Temperature- and pressure-dependent stopped-flow kinetic studies of jack bean urease. Implications for the catalytic mechanism

Article abstract of DOI:10.1007/s00775-012-0926-8

Urease, a Ni-containing metalloenzyme, features an activity that has profound medical and agricultural implications. The mechanism of this activity, however, has not been as yet thoroughly established. Accordingly, to improve its understanding, in this study we analyzed the steady-state kinetic parameters of the enzyme (jack bean), K_M and k_cat, measured at different temperatures and pressures. Such an analysis is useful as it provides information on the molecular nature of the intermediate and transition states of the catalytic reaction. We measured the parameters in a noninteracting buffer using a stopped-flow technique in the temperature range 15-35 °C and in the pressure range 5-132 MPa, the pressure-dependent measurements being the first of their kind performed for urease. While temperature enhanced the activity of urease, pressure inhibited the enzyme; the inhibition was biphasic. Analyzing K_M provided the characteristics of the formation of the ES complex, and analyzing k_cat, the characteristics of the activation of ES. From the temperature-dependent measurements, the energetic parameters were derived, i.e. thermodynamic ΔH^o and ΔS^o for ES formation, and kinetic ΔH^≠ and ΔS^≠ for ES activation, while from the pressuredependent measurements, the binding ΔV_b and activation ΔV6_cat ^≠volumes were determined. The thermodynamic and activation parameters obtained are discussed in terms of the current proposals for the mechanism of the urease reaction, and they are found to support the mechanism proposed by Benini et al. (Structure 7:205-216; 1999), in which the Ni-Ni bridging hydroxide-not the terminal hydroxide-is the nucleophile in the catalytic reaction. SBIC 2012.

Full text of DOI:10.1007/s00775-012-0926-8

J Biol Inorg Chem (2012) 17:1123–1134
DOI 10.1007/s00775-012-0926-8
ORIGINAL PAPER
Temperature- and pressure-dependent stopped-flow kinetic
studies of jack bean urease. Implications for the catalytic
mechanism
Barbara Krajewska
Małgorzata Brindell
•
Rudi van Eldik
•
Received: 30 April 2012 / Accepted: 14 July 2012 / Published online: 14 August 2012
Ó The Author(s) 2012. This article is published with open access at Springerlink.com
Abstract Urease, a Ni-containing metalloenzyme, fea-
tures an activity that has profound medical and agricultural
implications. The mechanism of this activity, however, has
not been as yet thoroughly established. Accordingly, to
improve its understanding, in this study we analyzed the
steady-state kinetic parameters of the enzyme (jack bean),
current proposals for the mechanism of the urease reaction,
and they are found to support the mechanism proposed
by Benini et al. (Structure 7:205–216; 1999), in which the
Ni–Ni bridging hydroxide—not the terminal hydroxide—is
the nucleophile in the catalytic reaction.
KM and k , measured at different temperatures and pres-
cat
Keywords Urease ꢀ Catalytic mechanism ꢀ Temperature
and pressure dependence ꢀ Thermodynamic and activation
parameters ꢀ Stopped flow
sures. Such an analysis is useful as it provides information
on the molecular nature of the intermediate and transition
states of the catalytic reaction. We measured the parame-
ters in a noninteracting buffer using a stopped-flow tech-
nique in the temperature range 15–35 °C and in the
pressure range 5–132 MPa, the pressure-dependent mea-
surements being the first of their kind performed for urease.
While temperature enhanced the activity of urease, pres-
sure inhibited the enzyme; the inhibition was biphasic.
Introduction
Ureases (urea amidohydrolases, EC 3.5.1.5) are high
molecular weight, multisubunit, Ni-containing metalloen-
zymes [1] that are found in numerous bacteria, plants, fungi,
algae, and some invertebrates, as well as in soils as a soil
enzyme [2, 3]. Bacterial ureases differ from plant and
fungal ones, typically homohexameric, in that they are
composed of heteromeric subunits. Yet, irrespective of their
origin, ureases fulfill one catalytic function: to hydrolyze
urea [2, 3]. The immediate products of this enzymatic
Analyzing K provided the characteristics of the formation
M
of the ES complex, and analyzing k , the characteristics of
cat
the activation of ES. From the temperature-dependent
measurements, the energetic parameters were derived, i.e.
o
o
thermodynamic DH and DS for ES formation, and kinetic
=
=
DH and DS for ES activation, while from the pressure-
dependent measurements, the binding DV and activation
reaction are NH and carbamate; however, the observed
3
b
6¼
products are NH3 and H CO , due to the spontaneous
2
DV volumes were determined. The thermodynamic and
3
cat
hydrolysis of carbamate (Scheme 1). These reactions cause
a significant increase in pH.
activation parameters obtained are discussed in terms of the
Notwithstanding that urease was the first enzyme ever
crystallized (1926) [4] and extensively studied over the
years, its catalytic mechanism still remains disputable
B. Krajewska (&) ꢀ M. Brindell
Faculty of Chemistry, Jagiellonian University,
Ingardena 3, 30-060 Krak o´ w, Poland
e-mail: krajewsk@chemia.uj.edu.pl
[
5, 6]. The elucidation of this mechanism is of importance
for counteracting undesirable effects generated by the
R. van Eldik (&)
enzyme. These include the product NH and an increase in
3
Department of Chemistry and Pharmacy,
Friedrich Alexander University Erlangen-N u¨ rnberg,
Egerlandstrasse 1, 91058 Erlangen, Germany
e-mail: vaneldik@chemie.uni-erlangen.de
pH, both capable of causing deleterious complications,
notably in medicine and agriculture [2, 3]. In medicine,
bacterial ureases may act as virulence factors that give rise
123

1
124
J Biol Inorg Chem (2012) 17:1123–1134
Scheme 1
to pathological conditions, such as peptic ulcer disease,
gastric cancer, and hepatic coma resulting from infection
of the gastrointestinal tracts (primarily with Helicobacter
pylori), as well as kidney stone formation and pyelone-
phritis, resulting from infection of the urinary tracts
Active site of urease
The active site of urease (Scheme 2) contains a binuclear
nickel center where nickel(II) ions, separated by a distance
˚
of 3.7 A, are bridged by a carbamylated lysine through its
(
chiefly with Proteus mirabilis and Ureaplasma urealyti-
O atoms. Ni(1) is further coordinated by two histidine
residues (through their N atoms), and Ni(2) by two histi-
dine residues (also through N atoms) as well as by an
aspartic acid residue (through its O atom). The Ni ions are
also bridged by a hydroxide ion (WB), which—along with
two terminal water molecules (W1 on Ni(1), W2 on Ni(2))
and another water (W3) located towards the opening of the
active site—form an H-bonded tetrahedral cluster that fills
the active-site cavity. As a result of the above ligations,
Ni(1) is pentacoordinate and Ni(2) is hexacoordinate. In
addition to the amino acid residues that are directly
involved in the architecture of the active site, functional in
the urease catalysis are also the residues that compose the
mobile flap of the site. Mainly through H-bonding, these
residues participate in substrate binding, stabilize the
catalytic transition state, and accelerate the reaction.
cum). In agriculture, by contrast, if the hydrolysis of fer-
tilizer urea by soil urease is too rapid, it can lead to the
unproductive volatilization of nitrogen, and may cause
ammonia toxicity and alkaline-induced plant damage.
