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TA BEAR

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3 1293 01021 9768

 

This is to certify that the
dissertation entitled

MECHANISTIC STUDIES OF Klebsiella aerogenes UREASE
AND ITS NICKEL METALLOCENTER ASSEMBLY PROCESS

 

presented by

IL-SEON PARK

has been accepted towards fulfillment
of the requirements for

 

Ph. D degree in my

.’/ If / 71’; / \
,’ . .r’ / (fl
" /(Vf/ , 7 m/zuuuvfrb‘l
V _ ,1

ROBERT P. HASINGER

Major professor

 

Date—mg...—

MSUi: an Affirmative Action/Equal Opportunity Institution 0-12771

 

LIBRARY
Michigan State
Universlty

 

 

 

PLACE ll RETURN BOXtonmavothb checkout“! yum
TO AVOID FINES Mum on or baton data duo.

DATE DUE DATE DUE DATE DUE

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

MSU loAn mmmw Opponmuy lthon
W1

MECHANISTIC STUDIES OF Klebsiella aerogenes UREASE AND
ITS NICKEL METALLOCENTER ASSEMBLY PROCESS

BY

lL-SEON PARK

A DISSERTATION

Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of
DOCTOR OF PHILOSOPHY

Department of Microbiology

1994

ABSTRACT

MECHANISTIC STUDIES OF Klebsiella aerogenes UREASE AND
ITS NICKEL METALLOCENTER ASSEMBLY PROCESS

By
lL-SEON PARK

Chemical modification and site-directed mutagenesis methods were used to
examine the mechanism of Klebsiella aerogenes urease. The enzyme is
inactivated by diethylpyrocarbonate (DEP) in a manner consistent with the
presence of at least one essential histidine per catalytic unit. Each of ten
conserved histidine residues (H96 in the 7 subunit, H39 and H41 in B, and
H134, H136, H219, H246, H312, H320 and H321 in the a subunit) was
substituted with alanine. The yH96A, BH39A, BH41A, aH312A and aH321A
mutant proteins possess activities and nickel contents similar to wild-type.
The aH134A, aH136A and aH246A proteins exhibit no detectable activity
and possess 550% of the nickel content of wild-type enzyme. The aH219A
protein is active and has nickel (~1.9% and ~80%, respectively, when
compared to wild-type protein), but exhibits a very high Km value (1100:40

mM compared to 23:02 mM for the wild-type enzyme). Finally, the aH320A
protein (Km=8.3:0.2 mM) only displays ~0.003% of the wild-type enzyme
activity, and has a normal nickel content. This mutant protein is not
inactivated by DEP. These results are consistent with aH134, aH136 and
aH246 functioning as nickel ligands, aH219 having some role in facilitating
substrate binding, and aH320 being the DEF-reactive general base that
facilitates catalysis. I

In vivo assembly of the urease metallocenter is known to require four
accessory proteins: UreD, UreE, UreF and UreG. In vitro activation of purified

K. aerogenes urease apoprotein has now been accomplished by simply
providing CO; (half-maximal activation at ~0.2% C02) in addition to nickel
ion. Activation requires that CO; be incorporated into urease in a pH-
dependent reaction “K. 2 9). I propose that CO; binding to urease
apoprotein generates a ligand that facilitates productive nickel binding. Site-
directed mutagenesis methods were used to overexpress ureD in the
presence of the other urease genes, and UreD was found to co-purify with
urease. The UreD-urease apoprotein complexes are activated in a C02-
dependent manner. And, as for urease apoprotein, the activation is
accompanied by non-productive nickel-binding. The presence of UreD,

however, appears to reduce the rate of this inactivation step.

To my family,
with respect and love.

ACKNOWLEDGMENTS

First of all, I would like to thank Dr. Hausinger for his intelectual guidance,
encouragenment and care with so much patience. I would also like to thank my
commettee members: Dr. Tom Deits, Dr. Lee Kroos, Dr. John Breznak, and Dr.
Mickel Bagdasarian. In addition, I would like to thank my collaborators: Mathew
Todd, Mann-hyung Lee, Rick Martin, Fumiyasu Fukumori, Vladimir Romanov,
Mary Beth Carr, Ruth Walace, Tim, Taha, and Linda. I thank the Microbiology
department, its office members and Dr. Ron Patterson for guiding me to
complete my degree.

Special thanks also go to Dr. Sang-Dai Park for being my first professor
and letting me have firm basis as a biologist. I am also indebted to Dr. Kyung-Ja
Ryu for her care and encouragement, and Dr. Yoo-Hun Suh for teaching me
molecular biology.

Finally, I thank my parents who always give me all support they can do at
any cost and my brothers and relatives for their encouragement and support as
always.

TABLE OF CONTENTS

 

 

 

 

 

 

 

 

 

 

 

 

 

 

List of Tables“ Vii
List of Figures Viii
Chapter 1 Introduction 1
Medical significance """""" 3
Agricultural significance 4
Urease structural properties 5
Urease active site 6
Urease hydrolysis mechanism 8
In vivo activation of urease apoprotein 9
Accessory genes for urease activation 10
In vitro activation of urease apoprotein and the probable role of
UreD 12
Eukaryotic ureases 12
Outline of this thesis 14
References 16

Chapter 2. Diethylpyrocarbonate reactivity of Klebsiella aerogenes urease:
effect of pH and active site ligands on the rate of inactivation "1°25

 

 

 

 

 

 

 

Abstract 26
Introduction 27
Materials and methods 28
Materials 28
Enzyme purification 28
Kinetics of urease inactivation 28
Spectroscopy 28
Recovery of enzyme activity by NH20H treatment """"""""""" 29

 

Results and Discussion 29

Chapter 3.

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Inactivation of urease by DEP 29
Effect of ligands on the rate of urease inactivation by
DEF 32
Effect of pH on the rate of urease inactivation 34
Reactivation of DEF-modified urease 37
Spectrophotometric analysis of urease reactivity with
DEF 37
References 42
Site-directed mutagenesis of Klebsiella aerogenes urease:
Identification of histidine residues that appear to function in
nickel ligation, substrate binding, and catalysis 43
Abstract 44
Introduction 45 ‘
Materials and methods 47
Materials 47
Site-directed mutagenesis 47
Enzyme purification 48
Assay of enzyme activity 49
Nickel quantitation 49
Thermal stability 49
Inactivation by DEF 50
Inactivation by IAM 50
Results 50
Initial characterization of mutant proteins 50
Thermal stabilities of mutant ureases 54
pH dependence of mutant urease activities 54
Inactivation of mutant ureases by DEF 57
Inactivation of mutant proteins by IAM 60

iv

Chapter 4.

Chapter 5.

Chapter 6.

 

 

 

 

 

 

 

 

 

 

 

 

Discussion 63
References 65
In vitro activation of urease apoprotein and role of UreD as a
chaperone required for nickel metallocenter assembly """""""""" 69
Abstract 70
Introduction 71
Materials and methods 72
Site-directed mutagenesis of the ureD ribosome binding
site 72
Cell growth and disruption 73
Purification of UreD-urease apoprotein complex """""""""""" 73
Polyacrylamide gel electrophoresis 74
Urease activity assays 74
Results 74
In vitro activation of urease apoprotein 74
Purification of UreD-urease apoprotein complex """""""""""" 75
Activation of UreD-urease apoprotein complexes """""""""" 81
Discussion 83
References 87

 

Carbon dioxide is required for in vitro assembly of the urease nickel

 

 

 

 

metallocenter I 89
Abstract 90
References and notes 98
Related studies, and prospects for further research 100

Characterization of aHis-219 and aHis-312 mutant proteins """"" 101

V

 

Function of UreD in urease apoprotein activation 104

 

 

 

 

 

Characterization of UreD 108
Possible role of UreF 114
Presence of a complex containing UreD, urease apoprotein, UreF,
and UreG 1 17
Urease apoprotein activation: possible scheme 121
References 125
126

 

Appendix

vi

LIST OF TABLES

 

Chapter 3.
1. Characteristics of wild-type and mutant Klebsiella aerogenes
ureases 53
2. Thermal stabilites of wild-type and mutant ureases """"""""""""""" 55
Chapter 6.

1. Characteristics of wild-type and selected mutant Klebsiella

 

aerogenes ureases 102

vii

Chapter 2.

Chapter 3.

Chapter 4.

51.0.01?

LIST OF FIGURES

. Pseudo-first—order urease activity loss in the presence of

 

DEF 30
Effect of DEP concentration on pseudo-first-order rate of

 

inactivation 31

Effect of competitive inhibitors on the rate of urease inactivation

 

 

by DEF 33
Effect of urea on the rate of urease inactivation by DEP """""""" 35
The pH dependence of DEP inactivation of urease 36

Reactivation of DEF-inactivated urease by hydroxylamine """"" 39

Spectroscopic analysis of urease modification by

 

 

DEF 40
Kinetic comparison of activity and absorbance changes during
modification by DEF 41

. Denaturing gel electrophoretic analysis of partially purified

 

 

 

urease proteins 52
The pH dependence of urease activity 56
Kinetics of urease inactivation by DEP 58

The pH dependence of DEP inactivation of wild-type and
aH219A ureases 59

Kinetics of urease inactivation by IAM o1

 

The pH dependence of IAM inactivation of wild-type and

 

aH219A ureases 62

viii

Chapter 5.

Chapter 6.

 

. In vitro activation kinetics of urease apoprotein 76
. SDS-polyacrylamide gel electrophoretic analysis of UreD and

 

urease fractions 77

. Phenyl-Superose resolution of UreD-urease complexes """""""" 79
. Native polyacrylamide gel electrophoretic analysis of urease

 

fractions 80

 

. Activation kinetics of the UreD-urease complexes 82

6. Western blot comparison of cell extracts before and after nickel

 

 

 

activation 84

. Simplified model illustrating the functional role for UreD in
urease apoprotein activation 85
1. In vitro activation of urease apoprotein 94
. In vitro activation of UreD-urease apoprotein complex """"""""""" 96

. The pH dependence of wild-type and mutant urease activity of

 

cell extracts 103

. In vitro activation of the three partially purified UreD-urease

 

apoprotein complexes 106

. Inactivation of the three UreD-urease apoprotein complexes by

 

 

incubation with nickel prior addition of NaHCOa 109
. Initial activation rates of UreD-urease apoprotein

complexes 1 10
. Characterization of UreD 112

 

. Site-Directed Mutagenesis of the ureF Ribosome Binding

 

Site 115

. Western blot comparison of fractions from Sepharose 6 column

 

chromatography of selected cell extracts 116

ix

 

8. Western blot comparison of cell extracts 118

9. Partial purification and characterization of the urease-containing

 

8 complexes 1 19
10. Activation of urease apoprotein associated with the B

 

complexes 122

CHAPTER 1

INTRODUCTION

2

A large amount of urea is biologically produced. For example, urea is a
product of biodegradation of nitrogenous compounds such as purines,
arginine, agmatine, allantoin, and allantoic acid (1). Mammals excrete urea in
urine as a detoxification product (2). Although urea is very stable in solution
(its half-life is 3.6 years in aqueous solutions at 38°C (3)), it is rapidly
hydrolyzed to ammonia and carbamate in the environment through the action
of the enzyme urease (urea amidohydrolase, EC 3.5.1.5) (4). Carbamate
spontaneously decomposes to form a second molecule of ammonia and
carbon dioxide which hydrates to carbonic acid. At normal physiological pH,
carbonic acid dissociates into a proton and bicarbonate and ammonia is
protonated to yield ammonium ion. Since two moles of ammonium ion and
one mole of bicarbonate are produced from the decomposition of one mole of

urea, the hydrolysis results in a pH increase (Scheme 1).

I"'20 H O
NHZCONHZ :9 NHZCOOH v4 co2 \__>\ H2003 # Hco3
Urease 4' + 4’
NH3 NH3 H”
2NH3+2H* Eéié 2NH:
Scheme 1

Ureases have been found in a wide range of bacteria, several fungi, a
few invertebrates, and a variety of plants (reviewed by Mobley and
Hausinger, 5). Until recently, the best-studied urease was that from jack
bean. This urease was the first enzyme to be crystallized (6) and also the

first shown to contain nickel (7). More recently, urease studies have focused

3

on the bacterial enzymes because of the ease of their genetic manipulation
and due to their medical importance. Microbial ureases are important in
human health, rumen ecology, and soil nitrogen management. This chapter
will present brief overviews of the significance, enzymology and
metallocenter biosynthesis of ureases. I will focus this introduction on the
ureases of bacteria, although examples from plant and other sources will be

mentioned where appropriate.

Medical significance. Pathogenic ureolytic microorganisms have
been isolated from humans and other animals where they can colonize the
urinary tract, the colon, or the stomach.

Human urine consists of 0.4 to 0.5 M urea (8) and the urinary tract
provides a favorable environment for many ureolytic microorganisms. The
hydrolysis of urea by urease from these organisms causes a rise in urine pH
leading to supersaturation and precipitation of magnesium and calcium salts
(8) to form stones. Approximately 15-20% of all kidney stones are associated
with infections by ureolytic microorganisms (9). Proteus mirabilis is the major
urease-producing uropathogen in humans (10). Other urease-producing
species associated with infection stones have been recognized, including
Pseudomonas, Klebsiella, and Staphylococcus spp. (9). Urease has also
been shown to be a virulence factor in urinary tract infections that result in
pyelonephritis, an inflammation of the kidney (reviewed in 5).

Ureolytic microorganisms in the colon or the urinary tract can produce
a large amount of ammonia, a toxic compound (11). Under normal conditions,
the ammonia can be removed by the liver. However, when the amount of
ammonia produced is too excessive or the liver is damaged, the elevated
level can cause hyperammonemia (12), ammonia encephalopathy (13), and

hepatic coma (14).

4

The urease-producing microbe Helicobacter pylori (formerly
Campy/obecter pyloridis) has been implicated in another type of infection.
This bacterium has been reported to colonize at intercellular junctions of the
mucosal lining of the mammalian stomach (15, 16). The urease of H. pylori
rapidly hydrolyzes urea from serum to form large amounts of ammonia. The
elevated level of ammonia causes a rise of pH in the immediate region of the
stomach and allow the bacteria to grow in a relatively neutral pH range. This
organism is associated with development of 8 gastritis (16, 17) and peptic

ulcers (18).

Agricultural significance. A major limitation to plant growth is the
lack of fixed nitrogen in the soil. Urea can be a good nitrogen source
because of its ease of application and its high nitrogen content (19).
Hydrolysis of urea by soil ureases is necessary before it can be used in the
plant. However, uncontrolled hydrolysis can result in the rise of soil pH and
ammonia toxicity. Urea hydrolysis may be able to be controlled by using
urease inhibitors (20, 21). Although the urease inhibitor
phenylphosphorodiamidate appears not to be effective for an increase in
production of corn (22), this approach has the potential to solve the problems
mentioned above .

Microbial ureases are also important in the nitrogen metabolism of
ruminant livestocks that contain a forestomach (23). Urea produced from
metabolism in liver or other tissues is recycled to the rumen through saliva or
the bloodstream. Several ureolytic strains of bacteria including
Staphylococcus, Klebsiella, Selenomonas, Succinovibrio, Bifidobacterium,
and Ruminococcus spp. (24, 25) hydrolyze urea to produce ammonia, an
important nitrogen source for the ruminal bacteria (26). The biomass of the

bacteria can be then utilized as a nutrient by the ruminant (reviewed in 5).

Urease structural properties. Bacterial ureases are heteropolymeric
proteins. For example, the ureases of Klebsiella aerogenes (27), Proteus
mirabilis (28, 29), Providencia stuartii (30), Selenomonas ruminantium (27,
31), Morgana/Ia morganii (32), Ureaplasma ureolyticum (33), Lactobacillus
reuten’ (34), Lactobacillus fermentum (35), and Bacillus sp. strain TB-90 (36)
consist of one large (a; M, = 60,000 to 75,000) and two distinct small
subunits (B and y; M, = 8,000 to 11,000). In contrast, the bacterial strain, H.
pylori possesses a urease of two subunits; one large subunit (M, = 66,000) is
homologous to the a subunit of the other bacterial ureases, whereas the
other subunit (8; M, = 29,500) looks like a fusion of the B and 7 subunits of
others (37). The stoichiometry of the subunits for bacterial ureases may not
be consistent in the different sources. K. aerogenes urease was reported to
possess a (043272); structure as deduced by native size and denaturing gel
scanning methods; however, a (aBy)3 structure has been established in
preliminary X-ray crystallographic analysis. Stoichiometries reported for
those of L. reuteri (34) and L. fermentum (35) were 0432)! and (0432”); for P.
stuartii (30), am for H. pylori (37), (123472 for Streptococcus mitior (38);
however, these enzymes are also likely to be trimers of dimers or trimers.

The urease structural genes have been cloned and sequenced from K.
aerogenes (39), Proteus vulgaris (40), P. mirabilis (29), U. ureolyticum (41),
Bacillus sp. strain TB-90 (36), H. pylori (42), Yersinia enterocolitica (43),
Rhizobium meliloti (44), Helicobacter felis (45), Bacillus pasteurii (46),
Staphylococcus xylosus (47), Lactobacillus fermentum (48) and jack bean
(49). In contrast to the bacterial enzyme, jack bean urease is homohexameric
(subunit Mr = 90,770; 73). Comparison of the amino acids and DNA
sequences of jack bean (49, 50) and bacterial ureases indicates the genes

are highly conserved through evolution. It is likely that the single plant gene

6

of the plant urease was formed via the gene fusion of the multiple structural

genes from bacteria.

Urease active site. Urease is the first enzyme shown to have nickel
(7). All ureases examined possess nickel as a cofactor (5, 51). K. aerogenes
urease apoprotein has been prepared from a nickel-deficient culture. In spite
of lacking enzyme activity, the apoprotein showed characteristics similar to
holoprotein in terms of the composition of urease subunits (52). Furthermore,
preliminary X-ray crystallographic analysis showed no difference between the
structures of apoprotein and holoprotein except the absence of nickel in the
former protein. These results are consistent with the nickel cofactor playing
an important role in enzyme catalysis rather than participating in a structural
role. Bacterial and plant ureases have been shown to possess 0.8-2.1 mol
nickel/mol large subunit (reviewed in 5). In combination with the analysis of
slow-binding inhibitors, the K. aerogenes (27, 53) and jack bean (7, 54)
ureases have 2 mol nickel/mol catalytic unit. In spectroscopic studies, the
interaction of the plant urease with B-mercaptoethanol was shown to produce
an increase in absorbance at ~320, ~380, and 425 nm due to a thiolated
anion->Ni(ll) charge transfer interaction. The change is associated with a K,
of 0.95 mM which is compatible with the K. value of 0.72 mM determined for
the competitive inhibitor (55). K. aerogenes urease also interacts with B-
mercaptoethanol to result in increased absorbance at 322, 374, and 432 nm
with a K, of 0.38 mM (53). This value is again consistent with the K. of 0.55
mM determined kinetically. The close relationship of spectroscopically
measured thiol binding to nickel of urease and the inhibitory effect of the thiol
on urea hydrolysis supports the assumption that urea binds to nickel during
the catalysis. These results again suggest that nickel ions are located

proximal to the active site and play a catalytic role (3, 55).

7
X-ray absorption spectroscopy (XAS) of jack bean urease (56, 57, 58)

has demonstrated that the nickel is coordinated by a mixture of nitrogen and
oxygen ligands, implying the involvement of amino acids containing those
atoms in their side chains. In a study involving photooxidation of jack bean
urease, it was demonstrated that histidine residues are destroyed when the
enzyme is illuminated by intense light in the presence of methylene blue. The
photooxidation was found to be suppressed in the presence of the tight-
binding urease inhibitor, acetohydroxamic acid (59). This early chemical
modification study suggests the presence of one or more histidine residue
located proximal to the active site. K. aerogenes urease apoprotein showed
reactivity with diethylpyrocarbonate (DEP), a histidine-selective modifier (52).
In the following site-directed mutagenesis studies (Chapter 3, 60), H134,
H136, and H246 residues of the a subunit of K. aerogenes urease were
substituted with alanine. The aH134A, aH136A and aH246A mutant proteins
have no detectable activities and possess one or fewer nickel ion bound per
active site (60). These results strongly suggest the ligation of nickel by these
histidine residues.

