Effect of prolonged exposure to organic solvents on the active site environment of subtilisin Carlsberg

Article abstract of DOI:10.1016/j.molcatb.2010.01.021

The potential of enzyme catalysis as a tool for organic synthesis is nowadays indisputable, as is the fact that organic solvents affect an enzyme's activity, selectivity and stability. Moreover, it was recently realized that an enzyme's initial activity is substantially decreased after prolonged exposure to organic media, an effect that further hampers their potential as catalysts for organic synthesis. Regrettably, the mechanistic reasons for these effects are still debatable. In the present study we have made an attempt to explain the reasons behind the partial loss of enzyme activity on prolonged exposure to organic solvents. Fluorescence spectroscopic studies of the serine protease subtilisin Carlsberg chemically modified with polyethylene glycol (PEG-SC) and inhibited with a dansyl fluorophore, and dissolved in two organic solvents (acetonitrile and 1,4-dioxane) indicate that when the enzyme is initially introduced into these solvents, the active site environment is similar to that in water; however prolonged exposure to the organic medium causes this environment to resemble that of the solvent in which the enzyme is dissolved. Furthermore, kinetic studies show a reduction on both V_max and K_M as a result of prolonged exposure to the solvents. One interpretation of these results is that during this prolonged exposure to organic solvents the active-site fluorescent label inhibitor adopts a different binding conformation. Extrapolating this to an enzymatic reaction we argue that substrates bind in a less catalytically favorable conformation after the enzyme has been exposed to organic media for several hours.

Full text of DOI:10.1016/j.molcatb.2010.01.021

Journal of Molecular Catalysis B: Enzymatic 64 (2010) 38–44
Contents lists available at ScienceDirect
Journal of Molecular Catalysis B: Enzymatic
journal homepage: www.elsevier.com/locate/molcatb
Effect of prolonged exposure to organic solvents on the active site
environment of subtilisin Carlsberg
Vibha Bansala,1, Yamixa Delgadoa, Ezio Fasolia, Amaris Ferrera, Kai Griebenowb,
Francesco Secundoc, Gabriel L. Barlettaa,∗
a University of Puerto Rico at Humacao, Department of Chemistry, Humacao 00791, Puerto Rico
b University of Puerto Rico at Rio Piedras, Department of Chemistry, Puerto Rico
c Istituto di Chimica del Riconoscimento Molecolare, v.M. Bianco, Milano, Italy
a r t i c l e i n f o
a b s t r a c t
Article history:
Received 6 August 2009
Received in revised form 21 January 2010
Accepted 25 January 2010
Available online 2 February 2010
The potential of enzyme catalysis as a tool for organic synthesis is nowadays indisputable, as is the
fact that organic solvents affect an enzyme’s activity, selectivity and stability. Moreover, it was recently
realized that an enzyme’s initial activity is substantially decreased after prolonged exposure to organic
media, an effect that further hampers their potential as catalysts for organic synthesis. Regrettably, the
mechanistic reasons for these effects are still debatable. In the present study we have made an attempt to
explain the reasons behind the partial loss of enzyme activity on prolonged exposure to organic solvents.
Fluorescence spectroscopic studies of the serine protease subtilisin Carlsberg chemically modified with
polyethylene glycol (PEG-SC) and inhibited with a dansyl fluorophore, and dissolved in two organic
solvents (acetonitrile and 1,4-dioxane) indicate that when the enzyme is initially introduced into these
solvents, the active site environment is similar to that in water; however prolonged exposure to the
organic medium causes this environment to resemble that of the solvent in which the enzyme is dissolved.
Furthermore, kinetic studies show a reduction on both Vmax and KM as a result of prolonged exposure to
the solvents. One interpretation of these results is that during this prolonged exposure to organic solvents
the active-site fluorescent label inhibitor adopts a different binding conformation. Extrapolating this to
an enzymatic reaction we argue that substrates bind in a less catalytically favorable conformation after
the enzyme has been exposed to organic media for several hours.
Keywords:
Enzyme catalysis in organic solvents
Enzyme storage stability in organic solvents
Active-site polarity
Subtilisin Carlsberg
Published by Elsevier B.V.
1. Introduction
quent exposure to organic media generally does not perturb the
enzyme’s structure [5,6], although there are some accounts in the
Although during the last 20 years enzymes in organic solvents
have been successfully used to catalyze a range of important reac-
tions, there are still some drawbacks to overcome before their
potential can be fully exploited. Among then is the fact that their
initial activity decreases during prolonged exposure to organic
solvents. This is particularly significant for lengthy reactions, for
the construction of bioreactors, or for reusing the biocatalyst.
Several studies have addressed issues such as shelf life, biore-
actor stability and operational stability of a variety of enzymes
suspended in organic solvents and in aqueous–organic solvent mix-
tures [1–3]. It is well documented that thedehydration step (usually
lyophilization), typically required prior to suspension in organic
media, causes minor structural perturbation [4], whereas subse-
literature that contradicts this assessment [7,8]. Nevertheless, the
mechanism of enzyme partial inactivation in organic solvents is
still unclear. Recent studies on the serine protease subtilisin Carls-
berg (SC) have shown that this detrimental effect was observed
in different solvents and under different experimental conditions,
suggesting that this effect is independent of the physicochemical
properties of the organic medium, its hydration state, and the reac-
tion temperature. It was also determined that the enzyme remains
structurally defined during prolonged exposure to organic solvents
[9,10]. In a separate study electron paramagnetic resonance spec-
troscopy (EPR) was used to gain vital information about the effect of
prolonged exposure to an organic solvent on the mobility of a spin
label bound to the active site of SC (which in turn could be extrap-
olated to a similarly bound substrate in the active site). Although
those results were inconclusive as to pinpointing the mechanis-
tic reason for this observed decrease of initial enzyme activity,
two different components were observed in the EPR spectra whose
percentages changed during the incubation period, giving the first
indication that perhaps the observed activity loss was due to an
∗
Corresponding author. Tel.: +787 850 0000x9055; fax: +787 850 9422.
