Biotechnol. Prog. 1993, 9, 234-241
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ARTICLES Optimization of Baker’s Yeast Alcohol Dehydrogenase Activity in an Organic Solvent Fangxiao Yang and Alan J. Russell**tJ Department of Chemical Engineering, University of Pittsburgh, Pittsburgh, Pennsylvania 15261, and Center for Biotechnology and Bioengineering, University of Pittsburgh, Pittsburgh, Pennsylvania 15261
Alcohol dehydrogenase from baker’s yeast (YADH) has been used to oxidize an unsaturated alcohol, 3-methyl-2-buten-1-01 (UOL), to the corresponding unsaturated aldehyde, 3-methyl-2-butenal (UAL), in heptane. The influence of total system water content, coenzyme NAD to enzyme molar ratio prior to catalyst preparation, and enzyme concentration on the conversion and redox reaction rates were investigated. The optimal water content was found to be 0.25-0.5% (viv), and a molar ratio of NAD to enzyme of 2.15 is recommended on the basis of economic and kinetic data. A maximal initial reaction rate for YADH of 0.02 mMlminlmg was observed under optimal conditions. The minimum cofactor turnover number was also determined under all conditions, and the effect of doping the catalyst with extra binding sites for NAD was investigated. The effect of substrate on enzyme stability has also been investigated. Interestingly, when the enzyme is preincubated in heptane, its initial activity can be increased 3-fold.
Introduction Dehydrogenation and hydrogenation reactions are of great importance in industry (2). In particular, the interconversion of unsaturated aldehydes from unsaturated alcohols has a special significance in commercial polymer and fine chemicalssynthesis (24). Unfortunately, active commercial metal catalysts often lack selectivity between C=C and C=O, and thus there is a need to develop alternative strategies. Enzymatic dehydrogenation reactions have potential application because of the extreme selectivity of proteins. Preparative production of optically active alcohols and ketones using alcohol dehydrogenases (ADH) has been carried out by a number of investigators (7, 9). Given the need to work with substrates that are insoluble in water, investigators have studied oxidoreductases suspended in organic solvents. Another significant advantage which this approach permits is that enzymes are insoluble in organic media, enabling easy recovery and reuse (11, 12,211. Economics, however, does not look favorably on oxidoreductase processes because of slow reaction rates, low product concentrations, short-livedcatalysts, and the high cost of the enzyme and cofactor. In order to overcome these drawbacks, the use of alcohol dehydrogenases in nonaqueous media has been evaluated by suspending the enzyme power directly in the organic solvent (28), immobilizing the enzyme on a solid support and suspending that in the solvent ( 5 , 7,8, 26), dissolving the enzyme in the aqueous core of a microemulsion (14,23), and crosslinking the enzyme to porous particles followed by suspension in the substratevapor phase (20). The problem of efficient cofactor recycling has been a challengefor many * Author to whom correspondence should be addressed at the Department of Chemical Engineering. Department of Chemical Engineering. 1 Center for Biotechnology and Bioengineering. +
years. There are many strategiesfor nicotinamide cofactor regeneration in aqueous media. These strategies can be classified into four general categories: enzymatic, electrochemical, chemical and photochemical, and biological. The efficiency for cofactor recycling can be expressed in total turnover number (TON,f). A practical goal is the achievement of 1000-100 000 synthetic cycles for each cofactor molecule (1, 3). Enzymes and a sacrificial cosubstrate can be utilized for cofactor recycling as described previously, and as indicated in the reaction scheme we utilize (3,15). Typical turnover numbers are in the range of 100-400 (15), although optimization of reactor configuration has been reported to yield values as high as 400000 (16, 17). Because both enzymes and nicotinamide cofactors are insoluble in nearly all organic solvents, most methods employed in aqueous media for cofactor recycling are not suitable for the same purpose in organic media. Enzymatic recycling in the presence of a second substrate, however, is still successful for cofactor turnover optimization. Klibanov et al. ( 7 ) employed immobilized HLADH for the reduction of 2-methylvaleraldehyde in ethyl acetate and reported a TON,fof greater than 1000 000 after a 7-day reaction. Other systems are reported to give values ranging from 64 to 300 (1, 3, 1517).
