Cutinase Activity and Enantioselectivity in Supercritical Fluids

2780 Oeiras, Portugal, and Centro de Engenharia Biolo´gica e Quı´mica, Instituto Superior Te´cnico,. Av. Rovisco Pais, 1096 Lisboa Codex, Portugal...
1 downloads 0 Views 75KB Size
Download PDF
Ind. Eng. Chem. Res. 1998, 37, 3189-3194

3189

Cutinase Activity and Enantioselectivity in Supercritical Fluids Nuno Fontes,† M. Conceic¸ a˜ o Almeida,† Ce´ lia Peres,† Sı´lvia Garcia,† Joa˜ o Grave,† M. Raquel Aires-Barros,‡ Cla´ udio M. Soares,† Joaquim M. S. Cabral,‡ Christopher D. Maycock,† and Susana Barreiros*,† Instituto de Tecnologia Quı´mica e Biolo´ gica, Universidade Nova de Lisboa, Quinta do Marqueˆ s, Apt. 127, 2780 Oeiras, Portugal, and Centro de Engenharia Biolo´ gica e Quı´mica, Instituto Superior Te´ cnico, Av. Rovisco Pais, 1096 Lisboa Codex, Portugal

We studied the performance of Fusarium solani pisi cutinase, immobilized on a zeolite, in supercritical fluids. The catalytic activity of the enzyme was strongly dependent on water activity, was unaffected by pressure up to 300 bar, and was higher in supercritical ethylene than in supercritical carbon dioxide. The enzyme was very selective toward one of the isomers of 1-phenylethanol, with an enantiomeric excess of virtually 100%, regardless of water activity, pressure, solvent, and temperature. We used the X-ray crystal structure of the enzyme and did a computer modeling of the structures of the transition states formed by the two enantiomers. The differences between these structures helped elucidate the preference for the (R)-enantiomer. Introduction The properties of supercritical fluids (SCF) which make them interesting solvents for extraction also make them attractive media for conducting enzymatic reactions, an area of research which began in 1985.14,40,46 Carbon dioxide is unquestionably the preferred SCF for a number of reasons, among which a close to ambient critical temperature, a moderately high critical pressure, nonflammability, and low cost. Adjustable physical properties are one of the most useful characteristics of SCF, allowing, for example, the manipulation of the solubilities of solutes. This adjustability is only very pronounced at temperatures and pressures not far from the critical point of the fluid, making the range of high compressibility close to that point the most interesting to explore. The rapid growth of the market for enantiopure compounds and the fact that nonaqueous conventional solvents are already being used in productionscale applications to affect the performance of enzymes27 opens interesting perspectives for the use of SCF as solvents for enantioselective biotransformations, allowing in particular a reduction of organic solvent residues. The cutinase used in the present work is an extracellular enzyme produced by the plant pathogen Fusarium solani pisi. Although it is often classified as a lipase, it lacks a lid33 and does not exhibit interfacial activation.29 It accepts a wide range of substracts, has considerable thermostability,29 and shows specificity toward smaller chain esters, fatty acids, and alcohols.8,31 There is still little information on its potential to catalyze enantioselective transformations,31,32 and this was one of the motivations for the present study. The substrate chosen, 1-phenylethanol, is a convenient choice for a kinetic resolution given the large bulk differences among the substituents attached to the chiral center. It has been used in a variety of asymmetric transformations21 and also in the modeling of enzyme enantioselectivity.10 * To whom correspondence should be addressed. Phone: 351-1-446 9441. Fax: 351-1-441 1277. E-mail: [email protected]. † Universidade Nova de Lisboa. ‡ Instituto Superior Te ´ cnico.