E-mail address: gabriel.barletta@upr.edu (G.L. Barletta).
Current address: University of Puerto Rico at Cayey, Department of Chemistry,
1
Cayey 00736, Puerto Rico.
1381-1177/$ – see front matter. Published by Elsevier B.V.
doi:10.1016/j.molcatb.2010.01.021

V. Bansal et al. / Journal of Molecular Catalysis B: Enzymatic 64 (2010) 38–44
39
effect localized at the active site [9]. Furthermore, two recent
studies shed additional light to this phenomenon: the first study
looked at the enzyme dynamics by measuring the H/D exchange by
NMR, and showed that globally, the enzyme becomes more rigid
upon storage in acetonitrile (ACN) and 1,4-dioxane [11]. As previ-
ously observed the enantioselectivity remained constant, which is
an indication that the active-site structure remained intact. More
importantly however, was the fact that there was no correlation
between enzyme flexibility and activity. The most active prepa-
ration (PEG-SC) was less flexible than the less active lyophilized
preparation, and both preparations were more flexible in ACN than
in 1,4-dioxane (the enzyme is more active in the second solvent)
[11]. These results suggested to us that decreased global flexibil-
ity might not be the only cause of the observed loss of activity
upon prolonged storage in organic media, and that other factors
must be involved. The second study looked at the dynamics of the
same enzyme in acetonitrile during 92 ns. This molecular dynam-
ics approach showed that a key residue in the active site change
its orientation, diminishing the ability of the oxyanion hole to sta-
bilize the tetrahedral intermediates [12]. This study also showed
that most internal water molecules are exchanged by the solvent
used (ACN), thus changing the environment (the polarity) of the
active site. Although the study represents a snapshot of the ini-
tial events after exposure to organic media, it is an indication that
additional changes (such as water removal and reorientation of
additional residues and side chains) might take place during pro-
longed exposure to organic media (Cruz et al., 2009). In an effort
to shed more light to the mechanism of enzyme partial inactiva-
tion upon prolonged exposure to organic solvents, we decided to
study the environment of the active site of SC during storage in
ACN and 1,4-dioxane—solvents in which SC exhibits different cat-
alytic properties, directly, by fluorescence spectroscopy. There is
a common characteristic shared by most enzyme-in-organic sol-
vents systems that limits the types of studies that can be carried
out with then, and that is the fact that they are virtually insoluble in
organic solvents. This is particularly relevant for the types of studies
we are about to discuss since suspended enzyme particles would
impede and obscure fluorescence experiments. To overcome this
problem we chemically modified our enzyme with polyethylene
glycol (PEG) of 5 kDa molecular weight, a technique which has been
shown to increase the solubility of several enzymes in some organic
solvents without affecting or altering their secondary and tertiary
structure [10,13–15]. However, the degree of solubility of a par-
ticular PEG-enzyme system varies among solvents. In our case, we
found the enzyme to be highly soluble in ACN, 1,4-dioxane, toluene
and dichloromethane. The last one was found to be detrimental to
enzyme structure (determined by 1H NMR), and toluene turned out
to be a bad choice for our spectroscopic experiments. Other sol-
vents tested render a partially soluble enzyme while in others the
PEG-enzyme precipitated during the long incubation period. This
restricted our choices to 1,4-dioxane and ACN, which are actually
good picks since previous studies found that SC is structurally stable
in these solvents [9]. Additionally, in these two solvents SC exhibits
vastly different activity and enantioselectivity (the enzyme is much
more active and enantioselective in 1,4-dioxane than in ACN).
Fluorescence spectroscopy is a powerful tool to study discrete
sites in proteins by looking at the emission of intrinsic or extrinsic
fluorophores. For these studies we chose two very basic analyses
based on steady-state fluorescence emission in organic solvents.
Similar studies (but in aqueous systems) have previously been
reported by Vaz and Schoellmann [16,17]. Variables such as quan-
tum yields and molar extinction coefficients were calculated for the
system with respect to the solvents being used. Our model enzyme
was studied with respect to two fluorophores: Trp residue present
at position 113 (Trp113) as the intrinsic fluorophore (the enzyme’s
only intrinsic fluorophore) and a dansyl group (D), bound to the
active site Ser221 (Ser221-D) of the enzyme, as the extrinsic fluo-
rophore. D binds exclusively at the active site serine residue of SC
with a concomitant loss of enzymatic activity (indicative that it
does bind to the active site) and hence serves as an effective probe
for the active site studies [17]. According to Turner and Brand the
emission maximum of adsorbed probes gives the most reliable esti-
mate of the binding site polarity [18]. The high solvent sensitivity of
both these fluorophores (Trp and D) makes them particularly use-
ful for studying their microenvironment on the enzyme such as the
polarity. We have thus used this system to study the effect of ACN,
1,4-dioxane and aqueous medium (sodium phosphate buffer) on
the active-site polarity of the enzyme. The fluorescence and kinetic
studies presented here and those previously published suggests
that over time, substrates prefer a binding orientation that is not
catalytically favorable.