The presence of two ionizable groups essential for urease activity was
proposed from the pH dependence studies with the K. aerogenes urease
enzyme (29). One group (pK,=6.5) must be deprotonated and a second
group (pK.=8.9) must be protonated for catalysis. Although the identity of
these residues was unknown at the time, the former residue was proposed to
act as a general base and the latter one as a general acid. In succeeding
experiments with the same urease it was found that the histidine-selective
reagent DEP rapidly inactivates the enzyme (Chapter 2, 61) in a modification
reaction that is associated with a pK. of 6.5, consistent with modification and
inactivation of the residue which acts as the general base. Substitution of

aH320 with alanine by site-directed mutagenesis resulted in a mutant protein

8

with very low activity (~0.003% of wild-type protein) that was apparently
resistant to DEP (Chapter 3, 60), consistent with this being the residue acting
as a general base in urea hydrolysis.

K. aerogenes urease is also susceptible to disulfide and alkylating
agents. The pH dependence of the inactivation kinetics is compatible with the
presence of a single essential thiol interacting with a second ionizable
residue together yielding macroscopic pK. values of <5 and 12 (62). Active
site labeling studies have identified the cysteine as residue 319 in the a
subunit of the urease protein (63). Three site-directed mutant proteins
(C319A, C3198, and C319D) showed a shift in pH optimum compared to
wild-type enzyme. Based on all these results, the authors proposed that the
cysteine residue ionically interacts with a second residue, together acting as
a proton donor (general acid) during catalysis (64).

Inhibitor studies and inactivator with the same enzyme indicate the
presence of a negatively charged residue at the active site. Uncharged or
positively charged thiolates were found to have lower Ki values than
inhibitors possessing a negatively charged carboxyl group (53). Similarly,
negatively-charged chemical reagents inactivate the enzyme more slowly
than uncharged or positively charged compounds (62). It is suggested that a
negative charge in the active site may repel the carboxylated inhibitors and

inactivators.

Urea hydrolysis reaction mechanism. A mechanism for the
hydrolysis of urea has been proposed for the jack bean urease (65) and the
mechanism is retained here (Scheme 2) with some elaboration. One nickel
ion in the active site is supposed to bind to the carbonyl oxygen of urease. A
general base (His-320) is thought to activate a hydroxide bound to the other

nickel ion and it nucleophilically attacks the urea carbon. A molecule of

9

ammonia is released with the action of a general acid (comprising Cys-319
and another unidentified residue) to leave a carbamate. As a final step of
catalysis, the carbamate dissociates from the active site and spontaneously

decomposes to ammonia and carbon dioxide.

 

 

 

 

 

 

 

 

Nl—o—c :u—‘o- -NI
fondant”
2H9}
Ni —o-c\ I—o Nl—o—c _°2

SchemeZ

In vivo activation of urease apoprotein. Even in the absence of
nickel, urease apoprotein is synthesized in both prokaryotes and eukaryotes
(66, 67). Thus, nickel ion appears not to be involved in transcriptional
regulation of urease. The addition of nickel to intact K. aerogenes cells was
shown to activate urease apoprotein. This slow and partial activation process
was found to be independent of protein synthesis, but sensitive to a proton
uncoupler, dinitrophenol, or an ATP synthase inhibitor, DCCD (52), indicating
that in vivo activation is an energy-dependent process. In vivo activation of
urease apoprotein also has been reported for several other bacteria

including P. mirabilis (68), a purple sulfur bacterium (69), and a

10

cyanobacterium (70), as well as eukaryotes such as several algae (71) and

soybean (67).

Accessory genes for urease activation in bacteria. Formation of the
nickel-containing urease holoprotein in vivo has been shown to require
several genes in addition to the structural genes encoding the urease
subunits. For example, the K. aerogenes urease operon has four additional
reading frames (ORF’s) which were classified as accessory genes by the
authors (39, 72). The ureD gene is located upstream of the ureA, ureB, and
ureC genes encoding the urease subunits, and the ureE, ureF, and ureG
genes follow the three structural urease genes. The same arrangement of
genes has been found in several other microorganisms including P. mirabilis
(73, 74), and a ureolytic strain of E.coli (75). The urease operon of Bacillus
sp. strain TB-90 has a slightly altered organization of its urease gene cluster,
where the location of ureD is downstream of ureG and additional genes, ureH
and ureI, are present (36). The urease gene cluster from Helicobacter pylori
was shown to have nine genes in three blocks (76). Among them, two may
have a regulatory role, two genes encode urease subunits and four genes
appear to encode accessory proteins analogous to the counterparts of K.
aerogenes. Transposon mutagenesis of the H. pylori urease gene cluster
allowed the authors to identify ureF, ureG and ureH (homologous to ureD in
K.aerogenes) as the essential accessory genes (76). A similar observation
was made in studies with deletion mutants of the urease gene cluster of
Bacillus sp. strain TB-90. The deletion of ureE caused somewhat impaired
urease activity, whereas, deletions in ureF, ureG, and ureD abolished the
enzyme activity (36). The importance of the accessory genes also has been
shown in K. aerogenes urease formation (72). Inactive urease protein was

synthesized in E.coli carrying the K. aerogenes urease gene cluster with

11

deletions in ureD, ureF, or ureG. The ureases from each deletion mutant
turned out to have little or no amount of nickel. The defects of the deletion
mutants, however, can be partially or fully repaired by complementation with
the plasmid possessing each deleted gene. Furthermore, the ureE deletion
mutant synthesizes urease enzyme with reduced activity which roughly
corresponds to the reduced nickel content of the purified urease enzyme. All
these results indicate that the accessory gene products are trans-acting
factors essential for the incorporation of a functional nickel metallocenter into
urease protein.

It is evident that elucidation of the functions of each accessory protein
is essential for understanding the mechanism and control of nickel
metallocenter formation. Although not fully understood, several roles of the
proteins have been speculated. UreE purified from K. aerogenes has been
shown to bind approximately six nickel ions per dimer with a K. of ~10 uM
(77). The authors suggested that UreE may bind nickel ion and act as the
nickel donor to the urease apoprotein. UreF is expressed in very low
amounts in E. coli containing K. aerogenes urease gene cluster. This protein
has not been identified or purified except as a part of a protein complex
(Chapter 6), which appears to have urease apoprotein, UreD, UreF and
UreG. It is possible that UreF may play its unknown role only as a part of the
complex. The function of UreG has not been determined, either. However, it
is intriguing that the protein was revealed to have a P-loop motif (78) that is
found in a variety of ATP- and GTP-binding proteins. This feature and the
energy dependence for in vivo nickel ion incorporation suggest that UreG
may bind ATP or GTP and couple its hydrolysis to the nickel incorporation

event

12

In vitro activation of urease apoprotein and the probable role of
UreD. In vitro activation of purified K. aerogenes urease apoprotein has been
accomplished by simply providing 002 (half-maximal activation at ~0.2%
C02) as well as nickel ion (Chapter 5). Based on the pH-dependent
activation (pKa 2 9) and the precedent example of ribulose 1,5-bisphosphate
carboxylaseloxygenase (reviewed by Hartman and Harpel, 79), it is proposed
that C02 binds to apoprotein to form a ligand for nickel.

Prior to the successful activation of purified urease apoenzyme, in vitro
activation of a UreD-urease complex was reported (80, Chapter 4). Since the
poor expression of ureD in E. coli harboring the K.aerogenes urease gene
cluster was suspected to be due to the weak ribosome-binding site, site-
directed mutagenesis methods were used to overexpress the gene. UreD was
found to co-purify with urease apoprotein in ion-exchange, size exclusion,
and hydrophobic column chromatographies. The UreD-urease apoprotein
complexes are able to be activated alone (Chapter 4), but also were shown
to be activated in a C02 concentration-dependent manner (Chapter 5). In
both the apoprotein and the UreD-apoprotein complex, activation appears to
be accompanied by a non-productive reaction that results in enzyme
inactivation. However, the presence of UreD evidently reduces the rate of
this inactivation step (Chapter 5, Chapter 6). Thus, it was proposed that the
UreD protein appears to function, at least in part, to prevent apoprotein from

binding nickel non-productively in the absence of C02.

Eukaryotic ureases. Urease activities are present in many
eukaryotes, including yeast (81), other fungi (82, 83), and plants. Urease
from soybean (glycine max) is a well studied eukaryotic enzyme
genetically. Four urease-related genes are identified. The Eu1 locus

encodes a urease structural gene that is expressed at very high levels in

13
the developing embryo (84, 85, 86). The partial amino acid sequences

deduced from cloned genes apparently match those of the purified jack
bean urease (87). An isozyme was found to be encoded by the EM locus
and is constitutively expressed at low levels in all soybean tissues (88). In
addition to the urease structural genes, accessory genes appear to be
required for the functional plant enzymes. The defect in Eu2 or Eu3 gene
loci result in deficient ureases (89). These loci appear to encode gene
products that are required for a maturation of functional urease; for
example, nickel ion incorporation.

Four complementation groups were identified for the formation of
active urease in Aspergillus nidulans (83). It has been found that ureA
encodes a urea transport protein, ureB is the single subunit urease
enzyme, and ureC is essential for the formation of active urease, but the
function is not known. The suppression of a ureD mutation by culture of
this organism in the presence of 0.1 mM nickel sulfate (82) indicates that
the gene product may play a role in the incorporation of the nickel
cofactor. Four loci are also reported to be required for functional urease
activity in Neurospora crassa (90) and in Schizosaccharomyces pombe
(83).

14
Outline of this thesis

The following chapters describe my studies on characterization of Klebsiella
aerogenes urease and in vitro activation of urease apoprotein. In Chapter 2, I
present evidence that urease possesses at least one essential histidine
residue per catalytic unit which may act as a general base [published in J.
Protein Chem. 12, 51 -56 (1993) (61)]. In Chapter 3, I describe the generation
of and characterization of ten histidine—+alanine mutants by using site-
directed mutagenesis. I identify histidine residues that appear to function in
nickel ligation, substrate binding, and catalysis [published in Protein Science.
2:1034-1041 (1993) (62)]. In Chapter 4, I describe studies related to in vitro
activation of UreD-urease apoprotein complexes. One, two, or three UreD
molecules are present per enzyme in different complexes and the UreD
protein dissociates from the enzyme during activation (published in PNAS
91:3233-3237 (1994) (80) with results related to the in vitro reconstitution of
urease apoprotein in cell extracts by Mary Beth Carr]. In Chapter 5, l
characterize apoprotein activation in the presence of 002 and nickel and
compare the properties of this process with that for UreD-urease apoprotein
complexes [submitted to Science]. Chapter 6 contains several sections. In
section A, I describe the characterization of H219NIQ and H320NIQ mutant
proteins. In section B, I present studies that further explore the possible role
of UreD in the in vitro activation of urease. In section C, partial purification
and characterization of UreD are described. In section D, I show evidence
consistent with the existence of at least two different forms of UreD-urease
protein complex containing the same number of UreD molecules bound per
urease apoprotein. In section E, I show the presence of protein complexes

containing urease apoprotein, UreD, UreF and UreG and describe its partial

1 5
characterization. In the last section, I suggest speculative models of urease
apoprotein activation.

Additional studies that l have not included here involve several protein
preparations for X-ray crystallographic analysis (urease holoprotein, H134A,
H219A, H320A, UreE, and the three, enriched UreD-urease apoprotein
complexes), variable temperature magnetic circular dichroism spectroscopy
(urease holoprotein, H134A), and X-ray absorption spectroscopy (urease
holoprotein, H134A). Finally, the construction of several plasmids that I have

made is appended.

10.

11.

12.

16
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88.

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24

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CHAPTER 2

Diethylpyrocarbonate Reactivity of Klebsiella aerogenes Urease: Effect of

pH and Active Site Ligands on the Rate of Inactivation

These studies were published in J. Protein Chem. 12, 51 -56, 1993.

25

26
ABSTRACT

Reaction of Klebsiella aerogenes urease with diethylpyrocarbonate (DEP) led to
a pseudo-first-order loss of enzyme activity by a reaction that exhibited
saturation kinetics. The rate of urease inactivation by DEP decreased in the
presence of active site ligands (urea, phosphate, and boric acid), consistent with
the essential reactive residue being located proximal to the catalytic center. The
pH dependence for the rate of inactivation indicated that the reactive residue
possessed a pKa of 6.5, identical to that of a group that must be deprotonated
for catalysis. Full activity was restored when the inactivated enzyme was treated
with hydroxylamine, compatible with histidinyl or tyrosinyl reactivity.
Spectrophotometric studies were consistent with DEP derivatization of 12 mol of
histidine/mol of native enzyme. In the presence of active site ligands, however,
approximately 4 mol of histidine/mol of protein were protected from reaction.
Each protein molecule is known to possess two catalytic units; hence, I propose

that urease possesses at least one essential histidine per catalytic unit.

27
INTRODUCTION

Urease is a nickel-containing enzyme that catalyzes the hydrolysis of urea to
form ammonia and carbamate; the carbamate spontaneously decomposes to
form a second molecule of ammonia and carbonic acid (Andrews et al., 1988;
Mobley and Hausinger, 1989). The best studied bacterial urease is that from
Klebsiella aerogenes: the native enzyme consists of three subunits (M =
60,304, 11,695, 11,086; Mulrooney and Hausinger, 1990) in apparent 098474
stoichiometry and contains two active sites, each of which possesses two tightly
bound nickel ions (Todd and Hausinger, 1989). In addition to the requisite bi-Ni
metallocenter, an essential cysteine residue has been suggested to be located
proximal to the K. aerogenes active site (Todd and Hausinger, 1991a). The pH
dependence of urease inactivation by disulfide and alkylating agents is
consistent with a single essential thiol interacting with a second ionizable
residue together yielding macroscopic pKa values of <5 and 12. Active site
labeling studies have identified the cysteine as residue 319 in the large subunit
(Todd and Hausinger, 1991b), but the identity of the second ionizable residue is
unknown. The pH dependence of urease catalysis is compatible with the
presence of two additional ionizable groups participating in the hydrolytic
reaction (Todd and Hausinger, 1987); one group (pKa a 6.55) must be
deprotonated and a second group (pKa a 8.85) must be protonated for catalysis.
Although the identity of these catalytic residues is unknown, histidine is a
reasonable possibility for the group of lower pKa. Furthermore, in a study
comparing properties of urease holoenzyme and apoprotein Lee et al. (1990)
noted that urease was inactivated by diethylpyrocarbonate (DEP). This result is
also consistent with the presence of an essential histidine residue. Here, I follow
up this observation to more carefully detail the DEP reactivity of urease and
assess whether a histidine residue may be present at the active site.

28
MATERIALS AND METHODS

Materials. DEP, obtained from Sigma Chemical Co., was dissolved in
ethanol immediately before use. DEP concentration was measured by reacting
an aliquot with 10 mM imidazole (pH 7.0) and monitoring the absorbance at 230
nm using an extinction coefficient of 3,000 M'1cm'1 (Miles, 1977).

Enzyme Purification. Urease was purified to a specific activity of >2500
units/mg from K. aerogenes carrying plasmid pKAU19 (Mulrooney et al., 1989)
by procedures described previously (Todd and Hausinger, 1989). One unit of
enzyme activity is defined as the amount of enzyme required to degrade 1 umol
of urea per min in the standard assay mixture which contained 50 mM urea, 0.5
mM EDTA, and 25 mM HEPES, pH 7.75, at 37°C. Linear regression analysis of
the released ammonia, determined by conversion to indophenol, versus time
yielded the initial rates. Protein was assayed by the method of Lowry et al.
(1951)

Kinetics of Urease Inactivation. Inactivation reactions contained 0.5 mM
EDTA, 80 mM HEPES, pH 7.0 buffer (or other buffers and pH values as
indicated for the pH dependence studies) plus the indicated concentrations of
reagent and inhibitor (if present). All reactions were performed at 37°C.
Aliquots were removed at the indicated time points and diluted 200-fold into
assay buffer. The stability of DEP in the various enzyme inactivation buffers was
assessed in control experiments, and enzyme inactivation reactions were carried
out over a sufficiently short period of time to minimize concerns over DEP
hydrolysis.

Spectroscopy. All spectra were obtained by using a Gilford Response
spectrophotometer. Spectral data were transferred to an IBM PC for calculation
of difference spectra.

29

Recovery of Enzyme Activity by NHZOH Treatment. Urease (140 nM)
was inactivated with DEP (50 pM) for 1 min in 0.5 mM EDTA, 80 mM HEPES, pH
7.0 at 37°C. Excess DEP was reacted with imidazole (final concentration of 0.5
mM), and hydroxylamine was added to a concentration of 0.5 mM. The enzyme
mixture was incubated at 37°C and aliquots were assayed for recovery of
activity.

RESULTS AND DISCUSSION

Inactivation of Urease by DEP. The effect of DEP concentration on
urease inactivation at pH 7 is shown in Figure 1. At each inhibitor concentration,
a pseudo-first-order loss of activity was observed; i.e., In VtNo (where Vt is the
velocity at time t and V0 is the initial velocity) is linear with time until less than
5% of the starting activity remains. Analogous studies were carried out at pH 6,
and the observed rate constants for both experiments were plotted versus DEP
concentration to yield the lines shown in Figure 2. At pH 6, high concentrations
of DEP clearly demonstrated partial saturation of the inactivation rate.
Saturation kinetics is expected for a reagent which reacts as illustrated in
Scheme 1.

g

k . .
E+ l ‘— El -1-> inactive enzyme
Scheme 1
For inactivators which follow this scheme (where k1 is much slower that the rate

of El dissociation), the apparent rate of urease inactivation (kapp) obeys
Equation 1.

30

 

 

 

 

O \\. I I i
-1 — \I ' . . —
’5 -2 — ‘ - -
é. ’ . .
.>, , .
5 '3 L . fl
9 o ‘
<>
-4 - o
O
_5 1 I I 1 1 I
0.0 0.2 0.4 0.6 0.8 1.0 1.2 1.4
Time (min)

Figure. 1. Pseudo-first-order urease activity loss in the presence of DEP.
The fraction of activity remaining is shown as a function of time for urease (140
nM) reacting with DEP at 5 (0), 10 (O), 20 (V), 30 (V), 40 (U), 50 (I), 60 (A),
70 (A) 80 (0), and 100 (9) (AM. All reactions were in 0.5 mM EDTA, 80 mM
HEPES buffer at pH 7.0 and 37°C.

31

 

 

 

 

L l l 4 J

0 50 100 150 200 250 300
DEP (,uM)

Figure. 2. Effect of DEP concentration on pseudo-first-order rate of
inactivation. The pseudo-first order rate constants calculated from Figure. 1 (e)
and analogous constants determined at pH 6.0 (0) were plotted as a function of

DEP concentration.

32

it,” = km (1)
[11+K.

 

At pH 6, Kd and k1 were estimated to be 327 1 36 uM and 8.3 1 0.6 min-1,
yielding a second-order rate constant (k1le) of 25,400 M‘1 min'1. Non-
linearity in the pH 7.0 data suggest that saturation also occurs here, however,
the extremely rapid inactivation rate precluded estimation of the individual
constants. Nevertheless, a value of 60,000 1 2,000 M'1 min'1 was calculated
for the second-order rate constant. These second-order rate constants surpass
all other DEP-dependent second-order inactivation rates tabulated for diverse
enzymes (Lundblad and Noyes, 1984), with the nearest similar rate being 10,380
M'1 min'1 at pH 7.2 for lactate dehydrogenase (Bloxham, 1981).

