THE ROLE OF URED IN NICKEL UREASE MATURATION AND SPECTROSCOPIC
INVESTIGATIONS INTO THE IRON UREASE
By
Mark Anthony Farrugia

A DISSERTATION
Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of
Biochemistry and Molecular Biology – Doctor of Philosophy
2015

ABSTRACT
THE ROLE OF URED IN NICKEL UREASE MATURATION AND SPECTROSCOPIC
INVESTIGATIONS INTO THE IRON UREASE
By
Mark Anthony Farrugia
Urease catalyzes the hydrolysis of urea into ammonia and carbamic acid, with the latter
compound decomposing into a second molecule of ammonia and carbonic acid. The enzyme
from Klebsiella aerogenes is composed of three subunits (UreA, UreB, and UreG) and
assembles into a trimer of heterotrimers (UreABC)3. The in vivo formation of its di-nickel,
carbamylated lysine-bridged active site requires four accessory proteins: UreD, UreE, UreF, and
UreG. The sequential binding of UreD, UreF, and UreG to urease or the binding of an accessory
protein complex consisting of UreD:UreF:UreG primes the enzyme for activation by nickel that
is delivered by UreE. Two of my projects focused on determining the role of UreD in this
activation process and characterizing the properties of a soluble UreD:UreF:UreG accessory
protein complex.
I characterized a soluble, urease accessory protein complex containing a translational
fusion of the maltose binding protein with UreD (MBP-UreD), UreF, and UreG (termed MBPUreDFG) that was formed in vivo. This complex bound nickel weakly and existed as a dimer of
heterotrimers in solution, with two UreF protomers located at the interfacial site. (MBPUreDFG)2 dissociated to the heterotrimer as it bound to urease apoprotein or holoenzyme. The
interaction with the apoprotein was disrupted by the presence of nickel and (UreG)2 dissociated
from (MBP-UreDFG)2 in buffer containing GTP, magnesium, and nickel.
I used mutagenesis approaches to examine the interaction of UreD with both urease and
UreF and to explore the function of this protein in the transfer of nickel into the active site of

urease. On the basis of a multiple sequence alighnment and a UreD homology model I generated
from the Helicobacter pylori UreH:UreF:UreG crystal structure (UreH is homologous to UreD),
26 residues were selected for substitution. None of the variants were affected in their
urease:UreD and UreD:UreF interactions in vivo. In vivo activation studies using UreD variants
produced in the context of the complete urease gene cluster identified the D63A, D63Q, S85K,
D142A, E176A, and E176Q UreD variants as being deficient in urease activation. The
substituted residues mapped to a buried water tunnel identified in silico which originates at the
putative UreF:UreD interface and exits at the opposite face of UreD. Purified urease activated in
vivo by these variants contained substoichiometric amounts of nickel and varied amounts of zinc
and iron per UreABC.
My final project focused on characterization of the iron-containing urease from
Helicobacter mustelae, UreA2B2, by resonance Raman spectroscopy. Previous studies showed
the diferrous active site of this enzyme is rendered inactive in the presence of O 2, yielding a µoxo bridged, diferric metallocenter. In contrast to earlier results, I observed downshifts of the νs
and νas(Fe-O-Fe) following anaerobic reduction in H218O and subsequent chemical oxidation
relative to analysis of enzyme in H216O, consistent with exchange of the µ-oxo atom. The νs(FeO-Fe) was downshifted by 10 cm-1 when the enzyme was incubated with urea or the slowbinding substrate phenyl phosphorodiamidate, but not following incubation with the inhibitor
acetohydroxamic acid or the chaotropic agent guanidinium chloride. Rapid bulk-solvent
exchange studies identified a urea- and solvent-sensitive mode at ~ 530 cm-1 which downshifted
to ~511 cm-1 in D2O or H218O and upshifted by 10 cm-1 in the presence of urea. This result was
consistent with a terminal Fe-OH stretch where the hydroxyl group is not displaced on substrate
binding. I also identified a slow substrate-binding form of UreA2B2.

I dedicate this dissertation to my father, Michael Farrugia, for his unending love and support.
Through his sacrifice and example he made me the man I am today, and for that I owe him the
world.

iv

ACKNOWLEDGEMENTS

I thank my mentor and role-model, Robert Hausinger, for his guidance, support, patience,
and for the numerous professional advancements he has helped me achieve, including my
premier first author-paper and several scholarship and fellowship awards.I thank my family for
providing me a wonderful network of love and support throughout my life and for always
showing interest in my work by asking “What exactly do you do, again?” constantly. I thank my
collaborators Drs. Kristin Parent, Brandon Ruotolo, Denis Proshlyakov, Roman Davydov, and
Edward Solomon, as well as their undergraduate and graduate students for their assistance and
input. Dr. Hausinger provided me the opportunity to mentor several undergraduate students,
Sneha Grandhis, Evan Hsueh, and Adib Abdullah, whose assistance was greatly appreciated. A
special thanks goes out to my lab colleagues, Scott Mulrooney, Tina Müller, Lee Macomber,
Jodi Boer, and Eric Carter, whose insight and inspiration was invaluable. I thank my wife,
Cynthia Lucidi, for her patience and support during my graduate career. Finally, I offer thanks to
the DuVall fellowship for helping to support my graduate education.

v

TABLE OF CONTENTS

LIST OF TABLES ....................................................................................................................... ix
LIST OF FIGURES .......................................................................................................................x
KEY TO ABBREVIATIONS...................................................................................................... xi
CHAPTER 1: Introduction ...........................................................................................................1
Urease Structure and Mechanism. ...........................................................................2
OVERVIEW OF UREASE ACTIVATION. .......................................................................4
UreD (UreH). ...........................................................................................................4
UreF. ........................................................................................................................5
UreG. ........................................................................................................................6
UreD(UreH):UreF:UreG. .........................................................................................7
UreE. ........................................................................................................................8
Iron-Containing Urease. .........................................................................................10
BIBLIOGRAPHY ..............................................................................................................11
CHAPTER 2: Analysis of a Soluble (UreD:UreF:UreG)2 Accessory Protein Complex and
Its Interactions with Klebsiella aerogenes Urease by Mass Spectrometry ..............................18
ABSTRACT .......................................................................................................................19
INTRODUCTION .............................................................................................................20
MATERIALS AND METHODS .......................................................................................22
Preparation of Protein Samples. .............................................................................22
General Properties of MBP-UreDFG.....................................................................23
Chemical Cross-Linking, Proteolysis, and MALDI-TOF-MS of MBP-UreDFG. 23
MS Analysis of Intact Protein Complexes. ............................................................25
RESULTS AND DISCUSSION ........................................................................................26
Characterization of the MBP-UreDFG Complex. .................................................26
MS Analysis of Intact Urease Complexes. ............................................................29
CONCLUSIONS................................................................................................................42
BIBLIOGRAPHY ..............................................................................................................44
CHAPTER 3: Characterization of Klebsiella aerogenes UreD Variants Reveals a Potential
Nickel-Transfer Tunnel for Urease Activation .........................................................................49
ABSTRACT .......................................................................................................................50
INTRODUCTION .............................................................................................................51
MATERIALS AND METHODS .......................................................................................54
UreD Homology Model Generation, Conservation Mapping, and Water Tunnel
Prediction. ..............................................................................................................54
In Vivo Activation of Urease by Variant UreDs. ...................................................56
Affinity Pull-Down Assays Using Soluble Cell-Free Extracts. .............................57
Protein Purification. ...............................................................................................58
Urease Activity Assays. .........................................................................................59
vi

Protein Analyses. ...................................................................................................59
RESULTS ..........................................................................................................................59
Selecting Residues for Substitution in UreD. ........................................................59
Effects of UreD Variants on the In Vivo Activation of Urease..............................67
Pull-Down Assays. .................................................................................................69
Analysis of the Metal Contents and Specific Activities of Urease Samples
Activated In Vivo by Variant UreDs. .....................................................................70
DISCUSSION ....................................................................................................................73
BIBLIOGRAPHY ..............................................................................................................78
CHAPTER 4: Further Insights into the Metallocenter Vibrational Structure of
Helicobacter mustelae UreA2B2 ..................................................................................................84
ABSTRACT .......................................................................................................................85
INTRODUCTION .............................................................................................................87
MATERIALS AND METHODS .......................................................................................91
Purification, Protein Concentration, and Activity Assays of UreA2B2. ...............91
Isotopic Exchange of the UreA2B2 Oxygen Bridge with Bulk Solvent. ..............91
The Effects of Substrate, Chaotropic Agents, and Inhibitors on the Raman
Spectrum of UreA2B2. ..........................................................................................92
Effect of pH on the Rate of Urea-Induced Shifts on UreA2B2. ............................93
Resonance Raman Spectroscopy. ..........................................................................93
RESULTS ..........................................................................................................................94
Resonance Raman Spectra, the Effect of pH, and Identification of Fast and Slow
Substrate-Binding Forms of UreA2B2 at 406.7 nm Excitation. ............................94
Anaerobic Chemical Reduction and Oxidation Facilitates the Complete BridgeExchange for UreA2B2..........................................................................................97
Urea-Induced Shifts are not due to Chaotropic Effects in UreA2B2. ..................100
The Effects of 15N- and 18O-Urea on the Resonance Raman Spectrum
of1UreA2B2. ........................................................................................................102
Rapid Bulk Isotopic Solvent Exchange Provides Evidence of Terminal Solvent
Modes in UreA2B2. .............................................................................................102
DISCUSSION ..................................................................................................................104
BIBLIOGRAPHY ............................................................................................................108
CHAPTER 5: Additional Studies .............................................................................................111
ADDITIONAL K. aerogenes UREASE STUDIES.........................................................112
Equilibrium Dialysis Studies of MBP-UreDFG Using Nickel. ...........................112
Effects of Urease Activation Conditions on MBP-UreDFG Stability. ................113
Interactions Between MBP-UreDFG and UreE. ..................................................118
Crystallization Attempts on MBP-UreDFG and UreABC:UreD.........................120
Native PAGE Analysis of Urease Apoprotein and Its Complexes with MBPUreDFG................................................................................................................124
Stability of MBP-UreDFG Following Tag Removal. ..........................................126
Metal-Catalyzed Oxidation of H144*UreE and MBP-UreDFG. .........................127
Negative Stain Electron Microscopy of Urease:Accessory Protein Complexes. 130
ADDITIONAL UreA2B2 STUDIES ...............................................................................132
vii

Additional Spectroscopic Studies of UreA2B2. ..................................................132
Boric Acid Inhibition of UreA2B2. .....................................................................136
Resonance Raman Spectroscopy on UreA2B2 with Increasing Amounts of
Urea. .....................................................................................................................139
BIBLIOGRAPHY ............................................................................................................141
CHAPTER 6: Concluding Remarks ........................................................................................145
CONCLUDING REMARKS ...........................................................................................146
BIBLIOGRAPHY ............................................................................................................149
APPENDIX .................................................................................................................................151
BIBLIOGRAPHY ............................................................................................................153

viii

LIST OF TABLES

Table 2.1: A list of measured mass determined from primary MS and tandem MS (MS/MS) data,
and sequence masses for the urease related species that appear in Figure 2.6A. ..............39
Table 3.1: Plasmids used in Chapter 3. ..........................................................................................56
Table 3.2: Multiple sequence alignment of UreD homologs used in Consurf conservation
analysis. ..............................................................................................................................60
Table 3.3: Variant forms of KaUreD, primers utilized, rational for the changes, and conservation
scores for the residues. .......................................................................................................67
Table 3.4: Specific activities of urease samples enriched from cells containing selected KaUreD
variants. ..............................................................................................................................72
Table 5.1: Crystal morphology and buffer conditions used in production of crystals from
solutions of UreABC:UreD..............................................................................................123
Table 5.3: Apparent Km and Vmax for UreA2B2 in various concentrations of boric acid. ...........138
Table A.1: Plasmids used in this dissertation. .............................................................................152

ix

LIST OF FIGURES

Figure 1.1: Urease structures. ..........................................................................................................3
Figure 1.2: Structure of UreH:UreF:UreG. ......................................................................................8
Figure 1.3: Hypothetical model of urease activation for K. aerogenes urease. ...............................9
Figure 2.1: Working model of urease activation. ..........................................................................20
Figure 2.2: MALDI-TOF MS analysis of chemically cross-linked MBP-UreDFG. .....................27
Figure 2.3: Interaction of MBP-UreDFG with urease. ..................................................................29
Figure 2.4: Comparing the stability and size differences observed between holo-enzyme and apoprotein complexes by IM-MS. ...........................................................................................31
Figure 2.5: IM-MS data for the urease-related species that appear in Figure 2.6. .........................32
Figure 2.6: Intact complexes of MBP-UreDFG and urease as probed through ESI-MS. ..............34
Figure 2.7: Mass spectrum recorded for MBP-UreD:UreF:UreG at pH 6.9. ................................35
Figure 2.8: A likely assembly pathway for the urease activation complex. ..................................35
Figure 2.9: Protein complex identification through precise mass measurements enabled by
tandem MS. ........................................................................................................................38
Figure 3.1: Homology model-guided mutagenesis of K. aerogenes UreD....................................65
Figure 3.2: Urease activity of cell-free extracts from cells containing KaUreD variants..............69
Figure 3.3: Interactions of KaUreGStr with other K. aerogenes urease proteins. ...........................70
Figure 3.4: Purified urease activated in vivo using variant KaUreDs. ...........................................72
Figure 4.1: Structural comparison of iron- and nickel-dependent ureases. ...................................89
Figure 4.2: Representative resonance Raman absolute and difference spectra of UreA2B2 (blue)
and sample that had been incubated with 60 mM urea (red). ............................................95
Figure 4.3: Rate of urea-induced Raman downshift at different pH values. .................................96
Figure 4.4: Evidence of slow substrate binding to UreA2B2. .......................................................97

x

Figure 4.5: The effect of anaerobic chemical reduction and oxidation of UreA2B2 in H218O
solvent, in the presence of absence of substrate. ...............................................................99
Figure 4.6: Examination of the basis for the urea-induced Raman shifts on UreA2B2. .............101
Figure 4.7: The effects of isotopically-labeled urea on the resonance Raman spectrum of
UreA2B2. .........................................................................................................................102
Figure 4.8: The effect of rapid bulk solvent exchange on 16O- or 18O-bridged UreA2B2 in the
presence or absence of substrate. .....................................................................................103
Figure 5.1: Nickel binding to MBP-UreDFG as determined by equilibrium dialysis. ................113
Figure 5.2: The effects of urease activation components on the stability of MBP-UreDFG and
MBP-UreD:urease apoprotein. ........................................................................................115
Figure 5.3: Size-exclusion chromatography of MBP-UreDFG exposed to various activation
conditions. ........................................................................................................................117
Figure 5.4: The effects of nickel and zinc on MBP-UreDFG:UreE interactions.........................119
Figure 5.5: Size-exclusion chromatography of MBP-UreDFG. ..................................................121
Figure 5.6: Blue native PAGE analysis of UreABC and UreABC:MBP-UreDFG complexes. ..125
Figure 5.7: Factor Xa cleavage of MBP-UreDFG. ......................................................................127
Figure 5.8: Mass spectra of pepsin digests from MCO-treated or untreated H144*UreE. ..........129
Figure 5.9: General shape comparison of the structures of K. aerogenes urease obtained by X-ray
crystallography (left (18)) and negative-stain electron microscopy (right). ....................131
Figure 5.10: EPR studies of UreA2B2 subjected to radiolytic reduction. ...................................134
Figure 5.11: Low-temperature MCD spectra of anaerobic, dithionite-reduced UreA2B2. .........136
Figure 5.12: Kinetic analysis for boric acid inhibition of UreA2B2. ..........................................138
Figure 5.13: Resonance Raman spectra for samples of UreA2B2 incubated with varied
concentrations of urea. .....................................................................................................140

xi

KEY TO ABBREVIATIONS
BS3, bis(sulfosuccinimidyl)suberate
CD, circular dichroism
CID, collision induced dissociation
CHES, 2-(cyclohexylamino)ethanesulfonic acid
EDTA, ethylenediaminetetraacetic acid
EPR, electron paramagnetic resonance spectroscopy
HEPES, 4-(2-hydroxyethyl)piperazine-1-ethanesulfonic acid
GTP, guanosine-5'-phosphate
ICP-AES, inductively coupled plasma-atomic emission spectroscopy
IPTG, isopropyl β-D-1-thiogalactopyranoside
MALDI-TOF MS, matrix-assisted laser desorption ionization time of flight mass spectrometry
MBP, maltose binding protein
MCD, magnetic circular dichroism
MES, 2-(N-morpholino)ethanesulfonic acid
MME, monomethyl ether
MPD, 2-Methyl-2,4-pentanediol
PEG, polyethylene glycol
TAPS, N-tris(hydroxymethyl)methyl-3-aminopropanesulfonic acid
Tris, 2-amino-2-(hydroxymethyl)-1,3-propanediol or (tris(hydroxymethyl)aminomethane)
SDS-PAGE, sodium dodecyl sulfate-polyacrylamide gel electrophoresis
SPG, succinic acid-phosphate-glycine
VTVH, variable-temperature, variable-field
xi

U, µmol of urea degraded min-1
UTR, untranslated region

xii

CHAPTER 1

Introduction

Sections of this chapter were adapted from the publication “Biosynthesis of the Urease
Metallocenter” in The Journal of Biological Chemistry 288:13178-13185 by Mark A. Farrugia,
Lee Macomber, and Robert P. Hausinger.

1

The metalloenzyme urease catalyzes the first step in urea hydrolysis, the conversion of
substrate to ammonia and carbamic acid, with the latter compound spontaneously decomposing
to a second molecule of ammonia and bicarbonate (12, 19). This enzyme is found in all plants as
well as a few Archaea and selected species of bacteria, fungi, and algae (18, 41, 53). Urease from
jack bean seeds was the first enzyme to be crystallized (59) and the first enzyme shown to
contain nickel that is essential to catalysis (19). Because urease activity is associated with several
pathogenic microorganisms, studies related to its inhibition have the potential to yield advances
in human health (6, 28, 40). Similarly, since urea is an important metabolite in plants and is often
used as a nitrogen fertilizer, the inhibition of urease from plants and in soils has important
impacts on agriculture (41, 60). In addition to studies of the enzyme itself, investigations into the
activation of urease have highlighted a network of accessory proteins involved in the cellular
uptake (27) and delivery of nickel into the urease active site (9, 35, 43-45). At the beginning of
my studies, the general protein network for urease in vivo maturation and the kinetic properties
and roles of urease in a variety of systems had already been determined (12, 42, 55). The
following paragraphs summarize pertinent aspects of this prior set of knowledge and selected
examples of more recent findings.
Urease Structure and Mechanism. Ureases from several organisms have been studied
in detail (22, 32, 37) including crystallographically-characterized examples from Klebsiella
aerogenes (30, 49, 50) and Sporosarcina (formerly Bacillus) pasteurii (4, 5) (each containing
three subunits in a (UreABC)3 configuration, Figure 1.1A), Helicobacter pylori (29) (for which a
fusion of genes corresponding to K. aerogenes ureA and ureB results in only two subunits
yielding a ((UreAB)3)4 structure, Figure 1.1B), and the jack bean enzyme (1) (with a single
subunit representing a fusion of all three bacterial subunits and forming back-to-back

2

homotrimers, ((α)3)2, Figure 1.1C). The metallocenter structures of these proteins are identical
(Figure 1.1D), with two Ni2+ ions bridged by a carbamylated Lys residue and water; one metal
(Ni#1) additionally coordinates two His residues and a terminal water molecule, while the
second Ni2+ ion (Ni#2) coordinates to two His residues, one Asp residue, and a terminal water
molecule. Aspects of the enzyme mechanism remain controversial (10, 12, 20, 21, 62), but most
proposals suggest that the urea carbonyl oxygen displaces the terminal water from the Ni#1
while a second Ni2+-bound water molecule (generally thought to be the bridging solvent) acts as
a nucleophile to achieve catalysis. The tetrahedral intermediate then decays, with a nearby His
residue serving as a general acid according to some proposals (50).

Figure 1.1: Urease structures. (A) Three-subunit bacterial ureases (UreA, red; UreB, blue;
UreC, green; with two more copies, yellow) assemble into a trimer of trimers (Protein Data Bank
code 1FWJ). (B) Two-subunit Helicobacter ureases (a fusion of the two small domains, blue;
large subunit, green; with two more copies, yellow) form a trimer of dimers, which interacts with
three more trimers (gray surface view) to form a dodecamer of dimers (code 1E9Z). (C) Single
3

Figure 1.1 (cont’d): subunit urease of fungi and plants (a fusion of all three domains, green;
with two more copies, yellow) forms a trimer that stacks back-to-back with a second trimer (gray
surface view) (code 3LA4). (D) Dinuclear Ni2+ metallocenter of urease (Ni2+, magenta; solvent,
red).
OVERVIEW OF UREASE ACTIVATION.
Our understanding of urease activation predominantly comes from investigations
involving the heterologous expression of K. aerogenes urease genes in Escherichia coli. The K.
aerogenes urease gene cluster (ureDABCEFG) encodes the urease structural enzyme subunits
UreA, UreB, and UreC along with the accessory proteins UreD, UreE, UreF, and UreG (35). The
accessory proteins are required for proper insertion of the two divalent nickel ions into each
nascent active site, which is located within UreC (24). Complete maturation of the active site
also includes carbamylation of a conserved lysine positioned between the two Ni 2+ ions, though
it is unclear whether this process is driven by the accessory proteins (47). Both the unmetallated
and metallated forms of K. aerogenes urease (termed apo- and holourease or apo- and
holoUreABC, respectively) associate as a trimer of heterotrimers (UreABC) 3 (30, 31). The roles
of the urease accessory proteins in synthesis of the enzyme’s buried metallocenters are described
below.
UreD (UreH). UreD forms a soluble complex with apourease in vivo (45). Twodimensional gel electrophoresis showed this complex exists with zero to three UreD bound per
(UreABC)3, with this association being lost upon activation (45). In vitro studies on the
apoUreABC:UreD complex highlighted a role for UreD in enhancing activation, as incubation of
apoUreABC with CO2/bicarbonate plus Ni2+ yields urease specific activity that is only ~15% of
that for wild-type holoUreABC whereas activity corresponding to ~30% of wild-type levels is
obtained for similar incubation of apoUreABC:UreD (45, 48). Chemical cross-linking (16) and
4

small angle X-ray scattering studies (51) demonstrated that UreD interacts with UreC and UreB
at the vertices of the trimer of heterotrimers. In vitro analysis of isolated UreD initially was
stymied by its insolubility when produced in the absence of the structural subunits, but this issue
was overcome by fusing it to maltose-binding protein (MBP) (13). In vitro equilibrium dialysis
studies using this fusion construct showed UreD binds Ni 2+ independent of its association with
the structural subunits of urease (13). Finally, while no crystal structure of the K. aerogenes
UreD exists, structures for the homologous UreH protein from H. pylori have been obtained in
complex with the cognate UreF or with both UreF and UreG bound (25, 26). Overall, previous
studies with UreD led to its hypothetical role as a scaffold protein that forms a complex with
apourease and binds the distal accessory proteins, altogether allowing for nickel transfer into the
nascent urease active site. The structural and functional roles of UreD in protein complexes with
urease are investigated further in Chapter 3 of this dissertation.
UreF. Similar to UreD, K. aerogenes UreF is insoluble when produced heterologously in
E. coli (43). The MBP-UreF (34) and UreE-UreF fusion proteins have been shown to be soluble,
with the latter protein used in studies detailing its role in the cellular activation of urease and in
interacting with UreABC:UreD (33). A complex containing UreABC:UreD:UreF can be directly
isolated from cells expressing the corresponding genes (43). Native gel electrophoresis of
UreABC:UreD:UreF revealed multiple species consistent with zero to three pairs of UreD:UreF
bound per (UreABC)3. Small angle x-ray scattering experiments suggested a close proximity
between UreD and UreF, with both accessory proteins binding in the vicinity of UreB (51).
Chemical cross-linking results support this configuration and also provide evidence for a
conformational change in urease within the UreABC:UreD:UreF complex (16); specifically,
UreB is proposed to undergo a hinge-like motion that enhances access to the nascent active site

5

(11). Following the in vitro activation of UreABC:UreD:UreF by incubation with
CO2/bicarbonate and Ni2+, the urease specific activity is similar to that obtained by activation of
UreABC:UreD; however, much lower concentrations of bicarbonate are required (43). UreF also
serves as the binding site for the UreG GTPase within the UreABC:UreD:UreF complex.
Mutagenesis studies of UreF were used to gauge its role as a GTPase activating protein and gave
evidence that the protein instead acts to gate the GTPase activity of UreG so as to promote
efficient coupling of GTP hydrolysis and metallocenter biosynthesis, thereby enhancing the
fidelity of urease activation (7). Whereas no evidence has been obtained for nickel binding to K.
aerogenes UreF, isothermal titration calorimetry studies using H. pylori UreF that was produced
heterologously in E. coli provided evidence of a nickel-binding role for UreF, suggesting a
potential role in nickel transfer into urease (61). In summary, UreF functions in urease activation
by connecting UreD to UreG and by regulating the GTPase activity of the latter protein to
facilitate nickel transfer into urease.
UreG. In contrast to K. aerogenes UreD and UreF, UreG is soluble in this and other
organisms, and the protein has been characterized from several sources (8, 44, 52, 63-65). K.
aerogenes UreG is a monomer that binds one equivalent of nickel or zinc (Kd ~5 µM for either
metal) (8), whereas the H. pylori protein dimerizes in the presence of zinc (Kd ~0.3 µM,
1/dimer), but not with nickel which binds weakly (Kd ~10 µM, 1.8/monomer). X-ray absorption
spectroscopy of the zinc-bound H. pylori protein reveals a trigonal bipyramidyl site including
two His and two Cys residues (36). This site is buried between the homodimers of UreG and
UreF in the (UreH:UreF:UreG)2 crystal structure from H. pylori (26). As expected from
sequence analysis, UreG is a GTPase; however, the free protein exhibits slow (52, 63, 64) or no
(44, 65) GTPase activity. Substituting a key residue in the P-loop motif of UreG from K.

6

aerogenes or H. pylori abolishes the cell’s ability to make active urease (38, 44), demonstrating
the importance of this GTPase activity when UreG is present in urease activation complexes. A
UreABC:UreD:UreF:UreG complex forms in E. coli cultures expressing the K. aerogenes urease
gene cluster in the absence of nickel (46).
UreD(UreH):UreF:UreG. As an alternative to sequentially adding each accessory
protein to the urease apoenzyme, the UreD:UreF:UreG heterotrimer may bind as a unit to urease.
A UreD:UreF:UreG complex forms in vivo when the corresponding K. aerogenes genes are
expressed independently of the structural components (44); however, this species is poorly
soluble and not well characterized. This solubility problem is overcome in the MBPUreD:UreF:UreG complex, and this species has been shown to bind urease (11, 23). MBPUreD:UreF:UreG contains two copies of each protomer according to gel filtration and mass
spectrometric studies (23). The properties of this complex, including its interactions with urease
and stability in various urease activation conditions, are detailed further in Chapter 2 and Chapter
5 of this dissertation. The structure of the analogous (UreH:UreF:UreG) 2 complex from H. pylori
reveals two UreG promoters binding to one face of the UreF dimer, with each UreG interacting
with both UreF protomers, and with each UreH interacting with a single UreF (Figure 1.2) (26).
GDP is bound opposite of UreF within each UreG, confirming that the former protein is not a
typical GTPase-activating protein. A potential nickel-binding site is deeply buried and bridges
the two UreG molecules, with each protomer providing His and Cys residues. Analysis of this
structure reveals a potential water tunnel extending from the buried nickel-binding site of UreG
through UreF and exiting at UreH, with the authors hypothesizing the tunnel functions in nickel
transfer (61).