2. Materials and methods
2.1. Materials
Subtilisin Carlsberg (alkaline protease from Bacillus licheni-
formis, EC 3.4.21.14) was purchased as lyophilized powder, the
solvents were all purchased in the anhydrous form (Aldrich
Sure/Seal bottles—water content below 0.005%) and used with-
out further drying, and sec-phenethyl alcohol were purchased
from Sigma–Aldrich (St. Louis, MO). Vinyl butyrate and 5-
(dimethylamino)-1-naphthalenesulfonyl fluoride (dansyl fluoride,
DF) were purchased from TCI America (Portland, OR). N-
Succinimidyl PEG (5 kDa) was ordered from NeKtar Therapeutics
(Huntsville, AL). Sephadex PD-10 desalting columns were pur-
chased from Amersham Biosciences.
2.2. Methods
2.2.1. Chemical modification of the enzyme with polyethylene
glycol (PEG)
Subtilisin Carlsberg was PEGylated with 10-fold molar excess
of 5 kDa PEG in sodium borate buffer (pH 9.2). The PEGylation
was allowed to proceed with constant stirring for 15 min at 20 ◦C
and then 3 h at 4 ◦C. After this the pH of the reaction mixture was
reduced to 5.5 with dilute HCl to stop the reaction and the solution
was dialyzed for 24 h in 25 kDa cutoff dialysis bag. The dialyzed
solution was finally lyophilized for 48 h.
2.3. Preparation of dansyl methoxide (D-OMe) solution
Fifty milligrams of (0.198 mmol) of dansyl fluoride were dis-
solved in 1 mL of methanol in a vial. 0.350 mL of a 5.6 M solution
of sodium methoxide was added and the solution let under stirring
for 15 min at room temperature. The suspension was centrifuged
at 5000 rpm for 5 min. The supernatant was separated and used for
further analysis.
2.4. Preparation of dansyl hydroxide (D-OH) solution
Fifty milligrams (0.198 mmol) of dansyl fluoride were dissolved
in 1 mL of isopropyl alcohol. 0.02 mL of the solution was added to a
1 mL of 1 M solution of sodium hydroxide in water and the solution
let under stirring at room temperature for 15 min.
2.5. Fluorescence spectroscopy
The enzyme was labeled with DF according to the method
of Vaz and Schoellmann [16]. Active site titration (using N-trans
cinnamoyl imidazole as a substrate) to monitor the labeling
effectiveness revealed 100% inhibition of enzyme activity, thus

40
V. Bansal et al. / Journal of Molecular Catalysis B: Enzymatic 64 (2010) 38–44
indicating 100% labeling with the dansyl group (D). Note that fluo-
rine is released during the reaction.
2.10. Protein concentration
Protein concentrations were assayed by the bicinchoninic acid
(BCA) method using bovine serum albumin as standard [23].
2.6. Steady-state fluorescence emission spectra
2.11. Water activity control
Corrected steady-state emission spectra of the enzyme with
and without dansyl were recorded with a Horiba Jobin Yvon
Fluoromax-3 spectrofluorometer equipped with a xenon arc lamp.
Trp fluorescence emission was collected from 310 to 600 nm by
exciting directly the Trp113 in the non-labeled as well as dansyl-
labeled SC (Ser221-D) at 295 nm. Ser221-D fluorescence emission
was collected from 370 to 700 nm by exciting directly the dan-
syl at 360 nm. The excitation and emission slit widths were 2 nm
for all spectra. Background fluorescence from the solvents was
recorded and subtracted from the protein spectra. Steady-state
measurements were performed in a 3.0 ml square fluorescence
cuvette with 10 mm excitation/emission path lengths (emission
collected through a perpendicular face). The absorbance of the
samples was lower than 0.05 at ꢀex = 295 nm to avoid inner-
filter effects. All measurements were performed at 20 ◦C. The
effect of long-term exposure of the enzyme to the organic sol-
vents was studied by collecting the emission spectra at regular
intervals of 24 h for 4 days. The molar extinction coefficients at
360 nm were determined for free DF dissolved in the respective
solvents.
Some of the activity determinations and fluorescence experi-
ments were performed under controlled water activity by adding
salt hydrate pairs to the samples (sodium acetate 3/0, water activ-
ity = 0.28) [24]. Equal weights (50 mg each) of each of the two
differently hydrated sodium acetate salts were added to 750 ␮l of
the solvent and allowed to equilibrate for 30 min and then added
13.0 mg of protein to the solvent–salt mixture and allowed to
equilibrate for another 60 min. Fluorescence spectra/activity were
determined thereafter.
3. Results
3.1. Enzyme activity in organic solvents
The transesterification activity of PEGylated SC was determined
in dioxane and in ACN during incubation at 25 ◦C under controlled
and uncontrolled water activity conditions (Fig. 1). As previously
reported, the enzyme activity decreased approximately 10-fold
during a 4-day incubation performed in neat organic solvents [10].