Various strategies have been utilized to combat these
complications. One of them is to disable urease through
the use of inhibitors [7–9].
Several classes of compounds are known to inhibit
ureases [3], including amides and esters of phosphoric acid
[
5, 10], thiols [11], hydroxamic acids [12], phosphinic and
thiophosphinic acids [13], boric and boronic acids [14, 15],
phosphate [16], heavy metal ions [17, 18], bismuth com-
pounds [19], quinones [20, 21], and to a lesser extent H O
2
2
[
22], as well as L-ascorbic and dehydroascorbic acid in the
3
?
presence of Fe ions [23]. Due to their toxicity, however,
only few of the compounds may classify as medicinal and
agricultural agents.
Remarkably, this active site was found to be almost
completely superimposable among ureases from different
sources, including bacterial ureases from Klebsiella aer-
ogenes [24], Bacillus pasteurii [5], and Helicobacter pylori
[25], and the plant urease from Canavalia ensiformis (jack
bean) [26]. This is important, in that the conserved active
site and consequently the same catalytic mechanism allow
Thus, further to theoretical knowledge of urease bio-
chemistry, a thorough understanding of the catalytic
mechanism of the enzyme is indispensable for devising an
effective, dependable and safe manner of controlling its
activity.
(a)
(b)
Scheme 2
1
23

J Biol Inorg Chem (2012) 17:1123–1134
1125
the generalization of experimental data to all ureases,
independent of their origin.
however, problematic remains the proton transfer between
the bridging hydroxide and the distal amide group of urea.
All things considered, the proposed mechanisms of
urease catalysis contain a number of controversies that
remain to be clarified, primary among them being the urea
binding mode and the identities of both the nucleophile and
the proton donor.
Proposed reaction mechanisms for urease-catalyzed
urea hydrolysis
The currently proposed mechanisms for the urease-cata-
lyzed hydrolysis of urea are those by Karplus et al. [27] and
by Benini et al. [5], developed for K. aerogenes and
B. pasteurii urease, respectively. The mechanisms assume
that, in the active site of urease (Scheme 2), urea binds to
the more electrophilic Ni(1) ion with the oxygen atom of its
carbonyl group, owing to which the carbonyl carbon
becomes more electrophilic.
Steady-state approach to enzyme kinetics
Though at high concentrations, substrate and product
inhibitions are seen, urease typically exhibits Michaelis–
Menten kinetics [30] throughout the general scheme [31]:
k1
k2
E þ S ꢀ ES ! E þ P
ð1Þ
In the mechanism proposed by Karplus et al. [27]
kꢁ1
(
Scheme 2a), urea binds to the active site in a monoden-
where k and k are the rate constants for the formation
1
-1
tate manner only to Ni(1), with a water molecule retained
on Ni(2). Further, acting as a nucleophile, the Ni(2)-
coordinated hydroxide attacks the carbonyl atom of the
urea molecule to form a tetrahedral intermediate, from
which upon the protonation of the leaving amide group,
and dissociation of the enzyme–substrate (ES) complex,
and k is the rate constant for the breakdown of the ES
2
complex to E and P. If the steady-state approximation is
employed, the initial reaction rate is expressed as:
dP
dt
k k S
1
vmax
S
NH and carbamate are released. The authors argue that
3
2
v0 ¼
¼
E ¼
;
ð2Þ
the general acid that donates protons to the leaving NH3
is His320, located in the mobile flap of the active site.
The proposed monodentate urea binding and the sug-
gested catalytic mechanism were supported by molecular
dynamics simulations [28] and by an isotope study of the
urease-catalyzed hydrolysis of formamide [29]. Nonethe-
less, several issues associated with this mechanism remain
unclear, including the identity of a general base that would
deprotonate the Ni(2) water at the optimum pH (*7.5) for
activity, and the role of His320, which would need to be
protonated at the enzyme’s optimum pH to be able to act
kꢁ1 þ k2 þ k1S
S þ KM
where vmax = k E is the maximum reaction rate attained at
2
the saturating substrate concentration. Here, k is the first-
2
order catalytic rate constant kcat (hence vmax = k E) and E
cat
is the total enzyme concentration. KM by contrast, is the
Michaelis constant, expressed as:
k2 þ kꢁ1
KM
¼
:
ð3Þ
k1
When k ꢂ k-1, i.e. the dissociation of ES back to E ? S
2
is faster than the formation of P, K becomes the equilibrium
M
as a general acid, even though it has a pK of *6.5. To
a
constant K for the ES dissociation ES ¢ E ? S:
D
explain this, the authors assumed a reverse protonation
mechanism; however, the mechanism suffers from having
only 0.3 % of the enzyme in the protonation state optimal
for the catalysis.
kꢁ1
k1
KM
¼
¼ KD:
ð4Þ
However, when k ꢃ k-1, the Michaelis constant
2
In the other mechanism (Scheme 2b), proposed by
Benini et al. [5], a urea molecule replaces the W1–W3 water
molecules and, aside from being bound to Ni(1) through its
oxygen, it also binds to Ni(2) through the nitrogen of its
nonleaving amide group, to form an overall bidentate
binding to the metal center. The authors propose that the
nucleophile that attacks the carbonyl carbon of urea is the
bridging hydroxide, which simultaneously acts as a general
becomes:
k2
k1
KM ¼
:
ð5Þ
Significance of temperature- and pressure-dependent
studies of enzyme kinetics
One pragmatic approach to elucidating enzyme mecha-
nisms is to analyze the steady-state kinetic parameters KM
and kcat for the enzyme, measured at different temperatures
acid that delivers protons to the leaving NH molecules.