In recent years, many articles have been published on the catalytic properties of horse liver alcohol dehydrogenase (HLADH), both in aqueous and in organic solvents. Tapia (27) and Ptitsyn and co-workers (19) have studied the mechanism of HLADH. Sadana and co-workers (22) have proposed a three-state deactivation model based on a consideration of the influence of various parameters, including temperature, pH, immobilization, chemical modifier (inhibitor or protector), and substrate and metal ion concentration, on the kinetics of inactivation. Gorisch has employed adenosine monophosphate (AMP)to protect
8756-7938/93/3009-0234$04.00/0 0 1993 American Chemical Society and American Institute of Chemical Engineers
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Biotechnol. Prog., 1993,Vol. 9,No. 3
Scheme I. Reaction Network
Table I. Relative Quantities of YADH and Cofactor Prior to Lyophilization enzyme. mg
NAD X lo2,mg 10 5 2.5 1.25 0.5 0.25 0.125 0.025
NADienzyme molar ratio 4.25 2.13 1.06
0.53 0.21 0.106 0.053 0.011
dehydrogenase
free HLADH and immobilized preparations, thus increasing the stability of the enzyme (6). Studies on HLADH behavior in organic solvents containing different amounts of added water have been performed by many investigators (5,7,8,26).Most of these studies employed immobilized HLADH. An extremely detailed study on the effect of added water on HLADH properties in organic media, at the molecular level, was carried out by Clark and coworkers, using spin-labeling techniques to probe active site changes as a function of added water (4, 25). The cost of HLADH is so high, however,that it prevents commercialization with the enzyme. It has been reported that an alternative enzyme from yeast (YADH), which is 1%of the cost of HLADH, is far less active and less stable and, thus, unsuitable (26). Indeed, compared to studies on HLADH, there is little information about the catalytic properties of YADH in organic solvents (5,261. However, the low cost of YADH, relative to HLADH and the NADPdependent alcohol dehydrogenase from the thermophilic bacterium Thermoanerobium brockii (10, 13), was a driving force in the optimization of the activity and stability of this enzyme in an important reaction. The work presented in this article discusses the optimization of reaction conditions for the oxidation of 3-methyl-2-butenl-ol (UOL) to 3-methyl-2-butenal (UAL) catalyzed by alcohol dehydrogenase from baker's yeast (YADH) in heptane.
and p-nicotinamide adenine dinucleotide (NAD) were obtained from Sigma Chemical Co. (St. Louis, MO). All organic chemicals were purchased from Aldrich Chemical Co. (Milwaukee,WI). All chemicalsused were of analytical grade. Enzyme Lyophilization. Typically, 5 mg of enzyme powder was dissolved in 1mL of 0.01 M phosphate buffer, pH 7.8. After the addition of 1 mL of NAD solution containing varying amounts NAD, in the same buffer, this solution was lyophilized for 48 h. The various amounts of NAD and the molar ratios of NAD to enzyme are listed in Table I. Water Content Measurements. The water contents of heptane and lyophilized enzyme were tested using Karl Fisher titration in a Fisher Coulomatic K-F titrimeter (Model 447). The "pure" heptane contained 0.0029% (v/ v) water, and the saturated water content for heptane is 0.0085% (viv). In the presence of 0.25 M acetone and 0.25 M UOL, however, the saturated water content for heptane as bulk solvent was increased to 0.090% (v/v). Lyophilized enzyme powder (50 mg) was dissolved in 5 mL of dimethylsulfoxide (DMSO),and the water contents of the neat DMSO and the enzyme-DMSO solutions were measured. The water concentration difference between the enzyme solution and DMSO was taken as the water content of enzyme. The water content of the lyophilized enzyme powder varied from 15 to 20% (w/w). Activity Measurements. Substrate solution (2 mL; 0.25 M 3-methyl-2-buten-1-01 and 0.25 M acetone in heptane) was added to a 4-mL reaction vial containing 5
Materials and Methods Materials. Alcohol dehydrogenase (EC 1.1.1.1)from baker's yeast (YADH, once crystallized and lyophilized) I I
120.0
acetone
isopropanol
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+
100.0
&2.5 m g / d YADH fresh substrate -025 @/mlwater added
+100mMIPOH + 150mM UOL + 150 mM acetone
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73 32
60.0
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250 mM UOL + 250 mM acetone + 2.5 pi/rnl water
0.0 0
I
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20
40
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F i g u r e 1. YADH-catalyzed interconversion of UAL, UOL, IPOH, and acetone at 30 "C and 300 rpm in heptane.