Under controlled water activity conditions (using saturated salt
solutions), the initial activity was low and it decreased further by
1.6-fold during incubation. A similar effect was observed in the case
of ACN. However, the activity in 1,4-dioxane was higher than that
in ACN (Fig. 1). We proceeded henceforth to explore the polarity
2.7. Data analysis
The corrected emission spectra from the D-free and D-labeled
SC were used to determine the emission maxima of Trp/D in the
enzyme in different solvents. The emission maxima of dansyl bound
to the active site of SC were further used to calculate the active-site
polarity on the basis of data published by Vaz and Schoellmann and
Kosower [17,19]. A linear correlation (y = −113.03x + 288.24) was
obtained from a plot between the wave number (ꢁ) and empirical
polarities (Z) listed by Vaz and Schoellmann (1976) for dansyl-
ethyl ester as a reference compound for SC [16], which was used
to convert the emission maxima obtained for Ser221-D in differ-
ent solvents into the microenvironment polarities on the Z-scale.
Absorption spectra were obtained using a Hewlett Packard 8453
UV–vis spectrophotometer.
2.8. Enzyme preparation and kinetic measurements
PEGylated SC powder was prepared by lyophilization from a
solution of 5 mg SC per mL of a 20 mM potassium phosphate
buffer, pH 7.8, for 24 h. The transesterification reaction between
sec-phenethyl alcohol and vinyl butyrate was used in all of the
initial activity and enantioselectivity measurements, and the reac-
tions were carried out at 25 ◦C under anhydrous conditions (except
in the case of the controlled water activity experiments). These
reactions were done as previously described [20]. The retention
times and instrument calibration of the “R” and “S” products were
obtained using samples of the pure enantiomers synthesized from
the corresponding alcohol enantiomers, and the activity was deter-
mined from the sum of both enantiomers of the product [21]. Vmax
and KM were measured following the reaction of phenyl alcohol
and vinyl butyrate and the data analyzed as previously described
by us [9].
2.9. Degree of PEGylation
Fig. 1. Initial rate of PEGylated subtilisin Carlsberg in organic solvents during
incubation at 25 ◦C (measured before 10% product conversion). PEGylated enzyme
concentration: 1.0 mg/mL. Close squares: no water activity control; open squares:
with water activity control (sodium acetate): (A) 1,4-dioxane; (B) acetonitrile.
The degree of PEGylation in terms of number of PEG molecules
per molecule of SC was found to be approximately 4 as determined
by the method of Habeeb et al. [22].

V. Bansal et al. / Journal of Molecular Catalysis B: Enzymatic 64 (2010) 38–44
41
Table 1
Emission maxima of all fluorophores used in the study (all measurements are within a maximum error of 0.91%).
ꢀexcitation (nm)
Emission maxima (nm)a
Days of storage in the respective solvents
0
1
2
3
4
Native SC Trp113 bufferb
PEG-SC Trp113 buffer
Free Trp113 ACN
360
356
337
333
295
360
339
334
341
330
341
333
341
330
Free Trp113 1,4-dioxane
PEG Ser221-D ACN
PEG Ser221-D 1,4-dioxane
PEG Ser221-D buffer
524/445c
510/450c
510
515/445
502/450
515/445
502/450
515/445
502/450
515/445
502/450
510
The emission maxima written in bold letters refer to wavelength at which emission intensities increase during storage.
a
Approximately 2 nm.
Not PEGylated enzyme.
Shoulder: blank spaces stand for no measurement/storage.
b
c
of the active site (by fluorescence spectroscopy) and the binding
kinetics of the substrate to gain a deeper understanding of the rea-
sons that led to the observed partial loss of activity after prolonged
exposure to organic solvents. We proceed to describe each study in
detail.
3.2. Fluorescence spectroscopy
SC was labeled at the active site Ser221 with dansyl fluoride
(Ser221-D). A 100% inhibition of enzyme activity was achieved
under optimal reaction conditions with a labeling ratio of one (see
Section 2). The fluorescence studies in buffer were performed using
two different preparations of the enzyme: the native SC powder
and PEGylated (5 kDa) SC. However, for the studies in ACN and
1,4-dioxane only PEGylated SC was used for the solubility reasons
Fig. 2. Emission spectra of PEGylated and native subtilisin Carlsberg in different
solvents (excitation at 295 nm). Green: 1,4-dioxane; red: buffer; blue: acetonitrile;
brown: native (not PEGylated) in buffer.
discussed in Section 1.
3.3. Polarity studies
3.5. Active-site polarity
According to Stryer, the polarity of the binding site of a fluo-
rophore can be evaluated in terms of two parameters: the emission
maximum and the quantum yield of fluorescence of bound dye.
Emission maximum is indicative of the overall dipolar character of
the solvent shell, while the quantum yield is related to more local-
ized deactivating processes [25]. We examined the polarity at the
enzyme surface and the active site by following the fluorescence
properties of Trp113 and Ser221-D, respectively. Shifts in emission
maxima and changes in quantum yields of both Trp113 and Ser221-D
were seen in H2O, ACN and 1,4-dioxane (Table 1).
