Asymmetric reductions in aqueous media: Enzymatic synthesis in cyclodextrin containing buffers

Article abstract of DOI:10.1016/S0957-4166(99)00137-8

The enzymatic reduction of hydrophobic ketones in cyclodextrin containing media is reported yielding the corresponding alcohols (S)-1-(2- naphthyl)-ethanol, (S)-(E)-4-phenyl-3-en-2-ol and 1,2,3,4-tetrahydro-2- (1hydroxyethyl)-1-oxonaphthalene in good yiel

Full text of DOI:10.1016/S0957-4166(99)00137-8

Pergamon
Tetrahedron: Asymmetry 10 (1999) 1681–1687
Asymmetric reductions in aqueous media: enzymatic synthesis in
cyclodextrin containing buffers
Thomas Zelinski,a Andreas Liese,b Christian Wandrey b and Maria-Regina Kula a,∗
aInstitute of Enzyme Technology, Heinrich-Heine-Universität Düsseldorf, D-52426 Jülich, Germany
bInstitute of Biotechnology 2, Forschungszentrum Jülich GmbH, D-52425 Jülich, Germany
Received 4 March 1999; accepted 9 April 1999
Abstract
The enzymatic reduction of hydrophobic ketones in cyclodextrin containing media is reported yielding
the corresponding alcohols (S)-1-(2-naphthyl)-ethanol, (S)-(E)-4-phenyl-3-en-2-ol and 1,2,3,4-tetrahydro-2-(1-
hydroxyethyl)-1-oxonaphthalene in good yield and high enantiomeric excess. The reaction was catalyzed by a
carbonyl reductase from Candida parapsilosis, NADH regeneration was carried out using either 2-propanol in a
substrate coupled mode or formate dehydrogenase from Candida boidinii and formate as hydrogen donor. In con-
tinuous experiments high stability of the enzymes in media containing heptakis-(2,6-di-O-methyl)-β-cyclodextrin
could be demonstrated. Space time yields of 120 g/(Ld) were obtained under not yet optimized conditions during
the production of (S)-1-(2-naphthyl)-ethanol. © 1999 Elsevier Science Ltd. All rights reserved.
The enzymatic conversion of hydrophobic compounds, poorly soluble in water, presents a unique
challenge in the production of fine chemicals.1,2 Addition of water miscible solvents to increase the
solubility of educts is problematic, as many enzymes are destabilized by changes in the dielectric constant
and perturbation of the water structure.3 Lipases in their natural environment act at aqueous–lipid
interfaces and may be employed with advantage in biphasic reaction media in the presence of a water
immiscible solvent.4–6 For dehydrogenases, which often denature at such interfaces, a bi-membrane
reactor7–10 and an emulsion reactor11 have been designed to facilitate the transfer of the educt into
the aqueous phase and the removal of the product after conversion. Application of enzymes in nearly
anhydrous media have been reported,12–14 but the reaction rate is usually much reduced and problems
arise if the pH needs to be adjusted in the course of the reaction or coupled reactions are desired.
Another possibility to increase the apparent solubility of hydrophobic compounds is the application of
cyclodextrins in homogeneous aqueous media.
Cyclodextrins are torus-shaped cyclic maltooligosaccharides of six (α), seven (β) or eight (γ) D-
glucose units linked by α-1,4-glycosidic linkages.15 By intramolecular hydrogen bonds between the
glucose units an apolar cavity is formed, which accepts hydrophobic guest molecules.16 The hydrophilic
∗
Corresponding author. E-mail: m.-r.kula@fz-juelich.de
0957-4166/99/$ - see front matter © 1999 Elsevier Science Ltd. All rights reserved.
PII: S0957-4166(99)00137-8