7

Figure 1.2: Structure of UreH:UreF:UreG. Shown are two views of the (UreH:UreF:UreG)2
complex from H. pylori (UreH, UreF and UreG in shades of yellow, gray and magenta,
respectively). A GDP molecule (cyan) is located in each UreG.
UreE. Finally, the nickel metallochaperone UreE has been characterized from several
organisms (2, 3, 9, 17). A crystal structure of a functional, truncated form of K. aerogenes UreE
revealed three copper-binding sites per dimer, two sites composed of His110 and His112 within
each protomer and a third interfacial site composed of His96 from each protomer (9, 56). These
copper-binding sites are thought to also bind the relavent target metal: nickel. Mutagenesis
studies demonstrated that only the interfacial site is required for UreE function (17). S. pasteurii
UreE’s crystal structure also highlights a dimeric quaternary structure with a Zn-occupied
interfacial site (presumably substituting for Ni 2+) (4, 57). Structures of several forms of H. pylori
UreE are known, including the nickel-bound species (codes 3NYO and 3TJ8) (2, 54) for which
the Ni2+ is coordinated at the interfacial site with an additional His residue provided by the Cterminus. Overall, the highly soluble UreE proteins are proposed to bind metal ions in the

8

cytoplasm and specifically deliver nickel to urease within the complex of other accessory
proteins.
In addition to the independent protein species, UreE forms UreG:UreE and
UreABC:UreD:UreF:UreG:UreE complexes that may be functionally important. For the H.
pylori components, two UreG protomers bind the UreE dimer, with the interaction stabilized by
Zn2+, but not Ni2+ (3). In contrast, one UreG monomer from K. aerogenes binds to its cognate
UreE dimer, with the interaction stabilized by either Zn 2+ or Ni2+ (8). NMR studies on a complex
of S. pasteurii UreG and UreE were used to map the interaction face, leading to the suggestion
that the UreG binding sites for UreF and UreEare shared (39). Alternatively, using the K.
aerogenes system, a UreABC:UreD:UreF:UreG:UreE complex can be directly isolated from
cells that synthesize a G11P UreB variant (51) or a Strep II-tagged variant of UreG when the
culture contains Ni2+ (8). From this abundance of information on the urease activation machinery
the current hypothetical pathway for urease activation can be illustrated (Figure 1.3).

Figure 1.3: Hypothetical model of urease activation for K. aerogenes urease. The trimer-oftrimers urease apoprotein (UreA, red; UreB, blue; UreC, green) either sequentially binds UreD
(yellow), UreF (gray), and UreG (magenta) or binds the UreDFG complex (only one protomer of
9

Figure 1.3 (cont’d): each protein is shown, but the isolated complex contains two protomers of
each). UreD and UreF protomers depicted are homology models derived from the H. pylori
UreH:UreF crystal structure (PDB 3SF5) while UreG is a homology model derived from
Methanocaldococcus jannaschii HypB (PDB 2HF9). The coordinates were generously provided
by Dr. Célia Carlini for use in this figure. Formation of the active enzyme requires CO2 to
carbamylate Lys217 at the native active site, GTP binding and hydrolysis by UreG, and nickel
delivery by dimeric UreE (cyan). It remains unclear whether the accessory proteins are released
as a UreDFG unit or as individual proteins.
Iron-Containing Urease. A new tool for characterizing urease came in the form of an
iron-containing protein, UreA2B2, from Helicobacter mustelae (15). This organism contains a
conventional H. pylori-like urease gene cluster (ureABEDFGH) which is induced by nickel ions
and a separate two-gene cluster (ureA2B2) that lacks nearby genes coding for accessory proteins
and is inversely regulated by nickel (58). Although the sequences of UreA and UreB are 57%
and 70% identical to UreA2 and UreB2 in H. mustelae, the former protein contains nickel and
the latter possesses divalent iron – thus accounting for its oxygen sensitivity. The crystal
structure of UreA2B2 reveals that its active site residues are conserved with those of the nickel
enzymes (15). Using the oxidized, inactive enzyme, ultraviolet-visible and resonance Raman
spectroscopic studies identified a diferric species with a putative µ-oxo bridge (14, 15). In
particular, resonance Raman studies identified a Fe(III)-O-Fe(III) symmetric vibrational mode
that was shown to be sensitive to the presence of urea, with the bridging oxygen also being
exchange inert to bulk solvent over a long timeframe (> days) (14). Overall, the inactive form of
iron urease allowed for the use of spectroscopic techniques that were previously unavailable for
use in the catalytically active, spectroscopically silent, nickel enzyme. My studies further
characterizing the iron urease are detailed in Chapter 4 and Chapter 5.

10

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11

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17

CHAPTER 2

Analysis of a Soluble (UreD:UreF:UreG)2 Accessory Protein Complex and Its Interactions
with Klebsiella aerogenes Urease by Mass Spectrometry

This chapter is modified from the Journal of the American Society of Mass Spectrometry 24(9):
1328-37 (2013) by Mark A. Farrugia, Linjie Han, Yueyang Zhong, Jodi L. Boer, Brandon T.

Ruotolo, and Robert P. Hausinger. Linjie Han and Yueyang Zhong carried out ion mobility mass
spectrometry in the Ruotolo laboratory and Jodi Boer provided protein samples.

18

ABSTRACT
Maturation of the nickel-containing urease of Klebsiella aerogenes is facilitated by the
UreD, UreF, and UreG accessory proteins along with the UreE metallochaperone. A fusion of
the maltose binding protein and UreD (MBP-UreD) was co-isolated with UreF and UreG in a
soluble complex possessing a (MBP-UreD:UreF:UreG)2 quaternary structure. Within this
complex a UreF:UreF interaction was identified by chemical cross-linking of the amino termini
of its two UreF protomers, as shown by mass spectrometry of tryptic peptides. A pre-activation
complex was formed by the interaction of (MBP-UreD:UreF:UreG)2 and urease. Mass
spectrometry of intact protein species revealed a pathway for synthesis of the urease preactivation complex in which individual hetero-trimer units of the (MBP-UreD:UreF:UreG)2
complex bind to urease. Together, these data provide important new insights into the structures
of protein complexes associated with urease activation.

19

INTRODUCTION
Urease, an enzyme that hydrolyzes urea to form ammonia and carbonic acid (10, 43),
plays important roles in nitrogen cycling and pathogenesis (29, 42). Like most bacterial ureases,
the Klebsiella aerogenes protein contains three subunits (UreA, UreB, and UreC) arranged in a
(UreABC)3 architecture with a dinuclear nickel active site in each of its UreC subunits (21). The
enzyme is synthesized by an elaborate biosynthetic pathway that requires four accessory proteins
(UreD, UreE, UreF, and UreG) along with carbon dioxide, GTP, and the metal ions (10, 14, 43).
A working hypothesis for K. aerogenes urease activation (Figure 2.1) shows the (UreABC)3
apoprotein (25) forming sequential complexes with UreD (32), UreF (30), and UreG (33) or
binding a pre-formed complex of UreD, UreF, and UreG (UreDFG) (31). The resulting preactivation urease complex carbamylates an active site lysine side chain to form a metal-binding
ligand (34) and acquires nickel ions from the UreE metallo-chaperone (26) in a process driven by
GTP hydrolysis (39). Completion of this process is accompanied by dissociation of the accessory
proteins.

Figure 2.1: Working model of urease activation. The urease apoprotein (orange UreA, sand
UreB, and yellow UreC) sequentially binds UreD (green), UreF (magenta), and UreG (cyan) or
binds a complex of these three accessory proteins to form a pre-activation complex. Synthesis of
active enzyme requires carbamylation of an active site lysine by CO 2, GTP hydrolysis by UreG,
20

Figure 2.1 (cont’d): and Ni2+ transfer from the UreE metallochaperone. Structures of the docked
complexes are derived from computational models (27).
The protein complexes in the urease activation pathway have been studied to varying
extents, but many questions about their structures and properties remain to be answered.
Chemical cross-linking of (UreABC)3:UreD followed by proteolysis and matrix-assisted laserdesorption ionization (MALDI) time-of-flight (TOF) mass spectrometry (MS) indicates that
UreD binds to UreB and UreC (12). Small angle X-ray scattering (SAXS) data from this species
are consistent with the UreD density being located at the vertices of the triangular urease
apoprotein (35). Corresponding cross-linking/proteolysis/MALDI-TOF-MS and SAXS analyses
of the (UreABC)3:UreD:UreF species show a UreF linkage to UreB and yield similar positional
results. Moreover, cross-linking and computational flexibility studies provide evidence that UreB
is loosely associated with UreC in this complex and can move in a hinge-like motion that
potentially enhances the nickel ions’ access to the active site (12, 35). (UreABC)3:UreD,
(UreABC)3:UreD:UreF, and (UreABC)3:UreDFG exhibit multiple bands on native gels, and
analysis of these bands on denaturing gels reveals peptide staining intensities consistent with
varied numbers of UreD, UreF, and UreG bound to the (UreABC)3 apoprotein. The structure of
K. aerogenes urease apoprotein is known (22), but no urease:accessory protein complexes have
been structurally elucidated. By contrast, the urease-free structures are known for Helicobacter
pylori UreF (24) and the complex of UreF with UreH (the UreD homologue in H. pylori) (15),
while UreG homology models have been generated by using the related HypB dimeric structure
from Methanocaldococcus jannaschii (16).
Whereas the sequential protein binding urease activation model (Figure 2.1, top) depicts
individual protomers of the accessory proteins binding to the urease apoprotein vertices,
evidence from homologous systems suggests that the pre-formed accessory protein complex may
21

be dimeric. For example, H. pylori UreF and UreH:UreF each possess an overall dimeric
structure (i.e., (UreF)2, (UreH:UreF)2) (15, 24). Furthermore, UreG is dimeric as purified from
some microorganisms (44, 45), although the K. aerogenes protein is monomeric (7, 31). It is
plausible that distinct quaternary structures of the urease accessory proteins are found in K.
aerogenes and H. pylori. Alternatively, it is possible that K. aerogenes also produces a dimeric
accessory protein complex, but it dissociates to the monomeric form when it binds to urease. In
addition, it is feasible that the complexes obtained by sequential binding of accessory proteins
are distinct from those generated by binding of UreDFG. To resolve these questions, I utilized
chemical cross-linking/proteolysis/MS methods to characterize a soluble UreDFG complex from
K. aerogenes, demonstrated that the accessory protein complex binds to urease, and used MS
tuned to retain fragile non-covalent contacts to examine the intact structures of various urease
complexes. The results obtained provide substantial support for the protein complex assembly
model (Figure 2.1 lower path) and present new insights into the stability of various urease
complexes.
MATERIALS AND METHODS
Preparation of Protein Samples. Urease holoenzyme was produced in E. coli BL21Gold(DE3) cells containing pKK17 (the pKK223-3 vector with the complete ureDABCEFG
gene cluster) that were grown in lysogeny broth (LB) with 300 µg ml -1 ampicillin plus 1 mM
NiCl2 and induced at an optical density (600 nm) of ~0.4 with 0.1 mM isopropyl βthiogalactopyranoside. The protein was purified as previously described (11). The (UreABC)3
apoprotein was purified in the same manner from the same E. coli cells that were grown in LB
medium without added NiCl2.

22

The Strep-tag II version of (UreABC)3:UreDFG was formed in cells transformed with
pKKG containing ureDABCEFG where ureG is fused to the codons for Strep-tag II (6). This
complex was purified by use of a Strep-tactin column followed by Superdex 200 gel filtration
chromatography, as previously described (6).
A hetero-trimeric MBP-UreDFG species, containing a maltose binding protein (MBP)
fusion of UreD (MBP-UreD) along with UreF and UreG, was obtained from E. coli BL21Gold(DE3) cells co-expressing pEC005 (with ureFG cloned into pACT3) and pEC002 (with
ureD cloned into pASK-IBA3plus), as previously described (9, 11). The complex was isolated
by sequential amylose, DEAE-Sepharose, and Superdex 200 column chromatography of cell-free
extracts. A version of this species containing the K165A variant of UreF was purified by similar
approaches using cells with pEC005-UreF-K165A (a modified pEC005) and pEC002 (6). MBPUreDFG or its K165A UreF version was mixed with excess urease apoprotein or holoenzyme,
incubated 60 min, and separated by chromatography on amylose resin to remove excess urease.
General Properties of MBP-UreDFG. The native molecular mass of MBP-UreDFG
was assessed by Superdex 200 gel filtration chromatography in 20 mM Tris-HCl buffer (pH 7.5)
containing 1 mM EDTA, 1 mM β-mercaptoethanol, and 25 mM NaCl and using thyroglobulin,
γ-globulin, ovalbumin, and myoglobin (Bio-Rad) as molecular mass markers. The apparent
molecular masses of denatured components in various samples were obtained by sodium dodecyl
sulfate-polyacrylamide gel electrophoresis (SDS-PAGE; 12% running gels) (23), followed by
staining with Coomassie brilliant blue. Integration of band intensities was assessed by gel
scanning (Alpha Innotech Corp., using AlphaEase FC software).
Chemical Cross-Linking, Proteolysis, and MALDI-TOF-MS of MBP-UreDFG.
MBP-UreDFG and its K165A UreF derivative were further characterized by a chemical cross-

23

linking approach. The samples (3.75 µM of the heterotrimer in 20 µl) were incubated with 160fold excess (600 µM) Bis(sulfosuccinimidyl)suberate (BS3) (Thermo Fisher Scientific) in 25 mM
HEPES (4(-2-hydroxyethyl)-1-piperazineethanesulfonic acid, pH 7.4) buffer containing 1 mM
EDTA, 1 mM dithiothreitol and 1% glycerol for one h, then quenched with 15 μl of 1 M glycine.
Untreated and BS3 treated samples were subjected to SDS-PAGE (12% gel) and the
protein bands of interest were excised with a clean razor blade and chopped into ~1-mm3 pieces.
The gel fragments were washed with 100 μl of 100 mM NH4HCO3 buffer (pH 8.4) for 5 min,
dehydrated at room temperature in 50 μl of 100% acetonitrile, and rehydrated in 50 μl of 10 mM
dithiothreitol in 100 mM NH4HCO3 at 56°C for one h. Dehydration and rehydration was
repeated, with final rehydration in 50 μl of 55 mM iodoacetamide in 100 mM NH4HCO3. The
mixtures were incubated at room temperature and in the dark for one h. Samples were repeatedly
washed with 50 μl of 100 mM NH4HCO3, dehydrated, rehydrated in 20 μl 50 mM NH4HCO3
containing 15 ng/μl of sequencing grade trypsin (Promega), and incubated overnight at 37°C.
Supernatants from the trypsin digest were saved, and the gel pieces were sonicated three times
with 50 μl of 60% acetonitrile/1% trifluoroacetic acid in an ice water bath for 5 min followed by
centrifugation. The solutions were pooled with the initial supernatant, dried to ~2 μl in a Speed
Vac, and dissolved in 20 μl of 2% acetonitrile/0.1% trifluoroacetic acid. The final samples were
desalted using C-18 pipette tip columns (Agilent Technologies).
MALDI-TOF MS was performed using a Shimadzu Axima plus instrument equipped
with a nitrogen laser operating at 337 nm. Samples (5 µl) were mixed with 5 μl of a saturated αcyano-4-hydroxycinnamic acid solution (Sigma) in 50% acetonitrile containing 0.1%
trifluoroacetic acid. Aliquots (1 μl) were spotted on a MALDI plate and allowed to dry in a hood.
Spectra were obtained in linear positive mode and externally calibrated with bradykinin fragment

24

(1-7, RPPGFSP, 756.86 Da), angiotensin I (1296.48 Da), and human ß-chain insulin (3495.95
Da). The mass values at the centroid peak positions are reported, i.e. the average value across the
isotopic peaks.
MS Analysis of Intact Protein Complexes. Samples containing UreABC, MBPUreDFG and its K165A UreF version, UreABC:MBP-UreDFG and its K165A UreF version, and
UreABC:UreDFG (where UreG is fused to Strep-tag II) with protein concentrations of 0.5
mg/ml were buffer exchanged into 200 mM ammonium acetate buffer using Micro Bio-Spin 6
columns (Bio-Rad, Hercules, CA) just prior to MS analysis. A quadrupole-ion mobility (IM)TOF MS instrument (Synapt G2 HDMS; Waters, Milford, MA) was used to collect the intact
protein complex data. Protein ions were typically generated by nano-electrospray ionization
(nESI) using a capillary voltage of 1.5-1.7 kV. Ions traveled through a mass-selective quadrupole
and entered an ion trap constructed from a traveling wave ion guide, pressurized at 3×10 -2 mbar
of argon. Ions were then released from the trap cell into the IM separator pressurized with 2.7
mbar of N2. Typical settings for the traveling waves within the IM drift region were 150 m/s
wave velocity and a wave height of 15 V (46). Ions underwent m/z separation in a TOF mass
analyzer, which was maintained at a vacuum of 1.7×10-6 mbar, and operated using a mass range
of 500 to 25,000 m/z. All other instrument parameters were tuned to preserve fragile noncovalent protein interactions (8, 19, 36).
Tandem MS experiments were carried out to increase the confidence of the protein
stoichiometries and compositions assigned from intact mass alone. Ions of interest were
generated in the same fashion as above and then a single m/z was selected in the quadrupole
mass filter. Charge states selected for such dissociation experiments varied from complex to
complex, but typically those signals that presented the least overlap with other protein complex

25

ions having similar m/z and those with the greatest intensity values were chosen for tandem MS.
Following isolation, ions were given excess kinetic energy upon accelerating into the ion trap
region prior to the IM drift cell by increasing the acceleration voltage to 200 V. These data are
presented with minimal smoothing and no background subtraction.
RESULTS AND DISCUSSION
Characterization of the MBP-UreDFG Complex. Of the protein complexes illustrated in
Figure 2.1, the least well studied is UreDFG due to its insolubility (31). To overcome this hurdle,
the maltose binding protein (MBP) fusion with UreD (MBP-UreD, known to be functional for
urease activation in the cell) was used to obtain a soluble hetero-trimeric complex abbreviated
here as MBP-UreDFG (9, 11). As an initial step in characterizing this species, the sample was
digested with thrombin to remove the MBP tag; however, the resulting MBP-free urease subunits
precipitated quickly after (t < 1 hr, data not shown), so further investigations focused on the
properties of the intact MBP-UreDFG complex. SDS-PAGE analysis of purified sample (Figure
2.2A, lane 1) revealed the expected three bands (72,892, 25,222, and 21,944 Da, respectively,
according to their sequences; note that UreG migrates anomalously during SDS-PAGE and is
positioned above UreF), and division of the integrated band intensities by their relative masses
indicated the components were present at nearly equivalent ratios. Gel filtration analysis
provided an apparent relative mass of ~275 kDa (data not shown), consistent with a (MBPUreDFG)2 quaternary structure (predicted sequence mass of 240,116 Da). Furthermore, there
was no evidence of an equilibrium involving a single heterotrimeric species or of dissociation
into the individual protein subunits.

26

Figure 2.2: MALDI-TOF MS analysis of chemically cross-linked MBP-UreDFG. (A) SDSPAGE analysis of samples. Lanes: M, molecular mass markers (in kDa); 1, MBP-UreDFG (19.2
µg); 2, MBP-UreDFG (19.2 µg) treated with BS3 (0.9 µg); 3, MBP-UreD:UreF(K165A):UreG
(9.0 µg); 4, MBP-UreD:UreF(K165A):UreG (9.0 µg) treated with BS3 (6.9 µg). Note that UreG
migrates anomalously slowly for its known mass. (B) MALDI-TOF MS of in-gel trypsin digests
of bands from panel A. Spectra: 1, UreF; 2, ~45 kDa band from cross-linked MBP-UreDFG; 3,
UreF(K165A); 4, ~45 kDa band of cross-linked MBP-UreD:UreF(K165A):UreG. Theoretical
digest fragments of UreF include m/z 921.0 (residues 214-220), 1290.4 (70-81, carbamidomethylated), 1363.6 (181-191, carbamidomethylated), 1683.0 (166-180, absent in samples for
K165A UreF), 1768.0 (109-122, carbamidomethylated), and 2010.3 (192-213). The fragment at
m/z 2211.3 is from trypsin. Cross-links of UreF peptides 2-7 plus 2-7 and peptides 2-7 plus 1-7
have predicted m/z 1523.4 and 1654.6, respectively.
To further characterize the (MBP-UreDFG)2 complex, purified sample was subjected to
chemical cross-linking with the amine-specific reagent bis(sulfosuccinimidyl)suberate (BS3)
followed by trypsin proteolysis and MALDI-TOF-MS. SDS-PAGE analysis of the complex

27

treated with BS3 demonstrated the formation of a species (~44 kDa) that was not present in the
untreated sample (Figure 2.2A, lane 2). A mixture of additional less intense bands corresponding
to intact protein masses greater than MBP-UreD were detected, but not further characterized.
The apparent molecular mass of the ~44-kDa species could be explained by UreF-UreF, UreFUreG, or UreG-UreG cross-links. Digestion of this band with trypsin and MALDI-TOF-MS
analysis of the resulting peptides identified the species as arising solely from UreF,
demonstrating a UreF-UreF cross-link. The MALDI-TOF-MS spectrum of tryptic peptides
derived from untreated UreF was compared to that for BS3-treated sample (Figure 2.2B, spectra
1 and 2). Both samples contained a fragment corresponding to residues 166-180 (m/z = 1683;
LVPFGQQAAQQLILR) arising from cleavage after K165, the only lysine in this protein; thus,
cross-linking did not involve this residue. The BS3-treated sample gave rise to two new tryptic
peptides that were both consistent with cross-linking that involves the amino-termini (with
partial removal of the initial Met); a peak near m/z of 1523 corresponded to two peptides
containing residues 2-7 (m/z = 692; STAEQR) and the linker region (m/z = 140) while that at m/z
of 1655 linked residues 1-7 (m/z = 823) to residues 2-7. Confirmation of the amino-terminal
linkages was obtained by similar studies with a variant complex containing K165A UreF, thus
lacking free lysine residues. A ~44 kDa band derived from BS3 cross-linking again was observed
by SDS-PAGE (Figure 2.2A, lanes 3 and 4). MALDI-TOF-MS analysis (Figure 2.2B, spectra 3
and 4) again revealed the presence of fragments with m/z of 1523 and 1655, but a fragment with
m/z of 1683 was absent, consistent with a lack of proteolytic cleavage after the altered residue.
These results demonstrate a close juxtaposition of the amino termini of two UreF peptides in
(MBP-UreDFG)2.

28

The addition of excess urease holoenzyme or apoprotein to (MBP-UreDFG)2 or its
UreF(K165A) derivative led to the formation of pre-activation complexes that were separated
from free urease by amylose affinity chromatography (Figure 2.3). This procedure allowed for
further structural studies of the pre-activation complex.

Figure 2.3: Interaction of MBP-UreDFG with urease. Lanes: M, molecular mass markers (in
kDa); 1, Urease apoprotein (8.8 µg); 2, MBP-UreDFG (4.7 µg); 3, 1/20th of the elution fraction
(10 mM maltose in 20 mM Tris buffer, pH 7.8, containing 85 mM NaCl and 1 mM EDTA) from
an amylose resin after applying a mixture of urease apoprotein and MBP-UreDFG (207 µg and
41.6 µg, respectively; incubated 1 h, in the same buffer), washing with maltose-free buffer, and
eluting with buffer containing maltose.
MS Analysis of Intact Urease Complexes. To examine the composition and connectivity of
protein complexes along the urease activation pathway in more detail, MS measurements were
conducted on numerous urease-related samples under conditions designed to retain fragile, noncovalent interactions in the gas-phase (4, 5, 18-20, 37, 40). Through the proper tuning of the
nESI process, as well as the correct managing of both pressure and focusing potentials within the
instrument, MS enables the measurement of heterogeneous multiprotein assemblies at low
micromolar concentrations with unprecedented precision and accuracy (17). Errors in this mass
measurement arise primarily from buffer material that adheres to the surface of protein ions
created by gentle nESI, resulting in expected positive deviations between experimentally
29

determined and expected molecular masses of <3% (4, 28). Once sufficient signal intensity is
achieved, tandem MS approaches, involving collision induced dissociation (CID), can be
employed on multiprotein complex ions (2, 3, 38). The fragment ions produced typically take the
form of highly-charged monomeric subunits and charge-reduced assemblies stripped of the
aforementioned monomers (13). If these data are combined with ion mobility (IM) spectrometry,
which enables the separation of protein complex ions based on their orientationally averaged size
and shape, then closely-related assemblies can be individually analyzed and identified (36, 41,
47). While the data shown here were acquired in an IM-MS mode, and in many cases the IM data
were used to guide the MS analysis and differentiate overlapping charge state distributions
resulting from assemblies of similar intact mass, only the resulting MS data and its analysis are
discussed here. Limited IM-MS data comparing the stability differences observed between holoenzyme and apo-protein complexes are available in Figure 2.4, with complexes containing holoenzymes demonstrating superior stability. Sample IM-MS data enabling a more-detailed analysis
of the tandem MS datasets is displayed in Figure 2.5.

30

Figure 2.4: Comparing the stability and size differences observed between holo-enzyme and
apo-protein complexes by IM-MS. (A) Mass spectra for holo-enzyme (top) and apo-protein
(bottom) urease samples containing MBP-UreD:UreF(K165A):UreG indicate that complexes
involving Ni2+ bound species show high signal intensity, demonstrating remarkably superior
stability. (B) Drift time distributions of the equal charge state of holo-enzyme/apo-protein
(UreABC)3 alone and in accessory protein-bound complexes provide side-by-side comparisons
of their sizes. Despite the slight difference in drift time (<1.5%), I observed a trend that the holoprotein complexes get increasingly more compact than the apo-protein forms upon binding of
accessory proteins. While these differences are negligible for U 3 complexes (bottom panel) they
are significant and reproducible for larger assemblies (top two panels). All centroid IM data
shown are derived from Gaussian fits to the observed arrival time distributions and carry a
relative standard deviation of 0.2%. Drift time, (UreABC)3, MBP-UreD, UreF(K165A) and
UreG are abbreviated as DT, U3, MD, F' and G, respectively.

31

Figure 2.5: IM-MS data for the urease-related species that appear in Figure 2.6. A contour
plot of m/z versus drift time is shown. A narrow window that contains 33+ ions of
(UreABC)3:MBP-UreD (No. 7) and 34+ ions of (UreABC)3:MBP-UreD:UreF missing one UreB
subunit is highlighted (dotted box). These two species have similar m/z (~5% difference) and
thus were likely co-isolated in quadrupole during MS/MS acquisition.
To test if the accessory protein complex binds to urease as the dimeric species or whether
one heterotrimer binds while the other dissociates, these components were incubated under
conditions where urease was in three-fold excess and the products were analyzed by MS (Figures
2.6A-E). Figure 2.6E serves as a framework for describing the results. MBP-UreDFG (species
No. 1) in a 200 mM ammonium acetate solution yielded the well-resolved charge state series in
the mass spectrum of Figure 2.6D. This species had an intact mass corresponding to the expected
value for (MBP-UreDFG)2 (240,482 ±21 Da measured and 239,856 Da expected). No indication
of a single heterotrimer, free UreF, or UreF:UreF dimer was detected (see also Figure 2.7). The
observation of this overall dimeric species corroborates the dimerization suggested by gel
filtration analysis and MALDI-MS data that had indicated UreF-UreF cross-linking (Figure 2.2).
Significantly, individual heterotrimeric units of this dimer bind to the core urease assembly
32

(UreA:UreB:UreC)3 (Figure 2.6A, No. 2) to create a (UreABC)3:MBP-UreDFG assembly
(Figure 2.6A, No. 3). A hypothetical pathway for synthesis of this species is illustrated in Figure
2.8.

33

Figure 2.6: Intact complexes of MBP-UreDFG and urease as probed through ESI-MS. (A),
(B), (C) and (D) are mass spectra recorded for various stabilized urease complexes discussed in
the text. (E) Topological diagrams of the complex stoichiometries identified by ESI-MS, where
structures are indicated by numbers 1-10. MBP-UreD, UreF, and UreG are abbreviated as D, F,
and G, except for complex No. 4 which was isolated directly from cells, lacks MBP-tagged
UreD, and has UreF containing Strep-tag II (labeled 4*). Depicted topologies are congruent with
34

Figure 2.6 (cont’d): the experimental observations and other structural information known for
these assemblies. High confidence data from this work strongly indicates that complexes 1 and 2
form primarily complex 3 in vitro. Additional protein complexes are also observed, but likely
result from the dissociation of higher-order complexes, and not from intermediates in the
assembly of urease activation complexes. Complexes No. 9 and No. 10 each have at least two
possible topologies that match the ESI-MS data (as shown).

Figure 2.7: Mass spectrum recorded for MBP-UreD:UreF:UreG at pH 6.9. The charge
states and schematic topologies for the complexes are indicated in matching colors. No free
MBP-UreD:UreF:UreG, monomeric UreF or dimeric UreF is observed under any solution
condition studied here, which strongly supports a urease activation complex assembly
mechanism that does not involve the addition of discrete UreF, and consequently UreG, proteins
to the core complex under in vitro incubation conditions. The spectra are magnified 5 fold above
4400 m/z. Maltose binding protein, UreD, UreF and UreG are abbreviated as MBP, D, F, and G,
respectively.

