Determination of the catalytic activity of binuclear metallohydrolases using isothermal titration calorimetry

Article abstract of DOI:10.1007/s00775-013-1079-0

Binuclear metallohydrolases are a large and diverse family of enzymes that are involved in numerous metabolic functions. An increasing number of members find applications as drug targets or in processes such as bioremediation. It is thus essential to have

Full text of DOI:10.1007/s00775-013-1079-0

J Biol Inorg Chem (2014) 19:389–398
DOI 10.1007/s00775-013-1079-0
ORIGINAL PAPER
Determination of the catalytic activity of binuclear
metallohydrolases using isothermal titration calorimetry
•
Marcelo M. Pedroso Fernanda Ely
•
•
Thierry Lonhienne Lawrence R. Gahan
•
•
David L. Ollis Luke W. Guddat Gerhard Schenk
•
Received: 15 August 2013 / Accepted: 12 December 2013 / Published online: 12 January 2014
Ó SBIC 2014
Abstract Binuclear metallohydrolases are a large and
diverse family of enzymes that are involved in numerous
metabolic functions. An increasing number of members
find applications as drug targets or in processes such as
bioremediation. It is thus essential to have an assay avail-
able that allows the rapid and reliable determination of
relevant catalytic parameters (kcat, Km, and kcat/Km). Con-
tinuous spectroscopic assays are frequently only possible
by using synthetic (i.e., nonbiological) substrates that
possess a suitable chromophoric marker (e.g., nitrophenol).
Isothermal titration calorimetry, in contrast, affords a rapid
assay independent of the chromophoric properties of the
substrate—the heat associated with the hydrolytic reaction
can be directly related to catalytic properties. Here, we
demonstrate the efficiency of the method on several
selected examples of this family of enzymes and show that,
in general, the catalytic parameters obtained by isothermal
titration calorimetry are in good agreement with those
obtained from spectroscopic assays.
Keywords Isothermal titration calorimetry ꢀ Kinetic
assays ꢀ Metalloenzymes ꢀ Organophosphate-degrading
enzymes
Introduction
Binuclear metallohydrolases form a large group of
enzymes that cover a wide range of metabolic functions
ranging from bone resorption to signal transduction and
DNA replication, to name but a few [1–3]. Several
enzymes from this family are current targets for drug
design and development, and others have found application
in biotechnology. As an example, organophosphate-
degrading phosphotriesterases have been shown to be very
efficient bioremediators owing to their ability to hydrolyze
and thus detoxify many commonly used pesticides and
some nerve agents (including sarin and VX) [3–6]. Both
drug design and the development of enzymes with tailored
catalytic properties require access to kinetic assays that
facilitate the measurement of catalytic parameters (i.e., kcat
,
Electronic supplementary material The online version of this
article (doi:10.1007/s00775-013-1079-0) contains supplementary
material, which is available to authorized users.
Km, and kcat/Km) and inhibition constants (i.e., Ki). Such
assays are frequently based on spectrophotometric mea-
surements that record the consumption of reagents and/or
the formation of products. However, often biologically
relevant reactants do not have spectroscopic properties that
facilitate simple and rapid assays, and hence researchers
frequently use synthetic (nonbiological) substrates with
specific chromophoric fingerprints to study enzymatic
reactions. An example is para-nitrophenol phosphate.
Many metallohydrolases are capable of cleaving the ester
bond of this molecule, and the yellow color of the product
para-nitrophenol makes rate measurements rapid and fac-
ile. However, for some applications such as the assessment
of substrate specificity or the development of a catalyst
M. M. Pedroso ꢀ F. Ely ꢀ T. Lonhienne (&) ꢀ
L. R. Gahan ꢀ L. W. Guddat ꢀ G. Schenk (&)
School of Chemistry and Molecular Biosciences,
University of Queensland,
St. Lucia, Brisbane, QLD 4072, Australia
e-mail: t.lonhienne@uq.edu.au
G. Schenk
e-mail: schenk@uq.edu.au
D. L. Ollis
Research School of Chemistry,
Australian National University,
Canberra, ACT 0200, Australia
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with increased reactivity towards a particular molecule, it
is desirable to use native reactants as substrates. It is thus
essential to make available an assay that facilitates rate
measurements that are independent of spectroscopic prop-
erties of reactant molecules.
pesticide-polluted environments [20]. Current research
aims to evolve mutants of these enzymes with improved
catalytic efficiency towards specific pesticides and nerve
agents [3, 5, 6, 21, 22]. Rv0805 is involved in cell signaling
pathways. It has been postulated that this enzyme, owing to
its significance in maintaining cellular cyclic adenosine
monophosphate (cAMP) levels, may be a target for the
treatment of tuberculosis [17, 23, 24].