The fluorescence emission of the active site bound dansyl flu-
orophore (Ser221-D) was monitored to estimate the active-site
polarity of SC in different solvents after different periods of incu-
bation at 25 ◦C (Table 1). The emission maxima of Ser221-D did
no show a considerable red shift on initial exposure to the sol-
vents studied here, as compared to our controls D-OMe and D-OH
(in the absence of the enzyme, Table 2). The emission maximum
of Ser221-D was more red-shifted in ACN (525 nm) than in buffer
(510 nm), while in dioxane it was similar to the one in buffer
(Table 1). This means that the polarity experienced by Ser221-D in
dioxane is similar to that experienced by Ser221-D in buffer. This
3.4. Polarity on the surface of the enzyme
Since the enzyme’s only Trp residue (Trp113) is on the sur-
face and therefore its emission maximum will be determined by
its interaction with solvent molecules and neighboring residues’
side chains [26,27], we were able to study the polarity at the sur-
face of the enzyme in the different solvents here used and the
effect of prolonged exposure to these solvents. The emission maxi-
mum of Trp113 in the enzyme (without the dansyl moiety) showed
a clear red shift in going from the organic to aqueous solvent
(330 nm in 1,4-dioxane to 360 nm in water, Table 1, Fig. 2). No dif-
ferences were seen between the emission maxima of the Trp113
and free Trp (unbound Trp) in any of the solvents indicating that
the Trp113 emission is determined only by its interaction with the
solvent (Table 1). Additionally, no substantial shift in emission max-
imum of Trp113 was seen during the 5-day incubation period in
organic solvents, which indicates that the surface of the protein
arrives at a fast equilibrium with the bulk of the solvent system
(Table 2).
Table 2
Fluorophore properties for tryptophan and modified dansyl moiety.
ꢀexcitation (nm)
Emission maxima (nm)
Free Trp buffer
Free Trp ACN
Free Trp 1,4-dioxane
295
360
335
330
FreD-F Buffer
FreD-F ACN
FreD-F 1,4-dioxane
360
360
360
524
586
540
FreD-OH Buffer
FreD-OH ACN
FreD-OH 1,4-dioxane
502
460
450
FreD-OCH3 Buffer
FreD-OCH3 ACN
FreD-OCH3 1,4-dioxane
500
450
450
Same color emission maxima are for comparison purposes only.

42
V. Bansal et al. / Journal of Molecular Catalysis B: Enzymatic 64 (2010) 38–44
Table 3
Active-site polarity of subtilisin Carlsberg based on fluorescence emission maxima
of dansyl fluoride bound to the active site Ser221
.
−1
Solvent
Buffer
ꢁF, ␮ (peak 1/2)
Estimated “Z” (Peak 1/2)
1.96
66.61
ACN Day 0
Day 1
Day 2
1.91
72.53
1.94/2.25
1.94/2.25
1.94/2.26
72.53/34.24
72.53/33.67
72.53/33.43
Day 3
Day 4
Dioxane Day 0
Day 1
Day 2
Day 3
1.94/2.26
1.96
1.99/2.22
2.00/2.23
2.00/2.23
2.03/2.23
72.73/33.67
66.61
66.61/37.06
66.61/36.50
66.61/36.50
66.61/36.50
Day 4
The values in bold are derived from emission peaks that increase in intensity during
the incubation.
the active site [28]. However, hydrolysis of the dansyl group is ruled
out since parallel studies showed Förster Resonance Energy Trans-
fer (FRET) (obtained by activating Trp113, located about 20 Å away
from the active site) throughout the incubation period (data not
shown). If the dansyl acceptor was being hydrolyzed and released
from the enzyme, the FRET effect would be drastically reduced or
even disappear as huge concentrations of the acceptor are required
for FRET to occur when the donor and acceptor are not on the
same molecule, or are free in solution [29]. Furthermore, if the
dansyl group is hydrolyzed and released from the active site, the
new fluorophore (dansyl hydroxide) should give rise to a new peak
in the absorption spectrum at 320 nm (Table 2), while we only
observe a small shift to 335 nm (from 343 nm) in the absorption
spectrum of the sample during incubation in 1,4-dioxane (Fig. 4).
This small shift in ꢀmax could also be the result of changes of the
environment felt by the Ser221-D. Table 2 also shows the emis-
sion maxima of free dansyl hydroxide (D-OH), dansyl fluoride (DF)
and dansyl methoxide (D-OCH3) in buffer, 1,4-dioxane and ACN for
comparison purposes. A possible explanation is that these different
populations (which are responsible for these two emission bands)
arise from local conformational changes due to the penetration of
solvent molecules into the active site, forcing the dansyl group to
reorient itself towards the solvent. It is also possible that water
from the hydration shell quenches the fluorescence towards the
beginning of the incubation period and the gradual displacement
of this water by the organic solvent results in an increase in fluo-
rescence [30]. Quantitative estimation of the active-site polarity on
the basis of emission maxima of active site bound fluorophores was
proposed by Vaz and Schoellmann using the “empirical polarity”
scales established by Kosower [16,19]. They reported that, for the
evaluation of microenvironment polarity using fluorescent probes,
Fig. 3. Fluorescence emission spectra of dansyl-PEG-subtilisin Carlsberg in (A) 1,4-
dioxane and (B) in acetonitrile. Day 0; day 1; day 2; day 3; day 4 (excitation at
360 nm).
finding can account for the fact the PEG-enzyme shows a higher
activity in dioxane than in most other organic solvents [9,10,15].
D-labeled PEGylated and native SC, on being excited at 360 nm in
buffer showed a clear peak of D emission at 510 nm (Table 1). Also,
no shift in emission maximum occurred when the labeled enzyme
was stored in buffer for 5 days. However, a distinctly different and
complex behavior was seen during incubation in the organic sol-
vents (Table 1). The Ser221-D emission spectrum in dioxane on
day 0 showed an almost well defined peak centered at 510 nm
as a result of excitation at 360 nm (Fig. 3A). After nearly 24 h, a
shoulder started to appear in the emission spectrum at 450 nm.