As shown by many authors, enzyme activity in nonaqueous solvents is usually particularly sensitive to the level of hydration of the enzyme2,3,48 which is related to the water activity, aW. The effect of pressure on enzyme activity is usually larger in nonaqueous liquid solvents than in aqueous media,11,25,38 and can be very significant close to the critical point of a SCF.6,9,11 Enantioselectivity has been found not only to be insensitive to variations in aW4,35 but also to depend on the latter parameter.42 Pressure has also been used to affect enzyme enantioselectivity.15,18 To help elucidate the behavior of cutinase in supercritical CO2, we examined the effect of aW, pressure, and temperature on the catalytic activity and enantioselectivity of the immobilized enzyme suspended in that solvent. To test the influence of the medium, we also obtained a few data points in ethylene. To try to explain the observed results on enantioselectivity, we used the crystal structure of the enzyme and analyzed structural differences between transition states for the two enantiomers. Experimental Section F. solani pisi cutinase, cloned and expressed in Escherichia coli WK-6, was a gift from Corvas International (Gent, Belgium). The recombinant enzyme was produced and purified in one of our laboratories according to a published method.29 The lyophilized enzyme obtained was dissolved in a 50 mM sodium phosphate buffer solution (10 mg cm-3 of enzyme) at pH ) 8 and immobilized onto a zeolite support (4-Å molecular sieves powder from Aldrich; 25 mg of enzyme/g of support).12 The amount of protein immobilized was determined using a modified Lowry method.12 It was experimentally confirmed that the enzyme did not desorb from the support during the course of reaction within the error associated with that method. The cutinase estereolytic activity was determined spectrophotometrically, by following the hydrolysis of p-nitrophenyl butyrate at 400 nm in the reaction mixture 0.56 mM p-nitrophenyl butyrate, 11.3 mM sodium cholate, and 340 mM tetrahydrofuran, in a 50 mM sodium phosphate buffer at pH ) 8 (pH before addition of reactants). Racemic 1-phenylethanol, (S)-(1)-phenylethanol, and

S0888-5885(97)00933-0 CCC: $15.00 © 1998 American Chemical Society Published on Web 07/03/1998

3190 Ind. Eng. Chem. Res., Vol. 37, No. 8, 1998

(R)-(1)-phenylethanol were from Aldrich and vinyl butyrate was from Fluka. All sodium salt hydrates and reagents used in the quantification of the amount of protein adsorbed onto the support were from Merck with the exception of the protein standard which was from Sigma and Hydranal Coulomat A and C Karl Fischer reagents were from Riedel de Haen. CO2, ethylene, and nitrogen were supplied by Air Liquide and guaranteed to have purities of over 99.995 or 99.95 mol % (ethylene). We used variable volume stainless steel cells equipped with a sapphire window. Details of the experimental apparatus are given elsewhere.3 To measure an initial rate, the cell was loaded with the immobilized enzyme preparation (16 mg cm-3), vinyl butyrate and the desired amount of water, and pressurized with the solvent to the desired volume, at the selected pressure and temperature. The lower aW values were obtained by adding dried molecular sieves (the kind used in the immobilization procedure) to the reaction system. After a 2-h period of equilibration, samples were taken and released directly into a Karl Fischer apparatus to determine the water content of the solvent at equilibrium. To proceed for reaction, an appropriate mixture of 1-phenylethanol, vinyl butyrate, and water was flushed into the cell with solvent. Periodically, stirring was stopped and samples were taken for analysis. More details of the experimental technique are given in ref 3. The amounts of the two substrates were changed so as to keep the respective mole fractions constant in experiments at different pressures. For the determination of the enantiomeric excess, samples were taken every 2 h and analyzed until the conversion reached a constant value. To convert the water content of the solvent at equilibrium with the enzyme into aW values, separate equilibration of the solvent (with or without the substrates) with mixtures of salt hydrates known to give a certain aW were made. The salt hydrate pairs used and the respective aW values at 35 °C were13 Na2HPO4‚2/0 (0.19), Na2HPO4‚7/2 (0.69), and Na2HPO4‚12/7 (0.90). The mixtures were allowed to equilibrate for about 24 h, after which six samples were taken and released into the Karl Fischer apparatus. Reaction conversion was followed by GC analysis and performed with a 6000 Vega Series 2 Carlo-Erba gas chromatograph with a Chromjet Spectra-Physics integrator. GC conditions were as follows: 15-m DB-5 capillary column from J&W Scientific; oven temperature program, 5 min at 100 °C/20 °C min-1 ramp/150 °C for 2 min; injection temperature, 250 °C; flame ionization detection (FID) temperature, 270 °C; carrier gas, helium (1 cm3 min-1); split ratio, 1:50. Chiral HPLC separations were performed with a 25-cm Daicel OD-H (4.6mm o.d.) column and a mobile phase of 5% 1-propanol and 95% n-hexane, at a flow rate of 1 cm3 min-1. The ester product was separated by chiral GC with a 30-m fused silica β-cyclodextrin capillary column from Supelco; oven temperature program, 30 min at 40 °C/20 °C min-1 ramp to 200 °C; injector temperature, 120 °C; (FID) temperature, 280 °C; carrier gas, helium (1 cm3 min-1); split ratio, 1:50. The computer-assisted generation of the transition states for both enantiomers was made on the basis of the structure of a complex of cutinase with the inhibitor N-hexylphosphonate ethyl ester30 (PDB accession code: 1XZL) using the programs Sybyl 6.2 from TRIPOS and Turbo-Frodo.47 The structures obtained in this way