Here, we describe an assay that uses isothermal titration
calorimetry (ITC), a method that records changes in heat
associated with bond breakage (i.e., hydrolysis), and as
such is applicable for measurements with any potential
substrate of a particular enzyme. We have chosen three
enzyme systems to illustrate the method, an organopho-
sphate-degrading phosphotriesterase from Agrobacterium
radiobacter (OpdA) [7–11], a glycerophospodiesterase
from Enterobacter aerogenes (GpdQ) [12–16], and a cyclic
nucleotide diesterase from Mycobacterium tuberculosis
(Rv0805) [17–19]. Both OpdA and GpdQ are promising
bioremediators; in particular, OpdA has already been
shown in field trials to be competent in detoxifying
Materials and methods
Materials
Escherichia coli BL21(DE3) and DH5a host cells were
purchased from Novagen. All chromatographic devices (fast
protein liquid chromatography system and relevant resins)
were from GE Healthcare, and all chemicals, buffers, and
NO2
O
S
O
S
P
O2N
O
O
O-
O
P
O
a
b
O
N+
-O
NH2
N
N
O
P
O
O
N
N
O
O
c
H
O
H
O
P
O
OH
O
O-
O
e
P
O
O
d
Fig. 1 Substrates used in this study: a demeton-S; b bis(para-nitrophenol) phosphate (bpNPP); c paraoxon; d trimethyl phosphate (TMP);
e cyclic adenosine monophosphate (cAMP)
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391
substrates (see Fig. 1), unless mentioned otherwise, were
purchased from Sigma Chemical Company.
recombinant protein expression was achieved by adding
(isopropyl b-D-1-thiogalactopyranoside, 1 mM final concen-
tration) and continuing incubation at 30 °C under constant
shaking (200 rpm) for another 24 h. The cells were harvested
by centrifugation, resuspended in lysate buffer containing
50 mM TrisHCl, pH 8.5, with 5 mM mercaptoethanol,
150 mM NaCl, and EDTA-free protease inhibitor. The cells
were lysed using a French press (1,000 psi). Since the Rv0805
constructcontainsanN-terminalhexahistidinetag,thesoluble
fraction was applied to a Ni(II) affinity column (Ni(II)-IMAC
resin), equilibrated with 50 mM TrisHCl, pH 8.5, with 5 mM
mercaptoethanol, 150 mM NaCl, and 20 mM imidazole. The
protein was eluted with an isocratic gradient using 50 mM
TrisHCl, pH 8.5, with 5 mM mercaptoethanol, 150 mM
NaCl, 200 mM imidazole, and 10 % glycerol. Rv0805 con-
centration was measured at 280 nm using e280 = 19,940
M-1 cm-1 per monomeric unit.
Protein expression and purification
The expression and purification of OpdA and GpqQ were
described in detail elsewhere and will only be briefly
summarized here [9, 14]. The OpdA gene was previously
cloned into the recombinant plasmid pETMCSI, which was
used to transform competent E. coli BL21 (DE3) cells. A
single colony was inoculated in terrific broth supplemented
with 1 mM CoSO4 and 50 lM ampicillin. The culture was
grown at 37 °C at 200 rpm for 48 h. Following centrifu-
gation (approximately 30 g wet mass) the cells were dis-
rupted using a French press at 1,000 psi. The soluble
fraction was applied to a MonoS HR column, equilibrated
with 50 mM N-(2-hydroxyethyl)piperazine-N0-ethanesul-
fonic acid (HEPES) buffer, pH 7.0, supplemented with
0.5 mM CoSO4; enzymes were eluted using a gradient of
0–1 M NaCl. The fractions containing activity were con-
centrated and loaded onto a Sephacryl S-200 column,
equilibrated with the same buffer as in the previous step,
and were eluted with buffer supplemented with 0.2 M
NaCl. The purified OpdA was analyzed by sodium dodecyl
sulfate–polyacrylamide gel electrophoresis, and its con-
centration was determined by measuring the absorption at
280 nm (e280 = 29,280 M-1 cm-1 per monomeric unit).
