NikR. This protein is a repressor of the nikABCDE gene cluster
encoding a Ni uptake system in Escherichia coli and other bacteria (22, 23). NikR is a tetramer
consisting of two DNA-binding domains and four metal-binding domains. The NikR tetramer
binds four Ni ions, each coordinated to a Cys and 3 His (one from a different chain) in a squareplanar geometry. NikR from Helicobacter pylori has additional five or six-coordinate Ni binding
sites, and is a global regulator rather than the more specific E. coli homolog (24).
Nur is a Ni-dependent regulator that controls expression of Ni- and Fe- superoxide
dismutases. It uses three His residues to bind one face of Ni while the remaining octahedral
coordination sites can bind other ligands; in the crystal structure those sites are occupied by
malonate and ethylene glycol from the crystallization buffer (25).
The Ni sensor RcnR regulates the Ni efflux transporter RcnA. Although no crystal
structure is available, site-directed mutagenesis and X-ray absorption spectroscopic results
indicate that Ni is coordinated by using 2 His, 1Cys, the amino terminus, a backbone nitrogen,

8

and another unknown ligand (26). Members of the ArsR/SmtB family also regulate Ni related
genes, although the structural details of the metal binding sites are not known (27).
Ni uptake and efflux systems
Much less is known about the structure of Ni binding sites in proteins that import or
export this metal ion. The NikABCDE transport system belongs to the ATP-binding cassette
transporters (28). NikA is a periplasmic protein that binds Ni using a single His residue and an
unidentified organic complex, modeled in the crystal structure as butane-1,2,4-tricarboxylate
(29). NikB and NikC are transmembrane proteins, and NikD and NikE are nucleotide-binding
proteins. The amino acid ligands used to transiently coordinate Ni as it is taken into the cell
remain undefined. In contrast, some potential Ni binding ligands are proposed for the NiCoT
family proteins (transporting Ni, Co, or both, depending on the protein) on the basis of
mutagenesis studies (30). Nevertheless, the mechanism for discriminating between these
metals and the structural details of the metal binding sites are largely unknown (3, 4).
In addition to importing Ni, cells must export this metal ion when in excess. Very little is
known about how the Ni efflux proteins bind the metal. For example, E. coli uses RcnA as a
nickel exporter, however no details about its metal binding site are known.
Metal delivery
Some Ni-containing enzymes require intracellular chaperones to deliver the Ni to the
active site (4, 31). For example, ureases generally require four accessory proteins for enzyme
activation with UreE serving as a metallochaperone that delivers the Ni (32). This protein binds
Ni using His residues which are sometimes localized in His-rich C-termini of the dimeric protein
(33-35). Mutagenesis studies have shown that the His-coordinated Ni binding site at the

9

interface of the UreE dimer (or in the UreE tetramer formed at elevated protein concentrations)
is needed for Ni transfer to the urease apoprotein (36). The UreG and UreD urease accessory
proteins also have been shown to bind Ni with the metal ligands still undefined, and urease
activation may involve a series of Ni transfers (as discussed later).
[NiFe] hydrogenases also require accessory proteins for their assembly. HypA binds one
molecule of Zn, which is likely structural, as well as one Ni atom at a second site, the ligands for
which are unknown (37, 38). HypB also binds Ni and Zn, and has different numbers of metal
binding sites depending on the organism (39-41). Some HypB sequences also have a His-rich
tail that can bind multiple Ni atoms (42). A His-rich metal binding domain of E. coli SlyD is
known to play a role in maturation of its cognate hydrogenase (43, 44) and the Ni-bound
structure is available for SlyD from a thermophilic microorganism (45), however not every SlyD
homolog possesses this domain.
CODH also has accessory proteins necessary for enzyme activation. A CooC-like protein
from Carboxydothermus hydrogenoformans has been shown to bind Ni (46, 47), but the
homolog in Rhodospirillum rubrum did not exhibit detectable Ni binding (48). CooJ contains a
multi-His metal-binding motif at its C-terminus and binds four Ni per monomer, but the
structural details of those binding sites are unknown (49).
Ni-binding sites also are important in proteins that function in extracellular metal
delivery within multi-cellular organisms. Although a structure is not available, extensive studies
have shown how serum albumin binds Ni at its amino terminus, including the use of the amino
terminal amine, two backbone amides, an Asp, and a His (50).

10

Figure 1.2: Ni-binding sites in Ni-sensor or Ni-delivery proteins. (A) E. coli NikR (2HZV) is a
tetrameric Ni-responsive repressor with four high affinity binding sites, each coordinated in
square-planar geometry by three His and one Cys between subunit pairs. (B) NikR from
Helicobacter pylori (3LGH) binds Ni in the same manner as the E. coli protein in some subunits
and forms a distinct five- or six- coordinate site bound to three His in other subunits. (C) Nur
(3EYY, Streptomyces coelicolor) is a Ni-responsive repressor that is structurally distinct from
NikR. The Ni is coordinated by three His, with malonate and ethylene glycol occupying
additional sites in the crystal structure shown. (D) NikA (3DP8, E. coli) is a periplasmic protein
that binds Ni to a single His, additionally using an organic metallophore that was modeled as
butane-1,2,4-tricarboxylate. (E) UreE (3NY0, Helicobacter pylori) is a metallochaperone
involved in urease maturation. The structure shown depicts Ni bound non-symmetrically to five
His derived from four different subunits. (F) SlyD (3CGM, Thermus thermophilus) is a peptidylprolyl isomerase with six His coordinating the metal. The E. coli protein has been implicated in
Ni metabolism during hydrogenase maturation.

11

Ni-substituted proteins
Many proteins bind Ni in place of the native metal (Figure 1.3). The first row transition
elements Fe, Ni, Cu, and Zn all have similar atomic radii and proteins use the same amino acids
to coordinate them. How proteins discriminate between different metals is an open question
in the biochemistry of metals. In peptide deformylase, Ni
but oxygen-labile, Fe

2+

2+

substitutes for the physiological,

and still allows the enzyme to turn over, albeit with reduced kinetics

(51). In other cases, Ni substitution for Fe results in a nonfunctional enzyme, as in the lysyl
hydroxylase JMJD6 and other α-ketoglutarate dependent dioxygenases (52, 53).

Indeed,

substitution of this metal into other members of this enzyme family leads to the hypoxia
2+

mimicking effects of Ni

(54).

2+

2+

DtxR, an Fe -sensitive regulator, also binds Ni

which

promotes its binding to the appropriate sites on DNA in vitro (55). Due to its relatively stable
Ni

2+

redox state, substitution of this metal into Fe or Cu proteins such as rubredoxin (56) or

azurin (57) eliminates their electron transfer activities.

Finally, there are examples of

adventitious Ni-binding sites that are unrelated to protein function, such as the site in ornithine
transcarbamylase that connects three subunits (58). Non-catalytic metal binding sites can
participate in protein stabilization.

12

Figure 1.3: Ni-substituted protein metallocenters. (A) Peptide deformylase (1BS7, E. coli) is an
2+

2+

Fe enzyme, but can also bind Ni using two His and a Cys to form a site capable of catalyzing
the removal of the formyl group from the N-terminus of polypeptide chains. (B) Lysyl
hydroxylase JMJD6 (3K2O, Homo sapiens), an α-ketoglutarate-dependent dioxygenase, is
2+

inhibited by Ni replacing Fe at its active site consisting of a two-His-one-carboxylate motif. (C
and D) DtxR (2TDX, Corynebacterium diphtheriae) regulates genes that encode proteins
important to iron uptake and storage. In vivo it is specific for iron, but nickel and cobalt bind at
two sites (one involving one His, one Met, one Glu, and the side chain and amide of an Asp; the
second comprised of two His and one Glu) in vitro and promote DNA binding. (E) Rubredoxin
2+

(1R0J, Clostridium pasteurianum) contains Fe coordinated by four Cys natively, but the Ni substituted protein has been crystallized. (F) Azurin (1NZR, Pseudomonas aeruginosa) is a Cucontaining electron carrier that was crystallized with Ni bound to two His (including one His
amide), one Cys, and one Met. (G) Ornithine transcarbamylase (2W37, Lactobacillus hilgardii)
binds metals, including Ni, using three His side chains at its three-fold symmetry axis that is
distant from the active site.

13

Ni-binding to His-tagged proteins
tagged
A common purification strategy is to create a fusion between the protein of interest and
a poly-His tag (59). The His-rich region serves as an affinity tag for binding to immobilized
rich
immobilizedmetal ion chromatography beads containing attached Ni (60). This tag, though useful for
.
purification, can cause protein aggregation due to the His residues from multiple proteins
from
coalescing to chelate nickel. Figure 1.4 shows the poly-His tail from three subunits of a tagged
His
protein coming together to bind Ni (61).

Figure 1.4: His-tag used for affinity purification. A commonly used protein purification
tag
procedure is to construct an expression system where the gene of interest is fused to a
where
sequence encoding a poly-His tag, allowing for isolation by immobilized
His
immobilized-metal ion
chromatography. The structure shown (1Q3I, a sodium, potassium ATPase) illustrates the
potassium-ATPase)
mode by which the His region of three tags can aggregate when binding Ni, with each metal ion
aggregate
coordinated by six His.

14

INTRODUCTION TO UREASE
Urease is typically a Ni-containing enzyme (although there is an Fe-containing example
(62)) found in plants, fungi, and bacteria. It was the first enzyme to be crystallized (63), as well
as the first shown to contain Ni (64). It catalyzes the hydrolysis of urea into ammonia and
carbamic acid, which spontaneously decomposes into carbonic acid and ammonia as shown
below.
H2N-C(O)-NH2 + H2O → NH3 + H2N-COOH
H2N-COOH + H2O → NH3 + H2CO3
Urease plays a significant role in several biological processes. Perhaps of greatest
importance, it is a virulence factor in many pathogenic organisms. For example, Helicobacter
pylori utilizes a high concentration of urease (up to 10% of the cellular protein (65)) to produce
ammonia which buffers the acidic environment of the stomach. H. pylori infection, occurring in
approximately half of the world’s population, can result in gastric ulcers in a small percentage
of cases which may develop into carcinomas if left untreated (66). Other organisms (e.g.
Proteus mirabilis) cause infection stones and pyelonephritis due to urease-derived increase in
pH, with consequent precipitation of struvite or apatite crystals, and elevated ammonia
concentrations (67). Urease also is involved in the plant nitrogen cycle, where a variety of
enzymes in plant tissues degrade fixed forms of nitrogen (proteins and nucleotides) to urea,
and then cytoplasmic urease releases ammonia which is used by the plant (68). Finally, urea is
a common fertilizer that is decomposed by plant ureases, but it comes with the inherent

15

problem of potentially uncontrolled hydrolysis by microorganisms in the soil, which can lead to
pH imbalance and ammonia toxicity (68).
Urease structure
Ureases from bacteria and plants have a similar tertiary structure and active site (Figure
1.1E), despite being derived from differing numbers of gene products (Figure 1.5).

K.

aerogenes urease has been well-studied and has a structure typical of many bacterial ureases
(13, 14) The gene products from ureA, ureB, and ureC produce the subunits that form the
urease enzyme (69); ureD, ureE, ureF, and ureG encode accessory proteins needed for
maturation. UreA and UreB are small (11.1 and 11.7 kDa, respectively), whereas UreC, which
contains the active site residues, is much larger (60 kDa). These subunits come together to
form a (UreABC)3 structure (Figure 1.6A) (13). In Helicobacter species, UreA is a fusion of the
two small subunits in K. aerogenes, and UreB is equivalent to K. aerogenes UreC. Helicobacter
proteins UreA and UreB form a homodimer, three of these dimers trimerize, and four of these
trimers create the supramolecular structure of ((UreAB)3)4 (Figure 1.6B) (70). Ureases from
eukaryotes only contain one polypeptide, which is a fusion of all three subunits from K.
aerogenes. These come together to form a trimer, which then dimerizes (Figure 1.6C) (71, 72).

16

Figure 1.5: Genetic structures of the urease gene clusters from K. aerogenes and H. pylori.
Three genes encoding the K. aerogenes enzyme subunits are show in blue, whereas accessory
genes are shown in magenta or, for the gene encoding a metallochaperone, in green. In H.
gene
pylori, sequences corresponding to the above small urease subunit genes are fused into a single
,
gene labeled ureA, the large subunit is named ureB, ureD is renamed ureH and shifted in
,
position, and a proton-gated urea channel gene (
gated
(ureI) is inserted.

A

B

C

Figure 1.6: Urease structures can form different supramolecular structures. A. Cartoon
representation of urease from Klebsiella aerogenes (1FWJ) with UreA in blue, UreB in magenta,
1FWJ)
UreC in yellow, and Ni atoms represented as green spheres. UreABC forms a trimer. B. Urease
from H. pylori (1E9Z) is presented with UreA in blue and UreB in yellow for one trimer of
dimers. This trimer comes together with three more trimers (shades of gray) to form a
together
((UreAB)3)4 structure. C. Jack bean urease (3LA4) is a single polypeptide (shown in blue, yellow,
or maroon). Three of these polypeptides come together to form a trimer similar to the K.
aerogenes structure, which then dimerizes (
(dimer partner in gray, figure is rotated 90°
compared to the other two).
Urease from K. aerogenes can be purified in a Ni-free (apoprotein) form, which also
free
form
lacks the carbamylated Lys residue (73, 74). Under conditions with high levels of nickel and

17

bicarbonate (100 µM and 100 mM, respectively, with the latter needed as a CO2 donor for the
carbamylated lysine at the active site), purified apoprotein can be activated to about 15% of the
fully active enzyme (purified from E. coli expressing the urease operon and supplemented with
Ni) (75). After apoprotein activation there are nearly two Ni atoms per UreC, indicating that
most of the active sites are not properly formed even though they have the correct number of
metal ions (76).
Urease activation
Maturation of urease requires four other genes that typically are found surrounding the
urease structural genes in bacteria: ureD (ureH in Helicobacter species), ureE, ureF, and ureG
(Figure 1.5). Deletions in K. aerogenes ureD, ureF, or ureG expressed in E. coli resulted in
inactive urease; deletions in ureE allowed for partially active urease (69). These genes encode
accessory proteins necessary for delivering the Ni and assembling the active site. The beststudied activation system is that from K. aerogenes, where the accessory proteins have been
purified and characterized individually, as well as in complexes. Figure 1.7 summarizes what is
known about the activation process.

18

Figure 1.7: Interactions among urease-related proteins. The urease apoprotein either
sequentially binds UreD, UreF, and UreG or a preformed UreD—UreF—UreG (abbreviated
UreDFG) complex. In the resulting pre-activation complex, the UreDFG heterotrimer acts as a
GTP-dependent molecular chaperone. UreE serves as a metallochaperone, delivering the Ni
atoms necessary for active site formation to the UreABC—UreDFG complex. CO2 is required to
form the carbamylated lysine metal ligand, and GTP hydrolysis (occurring in UreG) drives the
metallocenter assembly process to form active urease. Finally, the accessory proteins are
released.

19

UreD is insoluble when expressed alone, but a translational fusion with maltose binding
protein (MBP) allowed for the creation of the soluble MBP-UreD (77). MBP-UreD partially
complements a ureD knockout strain, indicating that it can functionally replace UreD. MBPUreD binds ~2.5 Ni (Kd 50 µM) or ~4 Zn (Kd 5 µM) per protomer. This fusion protein forms a
complex with (UreABC)3 when the genes are co-expressed, but surprisingly does not when the
purified proteins are incubated together. Experiments using cell-free extracts of cells coexpressing MBP-UreD with the other accessory proteins demonstrated interactions between
UreD and UreF, UreD and UreG, and UreD, UreF, and UreG. In vitro experiments confirmed the
interaction between MBP-UreD and UreF (as UreE-F) (77). A complex of (UreABC—UreD)3 also
can be purified, and it has greater in vitro activation competence (30% of available active sites)
than the urease apoprotein alone (15%) when incubated with high levels of Ni and bicarbonate
(78). This complex represents the urease apoprotein with UreD binding at the vertices of the
triangular structure according to small angle X-ray scattering (SAXS) (79) and chemical crosslinking (80) experiments. A crystal structure of the UreD homolog from H. pylori (UreH) has
been solved in complex with UreF as a (UreH—UreF)2 heterodimer (Figure 1.8). UreH has a
novel protein fold containing 17 β-sheets and 2 α-helices (81).

20

Figure 1.8: Crystal structure (3FS5) of (UreH
3FS5) (UreH—UreF)2 from H. pylori. The UreF dimer is shown
.
in shades of blue, and UreH is in shades of green.
UreF from H. pylori has been crystallized, both as a C terminal truncation (82) and as a
C-terminal
(UreH—UreF)2 complex (Figure 1.8 (81). This protein crystallizes as a dimer, and its N-terminal
1.8)
N
24 residues are disordered (these residues are not present in K. aerogenes UreF). It has a novel
all alpha-helical fold with highly conserved residues map
mapping to one face of the crystal dimer
(82). A modeling study of UreF from Bacillus pasteruii suggested that it may play a role as a
.
GTPase activating protein (GAP), stimulating the GTPase activity of UreG (83 K. aerogenes
83).
UreF is insoluble when expressed alone; whereas a MalE fusion is soluble, but has not been
n
characterized (84). A translational fusion of ureE and ureF (termed UreE-F) is also soluble and
.
F)
complements a UreF knockout strain, indicating that it is a functional UreF (85). Deletion
mutants demonstrated that the N and C-termini are essential for urease activation, and the
Nhighly conserved C-terminal 15 residues are essential for binding the activation complex. UreEterminal
UreE
F is able to bind (UreABC—UreD)3 in vivo and in vitro (85). A complex of (UreABC—UreD—
UreD)
.
(UreABC
UreF)3 can be purified from cells expressing all five genes; however, no complex is formed

21

when UreD is absent (86). This complex represents the UreD—UreF heterodimer binding to the
vertices of the (UreABC)3 apoprotein according to SAXS analysis (79). In vitro activation of this
species results in the same percentage of active urease as the complex containing only
apoprotein and UreD; however, the complex can be activated under lower concentrations of
bicarbonate due to less nickel-dependent inhibition (86).

Cross-linking studies of the

(UreABC—UreD—UreF)3 complex demonstrated a link between residues in UreB and UreC,
known in the crystal structure to be very distant from each other, leading to the hypothesis that
this complex allows for UreB to undergo a conformational change to allow access to the active
site during the activation process (79, 80).
UreG is soluble when expressed alone, and has been characterized in K. aerogenes, H.
pylori, B. pasteurii, and Mycobacterium tuberculosis (87-91). It belongs to the G3E class of
SIMBI small GTPases, characterized by a signature motif (ESGG at positions 104-107 in K.
aerogenes UreG), as well as a P-loop motif and a guanine specificity loop (92, 93). Other
members of this family include HypB, a structurally-characterized [Ni-Fe] hydrogenase
maturation protein (39), and MeaB, which is involved in delivering vitamin B12 to
methylmalonyl-CoA mutase (94). Despite having a sequence characteristic of a GTPase, UreG
from K. aerogenes and H. pylori do not have any GTPase activity as purified (87, 91); UreG from
B. pasteurii and M. tuberculosis have very low but detectable levels (88, 90). UreG from K.
aerogenes is monomeric (87), whereas the B. pasteruii and M. tuberculosis proteins are dimeric
(89, 90), and H. pylori UreG undergoes zinc-dependent dimerization (91). All UreG proteins are
known to bind metal ions, although the number and binding affinities vary depending on the

22

source organism (88, 90, 91). The UreABC—UreD—UreF—UreG complex from K. aerogenes has
been purified (95). This complex demonstrates GTP-dependent urease activation, and 60% of
the protein can be activated under optimal conditions of GTP, Ni, and bicarbonate
concentrations. Mutations in the P-loop of UreG eliminate the GTP-dependence and have very
low activity in vivo, highlighting the importance of GTP binding for urease activation. GMP-PNP,
a non-hydrolysable GTP analog, does not have the same enhancement effect, indicating that
hydrolysis of GTP is essential for urease activation (95).

No crystal structure is available for

UreG, possibly because of its intrinsically disordered nature according to nuclear magnetic
resonance (NMR), circular dichroism, and fluorescence spectroscopic analysis (89, 96). A crystal
structure for a related protein, HypB from Methanocaldococcus jannaschii, is known (39). This
crystal structure shows a dimeric protein when GTP is bound containing two types of zinc
binding sites: a mononuclear site in each subunit uses residues not conserved in UreG, and a
nonsymmetrical dinuclear binding site at the subunit interface, the ligands for which are
conserved in UreG.
In addition to the individually purified proteins and their complexes with apourease,
several complexes containing UreD—UreF—UreG have been isolated. When the ureD, ureF,
and ureG genes are expressed together, a predominantly insoluble UreD—UreF—UreG complex
is formed. This complex can be partially solubilized by low concentrations of detergent and
purified using ATP-linked agarose (87). The MBP-UreD—UreF—UreG complex is soluble, has
been purified, and forms a complex with (UreABC)3 in vitro (77, 97).

These complexes

demonstrate that UreD, UreF, and UreG may not bind sequentially to urease, but instead may
assemble into a pre-formed complex before binding to apourease.

23

UreE is the metallochaperone that delivers the Ni ions to the activation complex. The
etallochaperone
structures of a C-terminal truncation form of the protein from K. aerogenes (35 and the fullterminal
35),
length proteins from B. pasteruii (33) and H. pylori (34) have been solved with bound Cu, Zn,
and Ni respectively (Figure 1.2E, 1.9 In K. aerogenes, UreE binds approximately six Ni atoms
1.9).
per homodimer, mostly utilizing a His rich tail that is not essential for urease activation (98).
His-rich
Using a truncation mutant lacking the His
His-rich C-terminus (UreE H144*), it was shown that two
terminus
other His residues, 110 and 112, participate in binding Ni but these sites are not necessary for
,
Ni,
urease activation (36). By contrast, His 96 plays a critical role in transferring the metal to the
active site (99). Importantly, when UreE is incubated with the UreABC UreD—UreF—UreG
.
UreABC—UreD
complex under biologically relevant concentrations of Ni, bicarbonate, and GTP, urease activity
reaches wild-type levels (100).

Figure 1.9: Crystal structure (1GMW) of truncated UreE from K. aerogenes. Protomers are
1GMW)
.
shown in purple and green. Key Ni
Ni-binding residue His96 in each subunit is shown in yellow.
Eukaryotic urease activation
The mechanism of urease activation has been extensively investigated in the case of
he
bacterial ureases; however, many fungi and plants also possess this enzyme which must

24

undergo analogous activation processes. Genetic and biochemical studies related to urease
activation in eukaryotes are summarized below.
Fungal ureases contain a single type of subunit, but genetic studies reveal that multiple
genes are needed for urease expression. For example, early studies with Neurospora crassa
(101-103), Aspergillus nidulans (104, 105), and Schizosaccharomyces pombe (106) show that
four distinct loci are required for obtaining active enzyme. The best studied fungal urease
system is that in the fission yeast S. pombe where the enzyme was purified (107), the structural
gene (ure2) identified (108), and candidate genes encoding UreD (ure4), UreF (ure3), and UreG
(ure1) were identified (109). Of interest, the fungal UreG has 61% sequence identity to soybean
UreG (see below) including the presence of a His-rich N-terminus that is not found in bacterial
UreG sequences. The fungal UreF protein of S. pombe shares only 20% sequence identity with
the soybean protein, but the plant gene rescues the corresponding yeast mutant, whereas this
situation is not observed for UreD where the proteins share 30% sequence identity (109).
Plants can possess multiple urease isozymes as well as genes encoding several
maturation proteins.