Figure 1. Water activity in the solvent mixtures as a function of water mole fraction or water concentration. Full symbols, 80 bar; open symbols, 250 bar. 1/3, CO2; b/O, CO2 + vinyl butyrate; 9/0, CO2 + vinyl butyrate + 1-phenylethanol. 2, ethylene + vinyl butyrate + 1-phenylethanol. Data for CO2 at 35 °C and for ethylene at 15 °C.

were subjected to local energy minimization (residues around the transition state) in Sybyl using the TRIPOS force field plus Kolmann united atom charges (united atoms were used). A distance-dependent dielectric was used for electrostatics and interactions were truncated at 10 Å. Results and Discussion The catalytic activity of an enzyme in a nonaqueous solvent is best correlated with the parameter enzyme hydration which measures the amount of water directly associated with the enzyme. At water-partitioning equilibrium between enzyme and solvent, enzyme hydration is a function of the water activity, aW, which characterizes all of the phases present in the heterogeneous system. For immobilized enzyme preparations, it is not possible to distinguish between the hydration of the enzyme molecules and the hydration of the support. However, provided that aW is kept constant and that solvents do not differ significantly in polarity, enzyme hydration in the various media should be the same for a given aW.37,43 Fixing a certain aW value may be accomplished in a variety of ways: for example, separate preequilibration of enzyme and solvent with saturated salt solutions,16,48 direct addition to the reaction mixture of pairs of salt hydrates known to confer a certain aW,50 circulation of saturated salt solutions in a silicone tubing immersed in the reaction medium.22 We used the second technique indirectly to build a scale of aW versus [H2O] with which we could convert [H2O] in the reaction mixtures at water-partitioning equilibrium into aW. A similar approach was used by Almeida et al.1 and Bell et al.2 With the salt hydrate pairs that we had available, we obtained three experimental points on the scale. Additionally, the solvent mixtures were saturated with water, yielding points for aW ) 1. The results obtained are shown in Figure 1. The solubility of water in CO2 is 0.56 g dm-3 at 80 bar and 1.75 g dm-3 at 250 bar,7,49 values which agree with those derived from Figure 1 to (10%. The addition of the substrates causes a 30% increase in the solubility of water on going from CO2 to the ternary mixture at 80 bar. At this pressure, and as seen also in the figure, the addition of the alcohol to the mixture of CO2 and vinyl butyrate does not change the physical properties of the binary significantly, which should be due to the low concentration of alcohol used

Ind. Eng. Chem. Res., Vol. 37, No. 8, 1998 3191

Figure 2. Dependence of the catalytic activity of cutinase on water activity. 9, CO2 at 80 bar; ×, CO2 at 150 bar; 0, CO2 at 250 bar; 2, ethylene at 80 bar. Reaction conditions: T ) 35 or 15 °C (ethylene); [enzyme preparation] ) 16 mg cm-3; constant mole fractions of the substrates (e.g., [vinyl butyrate] ) 238 mM and [1-phenylethanol] ) 24 mM at 80 bar in CO2).

(we used mole fractions of vinyl butyrate slightly below 0.02 and of phenylethanol 10 times lower). Thus, the increase in water solubility of the ternary mixture over that of single CO2 can be attributed to the presence of vinyl butyrate. The solubility of water in vinyl butyrate is much higher than the solubility of water in CO2 at 80 bar, but is similar to that of water in CO2 at 250 bar. This explains why the experimental points obtained at 250 bar all lie on the same line. The linearity of the plots in the figure implies constant activity coefficients for water in the media. Included in Figure 1 is also a point obtained in ethylene at 80 bar and aW ) 0.9. This point was used to give an indication of the aW values at which a few activity measurements in ethylene were made. We used the data in Figure 1 to plot enzyme activity as a function of aW, as shown in Figure 2. To include in this figure our results at 150 bar, we calculated an activity coefficient for water from the inverse of the published water solubility in CO2 at that pressure.7,49 The data in Figure 2 define nearly bell-shaped curves. Evidence for such plots of cutinase activity versus aW were presented for both the free enzyme and the enzyme physically adsorbed onto a support.28 These authors showed that enzyme activity became zero before aW became 1. As seen in Figure 2, in our case the enzyme remains catalytically active even after saturation of the solvent with water. This may be related to the type of support used. In fact, for cutinase covalently bound to a support, the above authors observed a continuous increase in activity with increasing aW. Due to the fact that the reaction studied is a transesterification with no net production or consumption of water, aW in our case should remain constant throughout the reaction. However, we note that at high aW levels hydrolysis of both the substrate and the product esters will most likely occur to a certain extent, in each case with the formation of butyric acid. Indeed, we detected the presence of this acid at those conditions. Competing hydrolysis could thus cause a downward shift of aW during the course of the reaction at high aW values. The direct effect of pressure on a reaction rate constant, k, is determined by the activation volume, ∆Vq ) -RT (∂(ln k)/∂P)T, where P is the pressure, T the temperature, and R the gas constant. The variable plotted on the Y-axis of Figure 2 was converted into pressure-independent units by dividing by the molar volume of the single solvent at each pressure of interest.

