Zaks & Klibanov 1988

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’ h e JOURNAL OF BIOLOGICAL CHEMISTRY 
(0 1988 by Tbe American Society for Biochemistry and Molecular Biology, Inc. 
Enzymatic Catalysis in Nonaqueous Solvents* 
Vol. 263, No. 7. Issue of March 5, pp. 3194-3201, 1988 
Printed in U. S. A. 
(Received for publication, October 26, 1987) 
Aleksey Zaks and Alexander M. Klibanov 
From the Department of Applied Biological Sciences, Massachusetts Institute of Technology, Cambridge, Massachusetts 02139 
Subtilisin and a-chymotrypsin vigorously act as ca- 
talysts in a variety of dry organic solvents. Enzymatic 
transesterifications in organic solvents follow Michae- 
lis-Menten kinetics, and the values of V/Km roughly 
correlate with solvent’s hydrophobicity. The amount 
of water required by chymotrypsin and subtilisin for 
catalysis in organic solvents is much less than needed 
to form a monolayer on its surface. The vastly different 
catalytic activities of chymotrypsin in various organic 
solvents are partly due to stripping of the essential 
water from the enzyme by more hydrophilic solvents 
and partly due to the solvent directly affecting the 
enzymatic process. The rate enhancements afforded by 
chymotrypsin and subtilisin in the transesterification 
reaction in octane are of the order of 100 billion-fold; 
covalent modification of the active center of the en- 
zymes by a site-specific reagent renders them catalyt- 
ically inactive in organic solvents. Upon replacement 
of water with octane as the reaction medium, the spec- 
ificity of chymotrypsin toward competitive inhibitors 
reverses. Both thermal and storage stabilities of chy- 
motrypsin are greatly enhanced in nonaqueous sol- 
vents compared to water. The phenomenon of enzy- 
matic catalysis in organic solvents appears to be due to 
the structural rigidity of proteins in organic solvents 
resulting in high kinetic barriers that prevent the na- 
tive-like conformation from unfolding. 
All the vast knowledge accumulated in the area of mecha- 
nistic enzymology (Dixon and Webb, 1979; Walsh, 1979; 
Fersht, 1985) has been derived from studies of enzymes in 
aqueous solutions. It is easy to imagine how our understanding 
of enzymatic catalysis could be enhanced if a new fundamental 
variable was introduced in the experimentation, namely the 
solvent (i.e. the reaction medium). 
Although conducting enzymatic processes in nonaqueous 
media might seem to go against the conventional wisdom, at 
least two alternative approaches to that goal have been re- 
cently successfully developed (Waks, 1986). In the first ap- 
proach (Luisi, 1985; Martinek et al., 1986), enzymes are dis- 
solved in micropools of water which are emulsified in water- 
immiscible solvents; the microemulsion is stabilized by sur- 
factants that form “reverse micelles.’’ In the second approach 
(Klibanov, 1986), powdered enzymes are directly suspended 
in organic solvents. 
In reverse micelles, the enzyme is confined to a water pool 
which, in turn, is insulated from the organic solvent by a 
monolayer of surfactant. Therefore, inherent catalytic prop- 
erties of enzymes in reverse micelles are generally similar to 
* This research was supported by Grant CBT-8710106 from the 
National Science Foundation. The costs of publication of this article 
were defrayed in part by the payment of page charges. This article 
must therefore be hereby marked “advertisement” in accordance with 
18 U.S.C. Section 1734 solely to indicate this fact. 
those in aqueous solutions (Luisi, 1985; Martinek et al., 1986; 
Waks, 1986). In contrast, solid enzymes dispersed in organic 
solvents are directly exposed to the solvent and hence exhibit 
some remarkable novel properties compared to those in water 
(Klibanov, 1986), e.g. greatly increased thermal stability (Zaks 
and Klibanov, 1984; Wheeler and Croteau, 1986; Ayala et al., 
1986) and strikingly different substrate specificity (Zaks and 
Klibanov, 1984; Zaks and Klibanov, 1986). Also, many trans- 
formations that are impossible in aqueous solutions due to 
kinetic or thermodynamic reasons, can be readily catalyzed 
by enzymes in organic solvents (Klibanov, 1986; Zaks and 
Klibanov, 1985). Consequently, a number of interesting and 
useful enzymatic conversions in organic solvents have been 
accomplished including lipase-catalyzed regioselective acyla- 
tion of glycols (Cesti et al., 1985) and sugars (Therisod and 
Klibanov, 1986,1987) and interesterification of fats Yokozeki 
et aL., 1982; Macrae, 1983), lipase-catalyzed stereoselective 
transesterifications and esterifications (Kirchner et al., 1985; 
Langrand et al., 1985), polyphenol oxidase-catalyzed regios- 
pecific oxidation of phenols (Kazandjian and Klibanov, 1985), 
alcohol dehydrogenase-catalyzed stereoselective oxidoreduc- 
tions (Grunwald et al., 1986), and peroxidase-catalyzed oxi- 
dations used in biosensors (Kazandjian et al., 1986; Boeriu et 
al., 1986). 
In order to take full advantage of the novel opportunities 
afforded by nonaqueous enzymology, it is imperative to un- 
derstand such basic features and characteristics of this phe- 
nomenon as the dependence of enzymatic properties on the 
nature of the solvent, the amount of water required for catal- 
ysis, catalytic parameters and conformational stability of 
enzymes in organic solvents, etc. Elucidation of these issues, 
in addition to its biotechnological importance, should also 
provide profound insights into such questions as protein fold- 
ing and dynamics, the role of water in enzyme catalysis and 
stability, and protein intramolecular interactions and mobility 
(Levinthal, 1986). 
The present paper describes a detailed investigation ad- 
dressing the aforementioned questions. Bovine pancreatic a- 
chymotrypsin and Bacillus subtilis protease (subtilisin Carls- 
berg) were employed as model enzymes in organic solvents. 