The historically interesting jack bean (Canavalia ensiformis) system

includes two structural urease genes (110, 111), but no accessory genes have yet been
reported. The better-studied soybean (Glycine max) system also has two urease isozymes: an
embryo-specific form encoded at the Eu1 locus (112, 113) and a ubiquitously-expressed species
encoded at the Eu4 locus (114, 115). In addition, this plant contains several demonstrated
urease accessory genes. For example, soybean genes encoding UreD and UreF are orthologues
of the bacterial and fungal genes; in the latter case, the plant gene complements a mutant
involving the corresponding gene in S. pombe (109). The Eu3 locus was long known to exhibit

25

pleiotropic effects on both ureases (113), and more recently was shown to encode a protein
related to bacterial UreG proteins with a conserved P-loop motif and an added His-rich Nterminus that resembles the C-terminus of some bacterial UreE proteins and is probably
involved in nickel binding (116). This UreG was purified from seeds and as a recombinant
protein. It binds Zn very tightly, and Ni with less affinity. Zn also stabilizes a dimeric form of the
protein. As with bacterial UreGs, this UreG is intrinsically disordered according to NMR (117).
Eu2 encodes another protein necessary for activation of both ureases (113). This gene,
analogous to bacterial ureF, was duplicated and found on chromosome 2 and chromosome 14,
with the Ch2 copy being the dominant one (118). Homologues encoding UreD-, UreF-, and
UreG-like proteins are now known to exist in many other plant species, including tomato
(Lycopersicon esculentum), potato (Solanum tuberosum), and Arabidopsis thaliana (109, 119,
120). Of interest, the gene encoding potato UreG complements a K. aerogenes ureG mutation
(119) and insertions into each of the three accessory genes of A. thaliana abolished urease
activity (120).

26

REMAINING QUESTIONS
Even though the accessory proteins involved in urease activation are known, the details
of how they work together to assemble the active site are still elusive. There is no clear picture
of how UreD, UreF, UreG and UreE interact, or how the accessory proteins bind to (UreABC)3.
UreD and UreG bind Ni, but the ligands they use and how Ni is transferred from UreE into the
active site of urease remains elusive. One hypothesis is that UreE transfers the Ni to UreG,
which shifts it to UreD, and then into the active site (121). There are several possible roles for
GTPase activity in the activation process that need to be tested. It could be utilized to activate
bicarbonate for lysine carbamylation (as stated in several publications (117, 122), but without
any evidence), it might be needed for a series of metal transfers, it may play a role in “gating”
the active site from incorrect metal incorporation, or it could be needed for dissociation of the
accessory proteins after the activation is complete.
This dissertation seeks to answer some of the remaining questions about the functions of UreG
and UreF. In chapter 2, I present my work on UreG that describes the Ni- and Zn-binding
properties of wild-type and Strep-tagged versions of the protein, along with mutants involving
potential metal-binding residues. I also examine these UreG variants for their abilities to
disrupt the binding to the other urease proteins in cell-free extracts. Additionally I investigate
the Ni-dependent interaction between UreG and UreE. In chapter 3 I move to UreF, first
elucidating the roles played by conserved residues in protein:protein interactions, then looking
into UreF’s proposed role as a GTPase activating protein (GAP) for UreG by focusing on the
conserved residue Lys165. Finally, in chapter 4 I describe some additional studies I have done,
including my role in a collaborative effort trying to understand the role of UreB and

27

characterization of the complexes MBP-UreD—UreF—UreG and UreABC—UreD—UreF—
UreGStr.

28

REFERENCES

29

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39

121.

Boer, J. L., Quiroz-Valenzuela, S., Anderson, K. L., and Hausinger, R. P. (2010)
Mutagenesis of Klebsiella aerogenes UreG to probe nickel binding and interactions with
other urease-related proteins, Biochemistry 49, 5859-5869.

122.

Zambelli, B., Musiani, F., Benini, S., and Ciurli, S. (2011) Chemistry of Ni
Sensing, Trafficking, and Catalysis, Acc. Chem. Res. 44, 520-530.

2+

40

in Urease:

Chapter 2

Mutagenesis of Klebsiella aerogenes UreG to Probe Nickel Binding and Interactions with
Other Urease-Related Proteins

This chapter is adapted from Boer, J. L., Quiroz-Valenzuela, S., Anderson, K. L., and Hausinger, R.
P. (2010) Mutagenesis of Klebsiella aerogenes UreG to probe nickel binding and interactions
with other urease-related proteins, Biochemistry 49, 5859-5869.

The cloning, initial purification, and partial characterization of UreGStr using circular dichroism
spectroscopy, inductively coupled plasma-emission spectroscopy, pull-down studies with cellfree extracts and western blot analysis were performed by Soledad Quiroz-Valenzuela and
Kimberly L. Anderson.

41

ABSTRACT
UreG is a GTPase required for assembly of the nickel-containing active site of urease.
Herein, a Strep-tagged Klebsiella aerogenes UreG (UreGStr) and selected site-directed variants
of UreGStr were constructed for studying the in vivo effects on urease activation in recombinant
Escherichia coli cells, characterizing properties of the purified proteins, and analysis of proteinprotein interactions using cell-free extracts and in vitro. Whereas the Strep-tag had no effect
on UreG’s ability to activate urease, enzyme activity was essentially abolished in the K20A,
D49A, C72A, H74A, D80A, and S111A UreGStr variants, with diminished activity also noted with
E25A, C28A, and S115A proteins. Lys20 and Asp49 are likely to function in binding/hydrolysis of
GTP and binding of Mg, respectively. UreGStr binds one nickel or zinc ion per monomer (Kd = ~5
μM for each metal ion) at a binding site that includes Cys72, as shown by a 12-fold increased Kd
for nickel ions using C72A UreGStr and by a thiolate-to-nickel charge-transfer band that is
absent in the mutant protein. Based on UreG homology to HypB, a GTPase needed for
hydrogenase assembly, along with the mutation results, His74 is likely to be an additional metal
ligand. Pull-down assays in cell-free extracts revealed Asp80 as critical for stabilizing UreGStr
interaction with the UreABC—UreD—UreF complex. In vitro pull-down assays demonstrated
UreG binding to UreE, with the interaction enhanced by nickel or zinc ions.

The

metallochaperone UreE is suggested to transfer its bound nickel to UreG in the UreABC—
UreD—UreF—UreG complex, with the metal ion subsequently transferring to UreD, and then
into the nascent active site of urease in a GTP-dependent process.

42

INTRODUCTION
Urease, a nickel-containing enzyme found in plants and microorganisms, catalyzes the
hydrolysis of urea to form ammonia and carbamate, which spontaneously decomposes to carbon
dioxide and ammonia (1, 2).

Structures of several ureases (3-6) reveal dinuclear nickel

metallocenters deeply buried in structural subunits that coalesce with three-fold symmetry. With
the possible exception of the Bacillus subtilis enzyme (7), activation of urease has been shown to
require a series of accessory proteins to assemble the active site (1, 8). The best understood
urease activation system involves the ureDABCEFG genes of the enterobacterium Klebsiella
aerogenes expressed in Escherichia coli. This model urease system begins with the structural
subunits (UreA, UreB, and UreC) assembling into the urease apoprotein (UreABC)3 (9, 10), with
UreD, UreF, and UreG sequentially associating with the apoprotein to form the (UreABC—UreD)3
(11), (UreABC—UreD—UreF)3 (12), and (UreABC—UreD—UreF—UreG)3 (13) activation complexes
(Figure 2.1). Finally, in a process that requires GTP hydrolysis, CO2 incorporation as an active site
lysine carbamate, and the nickel-delivering metallochaperone UreE, the active site assembles, and
then the accessory proteins release from the active enzyme (14, 15). As described below, the
roles of UreD, UreF, and UreG in urease activation are poorly understood.

43

Figure 2.1: Simplified scheme of the urease activation process. Urease apoprotein (UreABC)3 is
synthesized with the nascent active site lacking nickel ions and with no carbamylation of its Lys217.
Urease accessory proteins UreD, UreF, and UreG bind the apoprotein in a sequential manner to
form the (UreABC—UreD—UreF—UreG)3 activation complex (abbreviated further for simplicity in
the figure). Urease activation requires carbamylation of Lys217 by CO2, provision of nickel ions by
the UreE metallochaperone, and GTP hydrolysis accompanied by release of the accessory proteins.

44

Studies of UreD are limited to the K. aerogenes system, along with a structure (3sf5) of
the H. pylori UreD homolog UreH in complex with its cognate UreF (16). The K. aerogenes
protein is insoluble when expressed alone, however a maltose binding protein (MBP)-UreD
fusion is soluble and complements a ΔureD urease cluster (17). Significantly, the UreD portion
of MBP-UreD binds nickel (~2.5 Ni per protomer, Kd ~50 µM) and this protein, when in the
UreABC—UreD—UreF—UreG complex, is proposed to transfer the metal ion into the nascent
urease active site.
K. aerogenes UreF, like UreD, is insoluble when synthesized separately from the other
urease components; however, UreE-UreF and MBP-UreF fusion proteins are soluble and
partially characterized (18, 19). In addition, two crystal structures of UreF from Helicobacter
pylori (PDB codes 3cxn and 2wgl) were solved (20) along with the UreF-UreH complex (16).
Computational studies of Bacillus pasteurii UreF led to a proposal that the protein functions as
a GTPase activating protein (21), but no direct evidence for such a role has been reported in any
system.
Purified recombinant UreG proteins (subunit Mr 22,000 - 23,000) of K. aerogenes, B.
pasteurii, Mycobacterium tuberculosis, and H. pylori are soluble and contain motifs found in
GTPases, although their GTPase activities are very low or non-detectable (13, 22-24). Mutation
of Lys20 or Thr21 in the GXGKT P-loop motif (a GTPase motif) of the K. aerogenes protein
abolishes its ability to activate urease (13). This region also is critical for in vitro activation of
the (UreABC—UreD—UreF—UreG)3 complex (14).

45

K. aerogenes UreG is reported to be

monomeric (13). In contrast, UreG proteins from B. pasteurii and M. tuberculosis are dimeric,
with the subunits joined by a disulfide bridge involving Cys68 in the B. pasteurii protein and
probably Cys90 in that from M. tuberculosis (23, 25). UreG from B. pasteurii binds two zinc ions
per dimer (Kd 42 μM) or four nickel ions per dimer (Kd 360 μM), and this interaction was
speculated to involve Glu64, Cys68 (i.e., the same residue as that participating in the disulfide),
and His70 as metal ligands (22), although no experiments were performed to confirm these
assignments. H. pylori UreG, a monomer as purified, dimerizes as it binds zinc ions (1.0 Zn per
dimer, Kd 0.33 µM) or remains a monomer as it binds nickel ions with lower affinity (2.0 Ni per
monomer, Kd 10 µM) (24). Cys66 and His68 are proposed as ligands for the zinc ion-binding
site, but the C66A, H68A, and C66A/H68A double mutant still binds zinc with only 10-fold lower
affinity and these mutant proteins still dimerize upon addition of the metal ions. Furthermore,
the presence of zinc, but not nickel, ions stabilizes a UreE-UreG complex using the H. pylori
proteins (26). No crystal structure is available for any UreG; however, the crystal structure of
the related protein HypB from Methanocaldococcus jannaschii is known (27). HypB is an
accessory protein that participates in the metallocenter assembly of [NiFe] hydrogenases (8,
28). The crystal structure reveals two types of zinc binding sites: a mononuclear site in each
subunit involving His100 and His104 (numbering derived from the HypB crystal structure;
corresponding His residues are not located at these positions in UreG sequences) and a nonsymmetrical dinuclear binding site at the subunit interface. The metal-binding residues of the
dinuclear site in M. jannaschii HypB (Cys95, His96, and Cys127) most likely correspond to Cys72,
His74, and either Ser111 or Ser115 (although Ser is not a typical metal-binding residue) in K.

46

aerogenes UreG (or Cys68, His70, and Ser107 or Ser111 in the B. pasteurii protein) (Figure 2.2).

Ka UreG
Bp UreG
Mj HypB

Ka UreG
Bp UreG
Mj HypB

Ka UreG
Bp UreG
Mj HypB

Ka UreG
Bp UreG
Mj HypB

Ka UreG
UreG
Bp UreG
Mj HypB

10
20
--------------------------MNSYKHPLRVGVGGPVGSGKTALL 24
.
.
------------------------------MKTIHLGIGGPVGSGKTTLV 20
.
.
.
.
MHLVGVLDIAKDILKANKRLADKNRKLLNKHGVVAFDFMGAIGSGKTLLI 50
30
40
50
60
70
EALCKAMRDTWQLAVVTNDIYTKEDQRILTEAGALAPERIVGVETGGCPH 74
.
.
.
.
.
KTLSEALKEEYSIAVITNDIYTREDANFLINENILEKDRIIGVETGGCPH 70
.
.
.
.
EKLIDNLKDKYKIACIAGDVIAKFDAERMEK----HGAKVVPLNTGKECH 96
80
90
100
110
120
TAIREDASMNLAAVEALSEKFGNLDLIFVESGGDNLSATFSPELADLTIY 124
.
.
.
.
.
TAIREDASMNFEAIEELKNRFDDLEIILLESGGDNLSATFSPELVDAFIY 110
.
.
.
.
.
LDAHLVG--HALEDLNLDEI----DLLFIENVGNLICPADFDLGTHKRIV 140
130
140
150
160
170
VIDVAEGEKIPRKGGPGITKSDFLVINKTDLAPYVGASLEVMASDTQRMR 174
.
.
.
.
.
VIDVSEGGDIPRKGGPGVTRSDFLMVNKTELAPYVGVDLDTMKNDTIKAR 160
.
.
.
.
.
VISTTEGDDTIEKHPGIMKTADLIVINKIDLADAVGADIKKMENDAKRIN 190
180
190
200
GDRPWTFTNLKQGDGLSTIIAFLEDKGMLGK
.
.
.
.
NGRPFTFANIKTKKGLDEIIAWIKSDLLLEGKTNESASESK
.
.
.
PDAEVVLLSLKTMEGFDKVLEFIEKSVKEVK

205
201
220

Figure 2.2: Multiple sequence alignment of K. aerogenes UreG, B. pasteurii UreG, and M.
jannaschii HypB. Clustal W (44) was used to make the initial alignment, followed by manual
modifications. Residues mutated in K. aerogenes UreG and the corresponding residues in the
other sequences are highlighted in yellow. The P-loop motif, signature motif of the SIMBI G3E
family, and guanine specificity loop are underlined.
UreE serves as a metallochaperone that delivers the nickel ions needed to form the
urease active site (29, 30). The structures of a truncated version of UreE from K. aerogenes and

47

the full-length protein from B. pasteurii have been solved with bound copper and zinc,
respectively (31, 32). The interaction of UreE with the other accessory proteins has not been
well characterized; however, UreE and UreG from H. pylori were suggested to interact on the
basis of yeast two-hybrid assays (33) and a UreE2UreG2 complex (formed with the isolated H.
pylori proteins) was observed in the presence of zinc, but not nickel, ions (26).
In this study, I describe a new purification method for UreG that utilizes a Strep-tag.
Using protein purified by this approach, I examine the metal binding capabilities of UreGStr and
a selection of its variants. Additionally, I assess the effects of those mutations on urease
activation and exploit the Strep-tagged protein to examine its interactions with other urease
components. These findings using the K. aerogenes urease activation system expressed in E.
coli reveal significant new insights, many of which are likely to be more generally applicable to
other urease systems.

48

EXPERIMENTAL PROCEDURES
Vector Construction, Cell Growth, and Purification of Strep-Tagged UreG. The ureG
sequence was cloned into pASK-IBA3plus and pASK-IBA5plus plasmids (IBA GmbH, Göttingen,
Germany) to create vectors pIBA3+G and pIBA5+G (Table 2.1) encoding UreG with a Strep-tag II
(a WSHPQFEK peptide; subsequently referred to as a Strep-tag) at the C- or N-termini,
respectively. First, a polymerase chain reaction (PCR) was performed using PfuTurbo® Hotstart
PCR Master Mix (Stratagene, USA), the plasmid pKAUG-1 as a template, and the primers 5’-TA
CTG TCC CGC GGG ATG AAC TCT TAT AAA CAC-3’ and 5’-T ACT GTC CTG CAG TTT GCC AAG CAT
GCC TTT-3’. The first primer contains a SacII restriction site and the second a PstI restriction
site (shown in italics) used to clone the fragment into pASK-IBA3plus. In a similar manner, the
primers 5’-T ACT GTC CCG CGG GG AAC TCT TAT AAA CAC CCG-3’ and 5’- T ACT GTC GGA TCC
CTA TTT GCC AAG CAT GCC-3’, containing restriction sites for SacII and BamHI respectively,
were used to clone the fragment into pASK-IBA5plus. The plasmids and PCR products were
digested with the corresponding restriction enzymes (New England Biolabs) and ligated to
produce plasmids pIBA3+G and pIBA5+G. These constructions were confirmed by sequencing
(Davis sequencing, Davis, CA, USA).

49

Table 2.1: Plasmids and E. coli strains used in this study
Plasmid
Description
pASK-IBA3plus

pEC007
pIBA3+G

Plasmid for creating fusion proteins
with a Strep-tag II (WSHPQFEK) at the
C-terminus
Plasmid for creating fusion proteins
with a Strep-tag II at the N-terminus
Modified pACT3 to encode UreE
Modified pASK-IBA3plus to encode

pIBA5+G

UreGStr
Modified pASK-IBA5plus to encode

pASK-IBA5plus

UreGStr
pIBA3+GK20A,
pIBA3+GE25A,
pIBA3+GC28A,
pIBA3+GD33A,
pIBA3+GD49A,
pIBA3+GE68A,
pIBA3+GC72A,
pIBA3+GH74A,
pIBA3+GD80A,
pIBA3+GS111A,
pIBA3+GS115A,
pIBA3+GD120A
pIBA3+GD127A
pKAU17
pKAUD2
pKAUG-1
pKK17
pKKG
pKKGK20A, pKKGE25A,
pKKGC28A, pKKGD33A,
pKKGD49A, pKKGE68A,
pKKGC72A, pKKGH74A,
pKKGD80A, pKKGS111A,
pKKGS115A, pKKGD127A

Modified pIBA3+G to encode the
K20A, E25A, C28A, D33A, D49A, E68A,
C72A, H74A, D80A, S111A, S115A,

E. coli
strain

Reference
IBA

IBA
DH5α
DH5α or
BL21 (DE3)

(17)
This work

DH5α or
BL21 (DE3)

This work

BL21 (DE3)

This work

D120A and D127A variants of UreGStr

K. aerogenes urease gene cluster in
pUC8
pKAU17 modified to overexpress ureD
Modified pKAUD2 containing only K.
BL21 (DE3)
aerogenes ureG.
K. aerogenes ureDABCEFG gene
DH5α
cluster inserted into pKK223-3
DH5α
Modified pKK17 encoding UreGStr
Modified pKKG encoding the K20A,
E25A, C28A, D33A, D49A, E68A,
C72A, H74A, H74C, H74N, D80A,
S111A, S115A, and D127A variants of
UreGStr

50

DH5α

(34)
(11)
(35)
(36)
This work
This work

Isolated colonies of E. coli BL21(DE3) (Stratagene) were transformed with the plasmids
and grown at 37 °C overnight in lysogeny broth (LB, or Lennox broth, Fisher Scientific)
supplemented with 300 μg mL

-1

supplemented with 300 μg mL

-1

of ampicillin. These cultures were used to inoculate 1 L of LB
of ampicillin. The cultures were grown at 37 °C with shaking

for 4 h and induced overnight with 0.2 µg mL

-1

anhydrotetracycline. The cells were harvested

by centrifugation and resuspended in 1 mL of buffer W (100 mM Tris-HCl, pH 8.0, containing
150 mM NaCl and 1 mM EDTA) per gram of cells and supplemented with 1 mM
phenylmethylsulphonyl fluoride as a protease inhibitor before sonication (Branson 450 sonifier,
5 repetitions, each of 2 min, at 3 W output power and 50% duty cycle). The disrupted cells
were centrifuged at 100,000 x g at 4 °C for 45 min and the cell-free supernatant was loaded
onto a 1 mL Strep-Tactin column (IBA, Germany) previously equilibrated in buffer W. This
column has an engineered streptavidin ligand that binds to the Strep-tag with high affinity. The
Strep-tagged UreG protein (UreGStr) was eluted with desthiobiotin according to the
manufacturer’s instructions. For comparative studies, native UreG was purified as previously
described (13). For further purification and to provide assurance that samples were completely
-1

reduced, the proteins were chromatographed at 1 mL min on a preparative Superdex-75 column
(65 cm x 2.0 cm diam., GE Healthcare) equilibrated in 50 mM HEPES buffer, pH 7.4, containing 200
mM NaCl, 1 mM EDTA and 1 mM dithiothreitol (DTT).
Fractions containing UreGStr, UreG, or mutant forms of these proteins were analyzed by
sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) (37) using gels prepared

51

with 12% running and 5% stacking acrylamide sections and stained with Coomassie brilliant
blue. The calculated molecular weights of UreA (11.1-kDa), UreB (11.7-kDa), UreE (17.6-kDa),
UreG (21.9-kDa), UreGStr (23.1-kDa), UreF (25.2-kDa), UreD (29.8-kDa), and UreC (60.3-kDa)
generally migrate during electrophoresis as expected with the exception of UreG and UreGStr
which behave as if they are larger than UreF. Molecular weight markers were obtained from
Bio-Rad (Hercules, CA). Protein concentrations were determined by using a commercial dyebinding assay (Bio-Rad, Hercules, CA) with bovine serum albumin as the standard.
UreE Purification. E. coli DH5α cells containing pEC007 (17), expressing full length UreE,
-1

were grown overnight in 10 mL LB supplemented with 50 µg mL

chloramphenicol. These

cultures were used to inoculate 1L of LB supplemented with 50 µg mL

-1

chloramphenicol and

grown to an optical density at 600 nm (O.D.600) of 0.4, induced with 0.5 mM isopropyl β-D-1thiogalactopyranoside (IPTG), and grown overnight at 37 °C. UreE was purified by using
previously published protocols (38).
Site-Directed Mutagenesis. pIBA3+G was mutated by using overlapping oligonucleotides
containing the desired mutation (see Table 2.2) during PCR performed with PfuTurbo® Hotstart
PCR Master Mix. The products were digested with DpnI for 1 h at 37 °C and used to transform
chemically competent E. coli DH5α cells. After confirmation by sequencing, the mutated
plasmids were transformed into E. coli BL21(DE3) competent cells (Stratagene, USA). All
mutant UreGStr proteins were expressed and purified as described for UreGStr.

