Stabilization of anionic and neutral forms of a fluorophoric ligand at the active site of human carbonic anhydrase I

Article abstract of DOI:10.1016/j.bbapap.2010.06.024

We synthesized a fluorogenic dansylamide derivative (JB2-48), which fills the entire (15? deep) active site pocket of human carbonic anhydrase I, and investigated the contributions of sulfonamide and hydrophobic regions of the ligand structure on the spectral, kinetic, and thermodynamic properties of the enzyme-ligand complex. The steady-state and fluorescence lifetime data revealed that the deprotonation of the sulfonamide moiety of the enzyme bound ligand increases the fluorescence emission intensity as well as the lifetime of the fluorophores. This is manifested via the electrostatic interaction between the active site resident Zn²⁺ cofactor and the negatively charged sulfonamide group of the ligand, and such interaction contributes to about 2.2kcal/mol (ΔΔG°) and 0.89kcal/mol (ΔΔG^?) energy in stabilizing the ground and the putative transition states, respectively. We provide evidence that the anionic and neutral forms of JB2-48 are stabilized by the complementary microscopic/conformational states of the enzyme. The implication of the mechanistic studies presented herein in rationale design of carbonic anhydrase inhibitors is discussed.

Full text of DOI:10.1016/j.bbapap.2010.06.024

Biochimica et Biophysica Acta 1804 (2010) 1965–1973
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Stabilization of anionic and neutral forms of a fluorophoric ligand at the active site of
human carbonic anhydrase I
Sumathra Manokaran a, Jayati Banerjee b, Sanku Mallik b, D.K. Srivastava a,
⁎
a
Department of Chemistry and Biochemistry, North Dakota State University, Fargo, ND 58105, USA
b
Department of Pharmaceutical Sciences, North Dakota State University, Fargo, ND 58105, USA
a r t i c l e i n f o
a b s t r a c t
Article history:
We synthesized a fluorogenic dansylamide derivative (JB2-48), which fills the entire (15 Å deep) active site
pocket of human carbonic anhydrase I, and investigated the contributions of sulfonamide and hydrophobic
regions of the ligand structure on the spectral, kinetic, and thermodynamic properties of the enzyme–ligand
complex. The steady-state and fluorescence lifetime data revealed that the deprotonation of the sulfonamide
moiety of the enzyme bound ligand increases the fluorescence emission intensity as well as the lifetime of
the fluorophores. This is manifested via the electrostatic interaction between the active site resident Zn2+
Received 9 December 2009
Received in revised form 23 June 2010
Accepted 25 June 2010
Available online 8 July 2010
Keywords:
cofactor and the negatively charged sulfonamide group of the ligand, and such interaction contributes to
Human carbonic anhydrase-I
Inhibitor design
Sulfonamide
Fluorophore
Dansylamide
‡
∘
about 2.2 kcal/mol (ΔΔG ) and 0.89 kcal/mol (ΔΔG ) energy in stabilizing the ground and the putative
transition states, respectively. We provide evidence that the anionic and neutral forms of JB2-48 are
stabilized by the complementary microscopic/conformational states of the enzyme. The implication of the
mechanistic studies presented herein in rationale design of carbonic anhydrase inhibitors is discussed.
© 2010 Elsevier B.V. All rights reserved.
Ligand binding
1. Introduction
sentative potent inhibitors of these enzymes [10,13–15]. The X-ray
crystallographic structures of several CA-isozymes in the presence of
Although enzymes are evolved to catalyze specific biological
reactions, their active site pockets appear to lack absolute discrimina-
tory features between their cognate physiological substrates/products
and a wide range of structurally diverse compounds (which exhibit no
structural resemblance to putative substrates, products, or intermedi-
ates of enzyme catalyzed reactions) [1–3]. The ability of enzymes to
accommodate structurally diverse ligands (due to the marked dynamic
flexibility) within their active site pockets have led to designing a
variety of therapeutic agents against pathogenic enzymes [4–6]. Given
this, it is conceivable that the X-ray crystallographic structure of an
enzyme bound inhibitor may not serve as an ideal template for
predictably designing highly potent enzyme inhibitors as potential
drugs [7–9].
Of 15 different isozymes of carbonic anhydrases (CAs; EC 4.2.1.1),
which are ubiquitously distributed zinc metalloproteins, and are
involved in a variety of physiological functions, carbonic anhydrase I
(hCA I) is representative of one of the cytosolic enzymes, which is
present in all animal kingdom [10–17]. Although different isozymes of
carbonic anhydrases have been found to be inhibited by a variety of
differently functionalized ligands, aryl sulfonamides serve as repre-
different aryl sulfonamide inhibitors have been solved to atomic
resolutions [17–23]. Both crystallographic as well as NMR data reveal
that the sulfonamide moieties of the ligands interact with active site
resident Zn2+ cofactor as well as with Thr-199 [13,14,22,24–29]. The
structural data further suggest that the active site Zn2+ electrostat-
ically interacts with the negatively charged nitrogen (produced via
deprotonation of the amide group) of sulfonamide moiety of the
ligand [25–28].
Since the aliphatic sulfonamides do not serve as potent inhibitors
of carbonic anhydrases [13,30], it follows that the wide active site
pocket of the enzyme needs to be occupied by bulky (viz., aromatic)
groups of the ligand structures. Based on the structure-activity
relationship, it has been apparent that both sulfonamide and
aromatic moieties of ligands contribute to the overall binding energy
of carbonic anhydrase–ligand complexes [31–33]. However, to the
best our knowledge, no systematic studies have been conducted to
assess the contributions of hydrophobic and electrostatic interac-
tions of ligand structures in stabilizing the corresponding enzyme–
ligand complexes. This issue is specifically addressed in this
manuscript.