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shell of the torus maintains solubility in water. The hydrolysis of deacetyl-lanatoside A to digitoxin by β-
glucosidase17 and the hydrolysis of p-nitrophenylbutyrate as well as triolein or olive oil by lipases18 was
enhanced by addition of cyclodextrins, especially heptakis-(2,6-di-O-methyl)-β-cyclo-dextrin (DIMEB).
The enzymatic oxidation of steroids by cholesterol oxidase19 or phenoloxidase20 in the presence of
cyclodextrins has been studied as well.
We recently analyzed, in detail, the enzymatic reduction of 2-acetylnaphthalene 1c in cyclodextrin
containing media using a carbonyl reductase from Candida parapsilosis (CPCR) and formate dehydro-
genase (FDH) from Candida boidinii for in situ NADH regeneration.21 DIMEB at 100 mM concentration
enhanced the apparent solubility of 1c about 150 fold. The reaction velocity of the coupled reaction
was unaffected by the presence of DIMEB, indicating that the kinetics of dissociation of the preformed
complex between 1c and DIMEB is not rate limiting.
Since the production of chiral alcohols is of high interest we extended our studies to other substrates
of CPCR and explored enzymatic reductions in continuous processes using different modes of NADH
regeneration, as illustrated in Fig. 1.
CPCR was originally screened as an NADH dependent β-ketoester-reductase.22 Later it was shown,
that the enzyme has a very broad substrate range and accepts, among other carbonyl compounds,
diketones and aromatic ketones as substrates.23,24 Due to the alcohol dehydrogenase activity of the
CPCR, NADH regeneration may be carried out in a substrate coupled reaction (line I, Fig. 1) using
isopropanol as hydrogen donor.25 Besides, the enzyme coupled mode (line II, Fig. 1) was employed with
FDH as a second enzyme and formate as the ultimate hydrogen donor. Table 1 summarizes the results
obtained for batch conversions in preparative scale.
Figure 1. General reaction scheme
In all cases high conversions were reached after several days as judged by TLC. Only in the case
of 2c, two weeks were needed for the conversion due to the low enzyme concentrations applied. The
corresponding alcohol was extracted with hexane and/or ether, respectively, and purified by column
chromatography over silica before further analysis. On a preparative scale 2c was obtained in high ee
(>99%) and 70% isolated yield. The enzymatic reduction of the ketones 1a and 1b by CPCR is reported
for the first time. The (S)-alcohol 2b has been obtained before by enantioselective esterification of the
corresponding racemate in 32% yield and ee >95% using Pseudomonas lipase.26 High chemoselective
catalytic hydrogenation (>99.9%) of the carbonyl group of 1b was achieved by Noyori et al. using a
RuCl2 (binap)(dmf)n-diamine–KOH system.27 Stereoselectivity was only moderate, yielding 70% ee of

T. Zelinski et al. / Tetrahedron: Asymmetry 10 (1999) 1681–1687
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Table 1
Enzymatic reduction of aromatic ketones yielding S-alcohols
the (R)-enantiomer. Using an oxazaborilidine catalyst, Corey and Helal28 reported 97% ee and 90% yield
for the (R)-enantiomer of 2b performing the reaction at −78°C in toluene. Based on the known sign of the
specific rotation the product obtained by reduction with CPCR is the (S)-enantiomer, which was isolated
with an ee of >99% as determined by gas chromatography. By NMR analysis it was recognized that the
isolated product 2b contained about 5% of the saturated alcohol 4-phenyl-2-butanol. Analysis of the educt
employed showed that 1b contained about 5% of the corresponding saturated ketone, which evidently is
also a substrate for CPCR. The ee of the saturated alcohol was not determined, but it is expected that this
reduction by the CPCR will also be performed at a very high level of enantioselectivity.
To the best of our knowledge, regioselective reduction of 1a in the side chain has never been achieved
before. From NMR data it is evident that the cyclic ketone group remained intact. An earlier detailed
kinetic study of the CPCR revealed a small and a large hydrophobic binding pocket of the enzyme
differing in volume and affinity towards potential substrates. The small binding site accepts only alkyl
groups (preferentially a methyl group) while the presence of an aromatic ring in the large binding
pocket led to low KM values and good reaction rates.24 Based on these results we conclude that the
high regioselectivity is a steric effect directing the positioning of the substrate 1a in the binding site.
We assume that the product of the enzymatic reduction has the (S)-configuration, in accordance with
all other products obtained so far using CPCR. Definite proof is expected from structure determination
by X-ray analysis. The product was isolated as an oil and could not be crystallized yet. The latter may
be related to the fact that two diastereomers were obtained in the ratio of 2:1 as determined by NMR.
During the regioselective reduction of the diketone a second stereogenic center at C2 is generated. The
composition of the stereoisomers could not be determined, although the silylated, acetylated and non-
derivatized products were analyzed on different GC phases.†
The reduction of ethyl-2-methyl-3-oxo-butanoate with CPCR led to the syn-product with (2R,3S)-
configuration in diasteromeric excess of 98%.29 With the diketone 1a the diasteromeric discrimination is
less pronounced, which may reflect a slower reracemization step at C2.
Continuous reductions of 2-acetylnaphthalene were performed to demonstrate the feasibility of ope-
rating a membrane reactor with DIMEB containing reaction media, to generate reliable data on enzyme
stability during prolonged times and to test the different modes of coenzyme regeneration under these
conditions. Fig. 2 illustrates the conversion–time course of the reaction over 120 h under various
residence times using FDH for coenzyme regeneration. The reactor was charged with 4.6 U/mL CPCR
and 4.4 U/mL FDH. After about 10 h a steady conversion of 52% was reached at a residence time τ of 1 h,
representing a space time yield of 43 g/(Ld). Increasing τ to 2 h a new steady state was established at 79%
†
Tested GC phases: Lipodex B and Lipodex E from Macherey & Nagel (Düren, Germany), Cyclodex β 1/P, α-2P, γ-1P,
γ-Lactone, β-2P from Chromatographie Service (Langerwehe, Germany) and Chiraldex β-PH, GTA and BP-H from Astec
(Whippany, NJ, USA).