Figure 2.8: A likely assembly pathway for the urease activation complex. This scheme is
most consistent with the MS data, in which the (UreD:UreF:UreG) 2 complex transiently binds to
(UreABC)3, and then dissociates to leave one UreD:UreF:UreG unit bound to the core urease
complex (i.e., No. 1 plus No. 2 yields No. 3, Figure 2.6E). Successive (UreD:UreF:UreG)2
binding is required to load additional copies of UreD:UreF:UreG onto (UreABC) 3, resulting in
complex No. 4 (Figure 2.6E), as observed for sample directly purified from the cell. One heterotrimer unit of (UreABC)3, UreD, UreF and UreG are abbreviated as U, D, F and G, respectively.

35

Figure 2.6A also provides evidence of other protein complexes that presumably represent
incompletely synthesized, partially decomposed, or aggregated assemblies. These include (but
are not limited to): the (UreABC)3 central complex bound to MBP-UreD:UreF:UreF (No. 6), to
one copy of MBP-UreD (No. 7), or to two copies of MBP-UreD:UreF (No. 10, alternatively
depicted as one copy of MBP-UreD and one copy of MBP-UreD:UreF:UreF), and complex No.
3 with an additional MBP-UreD (No. 8). For a different sample purified directly from the cells
(rather than generated by mixing purified proteins in solution), I detected a complex with two
copies of UreDFG (lacking MBP and with UreG fused to Strep-tag II) bound to urease (No. 4*,
Fig 2.6B). The addition of a second (UreDFG)2 to the complex marked as No. 3, indicated as a
second transient species in brackets in Figure 2.8, is likely needed to satisfy the formation of
several of these assemblies. Alternatively, complexes No. 7 and 8, as well as some possible
topologies of complexes No. 9 and 10 can be formed through the binding of excess MBP-UreD
to smaller assemblies. MS data reveal additional complexes which result from the
oligomerization of (UreABC)3 to form ((UreABC)3)2 (Figure 2.6A, No. 5) and that same
complex bound to two copies of MBP-UreD:UreF (No. 9, alternatively depicted as one MBPUreD and one MBP-UreD:UreF:UreF). Due to the low working concentration of the samples
used in the MS analysis of these intact complexes (<10 µM), it is unlikely that these larger
oligomers can be described as artifacts of the nESI process.
All of the identifications discussed above are supported with tandem MS data, where CID
generates fragment ions that enable the high-confidence assignment of the protein complex
composition and stoichiometry. Figure 2.9 shows selected tandem MS spectra, using the same
numerical identifiers as used in Figure 2.6 to refer to the assemblies originally isolated for gasphase dissociation. In all cases shown, multiple stripped protein populations are observed and

36

rationalized based on the monomer components known to exist within each sample and the intact
mass of the isolated protein complex ion (Table 2.1). In the cases of Figures 2.9B, C, D and E,
for example, two overlapping protein complex ion populations are co-isolated in each dataset due
to their similar intact m/z values (Figure 2.6A), and they produce fragment ion populations that
allow for their discrete identification. In contrast, fragment ions recorded in Figure 2.9A
correspond to complex No. 2 and a previously undetected putative truncated form present at low
relative abundance. I also observed signals in the tandem MS data corresponding to multiple
dissociated monomers, dominated by signals from UreB and UreF at low m/z in Figures 2.9B, C,
D and E. For example, the ejection energies for UreF and UreB from complex No. 9 are similar,
resulting in two prominent stripped protein ion populations (indicated by open purple circles and
squares) in Figure 2.9E. In contrast, UreF and UreB are ejected at significantly different collision
energies for complex No. 3 versus No. 6 or complex No. 8 versus No. 10, thus aiding the
interpretation of the recorded CID patterns (Figure 2.9C and D).

37

Figure 2.9: Protein complex identification through precise mass measurements enabled by
tandem MS. Tandem MS CID spectra recorded for the urease-related species that appear in
Figure 2.6B, acquired at a trap collision voltage of 200 V (A, B, C, D and E). CID spectra are
typically dominated by signals for ejected UreB and UreF monomers at low m/z, indicated by
star and diamond notations respectively, while the signals corresponding to stripped protein
complex ions at high m/z are magnified (magnification factors indicated on each spectrum) and
marked by color-coded circles. Detailed annotations are given in Table 2.1.

38

Table 2.1: A list of measured masses determined from primary MS and tandem MS
(MS/MS) data, and sequence masses for the urease-related species that appear in Figure
2.6A. The sequence masses correspond to the urease apo-protein complexes, which can be
converted to the urease holo-enzyme species by adding 496 Da (6 nickel ions and 3 CO2
molecules) to the intact masses shown. Such small differences in masses cannot be resolved for
the complexes shown here using our instrumentation. Good agreement is found between
measured masses derived from MS/MS data and the sequence masses of the assigned complexes
in each panel of Figure 2.9. The same marks are indicated such that the dissociated monomers
and stripped complexes can correspond to their peaks shown in Figure 2.9. (UreABC)3, UreD,
MBP-UreD, UreD, UreF and UreG are abbreviated as U3, B, MD, D, F and G, respectively.

Surprisingly, Figure 2.9B contains signals corresponding to both UreB and UreF though
the complex I intended to isolate (No. 7) contains only the former as a constituent. Using IM-MS
data to guide my interpretation, however, I discovered that the 33 + charge state of complex No. 7
and the 34+ charge state of (UreABC)3:MBP-UreD:UreF lacking one UreB subunit coexist in the
dataset and possess m/z values that cannot be distinguished by the quadruple MS (Figure 2.5).
The latter complex contains UreF, and thus accounts for the appearance of those signals in
Figure 2.9B.
Overall, the specific datasets shown clearly assign the stoichiometries and compositions
39

of the complexes indicated in Figure 2.6E. By summing the fragment ion population masses
observed in Figure 2.9, I obtained intact complex masses that closely agree with the sequence
masses for the protein constituents indicated (see Table 2.1). Nevertheless, protein complex
topology is not established in all cases. For example, in addition to the two versions shown for
species No. 9 one could alternatively position both MBP-UreD:UreF units (or the MBP-UreD
and MPB-UreD:UreF:UreF units) on the same urease trimer; however, a urease:MBPUreD:UreF:UreF:MBP-UreD:urease arrangement is excluded by the CID results. Similarly, two
versions are shown for species No. 10, whereas the CID data preclude a single MBPUreD:UreF:UreF:MBP-UreD unit bound to the urease. In other cases where many possible
protein complex topologies occupy the same molecular mass, results from multiple samples
under different conditions inform my assignments. A hypothetical assembly pathway that is most
consistent with the MS data is depicted in Figure 2.8.
The soluble (MBP-UreDFG)2 complex derived from the K. aerogenes proteins is an
overall dimer, compatible with the previously reported (UreF) 2 and (UreH:UreF)2 structures of
the H. pylori proteins (15, 24). Furthermore, the chemical cross-linking results demonstrate a
close juxtaposition (BS3 forms a crosslink of 11.4 Å) of the amino termini of the two UreF
protomers within this complex. This result also is consistent with the published structural results;
the first 25 residues of H. pylori UreF are unstructured, but the shorter K. aerogenes protein
lacks these residues and its dimer is predicted to have the two amino termini on the same protein
face with the first four residues of each protomer likely to have substantial flexibility. The
(MBP-UreDFG)2 complex binds to the urease apoprotein and, unexpectedly, to the urease
holoenzyme. The dimeric structure of (MBP-UreDFG)2 raised the question of whether the intact
species binds to urease or if it dissociates to form a heterotrimeric species upon binding.

40

The intact protein complex MS results display a number of features related to urease
structure and stability that confirm both the chemical cross-linking and SDS-PAGE data, as well
as previously reported findings. First, the loss of UreB from urease (Figure 2.9A) and several of
its complexes (e.g., Figure 2.9B-E) may relate to its proposed weak interaction with UreC,
providing access to the nascent active site (35). Second, even though UreF can be cross-linked to
UreB in the (UreABC):UreD:UreF complex (12), previous data indicated UreF to be relatively
weakly bound to the urease core complex (11); these findings suggest that UreF uses UreD as a
scaffold to bind to urease rather than having a significant region of direct interface. This weak
binding is observed both in the stoichiometry of the complexes recorded in the intact MS dataset
that are most-likely formed through UreF dissociation, specifically the conversion between
complexes No. 6 to 7 and No. 4 to 8 (Figure 2.6), and the tandem MS data, which showed
several instances of the preferential ejection of UreF over other proteins upon collisional
activation in the gas phase. This is specifically seen in the cases of complexes No. 3, 6, 8, 9, and
10, despite the constant presence of the much smaller UreB, the ejection of which is favored in
(UreABC)3 CID. Furthermore, the intact protein complex MS data strongly indicate the
formation of a UreF dimer within (MBP-UreDFG)2, primarily through the observation of
complex No. 6, which is also consistent with the MALDI-MS cross-linking data (Figure 2.2). In
general, MS data obtained with intact protein complexes also agree well with the gel filtration
analysis, in that the stoichiometry of the MBP-UreDFG complex determined through both
techniques is identical.
In addition to these aspects of commonality, intact protein complex MS analysis reveals
unique aspects of urease-related structures and enzyme activation that were previously unknown.
I found substantial evidence for protein complexes that are likely off-pathway in the formation of

41

activated urease. Complexes such as No. 6, 7, 8, and 10 all represent assemblies that could
possibly result from the decay of the core urease complex bound to one or two MBP-UreDFG. In
addition, complexes No. 5 and 9 appear to result from the dimerization of either (UreABC)3 or
(UreABC)3:MBP-UreD:UreF, and are also likely to represent off-pathway assemblies. Of
potential interest, the hexameric architecture of urease subunits in these species is reminiscent of
jack bean urease, which is a hexameric enzyme (1). To account for all of the subcomplexes and
decay products observed in the intact protein complex MS data, I invoked sequential (MBPUreDFG)2 additions to the core urease complex to generate complexes No. 3 and 4 (Figure 2.8),
both of which are likely to lie on the activation pathway for the enzyme. I presume that
incubation of (MBP-UreDFG)2 in great excess over (UreABC)3 would lead to addition of a third
MBP-UreDFG unit to the enzyme; however, the experiment was designed to test for the presence
of No. 3 rather than larger complexes that would less clearly address the question. Each of the
pre-activation complexes appears to undergo dissociation under the analytical conditions to
produce other partially assembled complexes that are unlikely to contribute to urease activation.
Significantly, however, (MBP-UreDFG)2 does not dissociate to yield MBP-UreD:UreF:UreG,
free UreF, or UreF:UreF dimer (Figure 2.7), suggesting that these species are not available and
do not add to urease during the in vitro incubation studies. The observed assemblies are present
over relatively long time-scales (hours/days), suggesting that the complexes identified in Figure
2.6 are all relatively stable forms of these proteins in vitro.
CONCLUSIONS
I present here unique data on the urease activation complexes formed between the
(UreABC)3 core assembly and a complex comprised of UreD, UreF, and UreG. The MS data of
the intact urease assemblies involved in enzyme activation are supported by measurements

42

involving SDS-PAGE, gel filtration, chemical cross-linking, and IM separations. First, I detected
clear evidence that a (UreDFG)2 complex with juxtaposed UreF subunits is the primary
biological unit generated by UreD, UreF and UreG under the experimental conditions. Secondly,
I detected the stepwise addition of UreDFG units to the core (UreABC) 3 assembly upon exposure
to (UreDFG)2 in solution, or from complexes isolated directly from cells. These
(UreABC)3(UreDFG) and (UreABC)3(UreDFG)2 assemblies are on route to the putative, fullyassembled pre-activation complex shown in Figure 2.1 through the addition of UreDFG units.
This result, therefore, lends credence to a urease activation mechanism in which UreDFG binds
to urease apoprotein (Figure 2.1, bottom track), but does not preclude the sequential protein
binding pathway (Figure 2.1, top). Importantly, these findings are inconsistent with the
repeatedly seen UreF:UreF interaction remaining intact. Finally, I also observed evidence for
many off-pathway complexes that appear in high abundance in the intact MS measurements,
suggesting either the relative fragility of the pre-activation urease complexes probed, or the
likely complexity of the activation process. Future work in this area will likely capitalize on the
data shown here to further probe the structures and stabilities of urease activation complexes, and
eventually construct a complete map of their assembly in vivo.

43

BIBLIOGRAPHY

44

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48

CHAPTER 3

Characterization of Klebsiella aerogenes UreD Variants Reveals a Potential NickelTransfer Tunnel for Urease Activation

49

ABSTRACT
Nickel-containing urease from Klebsiella aerogenes requires the accessory proteins
UreD, UreF, and UreG, along with the metallochaperone UreE, for proper active site metalation.
Earlier efforts had shown that UreE binds to UreG in the presence of nickel, that UreD, UreF,
and UreG form a complex, and that the UreD:UreF:UreG complex binds to urease via UreD. In
silico analysis of the homologous, structurally characterized UreH:UreF:UreG complex from
Helicobacter pylori identified a putative water tunnel originating from the likely nickel-binding
motif in UreG, through UreF, and exiting UreH, but no experimental support for this tunnel was
reported. A UreD homology model was created and computationally analyzed to reveal a
comparable tunnel with an identical origin at the presumed UreD:UreF interface and several exit
points on the opposite face of the protein. In vivo activation of urease within cells that produce
specific UreD variants which were designed to disrupt or block the putative 34.6 Å internal
tunnel resulted in greatly reduced urease specific activities relative to enzyme activated with
wild-type UreD, whereas cells producing UreD variants targeted to the alternate tunnel exits or
to the protein surface exhibited no appreciable effect on activity. Affinity pull-down studies
using cell-free extracts of cultures producing variant forms of UreD along with the other urease
components showed no loss of interaction between UreD and either urease or the UreF:UreG
accessory proteins. Metal-content analysis of urease enriched from extracts of the activitydeficient cultures demonstrated reduced nickel content for all samples, as well as an increase in
zinc and iron incorporation. These findings support the role of UreD as a direct facilitator of
nickel transfer into urease by using an internal water tunnel.

50

INTRODUCTION
It has been estimated that one-third of all proteins contain metals, with ~40% of
metalloproteins having metal centers that are essential for catalysis (2, 22, 43). These proteins
must overcome several challenges during their synthesis in order to incorporate the proper
metal(s) to become functional. An approach to circumvent mismetalation is to use accessory
proteins for binding the metal and transferring it into the nascent metalloenzyme active sites (11,
14, 18). Examples of this strategy are seen in the Nif proteins involved in biosynthesis of the
FeMoco cofactor of nitrogenase, the Hyp proteins used to generate the [NiFe]-hydrogenase
cofactor, the copper chaperone for superoxide dismutase required for copper transfer into the
Cu,Zn-superoxide dismutase (11, 24, 44), and the focus here -- the activation of nickelcontaining urease.
Urease catalyzes the hydrolysis of urea into ammonia and carbamate, with the latter
molecule spontaneously decomposing into a second molecule of ammonia and bicarbonate (18).
The enzyme is found in plants, as well as some bacteria, Archaea, algae, and fungi (16, 33, 41,
47). Most ureases use a dinuclear nickel metallocenter for catalysis, though a recent example of a
di-iron urease has been characterized (10). Most of our knowledge about the formation of the
nickel active site comes from studies of the urease activation machinery of Klebsiella aerogenes
and Helicobacter pylori, as detailed below.
The urease gene clusters from K. aerogenes (ureDABCEFG) (18) and H. pylori
(ureABIEFGH) (21) are similar, but the encoded proteins exhibit a few important distinctions.
The three structural subunits of the K. aerogenes enzyme (KaUreA, KaUreB, and KaUreC) are
closely related to the two subunits of the H. pylori protein (HpUreA is homologous to a fusion of
KaUreA and KaUreB, while HpUreB is homologous to KaUreC). The subunits assemble into a
51

trimer of heterotrimers, (KaUreABC)3, and a tetramer of trimeric heterodimers, ((HpUreAB)3)4,
respectively, as shown by their crystal structures (22, 25). In each case, the dinuclear active site
is located in the largest subunit and, in the case of the K. aerogenes enzyme, access to this site is
proposed to be gated by the flexible domain of UreB (8, 25, 40). KaUreD and HpUreH are
homologous and neither is soluble when overproduced as an independent protein (19, 38);
however, a maltose-binding protein (MBP) fusion to KaUreD (MBP-UreD) is soluble, binds two
equivalents of nickel, and partially complements a ΔureD loss of function phenotype in vivo (9).
KaUreF is insoluble when independently overproduced (4), but it is soluble and functional as the
UreE-UreF fusion protein (28). By contrast, HpUreF is soluble, a slightly truncated form was
structurally characterized (29), and isothermal titration calorimetry studies demonstrate its
nickel-binding ability (53), although no studies have been performed to determine if this metal
binding is functionally relevant. KaUreG exists as a soluble monomer that binds one equivalent
of nickel or zinc with similar affinities (5, 34), while HpUreG is a monomer in the absence of
metal, binds zinc with high affinity leading to dimerization, and binds nickel with low affinity
without facilitating dimerization of the protein (3). These proteins act as GTPases during urease
activation, although the precise role of GTP hydrolysis in this process remains unclear. UreG
Cys and His residues associated with a highly conserved Cys-X-His motif are hypothesized to
function as the metal binding site, but substitution of one or both of these residues does not
abolish metal binding (4, 54). Crystal structures are known for both KaUreE (48) and HpUreE
(46); these are dimeric nickel-binding proteins proposed to deliver nickel for urease activation
(15, 37). Metal-ion induced interactions between UreE and UreG have been observed for the
proteins from K. aerogenes, H. pylori, and Sporosarcina (formerly Bacillus) pasteurii (3, 5, 32).
Attempts to activate K. aerogenes urease in vivo in the absence of any one of its accessory

52

proteins result in the production of inactive enzyme lacking nickel (30, 36). Finally, a protongated urea permease (UreI) is produced by H. pylori (50), but not K. aerogenes.
Interactions among the accessory proteins and the enzyme subunits are critical for urease
activation. A UreD:UreF:UreG complex is formed in vivo when the three corresponding K.
aerogenes accessory genes are expressed separately from those encoding the urease structural
subunits, though direct studies of this complex were limited due to its low solubility (34). In
contrast, a soluble complex is formed with MBP-UreD, producing a (MBP-UreD:UreF:UreG)2
species that dissociates to a monomer of heterotrimers when bound to urease according to sizeexclusion chromatography and mass spectrometry studies (17). The crystal structure of the
analogous (UreH:UreF:UreG)2 H. pylori species is known (20). The surface of UreD/UreH that
interacts with urease has not been defined.
While the importance of the accessory proteins in the activation of urease is clear, the
exact mechanism of this activation process is still unknown, with two main hypotheses posited.
One proposal invokes a “hand-off” mechanism in which cytosolic nickel binds to the UreE
metallochaperone and passes to surface-exposed residues of UreG, then UreD, and finally into
the nascent active site of K. aerogenes urease apoprotein, all within the urease:UreD:UreF:UreG
complex (5, 9). Precedence for such a hand-off mechanism includes Cox17 and ScoI, which
utilize thiol groups to pass Cu(I) into cytochrome c oxidase (42). The second hypothesis involves
the initial delivery of nickel from UreE to UreG, followed by the use of a buried channel
connecting the proposed nickel-binding site of UreG, through UreF and UreH, directly into the
nascent active site of H. pylori urease (53). Examples of this strategy can be observed in the
metal ion transfer tunnel into the ferritin core (7), Fe-Fe hydrogenase synthesis via a channel
involving HydA (35) and membrane-bound uptake proteins for cations (6).
53

Here, the function of KaUreD in urease activation is examined. The effects of targeted
UreD side chain substitutions on the in vivo activation of urease, protein interactions with other
urease components, and metal content of the purified enzyme are determined. This work
provides further insights into how UreD functions in the activation of urease.
MATERIALS AND METHODS
UreD Homology Model Generation, Conservation Mapping, and Water Tunnel
Prediction. To help select residues for substitution in KaUreD, a homology model was prepared
by using the Protein Homology/analogY Recognition Engine (Phyre2.0) server (27) and the
crystal structure of HpUreH from the UreH:UreF:UreG complex (PDB code 4HI0) as a template.
A PSI-BLAST analysis (26) was performed for HpUreD, sequences with more than 15% and
less than 90% identity were identified, and the top 30 were selected along with KaUreD. A
multiple sequence alignment was prepared and the residue conservation scores were mapped
onto the KaUreD homology model using the ConSurf server (12). MOLE2.0 (45) was used to
predict water tunnels within the KaUreD homology model, and the effects of substituting
residues along this tunnel were analyzed for steric clashes in Pymol (1) and by MOLE2.0 to
determine if the water tunnels had been disrupted.
Plasmid Construction. To characterize the effects of point substitutions on the function
of KaUreD in vivo, three types of plasmids were constructed (i.e., pMF001L*, pKK17D*, and
pKKD*G, where * indicates the mutant versions).
An EcoRI/HindIII fragment of pEC002 (9) containing ureD was inserted into similarly
digested pUC8 to yield plasmid pMF001. This plasmid cannot be used to overproduce UreD
directly because it was found to lack a critical upstream region needed for overexpression, so an

54

EcoRI/AgeI fragment of pKK17 (15) (containing ureD and a 197-bp 5' untranslated region) was
substituted into similarly digested pMF001 to yield pMF001L. The accessory gene within
pMF001L was mutated by polymerase chain reaction with overlapping oligonucleotides
(Integrated DNA Technologies, Coralville, IA) containing the proper base-pair substitution(s)
and amplified with PfuTurbo® polymerase (Aligent Technologies). The resulting pMF001L*
products were digested with DpnI and transformed into E. coli MAX Efficiency® DH5α cells
(Life Technologies). Mutagenesis was confirmed by sequencing (Davis Sequencing, Davis, CA;
Michigan State University Genomics Core, East Lansing, MI).
To study the effects of ureD mutations within the context of the intact urease gene
cluster, the pMF001L* versions were digested and the desired ureD-containing fragments were
isolated as described above. These fragments were ligated into similarly treated pKK17 to yield
the analogous versions of plasmid pKK17D*. These plasmids were also sequenced to ensure the
proper insertions were present.
To examine the effects of substitutions in UreD on protein:protein interactions, wild-type
and variant EcoRI/AgeI ureD fragments from pMF001L* were ligated into the similarly digested
and isolated backbone of pKKG (5). The resulting pKKD*G plasmids contain the ureD versions
within the context of ureDABCEFGStr, where UreG has been modified with a C-terminal Streptag II (UreGStr). The validities of the resulting plasmids were confirmed by sequencing. A
summary of all plasmids used in these studies can be viewed in Table 3.1.

55

Table 3.1: Plasmids used in Chapter 3.
Plasmid used
pUC8
pKK17
pKK17D*, -V37L, -Y42D,
-E46A, -E46Q, -C48A, H49A, -H54A, -I59Y, D63A, -D63Q, -L65I, L65W, -S85K, -K86A, Y88V, -Y88F, -R89A, W111Y, -T128E, -D142A,
-R148M, -E153A, -E153Q,
-R163A, -E165A, -D169A,
-E176A, -E176Q, -T196K,
-R211A, -R233A
pKKG
pKKD*G, -D63A, -D63Q.
–S85K, -D142A, -E176A, E176Q, -R211A
pEC002
pMF001
pMF001L

Description
High-copy number, pBR322-derived vector
conferring AmpR
Wild-type K. aerogenes urease cluster
(ureDABCEFG) placed into pKK223-3

EcoRI-AgeI 5’UTR-ureD fragment from
pMF001L* ligated into similarly digested
pKK17

PstI-KpnI ureGStr fragment ligated into
similarly digested pKK17 resulting in
replacement of UreG with one containing a Cterminal Strep-tag II (ureDABCEFGStr)
EcoRI-AgeI 5’UTR-ureD fragment from
pMF001L* ligated into similarly digested
pKKG
pMAL-c2X derived vector for the
overproduction of maltose binding protein
fused at the N-terminus of UreD
EcoRI-HindIII ureD fragment from pEC002
ligated into similarly digested pUC8
EcoRI-AgeI 5’UTR-ureD fragment from
pKK17 ligated into similarly digested pMF001

Source
(51)
(15)

This study

(5)

This study

(9)
This study
This study

In Vivo Activation of Urease by Variant UreDs. E. coli BL21(DE3) competent cells
were transformed with pKK17D* or pKKD*G and plated on lysogeny broth (LB) agar plates
supplemented with 300 µg/mL ampicillin. A single transformant colony was used to inoculate 2
mL of LB medium supplemented with 100 µg/mL ampicillin and cultured overnight. Aliquots
(150 µl) of these overnight cultures were used to inoculate 15 mL of LB supplemented with 1
mM NiCl2 and 100 µg/mL ampicillin in 50 mL flasks, which were shaken at 200 RPM at 37 °C
until reaching an optical density at 600 nm (OD600) of 0.5, induced with 0.1 mM isopropyl β-D-

56

1-thiogalactopyranoside (IPTG), and grown overnight at 37 °C. Urease apoprotein was prepared
in a similar manner, but with the culture medium lacking supplemental nickel. Cells were
harvested by centrifugation and resuspended in 2.5 mL of 100 mM HEPES, pH 7.8, per g of wet
cell paste. Cells were lysed by sonication while placed in an ethanol ice bath using a Branson
450 sonifier with three 2-min cycles at 4 W output power and 50% duty cycle and 1 min of
temperature recovery between cycles. Cell lysates were clarified by centrifugation at 100,000 g
at 4 °C for one h. The soluble cell-free extracts were diluted 100-fold into 100 mM HEPES, pH
7.8, buffer for urease activity assays. Overproduction of urease subunits was confirmed by
sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE).
Affinity Pull-Down Assays Using Soluble Cell-Free Extracts. To test if activationdeficient UreD variants exhibit impaired interactions with urease or the other accessory proteins,
E. coli BL21(DE3) cells were transformed with pKKD*G constructs of interest. Cells were
cultured, induced, and harvested as described above, and frozen at -80 °C until being further
analyzed. Cells were thawed, resuspended in 1 mL of buffer W (50 mM NaH2PO4, pH 8.0,
containing 300 mM NaCl and 1 mM ethylenediaminetetraacetic acid (EDTA)) supplemented
with 0.1 mM phenylmethanesulfonylfluoride (PMSF), and lysed by sonication using 10 pulses at
1 W output power while placed on ice for five cycles with 15 s of recovery between cycles.
Unbroken cells and debris were removed by centrifugation at 22,000 g at 4 °C for one h. Soluble
cell-free extracts were loaded onto 2.5 mL of Strep-Tactin Superflow Plus resin (Qiagen) preequilibrated in buffer W and washed with 8 volumes of buffer W to remove unbound proteins.
Proteins were eluted with 8 volumes of buffer W containing 2.5 mM desthiobiotin. The soluble
cell-free extracts, wash fractions, and elution fractions were analyzed by SDS-PAGE.

57

Protein Purification. To determine the metal contents of urease produced by cultures
containing functionally-deficient UreD, E. coli BL21(DE3) cells were transformed with
pKKD*G constructs of interest. A single transformant colony was inoculated into 5 mL of LB
supplemented with 100 µg/mL ampicillin and cultured overnight. For each UreD variant used, 1
L of LB supplemented with 1 mM NiCl2 and 100 µg/mL of ampicillin was inoculated with 1 ml
of overnight culture and grown until reaching an OD600 of ~0.6. These cultures were induced
with 0.1 mM IPTG and grown overnight with shaking at 37 °C. Cells were harvested by
centrifugation and resuspended in 2.5 volumes of buffer containing 50 mM Tris-base, pH 7.4, 1
mM EDTA, and 1 mM β-mercaptoethanol (TEB), and supplemented with 0.1 mM PMSF. Cells
were lysed by sonication while immersed in an ethanol ice bath using a Branson 450 sonifier
with three 2 min cycles at 4 W output power and 50% duty cycle including 1 min of temperature
recovery between cycles. Soluble cell-free extracts were separated from unbroken cells and
debris by centrifugation at 100,000 g for one h at 4 °C. Supernatants were diluted 1:1 in TEB
buffer before being applied to a 90 mL Macro-Prep® DEAE Support (Bio-Rad) column preequilibrated in TEB buffer. The samples were washed with one volume of TEB buffer and eluted
by using a 0 M to 1 M NaCl gradient in TEB buffer. Fractions containing urease were pooled,
dialyzed into TEB buffer containing 25 mM NaCl, and concentrated to 1 mL by using an
Amicon® Ultra-15 10K centrifugal filter (Millipore). Each concentrate was injected directly onto
100 mL columns containing either Superdex 200 (General Electric) or Sephacryl 300 HR
(Sigma) resin pre-equilibrated in TEB buffer containing 25 mM NaCl. Proteins were eluted using
the same buffer, and the fractions containing urease were pooled and concentrated to 2.5 mL.
The pooled samples were analyzed for their purity by SDS-PAGE and assayed for urease
specific activity.