As the figure shows, the data obtained at different pressures are practically superimposable over the whole aW range, leading to a small in magnitude ∆Vq in CO2. ∆Vq values ranging from -60 to +60 cm3 mol-1 were reported for subtilisin Carlsberg in several organic solvents.25,38 ∆Vq values at least 1 order of magnitude higher than those in liquid solvents were presented for a lipase in fluoroform19 and for subtilisin Carlsberg in CO2 and in ethane.11 In both cases, this was mainly attributed to changes in the physical properties of the solvents, rather than to an intrinsic contribution of pressure to ∆Vq. We note that using the molar volume of the single solvent instead of that of the solvent mixture to correct our measured Vmax/Km could artificially lower the points obtained in CO2 and ethylene at 80 bar, perhaps moderately so given the low-solute mole fractions used. In fact, close to the critical point one might expect a local density enhancement of the solvent around the solute molecules, resulting in negative solute partial molar volumes.26 In any case, the generally low impact of pressure on cutinase activity requires further investigation. There is now strong evidence that CO2 has an adverse effect on the catalytic activity of some enzymes, free3,17 or immobilized,1,36 for which different explanations were presented.20,44 Figure 2 does not seem to conform to such evidence. As pointed out by Kamat et al.,20 the inhibitory effect of CO2 should depend on the response of the enzyme to surface modification by CO2 molecules, which could be lower in the case of cutinase. However, we note that the data points for ethylene were obtained at 15 °C, whereas those for CO2 were obtained at 35 °C. It is reasonable to conceive that an increase in temperature in ethylene should be accompanied by an increase in enzyme activity. At aW values close to the optimum, that is, at the top of the bell-shaped curve, differences in enzyme activity between the two solvents should be more clearly visible.3 In regard to enzyme enantioselectivity, we found that cutinase only converted the (R)-isomer of phenylethanol, yielding enantiomeric ratios of 100% for both the alcohol and the ester product, within the error associated with the measurements. The fact that the conversion of the alcohol did not go beyond 50% was already an indication that the enzyme was highly selective toward one of the isomers. This marked selectivity was insensitive to the solvent, to aW over the whole aW range, to pressure, and to temperature (25, 35, and 45 °C) whose potential for modulating enantioselectivity has been discussed recently.45 To try and understand this, we did a computer modeling of the structures of transition states involved in the conversion of the isomers. We considered the deacylation step of the transesterification mechanism; it is in step that enantioselectivity is determined, given that the chiral agent is the alcohol. Our rationale was to assume that the active site was equally accessible to both isomers and that enantiomeric discrimination should depend on differences in the stabilization of the tetrahedral transition state for deacylation.24 To determine the orientation of the acylating agent part in the transition state, there were two essential mechanistic requirements that had to be met:23,34,41 (a) the nucleophilic oxygen of serine 120 must be covalently bound to the carbonyl carbon of the reacting ester and (b) the anionic oxygen must be oriented toward the oxyanion hole (serine 42, glutamine 121). None of the two possible orientations of the acyl chain of the

3192 Ind. Eng. Chem. Res., Vol. 37, No. 8, 1998

Figure 3. Hypothetical transition states for transesterification of the two enantiomers of 1-phenylethanol by cutinase. Colors for atoms: carbon, gray; nitrogen, dark blue; oxygen, red; hydrogen, cyan. Only polar hydrogens are displayed. The phenyl and methyl groups of the transition states are colored purple and green, respectively.