Effect of Ligands on the Rate of Urease Inactivation by DEP. Ligands
which bind to the active site of urease were examined for their effect on the rate
of inactivation by DEP. At 40 uM DEP, increasing concentrations of both borate
and phosphate (competitive inhibitors of urease) were found to reduce the rate
of inactivation (Figure. 3). At elevated phosphate concentrations, kapp was less
than 20% that of the control; whereas, saturating levels of borate decreased
kapp only by approximately 40%. These results can be accomodated by the
model shown in Scheme 2 (where the ligands are represented by L).

E+ l Ké El £19 inactive enzyme
+
L

II KL (XKd 2
EL+| v——‘ ELI -> inactive enzyme

Scheme 2

33

 

12 // //

0 I I/[T T T T fl I// I
1.0
0.8F

0.6 F

 

0.4 -

0.2 - an M65

0.0 1 1 7/// 1 1 1 1 1 1//// 1
0.0 0.2 0.4 4.06.0 8111 4 8 12 200 300 400

 

 

 

kapp (fraction of control)

 

Boric acid (mM) Phosphate (mM)

Figure. 3. Effect of competitive inhibitors on the rate of urease inactivation
by DEP. The second-order rate constants for urease inactivation by DEP (40
pM) at pH 7.0 were determined in the presence of boric acid (panel A) or
phosphate (panel B) at the indicated concentrations. The values shown are
expressed as fraction of the inactivation rate in the absence of inhibitors.

34 .
The rate of inactivation for a reaction that follows Scheme 2 is shown in

Equation 2 (Carrillo et al., 1981 ).

= (k,KLaK, + k,iLiK.)iIi + K [WK
‘“’ aK.K.<K. + [I] " “

 

+ [1]) (2)

The demonstration that active site ligands can decrease the rate of enzyme
inactivation is consistent with DEP reacting with a residue proximal to the active
site of urease.

A more detailed analysis of the results shown in Figure 3 highlights a
problem with the simple model of Scheme 2. Values of 0.07:0.02 mM and 6:2
mM were graphically determined for the KL of borate and phosphate; yet, these
compounds were found to possess simple competitive inhibition (Ki) values of
0.223 mM and 50.4 mM at this pH value (pH 7.0). These results indicate that the
binding step associated with KL is distinct from that associated with Ki. Our data
do not allow us to discriminate among models that could account for this
behavior. The effects of ligands on urease inactivation by DEP contrasts with
the results observed for alkylating and disulfide reagents (Todd and Hausinger,
1991a) where the KL measured for the effect of ligands on inactivation matched
the K; of these inhibitors.

Urea (100 mM) also protected urease from modification by DEP (Figure
4). Although a portion of the urea was degraded under these experimental
conditions, there was sufficient buffer capacity to prevent significant pH change
and the urea concentration remained saturating during the short time required
for the analysis.

Effect of pH on the Rate of Urease Inactivation. The effect of pH on the rate
of urease inactivation by DEP is illustrated in Figure 5. The pKa estimated from
these data is 65:01. This value is identical to that of a urease residue which

35

 

 

 

 

0
-1
5’
s -2
E
-3
.4

 

0.0 0.2 0.4 0.6 0.8 1.0 1.2 1.4
Time (min)

Figure. 4. Effect of urea on the rate of urease inactivation by DEP. Urease
(140 nM) was inactivated by DEP (60 (M) in the presence (0) or absence (0)
of urea (100 mM) at 37°C in 0.5 mM EDTA, 80 mM HEPES buffer, pH 7.0. The
fractional activity remaining is shown as a function of time.

36

2.5

 

2.0 -

V
1.5- D.
o

kapp (min‘1)

8
0.5 '— .V

 

 

0.0 I I I

 

Figure. 5. The pH dependence of DEP inactivation of urease. The apparent
second-order rate constants for urease inactivation with 25 11M DEP are plotted
as a function of pH for reactions carried out at 37°C in 0.5 mM EDTA, 80 mM
buffer: MES (O), MOPS (O), HEPES (V), TAPS (V), Tricine (Cl). At pH values
less than 6, the reactions were carried out using the same conditions as above
except that the concentration of DEP was 125 uM and the values were adjusted
by using the equation (1 ).

37 .
must be deprotonated for catalysis (Todd and Hausinger, 1987). I propose that

the DEP-reactive residue is the general base required for urea hydrolysis.
Although DEP has traditionally been viewed as a histidine-selective
reagent, reaction with protein tyrosine, lysine, and other nucleophilic groups has
been reported (reviewed by Miles, 1977; Lundblad and Noyes, 1984). In order
to identify the nature of the amino acid that corresponds to the essential residue,
I probed the stability of the modified residue and examined the inactivation

reaction spectroscopically as detailed below.

Reactivation of DEP-modified Urease. Treatment of DEP-inactivated
urease with 0.5 mM NHZOH almost completely restored urease activity within 6
hr (Figure 6). The restoration of activity of DEP-treated urease with NHZOH
implies that the modification involved either histidine or tyrosine residues and
suggests that lysine modification is not involved in activity loss (Miles, 1977;
Lundblad and Noyes, 1984).

Spectrophotometric Analysis of Urease Reactivity with DEP. Urease
(9.4 11M) was incubated in 80 mM HEPES and 0.5 mM EDTA, pH 7.0, containing
250 uM DEP for 30 min. The change in absorbance was monitored versus a
reference cuvet containing urease in buffer without DEP (Figure 7). Modification
of histidine residues leads to an increase in absorbance at 242 nm with an
extinction coefficient of 3,200 M'1cm‘1 (Lundblad and Noyes, 1984), whereas
modification of tyrosine leads to a spectral decrease at 280 nm with an extinction
coefficient of 1,310 M'1cm‘1 (Mulhrad et al., 1967). Our spectroscopic results
were consistent with modification of ~12 histidine residues per mol of enzyme.

The rate of urease activity loss was compared to the kinetics of
absorbance change at 242 nm for enzyme (4.2 uM), in 25 mM HEPES (pH 7.0)
buffer, treated with 100 11M DEP (Figure 8, open circles). The experiment was
repeated in the presence of 400 mM phosphate, a competitive inhibitor of urease

(Figure 8, closed circles). Protection of approximately two histidine residues

38
correlated to protection of ~60% of urease activity, consistent with the presence

of nearly four histidines protected by phosphate binding in fully active enzyme.
In contrast, small changes observed at 280 nm did not correlate with changes in
urease activity, precluding the presence of an essential tyrosine. It has been
shown that there are two active sites per native urease molecule (Todd and
Hausinger, 1989), hence, I conclude that there is at least one essential histidine
residue proximal to each active site.

39

 

 

O)
O

.h
0

Activity (%)

N
O

 

 

 

I l l

0 5 10 15 20 25
Time (hr)

C

Figure. 6. Reactivation of DEP-inactivated urease by hydroxylamine. DEP-
inactivated urease was incubated with 0.5 mM hydroxylamine at 37°C and
activity was monitored over time. The results are reported as percent of control
(non-DEP-treated) urease subjected to the same conditions.

0.4
0.3
0
0
5
e 0.2
O
U)
.0
< 0.1
VI
0.0

40

 

 

 

I

 

 

230 240 250 260 270 280 290

Wavelength (nm)

Figure. 7. Spectroscopic analysis of urease modification by DEP. Urease
(9.4 (M) was incubated for 30 min with DEP (250 nM) in 0.5 mM EDTA, 80 mM
HEPES buffer, pH 7.0 at 25°C. The difference spectrum is shown for this
sample minus untreated urease in same buffer.

41

 

 

 

 

 

 

 

1.0
0.8
o 0.6
%.
> 0.4
0.2
A 0.0
Q.
E 0.08- "an... «6 U
N o (D
R’. 006 °' 5:3:
§ ' ‘4 3
- 22
g 0.04 ‘ .. ‘ ‘ . . ‘ :1:
a A A A A A ‘ ‘ (2 “5
3 0.02- =14:
<
0.00 1 1 1 1 1 O
0.0 0.5 1.0 1.5 2.0 2.5 3.0

Time (min)

Figure. 8. Kinetic comparison of activity and absorbance changes during
modification by DEP. Urease (4.2 uM) was incubated with DEP (100 nM) in 25
mM HEPES buffer, pH 7.0 at 25°C in the absence (0) or presence (0) of 400
mM phosphate. Changes in activity (A) and absorbance at 242 nm (8) were
monitored, and differences between the phosphate-free and phosphate-
containing samples were calculated (A).

42

REFERENCES

Andrews, R. K., Blakeley, R. L., and Zemer, B. (1988). In The Biainorganic
Chemistry of Nickel (Lancaster, J. R., ed) pp. 141 -165, VCH Publishers,
Inc., New York.

Bloxham, D. P. (1981). Biochem. J. 193, 93-97.

Carrillo, N., Arana, J. L., and Vallejos, R. H. (1981). J. Biol. Chem. 256, 6823-
6828.

Lee, M. H., Mulrooney, S. B., and Hausinger, R. P. (1990). J. Bacterial. 172,
4427-4431.

Lowry, O. H., Rosebrough, N. J., Farr, A. L., and Randall, R. J. (1951). J. Biol.
Chem. 193, 265-275.

Lundblad, R. L.,and Noyes, C. M. (1984). Chemical reagents for protein
modification, CRC Press, Boca Raton, Florida, pp. 105-125.

Miles, E. W. (1977). Math. Enzymol. 47, 431-442.
Mobley, H. L. T., and Hausinger, R. P. (1989). Microbiol. Rev. 53, 85-108.

Mulhrad, A., Hegyi, G., and Toth, G. (1967). Acta Biochim. Biophys. Acad. Sci.
Hung. 2, 19-29.

Mulrooney, S. B., and Hausinger, R. P. (1990). J. Bacterial. 172, 5837-5843.

Mulrooney, 8. B., Pankratz, H. 8., and Hausinger, R. P. (1989). J. Gen.
Microbial. 135, 1769-1776.

Todd, M. J., and Hausinger, R. P. (1987). J. Biol. Chem. 262,5963-5967.
Todd, M. J., and Hausinger, R. P. (1989). J. Biol. Chem. 264, 15835-15842.
Todd, M. J., and Hausinger, R. P. (1991 a). J. Biol. Chem. 266, 10260-10267.

Todd, M. J., and Hausinger, R. P. (1991b). J. Biol. Chem. 266, 24327-24331.

CHAPTER 3

Site-directed mutagenesis of Klebsiella aerogenes urease: Identification of
histidine residues that appear to function in nickel ligation, substrate

binding, and catalysis

These studies were published in Protein Science 21034-1041, 1993.

43

44
ABSTRACT

Comparison of six urease sequences revealed the presence of ten conserved
histidine residues (H96 in the 7 subunit, H39 and H41 in [3, and H134, H136,
H219, H246, H312, H320 and H321 in the ct subunit of the Klebsiella
aerogenes enzyme). Each of these residues in K. aerogenes urease was
substituted with alanine by site-directed mutagenesis and the mutant proteins
were purified and characterized in order to identify essential histidine
residues and assign their roles. The yH96A, BH39A, BH41A, otH312A and 0t
H321A mutant proteins possess activities and nickel contents similar to wild-
type enzyme, suggesting that these residues are not essential for substrate
binding, catalysis, or metal binding. In contrast, the otH134A, 0tH136A and 0t
H246A proteins exhibit no detectable activity and possess 53%, 6%, and
21% of the nickel content of wild-type enzyme. These results are consistent
with aH134, 0tH136 and 01H246 functioning as nickel ligands. The otH219A
protein is active and has nickel (~1.9% and ~80%, respectively, when
compared to wild-type protein), but exhibits a very high Km value (1100:40
mM compared to 23:02 mM for the wild-type enzyme). These results are
compatible with 0tH219 having some role in facilitating substrate binding.
Finally, the aH320A protein (Km=8.3:0.2 mM) only displays ~0.003% of the
wild-type enzyme activity, despite having a normal nickel content. Unlike the
wild-type and 0tH219A ureases this mutant protein was not inactivated by
diethylpyrocarbonate (DEP), consistent with 0tH320 being the DEP-reactive

general base that facilitates catalysis.

45

INTRODUCTION

Urease (EC 3.5.1.5), found in a variety of plants and a broad range of bacterial
species, is a nickel-containing enzyme that catalyzes the hydrolysis of urea to
form carbonic acid and two molecules of ammonia (Mobley 8. Hausinger, 1989).
The best studied microbial urease is that from the enteric bacterium, Klebsiella
aerogenes. The enzyme is comprised of three subunits [Mrs=60,304 (a), 11,695
(B), and 11,086 (7) (Mulrooney & Hausinger, 1990)] and possesses two nickel
ions per 043272 catalytic unit (Todd & Hausinger, 1987, 1989). Although K.
aerogenes urease has been crystallized and the crystals shown to diffract to less
than 2 A (Jabri et al., 1992), the three-dimensional structure of the enzyme has
not yet been elucidated. Nevertheless, several structural features of the
bacterial urease active site have been characterized and multiple essential roles
for histidine residues have been implicated. For example, the pH dependence of
enzyme activity is consistent with the presence of two chemical groups at the
active site that participate in catalysis as a general base (pKa=6.55) and a
general acid (pKa=8.85) (Todd & Hausinger, 1987). Chemical modification
studies with the histidine-selective reagent DEP were compatible with a histidine
residue serving as the general base (Park & Hausinger, 1993). Additional
chemical modification and site-directed mutagenesis studies demonstrated the
presence of an active site cysteine residue (C319 in the ct subunit) in the K.
aerogenes enzyme (Todd & Hausinger, 1991 b; Martin & Hausinger, 1992). The
pH dependence of urease inactivation by disulfide and alkylating reagents (Todd
& Hausinger, 1991 a) and the shift in pH optimum observed for C319A, C3198,

_ and C319D mutant proteins compared to wild-type enzyme (Martin & Hausinger,
1992) were interpreted in terms of the cysteine residue ionically interacting with

a second residue (X), together acting as a proton donor during catalysis.

46

Although the identity of X is unknown, the pH-dependent behavior of urease
inactivation by thiol-specific chemical reagents is reminiscent of studies
involving papain where a Cys-His ion pair has been characterized (Brocklehurst,
1987). Finally, Lee et al. (1990) compared the chemical reactivity of K.
aerogenes holoenzyme and apoprotein toward DEP and found that more
histidines are accessible to the reagent in the nickel-free protein. The enhanced
DEP-reactivity of apoprotein is consistent with histidine residues participating as
nickel metallocenter ligands in urease.

This study combines site-directed mutagenesis and enzyme
characterization methods to identify several essential histidine residues in K.
aerogenes urease and to define their roles. Comparison of the urease amino
acid sequences from jack bean (Takishima et al, 1988), Helicobacter pylori
(Clayton et al., 1990; Labigne et al., 1991), Ureaplasma urea/yticum (Blanchard,
1990), Proteus vulgaris (Morsdorf & Kaltwasser, 1990), Proteus mirabilis (Jones
8 Mobley, 1989) and K. aerogenes (Mulrooney & Hausinger, 1990) revealed the
presence of ten conserved histidines: H96 in the y subunit, H39 and H41 in B,
and H134, H136, H219, H246, H312, H320 and H321 in the ct subunit of the K.
aerogenes enzyme. I substituted each of the conserved histidine residues in the
K. aerogenes enzyme with alanine (a residue that contains a side chain which
can not function as a metallocenter ligand or general base or general acid, can
not participate in hydrogen bond interactions, and is smaller than the wild-type
residue so that it will not cause steric disruption of the structure), purified the
mutant proteins, and characterized their enzyme activities, nickel contents, and
reactivities with DEP and iodoacetamide (IAM). The results suggest that
histidine residues in the ct subunit may play key roles with H134, H136, and
H246 participating in the ligation of nickel, H219 facilitating the binding of

substrate, and H320 acting as a general base in catalysis.

47

MATERIALS AND METHODS

Materials. DEP, obtained from Sigma Chemical Co., was dissolved in
ethanol immediately before use. DEP concentration was measured by reacting
an aliquot with 10 mM imidazole (pH 7.0) and monitoring the absorbance at 230
nm using an extinction coefficient of 3,000 M‘Icm‘1 (Miles, 1977). IAM (Aldrich

Chemical Co.) was prepared in distilled water.

Site-directed mutagenesis. For yH96A, BH39A and BH41A, a 1.4-kbp
SacI-Smal fragment of the pKAU17 (Mulrooney et al., 1989) was subcloned into
M13 mp18, and mutagenized by the method of Kunkel et al. (1983). For 0t
H134A, otH136A, aH219A, aH246A, aH312A, otH320A and 0tH321A, a 1.1-kbp
BamH1-Sall fragment of the same plasmid was used. Uracil-containing, single-
stranded template DNA was prepared from E. coli CJ236 (dut1 ung1 thi-1
relA1/pCJ105[camr F']). Mutagenized phage were isolated in E. coli MV1193 (A
[lacI-praAB] rpsL thi endA spcB15 hst4 A[er-recA]306::Tn10[teI‘] F'[traD36
praAB+ lac/QIaCZAM15]). The following oligonucleotides were synthesized by
using an Applied Biosystems Model 394 DNA synthesizer at the Michigan State
University Macromolecular Structural Facility: AATCGGGTTGGCAACGGTGAC,
GAAATGGTAGGCCGAACCGAC, CTCGGCGAAAGCGTAGTGCG,
CTGAAGATCGCTGAGGACTGG, GGTCGCCCTGGCCAGCGACACC,
GGATCGATACCGCTATTCACTG, ACCCATATTGCCTGGATCTGT,
CCATCGATGAAGCTCTCGATATG, ATGGTCTGCGCCCATCTGGAC,
GTCTGCCACGCTCTGGACCCG. These primers were used to alter the ten
conserved histidine codons to encode alanine at each position. Site-directed

mutants were identified by DNA sequencing, and subcloned back into pKAU17

48

on a 1.1-kbp SacI-Mlul fragment for yH96A, BH39A, and BH41A and a 0.8-kbp
MluI-Bsml fragment for the other mutations. These regions were completely
sequenced by using Sequenase 2.0 (United States Biochemicals) and the
single-strand DNA method of Sanger et al. (1977) to ensure that no other
mutations had been introduced into M13. In one case, sequence analysis
demonstrated that a recombination event had occurred and an alternate clone
that possessed the desired muation was chosen for further studies. After
subcloning, the mutated sequences were again confirmed by double-strand DNA

sequencing methods (Sambrook et al., 1989).

Enzyme purification. Ureases were purified from E. coli DH5 carrying
pKAU17 or the site-directed mutants of pKAU17 by procedures described
previously (Todd & Hausinger, 1989), except that cells were grown in LB
medium containing 1 mM NiCI2. As noted by Lee et al. (1992) and Martin and
Hausinger (1992), the wild-type enzyme isolated from cells grown under these
conditions does not exhibit the maximum specific activity observed for enzyme
isolated from K. aerogenes [pKAU19] cells (2,500 units mg‘1, Todd & Hausinger,
1989). This decreased activity does not result from a difference in nickel
content, but rather may be related to problems arising from the high levels of
urease biosynthesis or to unknown host-dependent effects (Lee et al, 1992).
Because urease synthesis in cells containing the mutated plasmids is similar to
that in cells producing wild-type enzyme, it is reasonable to directly compare
their relative specific activities and nickel contents. Purification of inactive
mutant proteins (aH134A, aH136A, a219A, aH246A, and otH320A) was
monitored for urease-containing fractions by sodium dodecyl sulfate-
polyacrylamide gel electrophoresis using 10-15% polyacrylamide gradient gels

and the buffers described by Laemmli (1970). Sample purities were determined

49

by using a gel scanner (AMBIS Inc.) and the measured values were used for

correction of enzyme activities and nickel contents.