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Figure 2. Continuous reduction of 2-acetylnaphthalene (20 mM) in 100 mM TEA buffer pH 7.0 containing 0.5 mM NAD+,
1 M sodium formate, 2 mM DTT and 50 mM DIMEB. Initial catalyst/concentration 4.6 U/mL CPCR and 4.4 U/mL FDH
conversion and maintained at this level for 40 h. The space time yield dropped to 33 g/(Lh). Reducing τ
to 0.5 h the conversion dropped to 29%. After 81 h of operation, start conditions were re-established and
a new steady state at 46% conversion reached. From the drop in conversion a deactivation of the enzymes
of 1.2% per day can be estimated demonstrating that both enzymes are very stable in DIMEB containing
reaction media.
Increasing the ratio of CPCR activity to total enzyme activity should result in a higher conversion
being reached,23 which was not intended here, however a lower ratio was chosen in order to see the
enzyme deactivation more clearly. The small decay in conversion does not distinguish whether CPCR,
or FDH, or both deactivated slowly. In control experiments incubation of CPCR in the presence of 0.2
mM 2-acetylnaphthalene in 100 mM TEA buffer pH 7.0 resulted in 15% residual activity of the enzyme
after 24 h. Between 25 and 30% loss of activity can be attributed to oxidation of CPCR in the absence
of reducing agents. The observed high stability in the presence of DIMEB during prolonged operation
in a continuously operated stirred tank reactor may be due to the low concentration of the uncomplexed
ketone, which no longer affects the enzyme stability.
In a second continuous reduction (Fig. 3) the high stability of CPCR under operating conditions was
confirmed. The conversion–time curve is very stable, from the difference in conversion after 20 and
90 h a deactivation of 0.2% per day was determined for CPCR. Using 260 mM 2-propanol (2% v/v)
for NADH regeneration in the substrate coupled mode a maximal conversion of 82% was reached at
τ=2 h. Reducing τ to 0.5 h, 71% conversion was obtained, corresponding to a space time yield of 118
g/(Ld). Under these conditions the productivity was about 2.5 fold higher as observed in Fig. 2 with
NADH regeneration by FDH. Since conditions for both experiments are not yet optimized this result
should not be over interpreted. Nevertheless, it could be demonstrated that in DIMEB containing media
substrate coupled coenzyme regeneration is possible and space time yields well in excess of 120 g/(Ld)
are expected under optimized reaction conditions, especially by increasing the enzyme concentration in
the reactor.
1. Experimental
1.1. Chemicals
DIMEB, heptakis-(2,6-di-O-methyl)-β-cyclodextrin was obtained from Wacker-Chemie (Düsseldorf,
Germany), 2-acetylnaphthalene was purchased from Fluka (Buchs, Switzerland), benzylidenacetone