We recently investigated the detailed microscopic pathways for
the binding of dansylamide to hCA I versus hCA II, and provided
evidence that despite a marked structural similarity, the active site
pockets of these enzymes behave differently in accommodating the
common ligand [34]. However, one of the deficiencies of dansylamide
fluorophore has been the comparable quantum yields of its aqueous
Abbreviations: CA, Carbonic anhydrase; CA-I, human carbonic anhydrase I; hCA-II,
human carbonic anhydrase II; EDTA, Ethylenediaminetetraacetic acid; DMSO,
Dimethysulfoxide
⁎
Corresponding author. Tel.: +1 701 231 7831; fax: +1 701 231 7884.
E-mail address: dk.srivastava@ndsu.edu (D.K. Srivastava).
1570-9639/$ – see front matter © 2010 Elsevier B.V. All rights reserved.
doi:10.1016/j.bbapap.2010.06.024

1966
S. Manokaran et al. / Biochimica et Biophysica Acta 1804 (2010) 1965–1973
versus the enzyme bound forms, and thus it did not serve as an ideal
ligand for pursuing detailed mechanistic studies such as reported
herein [35]. Although a few other fluorescent probes for carbonic
anhydrases have been reported in the literature, they have not been
utilized toward the above goal presumably due to their challenging
synthetic protocols [36,37]. Given these, we synthesized a series of
conjugated fluorophores using dansylamide as the parent compound
[38], of which JB2-48 was deemed to be an ideal fluorophores for
delineating the contributions of sulfonamide and hydrophobic regions
in binding with hCA I using the fluorescence signals of the enzyme–
ligand complex.
are total enzyme (hCA I), total ligand (JB2-48), stoichiometry of the
enzyme–ligand complex and the dissociation constant of the enzyme–
ligand complex, respectively.
2.2.2. Fluorescence Lifetime measurements
Fluorescence Lifetime measurements were performed on a custom
designed Photon Technology International (PTI) Fluorescence-Life-
time Instrument. The excitation sources for measuring the time
resolved fluorescence decay were the Light Emitting diodes (LEDs)
with maximum power outputs at 280 nm and 340 nm, respectively.
Whereas the 280 nm LED was utilized to excite the intrinsic
fluorescence of the protein (primarily contributed by the tryptophan
residues), the 340 nm LED was utilized to excite JB2-48 fluorescent
probe. The emitted light was detected (at right angle of the excitation
source) by means of a stroboscopic emission monochromator
configured at an appropriate wavelength. The time resolved fluores-
cence decay curves were analyzed to obtain the lifetimes of the
fluorophores under different conditions by the aid of the PTI's
software, Felix 32.
2. Experimental procedures
2.1. Materials
Zinc sulfate, ampicillin , chloramphenicol and IPTG were purchased
from Life Science Resources, Milwaukee, WI ; yeast extracts, and
tryptone were purchased from Becton Dickinson, Sparks, MD; Aceto-
nitrile was from Aldrich Chemicals, Milwaukee,WI; HEPES, p-amino-
methylbenzenesulfonamide-agarose, p-nitrophenyl acetate and PMSF
were obtained from Sigma; All the chemicals needed for synthesis of
JB2-48 were purchased from Aldrich Chemical Company, Milwaukee,
WI. The E. coli expression system BL21codon plus DE3(RIL) was from
Stratagene, La Jolla, CA. All other chemicals were of reagent grade, and
were used without further purification.
2.2.3. Steady state kinetics and spectrophotometric Studies
All spectrophotometric studies were performed on the Molecular
Devices SpectraMax Plus microplate reader, equipped with a cuvette
holder and magnetic stirrer. The enzyme activity of hCA I was
determined by monitoring the hydrolysis of p-nitrophenyl acetate
substrate at 348 nm in 25 mM HEPES buffer, pH 7.0, containing 10%
DMSO. The Ki for JB2-48 was determined as described by Banerjee
et al. [42].
2.2. Methods
The pKa values of free JB2-48 were determined by monitoring the
increase in absorption of the compound at 336 nm as a function of the
increasing pH value.
The compound JB2-48 was synthesized as described by Banerjee et al.
[38]. The recombinant form of human carbonic anhydrase I (hCA I) was
cloned, expressed and purified as per the previously developed protocol
in our lab [34]. All experiments were performed in 25 mM HEPES buffer,
pH 7.0, containing 10% DMSO (the standard buffer). Following buffers,
containing 10% DMSO, were used for the pH-dependent experiments.
pH 5.0: 25 mM acetate buffer; pH 6.0–6.5: 25 mM MES buffer; pH 7.0–
7.5: 25 mM HEPES buffer, pH 8.0–9.0: 25 mM Tris-phosphate buffer.
2.2.4. Transient kinetic experiments
Transient kinetic experiments were performed on an Applied
Photophysics SX-18 MV stopped-flow system, equipped with both
absorption and fluorescence detecting photomultiplier tubes. The
dead time of stopped flow was 1.3 ms. For fluorescence measure-
ment, the light path was configured such that the fluorescence
photomultiplier detected the emitting light via the 2-mm path
length. The excitation wavelength was maintained at 336 nm, and a
395-nm cut-off filter was installed at the exit port of the cuvette to
measure the time dependent changes in the fluorescence intensity by
a photomultiplier tube. The stopped flow traces were analyzed by the
data analysis package provided by Applied Photophysics.
2.2.1. Steady-state spectrofluorometric studies
All steady-state spectrofluorometric studies involving dansylamide
derivative, JB2-48, were performed on a Perkin Elmer lambda 50-B
spectrofluorometer, equipped with a magnetic stirrer and thermostated
water bath. The stock solution of JB2-48 was prepared in 100% DMSO
and was diluted in 25 mM HEPES buffer, pH 7.0, containing 10% DMSO
during the course of the titration as well as kinetic experiments. The
emission spectra of JB2-48 in the absence and presence of hCA I was
acquired by fixing the excitation wavelength at 336 nm. The contribu-
tions of Raman/Rayleigh scattering from the emission spectra of
fluorophores were eliminated by subtracting the spectra of the buffer
(obtained under identical experimental conditions). The dissociation
constant of the hCA I–JB2-48 complex was determined by titrating a
fixed concentration of the enzyme (2 μM) with increasing concentra-
tions of the ligand in the standard HEPES buffer containing 10 % DMSO,
pH 7.0. The excitation and emission slits were 9 mm each. The decrease
in fluorescence emission intensity of the protein and the increase in the
fluorescence emission intensity of probe were monitored at 330 nm and
470 nm, respectively (λex=280 nm). The dissociation constant of the
hCA I–JB2-48 complex was determined by analyzing the binding
isotherm by Eq. (1) as elaborated by Qin and Srivastava [39].