These two proteases are not associated with biological mem- 
branes in nature, and their physiological role is to hydrolyze 
water-soluble proteins; hence they represent enzymes whose 
natura1 environment and function involve aqueous solutions 
(in contrast to membrane-bound or lipolytic enzymes that act 
on interfaces and thus are accustomed to a nonaqueous mi- 
lieu). The enzymatic mechanisms and properties of chymo- 
trypsin and subtilisin in water are well understood (Black- 
burn, 1976; Walsh, 1979; Fersht, 1985). In this work, we 
mechanistically examined the catalytic behavior of these en- 
zymes when they were directly dispersed as solids in non- 
aqueous solvents. 
EXPERIMENTAL PROCEDURES 
Materials-Crystalline bovine pancreatic a-chymotrypsin (EC 
3.4.21.1) (Type 11) and protease from Bacillus subtilis (subtilisin 
3194 
Enzymatic Catalysis in Organic Solvents 3195 
Carlsberg) were purchased from Sigma as lyophilized powders with 
specific activities of 49 and 10.5 unita/mg of solids, respectively. The 
concentration of the active centers in chymotrypsin (Schonbaum et 
al., 1961) and in subtilisin (Polgar and Bender, 1967), determined by 
spectrophotometric titration with N-trans-cinnamoylimidazole, was 
found to be 100 and 54%, respectively. 
In experiments involving immobilized chymotrypsin, the enzyme 
was covalently attached to CNBr-activated Sepharose 4B following 
the procedure of h e n and Ernback (1971). This method afforded 
approximately 200 mg of chymotrypsin attached to 1 g of support. 
All chemicals used in this work were obtained commercially and 
were of analytical grade. All solvents were of the highest purity 
commercially available and were used without further purification 
unless otherwise indicated. When needed, the solvents were driedby 
gentle shaking with 3A molecular sieves (Linde). 
Assays-The water concentration both in organic solvents and in 
enzymes was measured by the optimized titrimetric Fischer method 
(Laitinen and Harris, 1975). The sensitivity limit of this method in 
determining the water content in organic solvents under our condi- 
tions was approximately 0.02% (v/v). The term “dry organic solvent” 
is used in this work when no water was detected by the Fischer 
method. 
All enzymatic transesterification reactions in organic solvents were 
measured by gas chromatography following accumulation of the new 
ester. A 5-meter HP series 530-pm column coated with methyl silicone 
gum (Hewlett-Packard) (Nz carrier gas, 5 ml/min, detector and injec- 
tor port temperature 250 “C) was used. 
Kinetic Measurements-Protease-catalyzed transesterification re- 
actions in organic solvents were measured as follows. A suspension 
of the enzyme (typically 1 mg) in 1 ml of a given organic solvent 
containing the substrates (2-12 mM ester and 0.25-1.5 M alcohol) was 
placed in a 2-ml screw-cap vial. The vial was shaken on an orbit 
shaker at 250 rpm and 20 “C. Periodically, 0.5-pl aliquots were with- 
drawn and assayed by gas chromatography as described above. 
Hydrolysis of N-acetyl-L-phenylalanine ethyl ester catalyzed by 
chymotrypsin or subtilisin in water was measured potentiometrically 
using a Radiometer recording pH-stat system. In a typical experi- 
ment, 10 d of an aqueous solution of the ester (0.3-5.0 mM) contain- 
ing 0.1 M KC1 was placed in a thermostated cuvette of a pH-stat, 
equilibrated at 20 ‘C, and adjusted to pH 7.8. Then 25 r g of the 
protease was added, and the acid liberated as a resuit of enzymatic 
hydrolysis was automatically titrated with 50 mM NaOH. 
Thermal Inactivation-The time course of irreversible thermal 
inactivation of chymotrypsin in organic solvents was determined as 
follows. Powdered enzyme (1 mg) was dispersed in 1 ml of the solvent, 
and the suspension was placed in a 2-ml sealed ampule. The ampule 
was then immersed in a 100 ‘C bath. After a certain period of time, 
the ampule was removed from the bath, cooled, and opened. The 
enzyme was recovered by centrifugation, dried under vacuum, and 
assayed in the hydrolysis reaction in water as described above. 
RESULTS AND DISCUSSION 
Subtilisin-catalyzed Transesterificatwns in Organic Sol- 
uents-The natural reaction of subtilisin (Ottesen and Svend- 
sen, 1970) and chymotrypsin (Bender and Kezdy, 1965; Hess, 
1970; Blow, 1976) is the hydrolysis of peptide bonds within 
proteins; both enzymes also specifically hydrolyze low molec- 
ular weight substrates such as esters of N-acyl-L-amino acids. 
Since enzymatic hydrolysis involves water as a substrate, it is 
not an ideal process for nonaqueous enzymology. Both pro- 
teases can also utilize nucleophiles other than water, in par- 
ticular aliphatic alcohols, thus replacing hydrolysis with 
transesterification (Fersht, 1985). Since the latter does not 
require water, we selected enzymatic transesterification as a 
model process to be studied in organic solvents. 
In an initial experiment, 1 mg of commercial crystalline 
subtilisin was placed in 1 ml of dry octane containing 5 mM 
N-acetyl-L-phenylalanine ethyl ester and 1 M propanol. The 
suspension (enzymes are insoluble in nearly all organic sol- 
vents (Singer, 1962)) was shaken at 20 “C; periodically, ali- 
quots were withdrawn and analyzed by gas chromatography. 
This analysis revealed formation of N-acetyl-L-phenylalanine 
propyl ester (the product of the transesterification reaction) 
which increased with time. The initial rate of this process was 
determined to be 0.13 pM/min (no reaction was detected in 
the absence of the enzyme). Hence subtilisin is catalytically 
active in dry octane. 
We previously found that enzymatic activity of porcine 
pancreatic lipase in organic solvents was greatly increased 
when the enzyme was recovered from an aqueous solution of 
the pH optimal for the lipase activity (Zaks and Klibanov, 
1985). This effect, which was later confirmed for terpene 
cyclase (Wheeler and Croteau, 19861, is due to the fact that 
ionogenic groups of the enzyme acquire a certain ionization 
state in the aqueous solution of a given pH. This ionization 
state (and the enzymatic activity corresponding to it) is re- 
tained in the solid state and in organic solvents (Klibanov, 
1986). Following this rationale, subtilisin was dissolved in an 
aqueous buffer (20 mM phosphate), pH 7.8 (optimum of the 
subtilisin activity in water (Ottesen and Svendsen, 1970)), 
and then lyophilized. The rate of the transesterification re- 
action in octane catalyzed by 1 mg/ml of the “pH-adjusted 
sample of subtilisin was 9.8 pM/min, i.e. 75 times greater than 
that of the enzyme “straight from the Sigma bottle.” The pH- 
adjusted subtilisin was used in all subsequent experiments. 
