enzymatic catalysis in nonaqueous solvents*

’ 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 catalysts 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 enzymes by a site-specific reagent renders them catalytically inactive in organic solvents. Upon replacement of water with octane as the reaction medium, the specificity of chymotrypsin toward competitive inhibitors reverses. Both thermal and storage stabilities of chymotrypsin are greatly enhanced in nonaqueous solvents compared to water. The phenomenon of enzymatic 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 native-like conformation from unfolding.

All the vast knowledge accumulated in the area of mechanistic 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 recently successfully developed (Waks, 1986). In the first approach (Luisi, 1985; Martinek et al., 1986), enzymes are dissolved 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 properties 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 understand such basic features and characteristics of this phenomenon as the dependence of enzymatic properties on the nature of the solvent, the amount of water required for catalysis, 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 chymotrypsin 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 enzymes when they were directly dispersed as solids in nonaqueous 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 dried by 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 conditions 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 reactions 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 withdrawn 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 experiment, 10 d of an aqueous solution of the ester (0.3-5.0 mM) containing 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 molecular 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 proteases can also utilize nucleophiles other than water, in particular 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 solvents (Singer, 1962)) was shaken at 20 “C; periodically, aliquots 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 retained 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 reaction 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 fluoride (Fahrney and Gold, 1963) was completely inactive in octane, thus indicating the nonartifactual origin of the transesterification reaction. The rate of the enzymatic transesterification in octane was directly proportional to the concentration 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 transesterification 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 (propanol in our case) to form another binary complex, which then yields the new ester and the free enzyme. This mechanism (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 and propanol 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 enzyme, 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 chymotrypsin-catalyzed hydrolysis in methylene chloride containing 0.25% water, but no quantitative details were pro- vided. In contrast, we have undertaken a mechanistic investigation 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 transesterification 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 contained the ligand N-acetyl-L-phenylalanine instead of phosphate (both compounds were used in the same weight concentration, 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 transesterification 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 absence 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 transesterification 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 because 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 transesterification 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


(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 catalysis in organic solvents.

The very different activities exhibited 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 retained 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, chymotrypsin 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 transesterification 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 reaction 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 chymotrypsin 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 mixture 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 chymotrypsin 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 consequently 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 deacylation 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 chymotrypsin, as revealed by the formation of more propyl trans-cinnamate (curve b in Fig. U), thus bringing the total number of active chymotrypsin molecules to 90% of that 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 subsequent 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-cinnamoylimidazole 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 concentrations 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-phenylalanine ethyl ester and amyl alcohol catalyzed by chymotrypsin 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-phenylalanine 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 interesting 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 catalytic efficiency of chymotrypsin and subtilisin in octane. On the basis of the rate constant of the nonenzymatic transesterification 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 numerous 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,,,.....


Table IV) to inhibit the hydrolysis of N-acetyl-L-phenylalanine ethyl ester catalyzed by chymotrypsin in water. The conventional kinetic analysis (Webb, 1963) confirmed the competitive nature of the inhibition and yielded the corresponding 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 inhibition, 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 inhibition 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 transesterification 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 concentration in the reciprocal coordinates at different inhibitor concentrations. A standard kinetic analysis (Segel, 1975) afforded the Ki values. We have established that the inhibition by carboxylic compounds 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 medium (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 chymotrypsin 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 solution, 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, dioxane, 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 chymotrypsin and subtilisin vigorously function as catalysts in organic solvents. Both are extracellular proteases whose natural 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 phenomenon. 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 transition from water to a nonaqueous solvent due to high kinetic barriers. This explanation is consistent with numerous experimental 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 addition 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 sulfoxide 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-phenylalanine 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 dimethyl 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-phenylalanine ethyl ester. These data strongly suggest that chymotrypsin 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 enzyme, 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 conformation 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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