acylating agent determined by (a) and (b) involve significant steric hindrance. To fix this orientation, we looked at published structures of cutinase bound to ester inhibitors30 and found that the acyl chain was preferentially oriented toward the tunnel region defined by valine 184 and leucine 81, in a hydrophobic cavity limited by leucine 182. We assumed this was the case here. That left only one mode of binding of the alcohol part of the complex. There were basically two possible locations for the phenyl group: a hydrophobic pocket defined by leucine 187, valine 184, and tyrosine 119, and an open region close to the surface of the enzyme (where oxygen atoms of polar residues can be seen in red, in Figure 3). We chose the former, on the basis of the favorable van der Waals interactions established and the high degree of complementarity between the protein and the transition state. Again, we note that this construction does not lead to significant steric hindrance of the complex and thus does not cause significant conformational changes on the enzyme after energy minimization: serine 42 moves slightly and

establishes a hydrogen bond with the negative oxygen of the complex, which is also in hydrogen bond contact with the main chain nitrogen of glutamine 121. Valine 184 and leucine 189 adjust themselves to the phenyl group. All these hydrophobic residues have high mobility in the X-ray structure,30,33 and thus the movement of these groups in the way described is not energetically important. Another residue that moves is tyrosine 119. This residue is partially exposed and has plenty of space in the original structure to change its conformation. Its OηH group establishes a hydrogen bond with the backbone oxygen of histidine 188, an interaction maintained in the transition states for the (R)- and (S)isomers. If the planar phenyl group is thus fixed, the structural difference between the transition states for the two enantiomers lies in the position of the methyl group attached to the chiral center. In the case of the (R)isomer, the methyl group is inserted in the structure, in close contact with the ring of tyrosine 119, whereas in the case of the (S)-isomer, this group is oriented toward the entrance to the active-site cavity. This situation is illustrated in Figure 3. From this figure, a plausible explanation for the enzyme preference for the (R)-isomer emerges: the burial of the methyl group in the case of the (R)-isomer and the van der Waals interactions of this group with tyrosine 119 which immobilize and stabilize the transition state. A high degree of complementarity between the protein and the complex, as referred to above when discussing the position of the phenyl group, leads to a more efficient catalytic cycle. We note that the nonaqueous solvent molecules may play a role in the stabilization of the methyl group of the (S)-isomer. Even at aW ) 1, the enzyme does not experience a situation similar to that in an aqueous solution, given that the formation of a complete monolayer of water over a protein surface is thermodynamically unfavorable.37,43 In fact, evidence for substantial solvent penetration of the active site in the vicinity of the transition state has been presented for subtilisin.38 But apparently, these interactions are less effective from the standpoint of catalysis than those of the methyl group with protein residues in the (R)isomer conformation. Ke et al.24 presented a model for predicting enzyme enantioselectivity based on enzyme structure which relies on the thermodynamics of desolvation of the substrates in the relevant transition states. We cannot apply the model to the present study given that the two enantiomers are essentially equally desolvated in the transition state, the main difference being a hydrogen atom versus a methyl group oriented toward the entrance to the active site. But we note that among the factors referred to by the above authors which may limit the applicability of a structure-based approach to enzyme enantioselectivity, a most important one is the preservation of the crystal structure of the enzyme in the nonaqueous solvent, which the mode of preparation of our enzyme should ensure. Any disruption of the structure occurring during lyophilization5 should have been reversed upon solubilization in an aqueous buffer, and the enzyme preparation was not dried extensively after immobilization. As for conformational changes due to water stripping in the solvent, we note that cutinase enantioselectivity did not depend on enzyme hydration and that at higher hydration levels we would expect to avoid the above indirect solvent effect.