58

Urease apoprotein was purified from cells containing the pKK17 plasmid as described
previously (9).
Urease Activity Assays. Enzyme activity was measured by quantifying ammonia release
from urea using methods described by Weatherburn (50). The release of ammonia over time was
monitored by the formation of indophenol on the basis of its absorption of light at 625 nm. One
unit of urease activity is defined as the amount of enzyme required to hydrolyze 1 µmol of
urea/min at 37 °C. The standard assay buffer was 50 mM of HEPES, pH 7.8, with 50 mM urea.
Protein Analyses. All SDS-PAGE analyses were performed using 15% running gels and
5% stacking gels prepared with 1.5 M Tris, pH 8.8, and 1 M Tris, pH 6.8, buffers, respectively.
All gels were prepared using a 30% acrylamide:bis-acrylamide solution at 37.5:1 (Bio-Rad). All
protein concentrations were determined with a commercially available protein assay kit and
using bovine serum albumin as a standard (Bio-Rad). The metal contents of the purified urease
pools were analyzed by inductively couple plasma-atomic emission spectroscopy (ICP-AES)
(University of Georgia Chemical Analysis Laboratory).
RESULTS
Selecting Residues for Substitution in UreD. To investigate the function of UreD in the
activation of K. aerogenes urease, a series of ureD mutants were examined for their effects on
urease activity, metal content, and UreD:urease and UreD:UreF protein:protein interactions.
Residues targeted for substitution were based, in part, on a multiple sequence alignment (31) of
32 homologous sequences that identified the conserved residues in the protein (Table 3.2). No
metal-binding motifs were readily apparent within the alignment, nor were any metal-binding
motifs identified for UreH within the UreH:UreF:UreG structure (19, 20).

59

Table 3.2: Multiple sequence alignment of UreD homologs used in Consurf conservation
analysis. a
Nostoc sp. PCC 7120 UTEX 2576
Nostoc sp. PCC 7107
Nostoc sp.
Anabaena sp. 90
Anabaena cylindrica
Nostoc punctiforme sp.
Microchaete sp. PCC 7126
Chlorogloeopsis fritschii
Scytonema hofmanni UTEX 2349
Fischerella sp. PCC 9339
Fischerella sp. PCC 9431
Fischerella sp. JSC-11
Fischerella sp. PCC 9605
Scytonema hofmanni
Mastigocladopsis repens
Mastigocoleus testarum
Calothrix sp. 336/3
Rivularia sp.
Calothrix sp.
Leptolyngbya boryana
Microcoleus sp.
Coleofasciculus chthonoplastes
Synechocystis sp.
Chroococcidiopsis thermalis
Crinalium epipsammum
Gloeocapsa sp. PCC 7428
Oscillatoria nigro-viridis
Oscillatoria acuminata PCC 6304
Arthrospira platenis sp.NIES-39
Pseudomonas syringae
Klebsiella aerogenes
Helicobacter pylori

-------GWHGKLNLVYAD-RSNSTQLIYNHQQAPLKVQRPFYPE-----------AWHGKLNLIYAD-RQNSTQLIYNHHQAPLKVQRPFYPE-----------GWHGKLNLVYAD-RLGATELIYNHQQAPLKIQRPFYPE----------QGWHGLLNLVYAN-RQDSTQLIYNHHQAPLKVQRPFYPE----VNSPIDKNWHGRLNLVYAK-RQDSTQLIYNHHQAPFNIQRPFYPE-----------GWHGKLNLVYAD-RQGATQLIYNQQQAPLKVQRPFYPE---------TEGWHGQLNLVYAD-RHNSTQLIYNHHQAPLKVQRPFYPE----------QGWYGNLNLVYAR-TQGETQLIHSQSQAPLKVQRPFYPE-----------GWHGKLNLVYAD-RLGTTALIHNSHQAPLKVQRPFYPE------------WHGNLHLVYAD-RQNQTQLIFNQNQAPLKVQRPFYPE------------WYGNLDLVYAY-HQNQTQLIFNQNQAPLKVQRPFYPE------------WHGNLHLVYAY-RQNQTQLIFNQNQAPLKVQRPFYPE-----------GWHGKLNLVYAH-SQSGTQLIFNRNQAPLKVQRPFYPE-----------GWHGKLDLVYAQ-CQGKTTLIHNQNIAPLKVQRPFYPE---------PTGWYGKLSLVYAH-RQNTTTLIHNQNQAPLKVQRPFYPE------------WHGMLNLTYAS-RQGKTVIVEQQNQAPLKVQRPLYPEEQTDP
-------SWHGKLHLTYGS-HGGKTQLVSSQNQAPLKVQRPFYPE--------SSQSWHGKLNLVYNH-SQNKTALIQSRNQAPLKVQRPFYPE-----------SWHGKLDLLYAN-RQGITQLIHAHHQAPLKVQRPFYPE-------LNPTDWHGILQLGFEY-RQGKTQLIRNQGQAPLKVQRPFYPE--------SSSGWHGSLQLVYAH-DHNGTQLTHAQVQAPLKVQRSFYPE------------WQGSLELVYAN-DQGKTRLVRDRITSPLKVQRPFYPE------------WHGNLNLVYAQ-HQGKTQVIHSQMKAPLKVQRPFYPE---------PSTWHGNLDIVYAL-RNGKTQPISDRVQAPLKVQRPFYPE---------KSSWHGSLDLKFAC-RDGGSQVIKSQGKAPLKVQRPFYPE-----------SWHGSLNMVYTC-VDGATTVTHQQMQAPLKVQRPFYPE---------QTAWHGRLNLAYAN-RSGATQIIHNQMQAPLKVQRPFYPE----------QGWHGSLELVYAQ-SGNATQMVSAKATAPLKIQRPFYPE--------NPSEWHGILNLTYSQ-NHGITQVVDKYTTAPYKIQRPFYPP---------TPNWHAELELGYGR-FGDSTRPVQRRHKGPLRVQKHLYAE-----------GWQATLDLRFHQ-AGGKTVLASAQHVGPLTVQRPFYPE-------MNTYAQESKLRLKTKIGADGRCVIEDNFFTPPFKLMAPFYPK----. * :
.
* :
:*.

Nostoc sp. PCC 7120 UTEX 2576
Nostoc sp. PCC 7107
Nostoc sp.
Anabaena sp. 90
Anabaena cylindrica
Nostoc punctiforme sp.
Microchaete sp. PCC 7126
Chlorogloeopsis fritschii
Scytonema hofmanni UTEX 2349
Fischerella sp. PCC 9339
Fischerella sp. PCC 9431
Fischerella sp. JSC-11
Fischerella sp. PCC 9605
Scytonema hofmanni
Mastigocladopsis repens
Mastigocoleus testarum
Calothrix sp. 336/3
Rivularia sp.
Calothrix sp.

GEKVCHSVILHTAGGVVGGDRLSYNLHLQPNAQALITTAAAGKVYRSDG
-GEKVCHSVILHTAGGVVGSDRLSSHFHLQPNTQALITTAAASKIYRSNG
-GERVCHSVILHTAGGVVGGDRLSTKIHLQPHTQAVITTAAAGKIYRSNG
-GQEICHSVILHTAGGIVGGDRLSSHIHLQPDTNALITTAAAGKIYRSNG
-GQEVCHSVILHTAGGIVGGDRLSSDIHLEKDSQALITTAAAGKVYRSNG
-AEKVCHSVILHTAGGMVGGDRLSSNIHLQPQAQALITTAAASKIYRSNG
-GEKVCHSIILHTAGGIVGGDRLSSKIHLQPDAQAVITTAAASKIYRSNG
-GKNVCHSVILHTAGGIVGGDRLSCNFHLEPHAQALITTAAASKIYRTNG
-GQQVCHSIILHTAGGVVGGDRLSLNFHLQPNTQALITTAAASKIYRSNG
-GQKVCHSVILHTAGGVVGGDRLSYDFHLQPHAQALITTATAGKIYRTNG
-GQEVCHSVILHTGGGVVGGDRLSYDFHLQSDAQALITTATAGKIYRSNG
-GQEVCHSVILHTAGGVVGGDRLSYNFHLQPHAQALITTATAGKIYRSNG
-GQEVCHSVILHTAGGVVGGDRLSYHLHLQPNAQALITTAAAGKIYRSNG
-GQEICHSVILHTAGGVVGGDRLSYKVHLQPKAQALITTAAASKIYRSNG
-GQQVCHSVILHTAGGIVGGDRLSCNFHLQPNAQALITTAAASKIYRSNG
EGQKICHSIILHTAGGVVGGDRLSCNFDLQPNSQALITTAAASKIYRSNG
-GEAVCHSIILHTAGGVVGGDRLSTNIHLQPQSQVLLTTAAASKIYKSNG
-GQSVCHSVILHTAGGIVGGDRLSSDFHLQPDSKALITTAAANKIYRSNG
-GKAVCHSVILHTAGGVVGGDKLSCNFQLQPESQVLITTAAAGKIYRSNG

60

Table 3.2 (cont’d):
Leptolyngbya boryana
Microcoleus sp.
Coleofasciculus chthonoplastes
Synechocystis sp.
Chroococcidiopsis thermalis
Crinalium epipsammum
Gloeocapsa sp. PCC 7428
Oscillatoria nigro-viridis
Oscillatoria acuminata PCC 6304
Arthrospira platenis sp.NIES-39
Pseudomonas syringae
Klebsiella aerogenes
Helicobacter pylori

-GDEVCHSVIMHTAGGIVGGDRLTFDFHLASGSQALITTPAASKIYRTNG
-GSAVCHSVVLHTAGGIVGGDRLSQTIHLSPHAHALITTAAASKIYGSKG
-GQGVCHTVVLHTAGGIVGGDRLSQTIHLQEDSQALITTAAASKIYRSNG
-G-GVCHSVVLHTAGGIVGGDRNNLSFHLQPHSQSLITTATASKIYRSNG
-G-DICHTAILHTAGGIVGGDCLSINLQLQPRSQALVTTTAASKIYRSNG
-GREVCHSVILHTGGGVVGGDRLSLNFHLQPNTHALITSAAAGKVYRSNG
-GAEVCHSVILHTAGGVVGGDKLSLNFHLQQNAHTLITTAAASKIYRSNG
-GKDVCHSVILHTAGGVVGGDRLSGHFHLQPNAKALITTAAAGKIYRSSG
-GEGVCHSVILHTAGGIVGGDSLGQSIHVQENAHALITTAAAAKIYRSTG
-GEEICHSVALHTAGGMVGGDRLSQNLHLQADTKVLLTTAAASRVYRSTG
-GPQVCQHIIVHPPGGIAGGDRLDISAHVGPDAWAQLTSPGAAKWYRAAG
--EETCHLYLLHPPGGIVGGDELTISAHLAPGCHTLITMPGASKFYRSSG
--DDLAEIMLLAVSPGMMRGDAQDVQLNIGPNCKLRITSQSFEKIHNTED
..
:
*: .*
.:
:*
: : : .

Nostoc sp. PCC 7120 UTEX 2576
Nostoc sp. PCC 7107
Nostoc sp.
Anabaena sp. 90
Anabaena cylindrica
Nostoc punctiforme sp.
Microchaete sp. PCC 7126
Chlorogloeopsis fritschii
Scytonema hofmanni UTEX 2349
Fischerella sp. PCC 9339
Fischerella sp. PCC 9431
Fischerella sp. JSC-11
Fischerella sp. PCC 9605
Scytonema hofmanni
Mastigocladopsis repens
Mastigocoleus testarum
Calothrix sp. 336/3
Rivularia sp.
Calothrix sp.
Leptolyngbya boryana
Microcoleus sp.
Coleofasciculus chthonoplastes
Synechocystis sp.
Chroococcidiopsis thermalis
Crinalium epipsammum
Gloeocapsa sp. PCC 7428
Oscillatoria nigro-viridis
Oscillatoria acuminata PCC 6304
Arthrospira platenis sp.NIES-39
Pseudomonas syringae
Klebsiella aerogenes
Helicobacter pylori

--------LQARQTIEIKIDAGACLEWLPQETILFNGAIYRQDLRVELAT
--------LQARQTIDIQIDAGACLEFLPQETILFNGADYRQDLRVELAT
--------LQARQTIDIQIDAGACLEWLPQETILFNGAIYRQDLRVELAT
--------LPARQTVNIQVDSHACLEYLPQETILFNGGIYRQDLRVELAT
--------LPAKQTVNIQIGANACLEYLPQETILFNGAVYRQDLKVKLDT
--------LQARQTIQMQVDPGACLEWLPQETILFNDAIYRQDLRVELAT
--------LPATQTINLKIDTGACLEWLPQETILFNDGIYRQDLRVELAT
--------LRARQTINIQIDANACLEWLPQETIVFNSALYRQDLRVELAT
--------TQAKQSINIQVDAGACLEWFPQETIVFNGAIYRQDLRVELAT
--------MTAKQMIEIKVDDGACLEWLPQETIVFDGALYRQDINVKLAT
--------MIAKQVIEIKVDDGACLEWLPQETIVFDGALYQQDLRVNLAT
--------TIAKQTIEIKVDDHAYLEWLPQETIVFDGALYRQENIVKLAT
--------TTARQTIEIKVDAGACLEWIPQETIVFNSALYRQDLRVELTT
--------AQARQNIDIQVDAGASLEWLPQETIVFNGAIYRQDLRVELAT
--------TQAQQNIEIRVDSGACLELLPQETIVFNGAIYRQDLRVELAT
--------SQARQKIEINVDERACLEWLPQETIVFNAADYRQDLRVDLAN
--------LQARQDVTIHIDSHACLEYLPQETIVFNSANYRQDVRIHLAV
--------LQARQNIDIKIDKNACLEWLPQETIVFSGASFRQDIKIELAD
--------RQATQNINIEVSTGATLEWLPQETIVFDGAIYRQDTRINLAE
--------REAHQVIRVDVAEGACLEWLPLDSIVFNQAIYRQTMQINLAQ
KSRACPQGQPAKQTIQIQIDQAACLEWLPQETIVFNGAIYQQDLRVELAP
--------QRANQRIHIHVEAGACLEWLPQETIVFNGADYQQQMTVELAP
--------LLAKQNIQMQVDTDACLEWLPQETIVFDGAIYRQDLQVELAP
--------LQARQIVEIQIDEGACLEWLPQETIVFNGANYRQDLRVELAP
--------LEARQTVEMRVEAGACLEWLPQENIIFNDANYRQDLRVELAT
--------WEARQNIQVQVDSNACLEWFPQETIVFNGAIYRQDLRVELAP
--------LESQQNIDIQLDTGANLEWLPQETIVFDGAIYRQNLRVELAP
--------EVARQSIKINVDSGAICEWLPQESIIFNGAIYRQDLRIELAP
--------KTASQNVKIKLEKGAYLEYLPRETIIFNGAIYRQDLRVELAP
---------PAYQQLDLTVEAGATLEWLPQETIVFSAAQAELTTRIELQG
--------AQALVRQQLTLAPQATLEWLPQDAIFFPGANARLFTTFHLCA
G--------FASRDMHIVVGENAFLDFAPFPLIPFENAHFKGNTTISLRS
:
: :
* :: *
* * . .
. *

Nostoc sp. PCC 7120 UTEX 2576
Nostoc sp. PCC 7107
Nostoc sp.
Anabaena sp. 90
Anabaena cylindrica
Nostoc punctiforme sp.
Microchaete sp. PCC 7126
Chlorogloeopsis fritschii
Scytonema hofmanni UTEX 2349
Fischerella sp. PCC 9339
Fischerella sp. PCC 9431
Fischerella sp. JSC-11

GANFLGWEITRFGRSARGEKFYQGEWRSHTEIWQQGVPLWIDRQWLPGND
GACFLGWEITRCGRSARGEKFLEGKWRSHTEIWQQGVPLWIDRQFLPGNT
GANFIGWEITRFGRSARGEKFYQGEWRSHTEIWQEGVPLWIDRQYLPGSE
DASYLAWEITRFGRSARGEKFVQGEMRSHTEIWQNGIPLWIDRQIVPGSE
NSSFIGWEITRLGRSARGEKFLEGEMRSHTEIWQNGIPQWIDRQILPGSE
GASWLGWEITRFGRSARGEKFLQGEWRSHTEIWQQSVPLWIDRQCLRGSE
GASFLGWEITRFGRTARGEKFYSGEWRSHTEIWQHNIPLWIDRQWLPGNQ
GASYLGWEITRFGRTARGEKFVQGEWRSHTEIWQQGKPLWIDRQWLPGSD
KASYLGWEITRFGRSARGEKFVQGEWRNHTEIWQQGKPLWIDRQWLPGSE
TASYIGWEITRFGRSARGEKFLQGEWRSHTEIWQQDKPLWIDRQWLPGSE
KANYIGWEITRFGRSARGEKFLQGEWRSHTEIWQQNKPLWIDRQWLPASE
NSHYLGWEITRFGRSARGEKFLQGEWRSHTEIWQQGKPLWIDRQWLPGRE

61

Table 3.2 (cont’d):
Fischerella sp. PCC 9605
Scytonema hofmanni
Mastigocladopsis repens
Mastigocoleus testarum
Calothrix sp. 336/3
Rivularia sp.
Calothrix sp.
Leptolyngbya boryana
Microcoleus sp.
Coleofasciculus chthonoplastes
Synechocystis sp.
Chroococcidiopsis thermalis
Crinalium epipsammum
Gloeocapsa sp. PCC 7428
Oscillatoria nigro-viridis
Oscillatoria acuminata PCC 6304
Arthrospira platenis sp.NIES-39
Pseudomonas syringae
Klebsiella aerogenes
Helicobacter pylori

GASYLSWEITRFGRSARGEKFLQGEWRSNTEIWQQGKPLWIDRQWVPGSE
GANWIGWEITRFGRSARGEKFCLGEWRSHTEVWQMGVPLWIDRQWLPGSE
KASWIGWEITRFGRSARGEKFLQGEWRSHTEIWQQGVPLWIDRQLLPGSE
NASFLGWEINRFGRTARGEKFVQGNWRSHTEIYQEGIPLWIDRQWLPGSE
DASFLGWEITRFGRTARNEQFIQGEWRNYTEIWRDNQPLWIDRQYLPGDM
DANFIGWEITRFGRTARQEKFLTGEWRSNTEIWQNNKPLWIDRQYLPGSE
NANFIGWEITRFGRSARGEKFLHGNWKSNTEVWQQGKPLWVDRQWLPGSE
GANWLGWEITRFGRSARGEKFVEGNWRSRTEVWQAGKPIWIDRQWMPGSE
GASWLGWEITRFGRSARGERFLQGEWRSHTEVWQQGRPLWIDRQWLPGEE
GASWLAWEITRFGRTARKERFLQGDWRSHTEIWQQGHPLWIDRQWLPGGE
GAKWLGWEITRFGRTAKGERFLHGDWKSHTEVWQQGLPLWIDRQWLPAGE
GASWLGWEITRFGRSARGERFLQGEWRSYTEVWQQGQPLWIDRQWLPGEE
DATWMGWEITRLGRTARGEQFLQGNWRSHTEIWRQGDPLWIDRQWLPGGE
GATWTGWEITRFGRSARGEKFLQGNWRSHTEVWQQQRPLWIDRQQLRPDV
TARILLWEITRFGRSARGENFLSGEWRSHTEVWQENSPLWIDRQLLKGGE
DARFLLWEINRFGRSARGETFVQGEWRSQTEIWQQGRPLWIDRQHLQGSE
EATWLGWEITRFGRSARGENFGQGEWRSHTEIWQNEKPLWIDRQWLPGGE
DAKLMYWDVVALGRPASGERFTSGHFQSHVDIRRDGRLLWHERQRIIGGD
SSRLLAWDLLCLGRPVIGETFSHGTLSNRLEVWVDNEPLLVERLHLQEGE
SSQLLYSEIIVAGRVARNELFKFNRLHTKISILQDEKPIYYDNTILDPKT
:
::
** . * * .
. .:
:. :

Nostoc sp. PCC 7120 UTEX 2576
Nostoc sp. PCC 7107
Nostoc sp.
Anabaena sp. 90
Anabaena cylindrica
Nostoc punctiforme sp.
Microchaete sp. PCC 7126
Chlorogloeopsis fritschii
Scytonema hofmanni UTEX 2349
Fischerella sp. PCC 9339
Fischerella sp. PCC 9431
Fischerella sp. JSC-11
Fischerella sp. PCC 9605
Scytonema hofmanni
Mastigocladopsis repens
Mastigocoleus testarum
Calothrix sp. 336/3
Rivularia sp.
Calothrix sp.
Leptolyngbya boryana
Microcoleus sp.
Coleofasciculus chthonoplastes
Synechocystis sp.
Chroococcidiopsis thermalis
Crinalium epipsammum
Gloeocapsa sp. PCC 7428
Oscillatoria nigro-viridis
Oscillatoria acuminata PCC 6304
Arthrospira platenis sp.NIES-39
Pseudomonas syringae
Klebsiella aerogenes
Helicobacter pylori

AVFHSPHGLAGQPIVGSLVWLGSPISTEIIEKARNLGNTQGE-------DIFHSPHGLFGQPIVGSLLWLGHPVSTEIIEQVRSL-FTKHL-------EVFHSPHGLSGQPIAGNFIYLGSPVSKETIEKARSIFTPHAL-------EVFHSPHGLRDNPVVGSFVGVGFPISPEIINQARSLIIQNSD-------EVFHSPHGLAGYPVVGSLVWVGSSVSREIIEKARSLVTQNNL-------DIFHSPHGLAGKPIVGSLVWVGGAVSAEIVEKTRSLWNGEGE-------DVFHSPHGLSGKPIVGTFVWVGDVVSAEIVETARNLWNGEGE-------KIFHSPHGLNGQAIAGSLVWVQGAVSQDMIEKARDLWHGEGE-------EVFHSPHGLAGQPITGTLVWVGSSVSAEIVEKARLKWNGAGE-------EVFHSPHGLAGQPIVGSLVYIGQKISPELVHQARNLWKPTLT-------EVFHSPHGLAGQPIVGSLVYIGQEISPELVNKARNLGKPTLS-------EVFHSFHGLAGQPIVGSLVYIGREISPEIVEKARNLWQLPIT-------EIFYSPHGLAGQPVVGSLVWVGSAVSVEIIEKARNIWQPIPPSPSLPL-DVFHSPHGLAGQPVTGSLVYIGQEVSQELVHKARTLFLSPTPHSLLP--EVFHSPHGLAGQPLVGSLVYVGQEVSPELVEKVRSLWNG----------EIFYSPNGLAGQAVIGSLIFVGKTISKDIVEQVRSLWKSNPHMTLSRHAA
AVFHSPHGLNSQAIAGSFIYLGKQISPQLITQIRQMWGEN---------EVFHSPHALAGKPIVGTLIYIGKPVSPEIVQKIRTLFIPPS--------EIFYSPHALNGQPVVGTFIYIGSTVSPEIIEKASSYTQHSVLS------ENFASPHGLAGCPVVGSFAWVGQVVTPELVEKARELWAGSSG-------QVLDSPHGLAGKPIVASLAWVGQAVSPEMIEKARLIWA---TQER----AVLDSPHGLAGEPIVGTLIWMGPPVSSEIIDNARSLWT---ADQR----KIIDSPHGLAGLPIVGSLAWIGQPVEPEIVEKARVLFP---NNS-----AILNSPHGLAGHSIVASLTWIGCEVSPELVTKCRDVTCNVSTTIP----NIINSPHDLAGYPVIASFAFVGKAVSKDLIEKARNCWQAGEY-------KVIDSPHGLAGKSIIGSFVWIGQPVSADVVEKVR--MLSTVD-------KMLESPHGLAGKPVVATLAWVGEPVTAEFVEKVRDLPSEATIYPG----SAVSSNSALAGFPIVATLAWIGDPVTPELVQEARSLWEGRSSSS-----TILESPHGLGGWPVVATLTWVGEPVSKETLNHVRMLWGEHQS-------GLLDSPIGLDGKTVFATLLITG-EVDSELLEACRSLSSPVRG------------LSSIAERPWVG--TLLCYPATDALLDGVRDALAPLGLY------TDLNNMCMFDGYTHYLNLVLVNCPIELSGVRECIEESEGVDG-------:
.
:

Nostoc sp. PCC 7120 UTEX 2576
Nostoc sp. PCC 7107
Nostoc sp.
Anabaena sp. 90
Anabaena cylindrica

-----------AGVTSLEN-GFLCRYRGASTSEVRNWFTSVWQLLRGEFF
-----------TGVTQLEH-GLLCRYRGASTSEVRNWFTAVWQILRTSFL
-----------IGVTRLEN-GFLCRYRGASTSEVRHWFTSVWQMLRVDYF
-----------AGVTRLQH-GFLCRYRGNSTSQVRSWFTNIWQMLRVSCL
-----------TGVSRLQN-GFLCRYRGNSTSEVRNWFTNVWQILRVSLL

62

Table 3.2 (cont’d):
Nostoc punctiforme sp.
Microchaete sp. PCC 7126
Chlorogloeopsis fritschii
Scytonema hofmanni UTEX 2349
Fischerella sp. PCC 9339
Fischerella sp. PCC 9431
Fischerella sp. JSC-11
Fischerella sp. PCC 9605
Scytonema hofmanni
Mastigocladopsis repens
Mastigocoleus testarum
Calothrix sp. 336/3
Rivularia sp.
Calothrix sp.
Leptolyngbya boryana
Microcoleus sp.
Coleofasciculus chthonoplastes
Synechocystis sp.
Chroococcidiopsis thermalis
Crinalium epipsammum
Gloeocapsa sp. PCC 7428
Oscillatoria nigro-viridis
Oscillatoria acuminata PCC 6304
Arthrospira platenis sp.NIES-39
Pseudomonas syringae
Klebsiella aerogenes
Helicobacter pylori

-----------VGASRLQH-GLLCRYRGSSTSEVRNWFIDVWQLLRVSFL
-----------TGVTLLTH-GFLCRYRGDSTSEVRNWFMAVWQMLRISVW
-----------VGVTRLEH-GFLCRYRGSSTAEVRNWFISVWQLLRVSFL
-----------AGVTRLEN-GFLCRYRGSSTSEVRNWFIDVWQLLRMSFL
--------HSQIGVTRLEH-GLLCRYRGYSTTEVRNWFIGVWQLLRIFFL
--------HSQIGVTRLEH-GLLCRYRGSSTAEVRNWFIGVWQLLRMSFL
--------HPQIGVTRLEH-GLLCRYRGSSTAEVRNWFIGVWQLLRMSFL
------PLSSQIGVTRLEH-GFLCRYRGSSTAEVRHWFIGVWQLLRMSFL
--------TPSVGVTRLSC-GLLCRYRGSGTTEVRNWFTSAWQLLRQSVL
--------KGEAGVTRLSC-GLLCRYRGSTTSEVRNWFTAVWQLLRQSFL
LTPIPNSKSPNQGVTRLNN-GFLCRYRGNSTAEVRNWFINVWLLLRTSFL
------SPS-AVGVTALEH-GFLCRYRGNSTSEVRNWFISVWQLLRQSIG
-PPLPIPPS-SQGVTRIEN-GLLCRYRGDSTAKVRNWFISVWQLLRISFL
-------TQYSFGVTRLEH-GFLCRYRGSSTSEVRNWFTAAWQLLRQSLL
----------EIGVTRLSI-GLLCRYRGHSSSEARRWFLAVWQLIRVSYF
--------QGEAGVTQLMS-GLLCRYRGSSTSEVRNWFTEVWQLLRLSFL
--------QGEAGVTQTQAQGLLCRYRGSSTTEVRNWFTQVWQCLRLTYL
--------SSQGGVTRLPM-GLLCRYRGSSSTEVRNWFTEIWQLLRSPYL
--------ITNYGVTRLPH-GLLCRYRGSSSIEVRNWFTSVWQ--------------QGESGVTTLLE-GMLCRYRGYSTLEARNWFIRVWELLRLAYL
--------QGETGVTRLTT-GLLCRYRGDSTTEVRQWFTEVWHLLRLSSL
--------NSTVGVTRIPN-GLLCRYRGTSTTAARDWFVNIWQLLRLSFS
--------EGEAGVTRLTH-GLLCRYRGSSTPEVRNWFSEVWQLLRLSFI
--------EGEAGATQLLS-GLLCRYRGPSSQEAIAWFTQIWQLLRPNLS
------------DLTQLPG-LLVARCLADEALHARAWLIELWKLLRPAVL
-----------AGASLTDR-LLTVRFLSDDNLICQRVMRDVWQFLRPHLT
------------AVSETASSHLCVKALAKGSEPLLHLREKIARLVTQTTT
:
: : .