52

Table 2.2: Oligonucleotides used to generate UreG mutations
Purpose
Sequence
UreG mutation E25A
5' GTA AAA CCG CTC TGC TGG CGG CGC TGT GTA AAG CGA TG 3'
5' CAT CGC TTT ACA CAG CGC CGC CAG CAG AGC GGT TTT AC 3'
UreG mutation C28A
5' CT CTG CTG GAA GCG CTG GCA AAA GCG ATG CGC GAT AC 3'
5' GT ATC GCG CAT CGC TTT TGC CAG CGC TTC CAG CAG AG 3'
UreG mutation D33A
5' GTA AAG CGA TGC GCG CGA CCT GGC AGC TGG C 3'
5' G CCA GCT GCC AGGTCG CGC GCA TCG CTT TAC 3'
UreG mutation D49A
5’ GAC ATC TAT ACC AAA GAA GCG CAG CGC ATC CTC ACC GAA 3’
UreG mutation E68A
5’ GAA CGC ATC GTC GGT GTG GCG ACC GGC GGC TGC CCG CAT 3’
UreG mutation C72A
5’ GTC GGT GTG GAA ACC GGC GGC GCG CCG CAT ACG GCG ATC
CGC GAA 3’
UreG mutation H74A
5’ GAA ACC GGC GGC TGC CCG GCA ACG GCG ATC CGC GAA GAT 3’
UreG mutation H74C
5’ GAA ACC GGC GGC TGC CCG TGC ACG GCG ATC CGC GAA GAT 3’
UreG mutation H74N
5’ GAA ACC GGC GGC TGC CCG AAT ACG GCG ATC CGC GAA GAT 3’
UreG mutation D80A
5’ CAT ACG GCG ATC CGC GAA GCG GCC TCA ATG AAC CTC GCC 3’
UreG mutation S111A 5' GAA AGC GGC GGC GAT AAC CTG GCC GCC ACC TTC AGC CCG GAG
CTG 3'
5' CAG CTC CGG GCT GAA GGT GGC GGC CAG GTT ATC GCC GCC GCT
TTC 3'
UreG mutation S115A 5' AAC CTG AGC GCC ACC TTC GCC CCG GAG CTG GCG GAT CTG 3'
5' CAG ATC CGC CAG CTC CGG GGC GAA GGT GGC GCT CAG GTT 3'
UreG mutation D120A 5' C CGG AGC TGG CGG CGC TGA CCA TCT AC 3'
5' GT AGA TGG TCA GCG CCG CCA GCT CCG G 3'
UreG mutation D127A 5' CA TCT ACG TCA TCG CGG TGG CCG AAG GGG AG 3'
5' CT CCC CTT CGG CCA CCG CGA TGA CGT AGA TG 3'

Circular Dichroism (CD). Wild-type and Strep-tagged UreG proteins were purified and
concentrated up to 0.2 mg mL

-1

in 15 mM potassium phosphate buffer, pH 7.6, containing 1

mM DTT. A 100 μL sample was placed into a 1 cm path length cell and scanned using a Jasco J710 spectropolarimeter between 180 and 300 nm.

The data were analyzed with the

DICHROWEB server (39), and the best fit was obtained by using CDSSTR and set 4.
Analytical Gel Filtration Chromatography. Analytical hydrodynamic radius assays used
Sephacryl 300 HR (65 cm x 2.0 cm diam., Sigma). The buffer contained 50 mM HEPES, pH 7.4,
-1

with 200 mM NaCl and other additives as indicated, using a flow rate of 1 mL min .

53

In vivo Expression of UreGStr Variants in the Context of the Urease Operon. Plasmid
pKK17 (29), which contains the entire ureDABCEFG urease gene cluster under the control of the
tac promoter, was modified to encode UreGStr and its mutant forms by replacing a PsiI/KpnI
fragment to create plasmid pKKG and variants. For analysis of urease activity in cell extracts, E.
coli DH5α containing the desired plasmid was inoculated into 1 mL of LB supplemented with
-1

300 μg mL

of ampicillin and 1 mM NiCl2 (unless noted) and grown overnight at 37 °C with

agitation. A 0.25 mL aliquot of the culture was used to inoculate 25 mL of LB containing 100 μg
mL

-1

ampicillin plus 1 mM NiCl2 (unless noted) and grown for 2.5 h at 37 °C with agitation.

IPTG added to 0.1 mM was used to induce the expression of the operon overnight at 37 °C.
Cells were harvested by centrifugation for 10 min at 5,000 x g and 4 °C and resuspended in
either 1 mL of 25 mM HEPES buffer, pH 7.4, if performing urease activity assays or 750 µl of
buffer W if used for pull-down assays. Phenylmethylsulphonyl fluoride was added to 0.1 mM,
the cells were sonicated (Branson 450 sonifier, 5 repetitions, each of 45 sec, at 1 W output
power and 50% duty cycle), and the disrupted cells were centrifuged 10 min at 4 °C and 16,000
x g in a microcentrifuge. The soluble, cell-free extracts were used to test urease activity and
perform pull-down assays.
Urease Activity Assays. Urease activities were measured by quantifying the rate of
ammonia release from urea by formation of indophenol, which was monitored at 625 nm (40).
One unit of urease activity was defined as the amount of enzyme required to hydrolyze 1 µmole
of urea per min at 37 °C. The standard assay buffer consisted of 50 mM HEPES, pH 7.8, and 50
mM urea.

54

Metal Quantification. The metal contents of freshly purified UreG and UreGStr were
assessed by using inductively coupled plasma-emission spectrometry at the University of
Georgia Chemical Analysis Laboratory.
Metal Binding Analyses. Purified proteins were dialyzed overnight against 50 mM HEPES
buffer, pH 7.4, containing 200 mM NaCl, 1 mM EDTA, and 1 mM DTT, followed by dialysis
against 50 mM HEPES buffer, pH 7.4, containing 200 mM NaCl until the EDTA and DTT
concentrations were negligible.

Non-radioactive equilibrium dialysis experiments were

performed by using an equilibrium micro-volume dialyzer (Hoefer Scientific instruments).
Purified protein (400 µL of 10 µM) was dialyzed against 400 µL of various concentrations of
NiCl2 or ZnCl2 overnight at 4 °C by using a 3,500 Da molecular weight cut off membrane
(MWCO, Spectra-Por). Metal concentrations on both sides of the membrane were determined
by adding 100 µL of these solutions to 900 µL of 100 µM 4-(2-pyridylazo) resorcinol (PAR) made
in 50 mM HEPES (pH 7.4) with 200 mM NaCl, incubating for 10 min, and monitoring the
absorbance at 500 nm (41). The data were plotted and analyzed in Sigma Plot (Systat Software,
Inc.) by using Eq. 1, appropriate for samples containing a single type of binding site, where Y is
the number of metal ions bound per UreG subunit, Bmax is the maximum number of metal ions
bound per UreG peptide, [Mf] is the concentration of free metal ions, and Kd is the dissociation
constant.
Y = Bmax[Mf]/(Kd + [Mf])

(1)

Metal competition experiments were performed in a Rapid Equilibrium Dialysis plate (Pierce

55

Biotechnology, Rockford, IL). Purified UreGStr (300 µL of 25 µM) was dialyzed against 500 µL of
varying concentrations of nickel ions containing

63

Ni and the indicated concentrations of ZnCl2,

with shaking overnight at 5 °C and 300 rpm. Aliquots of the resulting samples (200 µL) were
added to 10 mL of Safety Solve (Research Products International Corp.) and

63

Ni contents were

determined by using a Beckman-Coulter LS6500 liquid scintillation counter. The data were fit
by using the following equation for competitive binding to a single type of binding site:
Y = Bmax[Ni]/{Kd(1 + [Zn]/Ki) + [Ni]}

(2)

The constants are as indicated above, and Ki is the inhibition constant for Zn.
UV/Visible Spectroscopy. Samples (1 mL) of the indicated concentrations of UreGStr,
C28A UreGStr, and C72A UreGStr in 50 mM HEPES, pH 7.4, containing 200 mM NaCl were
titrated with aliquots of 1-5 µL of 1 mM NiCl2. Absorption spectra were obtained after each
addition, and these were corrected for dilution.
Pull-Down Assays. Soluble cell-free extracts from E. coli DH5α containing pKKG grown
with and without supplemented Ni were loaded onto a 0.3 mL Strep-Tactin column equilibrated
in buffer W. Proteins were eluted according to the manufacturer’s instructions and analyzed by
using 13.5% SDS-PAGE.
For in vitro pull-down assays, UreE and UreGStr or variants were mixed in a final
concentration of 16 µM for each protomer with varying concentrations of NiCl2 or ZnCl2,
incubated on ice as indicated, applied to a 0.5 mL Strep-Tactin column, washed, and eluted

56

according to the manufacturer’s instructions. Eluted fractions were analyzed by using 12% SDSPAGE. Further analysis of the interaction between UreE and UreGStr was carried out by mixing
equal concentrations of each protein (40 or 150 µM protomer) and subjecting the mixture to
chromatography on Sephacryl S-300 in buffer containing or lacking 60 µM NiCl2.
Western Blot. Proteins were resolved by SDS-PAGE and transferred to an Immobilon-P
polyvinylidene difluoride membrane (Millipore, USA).

ExtrAvidin®–alkaline phosphatase

conjugate (1:2500 dilution, Sigma, USA) was used as a probe to bind to Strep-tagged forms of
UreG. BCIP®/NBT-Blue Liquid Substrate (Sigma, USA) was added to develop the color. To
detect UreE or urease, the membranes were incubated for 45 min with anti-UreE IgG (1:10,000
dilution) (38) or anti-urease antibody (1:5,000 dilution) (34) in TBS buffer (150 mM NaCl, 100
mM Tris, pH 7.4) containing 1% Tween 20. After washing the membranes four times with TBS
buffer, they were incubated for 45 min with anti-rabbit IgG conjugated to alkaline phosphatase
(Sigma, USA) that was diluted 30,000-fold. The membranes were washed again and BCIP®/NBTBlue Liquid Substrate was added to develop the color. Prestained molecular weight markers
were obtained from Bio-Rad (Hercules, CA).

57

RESULTS
Characterization of Strep-Tagged UreG. The native form of K. aerogenes UreG was
previously purified from recombinant E. coli cells by sequential use of two Mono-Q columns in
different buffers followed by gel filtration chromatography (13); however, the tendency of the
protein to elute from ion exchange resins over a large number of fractions led to low overall
yields. To overcome this problem and to facilitate a single-step purification of UreG variants,
we developed a new purification system exploiting a fusion peptide sequence that binds with
high affinity to Strep-Tactin resin. The Strep-tag II (42, 43) was designed specifically to allow
affinity purification without introduction of metal-binding residues as in the commonly used
His6-tag. The ureG sequence was cloned into plasmids pASK-IBA3plus and pASK-IBA5plus to
encode UreG fused with a Strep-tag at the C- and N-terminus, respectively. E. coli BL21(DE3)
cells transformed with the plasmid derived from the pASK-IBA3plus vector produced more
recombinant protein, so this plasmid was selected for further experiments. For comparative
analyses, native UreG also was obtained by using the previously described protocol (13).
Highly purified UreGStr was obtained by single-step chromatography on a Strep-Tactin
column and essentially homogeneous protein was available after subsequent gel filtration
chromatography in buffer containing 1 mM DTT (Figure 2.3A). The elution profile (Figure 2.3B)
was consistent with UreGStr being monomeric with a very small shoulder suggesting a trace of
dimeric protein. Significantly, the monomeric nature of this protein was retained regardless of
the presence or absence of nickel or zinc ions. By contrast to these metal ion-independent
results, the inclusion of nickel ions caused the C28A UreGStr variant to chromatograph primarily

58

as a dimer (Figure 2.3B, gray trace), as described further in a later section. For comparison to
UreGStr, native UreG exhibited a major monomeric species as well as a minor dimeric feature
by size exclusion chromatography (data not shown), with the dimer peak disappearing after
overnight dialysis in a buffer containing DTT. These results are consistent with the dimer being
an artifact of oxidation that occurs much more readily in the wild-type protein than in UreGStr.

59

Figure 2.3: Purification of UreGStr and size exclusion chromatography native size analysis. (A)
Purification of UreGStr. Lane 1: UreGStr after Strep-Tactin column purification. Lane 2: UreGStr
after Superdex-75 gel filtration chromatography. (B) Sephacryl S300HR size exclusion
chromatography.

UreGStr (1.0 mL) was loaded onto a 130 mL Sephacryl S-300 column

equilibrated with 50 mM HEPES buffer, pH 7.4, containing 200 mM NaCl plus 15 µM NiCl2 and
-1

chromatographed at a flow rate of 1 mL min . Black: UreGStr. Gray: C28A UreGStr. The
positions of molecular weight standards (BioRad) are indicated in kDa.

60

The presence of the Strep-tag did not affect folding of UreG according to CD
spectroscopy (spectra not shown). Fitting of the spectra indicated UreGStr (60% α helix, 18% β
strands, 4% turns, and 18% random coil for UreGStr) possessed essentially the same secondary
structure as native UreG (65% α helix, 15% β strands, 5% turns and 15% random coil for native
UreG), each with a normalized root mean square deviation of 0.001).
Targeting Residues for Mutagenesis.
residues for mutagenesis.

Several criteria were used to select UreGStr

First, we identified highly conserved residues by creating an

alignment using Clustal W (44) of the most diverse UreG and HypB sequences found in the NCBI
database along with other UreG sequences of interest. The hydrogenase-activating GTPases
are ~25% identical in sequence to UreG and both of these protein families function in assembly
of nickel metallocenters. Notably, residues conserved in these two protein families constitute a
much smaller number than the residues conserved in just UreG proteins (where the identities
typically are over 50%; see UreG sequence comparisons in (22, 23)). The UreG/HypB sequence
comparison highlights the P-loop motif (GSGKT at positions 17-21 in K. aerogenes UreG), the
signature motif (ESGG at positions 104-107 of UreG) for the SIMBI G3E family of GTPases (45),
and the guanine specificity loop (NKTD at positions 151-154 of UreG) (Figure 2.2). Second, we
examined the crystal structure of M. jannaschii HypB (27) which uses Cys95, His96, and Cys127
to coordinate a dinuclear zinc binding site; counterparts were identified in K. aerogenes UreG
(Cys72, His74, and perhaps either Ser111 or Ser115, although Ser is rarely observed as a metal
ligand). In addition, the HypB structure indicated multiple residues involved in MgGTP binding,
hinting at the comparable residues in UreG. Finally, we identified K. aerogenes UreG residues

61

corresponding to those hypothesized to be metal ligands in B. pasteurii UreG (22), as well as
some residues that were not as highly conserved but seemed likely choices for metal binding.
On the basis of these criteria, the following residues were targeted for mutagenesis.
Lys20, previously shown to be a critical P-loop residue (13), was changed to form K20A UreGStr.
+2

Asp49, equivalent to the Mg -coordinating Asp75 in M. jannaschii HypB (27), was changed to
generate D49A UreGStr. Glu68, the residue corresponding to a suggested metal ligand of the B.
pasteurii protein (22), was changed to obtain the E68A protein. Cys72, likely to correspond to
the Cys95 metal ligand at the dinuclear site of M. jannaschii HypB (27) and whose equivalent
was speculated to be a metal ligand in B. pasteurii UreG (22), was changed to create the C72A
variant. His74, likely to correspond to the His96 dinuclear center ligand of HypB (27) and
equivalent to the postulated His70 metal ligand in B. pasteurii UreG (22), was changed to
produce the H74A mutant protein. Asp80, corresponding to Asp98 of HypB (where it is
positioned between the dinuclear center and the GTP-binding site) and highly conserved in
both proteins, was changed to make D80A UreGStr. Ser111 and Ser115 that approximate the
Cys127 ligand of the dinuclear site in HypB were changed to fashion the S111A and S115A
proteins.

In addition, Cys28, the only other cysteine residue in the K. aerogenes UreG

sequence, along with Glu25 and Asp33, the two acidic residues closest to that cysteine, and
Asp120 and Asp127, two highly conserved aspartic acid residues, were all changed to alanine
residues.
Effect of UreGStr Variants on Urease Activity in Cell Extracts. The selected ureG mutants

62

were expressed as part of the urease operon, and the levels of the encoded UreGStr variants
were shown to be indistinguishable by Western blot (data not shown). The urease activities
measured in soluble extracts of cells producing UreGStr were essentially identical to those of
extracts from cells containing native UreG (Table 2.3). Similarly, cell-free extracts containing
D33A and E68A UreGStr possessed about 80% of the activity observed for extracts containing
the non-mutant UreGStr. The E25A, C28A, S115A, and D127A forms of UreGStr exhibited
somewhat diminished activities (5%, 13%, 30%, and 33%, respectively, of wild type urease
activity). In contrast, the cells producing K20A, D49A, C72A, H74A, D80A, and S111A variants of
UreGStr exhibited nearly undetectable levels of urease activity. For unidentified reasons, the
gene encoding D120A UreGStr was unable to be cloned into the urease gene cluster despite
repeated attempts.

63

Table 2.3: Effects of UreG Mutations on the Urease Activity in Soluble Cell-free Extracts
Sample
Specific activity
Specific activity
-1 a
(% of wild type)
(U mg )
UreG
UreGStr

129.6 ± 10.4
122.6 ± 9.8

100
94.6

K20A UreGStr

0.09 ± 0.01

0.07

E25A UreGStr

6.4 ± 1.9

4.94

C28A UreGStr

16.6 ± 1.3

12.8

D33A UreGStr

105.4 ± 10.5

81.3

D49A UreGStr

0.19 ± 0.04

0.15

E68A UreGStr

107.4 ± 8.6

82.9

C72A UreGStr

0.14 ± 0.02

0.11

H74A UreGStr

0.14 ± 0.04

0.11

D80A UreGStr

0.24 ± 0.15

0.19

S111A UreGStr

0.09 ± 0.01

0.07

S115A UreGStr

39 ± 10

D127A UreGStr

42.6 ± 6.2

30.1
32.9

a

Error values are standard deviation from triplicate biological samples,
including a minimum error associated with protein assays.

Metal Binding to UreG, UreGStr, and UreGStr Variants. Based on published metalbinding studies of UreG from other sources (22-25) and the dinuclear zinc metallocenter
structure of HypB, we tested whether UreG from K. aerogenes would bind nickel or zinc ions.
Freshly purified UreG and UreGStr were free of metal according to inductively coupled plasmaemission spectrometry. The nickel and zinc ion binding properties of UreG, UreGStr, and the
site-directed mutants were examined by using PAR, a colorimetric indicator (41), to monitor
metal concentrations after equilibrium dialysis (Table 2.4). As shown in Figure 2.4, UreGStr and
native UreG each bound approximately one nickel ion per monomer (1.0 ± 0.08 and 0.95 ± 0.09

64

per monomer, respectively), with UreGStr also binding 1.1 ± 0.08 Zn (zinc ion binding to native
UreG could not be reliably determined due to protein precipitation). Surprisingly, UreGStr
bound nickel ions with greater affinity (Kd = 5.0 ± 1.8 µM) than native UreG (Kd = 16 ± 3.1 µM)
(Figure 2.4A). The basis of this lower Kd is unclear and this difference in Kd could be a point of
potential concern; however, UreGStr is able to activate urease to wild-type levels (Table 2.3), so
the tagged version of the protein was used for further studies. The nickel-binding properties of
the variant proteins were studied by using the tagged constructs and selected results were
confirmed by using the non-tagged version.

65

Figure 2.4: Equilibrium dialysis analysis of metal binding to wild type and Strep-tagged UreG.
(A) Nickel ion binding to UreGStr (filled circles) and UreG (open circles). (B). Zinc ion binding to
UreGStr. The concentrations of metal ions in dialysis chambers containing protein and buffer
were assessed by reaction with PAR, and the differences of these values were used to calculate
the amounts of metal:protein complexes. Ligand binding fits to a single type of binding site are
indicated.

66

In order to examine whether nickel and zinc ions compete for the same metal binding
site of UreGStr, additional equilibrium dialysis experiments used
varying concentrations of nickel ions containing

63

63

Ni. When dialyzed against

Ni, UreGStr (25 µM) bound 1.15 ± 0.07 nickel

ions per monomer with a Kd of 2.7 ± 0.2 µM (Figure 2.5 A), in very reasonable agreement with
the PAR data. Using these baseline data, two types of competitive binding assays were
performed. First, UreGStr was dialyzed against varying concentrations of nickel ions containing
63

Ni and a constant concentration of zinc ions (10 µM) (Figure 2.5 A, dashed line). These results

clearly demonstrate competition between the metal ions; equation 2 provided a zinc ion Ki of
3.9 ± 0.3 µM. Second, UreGStr was dialyzed against a constant concentration of nickel ions (25
µM) containing

63

Ni along with varied concentrations of zinc ions (Figure 2.5 B). A zinc ion Ki of

2.7 ± 0.2 µM was determined, in close agreement with the first method. The Ki for zinc ion
competition of nickel ion binding is in good agreement with the Kd for Zn determined by the
PAR method (Figure 2.4 B).

67

Figure 2.5: Equilibrium dialysis analyses to assess the competition of nickel and zinc ions. (A)
Varying concentrations of nickel ions containing

63

Ni were examined for binding to 25 µM

UreGStr in the absence of added zinc ions (filled circles) or in competition with 10 µM ZnCl2
(open circles). (B) 25 µM UreGStr was mixed with 25 µM nickel ions containing
concentrations of zinc ions. The data were fit by using Eq. 3.

68

63

Ni and varied

PAR-based equilibrium dialysis experiments were carried out with all site-directed
variants. Unfortunately, zinc ions caused protein precipitation at concentrations higher than
100 µM with nearly all of the mutant UreGStr proteins, thus precluding their detailed
thermodynamic analyses. In contrast, the mutant proteins exhibited well-behaved nickel ion
binding curves. Table 2.4 provides the nickel ion Kd and Bmax for each mutant protein. Most
UreGStr variants behaved much like the control protein in terms of their thermodynamics of
nickel ion binding. That is, their Kd values were the same or only slightly larger than that of
UreGStr and they bound a single nickel ion per protomer. Nearly 4-fold increases in Kd were
measured with the E25A and D80A variants. The largest change in thermodynamic properties
was measured in the case of the C72A UreGStr variant, which exhibited a nickel ion Kd of 61 ±
13 µM, consistent with its involvement in metal binding. Parallel to the large increase in Kd for
C72A UreGStr compared to UreGStr, a similar large increase in Kd was demonstrated in the
mutant protein lacking the Strep-tag (59 ± 24 µM, data not tabulated). By contrast to the
results related to substitution of Cys72, the mutation affecting the only other Cys in the protein
(i.e., C28A UreGStr) behaved much like the control protein in terms of its thermodynamic
properties. Nevertheless, this protein did exhibit anomalous behavior. In particular, the C28A
variant formed predominantly a dimer in the presence of nickel ions as identified by gel
filtration chromatography experiments (Figure 2.3 B).