3
Upon the attack, a tetrahedral intermediate is formed that
breaks down into NH and carbamate. The authors ascribe a
3
[31] and pressures [32]. An analysis of K provides
M
minor role to His323 (His320 according to the residue
numbering used for K. aerogenes) in stabilizing the positive
charge of the leaving N in the transition state. In this
mechanism, the issue of reverse protonation is avoided,
information on how the system changes upon the formation
of the ES complex: E ? S ¢ ES, when the binding of the
substrate takes place, whereas an analysis of kcat provides
information on the activation process of the ES complex:
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1
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J Biol Inorg Chem (2012) 17:1123–1134
=
ES ? (ES–EP) , when bond reorganization leading to the
formation of the products occurs. Using temperature-
dependent measurements, the energetic characteristics of
the above reaction steps can be obtained: the thermody-
indicator assay [34] with use of phenol red (pK = 7.9 at
a
20 °C [34]). The assay makes use of an increase in the pH
of the reaction mixture caused by the formation of
ammonia during the reaction. The color of phenol red
exhibits a gradual transition from yellow to red over the pH
range 6.8 to 8.2, thus including the optimum pH of urease
activity at 7.0–7.5 [3]. The color transition is followed by
the development of absorbance at 560 nm, which was
reported to be linear between pH 6.8 and 7.7 [35]. To allow
the pH of the reaction mixture to change, we performed the
o
o
namic parameters DH and DS for the formation of ES,
and the kinetic parameters DH and DS for the formation
of the transition state (ES–EP) . By contrast, using pres-
=
=
=
sure-dependent measurements, information on volume
changes associated with the formation of ES, i.e. DV (the
b
binding volume), and with the formation of the transition
6¼
=
reactions in 5 mM HEPES (pK = 7.55 at 20 °C [36]). Of
state (ES–EP) , i.e. DV (the activation volume), can be
a
cat
key importance for the measurements performed in this
derived. Therefore, such an analysis can provide valuable
mechanistic information on the molecular nature of the
intermediate and transition states of the catalytic reaction.
For ureases, the results of temperature-dependent kinetic
analysis are scarce in the literature [30, 33], and intrigu-
ingly, disparate in value and sign—likely due to buffer
effects. In contrast, pressure-dependent analysis never has
been carried out for ureases. Therefore, given its experi-
mental potential, clearly as yet unexploited in the area of
urease research, we offer here the results of temperature-
and pressure-dependent kinetic analysis of the enzyme
study was that the pK values of both phenol red and
a
HEPES exhibit little variance with temperature and pres-
sure: for phenol red, DpK /DT = -0.006/°C [37] and
a
DpK /Dp = -0.0017/MPa [38]; for HEPES buffer, DpK /
a
a
DT = -0.014/°C [36] and DpK /Dp = 0.0008/MPa [39].
a
To choose the correct reaction time, we performed a
preliminary experiment which showed that the reaction
mixture reached a pH of 7.7 when the reaction was carried
out for 5 min at the highest urea concentration of 50 mM.
Consequently, the reaction time was set to be up to 200 s at
each urea concentration. The initial reaction rates v were
0
(
jack bean) activity performed to broaden the understand-
calculated from the slope of the linear section of the
dependence of the phenol red absorbance at 560 nm on time.
To express v in ammonia concentration units (mM NH /s),
ing of its underlying catalytic mechanism. We studied the
kinetics of the reaction using a stopped-flow technique at
temperatures between 15 and 35 °C, and at pressures
between 5 and 132 MPa—importantly—in a noninteract-
ing biological buffer (HEPES). The obtained thermody-
namic and activation parameters are discussed in terms of
the current proposals for the mechanism of this reaction.
0
3
the change in the absorbance at 560 nm was standardized
against the NH concentration assayed by the colorimetric
3
phenol-hypochlorite method [40], for which the calibration
curve was determined independently in 5 mM HEPES at pH
6.84 [41]. The dependence of the absorbance at 560 nm on
the NH concentration was linear up to 1.6 mM NH . By
3
3
contrast, the concentration of the enzyme urease in the
reaction mixture was assessed from the determined activity
of the Sigma product, using the activity of the pure enzyme
reported to be 6,200 units/mg enzyme on average [6] and the
enzyme molecular weight 545.34 kDa [42]. The calculated
Materials and methods
Materials
Urease (from jack beans, type III, nominal activity 45 U/
mg solid), urea (for Molecular Biology), and HEPES buffer
-
7
concentration amounted to 2.305 9 10 mM, and all
subsequent calculations performed in this study refer to the
concentration of urease hexamers.
(
SigmaUltra) were from Sigma (St. Louis, MO, USA).
EDTA and phenol red were from POCh (Gliwice, Poland).
HEPES buffer 5 mM, pH 6.84, was prepared by diluting a
stock 200 mM HEPES solution (pH 7.33) and adding
Stopped-flow measurements of urease kinetics
1 mM EDTA. Ultrapure water (resistivity 18.2 MX cm)
from a Simplicity 185 water purification system (Millipore,
Temperature-dependent kinetic measurements were per-
formed with a SX20 stopped-flow spectrometer from
Applied Photophysics Ltd. (Leatherhead, UK), whereas the
pressure-dependent measurements, with a custom-built high-
pressure stopped-flow reactor described previously [43]. For
the former, the instrument was maintained at atmospheric
pressure at five temperatures in the range 15–35 °C, and for
the latter, at six pressures in the range 5–132 MPa at 25 °C.
Both instruments were thermostated to ± 0.1 °C.
Billerica, MA, USA) was used throughout.
Urease assay
Given the fact that in the stopped-flow instrument the
reaction mixtures are enclosed within the instrument and
samples cannot be withdrawn for analysis, for the mea-
surements of the urease reaction rates we chose a pH
1
23

J Biol Inorg Chem (2012) 17:1123–1134
1127
To perform the measurements, the following solutions
were prepared in 5 mM HEPES buffer at pH 6.84 con-
taining 1 mM EDTA: urease 0.04 mg/mL, phenol red
0
0
0
0
0
.4
.3
.2
.1
.0
0
2
.0267 mg/mL, and urea at concentrations between 4 and
00 mM. Next, the phenol red solution was mixed 1:1 with
a urea solution of preselected concentration, and the
obtained solution was placed in one syringe of the stopped-
flow reactor while the other syringe was loaded with the
urease solution. The solutions were conditioned in the
reactor at a chosen temperature/pressure for 20 min before
being mixed 1:1 to initiate the reaction. After mixing, the
concentrations in the reaction mixture were: urease
-
3
0
.02 mg/mL, phenol red 6.675 9 10 mg/mL, and urea
between 1 and 50 mM. The measurements were performed
in triplicate and the results were averaged.
4
00
500
600
wavelength, nm
The reaction rates v measured at various temperatures
0
and pressures were further used to calculate the steady-
state parameters of urease, KM and vmax, which was done
by the nonlinear least-square fitting of the measured v0
values to the Michaelis–Menten equation (Eq. 2). The
catalytic constant kcat was obtained by dividing vmax by the
enzyme concentration.
Fig. 1 Typical UV–vis spectra of phenol red recorded during a
kinetic run for the urease-catalyzed hydrolysis of urea in 5 mM
HEPES buffer, pH 6.84, under ambient conditions with 50 mM urea
(
a PerkinElmer Lambda 35 UV–vis spectrophotometer was used)
reported to occur in the range 45–65 °C on average [46],
temperatures [35 °C were not used in the present experi-
ments to avoid thermal denaturation of the enzyme [47].