The shoulder at 450 nm became increasingly pronounced during
subsequent days of incubation and by day 4 the emission spec-
trum showed a broad peak spanning from 450 to 500 nm (Fig. 3,
Table 1). The quantum yield of Ser221-D also increased during the
incubation in dioxane as is noticed from the increase in D fluo-
rescence intensity during this period. These results would indicate
a decrease in the polarity of the environment around the fluo-
rophore during incubation in dioxane. A similar effect was observed
in ACN, where again the D-labeled PEGylated subtilisin, on being
excited at 360 nm, showed a clear peak at 524 nm on day 0 (Fig. 3B,
Table 3). After 24 h, another peak appeared at 445 nm, and on con-
tinued incubation in ACN, the longer wavelength peak was reduced
to a mere shoulder while the peak at 445 nm became the major
peak. These changes in the emission spectra for Ser221-D in organic
solvents could also be due to a transition between two differ-
ent states or subpopulations, the midpoint being approximately
at 470 nm for both dioxane and ACN. It can be hypothesized that
there is a gradual reduction of the fraction of the first subpopula-
tion (ꢀem = 510/524 nm) and a corresponding increase in the size
of the second subpopulation (ꢀem = 450/445 nm) during incubation
in dioxane and ACN. It has been previously suggested, for a sim-
ilar system (SC-Ser221-D) in THF, that the second emerging peak
resulted from free dansyl in the solution due to its leaching from
Fig. 4. Absorption spectra of dansyl-PEG-subtilisin C. during incubation in 1,4-
dioxane. Day 0 (green); day 1 (blue); day 2 (red); day 3 (brown); day 4 (black).

V. Bansal et al. / Journal of Molecular Catalysis B: Enzymatic 64 (2010) 38–44
43
3.7. Effect of incubation of Vmax and KM
In addition, both Vmax (min) and KM (mM) decrease dur-
ing 4-day incubation in these two organic solvents. In ACN at
day 0, Vmax = 5 0.6, and KM = 322 87. After 4 days of incuba-
tion, Vmax = 0.63 0.09, and KM = 126 28. In 1,4-dioxane at day 0,
Vmax = 15 3, and KM = 440 120. After 4 days of incubation in this
last solvent, Vmax = 1.7 0.4, and KM = 192 65. This clearly indi-
cates that changes are occurring at the enzyme’s active site during
this period. The drastic decrease in Vmax suggests that the efficiency
of the enzyme is jeopardized, and although a decrease in KM could
be due several reasons such as to minor and local (at the active site)
structural changes, or to a tighter binding of the substrate or poorer
stabilization of the tetrahedral intermediate, it clearly shows that
the catalytic machinery is been affected. As it will be discussed in
the following section, results obtained here and recently reported
will support two possible mechanisms.
Fig. 5. Emission spectra of dansyl-labeled subtilisin Carlsberg during incubation
in 1,4-dioxane under controlled water activity. Day 0; day 1; day 2; day 3; day 4.
Excitation at 360 nm.
4. Discussion
the use of the “empirical polarity” scales is more appropriate than
the use of polarity scales based upon dielectric constant. We have
deduced the ‘empirical polarities’ for the microenvironments of the
dansyl group in the active site of SC in different solvents at different
times of incubation on the basis of the Z-values obtained by Vaz
and Schoellmann using the model compound dansyl-ethyl ester
as a reference [16]. The results obtained are listed in Table 3. The
Z-values reflect upon the microenvironment experienced by the
fluorophore and such low Z-values as seen here can be accounted
for by the presence of non-polar side chains of some amino acid
residues in the active site as suggested by Vaz and Schoellmann
[16]. Initially, for at least one subpopulation of the Ser221-D fluo-
rophore the active site environment in ACN seems to be more polar
than in buffer, while in dioxane it is similar to that in buffer on day 0
and becomes less polar during the subsequent incubation (Table 3).
These results clearly demonstrate a decrease in active-site polarity
during incubation in organic solvents. After initial exposure of the
enzyme to organic solvents, the polarity of the active site is similar
to that in water, but during incubation it becomes similar to the
polarity of the respective organic solvent, which can be explained
by penetration of organic solvent into the enzyme active site.
When PEG-subtilisin C. was initially exposed to 1,4-dioxane,
the emission maximum of the dansyl group bound to its active
site was similar to that in buffer (510 nm), whereas the emis-
sion maximum in ACN shifted 14 nm to a higher wave number
(to 524 nm). However, during prolonged exposure to these two
organic solvents a new, shorter wavelength peak, indicative of a
drastic decrease in the polarity felt by the fluorophore begins to
emerge. Both Vmax and KM decreased during prolonged exposure to
these two solvents, also indicative of major changes taking place in
the active site, which could be a diminished ability to stabilize the
transition states, or minor structural distortions that might force
substrates to adopt a less catalytically active conformation. Previ-
ous H/D exchange results obtained from this laboratory show that
the enzyme becomes more rigid during prolonged exposure (96 h)
to these two solvents [11], and a theoretical study obtained after
92 ns in ACN suggest that SC exchange its internal water molecules
by the organic solvent, while a critical residue in the oxyanion hole
changes its orientation [12]. The implications of theses findings is
that while the enzyme becomes more rigid by the removal of its
internal water molecules over a 96-h period, the active-site polar-
ity should decrease (as suggested by our fluorescence data) and
disruption of the oxyanion hole might reduce the enzyme’s abil-
ity to stabilize the tetrahedral intermediates, decreasing both Vmax
and KM, as shown by the results here presented. Furthermore, a
decrease in the polarity of the active site and disruption of the
oxyanion hole could lead to reorientation of the bound substrate
[12] or allow the substrate to bind in a less restricted fashion, let-
ting it move more freely, as it has been shown on previous studies
involving EPR spectroscopy [9,31]. In short, one interpretation that
is emerging from these studies is that changes in active-site polar-
ity during prolonged exposure to organic solvents are the major
cause of the observed decrease in enzymatic activity.