Assay of enzyme activity. The urease activities for wild-type and mutant
proteins except otH219A and aH320A were assayed in 25 mM HEPES, pH 7.75,
0.5 mM EDTA, and 50 mM urea. The aH219A protein activity routinely was
measured by using 1 M urea; however, for monitoring activity loss during thermal
stability and chemical modification studies a Concentration of 100 mM urea was
used. The aH320A activity was assayed in 25 mM HEPES, pH 6.75 buffer
containing 0.5 mM EDTA and 50 mM urea. One unit of enzyme activity is
defined as the amount of enzyme required to degrade 1 mol of urea per min at
37°C. Linear regression analysis of the released ammonia, determined by
conversion to indophenol (Weatherbum, 1967), versus time yielded the initial
rates. Calculation of kinetic constants made use of the method of Wilkinson

(1961 ). Protein was assayed by the method of Lowry et al. (1951 ).

Nickel quantitation. The nickel content of purified urease was assayed
by using a computer-controlled Varian Spectra AA-400Z graphite furnace atomic
absorption spectrophotometer with Zeeman background correction as previously
described (Lee et al., 1992). For calculation of the number of nickel ions per

catalytic unit, a M of 105,866 for the 043sz unit was used.

Thermal stability. Cell extracts containing wild-type and mutant urease
proteins were incubated in 20 mM phosphate, pH 7.0 buffer containing 1 mM
EDTA, and 1 mM B-mercaptoethanol at 50°C, 60°C, and 70°C. At the indicated

times, aliquots were removed and the remaining activities were assayed.

50

Inactivation by DEP. Inactivation reactions were performed at 37°C in 1
mM EDTA, 50 mM HEPES, pH 7.0 buffer (or other buffers and pH values as
described) plus the indicated concentrations of DEP. For wild-type and 0tH219A
proteins, aliquots were removed at the indicated time points and diluted at least
100-fold into assay buffer. For the otH320A protein, aliquots (100 til) were
quenched with imidazole (final concentration of 5 mM) and incubated for 5 min
on ice prior to measuring enzyme activity. The stability of DEP in the various
enzyme inactivation buffers was assessed in control experiments, and enzyme
inactivation reactions were carried out over a sufficiently short period of time to
minimize concerns over DEP hydrolysis. The pKa values for inactivation of wild-
type and aH219A proteins were calculated by fitting data sets to the following
equation (Cousineau & Meighen, 1976) by using linear-least-squares methods:

1=1+[H*]

kapp (kapp )max (kapp )max Ks

 

 

Inactivation by IAM. Purified urease was incubated in 1 mM EDTA, 50
mM HEPES, pH 7.75 buffer (or other buffers and pH values as described) plus
the indicated concentrations of IAM at 37°C. Aliquots were taken at 30-min
intervals and assayed for urease activity as previously described (Todd &

Hausinger, 1991 a).

RESULTS

Initial characterization of mutant proteins. Escherichia coli cells

containing the K. aerogenes urease genes on plasmid pKAU17 or derivative

51
plasmids with His—>Ala substitutions were grown under culture conditions that

led to high level synthesis of the wild-type and mutant ureases. The wild-type
and mutant proteins were highly purified (Figure 1) and the specific activities,
Km values, and nickel contents were determined (Table 1). The yH96A, BH39A,
BH41A, 0tH312A and 0tH321A proteins have Km values, specific activities, and
nickel contents that are similar to the wild-type enzyme, indicating that these
histidine residues are not likely to be important for substrate binding, catalysis,
or nickel ligation. In contrast, the other five mutant proteins exhibit significant
changes in their properties, consistent with important roles for the aH134, 0:
H136, aH219, otH246, and otH320 residues. These mutant proteins fall into
three classes, each of which is described separately below.

The 0tH134A, 0tH136A and aH246A proteins are inactive and possess
approximately 50% of the normal metal content (aH134A) or the near absence of
nickel (aH136A and aH246A). These results are compatible with aH134, aH136
and aH246 functioning as nickel ligands in the enzyme. It remains unclear
whether the latter two residues may bridge the two metal atoms at the active site
so that neither nickel can bind in the absence of these double ligands, or if the
increased lability of one active site nickel atom by loss of a ligand leads to
lability of the second metal ion. Similarly, it is unknown whether the 50% nickel
content of the 0tH134A protein represents a case where one nickel is
incorporated into each catalytic unit randomly between the two nickel sites or
whether one of the two nickel sites is filled while the other remains unoccupied
due to the loss of an essential ligand.

The otH219A protein exhibits a very high Km value for urea coupled with a
large decrease in specific activity (although the enzyme activity could not be
assayed under saturating conditions, the calculated Vmax value was about 3%
of that found in the wild-type protein). The altered properties of the aH219A
protein apparently are not due to changes in nickel content because, at most,

20% of the nickel was lost in this mutant protein compared to wild-type enzyme.

52

1234 56 789101112
*

' L" 5.00.09...-

‘hff -

 

Figure. 1. Denaturing gel electrophoretic analysis of partially purified urease
proteins. Samples were run on a 10-15% polyacrylamide gradient gel and stained
with Coomassie brilliant blue. The percent purity for each sample was assessed by
using an AMBIS gel scanner. Lane 1: molecular weight markers (phosphorylase b,
M, of 92,500; bovine serum albumine, M, of 66,200; ovalumin, M, of 45,000;
carbonic anhydrase, M, of 31,000; soybean trypsin inhibitor, M, of 21 ,500; and
lysozyme, M, of 14,400). The protein designations and percent purities for the
samples are indicated for lane 2: yH96A (72%), lane 3: BH39A (96%), lane 4: B
H41A (90%), lane 5: 0tH134A (70%), lane 6: aH136A (80%), lane 7: 0tH219A (93%).
lane 8: aH246A (52%), lane 9: aH312A (>98%), lane 10: aH320A (>98%), lane 11:
0tH321A (98%), and lane 12: Wild-type (88%).

53

Table 1. Characteristics of wild-type and mutant Klebsiella aerogenes ureasesa

 

 

Km Specific activity Nickel content
Urease (mM) (U/mg) (%) (tit/catalytic unit) (%)
Wild-type 23:02 1900 100 2.1 100
yH96A 19:02 1700 90 2 95
BH39A 1.5102 1500 - 79 2.1 100
8H41A 14:02 1300 68 2.5 120
aH‘I34A - <0001 - 1.1 53
0H136A - <0001 - 0.13 5
0H219A 1 1001-4019 3512.0 1.9 1.7 80
01H246A - <0001 - 0.44 21
aH312A 16:02 1800 95 2.2 105
cH320A 8.3:0.2d 0051‘»? 0.0027 2.3 110
0H321A 20:02 1700 90 2.4 114

 

a Determined for enzyme purified by DEAE-Sepharose and Mono-Q column
chromatographies as illustrated in Fig. 1, except where indicated.
b Determined by using enzyme purified only with DEAE-Sepharose column
chromatography. .
C Determined by using 1 M urea instead of the standard assay.
d Determined for cell extract.

9 Determined by using pH 6.75 buffer instead of the normal 7.75 buffer.

54

Rather, the high Km value of the 0H219A protein suggests that the aH219
residue is somehow important to substrate binding. For example, the histidine
residue may facilitate urea binding by forming a hydrogen-bond with the
substrate. Alternatively, however, the results are compatible with a model in
which the imidazole group simply props open the substrate binding site and
maintains its accessibility.

The otH320A protein was found to have a very significant decrease in
specific activity and a moderate increase in Km value compared to wild-type
enzyme. The observed kinetic changes were not correlated to the protein's
nickel content, consistent with 0H320 having a role in urease activity other than

in substrate binding or nickel ligation.

Thermal stabilities of mutant ureases. Of those mutant ureases that
possess activity, all but the 0H312A protein exhibited significant reductions in
thermal stability compared to the wild-type enzyme (Table 2). These results are
consistent with the yH96, BH41, BH39, aH219, aH320, and 0H321 residues
participating in ion-pair or hydrogen-bond interactions in the native protein.
Loss of these interactions leads to protein destabilization at temperatures 2
50°C but not at the growth temperature of 37°C. The reason for the enhanced

stability of the 0H312A mutant protein over wild-type enzyme is unclear.

pH dependence of mutant urease activities. The pH optimum of the
wild-type enzyme (Figure 2A) at around pH 7.75 is retained in the active mutant
proteins yH96A, BH39A, BH41A, aH219A, 0H312A, and 0H321A, as illustrated
for the 0H219A protein (Figure 28). The results are consistent with the wild-type
general base and general acid groups being retained in these mutant proteins.

In contrast, the 0H320A protein exhibited a significantly shifted pH optimum at

55

Table 2. Thermal stabilites of wild-type and mutant ureasesa

 

 

Incubafion
50°C 60°C 70°C
Urease 1 hr 2 hr 1 hr 2 hr 1 hr 2 hr
VVild-type 79 70 57 45 13 3
yH96A 74 61 33 12 <1 <1
BH39A 66 53 9 <1 <1 <1
BH41A 64 51 45 21 <1 <1
0tH219A° 83 72 43 18 <1 <1
0H312A 80 76 85 70 42 22
0H320A° 85 74 8 <2 <3 <3
0tH321A 65 58 30 12 <1 <1

 

a Values are expressed as percent of that for control samples that were not
subjected to high temperature incubation. In each case, cell extracts were
incubated in 20 mM potassium phosphate buffer (pH 7.0) containing 1 mM
EDTA and 1 mM B-mercaptoethanal at the indicated temperatures for up to 2
hr and the activities were measured by using the standard assay conditions,
except where indicated.

b Activities for the 0H219A protein were measured by using a concentration of
100 mM urea.

‘3 Assays for the 0H320A protein were carried out at pH 6.75.

56

 

 

 

  

 

 

 

400 I l I
A
300 - WT —
O
200 - ' -
A 100 - V 4
O
E O I I I
51 B _ aH219A
g 9 _ o 1
.3 5 - . V 4
it
.9 3 " ‘
fig 0 I I I
m C aH320A
0.002 - —
0.001 — -
0.000
4 8 8 10 12

Figure 2. The pH dependence of urease activity. Analysis using A: Cell extracts
of E. coli DH5[pKAU17], B: Purified 0H219A enzyme, C: Cell extracts containing
the 0H320A mutant protein. The reaction mixtures contained urea (50 mM for
wild-type enzyme and H320A protein or 1 M for the 0H219A protein), 0.5 mM
EDTA and the following buffers at a concentration of 25 mM: ME8(O), HEPES
(O), CHES (V), and CAPS (V).

57

around 6.75 (Figure 2C). Hence, one of the residues serving as a general base
or general acid may have been mutated in the 0H320A protein or the mutation

may have indirectly shifted one of these pKa values.

Inactivation of mutant ureases by DEP. The properties of the 0H219A
and aH320A mutant proteins were consistent with these two histidine residues
playing important roles in the enzyme other than metallocenter ligation.
Because chemical modification studies of urease using DEP had provided
evidence for an essential histidine residue acting as a general base in the native
enzyme (Park & Hausinger, 1993), the DEP reactivities of these two mutant
proteins were assessed. While the wild-type enzyme was rapidly inactivated by
50 11M DEP (Figure 3), both the aH219A and 0H320A proteins appeared to
retain full enzyme activity after the same treatment (data not shown). At 1 mM
DEP, however, the aH219A protein was inactivated in a pseudo-first-order
process, whereas the 0H320A protein continued to be resistant to inactivation by
DEP. Further characterization of DEP inactivation of the 0H219A protein
demonstrated that the rate was pH-dependent (Figure 4) with the pattern for
inactivation of the 0H219A (pKa=6.8) and wild-type (pKa=6.5) proteins being
nearly identical. These data are inconsistent with 0H219 serving as the general
base for catalysis in the wild-type enzyme. Rather, the reduced level of DEP
reactivity observed for the aH219A protein may be due to the same features that
account for the high Km value of this enzyme. The structure of DEP shares
some similarity to that of urea, hence, the binding of this reagent to the active
site correspondingly may be reduced in affinity. Alternatively, if aH219 is
important for maintaining access to the catalytic site the reduced reactivity in the
mutant protein may derive from partial closure of this region. In contrast to the

DEP reactivity of the 0H219A protein, the resistance to inactivation by DEP

58

0.5 1 I r

 

 

Ln vt/vo

 

 

 

 

—2.0 ‘ 1
0.0 0.4 0.8 1.2

Time (min)

Figure 3. Kinetics of urease inactivation by DEP. Purified wild-type (0), 0t
H219A (V), and aH320A (0) proteins were treated with DEP (50 11M, 1 mM,
and 1 mM, respectively) in 1 mM EDTA and 50 mM HEPES (pH 7.0) buffer. The
natural logarithm of Vt/Vo (where Vt is the velocity at time tand V0 is the initial

velocity) is shown as a function of time.

59

 

 

 

 

 

 

 

1600 40
1200 4— -_ 30
’3
s ’-.:
5 800 ~~ -- 20 g
5 e:
.2
400 —~ -— 10
0 0
4 6 8 10

 

log kapp

 

 

 

 

Figure 4. The pH dependence of DEP inactivation of wild-type and 0H219A
ureases. A: Apparent second-order rate constants (M’1SBC'1) for urease
inactivation with DEP (left axis for wild-type enzyme and right axis for the
aH219A enzyme) are plotted as a function of pH. 8: The same data plotted as
log kapp versus pH. The reactions were carried out in 1 mM EDTA and 50 mM
concentrations of the following buffers: MES (O), MOPS (O), HEPES (A),
Tricine (D) or TAPS (A) for wild-type and MOPS (I) for H219A urease.

60

displayed by the 0H320A protein is compatible with 0H320 serving as the target

of DEP in the native enzyme; i.e., the general base that facilitates catalysis.

Inactivation of mutant proteins by IAM. To further explore the roles of
the 01H21 9 and 0H320 residues, the chemical reactivities of the mutant proteins
towards IAM were assessed. This reagent had been used previously (Todd &
Hausinger, 1991 a) to provide evidence that a thiol group is ionically coupled to
another residue, X, together acting as the proton donor in catalysis. Both the a
H219A and aH320A proteins were inactivated by incubation with IAM (Figure 5).
A much higher concentration of the alkylating reagent is required for inactivation
of the 0H219A urease than for wild-type enzyme or the 0H320A protein.
Because IAM structurally resembles urea, part of this effect may arise from
decreased affinity of the reagent for the active site (prior to reaction with the
cysteine residue) due to the same features that lead to the high Km value in this
protein. Again, however, the results are compatible with a model in which the
active site is simply less accessible in the aH320A protein. The pH dependence
of otH320A inactivation by IAM could not be examined because of the low activity
of the protein. In contrast, the pH dependence for aH219A protein was
measured and shown to be similar to that in the wild-type enzyme (Figure 6).
This result would not be expected if the aH219 residue was equivalent to X in
the Cys-X ion pair that has been proposed to occur in this protein (Todd &
Hausinger, 1991 a).

61

 

 

 

 

 

 

 

_4 1 1 1 T

 

0 30 6O 90 120

Time (min)

Figure 5. Kinetics of urease inactivation by IAM. Purified wild type (C), 0H219A
(V), and aH320A (0) proteins were treated with 40 mM IAM and purified or
H219A (V) protein was treated with 400 mM IAM in 1 mM EDTA and 50 mM
HEPES (pH 7.75) buffer. The natural logarithm of thV0 (where Vt is the velocity

at time tand V0 is the initial velocity) is shown as a function of time.

62

 

 

 

 

3 T I T I
2_ 00 _
8
1- _
a.
8 o- -
.14
m_1_ _
2 1:1 A
_2- I _
_3_ _
_4 1 1 1 1
4 6 8 10 12 14
pH

Figure 6. The pH dependence of IAM inactivation of wild-type and aH219A
ureases. Apparent second-order rate constants (M-1sec-1) for urease
inactivation with IAM (20 mM for wild type enzyme and 120 mM for the aH219A
enzyme) are plotted as a function of pH. The reactions were carried out either in
1 mM EDTA and 80 mM of acetate (1:1), MES (I), HEPES (A), CHES (A), CAPS
(<>), and phosphate (0) buffers for wild-type enzyme or in 1 mM EDTA and 50
mM HEPES (O) and CHES (O) buffers for the 0H219A urease.

63
DISCUSSION

Dixon et al. (1980) proposed an elegant model for the hydrolysis of urea by the
bi-nickel active site of jack bean (Canavalia ensifarmis) urease that serves as an
excellent framework for discussion of K. aerogenes urease site-directed
mutagenesis. In their model, one nickel ion is suggested to coordinate a water
molecule and a second nickel ion coordinates hydroxide ion. Urea is proposed
to displace the water molecule and bind in O-coordination to nickel as the Ni--'O-
C(-NH2)=NH2EB resonance structure with electrostatic stabilization by a nearby
carboxyl group. A general base is hypothesized to activate the nickel-
coordinated hydroxyl group that carries out a nucleophilic attack on the urea
carbon. The resulting tetrahedral intermediate is thought to decompose to form
carbamate and ammonia with the participation of a nearby thiol group acting as
a general acid. Subsequently, carbamate dissociates and spontaneously is
converted to carbon dioxide and a second molecule of ammonia. According to
the Dixon model, two nickel ions, a carboxyl group, a general base and a
general acid are required for the jack bean enzyme activity. In independent
studies with jack bean urease, Sakaguchi et al. (1983) provided evidence from
photo-oxidation studies of the enzyme in the presence of methylene blue and
active site-directed inhibitors that histidine residues play essential roles at the
catalytic site. Furthermore, Takishima et al. (1988) identified the reactive
cysteine residue in the enzyme and found it to be located in a region that was
rich in histidine residues.

In studies with K. aerogenes urease, known to be ~60% identical in
sequence to the jack bean enzyme (Mulrooney & Hausinger, 1990), the Dixon
model has undergone further elaboration. The chemical reactivity of the general

base (pKaES.5) that appears to facilitate catalysis (Todd & Hausinger, 1987) was

64

shown to be compatible with that of a histidyl group (Park & Hausinger, 1993).
The thiol group that participates in proton donation was identified (Todd &
Hausinger, 1991b; Martin & Hausinger, 1992) and shown to function as an ion
pair with another residue (X) that could reasonably be accounted for by a
histidine group (Todd & Hausinger, 1991 a). Finally, the enhanced DEP
reactivity of apoprotein over holoenzyme (Lee et al., 1990) is consistent with at
least partial metallocenter ligation by histidyl residues. The multiple potential
roles for histidine groups in the protein led to'the experiments described above.
My results are consistent with the participation of three histidyl residues
(aH134, aH136, and otH246) in nickel coordination, one histidine residue (0
H320) serving as the general base in catalysis, and one residue (aH219)
somehow facilitating substrate binding. The latter residue may stabilize urea
binding by hydrogen—bond formation [similar to the ionic stabilization of bound
urea by a postulated carboxyl group in the model of Dixon et al. (1980)], or
perhaps 0H219 may act by maintaining accessibility to the catalytic site. No
evidence was obtained for the presence of a conserved histidine acting as
residue X in a Cys-X pair that functions as a general acid. Further efforts toward
elucidating the three—dimensional structure of K. aerogenes urease by x-ray
crystallographic methods (of, Jabri et al., 1992) will establish the validity of

several of the roles for histidine residues that have been proposed here.

65

References

Blanchard, A. (1990). Ureaplasma urea/yticum urease genes; use of a UGA
tryptophan codon. Molec. Microbial. 4, 669-679.