Results and discussion
The urease saturation curves, v versus S, are presented
0
It was assumed in this work that the urease-catalyzed
hydrolysis of urea proceeds in two steps according to the
Michaelis–Menten mechanism (Eq. 1). The first step is the
formation of a stable urease–urea complex, governed by an
equilibrium constant equal to the inverse of KD (Eq. 4).
Importantly, when analyzing the data obtained for this step,
we followed the suggestion made in the literature [44]—
based on the insignificant variability of KM as a function of
in Fig. 2. They show that the reaction followed Michaelis–
Menten kinetics (Eq. 2) at each studied temperature. The
kinetic parameters derived from the curves, K
and kcat
M
(Table 1), were found to be consistent in magnitude with
those reported in the literature [44, 45, 48], thus proving
that the analytical conditions chosen for the present study
were correct. The results obtained show that, as is typically
observed for enzymatic reactions, increasing the tempera-
ture increased the reaction rate. Accordingly, the kcat values
pH [16, 45]—that urease features KM = K . By contrast,
D
the second step of the enzymatic reaction involves the
activation of the urease–urea complex and its subsequent
decomposition into products and free enzyme, and is
grew, increasing twofold between 15 and 35 °C. The K
M
value, on the other hand, in contrast to a previous report of
temperature independence [45], showed a slight increase
from 3.3 to 4.6 mM, thus indicating a small reduction in
the enzyme’s affinity for the substrate at higher tempera-
tures. Apparently, this reduction in affinity can be consid-
ered as resulting from the loosening of the active site
structure, whose strict architecture is required for the
catalysis.
governed by the catalytic rate constant k . The effects of
cat
temperature and pressure on the two governing constants
were determined and will be discussed.
In this study, the urease activity was assayed using
phenol red. Typical UV–vis spectra for the indicator
recorded during a kinetic run performed under ambient
conditions with 50 mM urea are presented in Fig. 1. The
rise in the absorbance at 560 nm over time was used to
obtain v0.
Effect of temperature on K
M
Since the binding step E ? S ¢ ES in enzymatic reactions
is defined by the equilibrium constant, determining the
temperature effects on the constant yields the classical
Temperature-dependent measurements
o
o
o
The kinetics of urease was studied under atmospheric
pressure at temperatures between 15 and 35 °C. Although
the optimum temperature for urease activity has been
thermodynamic functions DH , DS , and DG (the standard
enthalpy, entropy, and free energy change of the reaction).
The equilibrium constant (K) and the thermodynamic
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1
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J Biol Inorg Chem (2012) 17:1123–1134
0
0
0
.02
.01
.00
occur when the substrate forms the complex ES, such as a
movement of the mobile flap that opens up the active site of
urease for urea binding.
3
08 K
3
2
03 K
98 K
Effect of temperature on kcat
2
2
93 K
88 K
The temperature dependence of kcat provides insight into
the energetic characteristics of the activation process
=
ES ? (ES–EP) . This process (the catalytic step of the
reaction) represents the bond-breaking and/or bond-making
step of the reaction, leading to the formation of products. It
0
20
40
60
is characterized by the apparent activation energy E and
a
urea, mM
the changes in the thermodynamic activation functions
=
=
=
DH , DS , and DG .
The apparent activation energy E for the catalytic step
of the urease reaction was derived from the Arrhenius
0
Fig. 2 Plots of v versus urea concentration for the urease-catalyzed
hydrolysis of urea at temperatures between 15 and 35 °C
a
equation:
Table 1 Kinetic parameters of urease measured at different tem-
peratures and ambient pressure
Ea
kcat ¼ A eꢁRT
;
ð9Þ
3
9 10 (M)
-4
cat 9 10 (1/s)
T (K)
K
M
k
which gave Ea = 24 ± 1 kJ/mol. This value corresponds
well to the literature values reported for jack bean urease,
which are 18–30 kJ/mol on average [46]. To allow a
288
293
298
303
308
3.3 ± 0.2
3.7 ± 0.2
3.9 ± 0.2
4.1 ± 0.2
4.6 ± 0.1
4.5 ± 0.1
5.2 ± 0.1
6.1 ± 0.1
7.2 ± 0.1
8.7 ± 0.1
comparison with the literature data, the value of E for
a
1
mM urea was also calculated. The reaction performed
under these conditions has a lower E = 15 ± 1 kJ/mol,
a
which is due to the fact that v at a low urea concentration is
a composite of KM and k , whereas at high urea concen-
cat
tration the E is controlled solely by k . Interestingly, a
a cat
functions are related as expressed by the following
similar urea concentration effect on E was reported for
a
equations:
jack bean urease in THAM buffer [30], but the opposite
was observed in phosphate buffer [33], buffer effects being
the likely reason for these diverse observations.
o
DG ¼ ꢁRT ln K;
ð6Þ
ð7Þ
o
o
o
and DG ¼ DH ꢁ TDS ;
o
o
The rate constant kcat was further analyzed to determine
the activation enthalpy and entropy for the catalytic step.
To do this, the absolute rate theory was used, where the
temperature dependence of the rate constant is expressed
by the Eyring–Polanyi equation:
DH
RT
DS
hence ꢁ ln K ¼
ꢁ
:
ð8Þ
R
o
o
The DH and DS values can be derived from a linear
plot of -ln K versus 1/T based on Eq. 8, and DG based on
Eq. 7. Such a plot was constructed for the dependence of
o
6
6
k
DH 1
R T
kB DS
the inverse of the Michaelis constant 1/K for urease on
M
ln ¼ ꢁ
þ ln h
þ
;
ð10Þ
o
o
o
T
R
1
/T (Fig. 3a). The resulting parameters DH , DS , and DG
298
are compiled in Table 2. As shown, the thermodynamic
parameters for the formation of the urease–urea complex
have favorable values; the reaction is exothermic and
spontaneous, and accompanied by a gain in entropy. This
entropy gain, however, should be interpreted with caution,
since its value is rather small, and it may be a composite of
many different contributions that partially cancel each other
out. Nevertheless, it may be ascribed, for instance, to the
displacement of the ordered water cluster in the active site of
urease by a molecule of urea (Scheme 2). Such release of
ordered water from proteins has been demonstrated to be
responsible for an increase in entropy [49]. Likewise, it may
be ascribed to conformational changes in the enzyme that
where R is the gas constant, T is the absolute temperature, h
is the Planck constant, and k is the Boltzmann constant.
B
=
The activation enthalpy DH was extracted from the slope
of the Eyring plot, ln (k /T) versus 1/T, and the activation
cat
=
entropy DS was derived from the intercept.