69

Table 2.4: Thermodynamics of Nickel Ion Binding to
UreG, UreGStr, and its Variants
Protein
UreG

Kd (μM)
16 ± 3.1

Bmax
1.0 ± 0.08

UreGStr

5.0 ± 1.8

0.95 ± 0.09

E25A UreGStr

18 ± 5

0.75 ± 0.08

C28A UreGStr

8.5 ± 1.9

0.81 ± 0.06

D49A UreGStr

11 ± 2

0.94 ± 0.08

E68A UreGStr

7.7 ± 3.3

0.82 ± 0.09

C72A UreGStr

61 ± 13

1.21 ± 0.1

H74A UreGStr

12 ± 4

1.2 ± 0.1

D80A UreGStr

20 ± 9

1.1 ± 0.1

S111A UreGStr

6.6 ± 2.4

0.97 ± 0.1

S115A UreGStr

12 ± 4

1.1 ± 0.1

10 ± 2

0.92 ± 0.06

10 ± 5

1.5 ± 0.2

D120A UreGStr
D127A UreGStr

a

a

Data were obtained for this sample using a 15,000
MWCO membrane, whereas all other data used a
3,500 MWCO membrane.
To further investigate the nature of the nickel ion-binding site in UreG, NiCl2 was
titrated into solutions of UreGStr and the two UreGStr cysteine variants while monitoring their
UV-visible spectra. For UreGStr, a peak at 330 nm appeared with increasing Ni
(Figure 2.6 A). This feature is consistent with a thiolate-to-Ni

2+

2+

concentrations

charge-transfer transition (46);

however, the changes in intensity of this peak versus the concentrations of added nickel ions
(Figure 2.7 A) were inconsistent with the Kd obtained by equilibrium dialysis analysis indicating
that the two techniques are not reporting on the same event. No absorption feature was
detected when nickel ions were added to the C72A protein (data not shown), indicating that

70

Cys72 is primarily responsible for the ligand-to-metal charge-transfer band noted for UreGStr.
Titration of nickel ions into the C28A UreGStr protein yielded a feature at 303 nm (Figure 2.6 B).
The perturbation of the peak maximum compared to that of the non-mutated protein could be
indicative of a different ligand environment for the metal ion in the primarily dimeric C28A
UreGStr. The intensity changes observed for the C28A variant with varied nickel ion
concentrations (Figure 2.7 B) do not reflect the expectations from equilibrium dialysis, again
consistent with the two methods measuring non-equivalent nickel ion-binding events.
Regardless, it is clear that a Cys residue binds nickel ions in the C28A and native proteins, but
not in the C72A variant.

71

Figure 2.6: UV-visible spectral titrations of UreGStr and its C28A variant with NiCl2. (A)
Difference spectra obtained for 58 µM UreGStr titrated with nickel ions in 50 mM HEPES buffer,
pH 7.4, containing 200 mM NaCl. (B) Difference spectra of 26 µM C28A UreGStr titrated with
nickel ions in the same manner.

72

Figure 2.7: Difference in UV-visible absorbance spectra according to Ni concentration (A)
Difference in absorbance at 330 nm for UreGStr plotted against the total nickel ion
concentration. (B) Difference in absorbance at 303 nm for C28A UreGStr plotted against total
nickel ion concentration.

73

Pull-down Assays. We exploited the Strep-tag on UreGStr to examine the interactions of
UreG with other cellular proteins and to identify complexes that form in vivo. E. coli DH5α cells
containing the modified urease operon expressing UreGStr or mutants of this protein were
grown with or without added nickel ions, then soluble cell-free extracts were chromatographed
on Strep-Tactin columns and the proteins eluted with desthiobiotin-containing buffer. The
resulting samples were examined by SDS-PAGE, with three key results illustrated (Figure 2.8).
For most samples, UreGStr (the expected major band) associated with the urease structural
subunits (identified by their characteristic sizes and by Western blot analysis using anti-urease
antibodies, data not shown) along with bands migrating at positions expected for the UreD and
UreF accessory proteins. In addition, Western blot analysis using anti-UreE antibodies (data not
shown) identified UreE in all samples, but this protein was present in much smaller amounts for
cells grown in the absence of added nickel ions. In contrast to the other samples, added nickel
ions led to the D80A UreGStr forming a complex only with UreE and not associating with urease,
UreD, or UreF (Figure 2.8 A).
To further investigate the interaction between UreGStr and UreE, in vitro pull-down
studies were performed with the purified proteins. UreGStr and UreE (1:1 molar ratio of
protomers, 16 µM each) were mixed in buffer containing various concentrations of nickel or
zinc ions. After incubating approximately 10 min on ice, the samples were loaded onto StrepTactin columns, washed, and the bound proteins were eluted with desthiobiotin and examined
by SDS-PAGE.

An increasing ratio of UreE bound to UreGStr as the nickel or zinc ion

74

concentrations increased, with approximately 0.5 UreE protomer per UreG observed for 60 µM
or higher metal ion concentration according to densitometry measurements (Figure 2.8 B). To
test whether the amount of complex formation increased over time, a mixture of UreGStr, UreE,
and nickel ions was incubated on ice for up to 4 h before performing the pull-down experiment;
all incubation times exhibited the same amount of complex (data not shown). The resulting
UreGStr:UreE complex was further investigated by using gel filtration chromatography. When
equal protomer concentrations of the two proteins were combined and chromatographed on a
Sephacryl S-300 column in the absence of metal ions (Figure 2.9 A), a single feature was
observed corresponding to overlapping peaks of the monomer of UreGStr (Mr = 23.1 kDa) and
the dimer of UreE (Mr = 35.1 kDa). By contrast, when the experiment was repeated with 60 µM
NiCl2 added to the buffer a second peak with an apparent molecular weight of 168 kDa
appeared. Analysis of fractions from that peak by using SDS-PAGE revealed an approximate 1:2
UreGStr:UreE protomer ratio as calculated by using densitometry measurements (Figure 2.9 B).
Significantly, the ratio obtained here reflects the actual protomer ratio in the isolated complex,
whereas the ratio described above includes a combination of UreE that reversibly associated
with UreGStr as well as free UreGStr. The second peak eluting from this column was comprised
of predominantly UreGStr and chromatographed as the expected monomer. UreE alone forms
an even larger complex with an apparent molecular weight of more than 330 kDa when 60 µM
NiCl2 is present (data not shown).

75

Figure 2.8: Interactions of UreGStr with urease proteins using cell-free extracts and in vitro
interactions between UreGStr and UreE. (A) In vivo complexes formed with selected UreGStr
samples in E. coli DH5α cells. Soluble cell-free extracts were generated from cells grown in
medium lacking (-Ni) or containing (+Ni) nickel ions and expressing the urease operon encoding
non-mutated UreGStr or for cells expressing the operon encoding D80A UreGStr (+Ni). The
extracts were applied to Strep-Tactin columns, and the proteins eluted with desthiobiotin were
subjected to SDS-PAGE. (B) In vitro pull-down assays using purified UreGStr and UreE. The two
proteins (1:1 molar ratio of protomers, 16 µM each) were incubated with varying
concentrations of nickel or zinc ions (0 to 100 µM), loaded onto Strep-Tactin columns, eluted
with desthiobiotin, and subjected to SDS-PAGE.

76

Figure 2.9: Chromotography of UreGStr:UreE complex. (A) Sephacryl S-300 chromatography of
a mixture of UreGStr and UreE (1:1 molar ratio of protomers) in 50 mM HEPES buffer, pH 7.4,
containing 200 mM NaCl with (black) and without (gray) 60 µM NiCl2. Molecular weight
standards (BioRad) are indicated in kDa. (B) SDS-PAGE analysis of the two peak fractions from
panel C from the chromatograph including nickel ions.

77

The UreGStr variants also were mixed with UreE and subjected to pull-down
experiments. None of the UreGStr variants exhibited deficiencies in their abilities to form a
complex with UreE when 60 µM nickel ions were present, nor did any of the mutations form a
complex with UreE in the absence of metal.

78

DISCUSSION
Using a new procedure to purify K. aerogenes UreG, we generated significant findings
related to the protein’s functional quaternary structure, its metal ion binding properties, the
effects of selected mutations on activity and metal binding, and the formation of a complex
between this protein and its cognate UreE in a manner induced by metal ions. Notably, some
of these results obtained using K. aerogenes UreG exhibit stark differences compared to those
reported for UreG proteins from other organisms (22-24).
The use of a Strep-tag on UreG facilitated purification and allowed for protein
interaction studies via pull-down assays. While designed to not interfere with metal binding
analyses (42, 43), a major concern for the more widely used His6-tag, we found the Streptagged version of UreG bound nickel ions more tightly than the wild type protein. The basis of
the three-fold difference in Kd is unclear, but we note that the Strep-tag contains a His residue
which could play some role in metal binding or in slightly perturbing the protein conformation.
These results demonstrate that any tag might have unexpected effects. Significantly, the tag on
UreG does not interfere with its function in urease activation as shown by the ability of UreGStr
to activate urease apoprotein within cells to 95% of that of the wild-type protein, signifying that
the difference in the Kd doesn’t affect the role of UreG in vivo.
CD measurements confirmed that the Strep-tag did not interfere with the overall fold of
the UreG protein. Furthermore, both wild-type UreG and UreGStr were found to be highly
structured (only 18% and 15% random coil, respectively) compared to the intrinsically

79

disordered structures of the B. pasteurii, M. tuberculosis, and H. pylori proteins (30%, 45%, and
~50% random coil, respectively) (23-25). This result might imply that K. aerogenes UreG is
better suited for structural characterization efforts than UreG from other sources.
UreGStr is monomeric according to gel filtration experiments, and this state is
unaffected by the addition of nickel or zinc ions. This quaternary structure differs from the
dimeric UreG proteins of B. pasteurii or M. tuberculosis (23, 25) and from H. pylori UreG which
dimerizes in the presence of zinc, but not nickel ions (24). Several other members of the SIMBI
G3E family of small GTPases possess dimeric structures, while others are monomeric. For
example, HypB and MeaB (an editor for transferring vitamin B12 into methylmalonyl-CoA
mutase), crystallized as dimers, although - of potential interest - their dimer interfaces are
distinct, whereas YjiA (a protein of undefined function) is a monomer (27, 47-49).

Our

conclusion that UreG functions as a monomeric protein in K. aerogenes coincides with earlier
results indicating stoichiometric levels of UreD, UreF, and UreG in various urease complexes
generated in this system and with data demonstrating that UreD and UreF are stoichiometric
with the urease subunits (11-13, 17, 50).
The metal binding properties of K. aerogenes UreGStr also differ significantly from those
of UreG proteins isolated from other species. Equilibrium dialysis studies demonstrated that
nickel and zinc ions compete with similar affinities for a single metal ion-binding site on UreGStr.
While the dimeric B. pasteurii UreG similarly binds 1 zinc ion per protomer, it binds 2 nickel ions
per protomer with the affinities for the two metal ions differing by an order of magnitude (and
these affinities are approximately 10- and 100-fold less than for K. aerogenes UreGStr) (22). H.

80

pylori UreG binds only 0.5 zinc ions per protomer leading to dimerization, whereas it binds two
nickel ions per monomer without dimerization and with 20-fold lower affinity (24).

In

comparison to the B. pasteurii and H. pylori proteins, the small nickel ion Kd of UreGStr may be
compatible with its functional significance in transferring nickel ions to UreD in the UreABCUreDFG activation complex; however, one must be cautious in interpreting these
thermodynamic results since urease metallocenter assembly is, at least in part, a kinetic
process linked to GTP hydrolysis. Furthermore, we cannot rule out that the physiologically
significant metal binding site is comprised of residues from UreG and another urease-related
protein.
We identified Cys72 as a nickel ion ligand in UreGStr. Replacing this residue with Ala led
to a 12-fold increase in the nickel ion Kd, consistent with its participation in the metal binding
site. In addition, titration of nickel ions into UreGStr led to the formation of a 330 nm
2+

absorption attributed to a thiolate-to-Ni

charge-transfer transition which was not generated

when Cys72 was absent, implicating this residue as a nickel-coordinating ligand.

The

corresponding Cys68 residue in B. pasteurii UreG also was proposed as a metal ion-binding
residue; however, the same residue was identified as forming a disulfide bond that stabilized
the dimeric form of that protein (22, 25). Simultaneous function as a disulfide and as a metal
ion ligand is unlikely. The corresponding Cys66 residue in H. pylori UreG was proposed to be
involved in zinc ion binding on the basis of a 10-fold decreased affinity in the C66A variant (24),
but curiously the effects of this mutation on nickel ion binding were not examined. Our studies

81

of K. aerogenes UreG confirm that this conserved cysteine is involved in nickel ion binding and
show it does not form an essential disulfide bond.
Other residues comprising the metal ion-binding site of K. aerogenes UreGStr were not
identified with certainty by our mutagenesis and equilibrium dialysis studies, but some
inferences are possible. The E25A and D80A variants exhibited four-fold increases in nickel ion
Kd, and other substitutions had even smaller effects, consistent with nearby residues
compensating for the loss of some ligands. Nevertheless, it is notable that mutants expressing
the K20A, D49A, C72A, H74A, D80A, and S111A UreGStr proteins in the context of the complete
urease gene cluster were essentially inactive. Lys20 is in the P-loop and Asp49 corresponds to
2+

the Mg

coordinating residue of HypB, thus likely accounting for their essential roles. Based

on homology to the HypB structure, we propose that His74 and Ser111 are located close to
Cys72 and the former residue is likely to participate in metal binding (while we cannot eliminate
the possibility, Ser is much less likely to serve as a metal ligand). The residue corresponding to
His74 was mutated in H. pylori UreG, and the resulting H68A protein bound zinc ions with lower
affinity by an order of magnitude (24), again without analysis of the effects on nickel ion
binding. For B. pasteurii UreG, the metal-binding ligands were proposed to be Glu64, Cys68,
and His70, corresponding to Glu68, Cys72, and His74 of the K. aerogenes protein (22). The lack
of effect on urease activity for cells containing E68A UreGStr effectively rules out this Glu
residue as an essential metal-binding residue.
The only other cysteine in K. aerogenes UreGStr, Cys28, is not essential for urease

82

activation, but the urease activity decreased to 13% of non-mutant samples in cells containing
the C28A variant. Titration of nickel ions into the C28A variant generated a perturbed UV
spectrum, with nearly a two-fold increase in intensity and a shift of about 30 nm in the
absorption feature, indicating a slightly different metal coordinating environment. This change
may be associated with the protein’s ability to form a dimer in the presence of nickel ions.
In addition to the above new results obtained with purified UreGStr, we investigated the
interaction of this protein with other urease-related proteins.

Soluble extracts of cells

expressing the urease gene cluster with ureG modified to encode UreGStr were analyzed by
pull-down assays. The Strep-tagged version of UreG formed a complex that included all other
urease components, with the amount of bound UreE enhanced by the presence of nickel ions.
A similar UreABC—UreD—UreF—UreG—UreE complex was previously described for a sample in
which a hinge-like region of UreB was mutated, resulting in the trapping of this complex (50).
Those studies led to a model in which the accessory proteins function, in part, to shift the
position of the main domain of UreB to allow nickel ions and bicarbonate to gain access to the
nascent active site.
When cells expressing the UreGStr variants were examined in the context of the other
urease components, Asp80 was identified as being essential for stabilizing the binding of UreG
to the UreABC—UreD—UreF complex. Significantly, the D80A variant failed to generate the
UreABC—UreD—UreF—UreG—UreE complex; instead, it only interacted with UreE. Asp80 is
likely to be positioned at the interface between UreG and the UreABC—UreD—UreF complex.
On the basis of prior studies examining urease-related complexes formed with the K. aerogenes

83

proteins, UreG most likely binds to UreF (12-14, 17). The D80A UreGStr:UreE complex indicates
that UreE binds to UreG within the UreABC—UreD—UreF—UreG—UreE complex.

An

interaction between UreG and UreE was previously suggested by two-hybrid analyses of the H.
pylori proteins (33) and by direct biochemical analysis of these proteins from the same
microorganism (26).
Further investigation of the interaction between UreGStr and UreE used purified
proteins and in vitro pull-down assays to reveal stabilization of the complex by either nickel or
zinc ions. Zinc ion-dependent stabilization of a complex between these proteins was seen
previously with the H. pylori proteins (26), but in that case nickel ions were ineffective for
generating the complex. Moreover, the protein stoichiometries of the two complexes differed.
Whereas the H. pylori proteins formed a zinc-stabilized (UreG)2(UreE)2 complex, with a dimeric
UreG binding to the dimeric UreE, the K. aerogenes proteins formed a nickel- or zinc-stabilized
complex with one UreE dimer per UreGStr protomer, aggregated into a [UreGStr(UreE)2]3
complex of ~168 kDa. The Ni-stabilized interaction between K. aerogenes UreGStr and UreE,
coupled with the Ni-binding capabilities of UreG and UreD (17), support a model in which UreE
delivers nickel ions to UreG within the UreABC—UreD—UreF—UreG complex (Figure 2.10),
with the metal ion subsequently passed from UreG to UreD and then into the nascent active
site of urease. One or more of the sequential metal ion transfer steps is likely driven by GTP
hydrolysis, and the overall process, but not the individual proteins, is specific for nickel ions.

84

Figure 2.10: Model of UreG:UreE interaction in the urease activation process. UreE binds Ni
and delivers it to UreG.
In conclusion, this work describes a new approach to purify K. aerogenes UreG using a
Strep-tag, provides critical new insights into the interactions between this protein and nickel
and zinc ions, identifies Cys72 as a nickel ligand, demonstrates the necessity of Asp80 for
stabilizing UreG binding to UreABC—UreD—UreF, establishes UreG as the site of binding for
UreE, and supports a model for sequential metal ion transfer from UreE to UreG to UreD to the
urease active site.

85

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91

Chapter 3

Klebsiella aerogenes UreF: Identification of the UreG Binding Site and Role in Enhancing the
Fidelity of Urease Activation

This chapter is adapted from Boer, J. L., and Hausinger, R. P. (2012) Klebsiella aerogenes UreF:
Identification of the UreG binding site and role in enhancing the fidelity of urease activation,
Biochemistry in press.

92

ABSTRACT
The Ni-containing active site of Klebsiella aerogenes urease is assembled through the
concerted action of the UreD, UreE, UreF, and UreG accessory proteins. UreE functions as a
metallochaperone that delivers Ni to a complex of UreD—UreF—UreG bound to urease
apoprotein, with UreG serving as a GTPase during enzyme activation. The present study
focuses on the role of UreF, previously proposed to act as a GTPase activating protein (GAP) of
UreG.

Sixteen conserved UreF surface residues that may play roles in protein:protein

interactions were independently changed to Ala. When produced in the context of the entire
urease gene cluster, cell-free extracts of nine site-directed mutants had less than 10% of the
wild-type urease activity. Enrichment of the variant forms of UreF, as the UreE-F fusion
proteins, uniformly resulted in co-purification of UreD and urease apoprotein; whereas UreG
bound to only a subset of the species. Notably, reduced interaction with UreG correlated with
the low activity mutants. The affected residues in UreF map to a distinct surface on the crystal
structure, defining the UreG binding site. In contrast to the hypothesis that UreF is a GAP, the
UreD—UreF—UreG—urease apoprotein complex containing K165A UreF exhibited significantly
greater levels of GTPase activity than that containing the wild-type protein. Additional studies
demonstrated the UreG GTPase activity was largely uncoupled from urease activation for the
complex containing this UreF variant. Further experiments with these complexes provided
evidence that UreF gates the GTPase activity of UreG to enhance the fidelity of urease
metallocenter assembly, especially in the presence of the non-cognate metal Zn.

93

INTRODUCTION
Urease, an enzyme that is widely found in bacteria, plants, and fungi, hydrolyzes urea
into ammonia and carbonic acid, with the resulting ammonification and increase in pH having
important implications in medicine and agriculture (1-4). The best-studied urease system is
produced by recombinant Escherichia coli expressing the Klebsiella aerogenes urease genes,
ureDABCEFG. K. aerogenes urease contains three subunits, UreA, UreB, and UreC, that form a
trimer of trimers in the supramolecular structure (5). The three active sites in the native
enzyme each contain two Ni atoms bridged by a lysine carbamate and are deeply buried in the
protein. Assembly of these active sites requires nickel, bicarbonate, GTP hydrolysis, and
products of the other four ure genes (2, 4). The (UreABC)3 apoprotein can be purified alone (6)
or with combinations of the UreD, UreF, UreG, and UreE accessory proteins (7-11). With each
additional accessory protein, (UreABC)3 is more primed for activation, leading to greater in vitro
activity after incubation with Ni, bicarbonate, and GTP when UreG is present (Figure 3.1). As
summarized below, some properties of these complexes and the individual urease accessory
proteins have been discerned (these comments refer to the K. aerogenes urease components
unless stated otherwise); however, their exact roles remain unclear.

94

Figure 3.1: Simplified scheme of the urease activation process. Urease is synthesized as an
apoprotein, lacking Ni ions and with its active site Lys free of carbamylation. Accessory proteins
UreD, UreF, and UreG bind to the apoprotein and, through a process not completely
understood, the Lys is carbamylated by CO2, the metallochaperone UreE delivers Ni, GTP is
hydrolyzed, and the auxiliary proteins depart leaving active urease.
The (UreABC—UreD)3 complex has UreD bound at the vertices of the triangular
apoprotein according to small-angle X-ray scattering (SAXS) (12) and chemical cross-linking (13)
approaches. When ureD is expressed alone the resulting protein is insoluble, but a fusion of the
maltose binding protein (MBP) with UreD is soluble and functional (14). Surprisingly, the fusion
protein does not bind to (UreABC)3 in vitro, but it forms a complex with the urease apoprotein

95

when co-produced in vivo. Also of interest, MBP-UreD binds divalent metal ions (~2.5 Ni per
protomer, Kd ~50 μM; ~4 Zn per protomer, Kd ~5 μM). The structure of UreH (a UreD
homologue) from H. pylori was recently reported as part of a (UreH—UreF)2 complex (15).
UreH exhibits a novel fold consisting of 17 β-strands and 2 α-helices. The working hypothesis
for urease activation suggests that UreD enhances the activation efficiency of urease
apoprotein while also acting as a scaffold for the binding of the other auxiliary proteins (2, 4).
(UreABC—UreD—UreF)3 represents the apoprotein with heterodimers of UreD—UreF
binding at its vertices, according to SAXS analysis (12). Chemical cross-linking studies indicate a
conformational change takes place in the urease apoprotein within this complex and show a
link between UreB and UreF (13). Yeast 2-hybrid experiments carried out using the H. pylori
and Proteus mirabilis proteins also support an interaction between UreF and UreD/H (16, 17).
Although K. aerogenes UreF is insoluble when produced alone, the fusion proteins with UreF
linked to MBP (18) or to UreE (creating UreE-F) (19) are soluble. The gene encoding the fusion
protein complements a ureF deletion mutant and binds in vitro to UreABC—UreD or MBP-UreD
(14, 19). Truncation variants indicate that the C-terminal 15 residues of UreF are essential for
forming the activation complex, and the N-terminal 24 residues are needed for urease
activation, but not for binding to the activation complex (19). The crystal structure was solved
for truncated H. pylori UreF lacking its C-terminal 21 residues and with its first 24 residues
disordered (20). The H. pylori (UreH—UreF)2 structure contains the last 21 residues that the
structure of UreF alone was missing, but the UreF N-terminus also was unstructured in this
complex (15). The dimeric UreF protein exhibits an all-alpha-helical fold with highly conserved

96

residues mapping to one face. Notably, the dimer interface would need to be disrupted to form
(UreABC—UreD—UreF)3. A modeling study suggested a role of UreF as a GTPase activating
protein (GAP) for UreG, a small GTPase (21).
UreABC—UreD—UreF—UreG is formed by UreG binding to the above complex or, as
likely to occur in the cell, by UreD—UreF—UreG binding to the urease apoprotein. The
UreABC—UreD—UreF—UreG complex was shown to have increased urease activation potential
when incubated with Mg2GTP in addition to Ni and bicarbonate (10). A related complex is
produced by utilizing the Strep II-tagged version of UreG (UreGStr) to purify UreABC—UreD—
UreF—UreGStr (22). UreG itself is soluble and has been characterized from several organisms.
In K. aerogenes this protein is a monomer that binds one Ni or one Zn with Kd values of about 5
µM (22). UreGs from other microorganisms have slightly different properties. H. pylori UreG
dimerizes when Zn is bound, but remains monomeric when Ni is present or in the absence of
metal ion (23). UreG from Bacillus pasteurii and Mycobacterium tuberculosis is dimeric with the
subunits joined by a disulfide bridge (24, 25).