The GpdQ-encoding expression vector pCY76::GpdQ
was used to transform competent E. coli DH5a cells. The
cells were grown in 4 L of terrific broth, containing 50 lg/
mL ampicillin and 0.1 mM CoCl2, for 36 h at 30 °C. Then
the cells were harvested by centrifugation and lysed using a
French press at 1,000 psi. The soluble fraction was loaded
onto a HiPrep 16/10 DEAE column, equilibrated with
20 mM tris(hydroxymethyl)aminomethane hydrochloride
(TrisHCl) buffer, pH 8.0, and the proteins were eluted with a
linear NaCl gradient from 0 to 1 M. Fractions containing
activity against 2 mM bis(para-nitrophenyl) phosphate
(bpNPP) were dialyzed against 20 mM HEPES buffer, pH
8.0, containing 1.5 M (NH4)2SO4. The dialysate was then
loaded onto a HiLoad 26/10 phenyl Sepharose column,
equilibrated with 20 mM HEPES buffer, pH 8.0, containing
1 M (NH4)2SO4. Fractions containing phosphodiesterase
activity were concentrated to approximately 4 mL and loa-
ded onto a HiPrep 16/10 Sephacryl S-200 gel filtration col-
umn, equilibrated with 20 mM HEPES, pH 8.0, containing
0.15 M NaCl. GpdQ concentration was measured at 280 nm
using e280 = 39,880 M-1 cm-1 per monomeric unit.
Apoenzyme preparation and enzyme reconstitution
Most binuclear metallohydrolases display some promiscuity
with respect to the metal ions they may bind [2, 3, 17, 25–27];
hence, the metal ion composition of a purified enzyme is
often affected by the availability of metal ions in the purifi-
cation buffers. Since differences in metal ion compositions
may affect both the reactivity of a metalloenzyme and its
interaction with substrates (see, e.g., [8, 11, 12, 23]), care was
taken to prepare samples with well-defined metal content.
Apoenzymes were obtained by incubating approxi-
mately 3 mg of protein (i.e., OpdA, GpdQ, or Rv0805) in a
3-mL solution containing 5 mM EDTA, 5 mM 1,10-phe-
nanthroline, 5 mM 2,6-pyridine dicarboxylic acid, 5 mM
8-hydroxyquinone-5-sulfonic acid, 5 mM 2-mercap-
toethanol in 20 mM HEPES buffer, pH 7.0, at 4 °C. After
48 h for Rv0805 and 24 h for OpdA and GpdQ, the protein
sample was separated from the chelating solution using an
Econo-Pac 10DG gel filtration column equilibrated with
50 mM TrisHCl, pH 8.5, containing 100 mM NaCl. The
absence of metal ions in the protein solutions was con-
firmed by atomic absorption spectroscopy.
Subsequently, enzyme activities were reconstituted by
the addition of the desired metal ion and incubation for at
least 24 h at 4 °C. GpdQ and OpdA activities were
reconstituted with CoSO4, and Rv0805 activity was
reconstituted using MnCl2.
ITC data collection
The Rv0805 gene, cloned into the vector pET47B, was
purchased from GeneArt. The construct was transferred into
competent E. coli BL21(DE3) cells. The transformed cell
culture was incubated at 37 °C until an optical density of
approximately 0.5 was reached. Subsequently, induction of
ITC data were collected with an iTC200 system from
MicroCal. All data analysis was performed using the pro-
gram Origin. All measurements were conducted at 25 °C.
Catalysis was recorded by measuring the change in the
thermal power triggered by the enzymatic hydrolysis of the
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12.0
11.6
11.2
Here, i is the ith value of dQ=dt obtained after each
injection.