T. Zelinski et al. / Tetrahedron: Asymmetry 10 (1999) 1681–1687
1685
Figure 3. Continuous reduction of 2-acetylnaphthalene (20 mM) in 100 mM TEA buffer pH 7.0 containing 0.5 mM NAD+, 260
mM 2-propanol, 2 mM DTT, 50 mM DIMEB and 4.6 U/mL CPCR
from Merck (Darmstadt, Germany), 2-acetyl-1-tetralone from Aldrich (Steinheim, Germany) and used
as delivered. NAD+ and NADH were products from Biomol or Oriental Yeast (Tokyo, Japan). All other
chemicals, buffer salts and solvents were obtained in high quality from commercial sources.
1.2. Enzymes
Formate dehydrogenase (FDH) from Candida boidinii was supplied by Dipl.-Ing. Holger Gieren,
Institute of Enzymetechnology (Jülich, Germany) and prepared as described by Weuster-Botz et al.30
Carbonyl reductase (CPCR) from Candida parapsilosis was isolated as described by Peters et al.23 and
partially purified by fractional precipitation with poly(ethylene glycol) and anion exchange chromato-
graphy on Sepharose Q Fast Flow (Amersham Pharmacia, Freiburg, Germany) to a specific activity of
∼16 U/mg in the standard assay system as described by Zelinski and Kula.21 CPCR was stored in buffer
solution containing 50% glycerol, 1 mM DTT at −20°C.
1.3. Analytical procedures
NMR spectra were obtained on a Varian VXR 300 at 300 MHz (1H) and 75 MHz [13C] shifts
are reported as ppm relative to TMS as internal reference. Abbreviations: m=multiplet, d=doublet,
s=singulet, br=broad Perkin Elmer model 241 MC polarimeter with a 10 cm path length cell was used to
determine [α]D. Gas chromatographic analysis was carried out using equipment (GC9A) from Shimadzu
(Düsseldorf, Germany) and flame ionization for detection. In general, separation was performed on
a chiral Lipodex-E-γ-Cyclodextrin column (25 m×0.2 mm ID) from Macherey and Nagel (Düren,
Germany) operated with helium as carrier gas and injecting 1 µL samples. The column was thermostated
at 70°C, while injector and detector were kept at 250°C. Thin layer chromatography: Samples (up
to 5 µL) were applied on silica plates 60 F254, Merck (Darmstadt, Germany) and developed with n-
hexane:ethyl acetate (3:1). After drying in a warm air current the plates were viewed under UV light.
Capillary electrophoresis was performed as described by Zelinski and Kula.21
1.4. Batch conversions
Enzymatic transformations of 5.5 mmole 1a, 1b were carried out at room temperature with stirring
(120 rpm) in volumes of 50–300 mL containing 100 mM triethanolamine (TEA) buffer pH 7, 2 mM