All UreGs contain GTPase-specific motifs

including a P-loop, which is necessary for GTP binding, but low or undetectable GTPase activity
is found for the isolated proteins, consistent with nucleotide hydrolysis being coupled to urease
activation in the larger complex.
UreE is proposed to be the metallochaperone responsible for delivering Ni to the
activation complex to allow for maturation of the active site (26). A complex containing all of
the urease proteins has been detected (12, 22). Furthermore, Ni or Zn promotes binding

97

between UreE and UreG (22), also noted with the H. pylori proteins (27).
This chapter examines the effects of UreF variants on urease activity, the interactions of
UreF with other urease proteins, and the function of UreF in urease activation. Variant forms of
UreF were created on the basis of the H. pylori UreF structure and sequence alignments. The
competence of each variant protein was assessed for in vivo activation of urease. Pull-down
analysis of UreE-F constructs was carried out to test the effects of the mutations on in vivo and
in vitro interactions with other urease components. Finally, the properties of UreABC—UreD—
UreF—UreGStr and a key UreF variant complex were characterized, the proposed role of UreF
as a GAP was assessed, and an alternative hypothesis that UreF functions as a checkpoint for
proper metallocenter synthesis was examined.

98

EXPERIMENTAL PROCEDURES
Plasmids, oligonucleotides, and site-directed mutagenesis. A description of the plasmids
used in this study is provided in Table 3.1, and oligonucleotides used here are identified in Table
3.2. Plasmids pKK17 (encoding the complete ureDABCEFG gene cluster) (26), pKKEF (with the
UreE-F fusion protein encoded within the complete urease gene cluster), and pET-EF (encoding
a translational fusion of UreE and UreF) (19) were mutated by using overlapping
oligonucleotides containing the desired mutations and PfuTurbo Hotstart PCR mastermix
(Promega).

The products were digested using Dpn1 and transformed into chemically

competent E. coli DH5α cells (Invitrogen). The mutations were confirmed by sequencing (Davis
Sequencing, CA), the pET-EF based plasmids were transformed into C41(DE3) competent cells
(28), and all other plasmids were transformed into BL21 (DE3) competent cells. In order to
examine further the K165A mutation of UreF, plasmid pKK17-K165A was digested with AatII
and AvrII and the desired fragment was ligated into similarly digested pKKG (expressing the
Strep II-tagged version of UreG along with the other urease genes) (22) to create pKKG-UreFK165A. The same restriction enzymes were used to replace a fragment of pEC005 (containing
ureFG cloned into pACT3) (14) with the DNA encoding the K165A mutation, producing pEC005UreF-K165A. A DNA fragment encoding UreG T21A was inserted into both pKKG and pEC005 by
using AatII and RsrII, forming pKKG-T21A and pEC005-UreG-T21A.

Plasmids pKAUD2,

pIBA3+UreG, and pEC002, used for production of (UreABC-UreD)3, UreGStr, and MBP-UreD—
UreF—UreG, were described previously (7, 14, 22).

99

Table 3.1: Plasmid properties
Plasmid
Description
pKK17
Wild-type K. aerogenes urease cluster
(ureDABCEFG) inserted into pKK223-3
pKK17-P19A, -G21A, -Y23A, - Single site mutations of pKK17 for studying the
S26A, -E30A, -D60A, -E94A, - effects of the encoded UreF variants on urease
K165A, -F169A, -Q171A, activity
H214A, -E215A, -R220A, L221A, -F222A, -S224A
pKKEF
Same as pKK17, but with a translational fusion of
UreE and UreF
pKKEF-P19A, -G21A, -Y23A, - Single site mutations of pKKEF for in vivo pull-down
S26A, -E30A, -D60A, -E94A, - studies using the encoded proteins
K165A, -F169A, -Q171A, H214A, -E215A, -R220A, L221A, -F222A, -S224A
pET-EF
Translationally fused ureEF genes inserted into
pET21 for production of the UreE-F fusion protein
pET-EF-P19A, -G21A, -Y23A, - Single site mutations of pET-EF for production of
S26A, -E30A, -D60A, -E94A, - UreE-F variants for in vitro pull-down studies
K165A, -F169A, -Q171A, H214A, -E215A, -R220A, L221A, -F222A, -S224A
pKKG
Same as pKK17, but with a translational fusion of
UreG and a Strep II tag
pKKG-UreF-K165A
Single site mutation of pKKG encoding the K165A
UreF variant
pKKG-T21A
Single site mutation of pKKG encoding the T21A
variant of Strep II tagged UreG
pEC005
ureFG fragment cloned into pACT3 for production
of UreF and UreG
pEC005-UreF-K165A
Single site mutation of pEC005 that encodes the
K165A variant of UreF along with UreG
pEC005-UreG-T21A
Single site mutation of pEC005 that encodes the
T21A variant of UreG along with UreF
pKAUD2
Plasmid for production of (UreABC—UreD)3
pIBA3+UreG
pEC002

Reference
(26)
This study

This study
This study

(19)
This study

(22)
This study
This study
(14)
This study
This study
(7)

pASK-IBA3plus-derived plasmid for production of (22)
UreG with a Strep II tag fused to its C-terminus
ureD cloned into pMal-c2x for production of UreD (14)
fused at its C-terminus to MBP. Used along with
pEC005 for production of MBP-UreD—UreF—UreG.

100

Table 3.2: Oligonucloetides used in this study
Purpose
Sequence
UreF P19A
5' C AGC AGC AAC CTG GCG GTA GGG GGT TAC 3'
variant
5' GTA ACC CCC TAC CGC CAG GTT GCT GCT G 3'
UreF G21A
5' C AAC CTG CCG GTA GCG GGT TAC AGC TGG 3
variant
5' CCA GCT GTA ACC CGC TAC CGG CAG GTT G 3'
UreF Y23A
5' CCG GTA GGG GGT GCG AGC TGG TCC CAG 3'
variant
5' CTG GGA CCA GCT CGC ACC CCC TAC CGG 3'
UreF S26A
5' G GGT TAC AGC TGG GCG CAG GGG CTG GAG TG 3'
variant
5' CA CTC CAG CCC CTG CGC CCA GCT GTA ACC C 3'
UreF E30A
5' G TCC CAG GGG CTG GCA TGG GCT GTG GAA G 3'
variant
5' C TTC CAC AGC CCA TGC CAG CCC CTG GGA C 3'
UreF D60A
5' C TTT TTT ACC GTT GCC CTG CCG CTG TTC 3'
variant
5' GAA CAG CGG CAG GGC AAC GGT AAA AAA G 3'
UreF E94A
5' GG GAA ACT CGT GCC CTG CGG GAG GAA G 3'
variant
5' C TTC CTC CCG CAG GGC ACG AGT TTC CC 3'
UreF K165A
5' G ATG GCC GGC GTC GCG CTG GTC CCC TTC 3'
variant
5' GAA GGG GAC CAG CGC GAC GCC GGC CAT C 3'
UreF F169A
5' C AAG CTG GTC CCC GCC GGC CAG CAG GC 3'
variant
5' GC CTG CTG GCC GGC GGG GAC CAG CTT G 3'
UreF Q171A
5' CTG GTC CCC TTC GGC GCG CAG GCC GCC CAG CAG 3'
variant
5' CTG CTG GGC GGC CTG CGC GCC GAA GGG GAC CAG 3'
UreF H214A
5' C GCC TCT GCC CGG GCG GAA ACC CAA TAC TC 3'
variant
5' GA GTA TTG GGT TTC CGC CCG GGC AGA GGC G 3'
UreF E215A
5' C ATC GCC TCT GCC CGG CAT GCG ACC CAA TAC TCT CGA TTA TTC 3'
variant
5' GAA TAA TCG AGA GTA TTG GGT CGC ATG CCG GGC AGA GGC GAT G 3'
UreF R220A
5' CAT GAA ACC CAA TAC TCT GCG TTA TTC CGT TCC TAG AGC 3'
variant
5' GCT CTA GGA ACG GAA TAA CGC AGA GTA TTG GGT TTC ATG 3'
UreF L221A
variant
UreF F222A
variant
UreF S224A
variant

5' GAA ACC CAA TAC TCT CGA GCG TTC CGT TCC TAG AGC TTG 3'
5' CAA GCT CTA GGA ACG GAA CGC TCG AGA GTA TTG GGT TTC 3'
5' C CAA TAC TCT CGA TTA GCG CGT TCC TAG AGC TTG CG 3'
5' CG CAA GCT CTA GGA ACG CGC TAA TCG AGA GTA TTG G 3'
5' CT CGA TTA TTC CGT GCG TAG AGC TTG CGG CCG 3'
5' CGG CCG CAA GCT CTA CGC ACG GAA TAA TCG AG 3'

Protein purification. E. coli C41(DE3) cells producing UreE-F or its variations were grown
overnight in 10 mL lysogeny broth (LB) supplemented with 300 µg mL

-1

ampicillin. The cultures

were used to inoculate 1 L of Terrific Broth (TB, Fisher BioReagents) supplemented with 300 µg

101

mL

-1

ampicillin, grown to optical densities at 600 nm (OD600) of 0.4 to 0.6, induced with 0.5

mM isopropyl β-D-1-thiogalactopyranoside (IPTG), and grown overnight at 37 °C. Cells were
-1

harvested by centrifugation, resuspended (1 g mL ) in buffer A (20 mM Tris, pH 7.8, containing
500 mM NaCl and 60 mM imidazole), supplemented with 1 mM phenylmethylsulfonyl fluoride,
and sonicated (Branson 450 sonifier, five repetitions of 2 min each at 3 W output power and
50% duty cycle). Disrupted cells were centrifuged at 100,000 x g at 4 °C for one h and the cellfree supernatants were loaded onto a 5 mL Ni-nitrilotriacetic acid (NTA) column equilibrated
with buffer A. The column was washed with buffer A until the A280 reached baseline, and
bound proteins were eluted by using buffer B (20 mM Tris, pH 7.8, containing 500 mM NaCl and
1 M imidazole). Fractions containing UreE-F or its variations were combined and dialyzed
overnight into 20 mM Tris, pH 7.8, containing 100 mM NaCl, 1 mM EDTA and 1 mM
dithiothreitol (DTT). The proteins were further purified by gel filtration chromatography using a
Superdex-75 column (65 cm × 2.0 cm diameter; GE Healthcare) equilibrated in the same buffer.
E. coli cells producing UreABC—UreD—UreF—UreGStr or its variants were grown
overnight in 10 mL LB supplemented with 300 µg mL
inoculate 1 L of LB containing 300 µg mL

-1

-1

ampicillin. The cultures were used to

ampicillin, grown to an OD600 of 0.4 to 0.6, induced

with 0.1 mM IPTG, and grown overnight at 37 °C. Cells were harvested by centrifugation,
resuspended in buffer W (100 mM Tris pH 8.0, 150 mM NaCl, 1 mM EDTA), and sonicated (same
protocol as above). Disrupted cells were centrifuged at 100,000 x g at 4 °C for one h and the
cell-free supernatant was loaded onto a Strep-tactin column. The column was washed with

102

buffer W until the A280 was at baseline, and bound proteins were eluted with the same buffer
containing 2.5 mM desthiobiotin.

Fractions containing the protein of interest were

concentrated and loaded onto a Superdex-200 column equilibrated with 25 mM 4-(2hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) buffer, pH 7.4, containing 150 mM NaCl
and 1 mM tris(2-carboxyethyl)phosphine (TCEP). Before use in further assays, the protein was
dialyzed into buffer with no TCEP.
(UreABC—UreD)3 and UreGStr were produced using pKAUD2 and pIBA3+UreG,
respectively, and purified as previously described (7, 22).

MBP-UreD—UreF—UreG was

obtained from cells coexpressing pEC005 and pEC002 (14) and isolated as reported earlier (29).
Sodium dodecyl sulfate (SDS)-polyacrylamide gel electrophoresis (PAGE). SDS-PAGE was
performed by using standard buffers (30), 12% acrylamide running gels, and 4% stacking gels,
except in the case of UreABC—UreD—UreF—UreGStr analysis, when 15% running gels were
used.
Pull-down assays using cell-free extracts. Cells containing pKKEF and its variants were
-1

grown in 50 mL TB supplemented with ampicillin (300 µg mL ) to an OD600 of 0.4 to 0.6,
induced with 0.5 mM IPTG, and allowed to grow overnight at 37 °C. Cells were harvested,
resuspended in 2 mL buffer A, sonicated, and microcentrifuged (14,000 rpm). Cell-free extracts
were added to 200 μL of Ni-NTA resin, washed with 5 mL buffer A, and then eluted with 250 µL
buffer B. Eluted fractions were examined for the presence of other urease related proteins by
using 15% SDS-PAGE.
In vitro pull-down assays. Two distinct types of in vitro pull-down studies were carried

103

out. (i) UreE-F and its variants were tested for their abilities to bind to purified (UreABC—
UreD)3. The apoprotein complex (12 μM nascent active site, dialyzed into buffer containing 20
mM Tris, pH 7.8, and 100 mM NaCl) was combined with UreE-F and its variants (12 µM
protomer concentration in the same buffer) and incubated at room temperature for 20 min.
The protein mixtures were applied to 0.5 mL Ni-NTA columns, washed with 5 mL of 20 mM Tris,
pH 7.8, containing 100 mM NaCl, and eluted with the same buffer containing 1 M imidazole. (ii)
For examining interactions of UreF with UreG, samples of UreE-F or its site-directed variants
were combined with purified UreGStr (12 µM of each protomer) in 20 mM Tris, pH 7.8, buffer
and incubated for one h at 42 °C. The proteins were applied to 0.5 mL Strep-Tactin columns,
washed with 5 mL of 20 mM Tris buffer and eluted with 2 mM desthiobiotin in the same buffer.
Each in vitro pull-down assay was analyzed by 12% SDS-PAGE and gel scanning (Alpha Imager
2200), with the intensities of the bands divided by the molecular mass of each protein (MBPUreD, 72.9 kDa; UreC, 60.3 kDa; UreE-F, 42.8 kDa; UreD, 29.8 kDa; UreF, 25.2 kDa; UreGStr, 23.2
kDa; UreG, 21.9 kDa; UreB, 11.7 kDa; and UreA, 11.1 kDa; though UreA and UreB typically were
not included in the calculations due to the aberrant dye-binding behavior of these small
subunits) to assess the ratios of interactions.
Urease activity and protein assays. Urease activities were measured by quantifying the
rate of ammonia release from urea by formation of indophenol, which was monitored at 625
nm (31). One unit of urease activity is defined as the amount of enzyme required to hydrolyze
-1

1 µmole of urea min at 37 °C. The standard assay buffer contained 50 mM HEPES, pH 7.8, and
50 mM urea. Protein concentrations were determined by a commercially available protein

104

assay (Bio-Rad).
Urease activation assays.

UreABC—UreD—UreF—UreGStr and its variants were

activated for one h at 37 °C in a standard solution containing 100 mM HEPES buffer, pH 8.3,
containing 150 mM NaCl, 100 µM NaHCO3, 100 µM NiCl2, and the indicated amounts of GTP
(provided as its Li salt with two equivalents of Mg), unless otherwise noted. Experiments to
examine the effects of Zn in the activation solution used the concentrations indicated in the
figures. After activation, urease activity was measured by using the standard assay, except that
the buffer additionally contained 0.5 mM EDTA to prevent Ni-dependent inhibition. Time
course experiments were analyzed by using Sigma Plot (Systat Software, Inc.) and the following
equation, where Y is the measured urease activity in U/mg, Amax is the maximal urease activity
generated, t is the time in minutes, and t1/2 is the time needed to reach half of the maximal
activity.
Y = Amaxt/(t1/2 + t)

(1)

Statistically significant differences in urease activation were determined by a p value < 0.05
when using the Student’s t-test.
GTPase activity assays. GTPase activity was measured by monitoring the amount of
released phosphate using malachite green (32). Samples of UreABC—UreD—UreF—UreGStr or
variants (10 µM nascent active site) were incubated in standard activation solution with varying
amounts of Mg2GTP. After 1 and 2 h, 100 µL aliquots were boiled for 5 min and centrifuged to
pellet the precipitated protein; 90 µL of each supernatant was boiled and centrifuged again to

105

remove more protein; and 80 µL of each supernatant was added to 20 µL of malachite green
dye in a 96-well plate, mixed, and the absorbance monitored at 620 nm. The readings were
compared to a standard curve prepared with known amounts of phosphate. Assays without
protein were performed at each GTP concentration to control for GTP hydrolysis over time or at
the higher temperatures.

106

RESULTS
Targets for site directed mutagenesis. K. aerogenes UreF residues were targeted for
mutagenesis by using a previously published alignment (an abbreviated alignment is depicted in
Figure 3.2) of multiple UreF sequences (20) and the two available H. pylori UreF structures (15,
20). Many conserved residues lie close to the N- or C-termini, with the last 10 residues in the
sequence showing high identity. Although residues 1-24 are unstructured in both H. pylori UreF
crystal structures (15, 20), this has no consequence for K. aerogenes UreF which is shorter and
lacks this N-terminal region. Mapping the highly conserved UreF residues onto the H. pylori
crystal structure revealed a clustering on one face of the UreF dimer. (Of interest, the UreF
residues at the interface between H. pylori UreF and UreH were poorly conserved and there is
little overall sequence similarity between K. aerogenes UreD and H. pylori UreH). Based on the
alignment and the structure, 16 K. aerogenes residues were chosen for mutagenesis to Ala
(Figure 3.2 and 3.3). These included highly conserved residues near the N-terminus (P19, G21,
Y23, S26, and E30), several residues at the C-terminus (H214, E215, R220, L221, F222, and
S224), a few conserved potential hydrogen-bond forming residues in the middle (D60, E94 and
Q171), one hydrophobic residue (F169), and the only lysine (K165) in the protein. A Lys or an
Arg is present at this position in all known UreF sequences, and this positively-charged site is
the best candidate for participating in an “Arg-finger”-like manner (33-35) to stimulate GTP
hydrolysis in accord with the proposed GAP activity of UreF (21).

107

* *

K.
H.
P.
B.

aerogenes
pylori
mirabilis
pasteurii

-------------------------MSTAEQRLRLMQLASSNLPVGG
MDKGKSVKSTEKSVGMPPKTPKTDNNAHVDNEFLILQVNDAVFPIGS
-------------------------MMLAD--LRLYQLVSPSLPVGA
---MKWGFLMNNQREGSKNHSNIEATNTNPWLLHLIQIHDTAFPTGS

22
47
20
44

* *
*
*
K. aerogenes YSWSQGLEWAVEAGWVLDVAAFERWQRRQMTEGFFTVDLPLFARLYR
H. pylori
YTHSFGLETYIQQKKVTNKESALEYLKANLSSQFLYTEMLSLKLTYE
P. mirabilis FTYSQGLEWAIEKGWVCSAETLSDWLSAQMTGTLATLELPILRQLQT
B. pasteurii FAHSFGMETYIQESDISNEDDLKAFCDMYLRQNLASTDAIIAQEAYR

69
94
67
91

K.
H.
P.
B.

aerogenes
pylori
mirabilis
pasteurii

*
ACEQGDIAAAQRWTAYLLACRETRELREEERNRGAAFARLLSDWQPSALQQDLKKILGVEEVIMLSTSPMELRLANQKLGNRFIKTLQAMNEL
SLAKGDSDTVKYWCDFMVASRETKELRQEERQRGIAFARLLPQLGILAKENDLQGLIRLENICHAIKLSPETRKGSMMMGRQFLQTVQPLNNS

115
141
113
138

K.
H.
P.
B.

aerogenes
pylori
mirabilis
pasteurii

-----DCPPPWRSLCQQSQLAGMAWLGVRWRIALPEMALSLGYSWIE
DMG-EFFNAYAQKTKDPTHATSYGVFAASLGIELKKALRHYLYAQTS
-----ELDDTLQQRVKQTQLMAFALAAVHWHIDSEKLCCAYVWGWLE
ELFTIWCEKLKNKEIKSHYPVVYGIYTAMLGVDLRTSLETFLYSSIT

157
187
155
185

*
* *
K. aerogenes SAVMAGVKLVPFGQQAAQQLILRLCDHYAAEMPRALAAPDGDIGSAT
H. pylori
NMVINCVKSVPLSQNDGQKILLSLQSPFNQLIEKTLELDESHLCTAS
P. mirabilis NTVMSGVKLVPLGQSAGQKMLFALAEQIPAIVELSAHWPQEDIGSFT
B. pasteurii SLVQNGVRAIPLGQNSGVQTIFSLLPVIQETTSRVMTLDLEHLDNNS

204
234
202
232

K.
H.
P.
B.

aerogenes
pylori
mirabilis
pasteurii

**
*** *
PLAAIASARHETQYSRLFRS
VQNDIKAMQHESLYSRLYMS
PAQVIASSRHETQYTRLFRS
IGLEIASMKHEFLHSRLFIS

224
254
222
252

Figure 3.2: Alignment of UreF sequences from selected microorganisms. Helicobacter pylori,
Proteus mirabilis, and Bacillus pasteurii sequences are aligned with K. aerogenes. Residues
changed to alanine in the K. aerogenes protein are highlighted in bold and indicated by an
asterisk.