We found that subtilisin irreversibly inactivated by the 
active center-directed inhibitor phenylmethanesulfonyl fluo- 
ride (Fahrney and Gold, 1963) was completely inactive in 
octane, thus indicating the nonartifactual origin of the trans- 
esterification reaction. The rate of the enzymatic transester- 
ification in octane was directly proportional to the concentra- 
tion of subtilisin. 
On the basis of the classical mechanism of serine protease 
catalysis in water (Bender and Kezdy, 1965; Blackburn, 1976), 
the kinetic scheme of the subtilisin-catalyzed transesterifica- 
tion is likely to involve formation of a noncovalent enzyme- 
ester complex, which then transforms to an acyl-subtilisin 
intermediate with the concomitant release of alcohol product. 
The acyl-subtilisin then interacts with the nucleophile (pro- 
panol in our case) to form another binary complex, which 
then yields the new ester and the free enzyme. This mecha- 
nism (compulsory order without ternary complexes) results 
in a set of parallel lines when the reciprocal initial rates are 
plotted against the reciprocal ester concentrations at fixed 
concentrations of the alcohol (Cornish-Bowden, 1979). When 
the maximal velocities determined in these coordinates are 
plotted against the reciprocal concentrations of the alcohol 
substrate, again a straight line should be obtained (Cornish- 
Bowden, 1979). 
Kinetics of the reaction between N-acetyl-L-phenylalanine 
ethyl ester and propanol catalyzed by subtilisin in octane were 
analyzed as described above. The kinetic behavior observed 
was found to be in good agreement with that expected for 
Michaelis-Menten kinetics. The slope of the parallel lines in 
the double reciprocal coordinates afforded the true ratio of 
the maximal velocity (V) to the Michaelis constant for the 
ester (Kmcester)) which was determined to be 2.0. min“. 
Similar kinetic analysis was carried out for the subtilisin- 
catalyzed transesterification in 14 other organic solvents (Ta- 
ble I). In all of them, the enzymatic process obeyed Michaelis- 
Menten kinetics. The V/Kmceste,) values presented in Table I 
exhibit a strong dependence on the nature of the solvent and 
roughly correlate with the hydrophobicity of the latter; the 
best solvents are water-immiscible hydrophobic solvents, and 
the worst are highly hydrophilic water-miscible ones. 
Kinetics of Chymotrypsin Catalysis in Octane-To establish 
the generality of the effects observed for subtilisin, another 
enzyme, bovine pancreatic a-chymotrypsin, was examined as 
a catalyst in organic solvents. Although the behavior of chy- 
3196 Enzymatic Catalysis 
TABLE I 
Kinetic parameters of the reaction between N-acetyl-L-phenylalanine 
ethyl ester andpropanol catalyzed by subtilisin and chymotrypsin in 
organic solvents 
Both subtilisin and chymotrypsin (5 mg/ml) were pH-adjusted by 
lyophilization from 20 mM aqueous phosphate buffer, pH 7.8 (in the 
case of chymotrypsin the aqueous solution contained 0.25% of the 
ligand N-acetyl-L-phenylalanine instead of phosphate). Following 
lyophilization, subtilisin and chymotrypsin contained 2.1 and 2.5% 
(w/w) water, respectively (for chymotrypsin, at smaller water con- 
tents lower enzymatic activities in organic solvents were observed). 
All organic solvents contained less than 0.02% (v/v) water (the 
sensitivity limit of our detection method). Conditions: 1 mg/ml en- 
zyme, 2-12 mM N-acetyl-L-phenylalanine ethyl ester, 1 M n-propanol, 
the suspension was shaken at 20 “C. No reaction was detected in the 
absence of the enzyme regardless of solvent. Lyophilization did not 
appreciably affect the catalytic activity of chymotrypsin and subtilisin 
in water, thus excluding any irreversible enzyme inactivation during 
that Drocess. 
Solvent” ~” 
V/Km(e.br, 
Subtilisin Chymotrypsin 
rnin” X 10s 
Hexadecane 3900 4300 
Octane 2000 1700 
Carbon tetrachloride 340 96 
Butyl ether 240 48 
Toluene 150 120 
tert-Amyl alcohol 2100 38 
Ethyl ether 97 48 
2-Pentanone 59 12 
Pyridine 97 <o. 1 
Tetrahydrofuran 120 7.2 
Acetone 810 0.6 
Acetonitrile 150 0.4 
Dioxane 9.2 0.2 
Dimethylformamide 19 <o. 1 
Dimethyl sulfoxide <o. 1 (0.1 
The solvents are listed in the order of decreasing hydrophobicity 
(increasing hydrophilicity) which is reflected by the logarithm of the 
partition coefficient for a given solvent between octanol and water 
(log P ) . The values of log P for the solvents listed in the table were 
determined by Laane et al. (1987). 
motrypsin in aqueous solutions of water-miscible organic 
solvents has been kinetically studied (Clement and Bender, 
1963 and references therein), water was always the main 
component of the solvent. Dastoli et al. (1966) reported chy- 
motrypsin-catalyzed hydrolysis in methylene chloride con- 
taining 0.25% water, but no quantitative details were pro- 
vided. In contrast, we have undertaken a mechanistic inves- 
tigation of chymotrypsin catalysis in various organic solvents. 
We found that following pH adjustment (lyophilization 
from 20 mM aqueous phosphate buffer, pH 7.8), chymotrypsin 
was catalytically active in dry octane. The enzymatic trans- 
esterification reaction between N-acetyl-L-phenylalanine 
ethyl ester and propanol fits Michaelis-Menten kinetics, as 
revealed by the same kinetic analysis as described above for 
subtilisin. The value of V/K,,,(ester) found was 4.8. min”, 
i.e. about 2% of that for subtilisin under the same conditions. 