The Eyring plot for the reaction studied here is pre-
sented in Fig. 3b, and the resulting values are listed in
Table 3. As shown, the reaction is characterized by a
=
=
=
positive DH , a negative DS , and a positive DG . The
=
values of both the activation enthalpy DH (22 kJ/mol)
=
and the entropy DS (-80 J/K mol) are such that they are
unfavorable for lowering the activation free energy
=
DG
(45 kJ/mol) to accelerate the reaction. The large
98
2
1
23

J Biol Inorg Chem (2012) 17:1123–1134
1129
Fig. 3 Effect of temperature in
the range 15–35 °C on a the
Michaelis constant K and
b kcat/T (Eyring plot) for the
urease-catalyzed hydrolysis of
urea
(b) 5.8
(
a) -5.2
M
5.6
5.4
5.2
-
-
-
5.4
5.6
5.8
0
5.0
0.0032
.0032
0.0033
0.0034
0.0035
0.0033
0.0034
0.0035
/T, K-1
1/T, K-1
1
Table 2 Thermodynamic parameters for the formation of the urease–
urea complex in the urease-catalyzed hydrolysis of urea, obtained
from temperature-dependent measurements of K
does not involve the release of ordered water molecules
(Scheme 3).
M
The activation data obtained in this study (Table 3)
o
o
o
98
DH (kJ/mol)
12 ± 1
DS (J/mol K)
DG2 (kJ/mol)
diverge in value and sometimes in sign from those reported
in previous temperature-dependent studies of urease
carried out in THAM [30] and phosphate buffer [33],
which is likely due to buffer-dependent effects. Impor-
tantly, since HEPES is a noninteracting biological buffer
-
7 ± 3
-14 ± 2
Table 3 Activation parameters for the urease–urea complex in the
urease-catalyzed hydrolysis of urea, obtained from temperature-
dependent measurements of kcat
[
41, 50], the results collected in this study can be regarded
as being independent of buffer effects.
=
=
6¼
298
=
E
a
(kJ/mol)
DH (kJ/mol)
DS (J/K mol)
DG (kJ/mol)
Interestingly, the magnitudes of the k -derived DH
cat
=
and DS values for the urease reaction were found to be
comparable with those reported for the hydrolysis of
p-nitrophenyl sulfate catalyzed by arylsulfatase [51].
Therein, it was suggested that such values are supportive of
the notion that the activation process consists of an asso-
ciation or interchange rather than of a dissociation. This we
2
4 ± 1
22 ± 1
-80 ± 3
45 ± 2
=
negative value of DS also indicates that the transition
state is more orderly than the ground state of the reactants,
thus suggesting that the formation of the transition state
(
a)
(b)
(e)
(d)
(c)
Scheme 3
123

1
130
J Biol Inorg Chem (2012) 17:1123–1134
experimental results correctly, it is crucial to identify
whether the effects observed originated from the former or
the latter process [32, 53–55, 58].
Jack bean urease is a large protein made up of six identical
subunits, each of molecular mass 90.77 kDa, which are
assembled into a hexamer a [3]. The mass of the hexamer
6
with the 12 nickel ions included is thus 545.34 kDa [42].
In this context, to verify whether our results were an
effect of pressure on urease catalysis and not due to the
enzyme deactivation by denaturation, that is, weather ure-
ase was in the active form during the pressure measure-
ments, we took the following arguments into consideration:
1.
In a separate experiment, we checked whether sub-
jecting urease to high pressure for the overall duration
of the high-pressure measurements caused a loss of
activity through denaturation. In our stopped-flow
system, the enzyme solution once set in the apparatus,
was subjected to a stepwise increase in pressure
(5–132 MPa) until the end of the kinetic measurements
performed at each pressure for a series of urea
concentrations. It took ca. 2 h to complete the entire
experiment. Therefore, to check whether the enzyme
was or was not denatured within the duration of the
stopped-flow experiment, typical urease samples in
5 mM HEPES buffer were pressurized at 132 MPa for
Fig. 4 Schematic energy diagram for the urease-catalyzed hydrolysis
of urea
find applicable to both of the proposed mechanisms of urea
hydrolysis (Scheme 2).
Taken together, based on the results obtained in this study
and the literature data [31, 52], we propose that the urease-
catalyzed hydrolysis of urea follows the energy diagram
=
=
o
illustrated in Fig. 4, where DH = DH (k ) - DH (K ).
cat
M
Pressure-dependent measurements
2
h, and their activities were assayed immediately after
Elevated pressure affects enzymes in a complex manner by
perturbing the intra- and intermolecular weak, noncovalent
interactions that are involved in protein conformation and
solvation, and thus responsible for enzyme structure and
activity [32, 53–55]. Depending on its magnitude, the
pressure may affect protein structures at quaternary, tertiary,
and secondary levels with concomitant reductions in their
activity, finally leading to denaturation [56]. Thus, there is
an upper limit on the pressure that enzymes can endure in the
active form. While moderate pressures of 100–200 MPa
typically cause the dissociation of oligomeric proteins into
subunits, the tertiary protein structures are destroyed by
higher pressures of 400–800 MPa, and secondary structures
are unaffected by pressures as high as 1,000 MPa. By con-
trast, the primary structures, which are maintained by
covalent bonds, are not pressure sensitive at ordinary tem-
peratures [56, 57]. In addition to obvious structural changes,
pressure-induced disturbances of the intra- and intermolec-
ular interactions that occur in enzymes may also cause
modifications to the kinetics of enzyme-catalyzed reactions.
As a result, measuring enzyme kinetic parameters under
increasing pressure provides access to the analysis of the
elementary steps of enzyme reactions [32, 53–55].
depressurization. The results showed that the pressure
did not change the activities of the samples. Obviously,
this means that the pressure did not bring about an
irreversible loss of activity due to denaturation.
However, this could also mean that (1) the enzyme
dissociates into subunits under pressure but rapidly re-
associates without denaturation when depressurized,
(2) the enzyme dissociates permanently, but the
combined activity of the subunits is the same as that
of the hexamer, or (3) there is no dissociation of the
enzyme into subunits under applied pressure.
2. Importantly, it was shown in studies on the chemical
denaturation of urease to half-units [59] and subunits
[60] that the quaternary structure of urease is not
required for catalytic activity, and—more importantly–
that the subunit is not only the fundamental unit for the
quaternary structure of urease but also for its activity.
3. The crystal structures of ureases revealed that the active
sites are always located in the a subunits of the enzyme,
and are entirely independent [26]. This physical inde-
pendence of the sites thus supports the notion that the
monomeric form of jack bean urease should be active.
4. The fact that the Michaelis constant K of urease shows
M
As enzyme denaturation (subunit dissociation and/or
unfolding) may occur under pressure simultaneously with
the changes in enzyme kinetics, in order to interpret the
little variation with pressure (Table 4) apparently con-
firms that the 3D structure of the enzyme responsible for
its activity is practically unperturbed.