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100.0
*
02
0
1:
0
1
0.05
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F i g u r e 2. Effect of total system water content on Ui, 71, and UAL,,,,,, a t 30 "C and 300 rpm, with 5 mg of YADH, 0.1 mg of NAD, 0.25 M UOL, and 0.25 M acetone in heptane.
mg of enzyme-NAD lyophilized powder. The suspension was sonicated for 20 s, and then different amounts water were added if appropriate. The suspension was shaken a t 300 rpm and 30 "C. Periodically, 0.2-pL samples were withdrawn and analyzed with a gas chromatograph (GC). Heptane was selected as the organic solvent because of its suitability for biocatalytic reactions, its low price, and its low volatility. Product Analysis. A Hewlett-Packard Series I1 5890 GC was used to analyze the products, 3-methyl-2-butenal (UAL), and isopropyl alcohol (IPOH). A column (6 f t X l/8 in., stainless steel, 60/80 Carbopark B/1% SP 1O00, Supelco, Inc., Belletonte, PA) and FID detector were used. The temperatures for oven, injector, and detector were 180,200, and 200 O C , respectively. The carrier gas was helium, and the flow rate was 30 mL/min.
- ;
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Results and Discussion Enzyme Activity. Using the cofactor recycling system described above and in Scheme I, we first attempted to measure the apparent equilibrium concentration for this system, that is, the point a t which no further addition of enzyme will result in increased concentration of products (or reactants). Figure 1demonstrates that when UAL = 67 mM, UOL = 183 mM, IPOH = 67 mM, and acetone = 183mM, the system is a t equilibrium. Thus, the apparent equilibrium constant (Ka,app) for this set of reactants/ products, a t the temperature and initial concentrations given, is0.134. We should note that, since this equilibrium constant is actually the product of the two individual reaction equilibrium constants, it could vary with altered reaction conditions. To ensure that these concentrations represent equilibrium, we initiated a reaction containing >70 mM UAL (see Figure 1)and measured the approach of the system to equilibrium. Also apparent in Figure 1 is that a single batch of enzyme ( 1, activity increases substantially with increasing CIE. However, when C/E 1 could result in low TON,f. When C/E < 1, the enzyme is not significantly more stable than for C/E > 1(enzyme lifetime does not change significantly), and the activity of the enzyme does not decrease proportionally with the reduced level of cofactor (activity at C/E of 0.01 is much more than 1% of that for CIE > 1). Thus, the enzyme that is active (bound to cofactor) with restricted cofactor availability must be more active than enzyme-cofactor complexes formed in the presence of excess cofactor. Our data can be explained in two ways. Either the cofactor remains associated with a particular YADH molecule, each of which is for some reason more active, or the cofactor must dissociate away from the enzyme after product release, and this release both activates the enzyme and stabilizesthe cofactor. In the latter case, the increase in specific activity could be explained by the increased stability of cofactor when bound in one of the many free active sites, rather than solubilized in the water layer surroundingthe enzyme. This suggests a possible method to increase TON,,,, while maintaining a ui more characteristic of preparations where CE > 1. What will happen if we colyophilize YADH with another protein which can bind cofactor, but not substrate? To address the question posed above, we lyophilized YADH in the presence of lactate dehydrogenase under conditions where the NADiYADH molar ratio was 4.25, whereas the NAD/[LADH+YADHl ratio was 0.25. Unfortunately, the amount of LADH necessary to achieve a ratio of 0.01 prevents the use of very low ratios. When the total NAD binding sites for the enzyme preparation are increased in this manner, the activity of the enzyme increases 3-fold (1.42-7.43 IU/mg X lo2),as does the TON,f (114-355). The effect was not a result of differing water contents between the doped and undoped preparations, since a change the amount of water in the solvent did not suppress the activation of the sample via doping with excess NAD binding sites. In addition, LADH has no activity on
Biotechnol. Prog., 1993,Vol. 9,No. 3
238
Table 111. Maximal Values for [UAL],,,, Enzyme Lifetime, and Initial Rates maximal value Figure section [UALI,,, (mM) lifetime (h) TONCi u , (IU x 102/mg) 2 46.7 57 3.96 X lo2 2.99 3 O.l@c Hi0 8.13 57 83.9 x 102 0.51 3 0.25% H?O 46.8 69 115 X lo2 2.27 3 1.0"r H?O 28.5 29 66.3 X lo? 2.99 5 0 rpm 32.9 83 1.50 5 50 rpm 42.1 83 2.19 5 100 rpm 52.1 84 2.54 5 200 rpm 43.9 76 2.86 5 250 rpm 47.5 76 3.07 5 300 rpm 50.0 76 3.68 6 3.68 ' I The values listed in this table enable one to calculate the actual values of all data presented as percentages in other figures.