Brockelhurst, K (1987). Acyl group transfer-cysteine proteinases. In Enzyme
Mechanisms (Page, M. l. 8 Williams, A., Eds), pp. 140-158. Royal
Society of Chemistry, Burlington House, London.

Clayton, C. L., Pallen, M. J., Kleanthous, J., Wren, B. W. & Tabaqchali, S.
(1990). Nucleotide sequence of two genes from Helicobacter pylori
encoding for urease subunits. Nucleic Acids Res. 18, 362.

Cousineau, J. & Meighen, E. (1976). Chemical modification of bacterial
luciferase with ethoxyfomiic anhydride: evidence for an essential
histidyl residue. Biochemistry 15, 4992-5000.

Dixon, N. E., Riddles, P. W., Gazzola, C., Blakeley, R. L. & Zemer, B. (1980).
Jack bean urease (EC 3.5.1.5). V. On the mechanism of action of
urease on urea, formamide, acetamide, N-methylurea, and related
compounds. Can. J. Biochem. 58, 1335-1344.

Jabri, E., Lee, M. H., Hausinger, R. P. & Karplus, P. A. (1992). Preliminary
crystallographic studies of urease from jack bean and from Klebsiella
aerogenes urease. J. Mol. Biol. 227, 934-937.

Jones, 8. D., & Mobley, H. L. T. (1989). Proteus mirabilis urease: Nucleotide
sequence determination and comparison with jack bean urease. J.
Bacteriol. 171, 6414—6422.

Kunkel, T. A., Roberts, J. D. 8. Zakour, R. A. (1987). Rapid and efficient site-
specific mutagenesis without phenotypic selection. Methods Enzymol.

154, 367-382.

66

Labigne. A., Cussac, V., 8. Courcovx, P. (1991). Shuttle cloning and nucleotide
sequences of Helicobacter pylori genes responsible for urease activity.
J. Bacteriol. 173, 1920-1931.

Laemmli, U. K (1970). Cleavage of structural proteins during assembly of the
head of bacteriophage T4. Nature 227, 680-685.

Lee, M. H., Mulrooney, S. B., & Hausinger, R. P. (1990). Purification,
characterization, and in viva reconstitution of Klebsiella aerogenes
urease apoenzyme. J. Bacteriol. 172, 4427-4431.

Lee, M. H., Mulrooney, S. B., Renner, M. J., Markowicz, Y., & Hausinger, R. P.
(1992). Klebsiella aerogenes urease gene cluster: sequence of ureD
and demonstration that four accessory genes (ureD, ureE, ureF, and
ureG) are involved in nickel metallocenter biosynthesis. J. Bacterial.

1 74, 4324-4330.

Lowry, O. H., Rosebrough, N. J., Farr, A. L., & Randall, R. J. (1951). Protein
measurement with the Folin phenol reagent. J. Biol. Chem. 193, 265-
275.

Martin, P. R. & Hausinger, R. P. (1992). Site-directed mutagenesis of the active
site cysteine in Klebsiella aerogenes urease. J. Biol. Chem. 267,
20024-20027.

Miles, E. W. (1977). Modification of histidinyl residues in proteins by

diethylpyrocarbonate. Methods Enzymol. 47, 431-442.

Mobley, H.L.T., 8. Hausinger, R. P. (1989). Microbial urease: significance,
regulation, and molecular characterization. Microbial. Rev. 53, 85-108.

M0rsdorf, G, & Kaltwasser, H. (1990). Cloning of the genes encoding urease
from Proteus vulgaris and sequencing of the structural genes. FEMS

Microbial. Lett. 66, 67-74.

67

Mulrooney, S. B., 8 Hausinger, R. P. (1990) Sequence of the Klebsiella
aerogenes urease genes and evidence for accessory proteins
facilitating nickel incorporation. J. Bacteriol. 172, 5837-5843.

Mulrooney, S. B., Pankratz, H. 8., 8 Hausinger, R. P. (1989). Regulation of gene
expression and cellular localization of cloned Klebsiella aerogenes (K.
pneuminae) urease. J. Gen. Microbiol. 135, 1769-1776.

Park, IS, 8 Hausinger, R. P. (1993). Diethylpyrocarbonate reactivity of
Klebsiella aerogenes urease: effect of pH and active site ligands on
rate of enzyme inactivation. J. Prat. Chem. 12, 51-56.

Sakaguchi, K, Mitsui, K, Kubashi, K. 8 Hase, J. (1983). Photo-oxidation of jack
bean urease in the presence of methylene blue. J. Biochem. 93, 681-
686.

Sambrook, J., Fritsch, E. F ., and Maniatis, T. (1989). Molecular Cloning: a
Laboratory Manual. Cold Spring Harbor Laboratory, Cold Spring
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Takishima, K, Suga, T., 8 Mamiya, G. (1988). The structure of jack bean
urease. The complete amino acid sequence, limited proteolysis and
reactive cysteine residues. Eur. J. Biochem. 175, 151-165.

Todd, M. J. 8 Hausinger, R. P. (1987). Purification and characterization of the
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Todd, M. J. 8 Hausinger, R. P. (1989). Competitive inhibitors of Klebsiella
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site. J. Biol. Chem. 264, 15835-15842.

68

Todd, M. J. 8 Hausinger, R. P. (1991 a). Reactivity the essential thiol of
Klebsiella aerogenes urease. J. Biol. Chem. 266, 10260-10267.

Todd, M. J. 8 Hausinger, R. P. (1991b). Identification of the essential cysteine
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324-332.

CHAPTER 4

In Vitro Activation of Urease Apoprotein and Role of UreD as a Chaperone
Required for Nickel Metallocenter Assembly

These studies, less those shown in Fig. 1 and combined with urease activation
studies of cell extracts by Mary Beth Carr, were published in
Proc. Natl. Acad. Sci. USA 80, 396-3965, 1994.

69

70
ABSTRACT

The formation of active urease in Klebsiella aerogenes requires the presence of
three structural genes for the apoprotein (ureA, ureB, and ureC), as well as four
accessory genes (ureD, ureE, ureF, and ureG) that are involved in functional
assembly of the metallocenter in this nickel-containing enzyme. Partial activation
of urease apoprotein was observed after adding nickel ion to extracts of
Escherichia coli cells bearing a plasmid containing the K. aerogenes urease
gene cluster or derivatives of this plasmid with deletions in ureE, ureF, and
ureG. In contrast, extracts of cells containing a ureD deletion derivative failed to
generate active urease, thus highlighting a key role for UreD in the metallocenter
assembly process. Site-directed mutagenesis methods were used to
overexpress ureD in the presence of the other urease genes, and UreD was
found to co-purify with urease. A molecule of native urease apoprotein is
capable of binding 0, 1, 2, or 3 molecules of UreD, consistent with a trimeric
structure of urease catalytic units. The UreD-urease apoprotein complexes are
competent for activation by nickel, with the level of activity obtained being
directly related to the number of UreD bound per urease molecule. Activation of
the UreD-urease complexes is rapid and accompanied by UreD dissociation. I
propose that UreD is a chaperone protein which stabilizes a urease apoprotein
conformation that is competent for nickel incorporation.

71
INTRODUCTION

Urease (EC 3.5.1.5) is a nickel-containing enzyme that catalyzes the hydrolysis
of urea to form carbonic acid and two molecules of ammonia. Until recently, the
best characterized urease was that from jack bean (1 ). This enzyme was the first
ever crystallized (2) and the first shown to possess nickel (3). However, because
of the ease of cell growth and genetic manipulation in bacteria compared to
plants and the medical importance of bacterial urease, our understanding of the
procaryotic enzyme has outpaced knowledge of the plant enzyme (4).
lmportantly, the active site residues of plant and bacterial ureases are highly
conserved and both enzymes possess a novel bi-nickel metallocenter.

Assembly of the urease bi-nickel metallocenter appears to be a complex
process requiring the action of several accessory gene products. For example,
deletion analysis of the Klebsiella aerogenes urease gene cluster revealed that
one gene (ureD) located directly upstream and three genes (ureE, ureF, and
ureG) found directly downstream of the three genes (ureA, ureB, and ureC)
encoding the subunits of the bacterial urease are involved in the functional
assembly of the urease metallocenter (5, 6). Homologs of the accessory genes
have been found in several bacterial species (39., 7-9) and an essential role for
auxiliary genes in forming active urease has been confirmed in these and many
other bacteria. Genetic studies are consistent with a similar requirement for
urease accessory genes in the fungi Neurospora crassa (10),
Schizosaccharomyces pombe (11), and Aspergillus nidulans (12, 13). Finally,
two loci (Eu2 and Eu3) that appear to encode genes associated with urease
maturation factors such as nickel emplacement have been identified in soybean
plants (14). Thus, accessory genes involved in metallocenter'assembly may be a
universal component of urease activation.

The detailed roles for the urease accessory genes are unknown. The histidine-
rich K. aerogenes UreE protein has been purified and shown to reversibly bind
six nickel ions per dimer (15); hence, it is reasonable to suggest that UreE

72 .
serves as the nickel donor to urease apoprotein. UreG contains a P-loop motif

that is found in many nucleotide-binding proteins (16); thus, this protein may be
involved in coupling nucleotide hydrolysis to the assembly process. In this
regard, in viva evidence is consistent with an energy-dependence for urease
activation (17). Furthermore, the sequence of K. aerogenes UreG was shown
(18) to be approximately 25% identical to that of the Escherichia coli hypB gene
product (19). The hyp operon encodes a hydrogenase pleiotropic operon that is
required for functional activation of hydrogenase. A mutation in hypB was shown
to be complemented, in part, by the addition of high levels of nickel (20),
consistent with a role for this protein in nickel processing. Of potential
significance to the function of UreG, purified HypB protein has been shown to
bind guanine nucleotides and to hydrolyze GTP (21). In contrast to UreE and
UreG, the sequences for UreD or UreF offer no insight into their function and
these proteins have not been purified or characterized.

Here I demonstrate the ability to activate urease apoprotein in vitro and present
evidence supporting a model in which UreD serves as a urease-specific
chaperone during this process.

MATERIALS AND METHODS

Site-Directed Mutagenesis of the ureD Ribosome Binding Site. In order to
enhance expression of ureD, the ribosome binding site and initiation codon of
this gene were changed from QAQGGCACCQTG to QAQGGCACCATG by two
rounds of site-directed mutagenesis. A 0.82—kb EcoRl-Sacl fragment of plasmid
pKAU17 containing a urease gene cluster from K. aerogenes (22) was
subcloned into M13 mp18 and mutagenized by the method of Kunkel et al. (23).
Uracil-containing single-stranded template DNA was prepared from Escherichia
coli CJ236 (dut1 ungi thi-1 relA1IpCJ105[cam'F']). Using a
CACGGTGCCQTQCAATGTTGC oligonucleotide primer, the phage DNA was

73 .
mutagenized and mutant phage were isolated in E. coli MV1193([IacI-proAB]

rpsL thi endA spcB15 hst4 [srl-recA]306::Tn10[tet'] F'[traD36 praAB+
laclqlacZM15l). Uracil-containing single-stranded DNA was again prepared and
similarly mutagenized with the primer GTGGTAACAIGGTGCCCTC. The site-
directed mutants were identified by sequence analysis using Sequenase 2.0
(US. Biochemical Corp, Cleveland OH) and the single-strand DNA sequence
method of Sanger et al. (24). The region was completely sequenced to ensure
that no other mutations had been introduced into M13. The mutated EcaRl-Sacl
region was subcloned back into pKAU17 to generate pKAUDZ and the sequence
was confirmed by double-stranded DNA sequencing methods (25).

Cell Growth and Disruption. E. coli DH5 carrying pKAUDZ, pKAU17,
pKAU22AD-1 (ureD deletion derivative of pKAUZ2, ref. 6) or pKAUDABC
(ureEFG deletion derivatives of pKAU17, see appendix) were grown in LB
medium to stationary phase. The harvested cultures were resuspended in 20
mM phosphate (pH 7.0), 0.1 mM EDTA, and 0.1 mM 2-mercaptoethanol buffer
and disrupted by three passages through a French pressure cell (American
Instrument Co., Silver Spring MD) at 18,000 Ib/inz. Cell extracts were obtained
after centrifugation (100,000 x g for 60 min) at 4°C. The ability to activate urease
apoenzyme in E. coli [pKAUD2] extracts was stable for over 1 month when
samples were stored on ice and adjusted to 20% glycerol and 0.8 mM
dithiothreitol.

Purification of UreD-Urease Apoprotein Complex. E. coli DH5[pKAUD2] cell
extracts from a 2 L culture were chromatographed on a column (2.5 x 20 cm) of
DEAE-Sepharose at 4°C in PEG buffer [16 mM phosphate (pH 7.0), 0.8 mM
EDTA, 20% glycerol, 0.8 mM 2-mercaptoethanol, 0.8 mM dithiothreitol]. The
proteins were eluted in PEG buffer with a linear salt gradient to 1.0 M KCI, and
UreD was found to co-elute with urease apoprotein at ~0.45 M KCI. UreD-
containing fractions were pooled, dialyzed against PEG buffer, and applied to a
Mono-Q HR 10/10 column equilibrated in the same buffer. A linear salt gradient
to 1 M KCl in PEG buffer again eluted the UreD in the same fractions as urease.

74 -
A portion of the pooled fractions containing UreD was adjusted to 1 M KCI by

addition of PEG buffer containing 2 M KCI and applied to a phenyl-Superose HR
515 column equilibrated in this buffer. A linear gradient to 0 M KCI in PEG buffer
was used for elution. Samples were monitored for the presence of UreD and
urease apoprotein by using polyacrylamide gel electrophoresis.

Polyacrylamide Gel Electrophoresis. SDS-polyacrylamide gel
electrophoresis was carried out by using the buffers of Laemmli (26) and
included a 13.5% polyacrylamide running gel and a 4.5% polyacrylamide
stacking gel. Nondenaturing gels utilized the same buffers without SDS and
used 7.5% and 3% acrylamide running and stacking gels. Gels were stained with
Coomassie brilliant blue or, in the case of selected native gels, blotted onto
nitrocellulose, probed with anti-K. aerogenes urease antibodies (22), and
developed by using anti-rabbit lgG-alkaline phosphatase conjugates (27). Band
intensities were measured by using a gel scanner (Ambis, Inc.). For calculation
of the ratio of UreD to UreC, M, values of 30,000 and 60,300 were used for these
pepfides.

Urease Activity Assays. Urease activities were assayed in 25 mM Hepes (pH
7.75) and 0.5 mM EDTA buffer containing 50 mM urea. Linear regression
analysis of the released ammonia, determined by conversion to indophenol (28),
versus time yielded initial rates. One unit of activity is defined as the amount of
enzyme required to degrade 1 mmol urea per min at 37°C. Protein
concentrations were assessed by the Spectrophotometric assay of Lowry et al.
(29) or, in the case of samples containing Hepes buffer, the Bio-Rad protein
assay (Bio-Rad Laboratories, Richmond CA).

RESULTS

In Vitro Activation of Urease Apoprotein. E. coli [pKAU17] carries the wild-

type K. aerogenes urease gene cluster. The urease apoprotein in extracts from

75 .
these cells (grown in the presence of nickel-free medium) is partially activated by

incubation with NiCIz at 20~500 11M (Fig. 1). Although in viva activation of urease
apoprotein has been described [e.g., (17)], data obtained by Mary Beth Carr of
this lab (not shown) and that shown in Fig. 1 represent the first evidence for in
vitro activation of this enzyme. The highest specific activity values observed in
these studies (40 U/mg protein) account for activation of only about 10% of the
urease that is present [i.e., when grown in the presence of 1 mM NICI2 E. coli
[pKAU17] cell extracts typically possess specific activity values of 300-400 U/mg
(17, 30)]. The leveling off of activity is not due to simultaneous formation and
inactivation of active enzyme because purified urease holoprotein is completely
stable under the incubation conditions used for activation of the cell extracts.
Alteration of the activation conditions (varying the pH, buffer composition, ionic
strength, and NICI2 concentration) failed to further improve the extent of
activation and the process is hindered by elevated nickel concentrations (4 mM)
and high (> 8.0) or low (< 7.0) pH.

Analogous urease apoprotein activation experiments were carried out using
extracts from cells that were defective in ureD or ureEFG (Fig. 1). Surprisingly,
deletion of ureEFG (Fig. 1) has no significant effect on the in vitro activation
process. This finding is intriguing because AureF or AureG cultures possess no
detectable urease activity even when provided with 1 mM nickel ion in the
growth medium (6). Of greatest significance, urease apoprotein in cell extracts
from a AureD mutant was not competent for activation under these activation
conditions. These results highlight a key role for UreD in the urease activation 1
process.

Purification of UreD-Urease Apoprotein Complex. Characterization of UreD
fiom E. coli [pKAU17] is hampered by very low level expression of the ureD gene
(6). We circumvented this problem by using site-directed mutagenesis to
improve the ribosome binding site and to provide an ATG initiation codon for this ,
gene in plasmid pKAUD2. An intense band of UreD is observed on SDS-
polyacrylamide gels for extracts of E. coli [pKAUDZ] (Fig. 2, lane 2) that is not

76

 

 

 

 

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8 A

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0 i=5" ' ~ t 3F

0 30 60 90 120
Time (min)

Fig. 1. In vitro activation kinetics of urease apoprotein. Cell extracts (2 mglml)
from E. coli [pKAU17] (closed symbols) or AureD (x), AureEFG (open symbols)
deletion derivatives were incubated in 25 mM HEPES (pH 7.9)10.08 mM EDTA!
0.08 mM 2-mercaptoethanol containing 20 (square), 50 (triangle), 100
(diamond), or 500 (circle, x) uM NiClz at 37°C. Aliquots were withdrawn and
assayed for urease activity and protein.

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78 .
present in extracts of E. coli [pKAU17] (not shown). Using extracts from E. coli

[pKAUD2] cells that were grown in the absence of nickel, UreD was found to co-
elute with urease during two steps of ion exchange chromatography and during
hydrophobic interaction chromatography on phenyl-Superose HR 5/5. The latter
step resolves the UreD/urease species into four pools that were termed flow-
through (FT) and fractions 1, 2, and 3 (Fig. 3). UreD and urease apoprotein are
unlikely to be present as a complex in the FT fraction because these proteins
are resolved using phenyI-Sepharose CL-4B chromatography (not shown). In
contrast, fractions 1, 2, and 3 do appear to represent distinct UreD-urease
apoprotein complexes.

The ratio of UreD to urease apoprotein in the phenyI-Superose fractions is not
uniform as demonstrated by SDS-polyacrylamide gel analysis (Fig. 2, lanes 6-8).
Given that urease is a trimer of catalytic units (P. A. Karplus, personal
communication), our gel scanning results are consistent with the presence of
1.0-1.3, 2.2-2.5, and 3-3.2 molecules of UreD per molecule of native urease in
fractions 1, 2, and 3. Native gel electrophoresis of the four phenyl-Superose
pools (Fig. 4, lanes 4-7) clearly demonstrates that four distinct urease
apoprotein species are present. The FT fraction is comprised almost entirely of
form I, a species that migrates identically with urease apoprotein or urease
holoprotein (Fig. 4, lane 8). Fractions 1, 2, and 3, in contrast, are comprised
predominantly of urease apoprotein species of decreasing mobilities that are
labeled forms II, III, and IV (Fig. 4, lanes 5-7). Form Ill appears to predominate in
the DEAE-Sepharose and Mono—Q pools (Fig. 4, lanes 2 and 3), and is the major
fraction found on the phenyl-Superose column (Fig. 3). Mobility differences
among the four urease species on native gels are likely to arise primarily from
differences in native size. Gel filtration chromatography results are consistent
with this hypothesis; when the Mono-Q pool was chromatographed on a
Superose 6 column and the fractions analyzed by native gel electrophoresis, the
overlapping order of elution was form IV, form III, form II, and form I (data not

shown). Thus, we conclude that the four species observed upon phenyl-

79

 

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Fig. 3. Phenyl-Superose resolution of UreD-urease complexes. Starting with
extracts from E. coli [pKAUD2] cells that overexpress ureD, UreD was found to
co-elute with urease apoprotein during chromatography on DEAE-Sepharose
and Mono-Q resins. The latter pool was resolved into four fractions by
chromatography on phenyl-Superose. Elution was monitored by absorbance at
280 nm, and the pooled samples are denoted flow-through (FT) and fractions 1,
2, and 3.