1
23

J Biol Inorg Chem (2012) 17:1123–1134
1131
Table 4 Kinetic parameters of urease measured at different pressures
at 25 °C
0.02
3
9 10 (M)
-4
kcat 9 10 (1/s)
p (MPa)
K
M
5
MPa
5
5.0 ± 0.2
5.3 ± 0.3
5.5 ± 0.4
4.3 ± 0.4
4.7 ± 0.8
4.1 ± 0.8
6.4 ± 0.1
6.3 ± 0.1
6.0 ± 0.1
4.2 ± 0.1
3.1 ± 0.1
2.1 ± 0.1
4
0 MPa
1
4
7
1
1
0
0
0
.01
.00
7
1 MPa
0
1
1
01 MPa
32 MPa
1
01
32
0
20
40
60
Table 5 Binding and activation volumes in the urease-catalyzed
hydrolysis of urea, obtained from pressure-dependent measurements
performed at 25 °C
urea, mM
Fig. 5 Plots of v versus urea concentration for the urease-catalyzed
0
hydrolysis of urea at pressures between 5 and 132 MPa and 25 °C
=
6
p (MPa)
DV
(
b
(K
mL/mol)
M
)
M
DV (K /kcat)
DV (kcat
(
)
cat
(mL/mol)
DV p
mL/mol)
K ¼ Ko eꢁ RT
;
ð11Þ
\
40
-2 ± 2
-2 ± 2
5 ± 1
28 ± 1
9 ± 4
where DV is the reaction volume, i.e. the excess volume of
products over reactants. The volume can be derived from
the slope of the linear plot of lnK versus p (Eq. 11).
To determine the reaction volume for the binding step
[40
26 ± 3
5
.
Furthermore, it has been reported that pressure-
induced protein denaturation is always accompanied
by large negative DV values [55]; those for the
dissociation of oligomeric proteins are typically neg-
ative and relatively large between -50 and -200 mL/
mol [54], whereas our values clearly do not fall within
this range (Table 5).
E ? S ¢ ES of the urease reaction (DV ), a plot of the
inverse of the Michaelis constant 1/KM versus p was drawn
b
(
Fig. 6a), and the binding volume DV was found to be
2 ± 2 mL/mol (Table 5). Although close to zero, this
b
-
small negative value may suggest that the system shrinks
slightly upon ES formation, due for instance to the
expulsion of water from the active site into the bulk
Based on the discussion above, we concluded that the
(
Scheme 2). Most interestingly, however, the insignificant
variance of KM with pressure proves that pressures up to
32 MPa barely have an impact on the architecture of the
pressure applied in this study did not denature urease, and
that even if dissociated into subunits the enzyme retained
its activity. We used this as the foundation for a further
analysis of the effects of pressure on the kinetic parameters
of urease.
1
active site of urease, and that the binding of urea to the site
is practically unperturbed.
The saturation curves for urease obtained in the studied
pressure range 5–132 MPa at 25 °C are presented in Fig. 5.
The curves are consistent with Michaelis–Menten kinetics
Effect of pressure on kcat
As predicted by the absolute rate theory, the dependence of
(
Eq. 2) at each pressure. The corresponding KM and kcat
the reaction rate constant k on pressure p is expressed by
[32, 58]:
values are listed in Table 4. Figure 5 shows that increasing
the pressure reduced the reaction rate. Correspondingly, the
kcat values decreased with increasing pressure (approxi-
mately threefold between 5 and 132 MPa; Table 4), but,
remarkably, the values of KM hardly changed. This shows
that the studied range of pressures had no significant
impact on the affinity of urease for the substrate.
DV 6¼ p
k ¼ k eꢁ RT
;
ð12Þ
o
=
where DV is the activation volume. In principle, the
activation volume is the difference between the volume of
reactants and their volume in the transition state of the
=
reaction. For enzymatic reactions, DV refers to the cata-
lytic step ES ? (ES–EP) , and is determined experi-
mentally from the slope of the linear plot of ln kcat versus p
=
Effect of pressure on KM
(Eq. 12); however, as will be argued later, interpreting it
The effect of pressure on a chemical equilibrium at a con-
may not be as simple as defining it.
stant temperature is to shift the existing equilibrium (K ) to a
0
In point of fact, consisting of contributions from both the
=
new position, wherein the equilibrium constant K changes as
a function of pressure p as expressed by [32, 58]:
catalytic and binding step, the overall activation volume DV
has a value that depends on substrate concentration [32].
123

1
132
J Biol Inorg Chem (2012) 17:1123–1134
Fig. 6 Effect of pressure in the
range 0.1–132 MPa and at
b) 11.2
(
(
a) 6.0
2
5 °C on a the Michaelis
1
1
1
0.8
0.4
0.0
constant K and b kcat for the
M
5
5
4
4
.6
.2
.8
.4
urease-catalyzed hydrolysis of
urea. Points for 0.1 MPa were
taken from the temperature-
dependent measurements
performed at ambient pressure
at 25 °C (Table 1)
0
50
100
150
0
50
100
150
p, MPa
p, MPa
If the Michaelis–Menten mechanism is assumed for the
enzyme reaction (Eq. 1), the initial reaction rate expressed
by Eq. 2, when differentiated with respect to pressure,
becomes:
KM
6
¼
6¼
DV ¼ DV
ꢁ
DVb:
ð13Þ
cat
KM þ S
Equation 13 reveals that, upon decreasing the substrate
=
concentration S, the binding contribution to DV increases
=
6¼
up to the limit DV = DV - DV , whereas at the
cat
b
saturating substrate concentration the contribution of the
6¼
=
=
binding volume becomes negligible and DV = DV ,
cat
which is the case analyzed here. Note that the overall DV
can be obtained independently from the dependence of
Fig. 7 Schematic volume profile for the urease-catalyzed hydrolysis
of urea
ln (K /k ) on p (Eq. 11 divided by Eq. 12, Table 5).
M
cat
The plot of ln kcat versus p for the studied urease–urea
system is presented in Fig. 6b. The effect of pressure on
kcat was found to be biphasic. Though not quite typical,
such behaviour of enzymatic systems is not infrequent
site is not particularly perturbed by elevated pressures.
Therefore, in order to account for the observed effect of
pressure, we need to analyze the suggested reaction
mechanisms in more detail.