2.2.5. pH Jump Relaxation Studies
These experiments were performed by mixing a solution of the
hCA I–JB2-48 complex, maintained at one pH, with a concentrated
buffer at the other pH in the stopped-flow syringes, and recording the
fluorescence change as described above. For a low to high pH jump,
syringe A contained 2 μM hCA I+20 μM JB2-48 in 5 mM Acetate
buffer at pH 5.0 containing 10% DMSO and syringe B contained
200 mM Tris at pH 9.0 with 10% DMSO. Upon mixing, the pH of the
solution changed to be pH 9.0. For a high to low pH jump experiment,
syringe A contained 2 μM hCA I+20 μM JB2-48 in 5 mM Tris at pH 9.0
having 10% DMSO and syringe B contained 200 mM Acetate buffer at
pH 5 with 10% DMSO. Upon mixing, the pH of the mixture changed to
5.0.
2.2.6. Molecular modeling studies
qffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi
8
<
9
=
2
The molecular modeling studies were performed on a Silicon
Graphics-O2 molecular modeling workstation with the aid of Accelrys
software, InsightII(98). The coordinates for the X-ray crystallographic
structures of bovine CA II complexed with dansylamide (1okl.pdb)
and hCA I complexed with acetazolamide (1azm.pdb) were down-
loaded from Research Collaboratory for Structural Bioinformatics
ðE n + L + K Þ− ðE n + L + K Þ −4E n L
:
*
t* *
t
*
2
t
t
d
t
t
d
ΔF = ΔFmax
ð1Þ
*
;
Where ΔF and ΔFmax are observed and maximum fluorescence
changes for the binding of JB2-48 to hCA I, respectively. Et, Lt, n, and Kd

S. Manokaran et al. / Biochimica et Biophysica Acta 1804 (2010) 1965–1973
1967
(RCSB) Protein Data Bank. The backbones of these proteins were
superimposed, and the dansylamide was computationally transferred
from bovine CA II to hCA I. The structure of JB2-48 was built on the
backbone of the enzyme bound dansylamide with the aid of the
software Builder under the InsightII(98) platform.
3. Results
We synthesized JB2-48 as a fluorescent ligand for hCA I by
conjugating p-hydroxybenzaldehyde with p-aminonaphthalein sul-
fonamide [38] as outlined in the scheme of Fig. 1. The molecular
modeling data revealed that unlike its parent compound, dansylamide
(which occupied only 1/3 of the enzyme's active site pocket), JB2-48
filled nearly the entire (~15 Å deep) hydrophobic pocket of hCA I
(Fig. 1). Hence, JB2-48 could be envisaged to experience “full”
hydrophobicity of the enzyme's active site environment. We observed
that both free as well as the enzyme bound form of JB2-48 was fairly
stable at the room temperature, and showed no evidence of
degradation/hydrolysis during the course of any of the experiments
reported herein. We further observed that 10% DMSO had no
influence on the catalytic activity of the enzyme, and the enzyme
was stable at different pH values during the course of all experiments
reported herein (data not shown).
In a preliminary manner, we previously noted that the excitation
and emission maxima of free JB2-48 were 336 nm and 528 nm,
respectively, and that the emission maximum was blue shifted to
470 nm (with concomitant increase in the fluorescence intensity)
upon binding to hCA I [38]. This is in contrast to the emission
maximum of the enzyme-bound dansylamide as being equal to
458 nm [34]. In addition, the quantum yield of the enzyme bound JB2-
48 (as compared to the free ligand) is increased by about 40 fold [38],
and this feature allowed us to easily differentiate between the free
versus enzyme bound form of the ligand via spectrofluorometric and
transient kinetic approaches (see below).
We performed fluorescence spectroscopic studies of the above
species as a function of pH of the buffer medium (Fig. 2A). The inset of
Fig. 2A shows the representative fluorescence emission spectra
(λex =336 nm) of the enzyme bound JB2-48 at pH 5.0 and 9.0. We
confirmed that at these pH extrema, the catalytic activity of the
enzyme remains unaffected during the time regime of all spectral,
Fig. 2. pH-dependent changes in the fluorescence emission profile of JB2-48 in the
presence (panel A) and absence (Panel B) of hCA I. The main figure of Panel A shows
the increase in fluorescence emission intensity of the enzyme bound JB2-48
(λex =336 nm, λem =470 nm) as a function of pH. [hCA I]=10 μM, [JB2-48]=3 μM.
The solid line is the best fit of the data according to the Henderson–Hasselbalch
equation for the pKa value of 6.6 0.1. The inset shows the representative emission
spectra (λex =336 nm) of the enzyme bound JB2-48 complex at pH 5.0 and 9.0.
Panel B shows the increase in absorption (at 336 nm) of aqueous solution of JB2-48
solution as a function of pH. The solid smooth line is the best fit of the data according
to the Henderson–Hasselbalch equation for two pKa values of 6.5 and 10.2,
respectively.
Fig. 1. Synthetic scheme of JB2-48 and the molecular model for its occupancy at the active site pocket of hCA I.

1968
S. Manokaran et al. / Biochimica et Biophysica Acta 1804 (2010) 1965–1973
kinetic, and thermodynamic studies reported in this as well as in
subsequent sections. The data of Fig. 2A reveal that the fluorescence
emission intensity of the enzyme bound JB2-48 at pH 5.0 is
significantly lower than that observed at pH 9.0. The main panel of
Fig. 2A shows the pH dependent changes in the fluorescence emission
intensity of the enzyme–JB2-48 complex. The solid smooth line is the
best fit of the data according to the Henderson–Hasselbalch equation
for the pKa value of 6.6. The latter value is similar to the pKa derived
from the pH dependent changes in the binding affinity of dansylamide
with hCA I (pKa =6.3) and hCA II (pKa =6.4) [34].