The experimental determination of DHapp is achieved by
allowing a known amount of substrate to react to com-
pletion in a single-injection experiment (Fig. 3; a second
injection was routinely performed to verify reproducibility
of the data and exclude product inhibition). DHapp arises
from the integration of the power over time obtained by the
total digestion of the substrate as described by Eq. 4:
dQ1
dt
dQ2
dt
t¼1
Z
1
dQðtÞ
dt
DHapp
¼
dt;
ð4Þ
V ꢀ ½Sꢁtotal
t¼0
0
150
300
where [S]total is the initial concentration of substrate in the
reaction cell.
Time (min)
Fig. 2 Illustrative multiple-injection experiment. In subsequent
injections, substrate was added to the enzyme; each injection followed
after thermal equilibrium from the previous injection had been
reached
The reaction rates can be measured with two different
methods, i.e., single-injection or multiple-injection experi-
ments. The first method consists of a continuous rate mea-
surement after a single injection of the substrate in the cell
containing enzyme or vice versa. When the substrate has been
completely digested, the thermal power returns to the prein-
jection baseline. DHapp is determined by the integration of the
area under the peak (Eq. 4). From the analyses of the decay
curve resulting from the substrate decomposition, catalytic
parameters such as kcat and Km can be obtained on the basis of
the calculation of the remaining substrate concentration at any
time in the reaction course as shown in Eq. 5:
substrate after injection. The samples were buffered using
50 mM TrisHCl, pH 8.5, containing 100 mM NaCl, to
keep the ionic strength constant throughout the experiment.
ITC data analysis
Conversion of substrate to product by enzymes generates
heat corresponding to the enthalpic component (DH) of the
reaction. Therefore, the rate of an enzymatic reaction is
directly proportional to the rate of heat generation and can
be obtained as described previously [28] and summarized
in the following paragraph.
R
t
dQðtÞ dt
t¼0
DHappdtꢀ V
St ¼ St¼0
ꢂ
:
ð5Þ
The rate can then be plotted as a function of the sub-
strate concentration, generating a continuous kinetic curve.
The second method consists of a known concentration of
enzyme solution placed in the cell, and a number of con-
secutive injections with substrate are performed. In this
method the thermal power trace does not return to the origin
(like in the single-injection model), but instead reaches a
constant value corresponding to the saturation of the enzyme
by the substrate. Integration of the thermal power over time
can be used to construct a plot of the reaction rate versus the
total substrate concentration (as shown in Fig. 2) [28]. This
method assumes that no appreciable substrate degradation
occurs during the measurements.
The rate of heat generation of an enzymatic reaction is
defined as the thermal power, expressed by Eq. 1:
dQ
Power ¼
:
ð1Þ
dt
Practically, the thermal power of a reaction is given by
subtracting the baseline (no reaction) from the calorimetric
trace associated with substrate consumption (Fig. 2).
The amount of heat involved in the conversion of
n moles of substrate to product is given by Eq. 2:
Q ¼ n ꢀ DHapp ¼ ½Pꢁtotal ꢀ V ꢀ DHapp
;
ð2Þ
where V is the volume of the reaction cell, [P] is the
concentration of the product generated by the enzymatic
reaction, and DHapp is the molar enthalpy for the reaction,
which needs to be determined experimentally (see below).
Since the reaction rate is defined as the concentration
change of the product over time d½Pꢁdt, it can be obtained
by the derivative of Eq. 2, resulting in Eq. 3:
Rate versus substrate concentration profiles were
accordingly fitted to the Michaelis–Menten equation
(Eq. 6) [29]:
Vmax ꢀ S
v ¼
;
ð6Þ
Km þ S
where Vmax represents the maximum velocity, Km is the
Michaelis–Menten constant, S is the substrate concentra-
tion, and v is the initial velocity.
d½Pꢁ
dt
1
dQi
Rate ¼
¼
ꢀ
:
ð3Þ
V ꢀ DHapp dt
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393
a
b
0
0
-1
-2
-3
-4
-5
-5
-10
-15
0
2000
4000
6000
8000
10000
0
500
1000
Time (s)
Time (s)
10
5
c
d
0.0
-0.5
-1.0
-1.5
-2.0
0
-5
-10
-15
-20
0
100
200
300
0
100
200
300
400
500
Time (s)
Time (s)
12.5
12.0
11.5
11.0
10.5
10.0
9.5
e
f
12.0
11.5
11.0
10.5
10.0
9.0
0
2000
4000
6000
8000
0
1000
2000
3000
4000
Time (min)
Time (s)
Fig. 3 Single-injection experiment for the following enzyme/sub-
strate pairs: a OpdA and paraoxon, b OpdA and demeton-S, c GpdQ
and bpNPP, d GpdQ and TMP, e Rv0805 and bpNPP, and f Rv0805
and cAMP. The reaction proceeds until all substrate has been
converted to product, i.e., until the power returned to the baseline.