Ind. Eng. Chem. Res., Vol. 37, No. 8, 1998 3193

Conclusions A scale of aW as a function of [H2O] is a useful tool for building an enzyme activity versus aW relationship. The catalytic activity of cutinase was very sensitive to changes in aW, in agreement with results from other authors. The enzyme was less active in CO2 than in ethylene, confirming the notion of an adverse effect of CO2 on some enzymes documented in the literature. The catalytic performance of the enzyme was practically unaffected by pressure. Although pressures to 300 bar were not expected to result39 in a very large (in magnitude) intrinsic ∆Vq, a solvent-mediated contribution to the latter parameter, especially close to the critical point of the solvent, might exist, as observed in other cases. By constructing model structures of the transition states for the two enantiomers of the alcohol substrate, the preference of the enzyme for the (R)enantiomer could be understood. Acknowledgment This work was supported by Junta Nacional de Investigac¸ a˜o Cientı´fica e Tecnolo´gica (JNICT, Portugal) through the contract PRAXIS 2/2.1/BIO/34/94. C.M.S. acknowledges support from the JNICT Grant PBIC/C/ 2037/95 and the PRAXIS XXI Fellowship BPD/4151/94. Literature Cited (1) Almeida, M. C.; Ruivo, R.; Maia, C.; Freire, L.; Correˆa de Sampaio, T.; Barreiros, S. Novozym 435 Activity in Compressed Gases. Water Activity and Temperature Effects. Enzyme Microb. Technol. 1998, 22, 494-499. (2) Bell, G.; Janssen, A. E. M.; Halling, P. J. Water Activity Fails To Predict Critical Hydration Level for Enzyme Activity in Polar Organic Solvents: Interconversion of Water Concentrations and Activities. Enzyme Microb. Technol. 1997, 20, 471-177. (3) Borges de Carvalho, I.; Correˆa de Sampaio, T.; Barreiros, S. Solvent Effects on the Catalytic Activity of Subtilisin Suspended in Compressed Gases. Biotechnol. Bioeng. 1996, 49, 399-404. (4) Bovara, R.; Carrea, G.; Ottolina, G.; Riva, S. Water Activity Does Not Influence the Enantioselectivity of Lipase PS and Lipoprotein Lipase in Organic Solvents. Biotechnol. Lett. 1993, 15, 169-174. (5) Burke, P. A.; Griffin, R. G.; Klibanov, A. M. Solid-State NMR Assessment of Enzyme Active Center Structure under Nonaqueous Conditions. J. Biol. Chem. 1992, 267, 20057-20064. (6) Chaudhary, A. K.; Kamat, S. V.; Beckman, E. J.; Nurok, D.; Kleyle, R. M.; Hadju, P.; Russell, A. J. Control of Subtilisin Substrate Specificity by Solvent Engineering in Organic Solvents and Supercritical Fluoroform. J. Am. Chem. Soc. 1996, 118, 12891-12901. (7) Chrastil, J. Solubility of Solids and Liquids in Supercritical Gases. J. Phys. Chem. 1982, 86, 3016-3021. (8) Cunnah, P. J.; Aires-Barros, M. R.; Cabral, J. M. S. Esterification and Transesterification Catalyzed by Cutinase in Reverse Micelles of CTAB for the Synthesis of Short Chain Esters. Biocatal. Biotransform. 1996, 14, 125-146. (9) Erickson, J. C.; Schyns, P.; Cooney, C. L. Effect of Pressure on an Enzymatic Reaction in a Supercritical Fluid. AIChE J. 1990, 36, 299-301. (10) Fitzpatrick, P. A.; Steinmetz, A. C. U.; Ringe, D.; Klibanov, A. M. Enzyme Crystal Structure in a Neat Organic Solvent. Proc. Natl. Acad. Sci. U.S.A. 1993, 90, 8653-8657. (11) Fontes, N.; Nogueiro, E.; Elvas, A. M.; Correˆa de Sampaio, T.; Barreiros, S. Effect of Pressure on the Catalytic Activity of Subtilisin Carlsberg Suspended in Compressed Gases. Biochim. Biophys. Acta 1998, 1383, 165-174. (12) Gonc¸ alves, A. P. V.; Lopes, J. M.; Lemos, F.; Ramoˆa Ribeiro, F.; Prazeres, D. M. F.; Cabral, J. M. S.; Aires-Barros, M. R. Effect of the Immobilization Support on the Hydrolytic Activity of a Cutinase from Fusarium solani pisi. Enzyme Microb. Technol. 1997, 20, 93-101.