Nostoc sp. PCC 7120 UTEX 2576
Nostoc sp. PCC 7107
Nostoc sp.
Anabaena sp. 90
Anabaena cylindrica
Nostoc punctiforme sp.
Microchaete sp. PCC 7126
Chlorogloeopsis fritschii
Scytonema hofmanni UTEX 2349
Fischerella sp. PCC 9339
Fischerella sp. PCC 9431
Fischerella sp. JSC-11
Fischerella sp. PCC 9605
Scytonema hofmanni
Mastigocladopsis repens
Mastigocoleus testarum
Calothrix sp. 336/3
Rivularia sp.
Calothrix sp.
Leptolyngbya boryana
Microcoleus sp.
Coleofasciculus chthonoplastes
Synechocystis sp.
Chroococcidiopsis thermalis
Crinalium epipsammum
Gloeocapsa sp.PCC 7428
Oscillatoria nigro-viridis
Oscillatoria acuminata PCC 6304
Arthrospira platenis sp.NIES-39
Pseudomonas syringae
Klebsiella aerogenes
Helicobacter pylori

SRSRKRNRNRNRKRSRV
NRSRSRNRNRTRARSRNRNRERQRGRGRNR--GKGKQRGRGKGRGKQKV

63

Table 3.2 (cont’d):
a

The K. aerogenes UreD sequence shown does not include seven amino-terminal residues
(MLPPLKK). Residues substituted by mutagenesis experiments are highlighted in yellow. The
asterisk (*) symbols indicate positions which have a single, fully conserved residue, (:) indicates
conservation between groups of strongly similar properties and (.) indicates weakly similar
properties.
To specifically guide substitution of surface residues that may facilitate protein:protein
interactions or participate in a hand-off mechanism during nickel transfer, a UreD homology
model was prepared using the Phyre2.0 server with the pdb coordinates of UreH extracted from
the UreH:UreF:UreG crystal structure (20) serving as the template (Figure 3.1A). Alignment of
this homology model with UreH using PyMol (1) resulted in a Cα root mean square deviation of
0.469 Å. The extent of conservation derived from the sequence alignment was mapped onto the
homology model by using the Consurf server (12), revealing a region of highly conserved,
surface exposed residues on the face opposite to that of the UreF binding site (Figure 3.1C) (20).
This region might reasonably function as the UreD:urease binding interface. Conserved surface
residues predicted to lie in this region of the protein (Tyr42, Glu46, Cys48, His49, His54, Asp63,
Lys86, Tyr88, Arg89, Arg148, and Glu153) were substituted. In addition, a few surface residues
on the reverse face or near the UreF interface (Arg163, Glu165, Asp169, Glu176, Thr196,
Arg211, and Arg233) were chosen for substitution. Arg211 and Arg233 are predicted to form
polar contacts with UreF based on comparison to their homologous residues in the
HpUreH:UreF:UreG crystal structure (20).

64

Figure 3.1: Homology-model guided mutagenesis of K. aerogenes UreD. (A) Phyre 2.0
homology model of KaUreD (green) aligned to HpUreH (yellow) from the HpUreH:UreF:UreG
crystal structure. HpUreF (magenta) is depicted to define the HpUreH:UreF interaction site. (B)
Water tunnels predicted for the KaUreD homology model by MOLE2.0. The color of residues
corresponds to the associated tunnel exit (tunnel 1 = magenta, 2 = yellow, 3 = orange, and 4 =
teal). Blue residues are positioned at the branch point shared by all tunnels, while red residues
are located at the entrance point of the tunnel. (C) Two views of the KaUreD homology model
colored by conservation score as assigned using the Consurf server and depicted in a surface
representation. Dark blue, white, and dark magenta denote low, average, and high conservation.
Left image is a 180° y-axis rotation of the right image. Surface residues and buried residues not
associated with predicted water tunnels are labeled. All residues listed were selected for
mutagenesis and are noted with K. aerogenes/H. pylori numbering.
An additional group of residues selected for substitution were predicted to be at least
partially buried in the protein and may be important for urease activation if the tunnel hypothesis
is correct. Channels within the H. pylori UreH:UreF:UreG complex were previously predicted
using CAVER 3.0 (13, 20); similar channels were identified on the same structure using
65

MOLE2.0 (not illustrated), an alternative software program that scans static structures for the
presence of water tunnels. When this program was applied to the K. aerogenes UreD homology
model, analogous tunnels were predicted, initiating near Glu176 at the UreF interface and exiting
through any of four pathways (Figure 3.1B). The origin of these tunnels at the UreD:UreF
interface is unchanged with the H. pylori protein and similarities exist between tunnel 1 of the
UreD homology model and the exit tunnel predicted by CAVER 3.0 analysis of
UreH:UreF:UreG. However, the three novel exit tunnels in the UreD homology model (tunnels
2, 3 and 4 in Figure 3.1B) were not predicted by either CAVER 3.0 or MOLE2.0 analysis of the
UreH:UreF:UreG structure. To test for the importance of these channels in urease activation,
selected non-surface (internal) residues were changed. For example, Glu176 was substituted with
similarly sized and smaller residues, in both cases lacking a negative charge. Additionally,
several substitutions were designed to place bulky (Phe, Tyr, or Trp) or long (Lys or Glu)
residues at positions within or at the temini of the tunnels (e.g. Val37, Asp63, Leu61, Leu65,
Ser85, Trp111, Thr128, and Thr196). For each alteration, the substitute residue was placed into
the UreD homology model using Pymol and analyzed to ensure most rotomers did not have
steric clashes. Models containing potential tunnel-blocking changes were analyzed with
MOLE2.0 to assess whether the predicted blocking substitutions exhibited the desired effect of
eliminating the tunnel. Variants which blocked the water tunnels in silico were selected for
experimental study. The positions of residues selected for study and the corresponding H. pylori
residue numbers are illustrated in Figure 3.1B and C. The set of substitutions, primers utilized,
rationale for the mutations, and conservation score of the residues are listed in Table 3.3.

66

Table 3.3: Variant forms of KaUreD, primers utilized, rationale for the changes, and
conservation scores for the residues.a
UreD substitution

Primer used (5'  3')

Rationale

V37L
Y42D
E46A
E46Q
C48A
H49A
H54A
I59Y
D63A
D63Q
L65I
L65W
S85K
K86A
Y88V
Y88F
R89A
W111Y
T128E
D142A
R148M
E153A
E153Q
R163A
E165A
D169A
E176A
E176Q
T196K
R211A
R233A

GGCCCGCTGACCCTCCAGCGCCCGTTT
CTGACCGTCCAGCGCCCGTTTGATCCGGAAGAAGAG
CAGCGCCCGTTTTACCCGGAAGAAGCAACCTGTCACCTC
CAGCGCCCGTTTTACCCGGAAGAACAAACCTGTCACCTC
CGGAAGAAGAGACCGCTCACCTCTATCTGC
GAAGAAGAGACCTGTGCCCTCTATCTGCTTC
CACCTCTATCTGCTTGCACCGCCCGGCGGCATC
CCGCCCGGCGGCTACGTCGGCGGTGAT
CATCGTCGGCGGTGCAGAGCTGACAATTAG
GGCGGCATCGTCGGCGGTCAAGAGCTGACAATT
GGCGGTGATGAGATAACAATTAGCGCG
GGCGGTGATGAGTGGACAATTAGCGCG
ATGCCTGGCGCCAAAAAGTTTTACCGC
GCCTGGCGCCAGCGCGTTTACCGCAGCAGC
GGCGCCAGCAAGTTTGTCCGCAGCAGCGGCG
GGCGCCAGCAAGTTTTTCCGCAGCAGCGGCG
GGCGCCAGCAAGTTTTACGCCAGCAGCGGC
GCGACCCTGGAGTATCTCCCGCAGGAT
GCCCGGCTGTTCGAAACCTTTCATCTT
CTGCTGGCCTGGGCACTGCTCTGCCTTG
CTGCTCTGCCTTGGCATGCCGGTGATTGGCG
CCGGTGATTGGCGCAACCTTCAGCCACGGC
CCGGTGATTGGCCAAACCTTCAGCCACGGC
ACCCTCAGCAACGCCCTGGAGGTATGGGTG
CCTCAGCAACCGGCTGGCGGTATGGGTGGAC
CCGCTGGAGGTATGGGTGGCCAATGAGCCGCTGC
GCTGCTGGTCGCACGCCTGCACCTG
CCGCTGCTGGTCCAGCGCCTGCACCTG
CCCTGGGTCGGCAAATTGCTGTGCTAT
CTCGACGGGGTGGCCGACGCGCTGGCG
CCGCCTGCTGACGGTGGCTTTCCTCAGTGAC

Block tunnel 3
Alter surface
Alter surface
Alter surface
Alter surface
Alter surface
Alter surface
Block tunnel
Disrupt tunnel 1 exit
Disrupt tunnel 1 exit
Block tunnel 1
Block tunnel 1
Block shared cavity
Alter surface
Alter surface
Alter surface
Alter surface
Block tunnel 4 exit
Block shared cavity
Disrupt shared cavity
Alter surface
Alter surface
Alter surface
Alter surface
Alter surface
Alter surface
Disrupt tunnel entrance
Disrupt tunnel entrance
Block tunnel entrance
Alter surface
Alter surface

Conservation
Scoreb
8
9
1
1
8
9
9
5
9
9
8
8
5
9
9
9
8
3
4
9
9
9
9
6
8
3
9
9
4
8
9

a

Reverse complements of the primers listed were also used. See Figure 3.1C for an illustration of
residue conservation. Bases altered from the wild-type sequence are noted in underlined bold
italic font.
b

Conservation scores are displayed on a scale from 1 to 9, with 1 denoting low and 9 denoting
high conservation.
Effects of UreD Variants on the In Vivo Activation of Urease. To determine whether
the ureD mutations affected in vivo urease activation, cells containing pKK17D* were cultured
in LB in the absence or presence of 1 mM Ni2+. SDS-PAGE was used to confirm similar levels
of urease protein were produced in all cultures (data not shown). Substitutions of surfaceexposed residues which map either to the highly conserved face of the UreD homology model or
to the putative UreD:UreF interfacial site did not appreciably alter the activation of urease.
67

Substitutions involving four buried or partially buried residues (Glu63, Ser85, Glu142, Asp176)
were shown to lead to greatly reduced (< 30%) urease specific activities relative to that of cells
containing wild-type UreD when assayed using soluble cell-free extracts from these cultures.
These residues were predicted to surround a 34.6 Å water tunnel (tunnel 1) within UreD. In vivo
activation of urease using E176A or E176Q variants of UreD resulted in activities that were 27%
and 21% of those activated with wild-type UreD. KaUreD Glu176 corresponds to Asp174 in the
H. pylori protein, where it maps to the interfacial site between UreH and UreF but is not
involved in direct bonding to UreF. The D142A variant of UreD led to urease activity that is
24% of that obtained with the wild-type protein. This residue (corresponding to Glu140 in UreH)
is predicted to be buried, forming backbone-mediated contacts with the side-chain of Thr196 and
a hydrogen bond between the carboxylate oxygen and the backbone carbonyl oxygen of Leu143,
and contributes to the β-sheet tertiary structure. Since β-sheet formation and stability is
dominated by backbone hydrogen bonding, loss of the polar interaction is unlikely to severely
affect the overall structural stability of UreD. Ser85 is positioned within UreD, with its side
chain facing the shared branch-point for the water tunnels. The S85K variant, selected to avoid
steric clashes predicted by modeling for bulky residues, yielded cell-free extract urease activities
of 13% relative to the sample prepared from cells producing wild-type UreD. Asp63 resides on
the face of UreD lying opposite the likely UreF interface and is positioned at the exit point of the
34.6 Å tunnel. Changing this residue to Ala or Gln resulted in cell-free extracts with urease
specific activities that were 4% and 9%, respectively, of that obtained from samples with wildtype protein. Somewhat surprisingly, T196K UreD, containing a substitution designed to block
the entrance point of the tunnel at the UreD:UreF interface, had only a mild effect on urease
activation (76% of wild-type activation). These findings are illustrated in Figure 3.2.

68

Figure 3.2: Urease activity of cell-free extracts from cells containing KaUreD variants. E.
coli BL21(DE3) cells were transformed with plasmid pKK17D* (encoding ureD*ABCEFG),
grown in LB containing 1 mM NiCl2 (except for the sample producing urease apoprotein, which
was not supplemented with the metal), and soluble cell-free extracts were assayed for urease
activity. Error bars represent triplicate analysis of single experimental samples.
Pull-Down Assays. To identify whether urease:UreD or UreD:UreF protein:protein
interactions were disrupted for the UreD variants deficient in urease activation, the
urease:UreD:UreF:UreGStr complexes were examined from E. coli cells containing pKKGD*.
These constructs allowed for Strep-Tactin pull-downs by making use of UreG containing a Cterminal Strep-II tag (5). Previous studies had shown the intact complex can be isolated from
such cells cultured in LB medium that lacked additional Ni2+. The five ureD constructs found to
be deficient in urease activation were inserted into pKKG, with the genes encoding wild-type
and R211A UreD (providing 90% activity relative to wild-type) used as controls. In all cases, the
soluble cell-free extracts produced similar levels of urease (Figure 3.3, lanes labeled I), and the

69

UreGStr-containing fractions eluted from the Strep-Tactin resin (Figure 3.3, lanes labeled E)
contained the urease subunits and the UreD, UreF and UreG accessory proteins. This result
demonstrates that urease:UreD and UreD:UreF interactions are maintained when using the UreD
variants, and it provides compelling evidence that these variant UreD proteins are properly
folded.

Figure 3.3: Interactions of KaUreGStr with other K. aerogenes urease proteins. E. coli
BL21(DE3) cells were transformed with pKKD*G variants producing the KaUreD variants
noted above the lanes. Soluble cell-free extracts (I, for input) were chromatographed on StrepTactin resin with the unbound wash (W) and the desthiobiotin-eluted (E) fractions analyzed by
SDS-PAGE.
Analysis of the Metal Contents and Specific Activities of Urease Samples Activated
In Vivo by Variant UreDs. To examine whether ureD mutant cultures with reduced urease
activities (<30%) produced enzyme that was altered in its metal-content, the enzyme was
enriched from selected E. coli pKKD(wild-type, D63A/Q, S85K, D142A, E176A/Q)G cells
cultured in the presence of 1 mM Ni2+, assayed for urease specific activity, and subjected to
metal-content analysis by ICP-AES. pKKD(R211A)G, which resulted in in vivo activated urease
with 75% of the activity of urease activated by wild type UreD, was similarly analyzed to act as
a control mutation. Urease samples were enriched by chromatography on anion-exchange and
size-exclusion resins to > 90% purity as determined by SDS-PAGE analysis (Figure 3.4). The
resulting urease samples were assayed for their activities and yielded results (Table 3.4) that
70

generally paralleled the activities measured in cell-free extracts (Figure 3.2). For example, the
UreD variants giving rise to reduced activity in the cell-free extracts also exhibited reduced
specific activities for the enriched proteins. Detailed comparison of these results, however,
underscores differences. For example, urease activity in cell-free extracts produced by D63A
UreD exhibited only 3.7% of the wild-type activity, whereas the urease purified from the sample
exhibited 25.7% of the wild-type enzyme activity. The samples associated with E176A UreD
also yielded greater activity for the purified enzyme than expected from the cell-free extracts. In
contrast, the purified urease specific activities were lower than expected on the basis of cell-free
urease activities for the samples associated with D63Q and D142A UreD. On inspection of the
metal-content, the enriched ureases with low urease specific activity contained less nickel than
those activated with wild-type UreD and also contained varying levels of zinc and iron (Table
3.4), with the highest levels of zinc corresponding to the most inactive urease generated by
activation with D142A UreD.

71

Figure 3.4: Purified urease activated in vivo using variant KaUreDs. E. coli BL21(DE3) cells
were transformed with either pKK17D (lane 1) or pKKD*G (lanes 2-9) and cultured in the
absence or presence (- and +) of nickel. Cell-free extracts of these cultures were used for urease
purification by anion exchange and size-exclusion chromatography. The resulting pools
containing urease were analyzed by SDS-PAGE. The KaUreDs used in cellular urease activation
are as follows: lanes 1-2, WT; 3, D63A; 4, D63Q; 5, S85K; 6, D142A; 7, E176A; 8, E176Q; 9,
R211A.
Table 3.4: Specific activities of urease samples enriched from cells containing selected
KaUreD variants.a
Specific Activityb

KaUreD variant
(-), WT (pKK17D)

(+), WT (pKKDG)
D63A
D63Q
S85K
D142A
E176A
E176Q
R211A

U/mg protein
(% WT)
0
1692 (100)
434.2 (25.7)
55.71 (3.29)
487.3 (28.8)
14.14 (0.84)
1077 (63.7)
375.0 (22.2)
1226 (72.4)

Ni/UreABC
(% WT)
0.004 (4.0 x 10-5)

1.1 (100)
0.54 (49.1)
0.22 (20)
0.39 (35.5)
0.07 (6.4)
0.51 (46.4)
0.43 (39.1)
0.76 (69.1)

a

Zn/UreABC

Fe/UreABC

0.13
0
0.16
0.34
0.06
0.52
0.09
0.03
0.21

0.27
0.02
0.21
0.07
0.30
0.14
0.11
0.13
0.29

Metal-content values were obtained from a single biological replicate. Specific activities were
obtained from a single biological replicate and a triplicate of technical replicates. The urease
samples were chromatographically enriched from cell-free extracts of E. coli pKKD*G grown in

72

Table 3.4 (cont’d): LB containing 1 mM NiCl2, except for a sample of urease apoprotein that
was obtained from E. coli pKK17D grown in LB lacking nickel. Metal contents were determined
by ICP-AES.
b

U is defined as µmole of urea hydrolyzed · min-1.

DISCUSSION
The studies outlined here give insights into the function of K. aerogenes UreD during
urease activation. Previous equilibrium dialysis studies of MBP-UreD showed that the fusion
protein can bind upwards of 2.5 nickel per UreD promoter, with zinc competing for these sites
(9). Prior ICP-AES studies quantified increased nickel content for the urease:UreD complex than
for urease under activating conditions lacking bicarbonate, also supporting nickel binding to
UreD (39). Lastly, a putative buried water tunnel in the UreH:UreF:UreG crystal structure was
predicted to span from the likely nickel-binding site in UreG through UreF and UreH,
hypothesized to function in urease activation (53); however, no direct experimental support was
reported. Here, I present evidence consistent with UreD functioning directly in nickel trafficking
via an internal tunnel rather than acting as a scaffold protein that transfers the metal ion via
surface residues.
The mutagenesis studies detailed in this work focused on the effects of substituting 27
residues, most of which are highly conserved among UreD homologues, and characterizing the
consequences of these substitutions. Mutations designed to disrupt potential metal-transfer sites
on the protein surface removed charges while either retaining the overall dimensions of the
residues (e.g., E165Q) or resulting in size differences (e.g., K86A). None of the UreD surface
residue substitutions examined led to significant reductions of urease activity, with the largest
changes noted for the K86A and Y88V variants (81% and 72%, respectively). The mild loss of
urease activity associated with these substitutions led me to hypothesize that the surface residues
examined are not of great importance to the function of UreD.
73

A second set of mutations were introduced to disrupt one or more potential internal
channels of UreD. Various substitutions include the use of several long or bulky side chains
designed to block the tunnels predicted by MOLE2.0 analysis of the UreD homology model. The
substitutions likely to affect the exits for tunnels 3 and 4 (Figure 3.1B) resulted in no loss of in
vivo urease activation relative to wild-type UreD. In sharp contrast, UreD variants with altered
residues at the ends (E176A/Q and D63A/Q) and the section prior to the initial branch point
(D142A) of tunnel 1 exhibited greatly reduced urease activation. A UreD variant designed to
block the shared branch point (S85K) also possessed reduced urease activity. One UreD variant
designed to block the exit of tunnel 1 (I59Y) had no appreciable effect on urease activation;
however, since this residue is located on a disordered loop in the homology model, it is likely
that it adopts multiple conformations that would not be accurately depicted using analysis of a
static structure. Surprisingly, T196K UreD had no appreciable effect on urease activation in vivo,
though in silico analysis of the available rotomers predicted blockage of the tunnel entrance.
Molecular dynamics simulations of this variant, as well as the variants affecting UreD function,
are currently underway to better explain this disconnect. Unfortunately, no residues facing tunnel
2 could be modeled to block or disrupt the tunnel without severe steric clashes. Taken together,
these initial results are consistent with a tunnel within UreD functioning in urease activation, but
loss of urease activity also could arise from disruption of protein:protein interactions.
Additionally, the nature of the loss-of-urease function could arise from a mis-metallated or unmetallated enzyme produced by variant UreD. Thus, further studies were carried out to probe for
UreD:protein interactions and to assess the activity and metal content of urease activated in vivo
by variant UreD.

74

To characterize whether the UreD variants of interest were capable of correctly
interacting with their known binding partners, namely UreABC and UreF:UreG, pull-down
studies were carried out using UreGStr. Previous studies using the UreGStr-producing construct
demonstrated that urease activation in this system was comparable to that observed when using
non-tagged UreG (5). Strep-Tactin pull-downs of UreGStr from cell-free extracts of E. coli cells
containing pKKD*G and cultured in the absence of nickel resulted in fractions containing
UreGStr along with the urease subunits, UreD, and UreF. This result qualitatively shows that
binding interactions are not disrupted when using the selected UreD variants; thus, the reduction
in urease activity in cells producing these UreD variants is not due to loss of protein:protein
interactions. Additionally, the ability to enrich the large complex provides evidence for the
proper folding of the UreD variants. This finding is especially important in the case of the
proposed tunnel-blocking S85K variant, where the introduction of a positively-charged residue
buried within the protein was a point of concern.
To investigate whether the reduced urease activities correlated with nickel content for
cells producing variant forms of UreD, ureases from selected cultures were enriched by anionexchange and size-exclusion chromatography and their specific activities and metal contents
were determined. The urease activities of these samples roughly correlated with their nickel
contents. Of interest, samples deficient in urease activity and nickel content, including the urease
apoprotein control, possessed contaminating levels of zinc and iron. Most notably, ureases
activated with the D63Q and D142A UreD variants had the lowest urease activity and nickel
content as well as the highest zinc occupancy. Certain UreD variants generated urease with
activities less than one would expect from their nickel content (see D63A and D63Q in Table
3.4). This, the can be thought to arise from malformation of the metallocenter, as it is known that

75

in vitro incubation of K. aerogenes urease apoprotein with nickel ions and bicarbonate leads to
activation of ~15% of the active sites even though the protein becomes fully loaded with nickel
(39). Alternatively, the composition of metallocenters could be heterogeneous in these samples,
with some active sites containing two nickel ions while others contain a nickel ion and a
contaminating metal, which would result in no urease activity.
While specific alterations of UreD clearly lead to reductions in ability to insert nickel
properly into the nascent active site, the function of UreE, UreG, and UreF are presumably
unaltered, so acquisition of nickel by UreE, transfer to UreG, and passage through UreF should
remain unaffected. Thus, one must consider how can a system designed to specifically select and
position nickel for transfer through UreD allow for spurious metal incorporation (zinc and iron).
It is known that accessory protein dissociation from urease occurs following urease activation in
vitro (49); however, it has not been detailed if the process itself (nickel ejection and transport
through the accessory proteins) induces the dissociation or if the incorporation of nickel results
in a conformation of urease that disallows for accessory protein binding. Indeed, the latter
explanation is unlikely, as it is known that the soluble maltose binding protein-UreD:UreF:UreG
accessory protein complex can interact with holourease (17) and holoUreABC:UreD:UreF:UreG
complexes have been observed following in vivo activation of urease (unpublished data). Two
explanations for the presence of contaminating metals in urease are reasonable to propose. First,
the metal contamination observed in these samples can explained by a failure of UreD (within
the UreABC:UreD:UreF:UreG complex) to deliver nickel from UreE into urease apoprotein,
allowing opportunistic metals to enter the nascent urease active site following activation-induced
dissociation of the accessory proteins from urease. Alternatively, the natural dissociation of the
accessory proteins from urease could also allow for contaminating metals to bind at the active

76

site when insufficient nickel is present within the cell, as seen in the urease apoprotein sample. In
vitro activation trials on either UreABC:UreD:UreF:UreG or UreABC:UreD:UreF:UreGStr
complexes would help answer this lingering question. Overall these findings provide further
evidence that UreD has a direct role in facilitating metal transfer into urease and support the
concept of a nickel-transfer tunnel in the protein.
In summary, the studies presented here provide evidence for a direct metal-transfer role
involving a tunnel within UreD. These findings fit well with results from previous nickel-binding
studies of UreABC:UreD and MBP-UreD, as well as the computational studies of
UreH:UreF:UreG that demonstrated metal binding by UreD and hypothesized a functional tunnel
within UreF that connected to the UreG and UreH accessory proteins (9, 39, 53).

77

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CHAPTER 4

Further Insights into the Metallocenter Vibrational Structure of Helicobacter mustelae
UreA2B2

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ABSTRACT
The enzyme urease has been studied for nearly one-hundred years, but several questions
involving the mechanism and substrate coordination of this nickel metalloenzyme still remain
unanswered. The discovery of an oxygen-sensitive, di-iron urease from Helicobacter mustelae
(UreA2B2) offers the potential to address some of these questions by use of spectroscopic
investigations that are inaccessible to the nickel enzyme due to its spectroscopically-silent active
site. Previous resonance Raman investigations of oxidized UreA2B2 provided evidence for an
Fe(III)-O-Fe(III) site that is solvent-exchange inert, even when subjected to a reduction/oxidation
cycle. Additionally, the vibrations associated with oxidized UreA2B2 were found to be sensitive
to the addition of urea, indicative of either a binding event or protein conformational changes due
to chaotropic effects. Here, I report new resonance Raman studies of UreA2B2 that demonstrate
the bridging oxygen species is a µ-oxo species which does, in fact, exchange when the enzyme is
redox cycled. I show the urea sensitivities observed previously are reversed by buffer exchange
and are not due to chaotropic effects, but arise from substrate interaction at the di-ferric
metallocenter. Additionally, I provide evidence that a known inhibitor of conventional nickel
ureases, phenyl phosphorodiamidate, yields a downshift equivalent in magnitude to that observed
with urea, whereas the inhibitor acetohydroxamic acid has no effect on the spectrum. I present
observations of a terminal hydroxide ligand that is not completely displaced on urea binding,
supporting a model where urea does not coordinate to oxidized UreA2B2 via displacement of all
terminal ligands. The discovery of a slow-binding form of UreA2B2 is also discussed, and its
solvent sensitivities are shown to be comparable to those of the fast-binding form of the enzyme.
Finally, I highlight that neither 15N-labeled nor 18O-labeled urea generate additional downshifts

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relative to normal abundance urea. These studies further define the vibrational features of the
only iron-containing urease identified.

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INTRODUCTION
Urease catalyzes the hydrolysis of urea into a molecule of ammonia and carbamate (10),
which subsequently decomposes into a second molecule of ammonia and bicarbonate. The
enzyme is found in all plants (17, 21), as well as certain bacteria, algae, Archaea and fungi (6,
14, 15, 17). Urease is the first nickel-containing enzyme to be discovered (7) and has been
identified as a virulence factor in the ulcerative human-pathogen Helicobacter pylori (9). An
abundance of structural information exists on urease from jack bean seeds (2), Sporosarcina
(formerly Bacillus) pasteurii (3), H. pylori (11), and the archetype nickel-urease from Klebsiella
aerogenes (12). These structures reveal a highly conserved active site architecture containing
two nickel atoms bridged by a carbamylated lysine side chain and a µ-hydroxy solvent molecule,
with one nickel additionally coordinated by two histidine residues and a terminal solvent
molecule, and the other metal coordinated by two histidine residues, an aspartic acid residue, and
a terminal solvent molecule. Attempts to design experiments to probe the mechanism of this dinickel site, however, are stymied by the spectroscopically silent nature of these metal atoms.
This limitation partially spurred investigations into ureases that can utilize alternative, more
tractable metals in their catalysis.
UreA2B2 is a urease from the ferret pathogen Helicobacter mustelae. This organism
contains a typical Helicobacter-like nickel-urease gene cluster (ureABIEFGH) that is induced by
nickel, with a second gene cluster containing genes homologous to the urease structural genes
(ureA2B2) that are regulated by iron and negatively regulated by nickel (20). UreA2 and UreB2
exhibit high sequence conservation with UreA and UreB (57.4% and 69.5% identity,
respectively) (5). Structural and biochemical characterization of this enzyme reveals the same
coordinating residues and geometries as found in the nickel-containing ureases, but with iron in
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place of nickel (Figure 4.1) (5). Despite the active site similarities, the K. aerogenes apoenzyme
is activated in vitro by bicarbonate and nickel or (anaerobically) iron, whereas the partial
apoprotein of the H. mustelae protein is only activated by bicarbonate plus iron. Ultraviolet
(UV)-visible (vis) spectra of oxidized UreA2B2 reveal absorption features (320, 380, and 500
nm) reminiscent of the µ-oxo bridged, diferric centers of methemerythrin and oxyhemerythrin (8,
16). These electronic features are bleached upon chemical reduction under anaerobic conditions,
accompanied by the generation of active enzyme. The fully reduced enzyme exhibits no electron
paramagnetic resonance (EPR) spectrum in the standard perpendicular mode, as expected for the
diferrous state, while the oxidized sample also yields no EPR signal, consistent with
antiferromagnetic coupling between the two ferric ions. Together, these results identify
UreA2B2 as an iron-containing metalloenzyme, the first urease identified to function with this
metal, with a bridging oxo-species coupling the two metals in the oxidized sample. This diferric
form of the enzyme uniquely allows for vibrational spectroscopy characterization of the
metalloenter under various conditions by using resonance Raman methods.