80

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81 .
Superose chromatography and resolved by native gel electrophoresis arise from

the presence of different numbers of UreD (0, 1, 2, or 3) complexed to the native
urease enzyme.

Activation of UreD-Urease Apoprotein Complexes. When adjusted to 0.1
mM NICIz, urease apoprotein is activated to different extents in each of the four
phenyI-Superose pools (Fig. 5). Importantly, the greater the ratio of UreD to
urease apoprotein, the greater is the final specific activity of the preparation. The
small amounts of activation observed in the FT fraction are likely to arise from
trace contamination with the form Il species that contain UreD. The activation
process for the UreD-urease complexes is rapid. The highest level of activity
reached (~550 Ulmg protein) accounts for approximately 30% of full activation
for the urease apoprotein that is present in the predominantly form IV species
[assuming one UreD per catalytic unit (M, of 113,000) and a specific activity of
2,500 Ulmg protein for fully active urease (6)]. This level of activation is even
more impressive when one considers that full activation of urease is not
observed in E. coli [pKAU17] cells grown in the presence of 1 mM NICI2 [with
typical specific activity values of 1,500-1,900 Ulmg (6, 30)]. These results are
consistent with the presence of a UreD-urease complex playing a central role in
urease activation.

Upon treatment with nickel ion, the form II, III, and IV species observed in
native gels collapse to the form I species for each of the phenyl-Superose pools
(Fig. 4, lanes 9-12). These results are consistent with dissociation of UreD
during the activation process. A band corresponding to UreD is not observed in
the native polyacrylamide gels, perhaps due to the inability of the highly
insoluble protein to enter the gel.

Rapid and high level in vitro urease apoprotein activation (to ~200 Ulmg total
protein) similarly was observed in cell extracts of the E. coli [pKAUD2] cells upon
addition of nickel ion. If urease is estimated to account for approximately 20% of
the protein in these cell extracts (see Fig. 2, lane 1), this result allows me to

calculate a specific activity for urease of about 1,000 Ulmg in this sample.

82

 

 

 

 

 

 

 

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a a
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3
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>
:3 300- ~
< J
5% 200:
8
$100 a ~

0 1

0 20 40 60 80 100
Time (min)

Fig. 5. Activation kinetics of the UreD-urease complexes. The four phenyl-
Superose pools [FT (O) and fractions 1 (I), 2 (A), and 3 (7)] from Fig. 3 were
incubated at 37°C in the presence of 0.1 mM NICIz in 25 mM Hepes (pH 7.9),
0.15 M KCI and aliquots were removed and assayed for urease activity over
time.

83 .
Optimal activity was observed from pH 7.6 to 8.2 and using 0.5 mM NICIz.

Phosphate strongly inhibits the activation process at concentrations greater than
20 mM. Surprisingly, growth of E. coli [pKAUD2] in the presence of 1 mM NiCIz
yields cell extract activities that are very low (3-15 Ulmg), equivalent to 1-5% of
the activity found in E. coli [pKAU17] cell extracts. It is possible that this result
may relate, at least in part, to the low level expression of ureG that is observed
in these cells compared to cells containing pKAU17. Addition of nickel to these
low activity cell extracts results in rapid and high level activation, similar to the
extracts from cells obtained by nickel-free conditions.

Using anti-urease antibodies and Western blot analysis of cell extracts
electrphoresed through a native gel, the three UreD-urease complexes from E.
coli [pKAUD2] were shown to collapse almost entirely to the form I (UreD-free
apoprotein or holoprotein) species (Fig. 6, lanes 4-5) after incubation with 0.3
mM nickel. More significantly, the presence of at least two UreD-urease
complexes (forms II and III) was established in E. coli [pKAU17] cells that do not
overexpress ureD. Furthermore, nickel addition to these cell extracts also was
shown to lead to diminishment of the bands corresponding to these complexes
(Fig. 6, lane 2-3).

DISCUSSION

I propose that urease apoprotein activation occurs by the mechanism illustrated
in Fig. 7. [This simplified model does not indicate that two nickel are
incorporated per catalytic unit or that three catalytic units are present per native
enzyme]. UreD is proposed to be a chaperone protein that stabilizes a
conformation of urease that is competent for nickel incorporation. This role is
reminiscent of that suggested for MW in activation of apodinitrogenase by the
iron-molybdenum cofactor (31, 32) and for MelC1 in incorporating copper into
tyrosinase (33, 34). The in vitro source of nickel in the proposed urease

84

IV-

I— “N...- .1»

Fig. 6. Western blot comparison of cell extracts before and after nickel
activation. Samples were activated by incubation for 100 min in 0.3 mM NICIz, 25
mM Hepes (pH 7.9). Lanes: 1, purified urease; 2, cell extracts of E. coli
[pKAU17]; 3, same as 2 after activation; 4, E. coli [pKAUD2] cell extracts; 5,

same as 4 after activation.

85

 

  
 

n-

see-m

Fig. 7. Simplified model illustrating the functional role for UreD in urease

4 x
2

apoprotein activation.

86 .
metallocenter assembly mechanism is NiClz, whereas, nickel bound to UreE may

fill this role in viva (15). The roles of UreF and UreG in the activation process
remain unclear, however, it is reasonable to speculate that they may function in
assembly of the UreD-urease complex (step 1), in facilitating interactions
between the complex and UreE (step 2), by enhancing the level of activation
during UreD release (step 3), or in recycling of UreD (step 4). The energy-
dependence of activation that was observed in vivo (17), but not in vitro (above),
and the low level of activation observed in E. coli [pKAU17] cell extracts can be
interpreted in terms of this model. For example, it is possible that UreD recycling
requires energy, whereas single turnover of the UreD-urease complex does not.
The levels of in vitro activation that we observe are consistent with generation of
active enzyme only from the UreD-urease complexes, with no recycling of UreD.
It is possible that, in the absence of sufficient UreD, partially folded urease that
is newly released from the ribosome undergoes a final conformational change to
bury the nickel binding site. Our in vitro conditions are inadequate to allow the
folded apoprotein to revert to the partially unfolded state that binds UreD and is
capable of activation; however, this species does appear to be slowly activated
in viva (17). This working model may help to focus future efforts to understand

urease metallocenter assembly.

10.

11.
12.
13.
14.

15.

16.

17.

18.

87
REFERENCES

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Dixon, N. E., Gazzola, C., Blakeley, R. L. 8 Zemer, B. (1975) J. Am.
Chem. Soc. 97, 4131 -4133.

Mobley, H. L. T. 8 Hausinger, R. P. (1989) Microbial. Rev. 53, 85-108.
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Cussac, V., Ferrero, R. L. 8 Labigne, A. (1992) J. Bacterial. 174, 2466-
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Jones, 8. D. 8 Mobley, H. O. T. (1989) J. Bacterial. 171, 6414-6422.
Collins, C. M. 8 Gutman, D. M. (1992) J. Bacterial. 174, 883-888.

Benson, E. W. 8 Howe, H. B., Jr. (1978) Molec. Gen. Genet. 165, 277-
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Kinghom, J. R. 8 Fluri, R. (1984) Curr. Genet. 8, 99-105.

Mackay, E. M. 8 Pateman, J. A. (1982) Biochem. Genet. 20, 763-776.
Mackay, E. M. 8 Pateman, J. A. (1980) J. Gen. Microbial. 116, 249-251.
Meyer-Bothling, L. E., Polacco, J. C. 8 Cianzio, 8. R. (1987) Molec. Gen.
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Lee, M. H., Pankratz, H. 8., Wang, 8., Scott, R. A., Finnegan, M. G.,
Johnson, M. K., lppolito, J. A., Christianson, D. W. 8 Hausinger, R. P.
(1 993) Protein Sci. 2, 1042-1052.

Saraste, M., Sibbald, P. R. 8 Wittinghofer, A. (1990) Trends Biochem. Sci.
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Lee, M. H., Mulrooney, S. B. 8 Hausinger, R. P. (1990) J. Bacterial. 172,
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Lutz, A., Jacobi, A., Schlensog, V., Bdhm, R., Sawers, G. 8 Book, A.

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Waugh, R. 8 Boxer, D. H. (1986) Biochimie 68, 157-166.

Maier, T., Jacobi, A., Sauter, M. 8 Beck, A. (1993) J. Bacteriol. 175, 630-
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Mulrooney, S. B., Pankratz, H. 8. 8 Hausinger, R. P. (1989) J. Gen.
Microbial. 135, 1769-1776.

Kunkel, T. A., Roberts, J. D. 8 Zakour, R. A. (1987) Meth. Enzymalogy
154, 387-382. '

Sanger, F., Mickle, S. 8 Coulson, A. R. (1977) Proc. Natl. Acad. Sci. USA
80, 3963-3965.

Sambrook, J., Fritsch, E. F. 8 Maniatis, T. (1989) Molecular Cloning: a
Laboratory Manual (Cold Spring Harbor Laboratory, Cold Spring Harbor,
NY), 2nd Ed.

Laemmli, U. K (1970) Nature (London) 227, 680-685.

Blake, M. 8., Johnston, K. H., Russel-Jones, G. L. 8 Gotschlich, E. C.
(1984) Anal. Biochem. 136, 175-179.

Weatherbum, M. W. (1967) Anal. Chem. 39, 971-974.

Lowry, O. H., Rosebrough, N. J., Farr, A. J. 8 Randall, R. J. (1951) J. Biol.
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Park, IS. 8 Hausinger, R. P. (1993) Protein Sci. 2, 1034-1041.

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Chem. 267, 20100-20207.

CHAPTER 5

Carbon Dioxide is Required for in vitro Assembly of the Urease Nickel
Metallocenter

These studies were submitted to Science.

89

90
ABSTRACT

In vivo assembly of the urease metallocenter is known to require four accessory
proteins: UreD, postulated to be a urease-specific molecular chaperone, UreE, a
Ni(ll)—binding protein, and UreF and UreG whose functions remain obscure. In vitro
activation of purified Klebsiella aerogenes urease apoprotein has now been
accomplished by simply providing CO; (half-maximal activation at ~0.2% C02) in
addition to nickel ion. Activation requires that CO; be incorporated into urease in a
pH-dependent reaction (PK. 2 9). CO; concentration also affects the level of
activation for UreD-urease apoprotein complexes. I propose that CO; binding to
urease apoprotein generates a ligand that facilitates productive nickel binding.

91 ,
Urease, the first enzyme ever crystallized (1) or shown to possess nickel (2), is

found in selected plants, fungi, and bacteria where it plays important roles in
environmental nitrogen transformations and as a virulence factor in certain
pathogenic microbes (3). The protein contains a dinuclear Ni(ll) active site (4) where
each metal atom has a Ni(imidazole),(N,O),-,,, (x = 2 or 3) coordination environment
according to X-ray absorption spectroscopic analysis (5). In viva assembly of this
novel metallocenter in Klebsiella aerogenes (6) involves the participation of four
accessory gene products: UreD, UreE, UreF, and UreG (7). UreD has been
postulated to function as a molecular chaperone that stabilizes a urease apoprotein
conformation which is competent for nickel incorporation (8). Evidence consistent
with this hypothesis includes the ability to purify several forms of a UreD-urease
apoprotein complex [(urease)3UreDN, where N = 1, 2, or 3], the demonstration that
the complexes can be partially activated by addition of nickel ions (increasing levels
of activation correlate to increasing N), and the finding that UreD dissociates from
urease during activation. UreE has been proposed to serve as a nickel donor for
urease activation (9). This suggestion was based on the ability of the protein to bind
approximately six nickel ions per dimer (Kd ~10 M) with rather high specificity. The
roles for UreF and UreG in urease activation are unknown, but they are absolutely
essential for activation in viva. Of related interest, UreG is homologous in sequence
to HypB, a GTP-binding protein that is required for incorporation of nickel into
hydrogenase (10, 11). Here, I demonstrate that mrbon dioxide is required for in vitro
activation of urease apoprotein. l detail several properties of this system, propose a
mechanism for the in vitro process, and discuss how UreD may assist in urease
activation.

Purified K. aerogenes urease apoprotein can be activated in the absence of
any accessory protein by incubation with nickel ion in the presence of bicarbonate-
containing buffers (Fig. 1A). Bewuse bicarbonate is in equilibrium with dissolved
mrbon dioxide in solution, it becomes imperative to determine which species is the
actual activating factor. By using NaHCOa from stock solutions that were adjusted to

92 .
pH 4.2 or pH 8.5 [the dissolved CO; concentration is much greater in the former

(12)], urease activation was demonstrated to be carbon dioxide-dependent (Fig. 18).
Additional support for this conclusion was obtained from experiments in which
carbonic anhydrase was included in the activation mixture. The initial rapid
activation (complete by ~5 min) observed for sample amended with the low pH
NaHCOa stock solution is eliminated in the presence of carbonic anhydrase due to
immediate hydration of dissolved CO; at the pH of the assay (Fig. 1B). The
bicarbonate/C02 concentration affects the extent of activation (Fig. 1A); the maximal
specific activity (~310 Ulmg) in this illustratration accounts for activation of
approximately 12.5% of the urease apoprotein that was present, but activation of up
to 30% was observed in another apoprotein preparation. Half-rnaximal activation of
urease apoprotein at pH 8.3 occurs at approximately 10 mM bicarbonate
concentration, corresponding to about 0.063 mM or 0.2% CO; (13) [for comparison,
the CO; concentration in the atmosphere is approximately 0.03% (12)]. The
activating C02 molecule becomes part of the enzyme as shown by measuring
incorporation of 1“C-NaHCOt ( 14). The labeling accounted for incorporation of 0.48
bicarbonate wrbon atoms per protein molecule. The specific activity generated in
this experiment (241 Ulmg) is much less than 48% of that expected for fully active
enzyme (~2500 Ulmg, 15); thus, CO; incorporation does not equate to enzyme
activation and other factors are important. The nickel ion concentration dependence
for urease apoprotein activation was assessed using buffer containing 100 mM
NaHCOa (Fig. 1C). The activation rate appears to saturate at approximately 60 11M
nickel ion. This value is only ~six-fold higher than the protein concentration or only
3-fold higher than that needed to load the bi-nickel site of the protein. lmportantly,
prior incubation of urease apoprotein with nickel ion followed by addition of
bicarbonate failed to yield active enzyme. This non-productive interaction between
apoprotein and nickel ion may account for the low extent of activation described
above. Activation of urease apoprotein under an atmosphere of 0.3% C02 exhibited

93

Fig. 1. In vitro activation of urease apoprotein. Urease apoprotein was purified from
Escherichia coli DH5 cells bearing plasmid pKA022ureD—1, containing the K.
aerogenes gene cluster deleted in ureD, as previously described (7). Enzyme
activity was assayed in 25 mM HEPES, 0.5 mM EDTA, 50 mM urea (pH 7.75). The
specific activity of urease was calculated based on linear regression analysis of the
rate of released ammonia, determined by conversion to indophenol (21 ), and protein
concentration determination (22). One unit of urease activity is defined as the
amount of enzyme required to degrade 1 mmol of urea per min at 37°C. (A)
Apoprotein (0.8 mglml or 9.6 uM final concentration) was added to activation buffer
[100 mM Hepes (pH 8.3), 150 mM NaCl, 10 mM EDTA] containing 100 pM MCI; and
0 (O), 1 (I), 3 (A), 10 (O), 30 (A), 100 (El) and 200(0) mM NaHCOs at 35°C.
Aliquots were removed overtime to monitor for urease activity. The results are
representative of four independent experiments. (B) Apoprotein (as above) was
placed in activation buffer containing 200 M NICIz at 0°C. NaHC03 was
immediately added (final concentration of 1 mM) from 4 mM stock solutions that
were prepared at pH 4.2 (O) or 8.5 (A) and timed aliquots were removed for assay.
An identical low pH NaHC03 stock solution experiment was carried out in the added
presence of mrbonic anhydrase (0.2 mglml) (O). The figure represents the
combined results of three separate experiments. (C) Apoprotein (as above) was
incubated at 35°C in activation buffer containing 100 mM NaHCO;. and mm; was
added to a concentration of 10 (O), 20 (I), 30 (A), 40 (O), 60, (A), or 100 (El) 11M.
Aliquots were removed and assayed at the indicated timepoints. The results are
representative of four independent experiments. (D) Activation buffers containing
100 mM Hepes (O or O) or Ches (A or A), 100 M NiClz, and 0.2 mglml mrbonic
anhydrase (added to speed up the rate of equilibration) were incubated at 35°C in
the presence of 0.3% CO; atmosphere. After a 30 min incubation, apoprotein (as
above) was added and aliquots were removed for assay at 45 (open symbols) and
120 (closed symbols) min. The indicated pH values were measured at the
conclusion of the experiment. The results are representative of two independent
experiments.

8.A.(Ulmg)
i§

 

8.A.(U/mg)
i§

 

 

 

 

 

I

60

 

    

120 180 240 300
Time(min)

l 1 J
r I 1 fi

120 180 240 300
Time(min)

8.A.(Ulmg)

 

 

 

 

B
0.03 ,_ .. . Q .
O O
. O
0
0m - '
8..
O
:'
O
0.01 " .0 0.0
O
419.908
0.00 °’° i 1 t
0 3 6 12
Time(n'in)
200 -—
D
150 -
a
E
a 100 -
<2
a)
50 .l
0 4 i t t 1
6.5 7 7.5 8 8.5

95 .
a clear pH dependence (Fig. 1D) with buffers of increasing pH leading to greater

rates of activation. The pK. of the activation process is 9 or greater, higher pH
values could not be examined because of the excessive amounts of bicarbonate that
would be present in the reaction mixture.

Urease activation kinetics, in which the extent of reaction is governed by the
CO; concentration and the rate is controlled by the nickel ion concentration, can be
accomodated by either an interaction of CO; with apoprotein prior to metal ion
binding or formation of a metal ion/CO; complex that binds to the apoprotein. There
exists a well characterized biologiwl precedent for the former mechanism involving
ribulose-1,5-bisphosphate carboxylaseloxygenase (Rubisco). This magnesium-
dependent enzyme has been shown to be activated by reaction of a lysine residue
with 002, where the resulting carbamate serves as a metal ligand (reviewed in 16).
A similar activation step may occur in 1-aminocyclopropane-1-carboxylate (ACC)
oxidase, a ferrous ion-dependent enzyme (17). As we observe for urease, the
activation rates for these enzymes are enhanced at higher pH (18, 19). I propose
that metallocenter assembly into urease apoprotein occurs by a pH-dependent
reaction between a protein sidechain and an activating C02 molecule to generate a
ligand that facilitates productive nickel binding. Unlike Rubisco and ACC oxidase,
however, urease activation is not reversed by addition of metal ion chelators. We
speculate that urease activation may include a protein conformational change that
serves to trap the metallocenter.

The ability to partially activate urease apoprotein in the absence of any
accessory protein compelled me to re-evaluate the previous suggestion that UreD
functions as a molecular chaperone (8); thus, we examined the effect of CO; on
activation of a mixture of UreD-urease apoprotein complexes [(urease)3UreDN,
where N = 0, 1, 2, and 3; <N> a 1.6]. The extent of activation was shown to depend
on the bicarbonate concentration (Fig. 2A) and the rate was found to depend on
nickel ion concentration (Fig. 28) with concentration dependences and kinetics that
closely resemble data obtained for urease apoprotein alone (Fig. 1A and 1C).