Since JB2-48 contains two ionizable groups (viz., the sulfonamide
group of the naphthalene ring and the phenolic hydroxyl group), we
proceeded to determine their pKa values using free fluorophore. In
this endeavor, we noted that the absorption maximum of JB2-48
increases at 336 nm as a function of the pH of the solution. Using the
above signal, we titrated a fixed concentration of JB2-48 (3 μM) with
increasing aliquots of dilute NaOH and recorded the resultant pH
value (Fig. 2B). The data were analyzed according to the Henderson–
Hasselbalch equation (solid smooth line) for two pKa values of 6.5 and
10.2, respectively. Given that the pKa value of sulfonamide group in
“aqueous” benzenesulfonamide is 9.79 [35], we ascribe our experi-
mentally determined pKa's of 6.5 and 10.2 (Fig. 2B) to the phenolic
hydroxyl group and the sulfonamide moiety of JB2-48, respectively.
On comparison of the titration results of Fig. 2A and B, it appears
evident that the coordination of sulfonamide group of JB2-48 with the
active site resident Zn2+ of hCA I (forming a Lewis acid-base pair)
decreases its pKa value by about 3 units. This is in accord with the wide
spread view that at neutral pH, the Zn2+ cofactor electrostatically
interacts with the negatively charged nitrogen of aryl sulfonamide
ligands in different isoforms of carbonic anhydrases [24–28].
Although the above coordination is also likely to decrease the pKa
value of the phenolic hydroxyl group of JB2-48 (due to an extended
conjugation in the entire molecule), we could not reliably ascertain its
magnitude. However, such a decrease is unlikely to be significant
since the enzyme bound form of JB2-48 yields only one pKa of 6.6.
Hence, the pKa of phenolic hydroxyl group of JB2-48 (in the enzyme
bound form) remains unresolved from that given by the protonation/
deprotonation of the sulfonamide moiety. It should be reiterated that
the sulfonamide moiety of dansylamide fluorophore (which is devoid
of the phenolic hydroxyl group) also shows the pKa of 6.3 when bound
to hCA I [34]. Irrespectively, a comparative account of the pKa values of
JB2-48 in free and the enzyme bound forms clearly attests to the fact
that the sulfonamide as well as phenolic hydroxyl groups of the
enzyme bound fluorophore remain fully protonated and deproto-
nated (ionized) at pH values 5.0 and 9.0, respectively, and at these pH
extrema, the enzyme bound fluorophore exhibits minimum and
maximum fluorescence intensities, respectively.
In view of the above information, we could probe the contributions
of hydrophobic versus electrostatic interactions in the ground and
putative transition states upon binding of JB2-48 to hCA I, respectively
(see below). Fig. 3 shows the emission spectra of hCA I (λex=280 nm)
in the presence of increasing concentrations of JB2-48 at pH 7.0. We
ascribe the intrinsic fluorescence of hCA I to the enzyme resident
tryptophan residues. As shown in Fig. 3A, as the concentration of JB2-48
increases, whereas the fluorescence emission intensity of
the tryptophan residues (λem=330 nm) decreases (presumably due
to the static quenching), the emission intensity of the probe
(λem=470 nm) increases. Given the changes in fluorescence signals
at the above wavelengths, we titrated a fixed concentration of hCA I by
increasing concentrations of the fluorophore to determine the binding
constant of the enzyme–fluorophore complex. Fig. 3B shows the
titration profiles for the binding of JB2-48 with hCA I, obtained by
monitoring the intrinsic fluorescence of the enzyme (decrease in the
fluorescence intensity at 330 nm; open circles) as well as the probe's
signal (increase in the fluorescence at 470 nm; open triangle). The solid
smooth lines are the best fit of the data for the dissociation constants of
Fig. 3. Fluorescence spectral changes and binding isotherms for the interaction of
JB2-48 with hCA I. Panel A shows the fluorescence emission spectra of hCA-I in the
absence and presence of increasing concentrations of JB2-48. [hCA I]=2 μM. The
concentration of [JB2-48] for the emission spectra (low to high fluorescence
emission intensity at 470 nm region) has been 0, 0.1, 0.2, 0.4, 0.78, 1.9, 2.7, 4.2, 4.9,
6.3, 7.8, 9.9, 12.6, 15.3, 17.9, 19.3 μM, respectively; λex =280 nm. Note that the
titration of hCA I by JB2-48 results in a decrease in the fluorescence intensity at
330 nm, and increase in the fluorescence intensity at 470 nm. Panel B shows the
binding isotherms for the interaction of JB2-48 with hCA I determined at 330 nm
(ο; decreasing phase) and at 470 nm (Δ; increasing phase). λex =280 nm; hCA
I=2 μM. The solid smooth lines are the best fit of the data for the Kd values of 2.1
0.2 (330 nm plot) and 2.3 0.4 μM (470 nm plot), respectively.
the enzyme–probe complex as being equal to 2.1 0.2 and 2.3 0.4 μM
at 325 and 470 nm, respectively, at pH 7.0. We further determined the Ki
value of JB2-48 using p-nitriphenyl acetate as the enzyme substrate as
described by Banerjee et al. [42] and found its magnitude to be 2.7
0.5 μM. Note that all these values are comparable, and thus they
appear to originate from a common physical/binding step. We
performed similar experiment to determine the binding affinity of
JB2-48 for hCA I at pH 5.0 and 9.0, respectively. In these experiments, the
concentration of JB2-48 was increased until the enzyme appeared to be
nearly saturated. These values are summarized in Table 1. Using the
standard state as being equal to 1.0 M and pH=7.0, we translated the Kd
values of Table 1 to the standard free energy changes (ΔGº) for the
Table 1
Binding of JB2-48 to hCA I as a function of pHa.
pH
Kd
ΔGºb
5.0
7.0
9.0
6.0 μM
2.2 μM
0.14 μM
−7.1 kcal/mol
−7.7 kcal/mol
−9.3 kcal/mol
a
The concentrations of hCA I and JB2-48 at different pH values were as follows: pH 5.0:
[hCA I]=3.0 μM, [JB2-48]=0–35.8 μM pH 7.0: [hCA I]=2 μM, [JB2-48]=0–19.3 μM
pH 9.0: [hCA I]=3 μM, [JB2-48]=0–7.2 μM.
b
∘
The standard state of pH was taken as 7.0 in computing the ΔG values.