3.6. Water activity control experiments
To rule out the effect of water these fluorescence studies
were performed under controlled water environment by adding a
hydrated salt mixture of sodium acetate to the sample. The spectra
obtained in 1,4-dioxane are shown in Fig. 5. Unlike what was pre-
viously observed, the Ser221-D emission peak obtained after initial
exposure to this solvent was centered at 445 nm with a shoulder at
510 nm (without controlling the water activity we observed a peak
centered at 510 nm and a shoulder at 445 nm which increased dur-
ing incubation in this solvent, Fig. 3). During the incubation period
the peak at 445 nm showed an increase in intensity as compared
to the shoulder (at 510 nm). Results obtained from a similar water
activity control experiment in ACN were analogous to those in the
case of 1,4-dioxane (spectra not shown). It seems that the effect
of incubation seen in samples with water activity controlled was
accelerated. This in turn suggests that when the system is not pre-
equilibrated with respect to water the effect seen during incubation
could actually be the consequence of the equilibrium that enzyme
and solvent molecules are trying to reach, while in case of pre-
equilibration (with saturated salt solution), the emission spectra
at time zero reflect a system already at equilibrium. However, it is
important to point out that a pre-equilibrated system also shows
the blue shift in Ser221-D emission during incubation in both sol-
vents.
It is important to mention that efforts have been made by other
research groups to explain the solvent dependence of enzyme activ-
ity using similar techniques and systems than us. However, most
studies were not geared to address the effect of prolonged expo-
sure to organic solvents on an enzyme’s activity, and therefore they
concentrate on the initial effect of exposure to organic solvents.
Nevertheless, it is important to discuss some of these results since
the effects that are linked to enzyme activity are enzyme flexibility
and active-site polarity, both related to our work. Eppler et al. 19
F
NMR active-site polarity studies show that increasing the salt con-
centration (NaCl, NaF, KCl and KF) in hexane from 0 to 98% did not
augment enzyme polarity (while enzyme activity was increased),
concluding that changes in polarity do not affect enzyme activity
[31]. On a similar and more recent study however, it was concluded

44
V. Bansal et al. / Journal of Molecular Catalysis B: Enzymatic 64 (2010) 38–44
that active-site polarity and hydration do affect enzyme activity,
but that this effect was also dependent on the solvent used [32].
Actually, our results show that initially (when first introduced to
an organic solvent) the enzyme retains the polarity of the aqueous
solution from which it was last dissolved in, and changes in enzyme
polarity are a slow process which might involve penetration of the
solvent onto the enzyme fold. In a system whose water activity is
controlled, besides the equal hydration of enzyme and solvent, the
2 h incubation period (required for the experiment) would allow
for some solvent molecules to enter the enzyme fold, and therefore
change the active-site polarity as we observed.
We propose that prolonged exposure to organic solvents leads
to a decrease in enzyme dynamics and active-site polarity, and a
reorientation of the active site inhibitors. These affect the ionization
state of the catalytic triad residues as well as the transition states
and intermediates’ stability, resulting in reduced enzyme activity,
Research Resources, and the National Institutes of Health (SCORE).
The content is solely the responsibility of the authors and does not
necessarily represent the official view of the National Center for
Research Resources or the National Institutes of Health. In addition,
the authors would like to thank Dr. Carmelo Garcia and Dr. Roland
Oyola from the University of Puerto Rico at Humacao for their assis-
tance in the use of and for providing us access to the Fluorescence
Spectrophotometer.
References
[1] J.F. Amorim Fernandez, P. Halling, Biotechnol. Prog. 18 (2002) 1455–1457.
[2] C.O. Fagain, Enzyme Microbial Technol. 33 (2003) 137–149.
[3] E.A. Susimar Gonzalez Martinez, L. Vergara Cordero, A. Ferrer, I. Montanez-
Clemente, G. Barletta, Biotechnol. Prog. 18 (2002) 1462–1466.
[4] K. Griebenow, A.M. Klibanov, Proc. Natl. Acad. Sci. U.S.A. 92 (1995)
10969–10976.
[5] K. Griebenow, A.M. Klibanov, Biotechnol. Bioeng. 53 (1997) 351–362.
[6] K. Griebenow, A.M. Klibanov, J. Am. Chem. Soc. 118 (1996) 1195–11700.
[7] Y. Zheng, R.L. Ornstein, J. Am. Chem. Soc. 118 (1996) 4175–4180.
[8] A. Dong, J.D. Meyer, B.S. Kendrick, M.C. Manning, J.F. Carpenter, Arch. Biochem.