88

Figure 4.1: Structural comparison of iron- and nickel-dependent ureases. (A) The crystal
structure of H. mustelae ((UreA2B2)3)4 (PDB: 3QGA). One protomer of UreA2 is depicted as a
red cartoon and one protomer of UreB2 is depicted as a green cartoon. The remaining protomers
are depicted in dark grey. (B) The crystal structure of K. aerogenes (UreABC)3 (PDB: 1EJW).
One promoter each of UreA, UreB and UreC are depicted in red, yellow, and green cartoon,
respectively. The remaining promoters are depicted in dark grey. The active-site residues for
UreA2B2 (top) and UreABC (bottom) are shown on the right. Iron is depicted in orange, while
nickel is displayed in red.
Resonance Raman spectroscopic characterization of UreA2B2 reveals an intense Raman
shift at 500 cm-1 and an additional mode at ~790 cm-1 that are downshifted by addition of urea,
but are insensitive to rapid bulk solvent (H218O) exchange (4). These findings, together with the
EPR and structural studies highlighted above, suggested that the 500 cm -1 and 790 cm-1 modes
are the symmetric (νs) and asymmetric (νas) vibrational modes from a µ-oxo bridged, diferric
metallocenter. Surprisingly, the µ-oxo group also was reported to be solvent exchange-inert
during a reduction/oxidation cycle, which could be invaluable for designing experiments to
probe the catalytic mechanism of urease (e.g., testing whether the bridging solvent molecule is
the nucleophile in catalysis). The nature of the urea-induced vibrational downshifts was not
89

examined (4), and one could speculate that the shift arises from urea coordination at or near the
metallocenter or that it stems from a chaotropic effect of urea. Should this effect be defined as a
binding event, one could further test whether the presence of urea at the metallocenter of
UreA2B2 affects the previously identified H218O bulk solvent modes attributed tentatively as
terminal Fe-OH modes (4). This type of spectroscopic analysis of ligand-binding can also be
expanded to test whether known nickel-urease inhibitors generate UreA2B2 Raman shifts similar
to those induced by urea binding. In addition, the effect of urea isotopes on the Raman spectrum
of UreA2B2 could provide some evidence for how substrate coordinates to the metallocenter.
The work described here addresses the gaps in knowledge described above and further
characterizes the metallocenter of oxidized UreA2B2 using resonance Raman spectroscopy. My
studies identified a purification-dependent heterogeneity in UreA2B2 presenting as a slow
substrate-binding form of the enzyme and a fast substrate-binding form. In contrast to prior
results, I found that anaerobic reduction and chemical oxidation of UreA2B2 in the presence of
isotopic solvent led to exchange of the µ-oxo group; hence, identification of the nucleophile in
catalysis by assessing retention of this atom was not feasible. Additionally, I demonstrated that
the urea-induced mode shifts are not reproduced with a different chaotropic agent, and I
examined the effects of conventional urease inhibitors on the Raman spectra. Binding of either
15

N- or 18O-labeled urea to UreA2B2 led to no additional downshifts in the UreA2B2 Raman

spectrum, nor did the presence of D2O affect the primary bridging mode. This latter finding gives
further evidence that the bridging species is indeed oxo as opposed to hydroxo. My findings
presented here provide further insights into the nature of the metallocenter in this unique urease.

90

MATERIALS AND METHODS
Purification, Protein Concentration, and Activity Assays of UreA2B2. The UreA2B2
holoprotein was prepared as described previously (4), with minor modifications, from E. coli
BL21(DE3) cells transformed with pEC015 (5). Either Sephacryl S300-HR or Superdex-200
resin (GE healthcare) was used for gel filtration chromatography, with no obvious difference in
separations between the resins. Fractions containing UreA2B2 were analyzed by sodium dodecyl
sulfate polyacrylamide gel electrophoresis using 4% stacking and 12% running gels, and the
protein bands were visualized by Coomassie blue staining. Purified samples were subjected to
UV-vis spectroscopy to assess the presence of the antiferromagnetically-coupled diferric
metallocenter associated with features at 500, 380, and 320 nm. Urease activity of anaerobic,
reduced UreA2B2 was assayed as described previously (5).
Protein concentrations were assayed in triplicate by using a commercially available
colorimetric protein assay kit (Bio-Rad).
Isotopic Exchange of the UreA2B2 Oxygen Bridge with Bulk Solvent. To reinvestigate the exchange of the bridging oxo-species, chemical reduction and oxidation of
anaerobic UreA2B2 was performed in buffers containing HPLC grade H216O (Fisher Scientific)
or H218O (ICON Isotopes). Aerobically purified, UreA2B2 (3 mM) was exchanged into 250 mM
HEPES, pH 7.8, buffer containing 5 mM EDTA and gas exchanged by injecting into an argonfilled vial and incubating on ice overnight. Solutions of 50 mM HEPES, 1 mM EDTA, pH 7.8,
buffer, HPLC grade H216O, and H218O were made anaerobic with a minimum of 5 cycles of
vacuum (10 min)/argon replacement (10 min) on a Schlenk line. Powders of sodium dithionite
(Na2S2O4) and potassium ferricyanide (Sigma-Aldrich) were made anaerobic with 3 cycles of
vacuum (2 min)/argon replacement (2 min). The following steps, unless otherwise noted, were
91

performed in a 100% N2 glove bag (PlasLabs). Solutions of UreA2B2 (3 mM) were reduced with
5 mM of Na2S2O4 at room temperature for 2 h. Samples were diluted with 2 or 4 volumes of
solvent (H216O or H218O at 40 or 80% isotope enrichment, respectively), incubated for 30 min at
room temperature, and chemically oxidized with 3 mM potassium ferricyanide. To test whether
substrate turnover facilitated exchange of the bridging oxygen, samples of UreA2B2 were
reduced and isotopically enriched as described above then incubated with 50 mM urea for one h
at 37 °C. To confirm successful turnover, an aliquot was removed and assayed for urease activity
under anoxic conditions. Samples were desalted with sequential concentration and dilution in 28
mM TAPS, pH 8.4, buffer using Amicon® Ultra-0.5 centrifugal filters.
The effects of bulk isotopic solvent on UreA2B2 in the absence or redox cycling or
substrate were determined for rapid (min) exchange. Desalted samples were concentrated 10-fold
and diluted with 9 volumes of the appropriate solvent for brief time periods prior to Raman
analysis. To examine the effects of urea on the observed solvent-sensitive modes, samples of
UreA2B2 were incubated with varied concentrations of urea. In some cases, these experiments
were combined with solvent exchange, as noted above.
The Effects of Substrate, Chaotropic Agents, and Inhibitors on the Raman
Spectrum of UreA2B2. To determine if the resonance Raman vibrational modes observed in
this work and previously (5) are a result of a chaotropic effect or due to coordination to the
metallocenter, samples of untreated UreA2B2 or samples treated as described above were
incubated with 30 mM urea or 30 mM guanidinium chloride for 20 min before being analyzed by
resonance Raman spectroscopy. To test whether the urea-induced mode shift could be reversed
by the removal of urea, samples of UreA2B2 were incubated with 20 mM urea overnight and
analyzed by resonance Raman spectroscopy to confirm the presence of the urea-induced mode
92

shift. Samples were then exchanged 900-fold into 28 mM Tris-HCl, pH 7.4, and analyzed again
by resonance Raman spectroscopy. To determine if the rate of urea binding in UreA2B2 can be
increased following substrate-binding and removal via dialysis, a sample of this enzyme was
incubated overnight with 60 mM urea and dialyzed 1,000,000-fold into 280 mM TAPS, pH 8.4,
over an additional two days at 4 °C. In addition, the effects on UreA2B2 vibrational modes of
known inhibitors for the conventional nickel urease were examined: phenyl phosphorodiamidate
(PPD: ICN Biochemicals) or acetohydroxamic acid (AHA: Sigma-Aldrich). Samples of
UreA2B2 (300 µM) were incubated with 2 mM AHA, 2 mM PPD, or a buffer blank overnight on
ice before being analyzed by resonance Raman spectroscopy.
Effect of pH on the Rate of Urea-Induced Shifts on UreA2B2. The effect of pH on the
rate of urea-induced Raman shifts was determined by dialyzing 300 µl of 200 µM UreA2B2
against 300 ml of 28 mM CHES (pH 9.4), TAPS (pH 8.4), Tris-Base (pH 7.4), or MES (pH 6.4)
overnight. Samples were incubated with 26 mM urea briefly (< 1 min) before being analyzed by
resonance Raman spectroscopy.
Resonance Raman Spectroscopy. Resonance Raman analysis of UreA2B2 samples and
buffer components was performed using the 406.7 nm excitation wavelength (20 mW) of a Kr+
laser (model I-90, Coherent Inc., CA). Sample aliquots (90-105 µl) were subjected to Raman
spectroscopy using a spinning cell cuvette. Scattered light was collected at right angle geometry
and analyzed by using a single polychromator (model Triax 550, Jobin Yvon, NJ) containing a
liquid N2-cooled CCD detector (model Spectrum One, Jobin Yvon, NJ). Spectra were acquired in
10 s intervals over 3-60 min. Time periods during which the intensities of the excited modes
were stable were integrated to provide a time-averaged Raman spectra. A 2400 ln/mm grating
and slit width of 0.1 µm were used for all experiments. Difference plots were obtained by
93

subtracting one absolute spectrum from another. In some cases, a spline function was used to
normalize the baseline of the spectrum.
RESULTS
Resonance Raman Spectra, the Effect of pH, and Identification of Fast and Slow
Substrate-Binding Forms of UreA2B2 at 406.7 nm Excitation. I collected resonance Raman
spectra of oxidized UreA2B2 using 406.7 nm excitation and found it to be largely similar to the
previously described spectra obtained using 363.8 nm and 413.1 nm (4). The 406.7 nm excitation
was chosen to circumvent a photo-degradative phenomena observed from extended exposure of
the sample to the 363.8 nm laser (data not shown). Excitation of as-purified UreA2B2 revealed
the same νs(Fe-O-Fe) mode at 500 cm-1 and the νas(Fe-O-Fe) at ~782 cm-1 as observed in
previous work (4). Incubation of UreA2B2 with urea resulted in downshifts of these modes to
487 and 765 cm-1, respectively (Figure 4.2). The substrate incubation time required for the
complete downshift of the νs(Fe-O-Fe) mode was observed to be short (<5 min), and the
downshift was shown to be more rapid at pH 8.4 than at pH 7.4 (Figure 4.3). Overnight dialysis
into buffer pH’s of 6.4 and 9.4 resulted in protein precipitation. Additional studies revealed that 2
mM urea was sufficient to induce a complete downshift of the νs(Fe-O-Fe) (data not shown),
though typically the studies presented in this work were performed with concentrations in excess
of this value (60 mM). Curiously, however, several purifications of the enzyme presented with a
slow urea-binding phenotype characterized by a much slower rate of urea-induced downshift (>
3 hours, Figure 4.4). No obvious differences in protein purification, handling, or storage were
noted between the two phenotypes of UreA2B2. Overall, the resonance Raman spectra of the two
forms of the enzyme were shown to be identical in the positions and magnitudes of their Raman
shifts under all conditions tested and only differ in the time needed to obtain complete red- or
94

blue-shifts. No distinction will be made between the two forms of the enzyme for the remainder
of this chapter.

Figure 4.2: Representative resonance Raman absolute and difference spectra of oxidized
UreA2B2 (blue) and sample that had been incubated with 60 mM urea (red). Urea-sensitive
modes are highlighted by a difference plot (black trace). Noted Raman shift positions are
displayed in units of inverse wavelength (cm -1)

95

Figure 4.3: Rate of urea-induced Raman downshift at different pH values. Samples of
aerobic, oxidized UreA2B2 were supplemented with 26 mM urea and scanned by resonance
Raman spectroscopy immediately afterwards. Changes in intensity over time for the ureaunbound (500 cm-1, red trace) and urea-bound (490 cm-1, black trace) Raman shifts are shown for
UreA2B2 at (A) pH 7.4 and (B) pH 8.4. Time (s) represents time post-exposure to the laser
source and Raman intensity here represents relative abundance of each species at a given time.

96

Figure 4.4: Evidence of slow substrate binding to UreA2B2. Samples of oxidized, aerobic
UreA2B2 were incubated with 60 mM urea for three h (green) or overnight (red) or with 180
mM urea overnight (black trace) and analyzed by resonance Raman spectroscopy. A shoulder
present at 500 cm-1 indicates binding is incomplete after 3 h, while overnight incubation at the
same concentration of urea lacks this shoulder. The feature shown at 1000 cm -1 is an internal
vibration of urea.
Anaerobic Chemical Reduction and Oxidation Facilitates the Complete BridgeExchange for UreA2B2. The reported resistance to isotopic substitution of the µ-oxo bridge
during anaerobic redox cycling (4) was reexamined. Anaerobic samples of oxidized UreA2B2
were subjected to reduction with Na2S2O4 in the presence or absence of urea and confirmed to be
reduced by assaying for urease activity under anaerobic conditions (data not shown). Resonance
Raman spectra of UreA2B2 that had been reduced in H218O in the presence or absence of urea
and then oxidized with potassium ferricyanide revealed complete and symmetric downshifts of
the νs(Fe-O-Fe) from 500 cm-1 to 484 cm-1 and of the νas(Fe-O-Fe) modes from 783 cm-1 to 742
cm-1 (Figure 4.5A and B). These downshifted modes remained present following extensive
desalting of the oxidized sample in buffer prepared with H216O. The 16 cm-1 downshift of the νs
is in good agreement with 16O/18O-shifts observed in other µ-oxo bridged enzymes (16, 19), but
97

it contradicts the previously reported, exchange-inert behavior of UreA2B2. The νas(Fe-O-Fe)
mode shift was close to that observed in methemerythrin (16), but the weakness in the intensity
of this mode made attaining Raman shift numbers difficult. The complete downshifts of the
νs(Fe-O-Fe) and νas(Fe-O-Fe) modes allowed me to test the number of and coordination state of
oxygens incorporated into the bridging site by redox cycling UreA2B2 in 40% H218O solvent.
The difference spectrum for UreA2B2 cycled in H216O or 80% H218O versus UreA2B2 cycled in
40% H218O showed only two Raman shift positions, which supports assignment of the bridging
oxo species as a µ-oxo bridge involving one oxo species as opposed to a bis(µ-oxo) bridge,
which would present as three Raman shift positions in the mixed isotope spectrum (Figure 4.5A).
Incubation of 18O-substituted UreA2B2 with urea led to further downshifts of the νs and νas(FeO-Fe) modes that are equivalent in magnitude relative to those observed in the 16O-substituted
enzyme (Figure 4.5C).

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Figure 4.5: The effect of anaerobic chemical reduction and oxidation of UreA2B2 in H218O
solvent in the presence or absence of substrate.(A) The resonance Raman absolute spectra of
UreA2B2 redox cycled in H216O (red), 80% H218O (blue), or 40% H218O (green) in the absence
of substrate. 18O, Redox-exchange sensitive modes are displayed in a difference plot of H216O
and 80% H218O (black trace), while the binding-mode of oxygen is displayed in the difference
plots for H216O or 80% H218O and 40% H218O (yellow or magenta, respectively) (B)
H216O/H218O difference plot comparison of UreA2B2 redox cycled in the absence (top trace) or
presence (bottom trace) of 60 mM urea. The samples containing urea were shown to have urease
activity (data not shown). (C) The effect of urea on 16O- or 18O-bridged UreA2B2. A difference
plot of UreA2B2 redox cycled in H216O versus H218O (red) is compared to similarly treated
samples with 60 mM urea added following sample oxidation (blue).

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Urea-Induced Shifts are not due to Chaotropic Effects in UreA2B2. The effects of
urea on the resonance Raman spectrum of UreA2B2 were previously characterized (4), but that
work did not address if the mode shifts observed were due to conformational changes arising
from the denaturing effect of urea or to interaction with the metallocenter. To begin to address
this question, samples of UreA2B2 with urea-induced mode shifts were dialyzed extensively into
buffer lacking urea and analyzed again by resonance Raman spectrometry. The resonance Raman
spectra for the samples were upshifted from 490 cm -1 to 500 cm-1 (Figure 4.6, red, blue, and
green traces). Furthermore, samples of UreA2B2 were incubated with an equal concentration of
the alternative chaotropic agent guanidinium chloride and subjected to similar analysis. The
spectrum of this sample showed no shifts relative to untreated enzyme (Figure 4.6, black trace).
Finally, two other urea-like molecules were examined for their effects on the (Fe-O-Fe)
associated Raman shifts. These compounds were the known nickel-urease inhibitor, PPD, and
the well-characterized nickel- and iron-urease inhibitor AHA. Overnight incubation of UreA2B2
with 2 mM PPD resulted in a downshift in the νs(Fe-O-Fe), while similar treatment with AHA
showed no appreciable shifts in any of the metallocenter-associated vibrational modes (Figure
4.6, purple and yellow traces). Taken together, these results are consistent with the urea-induced
shifts arising from a binding event at the metallocenter as opposed to chaotropic effects.

100

Figure 4.6. Examination of the basis for the urea-induced Raman shifts on UreA2B2. The
absolute spectrum of UreA2B2 (red), after incubation with urea (blue), and when subjected to
dialysis in buffer lacking substrate (green). The absolute spectrum of UreA2B2 treated with the
chaotropic agent guanidinium chloride (black), the known UreA2B2 inhibitor AHA (black
dotted), and the known urease inhibitor PPD (yellow).
The Effects of 15N- and 18O-Urea on the Resonance Raman Spectrum of UreA2B2.
To determine if the urea-sensitive modes are a result of direct coordination of the metallocenter
to substrate, samples of UreA2B2 were incubated on ice with 60 mM unlabeled urea or 15Nlabeled urea for 3 h or with 60 mM 18O-labeled urea for 5 h before being analyzed by resonance
Raman spectroscopy. The absolute spectra of these samples all show a complete shift to the ureabound mode of UreA2B2, with no detectable differences among the different isotopic forms of
substrate, as illustrated in the double-difference plots (Figure 4.7).

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Figure 4.7: The effects of isotopically-labeled urea on the resonance Raman spectrum of
UreA2B2. Samples of UreA2B2 were incubated overnight with 60 mM of unlabeled urea (red),
15
N-labeled urea (blue) or 18O-labeled urea (green) and analyzed by resonance Raman
spectroscopy. Difference spectra were used to highlight N-isotope sensitive modes (grey) and Oisotope sensitive modes (black).
Rapid Bulk Isotopic Solvent Exchange Provides Evidence of Terminal Solvent
Modes in UreA2B2. To examine the effect of urea on the solvent isotope sensitive modes of
UreA2B2, bulk solvent exchange studies were performed. Rapid D2O or H218O exchange just
prior to Raman analysis identified solvent proton- and oxygen-isotope sensitivities for UreA2B2
(Figure 4.8, top panels). These solvent-sensitive modes were upshifted in their relative positions
by ~10 cm-1 in the presence of urea (Figure 4.8, bottom panels). Furthermore, comparison of the
Raman spectra for 16O-bridged and 18O-bridged UreA2B2 in normal water versus D2O resulted
in no additional downshifts in the νs(Fe-O-Fe) (Figure 4.8, top panels, black traces), providing
further evidence that this mode contains a bridging oxo group as opposed to a hydroxo group
(Figure 4.8, bottom panels). The deuterium and oxygen sensitivities of the ~530 cm-1 to 511 cm-1

102

downshift point to it being a stretching mode of terminal hydroxide coordinated to a ferric ion as
hypothesized previously (4) and the existence of this terminal solvent precludes a substrate
coordination scheme resulting in the displacement of both terminal solvent molecules in the
oxidized metallocenter (3).

Figure 4.8: The effect of rapid bulk solvent exchange on 16O- or 18O-bridged UreA2B2 in
the presence or absence of substrate. UreA2B2 was redox cycled in H216O or H218O, diluted
into 90% of H218O (left panels) or D2O (right panels), incubated in the absence (top panels) or
presence of 60 mM urea (bottom panels) and analyzed by resonance Raman spectroscopy. In all
panels, the difference between the isotopic bridges (black) is compared to the difference between
the bulk solvent-sensitive modes (blue). For the top panels, a trace showing the uncorrected bulk
solvent difference is displayed for reference (green).

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DISCUSSION
The studies presented here have furthered our understanding of the oxidized
metallocenter of this unique urease. The pioneering studies on this enzyme highlighted several
important properties (5), but the most intriguing finding was the solvent-exchange inert nature of
the bridging oxo-species in the diferric metallocenter (4). In particular, it was surprising that that
reduction of the enzyme, while successfully showing loss of the µ-oxo bridge chromophore by
UV-vis spectroscopy, did not allow for exchange of the bridging oxo-group with bulk solvent.
Here, we have shown that chemical reduction and oxidation in the presence of isotopic solvent
does allow for the complete exchange of the bridging oxo-species in UreA2B2 independent of
substrate turnover. It is difficult to explain why the previous studies did not observe oxygenisotope solvent exchange by redox-cycling the enzyme, as reduction of the metallocenter was
clearly observed in that work. In contrast to the former study, I used a chemical oxidant whereas
the prior work used molecular oxygen for oxidation, but one expects to observe the isotopic shift
in either case. Comparison of the 16O/18O Raman shifts for this enzyme and the µ-oxo bridged,
diferric methemerythrin shows an identical wavenumber downshift (16). This result provides
strong evidence that the oxidized state of the UreA2B2 metallocenter is indeed a µ-oxo bridge
instead of a µ-hydroxo bridge. Additional evidence against a hydroxo bridging species comes
from the difference spectra comparing 16O- versus 18O-bridged UreA2B2 which show no
additional mode shifts when comparing enzyme in H2O versus D2O. Because the bridging
oxygen exchanges with solvent when the enzyme is reduced, my initial plan to identify the
nucleophilic water molecule by monitoring for loss of this atom during turnover was abandoned.
The metallocenter structures of urease from K. aerogenes, H. pylori, and S. pasteurii
highlight a triad of water molecules positioned at the metallocenter, with one molecule serving as

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a bridging water and two acting as terminal ligands to each nickel (3, 11, 12). Structures of
AHA-bound urease from H. pylori (PDB 1E9Y) and diamidophosphate-bound urease from S.
pasteurii (PDB 3UBP, where diamidophosphate derives from the slow enzymatic hydrolysis of
PPD (1)) show that this solvent triad is displaced following inhibitor binding. I have shown that
PPD, but not AHA, generates a downshift in the νs(Fe-O-Fe) mode equivalent in magnitude to
the urea-induced downshift, even though it has been previously established that AHA is an
inhibitor of UreA2B2 (5). The exchange-inert µ-oxo bridge of UreA2B2 precludes displacement
of the bridging solvent molecule seen in both crystal structures, so it is still unclear how PPD,
AHA, or urea, coordinate to the diferric site. Nevetheless, I hypothesize that the bulk-solvent
sensitive modes, specifically the likely terminal Fe-OH mode at ~530 cm-1, is absent in samples
treated with either inhibitor. My results provide further evidence that the solvent-sensitive modes
are from a terminal hydroxide and that urea coordination at the metallocenter alters their
orientation, but does not lead to their displacement.
The surprising fast versus slow urea-binding phenotype of various UreA2B2 preparations
caused me to examine my purification protocols in detail. However, no obvious deviations were
noted when comparing my purification and handling protocols to the previously published work
(4). I tested several hypotheses in an attempt to generate fast substrate-binding UreA2B2 from
pEC015 expressing cells, including rapid sequential purification and freeze-thaw avoidance, but
these adjustments did not produce fast substrate-binding UreA2B2. I also attempted to convert
slow substrate-binding UreA2B2 into the fast substrate-binding UreA2B2 form by building on
the hypothesis that a tight-binding inhibitor was present during purification. Though I could not
identify a likely theoretical inhibitor that would be introduced during purification, I reasoned that
displacement of the inhibitor with substrate followed by buffer exchange to remove both

105

inhibitor and substrate could change the phenotype. Unfortunately, application of this treatment
on slow substrate-binding UreA2B2 did not result in an appreciable change in the rate of ureainduced downshifts. I propose additional tests to identify conditions for the generation of fast
substrate-binding UreA2B2 in the concluding remarks of this dissertation.
The rapid bulk H218O-solvent exchange studies presented in this work mirror the
extensive H218O-solvent exchange presented previously (4), with an observed trough at ~511 cm1

and peak ~530 cm-1. In this study, the effect of D2O was more extensively characterized, with

H2O/D2O difference plots showing a similar isotopic shift, with shifts comparable in position and
magnitude. The exact position of these shifts is difficult to discern due to the relative weak
intensity of these modes. However, the solvent oxygen- and proton-isotope sensitivity of this
feature, its weak intensity, and its clear separation from the νs(Fe-O-Fe) mode support its
assignment as a terminal solvent or hydroxide coordinated to the metallocenter. This assignment
can be likened to the high pH Raman spectrum for horseradish peroxidase which arises from a
terminal Fe-OH stretch (18). The peroxidase work identified this mode to be D2O sensitive, but
this sensitivity presented as an upshift, reasoned to be due to hydrogen bonding to a deprotonated
His residue, which is precluded in UreA2B2. Deuterium exchange for Fe-OH stretches that are
not participating in local hydrogen bonding are expected to result in downshifts of 20 cm -1 (13),
similar to what is presented in my studies. The upshift of these features in the presence of urea
could be caused by a slight displacement of the bound hydroxyl group on urea occupancy. This
would present as an altered bond length or angle within the Fe-OH molecule, resulting in an
upshift for both the H218O and D2O sensitive shifts. The new mode at ~ 486 cm-1 following
bridging oxo exchange with 18O can be interpreted as a new Fe-OH bending mode as suggested

106

previously, however its overlap with the main νs(Fe-O-Fe) and weak intensity make its
assignment difficult.
The reversibility of the urea-induced sensitivity in UreA2B2 and the lack of effect when
using an alternative chaotropic agent were crucial for distinguishing if the resultant mode shifts
were due to changes in the protein structure caused by a a denaturing effect or by substrate
binding to the metallocenter. One could imagine that the urea-induced downshifts are due to
local denaturation at the metallocenter, which could result in altered symmetric stretch angles of
the Fe(III)-O-Fe(III) metallocenter and downshifts in the Raman spectrum. However, no
denaturing effect would be expected from PPD treatment of UreA2B2, especially at the
concentrations tested. This finding supports the hypothesis that urea coordination at the active
site causes an altered orientation of the terminal solvent molecule, presumably bound at the nonsubstrate coordinating iron, which results in the observed Raman upshifts for the terminal water
mode. In conclusion, strong evidence is presented in this work that the urea-induced mode shifts
we observe for UreA2B2 are evidence of a binding-event at or near the metallocenter.

107

BIBLIOGRAPHY

108

BIBLIOGRAPHY

1.

Andrews, R. K., Dexter, A., Blakeley, R. L., and Zerner, B. 1986. Jack bean urease
(Ec 3.5.1.5) .8. on the inhibition of urease by amides and esters of phosphoric-acid. J.
Am. Chem. Soc. 108: 7124-7125.

2.

Balasubramanian, A., and Ponnuraj, K. 2010. Crystal structure of the first plant urease
from jack bean: 83 years of journey from its first crystal to molecular structure. J. Mol.
Biol. 400: 274-283.

3.