108

Figure 3.3: Mutagenesis of K. aerogenes UreF. The structure depicted is that of H. pylori UreF,
from the (UreH—UreF)2 complex (PDB code 3SF5), with the protomers colored in two shates of
gray and the residues changed to alanine in the K. aerogenes protein shown in green and
identified with the H. pylori residue number/ K. aerogenes residue number.
Mutants in ureF have variable effects on in vivo urease activity. E. coli cells expressing
the complete urease operon, but encoding the selected variants of UreF, were shown by SDS
SDSPAGE to produce levels of urease proteins that were essentially identical to those found with
identical
the wild-type accessory protein (data not shown). Cell free extracts of these cultures were
type
Cell-free
examined for urease activities and the results were shown to fall into three general categories
(Table 3.3 and Figure 3.4). The mutant ce containing P19A, Y23A, E30A, E94A, H214A, R220A,
3
cells
L221A, F222A, and S224A forms of UreF all had less than 10% of the wild type activity. These
wild-type
results highlight the critical in vivo role of the C-terminal region in urease activation, since
terminal
altering any one of these highly conserved residues significantly decreased the urease activity.
the
The mutant cells containing G21A, S26A, D60A, K165A, Q171A, and E215A UreF variants

109

retained between 10 and 70% of the wild type activity, indicating that they play important but
wild-type
impor
not vital roles individually in the urease activation process. Finally, the mutant containing the
F169A form of UreF exhibited wild
wild-type activity, demonstrating the non-essentiality of this
essentiality
residue. The F169A variant of UreF was not characterize further.
characterized

Figure 3.4: Effect of UreF variants on urease activity in cell free extracts. Percent activity
cell-free
values shown are relative to that of cell free extracts of cells containing the wild-type urease
cell-free
wild
gene cluster. Measured urease specific acti
activities are provided in Table 3.3

110

Table 3.3: Effect of UreF variants on urease activity in cell-free extracts
Variant of K.
Corresponding residue Urease activity
Urease activity
-1
aerogenes UreF
in H. pylori UreF
(% of wild-type)
[μmol min (mg
-1

Wild-type
P19A
G21A
Y23A
S26A
E30A
D60A
E94A
K165A
F169A
Q171A
H214A
E215A
R220A
L221A
F222A
S224A

protein) ]
130 ± 4
4.4 ± 1.2
47.4 ± 5.3
2.2 ± 0.4
68.5 ± 7.3
0.5 ± 0.1
54 ± 13
0.3 ± 0.04
24.8 ± 2.9
122 ± 23
58.1 ± 5.9
0.14 ± 0.04
26.2 ± 9.5
0.24 ± 0.02
10.0 ± 3.6
0.17 ± 0.01
11.5 ± 1.5

-P44
G46
Y48
S51
E55
E85
E119
K195
L199
Q201
H244
E245
R250
L251
Y252
S254

100
3.5 ± 1.0
37.6 ± 4.2
1.8 + 0.3
54.4 ± 5.6
0.4 ± 0.1
43 ± 10
0.24 ± 0.03
19.7 ± 2.3
97 ± 18
46.1 ± 4.7
1.1 ± 0.03
20.8 ± 7.5
0.19 ± 0.02
7.9 ± 2.9
0.13 ± 0.01
9.1 ± 1.2

UreF variants primarily affect binding of UreG in cell-free extracts. Using constructs that
produce variant forms of UreE-F in the context of the other urease proteins, the fusion proteins
were purified, along with accompanying proteins, from cell-free extracts by use of Ni-NTA resin.
These samples were analyzed by SDS-PAGE (Figure 3.5) and compared to the control sample
where “pull-down” analysis of UreE-F resulted in substantial levels of co-purified UreABC, UreD,
and, at lesser levels, UreG. Approximately equal levels of UreE-F were produced in all mutant
cells according to the results from pull-down analysis as monitored by denaturing gel
electrophoresis. The UreE-F protein containing the D60A substitution was not soluble and so
the corresponding pull-down sample could not be analyzed.

All other UreE-F pull-down

samples, regardless of the changes within UreF, contained UreABC and UreD. This result
provides confirmation of the proper folding of UreE-F variants. Similarly, UreG remained bound

111

to UreE-F versions containing S26A, Q171A, and E215A forms of UreF; notably, each of these
UreF variants resulted in cell-free extracts retaining at least 20% of wild-type activity.
Surprisingly, the UreE-F mutant samples corresponding to G21A and K165A UreF bound less
UreG than control samples despite the relatively high levels of urease activity in cell-free
extracts of these UreF variants (38 and 20% of wild-type UreF, respectively). The weaker
interaction with UreG observed in these pull-down studies apparently could be overcome
within the cellular milieu to allow substantial production of active enzyme, especially for the
conservative G21A substitution. In contrast, UreG was greatly diminished or absent in pulldown samples from the P19A, Y23A, E30A, E94A, H214A, R220A, L221A, F222A, and S224A
UreE-F variants; these results coincide with those for cell-free extracts of the corresponding
UreF variants that exhibited <10% of wild type urease activity. Mapping these residues onto
the UreF dimer, from the (UreH—UreF)2 crystal structure (15), identified a clear binding pocket
for UreG (yellow and magenta portions of Figure 3.6). These results highlight the importance of
both the amino- and carboxyl-terminal residues of UreF for binding UreG.

112

Figure 3.5: UreE-F pull-down analyses of cell-free extracts. These identify UreF mutations that
lead to reduction or absence of UreG binding. Extracts of cells containing pKKEF, bearing the
complete ure cluster with ureE and ureF fused, and its ureF variants were added to Ni-NTA
resin, washed, and eluted with 1 M imidazole. Eluted fractions were analyzed by SDS-PAGE
using a 15% acrylamide gel. The lane labeled M denotes the marker proteins (97.4, 66, 45, 31,
21.5, and 14.4 kDa).

113

Figure 3.6: UreF mutations affecting UreG binding. The two protomers of H. pylori UreF are
shown in different shades of grey in space fill mode (PDB code 3SF5). Residues are colored to
indicate the corresponding side chains of K. aerogenes UreF where mutation to Ala had no
effect on UreG binding (blue), mutations resulting in low levels of UreG bind
binding and <10%
urease activity (yellow), mutations causing reduced UreG binding but retention of >10% urease
activity (magenta), and mutation leading to only 21% activity but retention of UreG binding
(red). Residues that interact with UreH in H. pylori are colored orange or red.
UreF variants affect in vitro binding to (UreABC—UreD)3 and UreGStr. Prior studies had
shown that (UreABC—UreD)3 could bind, at low stoichiometry, to UreE F according to pullUreE-F
pull
down studies using a Ni-NTA column (
NTA
(19). That work reported the purified UreE-F as being
rted
UreE
predominantly monomeric according to gel filtration chromatography; however, re
re-evaluation
of the native size by the same approach (in similar buffer conditions, but containing 100 mM
NaCl rather than 200 mM NaCl) revealed the isolated protein to be primarily dimeric (~73 kDa),
with a small amount of larger molecular weight aggregates. Each of the variant UreE-F proteins
exhibited similar profiles, consistent with their presence as dimeric structures (and providing
evidence for proper folding), except for D60A UreE F that was insoluble and therefore not
,
UreE-F
studied.

Each purified UreE-F variant was combined with isolated (UreABC
F
(UreABC—UreD)3, the

114

mixtures were chromatographed on a Ni-NTA resin, and the extents of interaction between the
proteins were analyzed by SDS-PAGE (an example using E30A UreE-F mixed with (UreABC—
UreD)3 is illustrated in Figure 3.7A) and quantified by gel scanning (Table 3.4). The original
version of UreE-F bound (UreABC—UreD)3 such that 0.32 UreC protomer was associated per
UreE-F protomer, consistent with each (UreE-F)2 dimer binding 0.21 (UreABC—UreD)3
molecules under these conditions. The small amount of complex obtained in this in vitro study
(using equivalent concentrations of UreE-F and UreABC—UreD incubated at room temperature
for 20 min) is less than that observed for UreE-F pull-down studies from cell-free extracts
(Figure 3.5), consistent with cellular factors acting to enhance productive interaction. All the
UreE-F variants except for the L221A variant bound less (UreABC—UreD)3 than the fusion
protein containing wild-type UreF; nevertheless, some interaction was retained in all cases. The
P19A, G21A, and Y23A variants all bound about 70% of the amount of (UreABC—UreD)3
compared to the original UreE-F, while the other UreE-F variants bound less than 50% of that
bound by the control protein. These results demonstrate that none of the sites of substitution
are essential for stabilizing the interaction between K. aerogenes UreF and (UreABC—UreD)3, a
result that is nicely compatible with the H. pylori (UreH—UreF)2 structure (15).

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Table 3.4: Binding of UreABC—UreD by control and variant UreE-F samples
UreE-F variant Ratio of UreABC—UreD/UreE-F
Wild-type
0.32
P19A
0.21
G21A
0.22
Y23A
0.22
S26A
0.15
E30A
0.13
E94A
0.05
K165A
0.14
Q171A
0.13
H214A
0.05
E215A
0.13
R220A
0.11
L221A
0.34
F222A
0.15
S224A
0.08

a

a

Determined by in vitro pull-down studies with Ni-NTA and gel scanning comparison of bands
for UreC and UreE-F.
An analogous series of studies was carried out by using Strep-tactin resin and UreGStr to

examine the interactions of this protein with UreE-F and its variants. A representative gel
depicting the interaction of UreGStr with the E30A variant of UreE-F is shown in Figure 3.7B and
the measured ratios of protein binding for all variants are provided in Table 3.5. The Streptactin chromatography approach avoided the use of Ni, needed for the Ni-NTA resin, due to the
known interaction between UreG and UreE in the presence of this metal ion (22). Interaction
between UreGStr and UreE-F was maximized by incubating the mixture at 42 °C without NaCl
prior to chromatography, resulting in 0.63 UreE-F protomer bound per UreGStr. A much weaker
interaction between these proteins was noted at 37 °C (data not shown) and is consistent with
UreD facilitating the UreE-F/UreG interaction in the pull-down studies from cell-free extracts,

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shown in Figure 3.5. This in vitro interaction assay with its artificial conditions provided trends
that substantially confirmed the results from the UreE-F pull-down assays using cell-free
extracts. UreGStr bound at least 75% of the S26A, Q171A, H214A, E215A, R220A, and S224A
variant UreE-F proteins compared to control UreE-F. Similarly, 65-70% of control levels were
found for P19A, Y23A, and F222A UreE-F, indicating only minor roles for those residues in
stabilizing the binding between UreF and UreG. In contrast, the G21A, E30A, E94A, K165A, and
L221A UreE-F variants exhibited substantially weaker binding (< 50% compared to control UreEF) to UreGStr. I conclude the residues associated with the latter positions are located at the
UreF:UreG interface under these conditions, which represent only a subset of the interface
residues when (UreABC—UreD)3 is present as shown in Figure 3.5.
Table 3.5: Binding of control and variant UreE-F by UreGStr
UreE-F variant Ratio of UreE-F/UreGStr
Wild-type
P19A
G21A
Y23A
S26A
E30A
E94A
K165A
Q171A
H214A
E215A
R220A
L221A
F222A
S224A
a

a

0.63
0.42
0.30
0.43
0.54
0.18
0.28
0.26
0.55
0.50
0.49
0.55
0.25
0.42
0.52

Determined by in vitro pull-down studies using Strep-tactin resin and gel scanning

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Figure 3.7: In vitro pull-down assay examples. A. Pull-down assay for UreE-F mixed with
(UreABC—UreD)3. In this example, E30A UreE-F was mixed with (UreABC—UreD)3 (lane 1) then
bound to Ni-NTA resin and eluted, along with any associated proteins, by addition of imidazole
(lane 2). B. Pull-down assay for UreGStr mixed with UreE-F. In this example, UreGStr was mixed
with E30A UreE-F (lane 1) then bound to Strep-tactin resin and eluted, along with any
associated proteins, by addition of desthiobiotin (lane 2). For each panel, the lane labeled M
denotes the marker proteins (97.4, 66, 45, 31, 21.5, and 14.4 kDa).
Examination of UreF as a GTPase activating protein. In order to test the hypothesis that
UreF acts as a GAP for UreG, several protein complexes containing UreF and UreG were tested
for their GTPase activity by using a malachite green assay to detect product phosphate. First,
UreE-F was incubated with UreGStr to examine whether the isolated proteins could stimulate
GTPase activity (N.B., purified K. aerogenes UreG is essentially inactive (36)). Unfortunately,
control assays using UreE-F alone revealed the presence of trace levels of contaminating
GTPase in the protein preparation that was not eliminated by Ni-NTA or gel filtration
chromatography, so this effort was halted since only UreG of the accessory proteins should

118

have GTPase activity. The urease-free MBP-UreD—UreF—UreG complex (14, 29) also was
investigated as a platform for testing the GAP hypothesis.

This heterotrimeric complex

appeared to possess GTPase activity; however, the complex containing the P-loop (T21A)
variant of UreG, used as a negative control, also exhibited this activity. The assay result again
indicated trace contamination by a GTPase so work on that complex was abandoned. As an
alternate approach, studies were carried out with the previously described UreABC—UreD—
UreF—UreGStr species (22) after performing an additional step of purification involving gel
filtration chromatography.
UreABC—UreD—UreF—UreGStr and the complex containing the K165A UreF variant
were purified (Figure 3.8A) for testing whether the highly conserved, positively-charged residue
at position 165 functions like the Arg finger of a GAP (33-35). Consistent with the weakened
binding of UreG in the K165A UreE-F pull-down studies utilizing cell-free extracts (Figure 3.5),
the complex prepared with the UreF variant possessed less UreGStr when examined after size
exclusion chromatography. The complex also was prepared (gel not shown) using the T21A
variant of UreGStr as a negative control that would be incapable of hydrolyzing GTP (10).
Whereas the wild-type complex exhibited an initial increase in urease activity with increasing
levels of GTP in the activation mixture, followed by a decrease in activation at greater GTP
concentrations, the complex containing the T21A P-loop substitution in UreG resulted in
decreasing extents of urease activation (possibly due to sequestration of Ni) with increasing
GTP (Figure 3.8B). Significantly, the activities generated with these two complexes in the
absence of GTP were similar. By contrast, the complex containing K165A UreF exhibited a

119

decrease in activation competence with added GTP, but the overall level of urease activation
was significantly greater than for the wild-type complex. Analogous activity patterns were
obtained for samples subjected to activation for 2 h (data not shown); i.e., the complex formed
with K165A UreF demonstrated greater activation competence in a GTP-independent manner
compared to the wild-type complex (with or without GTP).
The extent of GTP hydrolysis for each urease activation sample was assessed after 1 h by
using the malachite green assay (Figure 3.8C). The complex containing T21A UreG released
little phosphate, with no further increase after 2 h of incubation, indicating the absence of
contaminating GTPase. The wild-type complex gave rise to an approximately linear increase in
phosphate concentration with increasing amounts of GTP. At small GTP concentrations, the
amount of phosphate formed was roughly proportional to the amount of active urease formed,
but at concentrations greater than 75 µM GTP the amount of phosphate formed continued to
increase while the activity diminished, consistent with an uncoupling of these processes. The
complex containing K165A UreF possessed greater GTPase activity than the complex containing
wild-type UreF at most GTP concentrations, demonstrating that K165 does not facilitate GAP
activity. The comparatively large amount of phosphate produced by the variant complex also
indicates that GTPase activity was significantly uncoupled from urease activation.

120

A

B

C

Figure 3.8: Effect of K165A UreF on GTP-dependent activation and phosphate release by
UreABC—UreD—UreF—UreGStr. (A) SDS-PAGE analysis of purified UreABC—UreD—UreF—
UreGStr and UreABC—UreD—UreF(K165A)—UreGStr after purification by affinity resin and gel
filtration chromatography. M denotes the marker proteins (97.4, 66, 45, 31, 21.5, and 14.4
kDa). (B) Urease activation assays for versions of UreABC—UreD—UreF—UreGStr. Proteins (10
µM) were incubated for one h at 37 °C in standard activation solution with the indicated
concentrations of Mg2GTP. Aliquots were removed and assayed for urease activity. (C)
Phosphate released by UreABC—UreD—UreF—UreGStr during activation as in (B). Symbols
used in (B) and (C): UreABC—UreD—UreF—UreGStr with wild-type UreF (♦), K165A UreF (▲),
or T21A UreGStr (■).
.

121

UreF functions to increase the fidelity of urease activation. The experiments described
above demonstrate that a defective UreF enhances the GTPase activity of UreABC—UreD—
UreF—UreGStr and cast doubt on the hypothesis that this protein serves as a GAP. Because the
GTPase activity of the complex containing K165A UreF appears to be less coupled to urease
activation than that with wild-type protein, we hypothesize that UreF plays a non-GAP role;
specifically, we propose that UreF is a gatekeeper that increases the fidelity of the activation
process. Related to this notion, activation of the isolated urease apoprotein is known to
generate activity in only ~15% of the nascent active sites even though the protein is fully
metallated and carbamylation takes place (37, 38); thus, complete activation must somehow
overcome the formation of the improperly formed metallocenters. Two experiments were
carried out to explore the hypothesis that UreF enhances the fidelity of urease activation.
A time course activation experiment was performed in Ni-limiting conditions (i.e. 50 µM
Ni versus 100 µM Ni, as used above) for the UreABC—UreD—UreF—UreGStr complex and the
same complex containing K165A UreF, using activation assays with and without 75 µM Mg2GTP
(Figure 3.9). The resulting data were fitted to equation 1 to calculate maximal activities and the
times needed to reach half-maximal activities.

These fits ignored the possible slight

diminishments in activation capacities for samples after incubation at 37 °C for 5 h. The
complex containing K165A UreF activated quickly (t1/2 37 ± 12 min) and reached a maximal
activity of 225 ± 16 U/mg. The presence of GTP led to slower activation (t1/2 77 ± 18 min) and a
smaller final activity level (177 ± 14 U/mg), possibly due in part to chelation of Ni by GTP. In

122

contrast, the wild-type complex was slow to activate whether GTP was absent or present (t1/2
83 ± 12 min and 80 ± 20 min, respectively), it formed little activity without GTP (118 ± 6 U/mg),
and with GTP it reached approximately the same maximum activity (228 ± 19 U/mg) as
observed for the complex containing K165A UreF in the absence of nucleotide. These results
are compatible with the hypothesis that UreF enhances the efficiency of the process by
improving the coupling to GTP hydrolysis, decreasing the rate of activation, and assuring the
maximal amount of properly formed metallocenter.

Figure 3.9: Comparison of activation time course for UreABC—UreD—UreF—UreGStr complex
and UreABC—UreD—UreF(K165A)—UreGStr. Proteins (0.5 µM) were incubated in standard
activation buffer except for only containing 50 µM Ni. The wild–type and K165A complexes
were incubated in the absence (o and , respectively) or the presence (• and , respectively) of
75 µM Mg2GTP. Fits using equation 1 are shown for each data set.
As a more direct approach to evaluate this potential gatekeeper role of UreF, we
assessed the effect of added Zn ions on the activation of UreABC—UreD—UreF—UreGStr

123

containing wild-type and K165A accessory protein. Zn is known to compete effectively with Ni
and hinder production of urease activity during activation of (UreABC)3, (UreABC-UreD)3, and
(UreABC-UreD-UreF)3 (7, 8, 38), but its effects on UreABC—UreD—UreF—UreG had never been
examined. As illustrated in Figure 3.10, the presence of Zn in the activation mixture decreases
the level of urease activity generated for both complexes studied whether or not GTP is
present. The complexes were activated at three concentrations of Zn (0, 2.5, and 5.0 µM) and
activities were measured at 1, 2, and 4 h to ensure maximal activation for analysis. After 2 h,
both the wild-type and the K165A variant complexes had reached full activity according to the
control assays without Zn, so that time point was analyzed. Whereas the activity of complex
containing wild-type UreF activated with GTP (solid black bar) was equivalent to the K165A
UreF-containing complex (with or without GTP) in the absence of Zn (hatched and gray bars,
respectively), the wild-type complex with GTP generated significantly more activity (p < 0.05)
than the other mixtures in the presence of low concentrations of Zn. These results indicate that
wild-type UreF allows the GTPase activity of UreG to partially protect against improper
metallocenter assembly. UreF thus plays an important role in coupling the GTPase activity to
urease activation, insuring proper di-Ni cluster formation.

124

Figure 3.10: Zn inhibition of UreABC—UreD—UreF—UreGStr (0.5 µM) activation under standard
conditions for complexes containing wild-type and K165A UreF. Protein complexes (0.5 µM)
were incubated for two h at 37 °C in standard activation solution with the indicated
concentrations of ZnSO4. The wild-type and K165A complexes were incubated in the presence
(black and hatched, respectively) or absence (diagonal and grey, respectively) of 75 µM
Mg2GTP. The asterisks indicate statistically significant differences (p < 0.05) of the sample
indicated versus the samples subjected to the same conditions.

125

DISCUSSION
This study has significantly enhanced our understanding of the UreF urease accessory
protein by clarifying its interactions with UreG and its role in urease activation. Prior efforts
had shown that free K. aerogenes UreF is insoluble (8) whereas a truncated version of the H.
pylori protein is soluble and was structurally defined (20). In addition, soluble forms of UreF are
known to be found in various complexes, including (UreH—UreF)2 from H. pylori (15) and the
following K. aerogenes species: MBP-UreD—UreF—UreG (14), (UreABC—UreD—UreF)3 (8),
and UreABC—UreD—UreF—UreG (9). The (UreH—UreF)2 structure defined how UreF interacts
with the UreD homologue and mutational studies in the same work provided evidence that Y48
and R250 in the H. pylori UreF protein (corresponding to Y23 and R220 in K. aerogenes UreF)
are critical for binding UreG (15); however, other specific residues involved in UreG binding
were not defined. There is no crystal structure available for any UreG sample, perhaps because
the isolated protein cannot be crystallized due to its intrinsic disorder according to NMR,
circular dichroism, and fluorescence spectroscopic analyses (39, 40), so modeling of UreF/UreG
interaction (15) can provide only a crude notion of the potential interface surfaces. The present
investigations extend knowledge of residues critical to this interface and provide new
experimental insights into the role of UreF in urease activation.
The current study targeted sixteen highly conserved UreF residues for mutagenesis as a
means to identify their importance in urease activation and UreG binding. Urease assays of
cell-free extracts from cultures containing the complete urease gene cluster showed that nine
of the mutants exhibited less than 10% of the wild type activity, including those producing the

126

Y23A and R220A UreF variants (comparable to the two H. pylori UreF variants mentioned
above), confirming their importance in urease activation. UreE-F pull-down assays using cellfree extracts showed that substitutions affecting each of these nine residues had reduced levels
of UreG binding, with six leading to the absence of associated UreG. The G21A and K165A
variants had somewhat greater urease activity (38 and 20 U/mg, respectively), but also had
2

reduced amounts of UreG. These important residues map to a 1300 Å surface on UreF (Figure
3.6), and are close to the binding surface for UreD, consistent with how the three proteins are
known to work together for urease activation. None of the site-directed mutations greatly
affected the interaction between UreF and UreABC—UreD, as expected on the basis of the
(UreH—UreF)2 crystal structure which showed that only one of the highly conserved residues
we chose to mutate (E215) is located at the UreD binding surface. It is intriguing that the UreG
binding site on UreF has so many highly conserved residues, whereas the UreD and urease
binding sites on this protein are not highly conserved. Possibly related to this finding, UreG is
the most highly conserved urease accessory protein whereas alignments of UreD or UreF
proteins each exhibit significantly less similarity.
We attempted to directly test the proposed role of UreF as a GAP (21) by comparing the
activation properties of UreABC—UreD—UreF—UreGStr complexes containing wild-type and
K165A UreF. K165 is the only Lys in K. aerogenes UreF and this position is always occupied by a
Lys or Arg residue in homologues; thus, this positively-charged residue could reasonably
correspond to the typical Arg finger motif that is found in GAP proteins (33-35). The Arg finger
of GAPs plays a dual role in generating the nucleophilic water molecule needed for hydrolysis

127

and stabilizing the transition state (34). Urease assays using cell-free extracts showed that K165
of UreF plays a non-essential role in urease activation, with 20% of the wild-type urease activity
retained when using a culture that synthesizes K165A UreF along with the other urease
proteins. In vitro assays using wild-type and variant UreABC—UreD—UreF—UreGStr complexes
unexpectedly demonstrate that the species containing K165A UreF results in enhanced levels of
urease activity. I don’t have a simple explanation for why the in vivo and in vitro results differ in
this manner; however, I note that the urease activation conditions are quite distinct for those
two processes. For the purified complex, activation of the K165A UreF-containing version is
inhibited, not enhanced, by GTP. More significantly, the variant complex exhibits greater
GTPase activity than the wild-type complex; thus ruling out the possibility that K165 serves in
the Arg finger role of a GAP. Although K165 does not function as an Arg finger, it remains
possible that R220, another highly conserved and positively-charged residue (in this case near
the critically important C-terminus), could serve in this manner. Our UreE-F pull-down assays
using cell-free extracts reveal the lack of UreG binding to the complex containing the R220A
variant of UreE-F, so it was not possible to test the effect of mutating this residue in UreABC—
UreD—UreF—UreGStr as was accomplished with the K165A UreF variant. If R220 could act as
an Arg finger, one would need to postulate that K165 modulates its effect so that the K165A
variant leads to greater GTPase activity. There are examples of GAP-GTPase interactions that
do not involve an Arg finger, for example the Rap-RapGAP complex (41, 42). Nevertheless, the
above studies greatly reduce the possibility that UreF acts as a GAP, thus encouraging us to
consider and test other hypotheses for its function.