However, this value increased 35-fold when chymotrypsin was 
lyophilized from an aqueous solution (pH 7.8) which con- 
tained the ligand N-acetyl-L-phenylalanine instead of phos- 
phate (both compounds were used in the same weight concen- 
tration, 0.25% (w/w), to assure the same consistency of the 
protein sample after lyophilization (Karel and Flink, 1979)). 
Enzymes in lyophilized samples do not necessarily have the 
same conformation as in aqueous solution (Baker et al., 1983). 
Hence, the activating effect of N-acetyl-L-phenylalanine may 
be explained by its binding to chymotrypsin prior to freezing, 
thereby keeping it in a catalytically active conformation. This 
in Organic Solvents 
hypothesis suggests that other ligands should have a similar 
effect. To test that, we replaced N-acetyl-L-phenylalanine 
with two other known ligands (inhibitors) of chymotrypsin:N- 
acetyl-D-phenylalanine and hydrocinnamic acid (0.25%). The 
resultant samples of chymotrypsin had V/Kmcester, in the trans- 
esterification reaction in octane of 2.6.10-3 and 1.5. 
min”, respectively, ie. indeed comparable to the enzyme 
“tuned in” with N-acetyl-L-phenylalanine (1.7. min-’) 
and much greater than the enzyme lyophilized from an 
aqueous phosphate buffer. The latter sample, however, can be 
activated to about the same level as that afforded by the 
ligands if 0.1% water is added to the reaction mixture in 
octane (the presence of 1 M propanol makes that amount of 
water soluble in octane). This is consistent with the view 
(Poole and Finney, 1983) that water “loosens up” the enzyme 
molecule, and hence it becomes sufficiently flexible to be 
induced into the active conformation by interaction with the 
substrate N-acetyl-L-phenylalanine ethyl ester. In the ab- 
sence of added water, the enzyme molecule is apparently so 
rigid (Rupley et al., 1983) that it cannot acquire the active 
conformation upon a reaction with the substrate, unless it 
has been previously tuned in (i.e. activated) by a ligand. 
Therefore, in order to maximize the chymotrypsin activity in 
organic solvents in subsequent experiments, either N-acetyl- 
L-phenylalanine was present during lyophilization or a certain 
amount of water (typically, 0.1%) was added to the reaction 
mixture. 
Phosphate was also replaced with the ligands in the prep- 
aration of subtilisin samples, but this treatment only doubled 
their enzymatic activity (as opposed to a 30-50-fold activation 
for chymotrypsin). Thus, subtilisin does not require to be 
predisposed to the catalytically active conformation by the 
ligand nearly as much as chymotrypsin. This fact is consistent 
with a known greater conformational stability of subtilisin 
compared to chymotrypsin (Ottesen et al., 1970). Presumably, 
this inherent conformational stability of subtilisin prevents 
conformational changes upon lyophilization which would be 
detrimental to its activity. 
Chymotrypsin preinactivated by phenylmethanesulfonyl 
fluoride (Fahrney and Gold, 1963) was completely inactive in 
octane, thereby proving that the enzyme’s active center is 
indispensible for catalysis in nonaqueous solvents. We also 
demonstrated that the chymotrypsin-catalyzed transesterifi- 
cation was not limited by diffusion of the substrates which is 
frequently the case in heterogeneous catalysis (Satterfield, 
1980). External diffusional limitations were ruled out because 
the rate of the enzymatic reaction was independent of the 
extent of shaking of the suspension (in the range from 160 to 
300 rpm). Internal diffusional limitations were dismissed be- 
cause ultrasonication of a suspension of chymotrypsin in 
octane (resulting in a reduction of an average enzyme particle 
from 270 to 5 bm, as revealed by direct microscopic exami- 
nation) had no appreciable effect on the enzymatic transes- 
terification rate. 
Chymotrypsin in Different Organic Solvents-pH-adjusted 
and ligand-activated chymotrypsin was employed as a catalyst 
of the transesterification reaction in organic solvents listed in 
Table 1. In all instances the enzymatic process followed the 
Michaelis-Menten scheme, and the determined values of V/ 
Kmceste,, are given in the last column of Table I. One can see 
that, similar to subtilisin, the catalytic activity is greater in 
hydrophobic solvents with the highest V/K,,,(e6ter) observed in 
hexadecane, while no activity was detected in the hydrophilic 
solvent dimethyl sulfoxide. However, chymotrypsin is more 
sensitive to the nature of organic solvent than subtilisin which 
seems to be consistent with its lower conformational stability 
Enzymatic Catalysis in Organic Solvents 3197 
(Ottesen et ai., 1970). Correlation of enzymatic activity with 
hydrophobicity of the solvent and distinct sensitivities of 
enzymes to that parameter were also reported for lipases (Zaks 
and Klibanov, 1985). Comparison of the two studies reveals 
no essential differences between lipolytic and nonlipolytic 
enzymes, thus pointing to the generality of enzymatic catal- 
ysis in organic solvents. 
The very different activitiesexhibited by chymotrypsin in 
various organic solvents may be due to the direct effect of the 
solvent on the enzyme or due to stripping of the essential 
water (Klibanov, 1986) from the enzyme by the solvent. To 
distinguish between these two mechanisms, chymotrypsin was 
incubated in some of the solvents listed in Table I for the 
duration of the kinetic measurements, and the amount of 
water remaining on the enzyme was then determined titri- 
metrically. The data obtained are presented in Table 11, and 
they afford a number of important conclusions. 