The integration of this curve gives rise to the value of DHapp (see the
text for details). A second injection of substrate was also performed,
illustrating that product inhibition plays no significant role in these
reactions
Results and discussion
spectrophotometric assays) and (2) the necessity to have an
assay available for reactants that are not amenable to
simple kinetic measurements (i.e., substrates that lack
chromophoric markers). For the enzyme OpdA, the sub-
strate commonly used for functional studies is the orga-
nophosphate pesticide paraoxon (Fig. 1); on hydrolysis the
product para-nitrophenolate is released, which has a
In establishing a method for a task such as measuring
catalytic parameters, representative systems need to be
carefully selected. For the purpose of this study, our
selection was guided by (1) the availability of comparative
experimental data obtained from alternative methods (i.e.,
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Table 1 Comparison of catalytic parameters obtained by isothermal titration calorimetry (ITC) and continuous or discontinuous spectroscopic
assays
Enzyme
Substrate
DHapp (kcal mol-1
)
ITC
Spectroscopic assays
kcat (s-1
)
Km (lM)
kcat (s-1
)
Km (lM)
OpdA
Paraoxon
Demeton-S
GpdQ
-4.0
-7.9
1,541 (3)
0.7 (0.08)
149 (2)
1,426a
0.8 (0.2)b
103
227 (23)
114 (36)
bpNPP
-16.6
-0.3
1.7 (0.7)
2.5 (0.5)
155 (10)
335 (21)
1.7c
*0.3d
120
TMP
*1,000
Rv0805
bpNPP
30,50-cAMP
-18.3
-10.0
1.6 (0.3)
350 (1.7)
39 (3)
1.7 (0.07)b
8.5 (2.1)b
221 (2)
67 (6)
10.8 (1.1)
bpNPP bis(para-nitrophenol) phosphate, 30,50-cAMP 30,50-cyclic adenosine monophosphate, TMP trimethyl phosphate
a
From Ely et al. [9]
b
This study
c
From Hadler et al. [14]
d
From Gerlt and Whitman [13]
characteristic pH-independent isosbestic wavelength at
347 nm (e = 5,176 M-1 cm-1) [30]. However, since
OpdA is a very efficient and commercially available bio-
remediator (http://www.orica-landguard.com), its catalytic
efficiency may be geared towards the hydrolysis of specific
pesticides or nerve agents such as demeton-S or VX,
respectively, using in vitro evolution techniques [8, 22, 31].
These substrates lack a chromophoric marker that would
facilitate a simple and continuous spectrophotometric
assay.
of these rates versus paraoxon concentration (calculated
using Eq. 5) displays Michaelis–Menten-type saturation
behavior and can be fit using Eq. 6, resulting in kcat and Km
values of approximately 1,500 s-1 and 150 lM, respec-
tively (Table 1). The data obtained with the spectroscopic
assays are thus in good agreement with the values obtained
here by ITC.