Benini, S., Rypniewski, W. R., Wilson, K. S., Miletti, S., Ciurli, S., and Mangani, S.
1999. A new proposal for urease mechanism based on the crystal structures of the native
and inhibited enzyme from Bacillus pasteurii: why urea hydrolysis costs two nickels.
Structure 7: 205-216.

4.

Carter, E. L., Proshlyakov, D. A., and Hausinger, R. P. 2012. Apoprotein isolation
and activation, and vibrational structure of the Helicobacter mustelae iron urease. J.
Inorg. Biochem. 111: 195-202.

5.

Carter, E. L., Tronrud, D. E., Taber, S. R., Karplus, P. A., and Hausinger, R. P.
2011. Iron-containing urease in a pathogenic bacterium. Proc. Natl. Acad. Sci. U.S.A.
108: 13095-13099.

6.

Cox, G. M., Mukherjee, J., Cole, G. T., Casadevall, A., and Perfect, J. R. 2000.
Urease as a virulence factor in experimental cryptococcosis. Infect. Immun. 68: 443-448.

7.

Dixon, N. E., Gazzola, T. C., Blakeley, R. L., and Zermer, B. 1975. Letter: Jack bean
urease (EC 3.5.1.5). A metalloenzyme. A simple biological role for nickel? J. Am. Chem.
Soc. 97: 4131-4133.

8.

Duff, L. L., Klippenstein, G. L., Shriver, D. F., and Klotz, I. M. 1981. Multiple
conformations at functional site of hemerythrin: Evidence from resonance Raman
spectra. Proc. Natl. Acad. Sci. U.S.A. 78: 4138-4140.

9.

Eaton, K. A., Brooks, C. L., Morgan, D. R., and Krakowka, S. 1991. Essential role of
urease in pathogenesis of gastritis induced by Helicobacter pylori in gnotobiotic piglets.
Infect. Immun. 59: 2470-2475.

10.

Farrugia, M. A., Macomber, L., and Hausinger, R. P. 2013. Biosynthesis of the urease
metallocenter. J. Biol. Chem. 288: 13178-13185.

11.

Ha, N. C., Oh, S. T., Sung, J. Y., Cha, K. A., Lee, M. H., and Oh, B. H. 2001.
Supramolecular assembly and acid resistance of Helicobacter pylori urease. Nat. Struct.
Biol. 8: 505-509.

12.

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.
109

13.

Loehr, T. M., and Plane, R. A. 1968. Raman spectra and structures of arsenious acid
and arsenites in aqueous solution. J. Inorg. Chem. 7: 1708-1714.

14.

Mobley, H. L., Island, M. D., and Hausinger, R. P. 1995. Molecular biology of
microbial ureases. Microbiol. Rev. 59: 451-480.

15.

Rees, T. A., and Bekheet, I. A. 1982. The role of nickel in urea assimilation by algae.
Planta 156: 385-387.

16.

Shiemke, A. K., Loehr, T. M., Sanders-Loehr, J. 1984. Resonance Raman study of the
µ-oxo-bridged binuclear iron center in oxyhemerythrin. J. Am. Chem. Soc. 106: 49514956.

17.

Sirko, A., and Brodzik, R. 2000. Plant ureases: roles and regulation. Acta Biochim. Pol.
47: 1189-1195.

18.

Sitter, A. J., Shifflett, J. R., and Terner, J. 1988. Resonance Raman spectroscopic
evidence for heme iron-hydroxide ligation in peroxidase alkaline forms. J. Biol. Chem.
263: 13032-13038.

19.

Sjoberg, B. M., Loehr, T. M., and Sanders-Loehr, J. 1982. Raman spectral evidence
for a µ-oxo bridge in the binuclear iron center of ribonucleotide reductase. Biochemistry
21: 96-102.

20.

Stoof, J., Breijer, S., Pot, R. G., van der Neut, D., Kuipers, E. J., Kusters, J. G., and
van Vliet, A. H. 2008. Inverse nickel-responsive regulation of two urease enzymes in the
gastric pathogen Helicobacter mustelae. Environ. Microbiol. 10: 2586-2597.

21.

Witte, C. P. 2011. Urea metabolism in plants. Plant Sci. 180: 431-438.

110

CHAPTER 5

Additional Studies

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This chapter summarizes a series of disparate studies of both the Klebsiella aerogenes
urease components and the Helicobacter mustelae UreA2B2 iron-containing urease. These
investigations do not fit well into the earlier chapters, but they include a few novel findings that
are worth recording to assist in future studies of these systems.
ADDITIONAL K. aerogenes UREASE STUDIES
Equilibrium Dialysis Studies of MBP-UreDFG Using Nickel. Prior studies had
reported that UreG and MBP-UreD bind nickel (0.5 and 2 Ni atoms per protomer, respectively)
(1 ,6), raising questions of whether these sites are retained and whether additional sites exist in
MBP-UreDFG. To assess the nickel-binding capacity of this complex, I dialyzed 200 µl samples
of MBP-UreDFG (20 µM) in 50 mM Tris buffer, pH 7.4, containing 25 mM NaCl against buffer
containing varied concentrations of NiCl2·6H2O (12.5, 25, 50, 100, 200, 400, 800 or 1600 µM)
overnight at 4 °C. I determined nickel concentrations for both the dialysis buffer and the dialyzed
protein by colorimetric assay using the metal-binding probe 4-(2-pyridylazo) resorcinol (Figure
5.1). Experimental samples were compared against a nickel standard curve.

112

Figure 5.1: Nickel binding to MBP-UreDFG as determined by equilibrium dialysis. Ni2+ per
MBP-UreDFG was determined by subtraction of [Ni2+] in the dialysis tubing from [Ni2+] in the
protein sample, divided by the concentration of protein.
The results show that increasing nickel concentrations led to increasing numbers of nickel
atoms weakly bound to MBP-UreDFG that seemed to approach a plateau; however, additional
replicates would be required to be able to report binding constants. Furthermore, the data are
somewhat compromised because protein samples experienced significant precipitation above 400
µM Ni2+. A preliminary conclusion from these data is that MBP-UreDFG bound greater numbers
of nickel atoms than expected for the separate MBP-UreD and UreG components, consistent
with nickel binding to UreF when present in the complex or to new binding sites being exposed
or created within the complex compared to the individual proteins. If these studies are repeated
by others, it may be informative to extend the analysis by addition of GTP/Mg2+ to determine if
these ligands have an effect on nickel binding to this complex.
Effects of Urease Activation Conditions on MBP-UreDFG Stability. The MBPUreDFG complex provided me with a platform to study the stability of the accessory protein
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complex when subjected to various conditions in the presence or absence of urease. The
accessory proteins are known to exist as a complex separate from urease (20) and activation of
urease results in dissociation of the accessory proteins from the structural subunits (22), but it is
unknown if the accessory proteins dissociate from one another during this process or dissociate
as a unit. To examine the protein:protein interactions of the accessory proteins under urease
activation conditions, I incubated MBP-UreDFG and the MBP-UreDFG:urease apoprotein
complex in buffers containing one or more of the following components: 100 µM NiCl 2·6H2O,
100 mM sodium bicarbonate, and 400 µM Li·GTP plus 800 µM MgCl2. These treated samples
were studied for their protein:protein interactions using an amylose-affinity resin pull-down
approach (Figure 5.2).

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Figure 5.2: The effects of urease activation components on the stability of MBP-UreDFG
and MBP-UreD:urease apoprotein. (A) Samples of 3 µM MBP-UreDFG were incubated in
buffer alone (untreated, UT) or buffer containing one or more of 100 mM sodium bicarbonate
(B), 100 µM NiCl2·6 H2O (N), and 400 µM Li·GTP plus 800 µM MgCl2 (G), and subjected to
amylose affinity chromatography. Samples of the reactions prior to chromatography (input, I),
the first wash step (W) and the final elution (E) with 10 mM maltose are displayed. (B) Samples
of MBP-UreDFG incubated with UreABC apoprotein were treated and chromatographed as
described above. Additional lanes: untreated MBP-UreDFG (1); untreated UreABC apoprotein
(2), protein standard (S).
As shown in Figure 5.2A, MBP-UreDFG formed a stable complex that elutes from
amylose resin with very little protein lost during the wash step when using the untreated sample.
This association was at least partially lost in all samples containing GTP/Mg2+, as evidenced by
the appearance of the UreG band in the wash fractions and the loss of intensity of the UreG band
in the elution fractions. This loss seemed to be enhanced when nickel was present, as is evident
when comparing the fractions from GTP/Mg2+/Ni2+ and GTP/Mg2+/Ni2+/bicarbonate samples to
the GTP/Mg2+/bicarbonate sample. These results fit well with what was discovered in the
Helicobacter pylori urease system (16). That study reported the ejection of UreG from a soluble

115

UreH:UreF:UreG complex in a GTP-dependent manner, with UreG dissociation enhanced by
the presence of Ni2+.
In Figure 5.2B, I present similar experiments using the complex of MBP-UreDFG with
urease. Again, the untreated sample is stable to the chromatographic step and elutes as an intact
complex with maltose. In contrast to what was observed for MBP-UreDFG in panel A, the MBPUreDFG:urease apoprotein complex was unaffected by GTP alone, but dissociated to urease and
MBP-UreDFG in the presence of Ni2+ or Ni2+/bicarbonate. For the case of
Ni2+/bicarbonate/Mg2+/GTP, the data are consistent with initial release of MBP-UreDFG and
subsequent UreG dissociation, but the sequence of steps was not tested in this study. The nickeldependent dissociation of urease from the initial complex cannot be related to the active-site
metal ion occupancy of urease because in Chapter 2 I demonstrated that MBP-UreDFG forms a
stable interaction with urease holoprotein in vitro. Rather, I propose that added nickel binds to
one or more sites on the MBP-UreDFG complex or on urease (other than at the urease catalytic
center) to cause a physical change in the quaternary state that leads to dissociation of urease.
The GTP-dependent dissociation of UreG from MBP-UreDFG raised the question of the
quaternary state of the released UreG. It is well established that K. aerogenes UreG purifies as a
monomer that is not affected by added nickel or GTP (1, 20), whereas nickel-loaded, GTPbound, H. pylori UreG exists as a dimer (16). To examine the quaternary state of UreG released
from GTP-treated MBP-UreDFG, I incubated 60 µM protein complex for one h at room
temperature in 50 mM Tris buffer, pH 7.4, containing 150 mM NaCl or with the same buffer
supplemented with GTP/Mg2+, GTP/Mg2+/Ni2+, or GTP/Mg2+/Ni2+/bicarbonate in concentrations
equivalent to those listed above. The samples were analyzed using a 100 ml Sephadex-200 gel
filtration column pre-equilibrated in 50 mM Tris buffer, pH 7.4, containing 150 mM NaCl, and

116

eluted in the same buffer (Figure 5.3A). Fractions corresponding to elution peaks were pooled,
concentrated 50-fold, and analyzed by sodium dodecyl sulfate-polyacrylamide gel
electrophoresis (SDS-PAGE) (Figure 5.3B).

Figure 5.3: Size-exclusion chromatography of MBP-UreDFG exposed to various activation
conditions. (A) Size-exclusion chromatographs of MBP-UreDFG samples incubated in 50 mM
Tris buffer alone (red trace) or buffer supplemented with GTP/Mg2+ (green), GTP/Mg2+/Ni2+
(magenta) or GTP/Mg2+/Ni2+/bicarbonate (black). A sample of purified UreGStr (blue trace) is
included as a reference. (B) A representative SDS-PAGE gel containing the labeled fractions
from the trace above. Untreated MBP-UreDFG (lane M) is compared against peaks 1-4 in the
treated samples. Protein standards are located in lane S.
As illustrated above, treatment of MBP-UreDFG with buffers containing combinations of
GTP and nickel resulted in the decomposition of (MBP-UreDFG)2 into two species: dissociated
UreG (Figure 5.3B, lane 3) and a species approximated by ((MBP-UreDF)2:UreGx)y (Figure
5.3B, lane 1; where ‘x’ was clearly substoichiometric and ‘y’ was not determined). A third
117

species eluted at roughly the same time as untreated MBP-UreDFG (Figure 5.3B, lane 2) and
contained all of the subunits, likely representing unaffected complex. Comparison of the elution
time for the dissociated UreG versus purified UreGStr (Figure 5.3A, blue trace) to the elution
positions of standard proteins yields estimated sizes of 50 kDa for dissociated UreG and 27 kDa
for UreGStr, which suggests that UreG is ejected from the MBP-UreDFG complex as a dimer. An
incomplete ejection of UreG is observed in peak 1 of the GTP- and nickel-treated samples
(Figure 5.3, black and magenta traces). Whereas UreG dissociation from the H. pylori complex
provided the monomeric protein in the presence of bicarbonate (16), I observed no difference in
the elution profiles when comparing GTP/Mg2+/Ni2 treatments in the presence or absence of
bicarbonate (Figure 5.3A, black and magenta traces). Perhaps of significance, my elution buffers
lacked GTP, Mg2+, Ni2+, or bicarbonate, so a more complete study could examine the effects of
these buffer components on the chromatographic behavior. Interestingly, the ejection of UreG
generated a larger complex with an approximate molecular weight of 880 kDa. A prior study
reported a large oligomeric complex for purified MBP-UreD via size-exclusion chromatography,
and the authors hypothesized this was caused by associations among the MBP-tags (6). My
results can be explained similarly, although further experiments with differentially tagged
complexes would help to address this issue. As a final comment, the fourth species observed at
the end of the elution with an estimated size of less than 5 kDa was not due to GTP as shown by
analysis of GTP-containing buffer. The identity of this peak is still in question.
Interactions Between MBP-UreDFG and UreE. Previous studies identified both a
complex containing urease apoprotein, UreDFGStr, and UreE (according to SDS-PAGE analyses
of Strep-tactin pull-down fractions) and a UreG:UreE interaction that was observed in vitro in
the presence of nickel and zinc (1). These findings raised the questions of whether UreE also

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interacts with MBP-UreDFG and whether the presence of divalent metals affects this interaction.
To answer these questions, I incubated samples of MBP-UreDFG and His144*UreE (a truncated
UreE missing its C-terminal 15 residues) in 50 mM Tris buffer, pH 7.4, containing 25 mM NaCl
or the same Tris buffer supplemented with 100 µM ZnCl2 or 100 µM NiCl2·6H2O. After 30 min
on ice, the mixtures were incubated with 50 µl of amylose resin (pre-equilibrated in the Tris
buffer) and placed at room temperature with mixing for an additional hour. The resin was
washed several times with 250 µl of the same buffer and eluted in 100 µl of Tris buffer
containing 10 mM maltose. Samples from each step in treatment were analyzed by SDS-PAGE
(Figure 5.4).

Figure 5.4: The effects of nickel and zinc on MBP-UreDFG:UreE interactions. MBPUreDFG (2 µM) was incubated with 10 µM UreE for one hour on ice in 50 mM Tris buffer, pH
7.4, and 25 mM NaCl (Untreated). This buffer was supplemented with 100 µM of NiCl2·6H2O
(Nickel-treated) or 100 µM ZnCl2 (Zinc-treated). The lanes include: S, protein standards; A,
MBP-UreDFG; B, H144*UreE; 1, amylose-resin inputs; 2-5, wash fractions; 1-4; 6, maltose
elutions.
The above study shows that UreE eluted in the wash step regardless of treatment
conditions, and it did not form a complex with MBP-UreDFG. The results also revealed a slight
dissociation of UreG from the MBP-UreDFG complex, with the ejection of UreG being
unaffected by the presence of divalent metals. The lack of binding between UreE and the

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complex is somewhat surprising given the prior findings and this result may be attributed to
steric interference by the MBP fusion tag; e.g., it could be reasoned that MBP obstructs the
UreE:UreG interfacial site. However, this conclusion is weakened by recent work suggesting that
UreE and UreF interact at the same site on UreG (19). To circumvent any concern about
interference by MBP, an alternative tag could be used; however, solubility of UreDFG is an issue
for complexes not containing the MBP-UreD fusion protein. It is also possible that the results are
skewed because it relies on pull-down of MBP-UreD; a confirmatory experiment could be
carried out using Strep-II tagged UreG in the complex or placing this tag on UreE. I would not
recommend using the natural affinity of UreE for Ni-NTA resin because this requires the
confounding presence of nickel for elution in the chromatographic step.
Crystallization Attempts on MBP-UreDFG and UreABC:UreD. Although the crystal
structure of the UreH:UreF:UreG complex from H. pylori is known (15, 16), the many
distinctions between that urease system and the one from K. aerogenes made it desirable to
subject MBP-UreDFG to crystallization trials. I purified samples of MBP-UreDFG using a
modified protocol (14). In particular, I used Q-Sepharose resin in place of DEAE-Sepharose for
the second chromatographic step, while retaining the same elution protocol. Following sizeexclusion chromatography, I analyzed fractions containing MBP-UreDFG for their purity by
SDS-PAGE, pooled and concentrated them to 1 ml, and incubated the sample on ice for 3 days.
To ensure the complex was maintained during this time-frame, I subjected the pooled
concentrate to size-exclusion chromatography a second time and examined it for purity (Figure
5.5).

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Figure 5.5: Size-exclusion chromatography of MBP-UreDFG. (A) MBP-UreDFG samples
were purified by amylose and Q-Sepharose resins prior to being concentrated and
chromatographed on Sephadex-200 size exclusion resin. The chromatographs using for the first
(left) and second (right) Sephadex-200 separations are displayed. (B) SDS-PAGE analysis of the
fractions from each of the labeled elution peaks above (1-3) compared to protein standards (S).
The MBP-UreDFG complex was stable over the short time-span examined and eluted as
a single species off the Sephadex-200 resin. My studies from Chapter 2 demonstrated that MBPUreDFG exists solely as a stable dimer of the hetero-trimeric species in the absence of the urease
structural subunits. I have noted dissociation of UreG from the MBP-UreDFG complex for
samples that were thawed for longer time-periods (weeks), but in collaboration with
crystallographer Dr. Jian Hu (Michigan State University), we concluded this complex was a
viable candidate for initial protein crystallographic screening.
Samples of MBP-UreDFG purified as described above were concentrated to 8 mg/ml and
exchanged into 5 mM Tris buffer, pH 7.4, containing 1 mM β-mercaptoethanol, 1 mM EDTA,
and 12.5 mM NaCl. Crystal screens used a Mosquito robot (TPPlabtech. Cambridge, MA) and
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Wizard™ Classic (1 and 2, as well as 3 and 4), and the Hampton™ Salt, Crystal Screen, Index,
and PEG 96-well crystal screen trays. Samples were prepared by the sitting-drop method, stored
at 21 °C for four months, and checked on a semi-weekly basis for crystal formation.
Unfortunately, I observed no crystals during this timeframe. Previous MBP-UreDFG
purifications yielded stable protein at concentrations of 38 mg/ml, so future crystal trials could
examine greater concentrations of MBP-UreDFG to see if that change facilitates crystal
formation.
A crystal structure of the UreABC:UreD complex also was desired. The mutagenesis
studies of UreD in Chapter 3 of this dissertation provided evidence for a buried water tunnel that
plays a role in metal transfer into urease, and a structure could demonstrate the existence of such
a channel. While biochemical and structural studies have given us some information on how
UreD interacts with urease (8, 23, 24), the structure could directly establish how UreD interacts
with urease apoprotein -- especially related to the nascent active site. To this end, I purified
UreABC:UreD apoprotein from nickel-depleted E. coli cells expressing pKK17, which encodes
the entire K. aerogenes urease gene cluster (ureDABCEFG). I determined that purification of
UreABC:UreD:UreF:UreG by weak anion exchange, strong anion exchange, and size-exclusion
chromatography yielded a fraction containing UreABC:UreD at high purity. The key to this
process is the size-exclusion procedure, as I found that use of a low flow rate (0.3 ml/min) over a
long column (100 ml bed volume) allowed for optimal separation of the various contaminating
protein complexes. I subjected a sample of this complex at 12 mg/ml to crystallization conditions
in the same manner as for MBP-UreDFG, but using only the Wizard™ Classic (1 and 2, or 3 and
4) trays. I observed various morphologies of crystal formation after 2 weeks, as listed in Table
5.1.

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Table 5.1: Crystal morphology and buffer conditions used in production of crystals from
solutions of UreABC:UreD.
Buffer conditionsa

Crystal morphology
Thin/long
Oval/large
Small/cubic
Star-like
Star-like
Thin/long
Large/irregular
Large/cubic
Medium/cubic
Thin/long

Thin/long
Star-like
Thin/long
Cubic/medium
Star-like
Star-like
Star-like
Oval/large

Wizard Classic 1 and 2
35% MPD, 160 mM imidazole, 100 mM HCl,
200 mM MgCl2, pH 8.0
20% PEG 3K, 100 mM sodium citrate, 100
mM citric acid, pH 5.5
20% PEG 3K MME, 100 mM Tris HCl, pH
7.0
20 % PEG 1K, 100 mM imidazole/HCl, 200
mM calcium citrate, pH 8.0
10% PEG 3K, 100 mM H2KPO4/Na2HPO4,
pH 6.2
10% PEG 8K 100 mM imidazole/HCl, 200
mM calcium acetate, pH 8.0
30% PEG 3K, 100 mM Tris base/HCl, 200
mM NaCl, pH 7.0
10% PEG 8K, 100 mM KH2PO4/Na2HPO4,
200 mM NaCl, pH 6.2
1260 mM ammonia acetate, 100 mM
HEPES/NaOH, pH 7.5
10% PEG 3K, 100 mM sodium
cacodylate/HCl, pH 6.5
Wizard Classic 3 and 4
8% PEG 4K, 100 mM sodium acetate/HCl,
200 mM CaCl2, pH 4.6
28% PEG 400, 100 mM HEPES/NaOH, 200
mM CaCl2, pH 7.5
20% PEG100K, 100 mM HEPES/NaOH, pH
7.5
5% PEG 1K, 100 mM Na2HPO4/Citric acid,
pH 4.2
50% PEG 200, 100 mM sodium
cacodylate/HCl, 200 mM MgCl2, pH 6.5
40% PEG 200, 100 mM sodium
cacodylate/HCl, 200 mM calcium acetate, pH
6.5
25% PEG 1500, 100 mM SPG, pH 8.5
30% PEG 2K MME, 150 mM potassium
bromide

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Table 5.1 (cont’d):
10% PEG 2K MME, 100 mM sodium
Cubic/small
acetate/acetic acid, 200 mM ammonium
sulfate, pH 5.5
20% PEG 3350, 100 mM sodium citrate/citric
Cubic/small
acid, 200 mM sodium citrate tribasic, pH 4.0
100 mM Bis-Tris propane/HCl, 200 mM
Star-like
sodium malonate dibasic, pH 8.5
a
Abbreviations used: MPD = 2-Methyl-2,4-pentanediol, PEG = polyethylene glycol, MME =
monomethyl ether, SPG = succinic acid/phosphate/glycine.
While the observation of multiple successful crystallization conditions was an exciting
finding, optimization will be required for two reasons. First, the identity of the proteins within
these crystals needs to be ascertained, since dissociation of UreD could result in crystallization of
free urease. A test for this would be to dissolve a large crystal of putative UreABC:UreD sample
and analyze it by SDS-PAGE. The presence of bands corresponding to UreD and the urease
subunits would confirm the crystal indeed contained UreD docked to urease. Second, the size
and number of crystals generated was insufficient for X-ray scattering studies that are required
for structure determination. Dr. Hu suggested that the buffer conditions could be optimized by
hand, and this could be a future endeavor. Furthermore, it could be helpful to obtain a higher
yield of UreABC:UreD by using a published protocol involving cells with a plasmid expressing
only ureDABC and containing a second copy of ureD (21). My initial efforts with this approach
failed to yield enhanced levels of the complex, but this direction is worth further investigation.
Overall, crystallization efforts have great potential and appear to be poised to yield exciting
information on how the accessory proteins dock to urease.
Native PAGE Analysis of Urease Apoprotein and Its Complexes with MBPUreDFG. I was interested in identifying the relative protein stoichiometries of assembly
complexes between urease and MBP-UreDFG as a precursor to the work presented in Chapter 2.

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As a first step, I prepared mixtures of UreABC apoprotein and MBP-UreDFG in 50 mM Tris, pH
7.4, 1 mM EDTA, and 1 mM β-mercaptoethanol buffer containing 25 mM NaCl. The samples
were incubated for 30 min at room temperature before being analyzed by blue native PAGE on
precast 3-12% Bis-Tris acrylamide gels (GE Healthcare) (Figure 5.6).

Figure 5.6: Blue native PAGE analysis of UreABC and UreABC:MBP-UreDFG complexes.
(A) Mixtures of UreABC apoprotein and MBP-UreDFG were subjected to blue native PAGE
analysis. UreABC (1 µM) was incubated in the absence (lane 1) or the presence of 1, 2, 3, 4 or 5
µM MBP-UreDFG (lanes 2-6). NativeMark protein standards (lane S) were used to estimate the
molecular masses of the complexes observed. (B) Band migration analysis of protein species
observed in A. Band distances and molecular weights were calculated using Alphaview software
on an Alphaimage HP gel scanning apparatus (Protein Simple). Bands W and X are two closely
migrating species.
The two lowest molecular weight bands observed on the gel fit well with the theoretical
masses of MBP-UreDFG and (UreABC)3. No band was observed for (MBP-UreDFG)2, but the
MBP-UreDFG was not examined in isolation as a comparison. It is possible that the complex
exists as a heterotrimer when subjected to the electrophoretic conditions even though ion
mobility mass spectrometry (IM-MS) and size-exclusion chromatography findings presented in
Chapter 2 demonstrate MBP-UreDFG exists in solution as a stable dimer. Four additional bands
were observed when UreABC was incubated with increasing amounts of MBP-UreDFG (bands
W-Z, above). An approximate molecular mass for each band was determined by using band
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migration analysis and gave evidence of an (UreABC)3:MBP-UreDFG complex (band W). This
result is in good agreement with the published findings in Chapter 2 of this work. Assignments
of (UreABC)3:(MBP-UreDFG):UreG, (UreABC)3:(MBP-UreDF)2:(MBP-UreDFG), and
((UreABC)3)2:(MBP-UreDFG)2 for bands X, Y and Z, respectively, fit well with the
experimental molecular weights determined in this study, but these structures are highly
tentative. Although neither complex X or Y was observed in our IM-MS study, the prior work
showed the urease complexes are complex in their compositions. Complex Z was observed in the
earlier study, and I speculate that it derives from two urease trimer-of-trimers bridged by a single
(MBP-UreDFG)2 unit. Finally, I note that while MBP-UreDFG was added in increasing
amounts, only bands W and X increased in intensity, with no apparent increase in free MBPUreDFG or the appearance of a band corresponding to (MBP-UreDFG)2. Overall, this study
provides additional evidence that (MBP-UreDFG)2 interacts with urease as a monomer of
heterotrimers in vitro and provides new information on novel complexes formed between urease
and the soluble accessory protein complex under non-activating conditions.
Stability of MBP-UreDFG Following Tag Removal. Proteolytic removal of MBP from
MBP-UreD was previously shown to result in protein precipitation and an inability to recover
free UreD from anion exchange chromatography (5). In analogous studies, I tested whether free
UreD:UreF:UreG could be liberated from the complex containing the MBP tag and whether the
sample is stable and soluble. Thus, I digested 1.3 mg of MBP-UreDFG with 0.04 mg of factor
Xa in 20 mM Tris buffer, pH 8.0, containing 100 mM NaCl and 2 mM CaCl2. After 3 h at room
temperature, I analyzed the digested samples by SDS-PAGE (Figure 5.7). The results clearly
demonstrate the successful cleavage of the linker between the MBP and UreD domains of MBPUreD. In an extension of this approach, I attempted to separate the digestion products by size-

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exclusion chromatography, but no protein elution peaks were observed at the expected positions
(data not shown). This result could have been a consequence of the low protein concentrations I
used in this experiment, but on the basis of prior studies with UreD (5) and UreD:UreF:UreG
(20) I attribute the protein loss to protein aggregation on the resin. Thus, I conclude that
UreD:UreF:UreG is unstable when liberated from its solubility tag. Future extensions of these
studies could include use of detergent-containing buffers in an effort to retain solubility of the
accessory proteins. The resulting sample might be useful for future GTPase activity studies by
UreG or for crystallization studies of the UreD:UreF:UreG complex.