(13) Halling, P. J. Salt Hydrates for Water Activity Control with Biocatalysts in Organic Media. Biotechnol. Technol. 1992, 6, 271276. (14) Hammond, D. A.; Karel, M.; Klibanov, A. M.; Krukonis, V. J. Enzymatic Reactions in Supercritical Gases. Appl. Biochem. Biotechnol. 1985, 11, 393-400. (15) Ikushima, Y.; Saito, N.; Arai, M.; Blanch, H. W. Activation of a Lipase Triggered by Interactions with Supercritical Carbon Dioxide in the Near-Critical Region. J. Phys. Chem. 1995, 99, 8941-8944. (16) Janssen, A. E. M.; Vaidya, A. M.; Halling, P. J. Substrate Specificity and Kinetics of Candida rugosa Lipase in Organic Media. Enzyme Microb. Technol. 1996, 18, 340-346. (17) Kamat, S.; Barrera, J.; Beckman, E. J.; Russell, A. J. Biocatalytic Synthesis of Acrylates in Organic Solvents and Supercritical Fluids: I. Optimization of Enzyme Environment. Biotechnol. Bioeng. 1992, 40, 158-166. (18) Kamat, S. V.; Beckman, E. J.; Russell, A. J. Control of Enzyme Enantioselectivity with Pressure Changes in Supercritical Fluoroform. J. Am. Chem. Soc. 1993, 115, 8845-8846. (19) Kamat, S. V.; Beckman, E. J.; Russell, A. J. Enzyme Activity in Supercritical Fluids. Crit. Rev. Biotechnol. 1995, 15, 41-71. (20) Kamat, S.; Critchley, G.; Beckman, E. J.; Russell, A. J. Biocatalytic Synthesis of Acrylates in Organic Solvents and Supercritical Fluids: Does Carbon Dioxide Covalently Modify Enzymes? Biotechnol. Bioeng. 1995, 46, 610-620. (21) Kanerva, L. T. Hydrolase-Catalysed Asymmetric and Other Transformations of Synthetic Interest. In Enzymatic Reactions in Organic Media; Koskinen, A. M. P., Klibanov, A. M., Eds.; Blackie Academic & Professional: Glasgow, U.K., 1996; pp 170-223. (22) Kaur, J.; Wehtje, E.; Adlercreutz, P.; Chand, S.; Mattiasson, B. Water Transfer Kinetics in a Water Activity Control System Designed for Biocatalysis in Organic Media. Enzyme Microb. Technol. 1997, 21, 496-501. (23) Kazlauskas, R. J. Elucidating Structure-Mechanism Relationships in Lipases: Prospects for Predicting and Engineering Catalytic Properties. TIBTECH 1994, 12, 464-472. (24) Ke, T.; Wescott, R.; Klibanov, A. M. Prediction of the Solvent Dependence of Enzymatic Prochiral Selectivity by Means of Structure-Based Thermodynamic Calculations. J. Am. Chem. Soc. 1996, 118, 3366-3374. (25) Kim, J.; Dordick, J. S. Pressure Affects Enzyme Function in Organic Media. Biotechnol. Bioeng. 1993, 42, 772-776. (26) Kim, S.; Johnston, K. P. Effects of Supercritical Solvents on the Rates of Homogeneous Chemical Reactions. In Supercritical Fluids: Chemical and Engineering Principles and Applications; Squires, T. G., Paulaitio, M. E., Eds.; ACS Symposium Series 329; American Chemical Society: Washington, DC, 1987; pp 42-55. (27) Koskinen, A. M. P. Enzymes in Organic Solvents: Meeting the Challenges. In Enzymatic Reactions in Organic Media; Koskinen, A. M. P., Klibanov, A. M., Eds.; Blackie Academic & Professional: Glasgow, U.K., 1996; pp 1-8. (28) Lamare, S.; Legoy, M. D. Working at Controlled Water Activity in a Continuous Process: The Gas/Solid System as a Solution. Biotechnol. Bioeng. 1995, 45, 387-397. (29) Lauwereys, M.; de Geus, P.; de Meutter, J.; Stanssens, P.; Matthyssens, G. Cloning, Expression and Characterisation of Cutinase, a Fungal Lipolytic Enzyme. In Lipases: Structure, Mechanism and Genetic Engineering; Alberghina, L., Schmid, R. D., Verger, R., Eds.; VCH: New York, 1990; pp 243-251. (30) Longhi, S.; Nicolas, A.; Creveld, L.; Egmond, M.; Verrips, C. T.; de Vlieg, J.; Martinez, C.; Cambillau, C. Dynamics of Fusarium solani Cutinase Investigated Through Structural Comparison Among Different Crystal Forms of its Variants. Proteins 1996, 26, 442-458. (31) Mannesse, M. L. M.; Cox, R. C.; Koops, B. C.; Verheij, H. M.; Haas, G. H.; Egmond, M. R.; van der Hijden, H. T. W. M.; de Vlieg, J. Cutinase from Fusarium solani pisi Hydrolyzing Triglyceride Analogues. Effect of Acyl Chain Length and Position in the Substrate Molecule on Activity and Enantioselectivity. Biochemistry 1995, 34, 6400-6407. (32) Mannesse, M. L. M.; de Haas, G. H.; van der Hijden, H. T. W. M.; Egmond, M. R.; Verheij, H. M. Chiral Preference of Cutinase in the Reaction with Phosphonate Inhibitors. Biochem. Soc. Trans. 1997, 25, 165-171. (33) Martinez, C.; De Geus, P.; Lauwereys, M.; Matthyssens, G.; Cambillau, C. Fusarium solani pisi Cutinase is a Lipolytic