The above results indicate that ITC is a convenient
technique to assess kinetic parameters for OpdA-catalyzed
reactions. We thus expanded our study to include more
‘‘difficult’’ substrates, i.e., substrates that lack a chromo-
phoric marker necessary for a simple spectrophotometric
detection of product formation and/or substrate consump-
tion. In the organophosphate pesticide demeton-S, as well
as in the nerve agent VX, hydrolysis occurs at a P–S bond
(Fig. 1). The resulting free thiol is not directly spectro-
photometrically detectable, and an assay would require a
coupled reaction, whereby, for instance, the thiol moiety is
reacted with 5,5‘-dithiobis(2-nitrobenzoic acid) [7]. With
use of ITC, the activity for demeton-S can be easily and
rapidly determined in the same manner as described above
for paraoxon. Since demeton-S is a rather slow substrate,
the OpdA concentration in the cell was increased to a final
concentration of 260 lM and the multiple-injection
method was used (in an initial single-injection experiment
DHapp was estimated with a 2 lM OpdA solution and
[S]final = 3.8 mM). In total, 38 consecutive injections of
1 lL of a 5 mM demeton-S solution were performed
(Fig. 5) to determine the catalytic parameters. The result-
ing values for kcat and Km are approximately 0.7 s-1 and
227 lM, respectively (Table 1). For comparison, the cat-
alytic rates obtained by ITC and the coupled assay are
To demonstrate the aptitude of ITC as a technique to
measure catalytic parameters, paraoxon and demeton-S
were used as OpdA substrates. The published catalytic
parameters determined for paraoxon with the spectropho-
tometric assay are listed in Table 1. Because of solubility
problems, paraoxon was placed into the cell with a final
concentration of 3 mM and in a single injection OpdA was
added. The final enzyme concentration was 1.64 nM. The
reaction was monitored for approximately 40 min (2,500 s)
until all substrate had been converted to product (Fig. 4).
The complete conversion (Fig. 4a) is necessary in order to
obtain the experimental value of DHapp (according to
Eq. 2). As controls to account for nonenzymatic drifts in
the baseline and thermal dilution effects, respectively, the
same experiment was repeated twice, once in the absence
of substrate and once without enzyme (see Figs. S1–S5, for
the latter controls; no measurable dilution effects were
recorded for the former control). The heat associated with
these two controls was subtracted from the overall reaction
heat. The resulting DHapp was thus estimated to be
-4.007 kcal mol-1. With use of Eq. 3, the experimental
data can be converted to catalytic rates (Fig. 4b). The plot
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395
a
a
0.1
0.0
0
-1
-2
-0.1
-0.2
0
1000
2000
Time (s)
0.8
b
0
1000
2000
Time(sec)
0.6
0.4
b
1500
1000
500
0
0.2
0.0
0.0
0.2
0.4
0.6
0.8
Demeton-S (mM)
0
Fig. 5 Hydrolysis of demeton-S by OpdA. a Experimental data
obtained from a multiple-injection experiment ([E]final = 260 lM;
[S]stock = 5 mM; injection volume 1 lL; 38 injections). b The
dependences of the catalytic rates on the substrate concentration
measured by ITC (circles) and a discontinuous spectroscopic assay
(triangles) were compared. The data sets were fit using Eq. 6
1
2
3
Paraoxon (mM)
Fig. 4 Hydrolysis of paraoxon by OpdA. a Experimental data
obtained from single-injection experiment ([E]tot = 1.64 nM
(V = 200 lL), [S]tot = 3 mM). b The dependences of the catalytic
rates on the substrate concentration measured by isothermal titration
calorimetry (ITC) (circles) and a continuous spectroscopic assay
(triangles) [9] were compared. The data sets were fit using Eq. 6
a
paraoxon, its hydrolysis can be easily monitored spectro-
photometrically because of the para-nitrophenolate leaving
group. TMP is a slow substrate that is not amenable to a
simple continuous spectrophotometric assay. GpdQ is not
prone to product inhibition (two consecutive single injec-
tions with each of the two substrates were performed and
both isotherms had virtually the same shape; Fig. 3c, d).
Using a single injection, we determined DHapp as described
above ([S]final = 4.3 and 3.6 mM for bpNPP and TMP,
respectively). Catalytic parameters for the hydrolysis of the
two substrates were determined by the multiple-injection
method (Figs. 6, 7). For the reaction with bpNPP, 30
injections of substrate (10 mM stock concentration) were
added to GpdQ (2.7 lM at the beginning of the experi-
ment; Fig. 6). The values obtained for kcat and Km,
approximately 1.7 s-1 and approximately 120 lM,
respectively (Table 1), are in excellent agreement with the
values obtained by measuring the release of the para-ni-
trophenolate product [14]. A similar approach (50 injec-
tions) was used to determine the kinetic parameters for the
reaction of GpdQ with TMP (Fig. 7). The values for kcat
and Km are approximately 2.5 s-1 and approximately
335 lM, respectively. Although the agreement with pre-
viously reported values (using a discontinuous assay) is not
shown in Fig. 5b. Although the agreement between the two
data sets is not as good as that observed for the reaction
with paraoxon (Fig. 4b), the discrepancy is acceptable
(approximately 10 % for the rates, 50 % for the Km values;
Table 1). The estimated standard deviations reported for
the ITC-based assays were generally smaller than those
obtained from the alternative assays. In the case of the
measurements with demeton-S, the two methods thus
provide parameters that are, within errors, comparable.