 

 

 

 

 
  

 

 

 

 

 

 

 

600--
A i
500-- .:/
-400“ / A
U) D
5 f s
2300*-/ 2
< <
"’ 200, a:
i
ioo-i'
0 t i i i i O T . 3 . fi
060120180240300 060120180240300
Time (min) Time (min)
71' C
5!
05‘-
43.
(D4-
“6
:23
(U
cr
2 + _,
1
O
o.

 

0 50 100 150 200

[NaHC 03] (mM)
Fig. 2. In vitro activation of UreD-urease apoprotein complex. A mixture of the UreD-
urease apoprotein complexes was enriched through the Mono 0 HR 10/10 step of
purification as previously described (8). The (A) bicarbonate dependence of
activation and (B) nickel dependence of activation for the protein complex (0.8
mglml) was carried out as described for the apoprotein alone (Fig. 1A and Fig. 1C,
respectively). The ratio of the specific activities that are generated during activation

of the UreD-urease apoprotein and the urease apoprotein are compared as a
function of bicarbonate concentration for 4 min (0) and 300 min (O) of activation.

97 .
Furthermore, 0.51 carbon molecules were incorporated into urease from 1‘C-

NaHCOe treatment of the ureD-urease apoprotein complex mixture, again in close
agreement with the urease apoprotein studies. However, the final level of activation
for the UreD-urease complex mixture is higher than for the urease apoprotein alone,
with the effect most pronounced at the lowest bicarbonate concentrations (Fig. 20).
Indeed, the indiwted ratios for activation are underestimated by a factor of 1.19 if
one converts Ulmg total protein to Ulmg urease protein for the experiments that
include UreD (20). In order to compare the effect of UreD on initial activation rates,
the activities at 4 min were compared for the UreD-urease apoprotein complex and
the apoprotein (Fig. 2C). The presence of UreD appears to accelerate the rate at
low bicarbonate concentrations, but at concentrations above ~50 mM the rate is
reduced in the presence of UreD. Like apoprotein, the UreD-urease apoprotein
complex was shown to form a non-productive nickel-urease species; however, the
presence of UreD appears to reduce the rate of this inactivation step. For example,
10 min incubation of urease apoprotein with 100 [M nickel ion prior to adding
bicarbonate reduces its ability to be activated to 8.8% of that in which nickel ion and
bicarbonate are provided simultaneously, whereas, UreD-urease apoprotein
complex that is treated in a similar manner retains 25% of its ability to be activated.
These results confirm the importance of UreD to urease activation at low
concentrations of CO; and suggest a mechanism for this protein. I propose that
UreD functions, at least in part, to minimize non-productive binding of nickel to
apoprotein that lacks bound C02; i.e., the protein species that is expected to
predominate at low C02 concentrations. This function, if verified by further studies,
may still be considered that of a molecular chaperone in which UreD controls the

proper sequence of activation steps so that CO; binds prior to nickel ion.

:PSPN.‘

.01

10.
11.
12.

13.

14.

98
REFERENCES AND NOTES

J. B. Sumner, J. Biol. Chem. 69, 435 (1926).

N. E. Dixon et al., J. Am. Chem. Soc. 97, 4131 (1975).

H. L. T. Mobley and R. P. Hausinger, Microbial. Rev. 53, 85 (1989).

R. P. Hausinger, in Biochemistry of Nickel (Plenum Publishing Corp., New
York, 1993), pp. 23-57.

S. Wang et al., Inorg. Chem. 33, 1589 (1994).

K. aerogenes urease is comprised of three subunits (M, 11,086, 11,695, and
60,304) arranged in a (0437):, structure containing three active sites (4).
Protein or urease concentrations indicated here refer to the (1.87 catalytic unit.
M. H. Lee et al., J. Bacteriol. 174, 4324 (1992).

IS Park, M. B. Carr, R. P. Hausinger, Proc. Natl. Acad. Sci. USA 91, 3233
(1994).

M. H. Lee et al., Protein Sci. 2, 1042 (1993).

A Lutz et al., Mol. Microbiol. 5, 123 (1991).

T. Maier et al., J. Bacteriol. 175, 630 (1993).

J. N. Butler, Carbon Dioxide Equilibria and their Applications (Addison-Wesley
Publishing Company, Reading, MA, 1982).

Calculation of the dissolved CO; concentration was based on the equation
[00,] = CT[H‘]2/(K.1K.2 + K.,[H‘] + [H‘]:), where cT is 10 mM, [H‘] is 5.01 x 10"
(for pH = 8.3), and values for pK.t and W.,; were 6.0 and 9.74 (interpolated
from Tables 2.1 and 2.2 of ref. 12), respectively. Conversion from mM to %
concentrations used Henry's law with pKH estimated to be 1.61 (interpolated
from the same source).

Urease apoprotein (0.5 mglml final concentration) was incubated in the
standard activation buffer containing 100 M NICIz and 50 mM NaHCOa (1.44
mCi/mmole) for 9 h, quenched with EDTA (5 mM final concentration), and

15.
16.
17.

18.
19.

20.

21.

99 .
chromatographed using a Mono Q 10l10 column as previously described

(15). Following chromatography, the enzyme specific activity was measured
and the specific radioactivity was determined using a scintillation counter with
measurement of the stock Nal-lCOa radioactivity used as a control.
Analogous experiments were carried out with a mixture of the UreD-urease
apoprotein complexes at 0.79 mglml.

M. J. Todd and R. P. Hausinger, J. Biol. Chem. 264, 15835 (1989).

F. C. Hartman and M. R. Harpel, Annu. Rev. Biochem. 63, 197 (1994).

J. G. Dong, J. C. Femandez-Maculet, S.‘ F. Yang, Proc. Natl. Acad. Sci. USA
89, 9789 (1992).

G. H. Lorimer, M. R. Badger, T. J. Andrews, Biochemistry 15, 529 (1976).

J. C. Femandez-Maculet, J. G. Dong, S. F. Yang, Biochem. Biophys. Res.
Commun. 193, 1 168 (1993).

For converting Ulmg total protein to Ulmg urease protein for the UreD-urease
complexes, the measured specific activity was multiplied by M,
(urease)3UreDN and divided by M2 (urease)3, where (urease): has M, 249,300
and UreD has M, 29,807.

M. W. Weatherbum, Anal. Chem. 39, 971 (1967).

O. H. Lowry et al., J. Biol. Chem. 193, 265 (1951).

CHAPTER 6

RELATED STUDIES, AND
PROSPECTS FOR FURTHER RESEARCH

100

1 01
A. Characterization of aHis-219 and otHis-312 mutant proteins

The 0His-219 residue was suggested to be important for substrate
binding, based on the high Km value of the 0H219A mutant protein (1 ). Two
possible mechanisms to account for this result include (i) stabilization of urea
binding by hydrogen-bond formation and (ii) maintenance of catalytic site
accessibility by protein configurational support involving the imidazole group.
Reductions in the rates of urease inactivation by alkylating agents and
diethylpyrocarbonate (DEP) are consistent with diminished active site
accessibility of the 0H219A mutant protein. To further examine the function of
this residue, l constructed and partially characterized glutamine (0H219Q) and
asparagine (0H219N) mutant proteins. Unlike the alanine mutant, these changes
may still enable hydrogen-bonding to take place, and these residues have more
of a resemblance to histidine in terms of shape and size. Both mutant proteins
have lower K.,. values (~ 175 mM for 0H219N or ~227 mM for 0H2190)
compared to the alanine mutant protein (~ 2100 mM) (Table 1); however, the
values are still higher than that of the wild-type protein (~ 2.25 mM). The
aH219N protein, like orH219A, had 10~20-fold effects on the rate, whereas the
0H219Q protein possessed a rate that was only ~1/2 that of the wild-type
enzyme. The optimal pH for catalysis was found to be ~6.2 in the 0H219N
protein and ~6.5 in the 0H2190 protein (Fig. 1A and 8), compared to a value of
7.75 in the wild-type and H219A samples. The role of the aHis-219 residue
remains unclear. Further characterization, e.g. selected chemical inactivation
studies of these mutant proteins, may be useful to allow us to further define the
function of this amino acid. Furthermore, additional synthesis of mutant proteins
with bigger amino acids, such as tryptophan, in place of the aHis-219 residue
may be necessary to confirm theeffect of amino acid side chain size on
substrate binding.

0His-320 was proposed to act as a general base in catalysis (1 ).

Evidence consistent with this hypothesis includes the rapid inactivation of the

102

Table 1. Characteristics of wild-type and selected mutant Klebsiella aerogenes

 

 

 

ureases'.
Km° kcat° Assay‘
Urease” (mM) Ulmg %wr pH, buffer
Wild-type 2.25:0.04 296:4 100% 7.75, HEPES
H219A 2092:634 14:3 5% 7.75, HEPES
H219N 175:8 11.6:0.2 4% 6.2,MES
H2190 227:21 134:6 45% 6.5,MES
H320A 10.9:0.6 0.0049:0.0002 0.0017% 6.2,MES
H320N 7.4:0.5 0.0043:0.0001 0.0015% 6.2,MES
H3200 10.6:1 .0 0.0024:0.0001 0,0008% 6.2,MES

 

‘ Determined for cell extracts. SDS-polyacrylamide gel electrophoretic analysis
revealed that the urease expression levels were nearly identical in all
constructs except H219N, that appears to have approximately half the urease
protein of the others.

” For site—directed mutagenesis, a 1.1-kbp BamH1-Sell fragment of the pKAU17
was used as described previously (Chapter 3). The following oligonucleotides
were used: GAAGATCCAAGAGGACTGG (H2190),
CTGAAGATCAATGAGGACTGG (H219N), GGTCTGCCAQCATCTGGAC
(H3200). GATGGTCTGCAACCATCTGG (H320N).

° Measured by using 50, 100, 130, 170, 250, and 1000 mM urea for the His-219
mutant proteins or 2, 2.5, 3.5, 5, 10, and 50 mM for other proteins.

‘ Assays were carried out in 25 mM buffer containing 0.5 mM EDTA.

103

12 1-

i

 

 

 

 

 

 

A T B
”8'”.
300‘ (PO 9“ .0.
~11. o - .
a 200 - E’- 5 °oc88
‘5 o 4 0° 0
a) o to
100 -~ 3-~
at“. q. 60 o 8.
OO .0. o
0 g .‘0 . 0 ° .
5 7 9 11 5 7 9 11
pH pH
0.006 T
000 C
o o
O
a 0004 0. ‘50
E . c
a E c.o
< O Omo
0 0002» AA
AA 454 ~£°°
A O
F 4164“ o o
A o
0 1 .2! 0—1
5 7 9 11
pH

Fig. 1. The pH dependence of wild-type and mutant urease activity of cell
extracts. The activities were measured for extracts of E. coli DH5 cells canying
pKAU17 or mutant plasmids that encode the following proteins: (A) wild-type
(Q) and the aH219Q mutant (0); (B) orH219A (Q) and 0H219N (O); (C)
0H320A (O), 0H32OQ (A), and aH320N (Q). The reaction mixtures contained
urea (50 mM for wild-type enzyme and H320A, Q, and N proteins or 1 M for the
0H219A, Q, and N proteins), 0.5 mM EDTA, and the following buffers at a
concentration of 25 mM: MES (pH 5.1~6.19), HEPES (6.0~9.1), and CAPS

(9.4~104).

104 .
wild-type enzyme by DEP (2), a histidine-selective modifier, and the resistance

of 0LH320A mutant protein to inactivation by DEP (1 ). Two additional mutant
proteins, aH320N and 0H320Cl, were synthesized to further examine this
proposal. All three mutant proteins (0H320A, 0tH32OQ, 0H320N) have similar Km
values, specific activities, and pH dependencies in catalysis (Table 1, Fig. 1C).
These results indicate that the histidine residue is probably not important for
steric or hydrogen bonding interactions, rather these results are indirectly
consistent with the role of aHis-320 as a general base for catalysis, as

suggested previously (1 ).
B. Function of UreD in urease apoprotein activation

Activation of urease apoprotein requires CD: as well as nickel (3). UreD
was previously shown to form three protein complexes with urease apoprotein
(4), and a mixture of these complexes is also competent for activation in a C02-
dependent manner (3). In both cases, the activation is apparently accompanied
by a non-productive inactivation process; however, the presence of UreD
appears to reduce the rate of this inactivation step. Based on this finding, the
UreD protein was proposed to function, at least in part, to keep apoprotein from
binding nickel non-productively in the absence of C02, as illustrated in Scheme
1, where k; is reduced compared to k1. This hypothesis was further tested by
characterizing the activation process for the partially resolved complex species.

A molecule of urease apoprotein can associate with 1, 2, or 3 molecules
of UreD (4). Each of the forms of UreD-urease apoprotein complex was enriched
by chromatography of the mixture on a phenyl-Sepharose column. The urease
activation kinetics look similar in all three pools (Fig. 2). At low concentrations (1
mM) of H00; (therefore, low concentrations of C02) the extent of activation for
each complex was limited even in the presence of saturating concentrations
(100 nM) of nickel ion (Fig. 2). Importantly, the presence of increasing numbers

of UreD per urease apoprotein leads to higher activation levels in the low

105

Inactivation Inactivation
Urease L) UreDt-.
apoprotein ap0pro 9'"
complex
Ni Y C02 Ni i C02
Holoprotein Holoprotein
+
UreD
(precipitated)
<Scheme1>

bicarbonate studies. In contrast, the level of activation reached near its
maximum in the presence of high concentration (100 mM) of H00; regardless of
concentration (20 vs. 100 nM) of nickel ion, and the effect of UreD number was

less pronounced. The results are consistent with CO; being the limiting factor in

106 _
Fig. 2. In vitro activation of the three partially purified UreD-urease apoprotein

complexes. The protein complexes were purified as previously described, with
minor modifications (4). E. coli DH5[pKAUD2] cell extracts from a 2 L culture
were chromatographed on a column (2.5 x 20 cm) of DEAE-Sepharose at 4°C in
PEDG buffer [18 mM phosphate (pH 7.0), 0.09 mM EDTA, 10% glycerol, 0.09
mM dithiothreitol]. The proteins were eluted in the same buffer with a linear salt
gradient to 1.0 M KCI. The protein complex-containing fractions were pooled,
dialyzed against PEDG buffer, and applied to a Mono-Q HR 10/10 column
equilibrated in the same buffer. The protein was eluted with a linear salt gradient
to 1 M KCI in PEDG buffer. The pooled fractions containing the mixture of
protein complexes was adjusted to 1 M KCI by addition of PEDG buffer
containing 2 M KCI and applied to a phenyl-Sepharose HP (Pharmacia) column
(2.6 x 10 cm) equilibrated in this buffer. A linear gradient to 0 M KCI in PEDG
buffer was used to partially resolve the protein complexes into three pools.
Samples were monitored for the presence of the UreD-urease apoprotein
complexes by using native polyacrylamide gel electrophoresis. Pool 1 (eluted at
0.23~0.19 M KCI) contains apoprotein (19% of total protein), 1UreD-apoprotein
(47%), 2UreD-apoprotein, (27%), and 3UreD-apoprotein (7%) (averaging to 1.00
UreD per apoprotein molecule). Apoprotein was undetectable in pool 2 ( eluted
at 0.11~0.08 M KCI) which possesses 1UreD-apoprotein (16% of total protein),
2UreD-apoprotein (58%), and 3UreD-apoprotein (26%) (averaging to 2.00 UreD
per apoprotein molecule). In pool 3 (eluted at 0 M KCI) only 2UreD-apoprotein
and 3UreD-apoprotein were detectable (14% and 86% of total protein,
respectively. averaging to 2.83 UreD per apoprotein molecule). Each pool was
again chromatographed on a Mono-Q HR 10/10 column, as described above.
The three species of UreD-apoprotein complex (0.2 mglml or 2.4 pM final
concentration; circle: pool 1, square: pool 2, triangle: pool 3) were added to
activation buffer [100 mM Hepes (pH 8.3), 150 mM NaCl, 10 mM EDTA]
containing 20 (open symbols) or 100 (closed symbols) uM NiClz and 1 or 100

mM NaHCOa at 35°C. Aliquots were removed over time to monitor for urease

107

 

 

 

 

 

 

 

 

 

 

 

1200 ~—
/ /\e 100"“
900 -— _ ”am
’5 i _.
é
a, 600 -- ,
< l T _
05 I?"
300 *1” mm
/ m jg Nat-CO;
O '2‘" ‘/z i T—

0 60 120 180 240 300
Time (min)

activity. Enzyme activity was assayed as described elsewhere (1 ). In order to
convert the specific activity values based on total protein (urease plus UreD) into
values of urease specific activity, the rates were multiplied by factors of 1.12 for
pool1, 1.24 for pool 2, and 1.34 for pool 3.

1 08
urease apoprotein activation and with the presence of UreD serving a more

important role at low C02 levels.

Each of the UreD-urease apoprotein complexes lost their ability to be
activated by incubation in the presence of 100 pM NiClz prior to addition of
bicarbonate. For example, an 80 min incubation with nickel almost completely
abolished the ability of these proteins to be activated (Fig. 3). A 10 min
incubation reduced the level of final activation to only 25~30% of the control
samples (Fig. 3). lmportantly, the number of UreD bound per apoprotein affected
the extent of inactivation with increasing protection observed with increasing
UreD molecules bound per apoprotein.

The initial activation rates of each complex were examined under four
different conditions. When the activation was carried out at saturated
concentrations of HCOa' (100 mM) and NiCl2 (100 nM), the rate in the pool 1
sample was the fastest (Fig. 4A). This suggests that an increasing number of
UreD per urease apoprotein may hinder the rate of activation under these
conditions. However, the rate of pool 3 activation becomes the fastest and that
of pool 1 the slowest (Fig. 4B) when high concentrations of NiClz (100 (M) are
used in the presence of low concentrations of CO; (no addition of NaHCOa). The
difference apparently increases (Fig. 40) when the concentration of NiClz is
even higher (200 nM). From these results it can be hypothesized that either (i)
the complex is better at incorporating C02 than the apoprotein alone under low
concentration of CO; or (ii) the presence of UreD may protect the urease
apoprotein from undergoing nickel-induced inactivation. A low concentration of
MCI: (20 nM) results in pool 1 exhibiting the fastest activation among the three
pools even in the presence of low CO; (Fig. 4D). This result is compatible with
hypothesis (ii); i.e., UreD may somehow prevent the nonproductive inactivation

of urease apoprotein.

C. Characterization of UreD

109

 

 

 

 

 

 

 

 

 

1500 «- Preincubation
~ ' 0 min
1200 ~— /
’6:
g 900~~
3
< 600 -— ,
(I) _
l’=" ‘ .
300 __ /{_ . _o 10 min
JL ‘ " ”:- I 80 min
0 ' l l J.—
0 60 120 180
Time (min)

Fig. 3. Inactivation of the three UreD-urease apoprotein complexes by
incubation with nickel prior addition of NaHCOa. The UreD-urease apoprotein
complexes (0.2 mglml; circle: pool 1, square: pool 2, triangle: pool 3, see Fig. 2
for details) were incubated with the activation buffer containing 100 (M NiClz for
0, 10, or 80 min. NaHCOa (100 mM, final concentration) was added at time 0 and
aliquots were removed over time to monitor for urease activity. Calculation of

Urease specific activities was done as described in Fig. 2.