[
53, 61]. In the case of urease, the kcat value of the enzyme
decreased over the whole of the pressure range studied. The
decrease up to 40 MPa was only to 94 % of the initial
value, but the decrease was threefold upon going from 40
It is clear from the analysis of the coordination spheres
of the Ni(1) and Ni(2) centers in the active site of urease
(Scheme 2) that the pentacoordinate Ni(1) is more elec-
6¼
to 132 MPa. The resulting activation volume DV chan-
cat
trophilic than the hexacoordinate Ni(2), which explains
why urea first binds via the O-donor to Ni(1) before its
amide group interacts with Ni(2). Furthermore, from the
presence of the negatively charged coordinated ligands
ged from 5 ± 1 mL/mol at p \ 40 MPa to 28 ± 1 mL/
mol at p [ 40 MPa (Table 5). Compared to the binding
volume DV obtained from K , these results prove that the
b
M
variation of enzyme activity with pressure is mainly due to
the catalytic step of the reaction. In summary, we propose a
volume profile for the urease-catalyzed hydrolysis of urea
(
Lys–NH–CO , bridging OH, and O–Asp), it can be
2
inferred that the pK of water–Ni(1) is considerably lower
a
than that of water–Ni(2), which is due to the extra negative
charge from O–Asp on Ni(2). However, more importantly,
(
Fig. 7), which schematically presents the volume changes
6¼
=
that occur along the reaction coordinate, where DV =
DV ? DVb.
as it is bound to two Ni centers, the pK of the bridging OH
a
cat
group is expected to be lower than that of the OH bound to
either Ni(1) or Ni(2). For that reason, the reaction path
outlined in Scheme 2b seems more convincing to account
for the catalytic hydrolysis, where the bridging hydroxo
group supplies the proton required to initiate the process
[5, 62]. In terms of the arguments outlined above, the
Various pressure-induced changes in enzyme systems
can be considered to be responsible for the nonlinear
pressure dependence of enzyme kinetic parameters. These
include, in addition to protein structural changes, com-
pressibility changes and changes in the rate-determining
step [53]. As argued earlier, urease retains its activity even
in the dissociated form, and we also showed that the active
experimentally observed pK values of 5.3 and 6.6 can be
a
viewed as being related to the carboxyl and imidazole
1
23

J Biol Inorg Chem (2012) 17:1123–1134
1133
groups at the active site [16], while the pK value of 9.1
a
which involves the release of three water molecules during
the binding of urea to the active site. In general, deproto-
nation equilibria are characterized by negative reaction
volumes due to an increase in electrostriction as a result of
charge creation. Since the studied reaction is accelerated by
the deprotonation of the carboxyl and imidazole groups in
the lower pH range, their deprotonation is expected to be
accompanied by a negative reaction volume, which could,
in principle, account for the weaker effect of pressure
observed in the low-pressure range (0.1–40 MPa), since the
expected volume increase due to the reaction A ? B will
be partially canceled out by the negative reaction volume
associated with the deprotonation process. However, at
higher pressures, this contribution is expected to be can-
celed out by the effect of the selected buffer, since the latter
is independent of pressure and will stabilize the pH of the
solution to prevent further deprotonation of these groups at
higher pressures in the range 40–132 MPa.
could be related to the deprotonation of the bridging OH
group [16]. The latter step then accounts for the proton
transfer following the nucleophilic attack of the bridging
OH on the urea amide group to form NH . The above
3
conclusions are in keeping with the urease pH–activity
profile with an optimum pH of *7.5 [16].
Given the results of this work on the catalytic activity of
urease, and those reported in the literature [5, 27, 62], we
fully support the overall reaction mechanism put forward
by Benini et al. [5], in which the bridging OH—not the
terminal W2—is a nucleophile (Scheme 2b). This mecha-
nism, proposed in [5, 62], is illustrated in Scheme 3 [62].
In the mechanism, in step A ? B urea enters the active site
when the flap is open, to bind to Ni(1) via the carbonyl
O-donor, which involves the release of three water mole-
cules. During step B ? C, the flap closes and the urea NH2
coordinates to Ni(2). This is followed by nucleophilic
attack by the bridging OH to produce the tetrahedral
intermediate D. A proton transfer then occurs in step
We therefore conclude from the above interpretation of
the high-pressure kinetic results obtained in this study that
the data corroborate the catalytic mechanism proposed by
Benini et al. [5], which is outlined in Scheme 2b and
described in more detail in Scheme 3 [62].
?
D ? E to form C–NH3 , which is stabilized by the neutral
imidazole of His323 from the active-site flap. Finally, C–N
bond cleavage occurs to release NH and carbamate in the
3
last step E ? A, which is accompanied by flap opening
and the uptake of four water molecules to yield A.
Acknowledgments This work was supported by DS WCh/43 from
the Faculty of Chemistry of the Jagiellonian University, Krak o´ w,
Poland (BK), and the Deutsche Forschungsgemeinschaft, Germany
In general, it should be kept in mind that interpreting the
activation volume is not always a straightforward task,
because the experimentally obtained value could be the
sum of three contributions: an intrinsic contribution that
arises from structural volume changes in the molecules due
to bond formation and bond scission processes; a solva-
tional contribution resulting from the rearrangement of
water molecules during the reaction, which is especially
pronounced when charge and dipole changes occur in the
reacting molecules; and a conformational contribution
associated with changes in the conformation of the enzyme
that accompany substrate binding and chemical steps. In
(
RvE). The research at JU was carried out within the Coordination and
Bioinorganic Physicochemistry Group (head: Prof. Gra z_ yna Stochel)
using equipment financed by the European Regional Development
Fund within the framework of the Polish Innovation Economy Oper-
ational Program (contract no. POIG.0 2.01.00-12-0 23/08).
Open Access This article is distributed under the terms of the
Creative Commons Attribution License which permits any use, dis-
tribution, and reproduction in any medium, provided the original
author(s) and the source are credited.
terms of the observed pressure dependence of k , the
cat
References
binding of urea and the release of three water molecules in
step A ? B can be accompanied by a significant overall
volume increase in the transition state. The flap closure
process in step B ? C, on the other hand, is expected to be
accompanied by a significant volume decrease. The sub-
sequent steps C ? D ? E, involving intramolecular
nucleophilic attack and proton transfer, are not expected to
be accompanied by significant volume changes. In the final
step E ? A, the volume increase associated with the
1
. Dixon NE, Gazzola C, Blakeley RL, Zerner B (1975) J Am Chem
Soc 97:4131–4133
2
3
4
5
. Follmer C (2008) Phytochemistry 69:18–28
. Krajewska B (2009) J Mol Catal B Enzym 59:9–21
. Sumner JB (1926) J Biol Chem 69:435–441
. Benini S, Rypniewski WR, Wilson KS, Miletti S, Ciurli S,
Mangani S (1999) Structure 7:205–216
. Hausinger RP, Karplus PA (2001) In: Wieghardt K, Huber R,
Poulos TL, Messerschmidt A (eds) Handbook of metalloproteins.
Wiley, Chichester, pp 867–879
6
?