Biophys. 334 (1996) 406–414.
Vmax and KM
.
5. Conclusion
[9] V.B. Betzaida Castillo, A. Ganesan, P. Peter Halling, F. Secundo, A. Ferrer, K.
Griebenow, G. Barletta, BMC Biotechnol. 51 (2006).
[10] B. Castillo, Y. Pacheco, W. Al-Azzam, K. Griebenow, M. Devi, A. Ferrer, G. Barletta,
J. Mol. Catal. B: Enzym. 35 (2005) 147–153.
[11] E. Fasoli, A. Ferrer, G.L. Barletta, Biotechnol. Bioeng. 102 (2009) 1025–1032.
[12] A. Cruz, E. Ramirez, A. Santana, G. Barletta, G. Lopez, Mol. Simulat. (2009)
1–8.
[13] F.C.G. Secundo, G. Vecchio, F. Zambianchi, Biotechnol. Bioeng. 64 (1999)
624–629.
[14] R. Bovara, G. Carrea, A.M. Gioacchini, S. Riva, F. Secundo, Biotechnol. Bioeng. 54
(1997) 50–57.
[15] R.J.S. Betzaida Castillo, A. Ferrer, G. Barletta, K. Griebenow, Biotechnol. Bioeng.
99 (2008) 9–17.
[16] W.L. Vaz, G. Schoellmann, Biochim. Biophys. Acta 439 (1976) 194–205.
[17] W.L. Vaz, G. Schoellmann, Biochim. Biophys. Acta 439 (1976) 206–219.
[18] D.C.B.L. Turner, Biochemistry 10 (1968) 10.
Previous studies have shown that an enzyme’s initial activ-
ity in organic solvents decreases after prolonged exposure to this
medium. It was also reported that this effect was unrelated to
structural perturbations of the enzymes under study, since their
structures remained “unharmed”. Considering those studies and
the fluorescence spectroscopic results presented here, we suggest a
mechanism that involves two possible orientations/conformations
of the active site bound fluorophore after prolonged exposure of
PEG-SC to ACN and 1,4-dioxane. The dominant conformation of
the inhibitor (or substrate), upon initial exposure to organic sol-
vents is the one that interacts to a greater extent with the active
side residues. During incubation there is a gradual transition of the
active site bound fluorophore to a more flexible and mobile confor-
mation that has less interaction with the active site environment.
These changes seem to be brought about by water stripping effect
and the movement of solvent molecules into the enzyme fold that
brings about polarity changes in the active site. In the case of a
bound substrate in the enzyme active site in an organic solvent, we
believe that the first substrate conformation and active-site polar-
ity (similar to that in aqueous solvent) favors catalysis more than
the later conformation that results after prolonged exposure to the
organic solvent. However, additional studies are needed to rule out
other possible mechanisms and to unequivocally ascertain the con-
tribution of enzyme dynamics, active-site polarity and substrate
reorientation to the phenomenon of enzyme partial inactivation
upon prolonged exposure to organic solvents.
[19] E.M. Kosower, JACS 80 (1958) 3253–3260.
[20] K. Griebenow, Y. Laureano, A.M. Santos, I. Montanez-Clemente, L. Rodriguez,
M.W. Vidal, G. Barletta, J. Am. Chem. Soc. 121 (1999) 8157–8163.
[21] A. Fersht, Enzyme Structure and Mechanism, 2nd ed., Freeman and Company,
New York, 1985.
[22] A.F.S.A. Habeeb, H.G. Cassidy, S.J. Singer, Biochim. Biophys. Acta 29 (1958)
587–593.
[23] P.K.K.R. Smith, G.T. Hermanson, A.K. Mallia, F.H. Gartner, M.D. Provenzano,
E.K. Fujimoto, N.M. Goeke, B.J. Olson, D.C. Klenk, Anal. Biochem. 150 (1985)
76–85.
[24] E.N. Vulfson, P.J. Halling, H.L. Holland, Enzymes in Nonaqueous Solvents, Meth-
ods and Protocols Humana Press, Totowa, New Jersey, 2001.
[25] L. Stryer, J. Mol. Biol. 13 (1965) 482–495.
[26] K.J.S.A. Willis, Biochemistry 28 (1989) 4902–4908.
[27] M.R. Eftink, in: C.H. Suelter (Ed.), Methods of Biochemical Analysis, John Wiley
& Sons, Inc., 1991, pp. 127–205.
[28] J. Broos, A.J.W.G. Visser, J.F.J. Engbersen, J. Am. Chem. Soc. 117 (1995)
12657–12663.
[29] J.R. Lakowicz, Principles of Fluorescence Spectroscopy, 3rd ed., Springer, 2006.
[30] N.C.B.R.N. Genov, Biochem. J. 238 (1986) 923–926.
[31] R.K. Eppler, E.P. Hudson, S.D. Chase, J.S. Dordick, J.A. Reimer, D.S. Clark, Proc.
Natl. Acad. Sci. U.S.A. 105 (2008) 15672–15677.
Acknowledgements
[32] E.P. Hudson, R.K. Eppler, J.M. Beaudoin, J.S. Dordick, J.A. Reimer, D.S. Clark, J.
Am. Chem. Soc. 131 (2009) 4294–4300.
The project described was supported by Grants P20 RR016470,
S06 GM-08216, and GM-08102 from the National Center for