128

The finding that UreABC—UreD—UreF—UreGStr activation is less coupled to GTPase
activity when using K165A UreF and the demonstration of increased urease activity resulting
from the substituted version of UreF led us to consider a gatekeeper role for this protein; i.e.,
UreF would help to ensure proper metallocenter assembly during urease activation. The
complexes with the two forms of UreF have the same final activation competence, but that
containing wild-type protein requires GTP while that with K165A UreF is inhibited by the
nucleotide.

The time course assays reveal a more rapid activation of the K165A UreF-

containing complex (lacking GTP) than noted in the complex with wild-type UreF (with or
without GTP), consistent with wild-type UreF hindering the rate of activation while insuring
greater fidelity of metallocenter biosynthesis. In contrast, the K165A variant allowed the
process to occur more quickly but possibly with less control. UreG’s GTPase activity appears to
play a role in controlling the rate of activation, and when this activity is uncoupled from
activation (i.e., when using K165A UreF) urease activity is generated much faster. It is likely
more important that the timing is tightly regulated in the cell compared to the idealized
environment of the test tube, since the K165A variant only has 20% of the wild type activity in
vivo. The GTPase activity could play a role in gating the timing of the Ni transfer to the active
site in order to prevent the metal from binding incorrectly (as in the 85% of the urease
apoprotein that becomes carbamylated and binds Ni, yet remains inactive when subjected to
activation conditions as mentioned earlier). This hypothesis is supported by the demonstration
that the complex formed with wild-type UreF (coupled to GTPase activity) is more resistant to
inhibition by Zn during activation than is the case for the variant complex. We propose that
UreF induces a conformational change in, or stabilizes the structure of, UreG that allows for the

129

latter protein’s hydrolytic capacity while ensuring a tight coupling between GTP hydrolysis and
proper metal assembly. Replacing K165 with Ala leads to greater uncoupling of the activation
and hydrolysis steps, thus yielding enhanced rates of activation under favorable conditions, but
allowing incorrect metal assembly when Zn is present, and possibly having additional negative
consequences in vivo. This model is compatible with the finding that GTPase activity is not
essential in vitro for partial activation, as has long been known for (UreABC)3, (UreABC—
UreD)3, and (UreABC—UreD—UreF)3 (8, 9, 38), but the full set of accessory proteins allows for
greater final levels of activation to occur with UreF increasing the fidelity of the process. Thus,
when all urease accessory proteins, including UreE, are present, urease can be activated fully
(11).
The suggestion that GTPase activity can be used as a checkpoint has been reported for
other proteins in the same family as UreG. For example, the GTPase activity of HypB, a protein
that functions in hydrogenase maturation in E. coli, is thought to gate Ni transfer into
hydrogenase with the participation of the Ni-binding protein SlyD (43). Similarly, MeaB,
another member of the small GTPase family, serves a gating function for incorporating
coenzyme B12 into methylmalonyl-CoA mutase (44). We propose that UreF works to improve
the coupling of UreG’s GTPase activity to Ni insertion into urease.
In conclusion, we have confirmed that mutations affecting a series of highly conserved
UreF residues have dramatic effects on in vivo urease activity. Using these same UreF variants
in UreE-F pull-down assays with cell extracts or by carrying out interaction studies with purified
UreF and UreG components, we have identified several UreF residues that make up the binding

130

surface for UreG. We have also evaluated a proposed role of UreF as a GAP and obtained no
evidence to support this hypothesis since a variant UreF gave rise to increased GTPase activity.
Finally, we provide evidence that UreF is needed for coupling GTPase activity to ensure proper
metallocenter assembly, where the gating function of UreF increases the fidelity of activation—
especially when competing Zn is present.

131

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Zambelli, B., Stola, M., Musiani, F., De Vriendt, K., Samyn, B., Devreese, B., Van
Beeumen, J., Turano, P., Dikiy, A., Bryant, D. A., and Ciurli, S. (2005) UreG, a chaperone
in the urease assembly process, is an intrinsically unstructured GTPase that specifically

2+

2+

binds Zn , J. Biol. Chem. 280, 4684-4695.
25.

Zambelli, B., Musiani, F., Savini, M., Tucker, P., and Ciurli, S. (2007) Biochemical studies

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functional conservation among the bacterial UreG family, Biochemistry 46, 3171-3182.
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Colpas, G. J., Brayman, T. G., Ming, L. J., and Hausinger, R. P. (1999) Identification of
metal-binding residues in the Klebsiella aerogenes urease nickel metallochaperone,
UreE, Biochemistry 38, 4078-4088.

27.

Bellucci, M., Zambelli, B., Musiani, F., Turano, P., and Ciurli, S. (2009) Helicobacter pylori
2+

2+

UreE, a urease accessory protein: specific Ni - and Zn -binding properties and
interaction with its cognate UreG, Biochem. J. 422, 91-100.
28.

Miroux, B., and Walker, J. E. (1996) Over-production of proteins in Escherichia coli:
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29.

Carter, E. L., Boer, J. L., Farrugia, M. A., Flugga, N., Towns, C. L., and Hausinger, R. P.
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30.

Laemmli, U. K. (1970) Cleavage of structural proteins during assembly of head of
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31.

Weatherburn, M. W. (1967) Phenol-hypochlorite reaction for determination of
ammonia, Anal. Chem. 39, 971-974.

32.

Baykov, A. A., Evtushenko, O. A., and Avaeva, S. M. (1988) A malachite green procedure
for orthophosphate determination and its use in alkaline phosphatase-based enzyme
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33.

Ahmadian, M. R., Stege, P., Scheffzek, K., and Wittinghofer, A. (1997) Confirmation of
the arginine-finger hypothesis for the GAP-stimulated GTP-hydrolysis reaction of Ras,
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34.

Resat, H., Straatsma, T. P., Dixon, D. A., and Miller, J. H. (2001) The arginine finger of
RasGAP helps Gln-61 align the nucleophilic water in GAP-stimulated hydrolysis of GTP,
Proc Natl Acad Sci 98, 6033-6038.

35.

Scheffzek, K., and Ahmadian, M. R. (2005) GTPase activating proteins: structural and
functional insights 18 years after discovery, Cell. Mol. Life Sci. 62, 3014-3038.

36.

Moncrief, M. B. C., and Hausinger, R. P. (1997) Characterization of UreG, identification
of a UreD-UreF-UreG complex, and evidence suggesting that a nucleotide-binding site in
UreG is required for in vivo metallocenter assembly of Klebsiella aerogenes urease, J.
Bacteriol. 179, 4081-4086.

37.

Park, I. S., and Hausinger, R. P. (1995) Requirement of carbon-dioxide for in vitro
assembly of the urease nickel metallocenter, Science 267, 1156-1158.

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Park, I. S., and Hausinger, R. P. (1996) Metal ion interactions with urease and UreDurease apoproteins, Biochemistry 35, 5345-5352.

39.

Neyroz, P., Zambelli, B., and Ciurli, S. (2006) Intrinsically disordered structure of Bacillus
pasteurii UreG as revealed by steady-state and time-resolved fluorescence
spectroscopy, Biochemistry 45, 8918-8930.

40.

Zambelli, B., Cremades, N., Neyroz, P., Turano, P., Uversky, V. N., and Ciurli, S. (2012)
Insights in the (un)structural organization of Bacillus pasteurii UreG, an intrinsically
disordered GTPase enzyme, Mol. Biosyst. 8, 220-228.

41.

Chakrabarti, P. P., Suveyzdis, Y., Wittinghofer, A., and Gerwert, K. (2004) Fourier
transform infared spectroscopy on the Rap-RapGAP reaction, GTPase activation without
an arginine finger, J. Biol. Chem. 279, 46226-26233.

42.

Scrima, A., Thomas, C., Deaconescu, D., and Wittinghofer, A. (2008) The Rap-RapGAP
complex: GTP hydrolysis without catalytic glutamine and arginine residues, EMBO J. 27,
1145-1153.

43.

Kaluarachchi, H., Zhang, J. W., and Zamble, D. B. (2011) Escherichia coli SlyD, more than
a Ni(II) reservoir, Biochemistry 50, 10761-10763.

44.

Padovani, D., and Banerjee, R. (2009) A G-protein editor gates coenzyme B-12 loading
and is corrupted in methylmalonic aciduria, Proc. Natl. Acad. Sci. U.S.A. 106, 2156721572.

136

Chapter 4

Additional Studies, Conclusions, and Remaining Questions

Portions of this chapter were adapted from Carter, E. L., Boer, J. L., Farrugia, M. A., Flugga, N.,
Towns, C., and Hausinger, R. P. (2011) The function of UreB in Klebsiella aerogenes urease
activation, Biochemistry, 50, 9296-9308.

137

ADDITIONAL STUDIES
The following sections describe experiments that were not incorporated into chapters 2
or 3. In addition to this text, I summarize all constructs I’ve generated in a succinct table
provided as an appendix.
Metal analysis of (UreABC*)3 and (UreAC)3
One of the questions surrounding urease activation is how Ni gets into the active site,
which is buried in the UreC subunit. Previous studies have indicated that UreB may undergo a
conformational change during the activation process allowing for greater access to the active
site (Figure 4.1). For example, chemical cross-linking studies demonstrated that Lys 382 in UreC
crosslinked to Lys 76 in UreB when UreD and UreF were present, whereas the crystal structure
of urease shows those two residues are distant from each other (1). Furthermore, SAXS
analyses showed that the UreD and UreF accessory proteins bind near UreB (2). In order to
investigate the role of UreB, UreB alone and the apoprotein complex (UreAC)3 were purified
and characterized. Part of this characterization included metal analysis of urease with and
without UreB present to see how much Ni was bound. Metal analysis was carried out on
activated (i.e., with added Ni and bicarbonate) (UreAC)3, (UreABC*)3 (formed by mixing purified
(UreAC)3 with a 5-fold excess of UreB and incubating at room temperature for 30 min before
activation), and the complex formed by activating (UreAC)3 followed by addition of UreB.

138

A

B

Figure 4.1: UreB of K. aerogenes urease blocks access to the active site. A. Urease structure
with UreA in blue, UreB in orange, UreC in yellow, and Ni atoms in green. B. Same as A with
UreB removed
To assess the nickel content of activated (UreABC*)3, I mixed 120 μL of (UreAC)3 (370
μM heterodimer in HT buffer (50 mM HEPES (pH 7.8) and 1 mM TCEP)) with 120 μL of UreB
(1450 μM in HT buffer) and incubated the mixture for 30 min at ambient temperature. The
mixture was diluted to 10 μM UreC protomer with 4.2 mL of standard activation buffer (100
mM HEPES (pH 8.3), 150 mM NaCl, 100 mM NaHCO3, and 100 µM NiCl2) and incubated at 37 °C
for 60 min. The sample was concentrated with a 10,000 MWCO Amicon centrifugal filter device
(prewashed with deionized water) to 300 μL and either immediately chromatographed on a 1
cm × 43 cm Sephacryl S300 HR column (GE Healthcare) equilibrated in 100 mM HEPES (pH 8.3)
or treated with 10 mM EDTA for 5 min at ambient temperature and chromatographed in the
same buffer with 1 mM EDTA. Fractions of interest were verified for protein content with
SDS−PAGE, allowed to equilibrate for at least 16 h at 4 °C, concentrated by use of 10,000
MWCO Amicon centrifugal filter devices, and analyzed in parallel with protein-free buffer for
metal content by inductively coupled plasma-atomic emission spectroscopy (ICP-AES) (Chemical
Analysis Laboratory, University of Georgia, Athens, GA). For studies focusing on the nickel

139

content of (UreAC)3 after activation, a similar methodology was employed as described above
except that (UreAC)3 (120 μL, 370 μM in HT buffer) was directly diluted to a final UreC
protomer concentration of 10 μM in activation buffer, followed by the same incubation,
concentration, and gel filtration steps with and without EDTA. Lastly, 4.3 mL of (UreAC)3 (UreC
protomer final concentration of 10 μM) was subjected to standard activation conditions at 37
°C for 60 min, mixed with 120 μL of UreB (1450 μM in HT buffer), incubated at ambient
temperature for 30 min, and analyzed for nickel content after concentration, with or without
EDTA treatment, and gel filtration chromatography as described above.
2+

To examine the effect of inclusion of UreB on Ni

incorporation into urease apoprotein,

samples that had been subjected to the standard activation conditions were analyzed for metal
content by ICP-AES (Table 4.1). For comparison, 0.53 equiv of Ni per heterotrimer was
associated with authentic (UreABC)3 apoprotein that was activated by use of standard
conditions and treated with EDTA (3), whereas 1.74 or 1.83 equiv of Ni was incorporated into
the sample that was not treated with chelator (3, 4). When using (UreABC*)3, obtained by
mixing UreB with (UreAC)3 before activation, the metal content of the EDTA-treated sample
was 1.00 Ni per heterodimer, whereas samples not treated with chelator possessed 2.56 equiv
of metal. In contrast, (UreAC)3 subjected to activation conditions bound only 0.10 equiv of
nickel if treated with chelator but still bound 2.57 equiv in the absence of EDTA. Finally,
(UreAC)3 that was activated and then mixed with UreB contained 0.18 equiv of nickel with

140

EDTA treatment and 2.78 equiv of metal without the chelator.
Table 4.1: Metal content of urease apoprotein samples subjected to activation conditions
b
Complex
Ni content - EDTA
Ni content + EDTA
(UreABC*)3

2.56 ± 0.27

(UreAC)3

0.10 ± 0.03

2.57 ± 0.36

(UreAC)3 then add UreB
a

1.00 ± 0.04
0.18 ± 0.04

a

2.78 ± 0.17

Activation conditions involved incubating the protein for 1 h at 37 °C in 100 mM HEPES (pH
b

8.3), 150 mM NaCl, 100 mM NaHCO3, and 100 µM NiCl2. Ni content per UreC subunit.
These results indicate that UreB plays a vital role in protecting the Ni in the active site
from chelation, as well as insuring it is incorporated correctly. Only when UreB is present
before activation is the Ni content still high after EDTA is added (1 equivalent of Ni per UreC
compared to 0.1 and 0.18 equivalents when UreB is not present or added after activation).
Adding UreB to activated (UreAC)3 is not sufficient; UreB cannot trap a pre-formed
metallocenter but must be present during the activation process.
Crystallization Attempts
In order to understand the urease activation process in greater detail, I made several
attempts to crystallize selected proteins. The structure of UreF was unknown when I began my
thesis research and there is still no structure for UreG, so both of these accessory proteins were
targets for crystallization. In addition, the urease variant containing G11P UreB was known to
be a low activity mutant (where G11 is in a flexible region of UreB proposed to be important for
the conformational change needed for activation) (2), and this protein was studied to further
examine the potential implications of UreB movement in urease activation. All microbatch
crystallization efforts were carried out using an Oryx4 robot from Douglas Instruments
overseen by Professor Michael Garavito.

141

UreE-F. Purified UreE-F (5) (9 mg/mL in 20 mM Tris, pH 7.8, 300 mM NaCl, 1 mM EDTA,
and 10% glycerol) was used for microbatch screening using two commercially available sets of
96 conditions each (Hampton Research). One condition (0.1 M HEPES, pH 7.5, 4.3 M NaCl)
produced needle-shaped crystals after 2 weeks at 20 °C. Twenty-four sitting drop conditions
were set up by varying the condition that produced crystals by varying the pH by 1 unit in each
direction and reducing the concentration of NaCl in 0.2 M increments to 2.0 M in order to get a
different crystal form, but those attempts were not successful.
UreG. Purified UreG (6) (10 mg/mL) in 100 mM Tris, pH 8.0, 150 mM NaCl and 1 mM
EDTA was used for microbatch crystal screening under the same initial screens as UreF, but no
crystals were found after one month under these conditions.
(UreAB(G11P)C)3. Purified enzyme (2) (18 mg/mL) in 20 mM Tris, pH 7.0, 1 mM EDTA,
and 1 mM DTT was used for microbatch crystal screening using the same microbatch screens,
which included the condition that was used to crystallize urease (7). One condition (0.2 M
sodium acetate trihydrate, 0.1 M Tris-HCl, pH 8.5, 30% w/v PEG 4000) resulted in small crystals
after one week at 20 °C. Sitting drop experiments were set up by varying the concentrations of
the sodium acetate trihydrate, Tris-HCl, and PEG 4000; however, the crystals in the original
condition were not reproducible.
XAS studies on UreG
UreG binds one Ni or one Zn ion per monomer (Chapter 2), but mutagenesis efforts to
determine the ligands that bind the metal were largely unsuccessful. As another method to
learn about the ligands for the metal binding site in UreG, I provided samples to Professor
Michael Maroney at University of Massachusetts for XAS (X-ray absorption spectroscopy)

142

analysis. XAS is a method that can determine the number and types of atoms that ligate the
metal as well as their distance from the metal.
UreG and UreGStr were purified as described in chapter 2. Proteins were dialyzed into
buffer containing 50 mM HEPES (pH 7.4) and 200 mM NaCl overnight. 1 mL samples of 100 µM
UreGStr or UreG were dialyzed overnight into 250 mL of the same buffer that also contained
200 µM NiCl2 or ZnCl2. The next morning the samples were concentrated to 0.5 mL and, in the
same step involving a G-25 column (GE Healthcare), excess metal was removed and 25%
glycerol was added. The eluate was concentrated to 70 µL by using 0.5 mL Amicon 10,000
MWCO ultracentrifugal filters. A portion (20 µL) of the concentrated protein was used to assess
protein and metal concentrations and 50 µL was flash-frozen by using liquid nitrogen and
stored at -80 °C until they could be shipped. Samples were analyzed for protein concentration
by using a Nanodrop spectrophotometer (the extinction coefficients of each protein are 22,430
cm

-1

-1

M for UreGStr and 16,740 cm

-1

M

-1

for UreG) and for metal content by using PAR (8).

The Zn samples had a significant amount of precipitation and were not concentrated enough to
send for XAS analysis. Table 4.2 describes the samples sent. XAS data have been collected for
the samples, but analysis is still in progress.
Table 4.2: Samples sent for XAS analysis
Sample
Protein Concentration
UreG
1560 µM
885 µM
UreGStr

Ni concentration
770 µM
600 µM

In vitro pull-down assays with UreE-F and MBP-UreD
In addition to the studies described in Chapter 3 that involved mixing UreE-F and its

143

variants with (UreABC—UreD)3 or UreGStr and pulling out the complexes that bind NTA or
Strep-tactin resin, respectively, to determine protein:protein interactions, I also carried out
amylose resin pull-down studies with MBP-UreD. These studies were done at 37 °C (where the
interaction between UreD and UreF was weak) and at 42 °C (where a ratio of 1.8:1 MBPUreD:UreE-F was seen) (9). Unfortunately, as described below, these studies were difficult to
interpret and did not add to our knowledge of the UreD—UreF interface.
Interactions between UreF and UreD were analyzed by using purified MBP-UreD and
UreE-F. The UreE-F fusion protein and its variants (10 μM) were incubated with 2 μM MBPUreD at 37 °C (a temperature that leads to a lesser amount of interaction) or 42 °C (a
temperature that was previously shown to promote maximal interaction) for one h, added to
amylose resin at 1/5 the volume of the assay, rocked for one h at room temperature, washed
with five fold the assay volume, and eluted in buffer containing 10 mM amylose (9). Assays
were analyzed by 12% SDS-PAGE and gel scanning (Alpha Imager 2200), with the intensities of
the bands divided by the relative molecular mass of each protein (MBP-UreD, 72.9 kDa; UreEF,
42.8 kDa) to assess the ratios of interactions.
The interactions between MBP-UreD and wild-type or variant UreE-F proteins were
analyzed at two temperatures. Samples incubated at 37 °C yielded a 0.4:1 ratio of MBP-UreD to
UreE-F using the wild-type fusion protein, and, surprisingly, the ratio increased in half of the
experiments when using mutant protein samples. Samples incubated at 42 °C yielded a 1.8:1
MBP-UreD to UreE-F ratio using the wild-type protein, and the ratio increased for all but three
of the mutants proteins (E94A was similar to wild type, whereas K165A and Q171A were slightly
lower) (Table 4.3). The large binding ratios were attributed to MBP-UreD aggregation; thus, no

144

further conclusions could be discerned from the data. The crystal structure of (UreH—UreF)2
revealed that E215 was the only residue changed to alanine that was involved in the UreD-UreF
interface, further confirming concerns about aggregation in MBP-UreD.
a

Table 4.3: Binding of control and mutant UreE-F to MBP-UreD
UreE-F
Ratio of UreE-F/MBP-UreD at Ratio of UreE-F/MBP-UreD at
mutation
37 °C
42 °C
wt
0.41
1.81
P19A
0.58
2.39
G21A
0.94
2.41
Y23A
0.65
2.64
S26A
0.51
2.66
E30A
0.18
2.49
E94A
0.99
1.89
K165A
1.22
1.24
Q171A
1.00
1.39
H214A
0.25
2.09
E215A
0.25
2.21
R220A
0.31
2.91
L221A
0.36
2.98
F222A
0.42
3.12
S224A
0.29
2.94
a

Determined by in vitro pull-down studies using amylose resin and gel scanning

Characterization of MBP-UreD—UreF—UreG
GTPase activity and contamination. In order to investigate the hypothesis UreF acts as a
GAP for the GTPase UreG (see chapter 3), MBP-UreD—UreF—UreG and MBP-UreD—UreF—
UreG(T21A) (as a control that cannot hydrolyze GTP) were purified and their GTPase activities
were characterized. Experiments detailed below showed this complex had a contaminating
GTPase that was not removed by the purification steps, so a His-tagged version of MBP-UreD—
UreF—UreG (with the His-tag located at the amino terminus of MBP) along with the T21A UreG
and K165A UreF variant complexes were tested. This complex also had GTPase contamination,

145

so another method for testing the hypothesis had to be found. This section details the
purification and testing of the two complexes.
MBP-UreD—UreF—UreG was purified as described previously from E. coli BL21 (DE3)
cells containing plasmids pEC002 (encodes MBP-UreD) and pEC005 (encodes UreF and UreG)
(9). His-MBP-UreD—UreF—UreG was purified from E. coli BL21 (DE3) cells containing pCDFMBP-UreD (created by Nicholas Flugga, a construct that adds a His-tag to the N-terminus of
MBP) and pEC005. Cells containing pCDF-MBP-UreD and pEC005 were grown at 37 °C in 1 L of
-1

-1

LB containing 100 µg mL of spectinomycin and 45 µg mL of chloramphenicol to an O.D.600 of
0.4 to 0.6, and induced with 0.5 mM IPTG overnight at 22 °C. Cells were harvested by
centrifugation, resuspended in buffer A (20 mM Tris, pH 7.8, containing 200 mM NaCl and 20
mM imidazole), sonicated, and centrifuged at 100,000 × g for 1 h. Cell-free extracts were
loaded onto a 5 mL Ni-NTA column, washed until the A280 reached baseline, and eluted with
buffer B (20 mM Tris, pH 7.8, containing 200 mM NaCl and 1 M imidazole). Protein was pooled
and dialyzed into TEB (20 mM Tris pH 7.8, 1 mM EDTA, and 1 mM β-mercaptoethanol) with 25
mM NaCl. Protein was then loaded onto a 20 mL amylose column, washed with TEB plus 25
mM NaCl, and eluted with buffer containing 10 mM maltose. Fractions containing the protein
complex were pooled and loaded onto a Superdex-200 column (65 cm × 2.0 cm diameter; GE
Healthcare) equilibrated in TEB plus 100 mM NaCl. Fractions eluting off this column were
tested for GTPase activity.
Complexes containing the K165A variant of UreF were made by digesting plasmid
pKK17-K165A with AatII and AvrII and ligating into a similarly digested pEC005 (9), forming

146

plasmid pEC005-UreF-K165A. Complexes containing T21A UreG were formed by digesting
plasmid pKAU17T21A (10) with AatII and RsrII and ligating into similarly digested pEC005 (9),
forming plasmid pEC005-UreG-T21A. These plasmids were double transformed with either
pEC002 (for forming MBP-UreD—UreF—UreG variant complexes) (9) or pCDF-MBP-UreD (for
forming His- MBP-UreD—UreF—UreG variant complexes).
To determine how much phosphate was produced during GTPase assays, a malachite
green reagent was utilized (11). A 100 µL fraction was added to 900 µL of buffer (100 mM
Hepes, pH 7.8, and 25 mM NaCl) containing 500 µM Mg2GTP and incubated at 37 °C for the
indicated amount of time, then 100 µL was removed and added to 700 µL of buffer, with 200 µL
of malachite green dye subsequently added. The samples were vortexed, transferred to 1 mL
cuvettes, and the absorbances at 620 nm were read after 5 min. Samples were compared to a
standard curve with known concentrations of phosphate.
For MBP-UreD—UreF—UreG-containing fractions from the Superdex-200 column,
malachite green assays were used to monitor phosphate release from Mg2GTP. Figures 4.2 and
4.3 show SDS-PAGE gels of the fractions tested for the wild-type and T21A UreG-containing
complexes, respectively. Tables 4.4 and 4.5 show the changes in phosphate concentration from
20 to 40 min in the GTPase assays incubated at 37 °C and the protein concentrations of the
fractions as determined by Bradford assay for the same complexes.