S. Manokaran et al. / Biochimica et Biophysica Acta 1804 (2010) 1965–1973
1969
enzyme–ligand interactions at 25 °C (ΔGº=−RT ln (1/Kd)), and these
derived values are presented as the last column of Table 1. A casual
perusal of the ΔGº values of Table 1 reveals that the difference in the
enzyme–JB2-48 binding energy (ΔΔGº) between the high and low pH
values is 2.2 kcal/mol. Since at the above high and low pH values, the
sulfonamide nitrogen of JB2-48 is expected to exist in fully ionized and
neutral forms, respectively, the calculated ΔΔGº value serves to be the
measure of the electrostatic energetic contribution between the active
site resident Zn2+ cofactor and the negatively charged sulfonamide
moiety of the fluorophore.
of 12.5 ns), the hCA I bound form of the fluorophore at pH 7.0 yields
two lifetimes of 13.5 and 27.3 ns, respectively. Clearly, JB2-48 does not
experience identical microscopic environment in 90% dioxane vis a vis
that present at the active site pocket of hCA I at the neutral pH.
However, since the shorter lifetime of the fluorophore in the presence
of the enzyme (13.5 ns) is similar to that observed in the presence of
90% dioxane (12.5 ns), we surmise that the shorter lifetime
component is given by the microscopic state of the enzyme which
harbors the protonated (i.e., non-ionic) form of the sulfonamide
moiety of JB2-48 (see Discussion). To test this hypothesis, we
compared the lifetime profiles of hCA I–JB2-48 complex at neutral
pH with those determined at low (pH=5.0; Fig. 4C) and high pH
(pH=9.0; Fig. 4D) values. Such comparison revealed that unlike at
neutral pH, the excited state fluorescence decay traces at low and high
pH values conformed to single lifetimes with magnitudes of 14 and
20 ns, respectively. Note that the latter values are similar to the
shorter (13.5 ns) and longer (27.3 ns) lifetime components for the
excited state decay data of the enzyme bound JB2-48 at pH 7.0
(Fig. 4B). Evidently, the prevalence of two lifetimes of the enzyme
bound JB2-48 at the neutral pH, but single lifetimes at lower and
higher pH values imply that ionized and neutral forms of sulfonamide
moiety of JB2-48 are stabilized by the corresponding microscopic
states of the enzyme. In one such state, the fluorescence properties of
the enzyme bound JB2-48 is primarily given by the ligand harboring
the neutral form of sulfonamide (i.e., at pH 5.0), where in the other
state it is given by the negatively charged sulfonamide moiety (i.e., at
pH 9.0) of the ligand.
3.1. Effects of solvent polarity and the enzyme's active site
microenvironment on the steady-state and fluorescence lifetimes of
JB2-48
Both structural as well as isothermal titration microcalorimetric
(ITC) data for the binding of ligands to hCA I revealed that the active
site pocket of the enzyme is predominantly hydrophobic [17,28]. To
probe as to the extent such hydrophobicity modulates the fluores-
cent properties of the enzyme bound JB2-48, we determined the
emission maxima (λex =330 nm) and lifetimes of the fluorophore in
the presence of increasing concentration (from 0 to 90%) of dioxane.
Since we used 10% DMSO to solubilize the fluorophore, we could
maintain the concentration of dioxane between 0% and 90%. As
shown in Table 2, as the concentration of dioxane increases, the
fluorescence emission maxima of JB2-48 progressively shifts to blue.
We further observed that at all concentrations of dioxane, the excited
state decay profile of JB2-48 (λex =340 nm, λem =470 nm; see
Experimental procedures) conforms to the single lifetime with
increasing magnitude as a function of the dioxane concentration.
This feature is consistent with our previous observation [38] that the
fluorescence emission intensity of the fluorophore increases with
increase in dioxane concentration. It should be pointed out that due
to solubility problem, we could not reliably perform the above
experiment in different solvents to ascertain the contributions of
hydrogen bonding and/or other microenvironment effect on the
spectral features of JB2-48.