It is seen that the activity of chymotrypsin in organic 
solvents (Table I) correlates with the amount of water re- 
tained by the enzyme in those solvents (Table 11): the more 
water, the greater the enzymatic activity. It should be pointed 
out that the amount of water on the enzyme after incubation 
in octane is the same as the enzyme had before its contact 
with the organic solvent (2.5%). Hence, octane does not strip 
any water from the enzyme but all other (more hydrophilic) 
solvents do. It is tempting to assume that a lower enzymatic 
activity in other solvents listed in Table I1 compared to octane 
is due to partitioning of the essential water from the enzyme 
into them. If this explanation is correct, then the activity 
could possibly be restored if the water is replenished. To test 
this hypothesis, we added 1.5% (v/v) water to acetone; the 
amount of water on chymotrypsin incubated in such a solvent 
reached nearly the same value as in octane. When the V/ 
Kmceste,) for chymotrypsin was measured in this solvent, it was 
found to be 1.2 min-l, i.e. about 2000 times greater than 
that in dry acetone and more than two-thirds of that in octane 
(Table I). Therefore, it appears that the much greater activity 
of chymotrypsin in octane than in acetone is mainly due to 
the fact that the latter solvent reversibly strips much of the 
essential water from the enzyme molecule, thereby inactivat- 
ing it. However, the direct effect of the solvent on the enzyme 
(e.g. through binding or changing the dielectric constant of 
the reaction medium) is an important factor as well. For 
instance, the amounts of water on chymotrypsin in octane 
and in toluene are almost the same within the experimental 
error (Table 11), and yet the enzyme is 14 times more reactive 
in the former solvent (Table I). 
It is worth mentioning that even 2.5% water in chymotryp- 
TABLE I1 
The amount of water on chymotrypsin following its incubation in 
different organic solvents 
pH-adjusted and activated with N-acetyl-t-phenylalanine (see 
text) samples of chymotrypsin (50 mg) were placed in organic solvents 
(50 ml), and the suspensions were shaken at 20 "C for 1 h. The 
enzyme was then removed by centrifugation, and the amount of bound 
water was determined by the Fischer method. All organic solvents 
contained less than 0.02% (v/v) water (the sensitivity of our detection 
method). Each water content value presented in the table is a result 
of three independent measurements. 
Solvent 
.____ 
Residual water 
content 
% (W/UI) 
Octane 2.5 k 0.10 
Toluene 2.3 -C 0.09 
Tetrahydrofuran 1.6 2 0.11 
Acetone 1.2 k 0.08 
Pyridine 1.0 & 0.08 
sin corresponds to no more than 50 molecules of water/ 
enzyme molecule (assuming that water is uniformly distrib- 
uted among chymotrypsin molecules). This amount of water 
is about 10 times less than needed to form a monolayer on 
the enzyme surface (Rupley et al., 1983). In other organic 
solvents the water content is even lower (Table 11). Thus, the 
enzyme is catalytically active despite being quite literally 
surrounded by organic solvent, in contrast to enzymes in 
reverse micelles which are dissolved in water pools dispersed 
in organic solvent (Luisi, 1985; Martinek et al., 1986). Simi- 
larly, subtilisin recovered from octane contained about 40 
molecules of water/enzyme molecule. 
Enzymatic Properties of Chymotrypsin and Subtilisin in 
Octane-In all the experiments described above, chymotryp- 
sin had been lyophilized from an aqueous solution of pH 7.8. 
It was of interest to examine how the pH adjustment affected 
the enzymatic activity in octane. To that end, we lyophilized 
chymotrypsin from aqueous solutions of different pH and 
studied the dependence of V/Kmceste,) for the enzymatic trans- 
esterification in octane on the pH of the aqueous solution. 
The results obtained are depicted in Fig. 1. One can see that 
the enzymatic reaction in octane very much depends on the 
pH of the last aqueous solution to which the enzyme was 
exposed, with the optimum being at pH 7.8 which coincides 
with the pH optimum for chymotrypsin activity in water 
(Bender and Kezdy, 1965). Importantly, when the different 
samples of chymotrypsin were assayed in the hydrolysis re- 
action in water at pH 7.8, they all had essentially the same 
activity. Thus, the pH memory exhibited by chymotrypsin (as 
well as subtilisin, see above) in octane disappears in water 
where, in contrast to organic solvents, the enzyme's ionogenic 
groups can readily change their ionization states. 
It was important to determine what fraction of chymotryp- 
sin and subtilisin molecules was catalytically active in organic 
solvents. To that end, we developed a method for the titration 
of the enzymes' active centers based on Bender's classical N- 
trans-cinnamoylimidazole technique (Schonbaum et al., 1961; 
Polgar and Bender, 1967). Chymotrypsin (30 mg/ml) was 
dissolved in 0.1 M aqueous acetate buffer, pH 5.0. Then 100 
p1 of acetonitrile containing 1 mg of the spectrophotometric 
titrant N-trans-cinnamoylimidazole was added, and the mix- 
ture was incubated for 2 min at room temperature. This 
resulted in the formation of catalytically inactive trans-cin- 
namoylchymotrypsin (Schonbaum et al., 1961) which was 
subsequently desalted by gel permeation chromatography at 
pH 4 and then lyophilized at pH 5. The acyl enzyme was 
suspended in octane containing 1 M propanol, and the time 
pH 
FIG. 1. The dependence of the enzymatic activity of chy- 
motrypsin in octane on the pH of the aqueous solution from 
which the enzyme was lyophilized. Chymotrypsin (5 mg/ml) was 
lyophilized from aqueous solutions of different pH containing 0.25% 
N-acetyl-L-phenylalanine (to activate the enzyme in octane, see text). 
Samples of the enzyme (1 mg) were then added to 1 ml of octane 
containing the substrates N-acetyl-L-phenylalanine ethyl ester and 
n-propanol, the suspensions were shaken at 20 "C and 250 rpm, and 
the values of V/Km(=-,, were determined as cotangents in the double 
reciprocal coordinates as outlined in the text. 
3198 Enzymatic Catalysis in Organic Solvents 
course of formation of propyl trans-cinnamate was followed 
by gas chromatography. As curve a in Fig. 2A shows, about 
two-thirds of all chymotrypsin molecules have been conse- 
quently deacylated. This was confirmed by redissolving the 
enzyme sample in aqueous acetate buffer and spectrophoto- 
metrically titrating (Schonbaum et al., 1961) those 65% of 
chymotrypsin molecules, 
In an attempt to understand why only about two-thirds of 
chymotrypsin molecules were catalytically active in octane, 
the following experiment was carried out. When the deacyla- 
tion of the enzyme in octane leveled off (after approximately 
5 h, see curve a in Fig. 2A), the enzyme was removed by 
centrifugation, washed with anhydrous ether, redissolved in 
water, immediately lyophilized, and again resuspended in 
octane containing 1 M propanol. This operation resulted in 
some additional gas chromatographic titration of chymotryp- 
sin, as revealed by the formation of more propyl trans-cin- 
namate (curve b in Fig. U), thus bringing the total number 
of active chymotrypsin molecules to 90% ofthat in water. 