As mentioned above, OpdA is a member of a large
family of phosphorolytic binuclear enzymes. The enzyme
GpdQ is another promising bioremediator, but it is also one
of the most promiscuous phosphatases known since it has
been shown to act on monoester, diester, and triester sub-
strates [7, 14, 15]. Among its many substrates, GpdQ is
able to degrade (1) TMP and methylphosphonate, which
are extremely stable aliphatic phosphoesters [13, 32], and
(2) the molecule EA 2192, the toxic by-product of VX
hydrolysis [33]. Here, we measured GpdQ activity by ITC
using the substrates bpNPP and TMP. The former is the
standard substrate used for functional studies; like
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1
0
0.0
-0.2
-0.4
-0.6
-0.8
-1.0
-1.2
-1.4
a
a
-1
-2
-3
-4
-5
-6
-7
-8
0
500
1000
1500
2000
0
500
1000
1500
Time (s)
Time (sec)
2.0
2.5
2.0
1.5
1.0
0.5
0.0
b
b
1.5
1.0
0.5
0.0
0.0
0.5
1.0
bpNPP (mM)
0
5
TMP (mM)
10
Fig. 6 Hydrolysis of bpNPP by GpdQ. a Experimental data obtained
from a multiple-injection experiment ([E]final = 2.7 lM; [S]stock
10 mM; 30 injections of 1 lL). b The dependences of the catalytic
rates on the substrate concentration measured by ITC (circles) and a
continuous spectroscopic assay (triangles) [14] were compared. The
data sets were fit using Eq. 6
=
Fig. 7 Hydrolysis of TMP by GpdQ. a Experimental data obtained
from a multiple-injection experiment ([E]final = 100 lM; [S]stock
100 mM; 50 injections of 0.5 lL). b The dependence of the catalytic
rates on the substrate concentration was determined by ITC and the
data set was fit using Eq. (6). Only estimates of the turnover number
and the Michaelis constant were previously reported [13]
=
as good as the that observed with the substrate bpNPP
(Table 1), the slowness of the reaction and the likely het-
erogeneity in the metal ion composition in the previous
study [13] may be major contributors to the observed
discrepancy.
experiment; Fig. 8). The catalytic parameters obtained are
again in excellent agreement with those measured with a
continuous spectrophotometric assay (Table 1).
The same approach was used to monitor the reaction of
Rv0805 with cAMP (Figs. 3f, 9; 16 injections, 1.0 lM
Rv0805 at the beginning of the experiment). The estimated
catalytic parameters are listed in Table 1. To compare
these values with those obtained from an alternative
method, we applied a discontinuous spectroscopic method
as described by Carter and Karl [35]. In brief, cAMP
(ranging from 10 lM to 1 mM) was mixed with enzyme
(0.21 lM final concentration). The reaction (Vtot = 30 mL)
was allowed to run for 25 min. Every 60 s, 1-mL aliquots
were removed and mixed with 100 lL of 20 mM EDTA to
quench the reaction. The mixture was then diluted to
ensure that the concentration of the reaction product,
assuming complete turnover, did not exceed 80 lM.
Rv0805 was included in this study because it is the
closest known sequence analogue of GpdQ, sharing about
20 % sequence identity [34], but has a largely different
substrate specificity. Rv0805 is involved in bacterial signal
transduction by regulating the intracellular concentration of
cyclic nucleotides [17–19]. The biologically relevant sub-
strates for Rv0805 such as cAMP and cyclic diguanylate
are again not amenable to a simple continuous spectro-
photometric assay (a discontinuous assay to measure the
rate of phosphate formation is used instead) [17, 19].