We attempted to ascertain what concentration of dioxane
qualitatively mimics the hydrophocity of the enzyme's active site
pocket. In this endeavor, we determined the fluorescence lifetimes of
the enzyme bound JB2-48 at neutral (pH 7.0), acidic (pH 5.0), and
alkaline (pH 9.0) pH values (Fig. 4). By coincidence, we noted that the
fluorescence lifetime of JB2-48 in 90% dioxane (12.5 0.1 ns; Table 2)
was similar to the shorter lifetime of the enzyme bound fluorophore
at pH 7.0 (see below). In Fig. 4, we show a comparative account of
fluorescence decay profiles of JB2-48 in the presence of 90% dioxane
(panel A), and JB2-48 bound to hCA I at pH 7.0 (panel B), pH 5.0 (panel
C), and pH 9.0 (panel D). The solid smooth lines are the best fit of the
data for one (panels A, B, and D) or two (panel B) lifetimes of the
fluorophores; the residuals of the fitted data are shown at the bottom
of the individual panel. A comparison of the lifetime data of Fig. 4A
and B revealed that unlike the excited state decay profile of JB2-48 in
the presence of 90% dioxane (conforming to the single lifetime profile
3.2. Effect of temperature on the fluorescence lifetimes of free and
enzyme bound JB2-48
Although our above experimental data led to the suggestion that
the differential fluorescence profiles (including lifetimes) of JB2-48
under different environmental conditions were contributed by the
hydrophobic and electrostatic interactions within the active site
pocket of hCA I, we did not rule out the possibility that such changes
originated from the restriction in rotational freedom between the two
aromatic rings of the fluorophore as delineated in the case of green
fluorescent protein [43]. However, such possibility seemed unlikely
since both parent fluorophore (dansylamide) as well as its different
derivatives [38] showed increase in fluorescence as well as the blue
shift in their emission maxima. In addition, our pKa measurement of
free JB2-48 (Fig. 2B) reveals that the phenolic hydroxyl group yields
an unusually low pKa (6.5 instead of 10 for phenol) value, presumably
because of delocalization of the electrons (formed upon deprotona-
tion of the phenolic hydroxyl group) via extended conjugation
between the two aromatic rings as well as the terminal sulfonamide
moiety. Hence, even the aqueous form of JB2-48 (at least at neutral
and basic pH values) is likely to be devoid of the rotational freedom as
noted with the individual fluorophoric units of green fluorescent
protein [43]. To substantiate or refute this conclusion, we determined
the fluorescence lifetimes of free JB2-48 as well as its enzyme bound
form at pH 7.0 as a function of temperature. As shown in Table 3, the
increase in temperature from 25 to 40 °C (the range where enzyme
remains fully active) has practically no effect on the lifetimes of either
free or the enzyme bound JB2-48. Hence, the enhancement in the
fluorescence intensity of JB2-48 is not due to restriction in the
rotational freedom (enhancing internal conversion) around the
“bridged atoms” connecting the two aromatic rings of the fluorophore
upon binding to the enzyme.
Table 2
Fluorescence emission maxima and lifetimes of JB2-48 in dioxanea.
% Dioxane
Emission maxima (nm)
Lifetime (ns)
0
528
522
515
nd
1.2 0.02
2.4 0.05
4.9 0.09
8.4 0.1
25
50
75
90
3.3. Transient kinetic studies for the binding of JB2-48 to hCA I
494
12.5 0.1
We performed transient kinetic studies for the binding of JB2-48
to hCA I at pH 5.0 and 9.0, respectively. This was accomplished by
mixing the above species via the stopped flow syringes, followed by
a
All solutions contained 10% DMSO. λex =336 for determining emission maxima and
340 nm (LED source) for determining lifetimes. nd—not determined.

1970
S. Manokaran et al. / Biochimica et Biophysica Acta 1804 (2010) 1965–1973
Fig. 4. Time-resolved excited state decay profiles of JB2-48 in the presence of dioxane as well as hCA I at different pH values. The time courses for the decay of the excited state of
4 μM JB2-48 (λex =340 nm, λem =470 nm) in the presence of 90% dioxane (Panel A) and 50 μM hCA I at different pH values (Panels B–D) The solid smooth lines are the best fit of
the data for one (Panels A, C, and D) and two (Panel B) lifetimes (τ) under different experimental conditions. Panel A (90% dioxane) τ=12.5 0.1 ns, Panel B (hCA I, pH 7.0)
τ
1 =13.5 0.1 ns, τ2 =27.3 0.4 ns, Panel C (hCA I, pH 5.0) τ=14 0.6 ns, and Panel D (hCA I, pH 9.0) τ=20 0.7 ns. The residuals of the fitted data are given at the bottom of
the lifetime traces of the individual panel.
monitoring the increase in fluorescence intensity of the probe
(λex =336 nm, cutoff filter=395 nm) as a function of time. Fig. 5
shows the stopped flow traces for the binding of JB2-48 with hCA I
under pseudo-first order condition ([JB2-48]NNhCA I]) at the above
pH values. The solid smooth lines are the best fit of the data for
the single exponential rate equation with rate constants of 0.08 and
0.37 s−1 at pH 5.0 and 9.0, respectively. These rate constants are
Table 3
Effect of temperature on lifetimes of JB2-48a.
∘
translated into the activation energies (ΔG ‡ =−RT ln (6.2×1012/k))
of 18.9 and 18.0 kcal/mol at the above pH values, respectively. In view
of our earlier argument, the difference in the putative transition state
Temperature
( C)
Free JB2-48
hCA I–JB2-48 complex
τ1 (ns)
∘
τ (ns)
τ2 (ns)
energies (ΔΔG ‡ =0.9 kcal/mol) for the binding of JB2-48 to hCA I
∘
25
30
35
40
1.2 0.02
1.3 0.03
1.3 0.03
1.2 0.03
13.5 0.1
12.7 0.004
11.6 0.1
12.8 0.2
27.3 0.4
29.1 0.01
28.6 0.01
28.7 0.3
between pH 5.0 and 9.0 serve as the quantitative measure of the
electrostatic contribution (involving the negatively charged sulfon-
amide group of JB2-48 and the Zn2+ cofactor) in stabilizing the ligand
in the transition state.
To further confirm that the magnitude (as well as the associated
transient rate constants) of fluorescence changes for the binding of
a
In 25 mM HEPES buffer, pH 7.0, containing 10% DMSO. λex=340 nm (LED) and λem
=
470 nm. [JB2-48]=35 μM for measuring the lifetimes of free fluorophore. [JB2-48]=4 μM
and [hCA I]=50 μM for measuring the lifetimes of the enzyme bound fluorophore.

S. Manokaran et al. / Biochimica et Biophysica Acta 1804 (2010) 1965–1973
1971
Fig. 6. pH jump relaxation kinetic studies of hCA I bound JB2-48. The stopped-flow
traces for the change in fluorescence of the hCA I–JB2-48 complex upon mixing (the
after mixing concentrations of hCA I and JB2-48 being equal to 1 μM and 20 μM,
respectively) with buffers of selected pH values are shown. The pH jump from 5 to 9
(increase in fluorescence) was accomplished by mixing the hCA I–JB2-48 complex in
5 mM acetate buffer (pH 5.0) with 200 mM Tris buffer (pH 9.0). The pH of the mixture
was found to be 9.0. The pH jump from 9 to 5 (decrease in fluorescence) was
accomplished by mixing the hCA I–JB2-48 complex in 5 mM Tris buffer (pH 9.0) with
200 mM acetate buffer (pH 5.0). The pH of the mixture was found to be 5.0. The solid
smooth lines are the best fit of the data according to the single-exponential rate
equation with the rate constants of 0.67 0.003 s−1 and 0.078 0.0005 s−1 for the pH
jump experiments from 5 to 9 and from 9 to 5, respectively.