110

120 '1

1

S.A(U/mg)
S.A.(UImg)

 

 

 

 

 

Time (min)

1

100 a

S.A.(UImg)
S.A.(UImg)

 

 

 

 

 

Time (min) Time (min)

Fig. 4. Initial activation rates of UreD-urease apoprotein complexes. The UreD-
Urease apoprotein complexs (0.2 mglml; O, pool1, I, pool 2, A, pool 3, see Fig.
2 for details) were added to the activation buffer containing (A) 100 (M NiCIz and
1 00 mM NaHCO:, (8)100 uM NiC'z and no added NaHCOa, (C) 200 pM NiClz
and no added NaH003, or (D) 20 uM NiC'z and no added NaHCOa, all at 35°C.

Aliquots were removed over time to monitor for urease activity. Specific activity

Corrections were done as described in Fig. 2.

1 1 1 .
UreD has been purified only as one of three UreD-urease apoprotein

complexes (4). Further characterization of UreD is required to test the validity of
its suggested function as a molecular chaperone in apoprotein activation (3, 4).
In this section, I describe the partial characterization of UreD and my efforts to
examine the formation of the UreD-urease apoprotein complex.

pKAUDG (see Appendix) was constructed to contain only the ureD gene
without the urease structural genes or other accessory genes. Samples
prepared from Eco/i DH5 containing this plasmid possess large amounts of
UreD, which, however, was found only in the insoluble fraction (Fig. 5A, lane 3).
The presence of other accessory gene products (UreE, UreF, and UreG) does
not appear to be helpful for maintaining solubility of UreD (Fig. 5A). Several
detergents tested (such as NP-40, Tween-20, and Triton X-100) at up to 3%
concentration do not solubilize the protein. The protein was found to be capable
of solubilization in the presence of urea (~6 M) or guanidine-HCI (~4 M) (Fig.
5B). This result suggests that UreD forms an inclusion body when synthesized in
large amounts. It is unclear whether low level synthesis of UreD may result in a
soluble protein. The denaturant (8 M urea or 4 M guanidine HCl)-solubilized
protein was diluted into buffer alone or into buffer containing 1% detergent and
found to be insoluble under these conditions (data not shown)

To test whether another accessory gene (ureE, ureF or ureG) is required
for formation of the UreD-urease apoprotein complex, plasmid pKAUDABC
containing only ureD, ureA, ureB and ureC (see Appendix) was constructed.
Eco/i containing this plasmid was shown to possess UreD-urease species (Fig.
SC), indicating that formation of the UreD-urease apoprotein complexes does not
require other accessory genes. The cell extracts prepared from E. coli
[pKAUDABC] appear to have a similar level of ability to be activated in vitro as
that from wild-type extracts (Chapter 4, Fig. 1). This result suggests that other
accessory proteins (UreE, UreF, and UreG) are not essential for forming a

urease apoprotein species that is capable of activation in vitro.

112

Fig. 5. Characterization of UreD. (A) Samples were prepared from Eco/i cells
containing pKAUD6 (lane 2, 3), pKAUG1 (lane 4, 5), pKAUDG1 (lane 6, 7), or
pKADEFG (lane 8, 9) (see Appendix for maps). Whole disrupted cells (lane 3, 5,
7, 9) or the supematants (lane 2, 4, 6, 8) of the cell extracts after centrifugation
at 15,000 g for one hour were analyzed on SDS- polyacrylamide (12%) gel
electrophoresis. Lane 1 is molecular weight markers (phosphorylase b, Mr
92,500; bovine serum albumin, Mr 66,200; ovalbumin, Mr 45,000; carbonic
anhydrase, Mr 31 ,000; soybean trypsin inhibitor, Mr 21,500; and lysozyme, M,
14,400). (B) UreD present in E. coIIIpKAUD6] cells was tested for solublization in
buffer containing 25 mM Hepes (pH 7.9), 150 mM NaCI, 1 mM EDTA, and 10%
glycerol supplemented with 3, 4, 5, 6, 7, and 8 M urea (lane 2~7), or 1, 2, 3, 4, 5,
and 6 M guanidine HCl (lane 8~13). After incubation at 0°C for 30 min, the cell
extracts were centrifuged for 5 min at 15,0009. The supematants were analyzed
on SDS- polyacrylamide (13.5%) gel electrophoresis. Lane 1 is molecular weight
markers. Disrupted cells are shown in lane 14. (C) Western blot comparison of
cell extracts of E. coli [pKAU17] (lane 1), E. coli [pKAUDABC] (lane 2) and cell
extracts of E. coli [pKAUD2] (lane 3). Cell extracts were run on 6% native gel
and probed with anti-urease antibody (5).

113

 

-3D- Apo
--ZD -Apo
-1D- -Apo
-Apo

1 14
D. Possible role of UreF

The ureF gene is absolutely essential for the formation of active urease in
vivo (6). This gene is expressed at very low levels, apparently due to a weak
Shine-Dalgamo (SD)-sequence (ribosome-binding site) in front of the structural
gene (7). I made two site-directed mutants (pKAUF1 and pKAUF2, Fig. 6) which
have changes in their ribosome binding site regions leading to matches with 3 or
5 positions compared to the consensus SD-sequences. However, the desired
high level expression of UreF was not observed in cell extracts from these
mutants. Although the reason for continued low level expression of ureF is not
clear, there is still a prospect for the high level synthesis of UreF if the
environment around the ureF gene is altered, e.g., if the other genes are
deleted.

While comparing UreD-urease complex samples from cells containing the
intact gene cluster and various deletion mutants, a difference was observed that
provides evidence for a role of UreF in complex formation. The major UreD-
urease apoprotein complex in cell extracts prepared from E. coli [pKAU17] elutes
earlier from a Sepharose 6 column than does the apoprotein, consistent with a
larger hydrodynamic radius for UreD-urease than for urease alone (data not
shown). Similar elution patterns are observed for extracts from cells canying
pKAUAE-1 (Fig. 7A) or pKAUAG-1 (Fig. 7C) (i.e., ureE and ureG deletion
mutants). Surprisingly, the UreD-apoprotein complex in extracts from cells
carrying pKAUAF, a ureF deletion mutant, elutes at the same position as the
apoprotein (Fig. 7B). This result suggests that the UreF protein may be involved
in conversion of a highly compact form of UreD-urease apoprotein complex to a
less compact structure where these forms are not resolved by native
polyacrylamide gel electrophoresis. It is possible that such a conformational I
change is related to the decreased level of in vitro urease activation observed in
E. coli (pKAUAF) cell extracts compared to a control sample or to samples

possessing ureD or ureG deletions (4).

115

pKAU1 7 CAGCCACTAGCATGTC‘"
pKAUF1 CAGAQACTAGCATGTC'"
pKAUF2 CAGGAGCTAGCATGTC‘"

Fig. 6. Site-Directed Mutagenesis of the ureF Ribosome Binding Site. In order to
enhance expression of ureF, the ribosome binding site of this gene was changed
as shown above (underlined sequences) by site-directed mutagenesis methods.
A 1.8-kb BamH1-Avril fragment of plasmid pKAU17 containing the urease gene
cluster from K. aerogenes was subcloned into M13 mp18 and mutagenized by
the method of Kunkel et al. (8). Uracil-containing single-stranded template DNA
was prepared from Escherichia coli CJ236 (dut1 ung1 thi-1
relA1IpCJ105[cam'F']). Using GACATGCTAGTQICTGTGAGCGTG (pKAUF1)
or GTCGACATGCTAGC_TQCTGTGAGCGTGGTG (pKAUF2) oligonucleotide
primers, the phage DNA was mutagenized and mutant phage were isolated in E.
coli MV1 193([IacI-proAB] rpsL thi endA spcB15 hst4 [sfl-recA]306::Tn10[tet‘]
F'[traD36 proAB‘ Iacf'lacZM15]). The site-directed mutants were identified by
sequence analysis using Sequenase 2.0 (US. Biochemical Corp., Cleveland
OH) and the single-strand DNA sequence method of Sanger et al. (9). The
region was completely sequenced to ensure that no other mutations had been
introduced into M13. The mutated BamH1-Aatll region was subcloned back into
pKAU17 to generate pKAUF1 and pKAUF2 and the sequence was confirmed by
double-stranded DNA sequencing methods. Putative ribosome-binding sites and
the start codon are highlighted.

116

Volume(m|)
28 36 44 52 60 68 76

 

Fig. 7. Western blot comparison of fractions from Sepharose 6 (Pharmacia)
column chromatography of selected cell extracts. Cell extracts were prepared
from (A) Eco/i [pKAU17AE-1], (B) [pKAU17AF], and (C) [pKAU17AG-1] and
applied to a Sepharose 6 column (2.6 x 50 cm). Fractions (4 ml) were collected
and aliquots were analyzed on a 6% native gel, transferred to nitrocellulose
membrane, and detected by using anti-urease antibody and an alkaline

phosphatase-conjugated goat anti-rabbit-lgG.

 

117

E. Presence of a complex containing UreD, urease apoprotein, UreF, and
UreG

In addition to the three UreD-urease apoprotein complexes described
earlier (4), faint bands of much lower mobility that contain anti-urease antibody
cross-reactive material are detected in cell extracts from E. coli DH5 containing
pKAU17 (wild-type), pKAU17AE-1 and pKAU02 (abbreviated as 0*, leading to
high level synthesis of UreD) in western blotting studies using anti-urease
antibody (lane 1 and 3 of Fig. 5C, and lane 1 and 3 of Fig. 8). The bands are
designated B in the figures because they comprise a second or B series of
complexes that are distinct from the faster migrating UreD-urease complexes
already discussed. lmportantly, analogous bands were not detected in the
ureEFG deletion mutant (lane 2, Fig. 5C) or in ureD, ureF, or ureG deletion
mutants (lane 2, 4, 5 of Fig. 8). It is very intriguing that cells found to be
activation-competent in vivo possess the B complexes, whereas ureD, ureF, or
ureG deletion mutants, which are not competent for in vivo activation, do not.

The B complexes were enriched from extracts of cells containing pKAU17
and pKAUD2 by using DEAE-Sepharose and Mono-Q F PLC column
chromatographies. The species do not separate from the UreD-urease
apoprotein complexes when using Sepharose 6 size-exclusion column
chromatography. This was unexpected because the B complexes migrate much
more slowly than the UreD-urease apoprotein complexes during the native
polyacrylamide gel electrophoresis. Although overlapping, the peak fractions of
the UreD-urease complexes elute before the peak fractions of the B complexes
(Fig. 9A). When the same fractions are examined by denaturing gel
electrophoresis (Fig. 9B), it is clear that the fractions containing the B complexes
possess several bands in addition to those corresponding to urease and UreD.
One of these additional bands was demonstrated to be UreG by a western

blotting experiment using anti-UreG antibody (not shown). The other band below

 

118

.. ~~ -- -B

- 2D-Apo
.' “I «In - - 1D-Apo
h ‘ - M O. ‘ - Apo
ML —-

 

Fig. 8. Western blot comparison of cell extracts. Lane 1, E. coli [pKAU17]; Lane
2, E. coli [pKAU22AD-1 ]; Lane 3, E. coli [pKAU17AE-1]; Lane 4, E. coli
[pKAU17AF]; Lane 5, E. coli [pKAU17AG-1]; Lane 6, E. coli [pKAUF1] (see Fig.
6). The cell extracts were analyzed on a 5% native gel, transferred to a

nitrocellulose membrane, and probed with anti-urease antibody.

119

Fig. 9. Partial purification and characterization of the urease-containing B
complexes. Cell extracts from E. coli DH5 containing pKAU17 or pKAUD2 were
chromatographed on a column of DEAE-Sepharose at 4°C in PEDG buffer. The
proteins were eluted in the same buffer with a linear salt gradient to 1.0 M KCI.
The B protein complex-containing fractions were pooled, dialyzed against PEDG
buffer, and applied to a Mono-Q HR 10l10 column equilibrated in the same
buffer. The proteins were eluted with a linear salt gradient to 1 M KCI in PEDG
buffer with the 8 complexes eluting at ~05 M KCI. (A) Native polyacrylamide gel
electrophoretic analysis of Mono-Q fractions eluting from 0.43 to 0.51 M KCI
from D+ cells possessing elevated levels of UreD. (B) SDS-polyacrylamide gel
electrophoretic analysis of the same Mono-Q fractions. M represents molecular
weight markers (see Fig. 5). (C) Nondenaturing polyacrylamide gel
electrophoretic analyses of Mono-Q pool of B-complex-containing fractions from
E. coli DH5 [pKAUD2] (lane 1) or E. coli DH5 [pKAU17] (lane 2). (D) SDS-
polyacrylamide gel electrophoretic analysis of selected pools containing B
complexes. Lane 1, molecular weight markers. Lane 2, 3, and 4: cell extracts,
DEAE-Sepharose pool, and Mono-Q pool of E. coli DH5 [pKAU17], respectively.
Lane 5, 6, and 7: cell extracts, DEAE-Sepharose pool, and Mono-Q pool of E.
coli DH5 [pKAUD2]. (E) 2D gel electrophoretic analysis of the B complex. The
native gel portion containing the B complexes of the Mono-Q pool of E. coli DH5
[pKAU02] was excised, denatured, and eletrophoresed using an SDS-
polyacrylamide gel.

 

A- c. 1 2
y-r1
. ) .— -7
i}? B
rt )8
Lao-Apo-
—2D-Apo-
-1D-Apo— —-
- Apo - .- —-
M 1 2 3 4 5 6 7 8‘
B l
I -—.‘.u.—dv—-Urec
’

= UreA, B

   

121
UreG on the gel was excised and shown to have an N-terminal amino acid

sequence that agrees with that deduced for UreF from the DNA sequence (7).
As found for extracts of cells carrying pKAUD2, the pooled fractions from cells
containing pKAU17 exhibit bands corresponding to the B complexes (Fig. 9C)
and these fractions also appear to possess the UreF and UreG peptides (Fig.
9D). When the bands corresponding to the B complexes were excised from a
native gel and re—electrophoresed on a denaturing gel, both the UreG and UreF
bands were present as well as the urease subunits and UreD (Fig. 9E). Thus, I
suggest that the B complexes are composed cf UreD, urease apoprotein, UreF,
and UreG.

Urease apoprotein that is associated with the B complexes is apparently
capable of activation (Fig. 10). Upon incubation of the D+ Mono-Q fraction with
100 uM NiCIz (when this experiment was carried out, the involvement of CO; in
the activation process was not known), the protein was found to undergo a
change such that when analyzed by native gel electrophoresis the mutiple B
complex bands collapsed (Fig. 10A) in a manner which is reminiscent of UreD-
urease apoprotein activation (Chapter 4, Fig. 4). Enzyme activity was measured
in individual gel slices (Fig. 10B). As expected, significant levels of activity were
associated with the region associated with the UreD-urease apoprotein
complexes (region 1, 2, 3, and 4 in Fig. 10A) (Fig. 10B). Significant activity was
also detected in region 6 containing the B complex (Fig. 108). Enzyme activation
in the region appears to be complete at 30 min (Fig. 100). The K... value
determined for the enzyme in the slice of gel was ~10 mM which is a little higher
than that of urease (~3 mM, measured in the same way, or ~2.25 mM, measured
by using urease in solution). ATP, GTP, or high salt (up to 1 M KCl) does not
dissociate the B complex into its components (data not shown).

Purification of the complex will be essential for its further characterization.

F. Urease apoprotein activation: possible scheme

122

Activity (U/ml)
Activity (U/ml)

 

 

 

.. O l i l
123456789 0 50100150

Region Time (min)

 

Fig. 10. Activation of urease apoprotein associated with the B complexes. (A)
The Mono-Q pool of E. coli DH5 [pKAU02] was incubated for 0 or 100 min (lane
1 and lane 2, respectively) at 37°C in the presence of 0.1 mM NiCIz and

analyzed by native gel electrophoresis. (B) The same sample was incubated as

 

described in (A) for 30 min and the urease activities were measured for 9 gel

slices, cut as shown in A. (C) The effect of incubation time on the level of activity
was measured in the gel slice corresponding to region 6.

123 .
Urease apoprotein alone can be activated in vitro by providing nickel in

the presence of C02. UreD-urease apoprotein is also activation-competent in
vitro, but the presence of UreD and urease subunits is not sufficient for forming
active enzyme in the cell. The activation of urease in vivo absolutely requires
UreF and UreG, as well as UreD (6). It is reasonable that conditions in vivo
should be different from that found in vitro; for instance, the concentration of
nickel or CO; may be limited in vivo. The accessory gene products may serve to
provide these activating factors. For example, UreE is known to bind nickel ion
with high specificity suggesting that the protein may be the nickel donor. The B
complexes are composed of UreD, UreF, UreG and urease apoprotein. It is
intriguing that the 8 complexes have been found only in the activation-competent
wild-type, UreD-overexpressor (D’), and UreE deletion mutant cells. From these
results, I propose that the B complexes are required for in vivo activation of
urease, as illustrated in Scheme 2.

Inactivation

Urease
apoprotein

Ni f 002

Holoprotein

In vitro

Ni$k1 UreD

124

Inactivation

Mal” UreF, UreG

UreD- 5 B complex
apoprotein (UreD-apoprotein-

complex UreF-UreG)
Ni k C02 UreE-Ni i 002
Holoprotein Holoprotein
+ +
UreD UreD, UreE,
(precipitated) UreF, UreG
In vitro In vivo?

<Scheme 2>

125
REFERENCES

. Park, I. -S. and R. P. Hausinger. 1993. Site-directed mutagenesis of
Klebsiella aerogenes urease: identification of histidine residues that appear to
function in nickel ligation, substrate binding, and catalysis. Protein Sci.
2:1034-1041.

. Park, I. -S., and R. P. Hausinger. 1993. Diethylpyrocarbonate reactivity of
Klebsiella aerogenes urease: effect of pH and active-site ligands on rate of
enzyme inactivation. J. Prot. Chem.12:51-56.

. Park I. -S and R. P. Hausinger. 1994. Carbon dioxide is required for in vitro
assembly of the urease nickel metallocenter. Submitted to Science

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

. Mulrooney, S. B., H. S. Pankratz, and R. P. Hausinger. 1989. Regulation
of gene expression and cellular localization of cloned Klebsiella aerogenes (K.
pneumoniae) urease. J. Gen. Microbiol. 135:1769-1776.

. Lee, M. H., S. B. Mulrooney, M. J. Renner, Y. Markowicz, and R. P.
Hausinger. 1992. Klebsiella aerogenes urease gene cluster: sequence of
ureD and demonstration that four accessory genes (ureD, ureE, ureF, and
ureG) are involved in nickel metallocenter biosynthesis. J. Bacteriol.
174:4324-4330.

. Mulrooney, S. B., and R. P. Hausinger. 1990. Sequence of the Klebsiella
aerogenes urease genes and evidence for accessory proteins facilitating
nickel incorporation. J. Bacteriol. 172:5837-5843.

. Kunkel, T. A., J. D. Roberts, and R. A. Zakour. 1987. Rapid and efficient
site-specific mutagenesis without phenotypic selection. Methods Enzymol.
154: 367-382.

. Sanger, F., S. Mickle, and A. R. Coulson. 1977. DNA sequencing with chain-
terminating inhibitors. Proc. Natl. Acad. Sci. USA. 80: 3963-3965.

Appendix

pKAU17, pKAUD2 and their deletion mutants

Eco R1 AfI ll Bam H l Ksp l
Nru I Ksp l Bsm l Ksp l Rsr Il Hind lll

Inc—imma-

03* l .1
D ** L 1.
D5“ L —} L
DE“ I .k 1'
G1* I: L
DG1ttl . L
DEFG‘I‘l . L

A 02* B

EFG* E} L
BRH“ G fil—lll; 'L
DZABC’L fi- i
DABC‘ I -L 1'

02¢":M [ H

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

uouJJJJuJJJUU

02:16” 1 fl

* Derived from pKAU17
** Derived from pKAUD2
: Deletion

-: pUCB
l:| : Urease gene

 

126