The additional titration may be explained by the hypothesis 
that a small portion of the trans-cinnamoyl enzyme molecules 
in suspension in octane was not accessible to propanol due to 
protein-protein contacts. This masking of some active centers 
could not be alleviated by ultrasonication (see above) but 
understandably disappeared upon dissolving the protein in 
water, which exposed some new active centers upon subse- 
quent resuspension in octane. An alternative explanation of 
this titration pattern is that a minor fraction of chymotrypsin 
molecules exists in a reversibly inactive conformation. 
Analogous active center titration experiments were con- 
ducted with subtilisin. As one can see in Fig. 2B, the first 
deacylation again revealed 65% of the active centers, and the 
subsequent one increased that number to 88% of that in 
water. Hence both chymotrypsin and subtilisin display most 
of their active centers in octane. 
Catalytic Efficiency of Enzymes in Octane-The enzyme 
active center concentrations (E] , determined in the preceding 
section aIIow for the calculation of the enzyme specificity 
hours hours 
FIG. 2. Titration of the active centers of chymotrypsin (A) 
and subtilisin (B) in octane. The enzymes were dissolved in water, 
and their active centers were acylated with N-trans-cinnamoylimi- 
dazole as described in the text. Lyophilized powders of the N-trans- 
cinnamoylated chymotrypsin and subtilisin (10 mg in both cases) 
were added to 1 ml of octane containing 1 M propanol (and 0.4% 
water in the case of chymotrypsin to activate it, see text). The 
suspensions were shaken at 20 "C and 250 rpm, and the concentra- 
tions of the propyl tram-cinnamate formed were measured by gas 
chromatography as a function of times (curves a). The concentrations 
of the active centers depicted on the ordinate axes are the molar 
ratios of the concentration of propyl trans-cinnamate released to the 
initial concentration of the tram-cinnamoylated enzymes. When the 
enzymatic deacylation in octane practically stopped, the enzyme 
samples were recovered, thoroughly washed with anhydrous ether, 
redissolved in water, immediately lyophilized, and placed in octane 
solutions containing propanol that were identical to those used the 
first time. Curves b correspond to the production of propyl trans- 
cinnamate in these "secondary" suspensions. 
factor kJKm from V/Km values. Table I11 presents kat/& 
for the transesterification reaction between N-acetyl-L-phen- 
ylalanine ethyl ester and amyl alcohol catalyzed by chymo- 
trypsin and subtilisin in octane. The values of kat/K,,, for the 
enzymic hydrolysis of the ester in water (pH 7.8, 20 "C) were 
also experimentally determined, 4.0. IO4 and 1.3 I lo' M" . s-l, 
respectively. The second order rate constant kCat/Km refers to 
the reaction of the free enzyme with the free substrate (Fersht, 
19851, and hence it is independent of the nature of the 
nucleophile (amyl alcohol or water). Therefore, we compared 
the aforementioned values for ksJK,,, in water and in octane 
and found that with the specific substrate N-acetyl-L-phen- 
ylalanine ethyl ester the enzymes are 104-106 times more 
efficient in water than in the nonaqueous solvent. Presumably 
this is because the conformation and/or flexibility of the 
enzyme active center in octane (and other organic solvents) 
is much less favorable for catalysis than in water. It is inter- 
esting to note, however, that in the reaction with the nonspe- 
cific substrate N-acetyl-L-serine methyl ester chymotrypsin 
is more reactive in octane than in water (Zaks and Klibanov, 
1986). 
We also employed another approach to evaluate the cata- 
lytic efficiency of chymotrypsin and subtilisin in octane. On 
the basis of the rate constant of the nonenzymatic transester- 
ification in octane (k,,o,,,,) (Table III), we calculated the rate 
enhancement (acceleration) effects afforded by chymotrypsin 
and subtilisin. The rate enhancements (the ratio of kcat&,, to 
k,,,,,,e,,z) for the two enzymes, listed in Table 111, are of the 
order of 10'o-lO1l, that is the enzymes exhibit a remarkably 
high catalytic activity in octane (although lower than in 
water). These data suggest that the conformations of the 
enzymes in octane, while not identical to those in water, are 
not radically different from them either, i.e. the enzymes do 
not unfold when placed in octane (and other organic solvents, 
see Table I). 
Inhibition of Chymotrypsin in Octane and in Water-Chy- 
motrypsin is reversibly and competitively inhibited by nu- 
merous aromatic compounds, with the more hydrophobic 
compounds being more potent inhibitors (Wallace et al., 
1963). We have employed several such molecules (shown in 
TABLE I11 
Kinetic parameters of enzymatic and nonenzymatic reactions of 
transesterification (N-Ac-L-Phe-OEt + amyl alcohol -+ N-Ac-L-Phe- 
OAmyl + EtOH) in octane 
Enzyme L, lK," kmmmn: 
M' . 8" M' . s-1 
Acceleration 
effect' 
Chymotrypsin 0.7 1.1. lo-" 6.4.10'' 
Subtilisin 1.8 1.1. lo-" 1.6.10" 
"Initial rates of the enzymatic reactions were measured as de- 
scribed under "Experimental Procedures" at the concentration of the 
enzymes and n-amyl alcohol (which was used instead of propano! to 
avoid evaporation at high temperatures, see Footnote b) of 1 mg/ml 
and 1 M, respectively; the ester concentrations were varied from 2 to 
30 mM. The values of kat/& were calculated by dividing V/K,(-, 
by the concentration of the active enzyme [ E 10 determined from Fig. 
2. Both chymotrypsin and subtilisin were lyophilized from an aqueous 
solution of pH 7.8 prior to use as described under "Experimental 
Procedures," except in the case of subtilisin, 0.25% phosphate was 
used instead of the ligand N-acetyl-L-phenylalanine (see text). Octane 
contained less than 0.02% (v/v) water (the sensitivity limit of our 
assay). 