these substrates (data not shown),and doping YADH with subtilisin (a protein which does not bind NAD(H)) in the same molar ratios has no activation effect on catalyst activity or stability. Table I1 summarizes these experiments. Interestingly, if the LADH enzyme powder is added directly to the reaction mixture rather than colyophilized with YADH, there are still small increases in the stability and activity of the enzyme. This is further evidence that hydrated enzyme particles in organic solvents will aggregate (30). Indeed, YADH and NAD, which have been lyophilized separately, exhibit activity in the reaction we are studying when both are added to the reaction mixture (see Table 11). Although this has not been observed for HLADH (5), the result is not surprising. The hydrated cofactor and enzyme particles aggregate in hydrophobic solvents, and the particle is then equivalent to a YADHi NAD particle after the first turnover of substrate to product (product and cofactor release are mechanistically required for turnover in water). Since it has already been established that there is multiple turnover, the cofactor must be able to dissociate away from the enzyme active site into the associated water layer (the cofactor cannot be solubilized by heptane and would not be expected to leave the environ of the enzyme particle). A schematic representation of our proposed hypothesis for how cofactor
is recycled by YADH is given in Figure 8. Cofactor stabilization is of much interest, and it is clear that in the system we describe, strategies to improve NAD lifetime should be based on the assumption that the cofactor is not static within the enzyme active site. Indeed, the stabilization of cofactor by the addition of cofactor binding sites, referred to above, would be the result of the interaction between cofactor in sections 3 and 6 of Figure 8 with the non-YADH cofactor binding sites. Table I11 describes the maximal values for [UAL],,,, enzyme lifetime, and initial rate for each figure. These data enable one to calculate actual values, in addition to being able to determine trends within the data. Enzyme Stability. While activity is important in determining the productivity of a catalyst, the scale-up of a reaction will depend on protein stability. It is known that aldehydes can react with free lysine groups on the surface of enzymes, and as such biocatalytic processes utilizing alcohol dehydrogenase, in either aqueous or nonaqueous systems, may be limited by denaturation of the protein. We have measured the stability of YADH/ NAD particles as a function of catalyst water content, substrate, and method of enzyme preparation. As described above, most of the reactions we have studied do not utilize sufficient enzyme to reach equilibrium, and therefore the stability of the catalyst will be related to T I
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Incubation Time &s) Figure 4. Effect of incubation time and water content on stability of YADH in heptane. Activity is measured a t 0.25 M acetone, 0.25 M UOL, 2.5 mg/mL YADH, and 0.025 mg/mL NAD a t 30 "C and 300 rpm.
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Biotechnol. Prog., 1993, Vol. 9, No. 3
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and [UAL],,,. This measure of stability (described in detail in the previous section) is relevant for active enzyme, that is, enzyme in the presence of aldehyde. Interestingly, once the enzyme has lost activity, such as that enzyme described by Figure 1 after 54 h, much of the enzyme cannot be resolubilized in aqueous media. This indicates that the enzyme has been denatured or chemically modified (presumably by the aldehyde). A more classical method for monitoring enzyme stability is to preincubate the biocatalyst in solvent and then measure activity at increasing times of solvent exposure.