4. Discussion
The experimental data presented in the previous section lead to the
following conclusions: (1) Due to its size and extended conjugation,
JB2-48 occupies nearly the entire active site pocket (about 15 Å deep)
of hCA I, and this feature allows for deciphering the hydrophobic
versus electrostatic contributions on the spectral, kinetic, and
thermodynamic features of the enzyme–ligand complex. (2) Whereas
the pKa of sulfonamide moiety of free JB2-48 is 10.2 that of the phenolic
hydroxyl group is 6.5. (3) Depending upon the pH of the buffer media,
the enzyme bound JB2-48 exists in either neutral (with sulfonamide
group being fully protonated) or anionic form (pKa =6.6), and these
forms are stabilized by the cognate microscopic states of the enzyme.
(4) The neutral and anionic forms of the enzyme bound JB2-48 are
distinguishable by their fluorescent lifetimes; whereas the fluorescent
lifetime of the neutral form falls in the range of 12–14 ns, that of the
anionic form is greater than 20 ns. Consistently, the anionic form of the
enzyme bound JB2-48 is more fluorescent than its neutral counterpart.
(5) Due to added electrostatic interaction (between the enzyme
resident Zn2+ cofactor and the negatively charged sulfonamide
group), the anionic form of JB2-48 is stabilized at the active site of
hCA I by about 2.2 kcal/mol energy than its neutral form. (6) The rate
of fluorescence changes accompanying the transient course of binding
of JB2-48 to hCA I is about 5 fold faster with anionic than neutral form
of the ligand, unraveling the fact that the putative transition state is
about 0.9 kcal/mol more favorable with the anionic form of the ligand
than the neutral form. (7) The similarity in the rate constants for the
association of JB2-48 with hCA I (at acidic and basic pH values) with
those derived from the pH jump experiments lead to the suggestion
that the active site pocket of the enzyme undergoes slow restructuring
during the course of the ligand binding.
Fig. 5. Transient kinetics for the binding of JB2-48 to hCA I at acidic and basic pH
values. The stopped flow traces for the mixing of JB2-48 with hCA I (λex =336 nm,
“cutoff” filter=395 nm) at pH 5.0 and 9.0 are shown in Panels A and B, respectively. The
after-mixing concentrations of hCA I and JB2-48 were 1 and 30 μM, respectively. The solid
smooth lines are the best fit of the data according to the single exponential rate equation
for the relaxation rate constants (kobs) of 0.08 0.004 and 0.37 0.001 s−1 at pH 5.0 and
9.0, respectively.
JB2-48 to hCA I between the low (Fig. 5A) and high (Fig. 5B) pH values
were given by the neutral versus the ionized forms of the sulfonamide
moiety of the ligand, respectively, we performed the “pH jump”
experiment. In this endeavor, we mixed hCA I–JB2-48 complex present
in 5 mM Tris buffer at pH 9.0 with 200 mM acetate buffer at pH 5.0 (the
pH of the mixture emerged out to be 5.0) via the stopped flow syringes,
and monitored the time course of the fluorescence changes. Fig. 6
shows the time dependent decrease in fluorescence of the hCA I–JB2-
48 complex upon pH jump from 9 to 5. An essentially identical
experiment was performed for the mixing of hCA I–JB2-48 complex
maintained at pH 5.0 with 200 mM Tris buffer of pH 9.0 (the pH of the
mixture was 9.0). Note that the opposite pH jump (i.e., from 5.0 to 9.0)
resulted in the time dependent increase (Fig. 6) in fluorescence of the
enzyme–probe complex. Both the traces of Fig. 6 conformed to the
single exponential rate equation with rate constants of 0.078 and
0.67 s−1 for the pH jump from 9 to 5 and that from 5 to 9, respectively.
On the basis that the protonation/deprotonation reactions are
diffusion limited [40], the origin of the above rate constants must lie
in the slow change in the electronic structure of the hCA I bound JB2-
48, which is likely to originate from the readjustment/packing of the
enzyme–ligand complex. This is further supported by the fact that the
transient rate constants for the association of JB2-48 to hCA I at pH 5.0
and 9.0 (Fig. 5) are comparable to those obtained from the pH jump
experiments from 9 to 5 and from 5 to 9 (Fig. 6), respectively (see
Discussion).
This is the first demonstration (to the best of our knowledge) that
the fluorescence spectral changes upon binding of JB2-48 to hCA I is
contributed both by the hydrophobic environment of the enzyme's
active site pocket as well as the electrostatic interaction between the
active site resident Zn2+ cofactor and the negatively charged
sulfonamide group of the ligand. Our temperature dependent lifetime
data of both free and the enzyme bound JB2-48 (Table 3) as well as the

1972
S. Manokaran et al. / Biochimica et Biophysica Acta 1804 (2010) 1965–1973
pKa values of free fluorophore eliminates the possibility that the
enhancement in fluorescence of JB2-48 (upon binding to the enzyme
site) is due to restriction in the rotational freedom between the two
aromatic rings as observed with green fluorescent protein [43]. We
believe the fluorescence changes upon binding of JB2-48 (vis a vis
dansylamide; [34]) to hCA I is more pronounced due to an extended
conjugation of the π electrons between the two aromatic rings. The
fact that the fluorescence intensity of JB2-48 increases with decrease
in the solvent polarity [38] attests to the contribution of the
hydrophobic active site environment of hCA I in enhancing the
fluorescence emission intensity of the ligand. However, since the
fluorescence emission intensity of the enzyme bound JB2-48 is
significantly higher at neutral and basic pH values (as compared to
that obtained in the presence of 90% dioxane) implies that aside from
hydrophobicity, some additional factor(s) of the enzyme site phase is
involved in enhancing the fluorescence intensity of the ligand. Based
on the structural as well as the spectroscopic data [18–20,26,27,30], it
appeared evident that such an “additional factor” is the electrostatic
interaction between the active site resident Zn2+ and the negatively
charged sulfonamide group of the ligand. Hence, it is not surprising
that the decrease in pH diminishes the fluorescence emission
intensity of the enzyme bound JB2-48 (Fig. 2).