Figure 5.7: Factor Xa cleavage of MBP-UreDFG. Lane S depicts protein standards.
Samples of MBP-UreDFG were incubated three h at room temperature in calcium-containing
buffer that lacked (lane 1) or contained (lane 2) 3% w/w factor Xa.
Metal-Catalyzed Oxidation of H144*UreE and MBP-UreDFG. Previous studies that
attempted to probe the metal-binding residues of K. aerogenes UreD focused on targeting
residues for mutagenesis, but that process was made difficult by the relatively poor sequence
homology with H. pylori UreH and no side chain ligands were identified (5). In Chapter 3, I
expanded the alignment parameters to include 30 additional UreD homologs, but this effort also
left the identity of metal ligands unclear. Here, I describe my attempts to probe for metalcoordinating residues by applying the metal-catalyzed oxidation (MCO) labeling technique (2).
This method uses Fenton-like chemistry to generate reactive oxygen species that are localized
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near the coordinated metal ions and has been successfully used in conjunction with mass
spectrometry to probe for metal-coordinating sites on angiotensin I, bacitracin A (2), copper-zinc
superoxide dismutase (3), and the prion protein (25). As a proof-of-concept, I chose to use this
technique on the well-characterized nickel-metallochaperone H144*UreE (9). This truncated
protein has two distinct metal-binding sites: one is found at the interfacial site of the homodimer
with the metal coordinated by using His96 from each protomer, and a second site at the
periphery of each protomer that uses His110 and His112 for metal binding (17). Samples of
H144*UreE (100 µM) were incubated for one h at room temperature in 50 mM Tris buffer, pH
7.4, containing 85 mM NaCl or the same buffer supplemented with 10 mM of ascorbate and/or 1
mM of persulfate in a final volume of 20 µl. To test for metal-dependent oxidation, I added 100
µM of NiCl2·6H2O, CuSO4, or (NH4)2Fe(SO4)2·6H2O to samples, treated them with ascorbate
and persulfate, and incubated the mixtures for one h at room temperature. I adjusted each sample
to pH 2.0 with glacial acetic acid and digested overnight at 37 °C with 1 µg of gastric porcine
pepsin. I dried the digests in a speed-vac/SC110A centrifuge (Savant), resuspended them in
water containing 0.1 % trifluoroacetic acid and 2 % LC-MS grade acetonitrile (Sigma), desalted
using C-18 desalting tips (Thermo Scientific), and diluted 1:1 with α-cinnamic acid matrix. I
used 1 µl of these dilutions for analysis by MALDI-TOF MS (Figure 5.8).

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Figure 5.8: Mass spectra of pepsin digests from MCO-treated or untreated H144*UreE.
Pepsin digests are shown for H144*UreE that was incubated in buffer lacking (red trace) or
containing (blue trace) 1 mM ascorbate and 10 mM ammonium persulfate. Also shown for
comparison are digests of protein incubated with the reductant/oxidant supplemented buffer
containing Cu2+ (green trace), Ni2+ (purple trace), or Fe2+ (black trace). Peaks identified by
theoretical digest: 553.05 m/z (residues 44-48), 619.01 m/z (residues 119-123), 666.75 m/z
(residues 43-48, 1 missed cut), 687.57 m/z (residues 131-136), 720.17 m/z (residues 131-136,
two oxidation events), 780.97 m/z (residues 43-49, 2 missed cuts), 982.72 m/z (residues 131-139,
three missed cuts), 1004.96 m/z (residues 131-139, three missed cuts, 1 Na+), 1080.77 m/z
(residues 107-114, 1 missed cut), 1096.43 m/z (residues 107-114, 1 missed cut, 1 oxidation
event), 1286.16 m/z (residues 91-101, 2 missed cuts), and 1309.54 m/z (residues 89-99, 2 missed
cuts).
The above results identified a novel peak (m/z of 1096) that was consistent with oxidation
of the H144*UreE pepsin fragment 107-114, which contains the characterized UreE metalbinding site, following treatment of the protein with reductant/oxidant which is enhanced in
intensity when this treatment includes Cu2+, Ni2+, or Fe2+. The MCO effect is most notable in the
presence of ferrous ions, a highly effective reagent for Fenton chemistry (Figure 5.8, black
trace). Overall, this proof-of-concept study confirmed that redox treatment of the

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metallochaperone UreE in the presence of various divalent metals selectively oxidized a specific
UreE pepsin fragment that is known to contain a well characterized metal binding site.
I extended the MCO approach to studies of MBP-UreD, MBP-UreDFG, and UreG. I
examined both pepsin and trypsin digests, and I varied the ascorbate (0.1, 1 or 10 mM) and
persulfate (0.1, 1 or 10 mM) concentrations. Treated and untreated samples were separated by
SDS-PAGE, the bands corresponding to the individual protein subunits were excised and
digested in-gel by trypsin as described previously (14), or with 1 mg of porcine pepsin in 10 µl
of 100 mM ammonium bicarbonate, pH 2.0. Peptide fragments were extracted and analyzed by
MALDI-TOF MS as described previously (14); however, no novel peaks were noted in any
subunit for any condition tested when comparing metal-treated samples to untreated samples.
The low affinity for nickel seen in the equilibrium dialysis study described in this chapter may
account for the lack of metal-catalyzed oxidation in the MBP-UreDFG sample, but this result
does not account for the absence of an oxidation peak in the MBP-UreD and UreG samples, as
each protein has been shown to have relatively high affinity for nickel (1, 6).
Negative Stain Electron Microscopy of Urease:Accessory Protein Complexes. One of
the biggest remaining questions regarding urease activation is where the accessory proteins dock
to the enzyme. Previous small angle X-ray scattering and chemical cross-linking/mass
spectrometry studies provided evidence that UreD interacts with both UreB and UreC, where the
interaction was localized near the vertices of (UreABC) 3 (8, 23). Further structural studies on
accessory protein-bound urease complexes were needed to solidify this docking assignment. To
this end, I began a collaboration with Dr. Kristin Parent (Michigan State University) to study
urease and urease:accessory protein complexes by negative stain electron microscopy. The small
size of the urease complexes precluded the use of the more sensitive cryo-electron microscopy,
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but I hypothesized that by comparing ensemble images of accessory-protein unbound and bound
samples of urease we could determine more accurately how UreD was oriented on the enzyme.
Furthermore, comparison to UreABC:UreD:UreF:UreG could provide us with information on
how the distal accessory proteins orient themselves when docked to the enzyme, which would
provide crucial information regarding whether the accessory proteins adopt a different
conformation relative to that seen in the (UreH:UreF:UreG)2 structure from H. pylori (16). To
this end, I spotted 3 µl of a 0.2 mg/ml of UreABC, UreABC:UreD, UreABC:UreD:UreF or
UreABC:MBP-UreD:UreF:UreG complexes onto a carbon grid and allowed the samples to dry.
Excess protein not adsorbed to the grid was washed away with deionized water and the gridded
protein was stained with 0.5% phosphotungstic acid. This stain is atypical in its use in modern
electron microscopy, as it tends to give a dark, intense background. However, the more standard
stains were found to cause decomposition of the protein complexes. Grids were scanned on a
JEOL 2200-FS electron microscope. An ensemble image of particles resembling the expected
triangular shape of urease was prepared and compared to the crystal structure (Figure 5.9).
Overall, the averaged ensemble image overlays well with the crystal structure, providing us with
a baseline with which to compare accessory protein-docked urease complexes.

Figure 5.9: General shape comparison of the structures of K. aerogenes urease obtained by
X-ray crystallography (left (18)) and negative-stain electron microscopy (right).
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Unfortunately, similar studies on samples of (UreABC)3 docked with UreD, MBPUreD:UreF:UreG, or UreD:UreF yielded grid images that lacked particle homogeneity. In
general, these complexes presented a large amount of background particles that were smaller in
size compared to urease itself. The most peculiar finding was an absence of urease-like particles
in most grids scanned. This result may be explained by a loss of stability of (UreABC) 3 when
accessory protein are docked and in the presence of the dye; however, the UreABC:MBPUreD:UreF:UreG complex was stable under standard conditions by IM-MS (14). Several
experimental adjustments can be used in attempts to rectify these problems. For example, the
sample could be stabilized for accessory protein-docked urease by chemical cross-linking.
Samples of UreABC:UreD or complexes with additional accessory proteins could be treated with
the various cross-linkers as previously described (8), separated by Blue-Native PAGE, and bands
corresponding to (UreABC)3:(accessory protein)x could be extracted, purified and analyzed by
negative-stain electron microscopy as described above. I hypothesize the Blue native PAGE
would enrich for specific accessory protein-docked urease species, and the cross-linking would
help with stabilization of the enzyme.
ADDITIONAL UreA2B2 STUDIES
Additional Spectroscopic Studies of UreA2B2. While ample evidence has been
presented supporting a diferrous active site in active UreA2B2 and a µ-oxo bridged, diferric
center in oxidized UreA2B2, both in this dissertation and in founding studies on the enzyme (7),
it was unclear whether a mixed-valent (Fe(II)/Fe(III)) state was accessible. To this end, I
collaborated with Dr. Roman Davydov (Northwestern University) to attempt the generation of
mixed-valent UreA2B2 by radiolytic reduction under low temperature (frozen) conditions.
Significantly, electron paramagnetic resonance (EPR) spectra of mixed-valent species can
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discern between a µ-oxo and µ-hydroxo bridge in the starting diferric centers of enzymes or
synthetic inorganic complexes (10, 11, 12, 13). For example, mixed-valent di-iron centers
generated by radiolytic reduction of µ-oxo bridged diferric enzymes yield EPR spectra that
possess low anisotropy (Δg ~ 0.1) and exhibit an average g-value near 1.9 at 77 K, whereas
samples that are warmed (annealed) briefly following cryoreduction either exhibit EPR spectra
with no change in these values or they have increased anisotropy and a decrease in average gvalue (12). Most tellingly, similar studies with µ-hydroxo bridged species lead to spectra
exhibiting higher anisotropy (Δg ~ 0.2) and a lower average g-value (~ 1.8) at 77 K, with no
change in these values after annealing.
Using samples that I provided, Dr. Davydov observed that radiolytic cryoreduction of
UreA2B2 produced small quantities of the mixed-valent state that exhibited EPR spectral
features at 77 K indicative of a µ-oxo bridged diferric state for the starting sample, with g-values
of 1.945, 1.9 and 1.84 (Figure 5.10). These values did not change after annealing at temperatures
of 155 and 173 K. Overall, this study gives further evidence that UreA2B2 possesses a µ-oxo
bridged, diferric center in its oxidized state.

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Figure 5.10: EPR studies of UreA2B2 subjected to radiolytic reduction. Samples of
UreA2B2 were reduced by γ-radiation at 77 K and analyzed by EPR spectroscopy at 77 K (green
trace) without additional treatment or after brief annealing at 155 K (red trace) or 173 K (blue
trace). The feature noted by (*) at 77 K is a hydrogen-specific feature generated from irradiation
of the sample holder and does not originate from sample. This figure was generously provided by
Dr. Davydov (Northwestern University).
I also continued a collaboration initiated by Dr. Eric Carter with the laboratory of Dr.
Edward Solomon (Stanford University) to study the dithionite-reduced state of UreA2B2 by
circular dichroism (CD), magnetic circular dichroism (MCD), and variable-temperature,
variable-field magnetic circular dichroism (VTVH MCD) spectroscopic methods. The diferrous
state was assumed to possess a µ-hydroxo bridging mode, mirroring the situation that is thought
to exist in the nickel enzyme. This weakly electronically coupled site should be poorly
observable by MCD spectroscopy. As background, prior EPR studies of the fully reduced,
anaerobic form of UreA2B2 showed it to be EPR-silent when using the conventional
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perpendicular mode instrument as expected for two ferrous ions. Parallel mode EPR is capable of
showing a signal near g ~ 16 arising from a coupled diferrous center, but such studies were not
conducted.
Our initial MCD studies of reduced UreA2B2 highlighted strong MCD features at lowtemperatures, with these features decreasing as temperature increased (Figure 5.11). These
results provide evidence against a µ-hydroxo bridge in the fully reduced state and could suggest
a bridging water molecule. Preliminary pH and deuterium-sensitivity studies were performed by
analyzing reduced UreA2B2 at buffer pDs of 7.4, 8.4, and 9.4. The results (not illustrated)
showed no change in the observed features by either CD or MCD spectroscopy. A µ-hydroxo
bridged site would be expected to show sensitivity to bulk deuterium, thus giving further
evidence against a (Fe(II)-OH-Fe(II)) site. Rather, these findings also support a µ-aqua bridge
which would result in a loss of coupling between the ferrous ions. Preliminary VTVH MCD
studies were carried out (not shown) and appear to identify these same strong features associated
with weak J-coupling. Therefore, these studies provide evidence that the resting-state of
UreA2B2 is that of a diferrous, µ-aqua bridged active site.

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Figure 5.11: Low temperature MCD spectra of anaerobic, dithionite reduced UreA2B2.
Changes in molar absorptivity were observed at 2.2 K (red trace), 5 K (blue trace), and 15 K
(green trace). This figure was prepared and generously provided by Lars Boettger from the lab of
Edward Solomon (Stanford University).
The effect of the nickel urease inhibitor boric acid (4) on the CD and MCD spectra of
UreA2B2 was examined to gain further insights into the electronic coupling of the two metals at
the metallocenter with a bound substrate analog. The preliminary results show differences
between reduced UreA2B2 with or without boric acid (not shown), but additional studies need to
be carried out. I’ve provided additional samples to the Solomon group and further analyses are
planned in the near future.
Boric Acid Inhibition of UreA2B2. Although boric acid inhibition of nickel urease has
been kinetically analyzed (4), the kinetic inhibition properties of this compound with the iron
enzyme were unknown. Our collaboration with Dr. Solomon to investigate the electronic
properties of the borate-inhibited enzyme necessitated kinetic characterization of the inhibitor
mechanism and thermodynamics. To this end, I assayed for the mode and kinetics of inhibition
for boric acid under anaerobic reducing conditions. UreA2B2 in 50 mM HEPES, 1 mM EDTA,
pH 7.8, buffer was made anaerobic by injecting it into a 10 ml argon-filled vial using an argon136

purged gas-tight syringe (Hamilton) and allowing gas exchange to occur overnight on ice.
Powders of ferrous sulfate and boric acid were made anaerobic with 3 vacuum/argon purge
cycles at 2 min per step using a Schlenk line. A 50 mM HEPES, 150 mM NaCl, pH 7.8, assay
buffer was made anaerobic with 5 vacuum/argon purge cycles at 10 min per step. Enzyme stop
solution containing 5% phenol and 0.025% sodium nitroprusside was made anaerobic in a
similar manner. All samples were transferred into a 100% N2 chamber maintained at < 4 ppm O2.
Reaction tubes containing 0, 0.6, 2, 6, 20 or 60 mM boric acid and 2, 6, 20, 60, 200 or 600 mM
urea were prepared in 50 mM HEPES, 150 mM NaCl, pH 7.8, buffer. UreA2B2 was diluted to
0.8 µg/ml and reduced with 1 mM of ferrous sulfate over 2 h at room temperature. Following this
process, standard urease assay conditions were performed (27), but using anaerobic conditions.
A plot of the specific activities versus substrate concentrations is depicted in Figure 5.12. The
calculated Kmapp and Vmaxapp values are displayed in Table 5.3.

137

Figure 5.12: Kinetic analysis for boric acid inhibition of UreA2B2. Urease assays were
performed anaerobically after prior reduction of the enzyme in solutions lacking (open circles,
red trace) or containing boric acid at 0.6 (closed circles, blue trace), 2 (open square, green trace),
6 (closed square, magenta trace), 20 (open triangle, black line) or 60 mM (closed triangle)
concentrations and the substrate concentrations indicated.
Table 5.3: Apparent Km and Vmax for UreA2B2 in various concentrations of boric acid.
[Boric acid] (mM) Kmapp (mM) Vmaxapp (µmol urea degraded ∙·min-1·∙ mg protein-1)
0

3.3

23.2

0.6

4.4

21.22

2

6.98

20.13

6

27.09

25.11

20

103.2

37.8

60

197.4

36

138

The data presented here represent a single study, so caution must be exercised when
attempting to interpret the results. Nevertheless, the results are generally consistent with
competitive inhibition (Ki ~ 1.23 ± 0.25 mM; the increased activity at the largest urea
concentrations for 20 and 60 mM borate may be aberrant) which is an order of magnitude
smaller than for the nickel enzyme from K. aerogenes (26). Also of interest, these reduction and
reaction conditions yielded greater UreA2B2 activities than what was published previously (2
U/mg activity) (7). This result likely is due to the alternate reductant that I used, however, it is
also possible that the enzyme sample utilized in this study differed from that used previously
(e.g., see the evidence for two enzyme forms described in Chapter 4).
Resonance Raman Spectroscopy on UreA2B2 with Increasing Amounts of Urea. In
order to determine the necessary amount of urea to add to UreA2B2 to generate a complete
downshift in the νs(Fe(III)-O-Fe(III)) mode, resonance Raman spectra were obtained from
samples of UreA2B2 incubated briefly (<2 min) with increasing concentrations of urea prior to
analysis (Figure 5.13). The spectra showed a gradual shift toward a nearly complete downshift of
the mode with increasing concentrations. My studies point to this shift being caused by urea
binding at the metallocenter of UreA2B2, so this can be used as a rough assessment for the Kd of
urea for oxidized UreA2B2. From this study, an estimated Kd between 10 and 30 mM urea was
estimated. However, we later learned around 10 minutes of incubation is required for a complete
downshift of the νs(Fe(III)-O-Fe(III)) mode at 2 mM urea. Thus, this study would have to be
repeated with a longer incubation time to ensure binding has reached completion. Overall, this
technique presents a novel way for determining the binding constants of urea for oxidized
UreA2B2.

139

Figure 5.13: Resonance Raman spectra for samples of UreA2B2 incubated with
varied concentrations of urea. 285 µM UreA2B2 was incubated with 0 (red trace), 0.1 (blue),
0.3 (green), 1 (yellow), 3 (black), 10 (red, dotted), 30 (blue, dotted), or 100 mM (green, dotted)
for less than 2 minutes at room temperature before being analyzed by resonance Raman
spectroscopy.

140

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2.

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4.

Breitenbach, J. M., and Hausinger, R. P. 1988. Proteus mirabilis urease. Partial
purification and inhibition by boric acid and boronic acid. Biochem. J. 250:917-920.

5.

Carter, E. L. 2012. Investigations into urease maturation and metal ion selectivity.
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Carter, E. L., and Hausinger, R. P. 2010. Characterization of the Klebsiella aerogenes
urease accessory protein UreD in fusion with the maltose binding protein. J. Bacteriol.
192: 2294-2304.

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Carter, E. L., Proshlyakov, D. A., and Hausinger, R. P. 2012. Apoprotein isolation
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Chang, 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.

9.

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.

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the tyrosyl radical on the reduction and structure of the Escherichia coli ribonucleotide
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Dewitt, J. G., Bentsen, J. G., Rosenzweig, A. C., Hedman, B., Green, J., Pilkington,
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interactions with Klebsiella aerogenes urease by mass spectrometry. J. Am. Soc. Mass.
Spectrom. 24: 1328-1337.

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2011. Assembly of preactivation complex for urease maturation in Helicobacter pylori:
crystal structure of UreF-UreH protein complex. J. Biol. Chem. 286: 43241-43249.

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Fong, Y. H., Wong, H. C., Yuen, M. H., Lau, P. H., Chen, Y. W., and Wong, K. B.
2013. Structure of UreG/UreF/UreH complex reveals how urease accessory proteins
facilitate maturation of Helicobacter pylori urease. PLoS Biol. 11: e1001678.

17.

Grossoehme, N. E., Mulrooney, S. B., Hausinger, R. P., and Wilcox, D. E. 2007.
Thermodynamics of Ni2+, Cu2+, and Zn2+ binding to the urease metallochaperone UreE.
Biochemistry 46: 10506-10516.

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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.

19.

Merloni, A., Dobrovolska, O., Zambelli, B., Agostini, F., Bazzani, M., Musiani, F.,
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proteins UreE and UreG. Biochim. Biophys. Acta 1844: 1662-1674.

20.

Moncrief, M. B., 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.

21.

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.

22.

Park, I. S., and Hausinger, R. P. 1995. Evidence for the presence of urease apoprotein
complexes containing UreD, UreF, and UreG in cells that are competent for in vivo
enzyme activation. J. Bacteriol. 177: 1947-1951.

143

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Quiroz-Valenzuela, S., Sukuru, S. C., 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.

24.

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461-475.

25.

Srikanth, R., Wilson, J., Burns, C. S., and Vachet, R. W. 2008. Identification of the
copper(II) coordinating residues in the prion protein by metal-catalyzed oxidation mass
spectrometry: evidence for multiple isomers at low copper(II) loadings. Biochemistry 47:
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26.

Todd, M. J., and Hausinger, R. P. 1989. Competitive inhibitors of Klebsiella aerogenes
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144

CHAPTER 6

Concluding remarks

145

CONCLUDING REMARKS
My dissertation provides insights into the interactions of the accessory proteins with
urease, the role of UreD in the activation of urease, the mechanism of urease activation, and the
spectroscopic properties of the oxidized UreA2B2 metallocenter. These studies pave the way for
additional investigations by other researchers. Below, I summarize my results in greater detail
and describe potential future studies that could be performed.
My analyses of the (MBP-UreDFG)2 complex and its interaction with urease are
described in Chapters 2 and 5. This work detailed the purification and characterization of a
soluble dimer of heterotrimeric urease accessory protein complex from Klebsiella aerogenes and
identified the GTP-dependent conditions that result in the dissociation of UreG. Of great
importance, ion mobility mass spectrometry was used to demonstrate the complex dissociates to
a monomer of heterotrimers on binding to urease. At this time, no studies have identified
whether dimerization of the heterotrimer is functionally important. Prior studies on Helicobacter
pylori (UreH:UreF:UreG)2 had identified Tyr183 as a residue required for the formation of the
dimer of heterotrimers (3), and in that case urease activation was hindered with the variant
protein. Future efforts could create an MBP-UreD:UreF(Y155D):UreG complex (where Tyr155
is homologous to Tyr184 in HpUreF), and this purified species could be used to characterize its
ability to form a dimer of heterotrimers, test if it activates urease in vivo, and ascertain whether
stability is affected in the presence of GTP/Mg2+/Ni2+. My mutagenesis studies to substitute
individual surface residues of UreD, described in Chapter 3, did not prove to be useful for
mapping interactions surfaces, but this issue remains a point of interest. Additional future studies
can use the soluble accessory protein complex with wild-type components along with urease to
explore the urease:accessory protein interfacial site. Prior experiments identified cross-links
146

between UreB:UreD and UreD:UreC (2), and small angle X-ray scattering studies provided
evidence that UreD binds at the vertices of the triangular (UreABC) 3 (4). I propose the use of
hydrogen/deuterium exchange mass spectrometric studies for characterizing the exposed surfaces
of UreABC apoprotein, MBP-UreD, MBP-UreDFG, UreABC:MBP-UreD, and UreABC:MBPUreDFG. A reduction in deuterium exchange into specific residues of urease or MBP-UreD for
the complexes versus the non-complexed proteins would identify the potential binding site,
which could be further characterized by mutagenesis studies.
Chapter 3 defines a clear nickel-transfer role for UreD in the activation of urease and
provides evidence for a buried water tunnel being used for this purpose. While the evidence
presented in this chapter is strong, it does not address the identity of any nickel-coordinating
residues. To this end, one could compare the nickel-binding properties of wild-type and variant
forms of MBP-UreD (specifically the UreD variants that affect urease activation) by X-ray
absorption spectroscopy for changes in coordination number or ligating residues. Variants found
to have changes could be further studied to characterize nickel binding affinity and the number
of bound nickels per MBP-UreD via equilibrium dialysis or isothermal titration calorimetry. The
effects of UreD substitutions on urease activation in vitro should also be investigated to
determine if these changes affect the rate of activation or increase the susceptibility toward
contaminating metals. Finally, the feasibility of the water tunnel functioning in cation transfer is
not fully addressed in Chapter 3. Ongoing studies by our collaborators in the Feig group are
performing in silico molecular dynamics studies of the wild-type UreD homology model and the
variant proteins. Additional analysis could examine the energetics of a transfer tunnel using the
optimized UreD homology model to estimate whether nickel transfer is energetically favored and
which portions of the protein lead to energetic barriers. I recognize the hazards with carrying out

147

such simulations on a homology model, but at this time we don’t have a crystal structure
coordinates of the protein.
The studies on UreA2B2 in Chapters 4 and 5 provide more information on the oxidized
site of iron urease, but several aspects of the metallocenter were not fully addressed. The
assignment of a terminal Fe-OH stretch for the D2O/H218O-sensitive feature at ~530 cm-1 could
be strengthened by repeating these studies at varied pH, based on the assumption that low pH
studies may result in the loss of this feature due to protonation. The issue of protein precipitation
was noted in Chapter 4 at lower (6.4) and higher (9.4) pH, but this problem can be circumvented
by using rapid pH changes with concentrated buffers at the desired final pH. The observation that
phenyl phosphorodiamidate (PPD), but not acetohydroxamic acid, causes a downshift in the 500
cm-1 feature gives evidence that PPD coordinates to the diferric site of UreA2B2 in a manner
similar to urea, but this interpretation needs to be further explored. Rapid bulk-solvent exchange
of 16O- or 18O-bridged UreA2B2 can be undertaken in the absence or presence of PPD or urea
and analyzed by resonance Raman spectroscopy to determine if the observed isotopic-solvent
sensitivities are similar between the molecules or if they differ. A loss of the terminal solvent
mode would imply the binding of PPD in oxidized UreA2B2 adopts a similar orientation to that
of PPD shown in the Sporosarcina pasteurii structure (1), while it also gives evidence that ureabinding does not result in the displacement of terminal waters at both irons of the metallocenter.
In total, my results have led to significant advances in understanding of both the
conventional nickel urease activation machinery and the spectroscopic properties of iron urease.
Nevertheless, many questions about these systems remain to be answered and investigations to
extend our understanding of urease and its activation remain an exciting area for future research.

148

BIBLIOGRAPHY

149

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Benini, S., Rypniewski, W. R., Wilson, K. S., Miletti, S., Ciurli, S., and Mangani, S.
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2.

Chang, Z., Kuchar, J., and Hausinger, R. P. 2004. Chemical cross-linking and mass
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Chem. 279: 15305-15313.

3.

Fong, Y. H., Wong, H. C., Yuen, M. H., Lau, P. H., Chen, Y. W., and Wong, K. B.
2013. Structure of UreG/UreF/UreH complex reveals how urease accessory proteins
facilitate maturation of Helicobacter pylori urease. PLoS. Biol. 11: e1001678.

4.

Quiroz-Valenzuela, S., Sukuru, S. C., Hausinger, R. P., Kuhn, L. A., and Heller, W.
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mutagenesis, and small-angle X-ray scattering. Arch. Biochem. Biophys. 480: 51-57.

150

APPENDIX

151

Table A.1: Plasmids used in this dissertation.

Plasmid used
pUC8
pKK17
pKK17D*, -V37L, -Y42D,
-E46A, -E46Q, -C48A, H49A, -H54A, -I59Y, D63A, -D63Q, -L65I, L65W, -S85K, -K86A, Y88V, -Y88F, -R89A, W111Y, -T128E, -D142A,
-R148M, -E153A, -E153Q,
-R163A, -E165A, -D169A,
-E176A, -E176Q, -T196K,
-R211A, -R233A
pKKG
pKKD*G, -D63A, -D63Q.
–S85K, -D142A, -E176A, E176Q, R211A
pEC002
pEC005
pEC005-UreF-K165A
pMF001
pMF001L
pEC015

Description
High-copy number, pBR322-derived vector
conferring AmpR
Wild-type K. aerogenes urease cluster
(ureDABCEFG) placed into pKK223-3

EcoRI-AgeI 5’UTR-ureD fragment from
pMF001L* ligated into similarly digested
pKK17

PstI-KpnI ureGStr fragment ligated into
similarly digested pKK17 resulting in
replacement of UreG with one containing a Cterminal Strep-tag II (ureDABCEFGStr)
EcoRI-AgeI 5’UTR-ureD fragment from
pMF001L* ligated into similarly digested
pKKG
pMAL-c2X derived vector for the
overproduction of maltose binding protein
fused at the N-terminus of UreD
KpnI-XbaI ureFG fragment cloned into
similarly digested pACT3
Single-site mutation of pEC005 that encodes
the K165A variant of UreF along with UreG
EcoRI-HindIII ureD fragment from pEC002
ligated into similarly digested pUC8
EcoRI-AgeI 5’UTR-ureD fragment from
pKK17 ligated into similarly digested pMF001
BamHI-PstI ureA2B2 fragment cloned into
similarly digested pEXT20

152

Source
(6)
(5)

This work

(2)

This work
(3)
(3)
(1)
This work
This work
(4)

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