The results of such an experiment are presented in Figure 4. The most striking fact is that after 144 h of incubation in heptane YADH can still retain 100% initial activity, depending on water content. When the same enzyme catalyzes a reaction in heptane, it ceases to function after only 50 h. The trends for the effect of solvent incubation, in the absence of substrate, on activity, enzyme lifetime (data not shown), and [UALI,,, (data not shown) emphasize that it is either the substrate or product of the active enzyme which causes the irreversible inhibition described in Figure 1and that the enzyme itself is relatively stable at all water contents. A closer look at the data for activity in Figure 4 reveals that preincubation in heptane can actually activate the enzyme. For 0.25 and0.1% water contents, the ui is more than doubled by preincubation for 10 h. There are a number of possible mechanisms for this activation phenomenon (which is 20 times greater than experimental error). The most plausible explanation is related to slow morphological changes in the enzyme particle during preincubation, which in turn could affect the activity/ stability of the catalyst and/or the mass-transfer characteristics of the catalyst. We have seen a similar activation effect for a number of other enzymes in organic solvents, and thus we hypothesize that the cause is related more to the use of powdered suspensions of enzymes than to a specific effect of solvent on YADH. One would expect that a change in morphology would be affected by varying the degree of agitation during incubation. Figure 5 and 6 describe how the preincubation activation was affected by altering the shaking speed during incubation and then determining whether preincubation could still activate the enzyme. Surprisingly, with no external agitation the reaction can still proceed, although the rate of reaction is dependent on the time of preincubation (Figure 6,O rpm data). Only at >250 rpm is the enzyme activated by preincubation in heptane. Once again, the trends in the effect of preincubation on enzyme lifetime and [UALI,,, mirror those in ui (Figure 5). Experiments with other substrates (see discussion following for allyl alcohol) show that much higher activities are possible for YADH. Thus, the reaction being studied here is unlikely to be diffusionally limited, and enzyme activation caused by altered mass-transfer properties in the catalyst cannot be supported by the data. Rather, the activation effect (which we have not attempted to optimize) is more likely to be a direct effect of solvent on the enzyme particles, and the effect is dependent on the degree of agitation because that will also alter morphology and thus the degree of interaction between enzyme and solvent. Given the sensitivity of YADH to its substrate-product mixture, we tested another pair of substrates for their inhibitory effects in an identical experiment. Using allyl alcohol and butyraldehyde as substrates, the activity of the enzyme was determined as a function of water content (Figure 7). Although the initial activity (measured either as activity or [acrolein]),,, of the enzyme far exceeds that for UOL and acetone, the enzyme is inactivated rapidly as indicated by short enzyme lifetimes (data not shown). The details of YADH activity and stability with this substrate pair will be published in a separate article. Clearly, these results indicate that YADH stability in heptane is related to substrate/product concentration and direct effects of the solvent on the morphology of the enzyme particle. Scale-up and optimization of processes utilizing YADH will have to address these issue.
Bbt8&?d. Prog.., 1993, Vd. 9,
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Figure 6. Effect of shaking speed on activity of YADH as a function of preincubation in heptane. Activity is measured at 0.25 M acetone, 0.25 M UOL, 2.5 mg/mL YADH, and 0.025 mg/mL NAD at 30 "C. 90.0
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Conclusions
pi .................. ......... ..... ..... ..... ...... ... ....... .......... ........ ....... ...... ....... .. ....... .. ....... ... ....... ........ ... .. ........ ......... .... ......... ................ .................. e . .
For the baker's yeast alcohol dehydrogenase catalyzed oxidation of 3-methyl-2-buten-1-01to 3-methyl-2-butena1, with concomitant acetone reduction to isopropyl alcohol, in heptane, the optimalactivityand maximal final product concentration were observed at an added water concentration of 0.25-0.5 % (v/v)for the amount of enzyme added. The data also show that the optimal molar ratio of coenzyme NAD to enzyme is approximately 2.15, on the basis of the need for high activity and conversion. Rates of reaction with YADH were on the order of 0.02 mM/ min/mg, which compared favorably to those with horse liver alcohol dehydrogenase (which costs 300 times more than YADH) and the NADP-dependent alcohol dehydrogenase from Thermoanaerobium brockii (which costs 400 times more than YADH). We have also demonstrated that, during YADH catalysis in heptane, the cofactor dissociates from the enzyme (see Figure 8) and that the stability of cofactor (TONd) can be increased by providing extra cofactor binding sites. Indeed, while activity is
..... ..-.. ..... ....... ... ...... ........ ...... ...... ... ...... ....... ... ...... ....... .. ... ....... .......... .... ........ .............. ................ .................. - . I
Figure 8. Schematic representation of cofactor recycling by YADH suspended in organicsolvents. Notethe releaseof cofactor from the enzyme active site in steps 3 and 6. Details are given in the text.
maintained a t an optimal level (C/E> l),TONd can be increased in the presence of lactate dehydrogenaseto levels characteristic of C/E < 1. Finally, the stability of the enzyme in heptane is dependent on the substrate being utilized and can be increased significantlyby preincubation of the suspended enzyme in substrate-free solvent.