In view of the fact that the pKa value of the enzyme bound JB2-48
is equal to 6.6, it is conceivable that the enzyme bound form of the
above ligand would remain in fully protonated and ionized states at
pH 5 and 9, respectively. This feature is further corroborated by the
lifetime measurements of JB2-48 under different experimental
conditions (Fig. 4). Since the lifetime of “free” JB2-48 in 90% dioxane
(τ=12.5 ns) is similar to the lifetime of the enzyme bound JB2-48 at
pH 5.0 (τ=14 ns), it supports the notion (rather quantitatively)
that the fluorescence profile of the enzyme bound ligand at pHbpKa
is dominated by the hydrophobic interaction within the enzyme's
active site pocket. This is in contrast to the lifetime of the enzyme
bound JB2-48 at pH 9.0 being equal to 20 ns (Table 1), supports our
hypothesis that at pHNpKa, the fluorescence profile of the enzyme
bound ligand is contributed by a combination of hydrophobic and
electrostatic interactions. Hence, it is not surprising that at pH 7.0
(i.e., pH≈pKa), the enzyme bound JB2-48 yields two lifetimes of
13.5 ns (τ1) and 27.3 (τ2 ), of which τ1 and τ2 are similar to the
lifetimes of the enzyme bound ligand at pH 5.0 (or the free ligand in
the presence of 90% dioxane) and at pH 9.0, respectively. Since
under physiological conditions CAs are known to bind one molecule
of arylsulfonamide ligand per monomeric unit of the enzyme
[15,17–22,28,29,41], it appears plausible that τ1 and τ2 are
associated with the two alternative microscopic states of the
enzyme (see the cartoon of Fig. 7).
enzyme. In pursuit of answering this question, we note that the
observed rate constants during the pH jump experiments (Fig. 6) are
several orders of magnitude lower than those expected for the simple
protonation/deprotonation (which are considered to be the diffusion
limited process; 49) step of the sulfonamide group of the enzyme
bound ligand. Evidently, the protonation/deprotonation of the
enzyme-bound ligand may not be the rate limiting step of the
fluorescence changes during the pH jump experiments. The overall
rate is neither expected to be contributed by the rate of decay of the
excited state of the enzyme bound fluorophores since the lifetimes of
the enzyme bound JB2-48 falls in the range of nanoseconds. However,
one can argue that the slower rate constant for the increase in the
fluorescence of the enzyme bound JB2-48, particularly during the
9→5 pH jump experiment (kobs =0.08 s−1), is due to slow (rate
limiting) breakdown of the Zn2+-(anionic) sulfonamide bond of the
ligand prior to the diffusion limited protonation step. Unfortunately,
the above possibility falls short in explaining the comparably lower
rate of fluorescence changes (kobs =0.67 s−1) during the 5→9 pH
jump experiment. Aside from these, the observed rate constants for
the association of JB2-48 to hCA I at pH 5 (kobs =0.08 s−1) and 9
(kobs =0.37 s−1) (Fig. 5) are similar to the rate constants for the 9→5
and 5→9 pH jump experiments, respectively. Such a similarity in the
rate constants has been somewhat surprising since these experiments
are mechanistically different. In one case, the transient course of the
fluorescence changes (as a function of pH) occur due to the
association of the ligand to the enzyme, but in other case, the overall
process is manifested via the protonation/deprotonation of the
enzyme bound ligand. These arguments, in conjunction with our
earlier transient kinetic studies for the binding of dansylamide to hCA
I and hCA II [34], prompt us to propose that the two alternative
microscopic states of the hCA I (Fig. 7) are representative of two
alternative conformational states of the enzyme which stabilize
neutral and anionic forms of the ligand, and the transition between
the above states serve as the rate limiting step during the pH
dependent experiments. It should be emphasized that the conforma-
tional changes during the ligand binding and/or the pH jump
experiments need not be extensive to be deciphered via the
fluorescence techniques—they can just be subtle and/or dynamic in
nature to elicit the fluorescence changes reported herein.
The mechanistic conclusion derived herein has potential to find
applications in the rationale design of hCA inhibitors as therapeutic
agents. It appears evident that the potency of an arylsulfonamide
inhibitor of hCAs can be increased by incorporating non polar groups
toward the exterior region of the active site pocket as well as by
introducing the electronic withdrawing groups on the aromatic ring
(s) of the inhibitor [13,14,28,31–33]. The latter groups would facilitate
the deprotonation of the sulfonamide moiety (of the inhibitor) such
that it's anionic form would interact more strongly with the Zn2+
cofactor, and the overall structure would be stabilized by the cognate
The question arose whether the two microscopic states of the
hCA I, harboring neutral versus anionic forms of JB2-48, respectively,
are representative of the alternative conformational states of the
Fig. 7. Cartoon showing the stabilization of neutral and anionic forms of JB2-48 at complementary microscopic states of hCA I. The factors affecting the reversible transition between
the two forms of the enzyme–ligand complexes are shown.

S. Manokaran et al. / Biochimica et Biophysica Acta 1804 (2010) 1965–1973
1973
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This research was supported by the NIH grants CA113746 and
CA132034 and the National Science Foundation grant DMR-0705767
to D.K.S and S.M. Sumathra Manokaran was supported by the National
Science Foundation—Experimental Program to Stimulate Competitive
Research Award [grant number EPS-0814442].
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