*At 20 'C, the nonenzymatic reaction was too slow to measure. 
Therefore, transesterification was studied in the temperature range 
from 80 to 110 "C and then extrapolated to 20 "C using the Arrhenius 
dependence. 
e Defined as the ratio of k.,/K, to k,,,..... 
Enzymatic Catalysis in Organic Solvents 3199 
Table IV) to inhibit the hydrolysis of N-acetyl-L-phenylala- 
nine ethyl ester catalyzed by chymotrypsin in water. The 
conventional kinetic analysis (Webb, 1963) confirmed the 
competitive nature of the inhibition and yielded the corre- 
sponding inhibition constants Ki. As one can see in Table IV, 
an increase in hydrophobicity from benzene to toluene to 
naphthalene indeed results in lower inhibition constants in 
water (ie. in a higher affinity). Furthermore, for each com- 
pound the introduction of a carboxyl group diminishes inhi- 
bition, thereby increasing Ki. This phenomenon can be readily 
explained in terms of the hydrophobic inhibition model for 
the addition of a hydrophilic moiety reduces the incentive for 
the inhibitor to leave water and partition into the active 
center of chymotrypsin. 
Consider now the implications of this model for the inhi- 
bition of chymotrypsin in octane instead of water. First, an 
increase in hydrophobicity should no longer result in a higher 
affinity, as hydrophobic interactions, the driving force of the 
enzyme-inhibitor binding in water, will not exist in octane. 
Second, the introduction of a carboxyl group should enhance 
the inhibition, since the inhibitor will tend to "hide" from 
octane by partitioning into the active center of the enzyme. 
We have found that all six compounds listed in Table IV 
are competitive inhibitors of chymotrypsin in the transester- 
ification of N-acetyl-L-phenylalanine ethyl ester in octane. 
The inhibition constants, determined by a standard kinetic 
analysis (Segel, 1975),are presented in Table IV. One can see 
that the prediction made above holds. While in water Ki for 
naphthalene is 50 times better than for benzene, in octane 
the two are nearly the same. Moreover, in sharp contrast to 
the situation in water, in octane all carboxyl-containing com- 
TABLE IV 
Competitive inhibition of chymotrypsin in water and octane 
Inhibitor 
lnhibition constant K; 
In wateP In octane6 
mM 
Benzene 21 1000 
Benzoic acid 140 40 
Toluene 12 1200 
Phenylacetic acid 160 25 
Naphthalene 0.4 1100 
I-Naphthoic acid 7.2 3 
The aromatic compounds listed in the table were used to inhibit 
the hydrolysis of N-acetyl-L-phenylalanine ethyl ester catalyzed by 
chymotrypsin (lo" M) in an aqueous solution containing 0.1 M KC1 
and 5% dimethyl sulfoxide (to dissolve the inhibitors), pH 7.8, at 
20 "C. At each given concentration of the inhibitor (in the range from 
0.1 to 200 mM dependent on the affinity), the dependence of the 
initial rate of the enzymatic hydrolysis on the ester concentration 
(0.8-8 mM) was studied in reciprocal coordinates. The resultant 
straight lines afford the K, values following a standard analysis 
(Webb, 1963). 
The aromatic compounds listed in the table were used to inhibit 
the transesterification reaction between N-acetyl-L-phenylalanine 
ethyl ester and n-propanol catalyzed by chymotrypsin in octane. A 
given concentration of the inhibitor (in the range from 1 mM to 1.2 
M depending on the affinity) was added to a suspension of pH- 
adjusted chymotrypsin (1 mg/rnl) in octane containing 1 M propanol, 
0.1% water (to activate the enzyme, see text), and 2-12 mM ester. 
The suspension was vigorously shaken at 20 "C. The initial rates of 
the enzymatic transesterification were plotted against the ester con- 
centration in the reciprocal coordinates at different inhibitor concen- 
trations. A standard kinetic analysis (Segel, 1975) afforded the Ki 
values. We have established that the inhibition by carboxylic com- 
pounds is not complicated by acidification of the reaction medium, 
as 100 mM heptanoic acid had no appreciable effect on the enzymatic 
transesterification in octane. 
pounds are much more potent inhibitors than the parent 
molecules. The effect is particularly striking for naphthalene 
whose affinity to chymotrypsin in water is 18 times higher 
but in octane 370 times lower than that of 1-naphthoic acid. 
The data in Table IV demonstrate that better enzyme 
inhibitors in water become poorer inhibitors in octane and 
vice versa. This phenomenon is both phenomenologically and 
mechanistically similar to a reversal of substrate specificity 
of chymotrypsin, subtilisin, and pig liver carboxyl esterase 
upon replacement of water with octane as the reaction me- 
dium (Zaks and Klibanov, 1986). 
Stability of Chymotrypsin in Organic Solvents-Stability, in 
particular thermostability, is an important functional char- 
acteristic of an enzyme (Klibanov, 1983). It has been reported 
that lipases (Zaks and Klibanov, 19841, terpene cyclase 
(Wheeler and Croteau, 1986), and cytochrome oxidase and 
ATPase (Ayala et al., 1986) are much more thermostable in 
organic solvents than in water. This phenomenon can be 
readily explained by our recent findings (Ahern and Klibanov, 
1985; Zale and Klibanov, 1986) that all the processes causing 
irreversible thermal inactivation of enzymes require water. In 
the present work, we examined thermostability of chymotryp- 
sin in organic solvents under various conditions. 
Chymotrypsin was lyophilized from an aqueous solution, 
pH 7.8, containing 0.25% ligand N-acetyl-L-phenylalanine 
and then placed (1 mg/ml) in dry octane. The suspension was 
incubated at 100 "C, and periodically the enzyme was assayed 
in water at 20 "C. The half-life of chymotrypsin was found to 
be 4.5 h. Thus, chymotrypsin, similar to other enzymes, is 
much more stable in an organic solvent than in water, e.g. the 
half-life of the enzyme in aqueous solution at pH 8.0 even at 
55 "C is less than 15 min (the decay was due to a monomo- 
lecular thermoinactivation and not due to autolysis) (Marti- 
nek et al., 1977). 