Biotechnol. Prog., 1993,Vol. 9,No. 3
Acknowledgment We thank Professor Donna Blackmond, Akshay Waghray, and Ron Mahlab for their help in this work. This work has been funded by PPG, Union Carbide Corporation, and a National Science Foundation Presidential Young Investigator Award to A.J.R. (BCS 9057312). Literature Cited (1)Bowen, R.; Pugh, S. Redox Enzymes in Industrial Fine Chemicals Synthesis. Chem. Znd. (London) 1985, (No. 10, May 20), 323-326. (2) Caroling, G.; Rajaram, J.; Kuriacose, J. C. Selective Oxidation
of Unsaturated Alcohols and Primary Alcohols by RuC1:dNMethylmorpholine N-oxide (NMO) System: A Kinetic Study. J . Mol. Catal. 1990. 58. 235-243. (3)Chenault, H. K.; Whitesides, G. M. Regeneration of Nicotinamide Cofactors for Use in Organic Synthesis. Appl. Biochem. Biotechnol. 1987, 14, 147-197. (4) Clark, D. S.; Skerker, P. S. Randolph, T. W.; Blanch, H. W.; Prausnitz, J. M. Conformational Analyses of Enzymes and Substrates in Processing Environments. Ann. N . Y.Acad. Sci. 1988, 542, 16-29. (5) Deetz, J. S.; Rozzell, J. D. Catalysis by Alcohol Dehydrogenase
in organic Solvents. In Biocatalysts for Industry; Dordick, J. S., Ed.; Plenum Press: New York, 1991; Chapter 9. (6) Gorisch, H.; Scheider, M. Stabilization of Soluble and Immobilized horse Liver Alcohol Dehydrogenase by Adenosine 5’-Monophosphate. Biotechnol. Bioeng. 1984,26, 998-1002. (7) Grunwald, J.; Wirz, B.; Scollar, M. P.; Klibanov, A. M. Asymmetric Oxidoreductions Catalyzed by Alcohol Dehydrogenase in Organic Solvents. J . Am. Chem. SOC.1986, 108, 6132-6734. (8) Guinn, R. M.; Skerker, P. S.; Kavanaugh, P.; Clark, D. S.
Activity and Flexibility of Alcohol Dehydrogenase in Organic Solvents. Biotechnol. Bioeng. 1991, 37, 303-308. (9) Jones, J. B. Enzymes in Organic Synthesis. Tetrahedron 1986, 42, 3351-3403. (10) Keinan, E.; Hafeli, E. K.; Seth, K. K.; Lamed, R. Thermostable Enzymes in Organic Synthesis. 2. Asymmetric Re-
duction of Ketones with Alcohol Dehydrogenase from Thermoanaerobium brockii. J . Am. Chem. SOC.1986, 108, 162169. (11) Klibanov, A. M. Enzymatic Catalysis in Anhydrous Organic Solvents. Trends Biochem. Sci. 1989, 14, 141. (12) Klibanov, A. M. Enzymes That Work in Organic Solvents. CHEMTECH 1986 (June), 354. (13) Lambed, R. J.; Keinan, E.;Zaikus, J. G. Potential Application
of an Alcohol-AldehydeiKetone Oxidoreductase from Thermophilic Bacteria. Enzyme Microb. Technol. 1981, 3, 144148. (14) Larsson, K. M.; Adlercreuts, P.; Mattiasson, B. Studies on
Horse Liver Alcohol Dehydrogenase in a Microemulsion System. Ann. N.Y. Acad. Sci. 1990, 613, 791-795.
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(15) Lee, L. G.; Whitesides, G. M. Enzyme-Catalyzed Organic
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Accepted February 1, 1993.