147

Figure 4.2: Fractions of MBP-UreD—UreF—UreG tested for GTPase activity. SDS-PAGE analysis
of MBP-UreD—UreF—UreG fractions eluted from a Superdex-200 column.
Table 4.4: GTPase activities of MBP-UreD—UreF—UreG samples. Fractions assessed for
GTPase activity can be visualized in Figure 4.2. Protein concentrations were determined by
Bradford assays. Incubations contained 100 µL of sample and were incubated at 37°C for up to
40 min. Malachite green assays were performed after 20 and 40 min and the differences in
phosphate concentrations (ΔPO4) between the pairs of analyses were recorded. If the
difference was calculated to be negative, the ΔPO4 concentration was rounded up to zero.
-1
[Protein]
Δ [PO4]
[PO4]/(mg ml )
Fraction
blank
21
25
26
27
28
29

(20 to 40 min) (µM)
0
24
0.55
0
8.07
4.14
1.09

-1

(mg mL )
0
0.06
0.08
0.31
0.45
0.27
0.04

0
400.00
6.88
0.00
17.93
15.33
27.25

148

Figure 4.3: Fractions of MBP-UreD—UreF—UreG(T21A) tested for GTPase activity. SDS-PAGE
analysis of MBP-UreD—UreF—UreG(T21A) fractions eluted from a Superdex-200 column.
Table 4.5: GTPase activities of MBP-UreD—UreF—UreG(T21A) samples. Fractions assessed for
GTPase activity can be visualized in Figure 4.3. Protein concentrations were determined by
Bradford assays. Assays contained 100 µL of sample and were incubated at 37 °C for up to 40
min. Malachite green assays were carried out after 20 and 40 min and the differences in
phosphate concentrations between pairs of samples were calculated. If the difference was
negative, the ΔPO4 concentration was rounded up to zero.
-1
[Protein]
Fraction
[PO4]/(mg ml )
Δ[PO4]
Blank
20
21
22
23
24
25
26
27
28
29

(20 to 40 min) (µM)
0
5.33
11.58
8.75
0
0
5.85
9.34
5.22
4.76
0

-1

(mg mL )
0
0.02
0.10
0.15
0.13
0.12
0.21
0.45
0.34
0.16
0.06

0
266.50
115.80
58.33
0.00
0.00
27.86
20.76
15.35
29.75
0.00

As seen in Figures 4.2 and 4.3 and Tables 4.4 and 4.5, the maximal GTPase activity did
not coincide with the fractions containing the most MBP-UreD—UreF—UreG complex and the
sample containing the T21A UreG mutation (that should be incapable of hydrolyzing GTP) had

149

significant activity. These results confirm the presence of a contaminating GTPase in these
samples. In order to overcome this concern, His-MBP-UreD—UreF—UreG as well as that
complex containing the K165A UreF or the T21A UreG mutations were purified and tested for
GTPase activity in a similar way as the non His-tagged complex, except that the phosphate
concentration was measured after 1 h incubation with 300 µM Mg2GTP (Figure 4.4 and Table
4.6). Unfortunately, the samples had the same type of GTPase contamination that did not
correlate with the major protein peaks and was similar in the control complex containing T21A
UreG. Additionally, UreG tended to dissociate from the T21A and K165A complexes during the
final step of purification, adding to the difficulty of accurately measuring GTPase activity. These
studies highlight the difficulty of measuring very small amounts of GTPase activities in samples
that contain contaminating amounts of highly active GTPases.

Figure 4.4: Fractions of His-MBP-UreD—UreF—UreG, His-MBP-UreD—UreF—UreG(T21A), and
His-MBP-UreD—UreF(K165A)—UreG that were tested for GTPase activity. SDS-PAGE analysis of

150

(Figure 4.4 con’t) fractions eluted from a Superdex-200 column, with lanes labeled with fraction
number above the brackets and protein below (wt is the wild-type complex, T is the form
containing T21A UreG, and K is the species containing K165A UreF.)
Table 4.6: GTPase activity of His-MBP-UreD—UreF—UreG and variants. Phosphate contents
after incubating selected fractions from Superdex-200 containing His-MBP-UreD—UreF—UreG
for 1 h at 37 °C with 300 µM GTP.
-1
Fraction # - protein content [PO4] after 1 h
[protein]
[PO4]/(mg ml )
Blank
20 – wt
20 - T21A
20 - K165A
24 – wt
24 - T21A
24 - K165A
25 - wt
25 - T21A
25 - K165A
31 – wt
31 - T21A
31 - K165A
35 – wt
35 - T21A
35 - K165A

-1

(mg mL )
0
0.62
0.53
0.72
0.36
0.30
0.69
0.49
0.17
0.45
0.03
0.15
0.15
0.13
0.06
0.24

(µM)
12.05
158.70
93.90
141.27
22.99
16.87
30.13
21.97
19.47
42.09
12.98
12.70
12.89
13.44
12.05
14.18

GTP-linked dissociation of MBP-UreD—UreF—UreG.

0
255.97
177.17
196.21
63.86
56.23
43.67
44.84
114.53
93.53
432.67
84.67
85.93
103.38
255.97
177.17

In order to test whether GTP

causes the dissociation of components within the MBP-UreD—UreF—UreG complex, pull-down
assays using amylose resin were carried out in the presence and absence of 400 µM GTP. MBPUreD—UreF—UreG (2 µM) was incubated in buffer containing 100 mM Hepes, pH 7.8, and 25
mM NaCl, with or without 400 µM GTP for 1 h at 37 °C. Those samples were added to 150 µL of
amylose resin which had been equilibrated in the same buffer, rocked at room temperature for
1 h, washed with five fold the assay volume, and eluted in 10 mM maltose. Figure 4.5 shows
the results of these two assays. GTP had no effect on the amounts of UreF and UreG bound to

151

the MBP-UreD, indicating that incubation with GTP does not cause the accessory proteins to
dissociate from each other in this time frame.

Figure 4.5: Effect of adding GTP to MBP-UreD—UreF—UreG. Lanes: 1-4, no GTP added to
starting material, first wash, final wash, and elution; lanes 5-8 are the same as 1-4 except for
the assay with GTP.
Characterization of UreABC—UreD—UreF—UreGStr
Superdex-200 profile. As a final purification step, UreABC—UreD—UreF—UreGStr was
concentrated and chromatographed on a Superdex-200 column equilibrated in 100 mM Hepes,
pH 7.8, 150 mM NaCl, and 1 mM β-mercaptoethanol. Three peaks and a shoulder were evident
in the chromatogram. The largest molecular weight peak and the shoulder correspond to the
full complex, the middle peak to (UreABC)3, and the third to UreGStr (Figure 4.6).

152

A

B

Figure 4.6:

Gel filtration profile of UreABC—UreD—UreF—UreGStr.

A. Chromatograph

produced using concentrated UreABC—UreD—UreF—UreGStr. B. SDS-PAGE of fractions from
the labeled peaks on the chromatograph. Peaks 1 and 2 correspond to the complex, peak 3 to
(UreABC)3, and peak 4 to UreGStr.

153

Native gel for UreABC—UreD—UreF—UreGStr. To compare the quaternary structure of
UreABC—UreD—UreF—UreGStr with that of the non-tagged complex (10), I used a Blue native
gel (3-12% gradient; Invitrogen). The buffers recommended by Invitrogen were utilized and the
gel was run at 150 V for 90 min. The gel (Figure 4.7) showed that the Strep-tagged complex’s
smallest molecular weight form migrated at the same size as the non-tagged complex, but
some of the larger molecular weight bands were not present in the tagged complex. The Streptag may interfere with larger complex formation, or cause the complex to be in a slightly
different conformation that does not allow for the same oligomeric states. This change in
quaternary structure does not affect the ability of the tagged complex to activate, however.

Figure 4.7: Native gel of UreABC—UreD—UreF—UreGStr and UreABC—UreD—UreF—UreG.
Lane 1: Purified UreABC—UreD—UreF—UreGStr. Lane 2: Same as lane 1, but protein was
concentrated three fold. Lane 3: UreABC—UreD—UreF—UreG. Lane 4: Same as 3, but protein
was concentrated two fold.

154

Stability of UreABC—UreD—UreF—UreGStr.

To quantify how much phosphate

precipitates with the UreABC—UreD—UreF—UreGStr protein as a control for experiments in
Chapter 3, the complex (10 µM) was mixed with 100 µM PO4 and boiled for 3 min. Unlike the
samples that had been activated with Ni, bicarbonate, and various concentrations of GTP, this
sample did not precipitate.

This complex was boiled for 7 additional min, and still no

precipitation was noted. This surprising finding was examined further by comparing samples
that had been activated for 1 h with buffer (100 mM Hepes, pH 8.3, 150 mM NaCl), 100 µM Ni
and 100 µM bicarbonate (with and without 200 µM GTP) to a sample that sat at 37 °C for 1 h in
buffer without Ni, bicarbonate, or GTP. All samples were boiled for 5 min, centrifuged, and the
supernatant solutions were electrophoresed (Figure 4.8). The non-activated complex retained
all proteins in the supernatant fraction after boiling, whereas similar treatment of the activated
complex apparently lead to UreD and UreF precipitation, as well as insolubility of most of the
UreC. This experiment demonstrates the surprising thermal stability of the UreABC—UreD—
UreF—UreGStr pre-activation complex, although the reason for this behavior is unknown.

155

Figure 4.8: Thermal stability of UreABC—UreD—UreF—UreGStr. Proteins were activated at 37
°C for 1 h under varying conditions, boiled for 5 min, centrifuged, and the supernatants were
analyzed by SDS-PAGE. Lane 1: Complex activated with 100 µM NiCl2 and 100 µM NaHCO3.
Lane 2: Complex activated with 100 µM NiCl2, 100 µM NaHCO3, and 200 µM Mg2GTP. Lane 3:
Complex activated in only buffer. Lane 4: Complex activated in only buffer, with a smaller
amount of protein loaded onto gel. Lane 5: Non-activated control.

156

CONCLUSIONS AND REMAINING QUESTIONS
This thesis has succeeded in answering some of the prior questions about urease
activation, but in answering these points other questions have arisen.

In chapter 2, I

characterized a Strep-tagged version of UreG, as well as 12 variant proteins in an attempt to
identify the metal-binding residues. Using UreGStr I showed that the protein binds one Ni or
one Zn at the same site with a similar Kd (~3 µM) based on competition assays with

63

Ni. Out

of all the mutations tested, C72A was the only one that had an effect on metal binding, as
shown by equilibrium dialysis and UV-visible spectroscopy. Other efforts are underway (XAS
analysis) to identify the other residues involved in chelating the metal, but the question
remains open. I also demonstrated that UreG and UreE interact in the presence of metal,
which led to the hypothesis that UreE transfers Ni to UreG, which passes it on to UreD and
finally into the active site. This hypothesis needs to be tested in more depth in order to fully
understand the role metal binding in UreG plays in the overall activation process.
In chapter 3, I described 16 alanine mutations of highly conserved residues in UreF. Of
great significance, I identified several residues that play a role in the binding between UreF and
UreG. Since we do not know the structure of UreG, one open question (difficult to answer
without the UreG crystal structure, although potentially accessible to cross-linking analysis) is
which residues on UreG interact with counterparts on UreF? In chapter 2 I showed Asp80 plays
a role in the interaction of UreG with the UreABC—UreD—UreF complex, but there is certainly
more than one residue involved in that interaction. It is challenging to choose other residues in
UreG for mutagenesis when specifically looking for that interaction since so many residues are

157

highly conserved. A broader question is why the binding site for UreG on UreF is so highly
conserved, yet the interface between UreF and UreD is not, as shown in the crystal structure.
In chapter 3 I also described how UreF acts as a “gate” and, using residue K165, allows
for GTP hydrolysis to be connected to urease activation, slowing down the activation process
and leading to protection against incorrect assembly with Zn. This discovery substantially
answers the question of what role GTPase activity plays in urease activation, but it also opens
up several more questions. Is the metal binding site in UreG important for this gating? Since
UreG binds Ni and Zn with approximately equal affinity when purified alone, can it distinguish
between the two metals when it is in complex with UreD and UreF? Is UreF directly involved in
determining the correct metal? Is Zn the only metal for which there is protection, or do other
transition metals such as Fe, Cu, and Co have the same effect? I demonstrated K165 is not the
GAP residue in UreF, but that does not completely rule out a GAP role for UreF. In particular,
R220 is a highly conserved positively-charged residue that could also play that role, a possibility
that is difficult to test because UreG does not bind to the UreABC—UreD—UreF(R220A)
complex.
In conclusion, my studies have greatly furthered our knowledge about UreG and UreF
and their roles in urease activation. There is now a clearer picture of the interaction between
UreG and UreE, and between UreG and UreF. UreG’s metal binding properties have been
partially characterized. The role of GTPase activity in urease activation has been solved, at least
in part; it is a “gate” guarding against incorrect metal incorporation in conjunction with UreF.
UreF’s role in connecting the GTPase to the activation process has been elucidated to a large
extent. Thus, my studies have provided important insights for understanding the overall urease

158

activation process, and this new understanding may be relevant to metallocenter activation in
other enzymes.

159

REFERENCES

160

REFERENCES
1.

Chang, Z. Z., Kuchar, J., and Hausinger, R. P. (2004) Chemical cross-linking and mass
spectrometric identification of sites of interaction for UreD, UreF, and urease, J. Biol.
Chem. 279, 15305-15313.

2.

Quiroz-Valenzuela, S., Sukuru, S. C. K., Hausinger, R. P., Kuhn, L. A., and Heller, W. T.
(2008) The structure of urease activation complexes examined by flexibility analysis,
mutagenesis, and small-angle X-ray scattering, Arch. Biochem. Biophys 480, 51-57.

3.

Yamaguchi, K., Cosper, N. J., Stalhandske, C., Scott, R. A., Pearson, M. A., Karplus, P. A.,
and Hausinger, R. P. (1999) Characterization of metal-substituted Klebsiella aerogenes
urease, J.Biol. Inorg. Chem. 4, 468-477.

4.

Park, I. S., and Hausinger, R. P. (1996) Metal ion interactions with urease and UreDurease apoproteins, Biochemistry 35, 5345-5352.

5.

Kim, J. K., Mulrooney, S. B., and Hausinger, R. P. (2006) The UreEF fusion protein
provides a soluble and functional form of the UreF urease accessory protein, J. Bacteriol.
188, 8413-8420.

6.

Moncrief, M. B. C., and Hausinger, R. P. (1997) Characterization of UreG, identification
of a UreD-UreF-UreG complex, and evidence suggesting that a nucleotide-binding site in
UreG is required for in vivo metallocenter assembly of Klebsiella aerogenes urease, J.
Bacteriol. 179, 4081-4086.

7.

Jabri, E., Carr, M. B., Hausinger, R. P., and Karplus, P. A. (1995) The crystal structure of
urease from Klebsiella aerogenes, Science 268, 998-1004.

8.

Hunt, J. B., Sue H. Neece, and Ann Ginsburg. (1985) The use of 4-(2-pyridylazo)
resorcinol in studies of zinc release from Escherichia coli aspartate transcarbamoylase,
Anal. Biochem. 146, 150-157.

9.

Carter, E. L., and Hausinger, R. P. (2010) Characterization of Klebsiella aerogenes urease
accessory protein UreD in fusion with the maltose binding protein, J. Bacteriol. 192,
2294-2304.

10.

Soriano, A., and Hausinger, R. P. (1999) GTP-dependent activation of urease apoprotein
in complex with the UreD, UreF, and UreG accessory proteins, Proc. Natl. Acad. Sci.
U.S.A. 96, 11140-11144.

11.

Baykov, A. A., Evtushenko, O. A., and Avaeva, S. M. (1988) A malachite green procedure
for orthophosphate determination and its use in alkaline phosphatase-based enzyme
immunoassay, Anal. Biochem. 171, 266-270.

161

Appendix A

162

Table A.1: Plasmids used in this work.
Plasmid

Description

Reference

pASK-IBA3plus

Plasmid for creating fusion proteins
with a Strep-tag II (WSHPQFEK) at
the C-terminus

IBA

pASK-IBA5plus

Plasmid for creating fusion proteins
with a Strep-tag II at the Nterminus

IBA

pEC007

Modified pACT3 to encode UreE

(1)

pKK17

K. aerogenes ureDABCEFG gene
cluster inserted into pKK223-3

(2)

pKAUG-1

Modified pKAUD2 containing only
K. aerogenes ureG.
Modified pASK-IBA5plus to encode

(3)

pIBA5+G

This work

UreGStr
pIBA3+GK20A,
pIBA3+GE25A,
pIBA3+GC28A,
pIBA3+GD33A,
pIBA3+GD49A,
pIBA3+GE68A,
pIBA3+GC72A,
pIBA3+GH74A,
pIBA3+GD80A,
pIBA3+GS111A,
pIBA3+GS115A,
pIBA3+GD120A
pIBA3+GD127A
pKKG

Modified pIBA3+G to encode the
K20A, E25A, C28A, D33A, D49A,
E68A, C72A, H74A, D80A, S111A,
S115A, D120A and D127A variants

Modified pKK17 encoding UreGStr

This work

pKKGK20A, pKKGE25A,
pKKGC28A, pKKGD33A,
pKKGD49A, pKKGE68A,
pKKGC72A, pKKGH74A,
pKKGD80A, pKKGS111A,
pKKGS115A, pKKGD127A

Modified pKKG encoding the K20A,
E25A, C28A, D33A, D49A, E68A,
C72A, H74A, H74C, H74N, D80A,
S111A, S115A, and D127A variants

This work

This work

of UreGStr

of UreGStr

163

Table A.1 (con’t)
pKK17-P19A, -G21A, -Y23A, S26A, -E30A, -D60A, -E94A, K165A, -F169A, -Q171A, H214A, -E215A, -R220A, L221A, -F222A, -S224A
pKKEF
pKKEF-P19A, -G21A, -Y23A, S26A, -E30A, -D60A, -E94A, K165A, -F169A, -Q171A, H214A, -E215A, -R220A, L221A, -F222A, -S224A
pET-EF
pET-EF-P19A, -G21A, -Y23A, S26A, -E30A, -D60A, -E94A, K165A, -F169A, -Q171A, H214A, -E215A, -R220A, L221A, -F222A, -S224A
pKKG-UreF-K165A
pKKG-T21A

pKAUD2
pEC005
pEC005-UreF-K165A

pEC005-UreG-T21A

pEC002

pCDF-MBP-UreD

Single site variants of pKK17 for
studying the effects of these UreF
variants on urease activity

This work

Same as pKK17, but with a
(4)
translational fusion of UreE and UreF
Single site variants of pKKEF for in
This work
vivo pull-down studies

Translationally fused ureEF genes
inserted into pET21
Single site mutants of pET-EF for
production of UreE-F variants for in
vitro pull-down studies

(4)
This work

Single site mutant of pKKG encoding This work
the K165A UreF variant
Single site mutant of pKKG encoding This work
the T21A variant of Strep II tagged
UreG
Plasmid for production of (UreABC— (5)
UreD)3
ureFG fragment cloned into pACT3
for production of UreF and UreG
Single site mutant of pEC005 that
encodes the K165A variant of UreF
along with UreG
Single site mutant of pEC005 that
encodes the T21A variant of UreG
along with UreF
ureD cloned into pMal-c2x for
production of UreD fused at its Cterminus to MBP. Used along with
pEC005 for production of MBPUreD—UreF—UreG.
malE-ureD cloned into pCDF-1b to
produce MBP-UreD with an Nterminal His6 tag

164

(1)
This work

This work

(1)

(6)

References

165

REFERENCES
1.

Carter, E. L., and Hausinger, R. P. (2010) Characterization of Klebsiella aerogenes urease
accessory protein UreD in fusion with the maltose binding protein, J. Bacteriol. 192,
2294-2304.

2.

Colpas, G. J., Brayman, T. G., Ming, L. J., and Hausinger, R. P. (1999) Identification of
metal-binding residues in the Klebsiella aerogenes urease nickel metallochaperone,
UreE, Biochemistry 38, 4078-4088.

3.

Park, I. S. (1994) Thesis, in Microbiology, Michigan State University, East Lansing, MI.

4.

Kim, J. K., Mulrooney, S. B., and Hausinger, R. P. (2006) The UreEF fusion protein
provides a soluble and functional form of the UreF urease accessory protein, J. Bacteriol.
188, 8413-8420.

5.

Park, I. S., Carr, M. B., and Hausinger, R. P. (1994) In vitro activation of urease
apoprotein and role of UreD as a chaperone required for nickel metallocenter assembly,
Proc. Natl. Acad. Sci. U.S.A. 91, 3233-3237.

6.

Carter, E. L., Boer, J. L., Farrugia, M. A., Flugga, N., Towns, C. L., and Hausinger, R. P.
(2011) The function of UreB in Klebsiella aerogenes urease, Biochemistry 50, 9296-9308.

166