As for GpdQ, the ITC method was initially tested using
bpNPP as a substrate (Figs. 3e, 8). Catalytic parameters
were determined with the multiple-injection approach (45
injections, 4.0 lM Rv0805 at the beginning of the
123

J Biol Inorg Chem (2014) 19:389–398
397
a
a
0.0
-0.1
-0.2
-0.3
-0.4
-0.5
-0.6
-0.7
0
-2
-4
-6
-8
0
200
400
600
800
1000
1200
0
500
1000
1500
2000
2500
Time (s)
Time (s)
b
b
10
5
1.5
1.0
0.5
0.0
0
0.0
0.5
1.0
1.5
2.0
0.0
0.2
0.4
0.6
0.8
bpNPP (mM)
cAMP (mM)
Fig. 8 Hydrolysis of bpNPP by Rv0805. a Experimental data
obtained from a multiple-injection experiment ([E]final = 20 lM;
[S]stock = 14 mM; 45 injections of 0.7 lL). b The dependences of the
catalytic rates on the substrate concentration measured by ITC
(circles) and a continuous spectroscopic assay (triangles) were
compared. The data sets were fit using Eq. 6
Fig. 9 Hydrolysis of cAMP by Rv0805.
a
Experimental data
obtained from a multiple-injection experiment ([E]final = 1 lM;
[S]stock = 10 mM; 16 injections of 1 lL). b The dependences of
the catalytic rates on the substrate concentration measured by ITC
(circles) and a discontinuous spectroscopic assay (triangles) were
compared. The data sets were fit using Eq. 6
Subsequently, malachite green was added to the aliquot and
after 15 min the absorption was measured at 620 nm. The
catalytic parameters obtained (Table 1) are in good overall
agreement with those from the ITC experiment, but the
latter was significantly more rapid.
precise biological role(s) is(are) often poorly understood,
obscured by a lack of knowledge about the identity of
their preferred substrates. Furthermore, the range of sub-
strates may frequently be very broad (‘‘promiscuous’’ use
of substrates). A uniform approach to assess/compare a
variety of substrates is thus desirable to probe both the
reaction mechanism of hydrolysis and the biological
function(s) of these enzymes. Here, ITC was demon-
strated to be very powerful, reliable, and rapid to assess
the catalytic properties of several binuclear metallohy-
drolases towards various substrates. It is anticipated that
this approach can be widely applied by bioinorganic
chemists/enzymologists, thus contributing significantly to
our understanding of the function and mechanism of such
systems, but it may also be useful in the development of
(1) variants of such enzymes with altered catalytic prop-
erties (e.g., different substrate preferences) or (2) specific
inhibitors as future drug leads.
In summary, ITC has been shown to be a simple,
efficient, and reliable method to measure enzymatic
activities for a representative group of binuclear metal-
lohydrolases. The catalytic parameters obtained are lar-
gely in good agreement with comparable values obtained
from spectrophotometric assays. ITC is not envisioned to
replace conventional spectrophotometric assays for func-
tional studies that can use substrates with convenient
chromophoric markers. Instead, ITC may provide a con-
duit to assess the interactions between enzymes and their
natural substrates, in particular if the latter are not ame-
nable to rapid, simple, and continuous spectrophotometric
assays. Binuclear metallohydrolases are a large family of
enzymes with a growing number of members. Their
123

398
J Biol Inorg Chem (2014) 19:389–398
´
15. Hadler KS, Mitic N, Ely F, Hanson GR, Gahan LR, Larrabee JA,
Ollis DL, Schenk G (2009) J Am Chem Soc 131:11900–11908
Acknowledgments This work was financially supported by the
Australian Research Council, Discovery Projects Scheme
(DP120104263). G.S. also acknowledges the receipt of an ARC
Future Fellowship (FT120100694). M.P. is supported by an Interna-
tional Postgraduate Research Scholarship and University of Queens-
land International Living Allowance Scholarship.
´
16. Hadler KS, Mitic N, Yip SH-C, Gahan LR, Ollis DL, Schenk G,
Larrabee JA (2010) Inorg Chem 49:2727–2734
17. Shenoy AR, Capuder M, Draskovic P, Lamba D, Visweswariah
SS, Podobnik M (2007) J Mol Biol 365:211–225
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20. Sutherland TD, Horne I, Weir KM, Coppin CW, Williams MR,
Selleck M, Russell RJ, Oakeshott JG (2004) Clin Exp Pharmacol
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