When chymotrypsin was lyophilized from an aqueous so- 
lution, pH 3.0, containing 0.25% N-acetyl-L-phenylalanine, 
its half-life in octane at 100 "C was determined to be half an 
hour, i.e. one-ninth of that for the enzyme sample lyophilized 
from pH 7.8. Thus, the "pH memory" of chymotrypsin applies 
not only to its catalytic activity but also to thermal stability. 
We then investigated thermal stability in other organic 
solvents. The half-lives of the enzyme (lyophilized from pH 
7.8 as outlined above) in butyl ether, tert-amyl alcohol, diox- 
ane, and pyridine were 130,8.0,3.5, and 1.5 min, respectively. 
That is, thermostability of chymotrypsin strongly depends on 
the nature of the solvent and is higher in hydrophobic than 
in hydrophilic ones (similar conclusions have been recently 
reported by Reslow et a1. (1987)). However, even in such 
unfavorable solvents as dioxane and pyridine, the enzyme is 
far more stable than in water. 
In addition to thermal stability, we also studied storage 
stability of chymotrypsin in octane versus water. At 20 "C, 
the enzyme (1 mg/ml) dissolved in 0.05 M aqueous phosphate 
buffer (pH 7.8) lost half of its catalytic activity after 1 week. 
At the same time, when chymotrypsin (lyophilized from 0.25% 
aqueous phosphate, pH 7.8) was suspended (1 mg/ml) in dry 
octane, full enzymatic activity was retained even after 6 
months of incubation at 20 "C. Hence replacement of water 
with organic solvents greatly enhances enzymes stability both 
at high and ambient temperatures. 
Why Do Enzymes Not Inactivate in Organic Solvents?-The 
data presented above unequivocally demonstrate that chy- 
motrypsin and subtilisin vigorously function as catalysts in 
organic solvents. Both are extracellular proteases whose nat- 
ural habitat does not involve nonaqueous media. Keeping in 
mind that several other unrelated enzymes were also found 
3200 Enzymatic Catalysis in Organic Solvents 
to be catalytically active in organic solvents (Klibanov, 1986), 
it appears that nonaqueous enzymology is a general phenom- 
enon. Since the notion that enzymes need aqueous solutions 
to work is one of the central dogmas of biochemistry, this 
conclusion poses an unsettling question of how it is possible. 
There are two alternative reasonings for enzymatic catalysis 
in organic solvents, thermodynamic (i.e. the enzyme “does 
not want” to unfold upon transition from water to aii organic 
solvent) or kinetic (ie. the enzyme is “unable” to unfold upon 
the transition from water to an organic solvent because of 
insurmountable kinetic barriers). The thermodynamic con- 
cept can be readily dismissed. It is inconceivable that the 
delicate balance of all noncovalent interactions maintaining 
the thermodynamically favored enzyme conformation (Schulz 
and Schirmer, 1979) is the same in such diverse solvents as 
octane and tert-amyl alcohol or ethyl ether and pyridine 
(where subtilisin exhibits identical activities, Table I). 
Therefore, the most likely explanation is that enzymes are 
catalytically active in organic solvents because they cannot 
radically change their native conformation upon the transi- 
tion from water to a nonaqueous solvent due to high kinetic 
barriers. This explanation is consistent with numerous exper- 
imental facts. The phenomenon of “pH memory” of enzymes 
discussed by Klibanov (1986) and in this study points to the 
existence of kinetically trapped enzyme structures in organic 
solvents. This is also confirmed by the observation that ad- 
dition of a high concentration (up to 0.1 M) of strong acids to 
lipase in hexane does not appreciably inactivate the enzyme 
(Kirchner et al.,1985). In addition, a high rigidity of enzymes 
in organic solvents is reflected by their greatly enhanced 
thermal stability (Zaks and Klibanov, 1984). 
The existence of high kinetic barriers was also supported 
by the following indicative experiment. Solid pH-adjusted 
chymotrypsin was dissolved in dry dimethyl sulfoxide which 
is a unique organic solvent in that it dissolves proteins (Singer, 
1962). The conformation of chymotrypsin in dimethyl sulf- 
oxide is radically different from that in water (the enzyme 
molecule is thought to be “turned inside out”) (Klyosov et al., 
1975), and thus it (as well as other enzymes (Zaks and 
Klibanov, 1985)) is completely devoid of catalytic activity 
(Table I). When this solution of chymotrypsin was diluted 
50-fold with acetone containing 3% water (to activate the 
enzyme, see above) and the substrates N-acetyl-L-phenylala- 
nine ethyl ester and propanol, no transesterification reaction 
was detected. However, when solid pH-adjusted chymotrypsin 
(the same amount as ended up in the aforementioned acetone 
solution) was directly added to acetone containing 3% water, 
2% dimethyl sulfoxide (to make it identical with the previous 
acetone solution), and substrates, the rate of the enzymatic 
transesterification was at least 10,000-fold higher than in the 
first acetone solution. Thus, the same enzymatic systems 
prepared by different means display very different catalytic 
behaviors. Irreversible inactivation of chymotrypsin by di- 
methyl sulfoxide can be excluded because when that solution 
was diluted with water 50-fold, the enzyme fully regained its 
catalytic activity in the hydrolysis of N-acetyl-L-phenylala- 
nine ethyl ester. These data strongly suggest that chymotryp- 
sin in acetone is severely kinetically restricted. 
We have also established that the kinetic barriers resulting 
in the retention of chymotrypsin’s native conformation upon 
replacement of water with nonaqueous reaction media do not 
stem from protein-protein contacts in a suspension of the 
enzyme in organic solvents. Chymotrypsin was covalently 
attached to CNBr-activated Sepharose. The immobilized en- 
zyme, where the protein molecules are spatially separated 
from each other, was as catalytically active in the transester- 
ification reaction in octane (0.1% water was also added for 
activation) as its free predecessor under the same conditions. 
Hence, chymotrypsin is locked into the native-like confor- 
mation in organic solvents by intramolecular interactions. 
It is hoped that the concepts and experimental approaches 
developed in this study will be conducive to further explora- 
tion of nonaqueous enzymology. 
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