3194 Ind. Eng. Chem. Res., Vol. 37, No. 8, 1998 Enzyme with a Catalytic Serine Accessible to Solvent. Nature 1992, 356, 615-618. (34) Martinez, C.; Nicolas, A.; van Tilbeurgh, H.; Egloff, M.-P.; Cudrey, C.; Verger, R.; Cambillau, C. Cutinase, a Lipolytic Enzyme with a Preformed Oxyanion Hole. Biochemistry 1994, 33, 83-89. (35) Martins, J. F.; Nunes da Ponte, M.; Barreiros, S. Lipase Catalyzed Esterification of Glycidol in Organic Solvents. Biotechnol. Bioeng. 1993, 42, 465-468. (36) Marty, A.; Chulalaksananukul, W.; Willemot, R. M.; Condoret, J. S. Kinetics of Lipase-Catalyzed Esterification in Supercritical CO2. Biotechnol. Bioeng. 1992, 39, 273-280. (37) McMinn, J. H.; Sowa, M. J.; Charnick, S. B.; Paulaitis, M. E. The Hydration of Proteins in Nearly Anhydrous Organic Solvent Suspensions. Biopolymers 1993, 33, 1213-1224. (38) Michels, P. C.; Dordick, J. S.; Clark, D. S. Dipole Formation and Solvent Electrostriction in Subtilisin Catalysis. J. Am. Chem. Soc. 1997, 119, 9331-9335. (39) Mozhaev, V. V.; Heremans, K.; Frank, J.; Masson, P.; Balny, C. High-Pressure Effects on Protein Structure and Function. Proteins 1996, 24, 81-91. (40) Nakamura, K.; Min Chi, Y.; Yamada, Y.; Yano, T. Lipase Activity and Stability in Supercritical Carbon Dioxide. Chem. Eng. Commun. 1986, 45, 207-212. (41) Nicolas, A.; Egmond, M.; Verrips, C. T.; de Vlieg, J.; Longhi, S.; Cambillau, C.; Martinez, C. Contribution of Cutinase Serine 42 Chain to the Stabilization of the Oxyanion Transition State. Biochemistry 1996, 35, 398-410. (42) Orrenius, C.; Norin, T.; Hult, K.; Carrea, G. The Candida antarctica Lipase B catalyzed Kinetic Resolution of Seudenol in Nonaqueous Media of Controlled Water Activity. Tetrahedron: Asymmetry 1995, 6, 3023-3090. (43) Parker, M. C.; Moore, B. D.; Blacker, A. J. Measuring Enzyme Hydration in Nonpolar Organic Solvents Using NMR. Biotechnol. Bioeng. 1995, 46, 452-458.

(44) Perrut, M. Enzymatic Reactions in Supercritical Carbon Dioxide. In High Pressure and Biotechnology; Balny, C., Hayashi, R., Heremans, K., Masson, P., Eds.; Colloque INSERM 224; John Libbey: Montrouge, France, 1992; pp 401-410. (45) Philips, R. S. Temperature Modulation of the Stereochemistry of Enzymatic Catalysis: Prospects for Exploitation. TIBTECH 1996, 14, 13-16. (46) Randolph, T. W.; Blanch, H. W.; Prausnitz, J. M.; Wilke, C. R. Enzymatic Cartalysis in a Supercritical Fluid. Biotechnol. Lett. 1985, 7, 325-328. (47) Roussel, A.; Cambillau, C. Turbo-Frodo. In Silicon Graphics Geometry Partner Directory; Silicon Graphics: Mountain View, CA, 1989; pp 77-78. (48) Wehtje, E.; Adlercreutz, P. Water Activity and Substrate Concentration Effects on Lipase Activity. Biotechnol. Bioeng. 1997, 55, 798-806. (49) Wiebe, R.; Gaddy, V. L. Vapor Phase Composition of Carbon Dioxide-Water Mixtures at Various Temperatures and at Pressures to 700 Atmospheres. J. Am. Chem. Soc. 1941, 63, 475-477. (50) Zacharis, E.; Omar, I. C.; Partridge, J.; Robb, D. A.; Halling, P. J. Selection of Salt Hydrate Pairs for Use in Water Control in Enzyme Catalysis in Organic Solvents. Biotechnol. Bioeng. 1997, 55, 367-374.

Received for review December 30, 1997 Revised manuscript received April 9, 1998 Accepted April 9, 1998

IE970933C