-
release of NH4 and NH COO should mostly be canceled
2
7. Follmer C (2010) J Clin Pathol 63:424–443
8. Kosikowska P, Berlicki Ł (2011) Expert Opin Ther Pat
out by the volume decrease associated with the uptake of
four water molecules, so this step is not expected to con-
tribute meaningfully to the observed pressure effect. Thus,
the significant decrease in kcat with increasing pressure can
mainly be ascribed to the first step of the catalytic cycle,
2
. Mulvaney RL, Bremner JM (1981) Soil Biochem 5:153–196
1:945–957
9
1
0. Dom ´ı nguez MJ, Sanmart ´ı n C, Font M, Palop JA, San Francisco S,
Urrutia O, Houdusse F, Garc ´ı a-Mina J (2008) J Agric Food Chem
56:3721–3731
123

1
134
J Biol Inorg Chem (2012) 17:1123–1134
1
1
1
1
1
1. Benini S, Rypniewski WR, Wilson KS, Ciurli S, Mangani S
37. Kolthoff IM (1921) Recl Trav Chim Pay-B 40:775–785
38. Raghuraman B, Gustavson G, Mullins OC, Rabbito P (2006)
AIChE J 52:3257–3265
39. Kitamura Y, Itoh T (1987) J Solution Chem 16:715–725
40. Weatherburn MW (1967) Anal Chem 39:971–974
41. Krajewska B, Chudy M, Drozdek M, Brz o´ zka Z (2003) Elec-
troanalysis 15:460–466
(
1998) J Biol Inorg Chem 3:268–273
2. Odake S, Morikawa T, Tsuchiya M, Imamura L, Kobashi K
1994) Biol Pharm Bull 17:1329–1332
(
3. Vassiliou S, Grabowiecka A, Kosikowska P, Yiotakis A, Kafarski
P, Berlicki Ł (2008) J Med Chem 51:5736–5744
4. Krajewska B, Zaborska W, Leszko M (1997) J Mol Catal B
Enzym 3:231–238
5. Reddy KRC, Kayastha AM (2006) J Enzym Inhib Med Chem
42. Takishima K, Suga T, Mamiya G (1988) Eur J Biochem
175:151–165
2
1:467–470
43. van Eldik R, Gaede W, Wieland S, Kraft J, Spitzer M, Palmer DA
(1993) Rev Sci Instrum 64:1355–1357
44. Lopreore C, Byers LD (1998) Arch Biochem Biophys
349:299–303
45. Dixon NE, Riddles PW, Gazzola C, Blakeley RL, Zerner B
(1980) Can J Biochem 58:1335–1344
46. Krajewska B (2009) J Mol Catal B Enzym 59:22–40
1
1
1
6. Krajewska B, Ciurli S (2005) Plant Physiol Biochem 43:651–658
7. Krajewska B (2008) J Enzym Inhib Med Chem 23:535–542
8. Zaborska W, Krajewska B, Leszko M, Olech Z (2001) J Mol
Catal B Enzym 13:103–108
9. Zhang L, Mulrooney SB, Leung AFK, Zeng Y, Ko BBC,
Hausinger RP, Sun H (2006) Biometals 19:503–511
0. Zaborska W, Krajewska B, Kot M, Karcz W (2007) Bioorg Chem
1
2
2
ˇ
47. Illeov a´ V, Polakovi cˇ M, Stefuca V, A cˇ ai P, Juma M (2003) J
Biotechnol 105:235–243
3
5:233–242
1. Krajewska B, Zaborska
5:4144–4151
W
(2007) Bioorg Med Chem
48. Blakeley RL, Webb EC, Zerner
8:1984–1990
B (1969) Biochemistry
1
2
2
2. Krajewska B (2011) J Mol Catal B Enzym 68:262–269
3. Krajewska B, Brindell M (2011) J Enzyme Inhib Med Chem
49. Truong K, Su Y, Song J, Chen Y (2011) Biochemistry
50:5757–5766
2
6:309–318
4. Jabri E, Carr MB, Hausinger RP, Karplus PA (1995) Science
68:998–1004
50. McGlothlin CD, Jordan J (1976) Anal Lett 9:245–255
51. Stawoska I, Gaw e˛ da S, Bielak-Lakomska M, Brindell M,
Lewi n´ ski K, Laidler P, Stochel G (2010) J Coord Chem
63:2472–2487
52. Jespersen ND (1975) J Am Chem Soc 97:1662–1667
53. Masson P, Balny C (2005) Biochim Biophys Acta 1724:440–450
54. Boonyaratanakornkit BB, Park CB, Clark DS (2002) Biochim
Biophys Acta 1595:235–249
2
2
2
5. Ha N-C, Oh S-T, Sung JY, Cha KA, Lee MH, Oh B-H (2001)
Nature Struct Biol 8:505–509
6. Balasubramanian A, Ponuraj K (2010) J Mol Biol 400:274–283
7. Karplus PA, Pearson MA, Hausinger RP (1997) Acc Chem Res
2
2
3
0:330–337
2
8. Manunza B, Deiana S, Pintore M, Gessa C (1999) Soil Biol
Biochem 31:789–796
55. Mozhaev VV, Heremans K, Frank J, Masson P, Balny C (1996)
Proteins Struct Funct Genet 24:81–91
2
3
3
3
3
3
3
9. Marlier JF, Cleland WW (2006) Biochemistry 45:9940–9948
0. Wall MC, Laidler KJ (1953) Arch Biochem Biophys 43:299–306
1. Segel IH (1975) Enzyme kinetics. Wiley, New York
2. Morild E (1981) Adv Protein Chem 34:93–163
3. Laidler KJ, Hoare JP (1950) J Am Chem Soc 72:2489–2494
4. Ru ´ı z-Herrera J, Gonzalez J (1969) Anal Biochem 31:366–374
5. Sheppard NF Jr, Guiseppi-Elie A (1999) Chapter 71. In: Webster
JG (ed) The measurement, instrumentation and sensors hand-
book. CRC, Boca Raton
56. Silva JL, Foguel D, Royer CA (2001) Trends Biochem Sci
26:612–618
57. Balny C (2004) J Phys Condens Matter 16:S1245–S1253
58. Northop DB (2002) Biochim Biophys Acta 1595:71–79
59. Contaxis CC, Fishbein WN (1971) J Biol Chem 246:677–685
60. Hirai M, Kawai-Hirai R, Hirai T, Ueki T (1993) Eur J Biochem
215:55–61
61. Dallet S, Legoy M-D (1996) Biochim Biophys Acta 1294:15–24
62. Zambelli B, Musiani F, Benini S, Ciurli S (2011) Acc Chem Res
44:520–530
3
6. Good NE, Winget GD, Winter W, Connoly TN, Izawa S, Singh
RMM (1966) Biochemistry 5:467–477
1
23