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dithiotreitol (DTT), 250 mM sodium formate, 1 mM NAD+, 100 mM DIMEB and 1 U/mL CPCR and 3
U/mL FDH. Reduction of 1c was performed in the same buffer at 20 mM 1c, 1 M sodium/formate, 0.5
mM NAD+, 50 mM DIMEB, at low enzyme concentrations of 0.007 U/mL CPCR and 0.035 U/mL FDH
for two weeks. At the end of the reaction the complete mixture was ultrafiltrated in an Amicon cell using a
YM 10 membrane (Witten, Germany) and the retentate washed with TEA buffer to separate the enzymes.
The filtrates were extracted several times with hexane and/or ether, the solvent dried over anhydrous
sodium sulfate and removed by rotary evaporation. The resulting oils were purified by chromatography
over silica gel 60 (Merck, Darmstadt, Germany) with n-hexane:ethylester (1:4) for development.
1.5. Continuous conversions
A 10 mL thermostated (25°C) enzyme membrane reactor was used as described by Wandrey and
Kula.31 The reactor was equipped with an ultrafiltration membrane YM-10, Amicon (Witten, Germany)
cut of 10 kDa, and operated with a magnetic stirrer at about 150 rpm. The reactor was sterilized before
use percolating a 0.1% solution of peracetic acid for 16 h and then flashed with sterile water before
introducing the substrate solution containing 20 mM 2-acetylnaphthalene, 0.5 mM NAD+, 1M sodium
formate, 2 mM DTT and 50 mM DIMEB in 100 mM TEA buffer pH 7.0 through a microporous
membrane (cellulose nitrate, 0.1 µm pore size, Sartorius AG, Göttingen, Germany). Enzymes were
injected into the inlet stream before passage through the microporous filter. In the case of the substrate
coupled NADH regeneration FDH was omitted and 260 mM 2-propanol added to the reaction medium.
An alternating piston pump (Pharmacia P-500, Freiburg, Germany) was used for constant delivery of
substrate solution to achieve the desired residence time. The microporous membrane in the inlet stream
and the ultrafiltration membrane in the reactor defined and maintained the sterile area. The ultrafiltration
membrane retains the enzymes in the reactor, while buffer and low molecular weight compounds of
the reaction medium can pass. The reactor effluent was collected using a fraction collector. Single
fractions were extracted manually with 0.5 volumes of toluene and educt and product were determined by
analyzing a suitable aliquot by capillary electrophoresis and/or gas chromatography as described above.
1.6. NMR analysis: 1,2,3,4-tetrahydro-2-(1-hydroxyethyl)-1-oxonaphthalene
1H NMR (300 MHz): major: δ 8.01 (d, J=7.7 Hz, 1H, H-8), 7.52–7.54 (m, 1H, H-6), 7.29–7.35 (m, 1H,
H-7), 7.25 (d, J=7.7 Hz, 1H, H-5), 4.39 (m, 1H, H-2), 4.5–4.3 (br, 1H, OH), 3.04 (m, 2H, H-4), 2.67–2.59
(m, 1H, H-9), 2.14–2.28 (m, 1H, H-3a), 2.11–1.98 (m, 1H, H-3b), 1.29 (d, J=7.7 Hz, 3H, H-10), minor:
δ 8.01 (d, J=7.7 Hz, 1H, H-8), 7.52–7.54 (m, 1H, H-6), 7.29–7.35 (m, 1H, H-7), 7.25 (d, J=7.7 Hz, 1H,
H-5), 4.17 (m, 1H, H-2), 3.25–3.1 (br, 1H, OH), 3.04 (m, 2H, H-4), 2.41–2.50 (m, 1H, H-9), 2.14–2.28
(m, 1H, H-3a), 1.74–1.91 (m, 1H, H-3b), 1.29 (d, J=7.7 Hz, 3H, H-10). 13C NMR (75 MHz): major: δ
200.8 (C-1), 144.3 (C-4a), 132.8 (C-8a), 132.8 (C-6), 128.8 (C-7), 127.2 (C-5), 126.6 (C-8), 66.9 (C-9),
53.4 (C-2), 29.0 (C-4), 23.8 (C-3), 19.3 (C-10), minor: δ 202.3 (C-1), 144.3 (C-4a), 133.6 (C-8a), 132.3
(C-6), 128.7 (C-7), 127.4 (C-5), 126.7 (C-8), 68.2 (C-9), 54.1 (C-2), 28.9 (C-4), 25.6 (C-3), 20.2 (C-10).
1
Major to minor diastereomer observed in a ratio of 2:1. (S)-(E)-4-Phenylbut-3-en-2-ol [81176-43-4] H
NMR (300 MHz): δ 7.39–7.18 (m, 5H), 6.56 (d, J=16.0 Hz, 1H), 6.25 (dd, J=16.0 Hz, 8.0 Hz, 1H), 4.48
(m, 1H), 1.77 (s br, 1H), 1.37 (d, J=6.4 Hz, 3H). (S)-1-(2-Naphthyl)ethanol 1H NMR (300 MHz CDCl3):
δ=7.87–7.46 (m, 7H), 5.09 (q, 1H-c-10), 1.76 (brs OH), 1.60 (d, 3H-1C-20) ppm, the data correspond to
literature values.32

T. Zelinski et al. / Tetrahedron: Asymmetry 10 (1999) 1681–1687
1687
Acknowledgements
We thank H. Gieren for the supply of FDH and ‘Deutsche Forschungsgemeinschaft’ SFB 380 for
financial support.
References
1. Enzyme Catalysis in Organic Chemistry; Drauz, K.; Waldmann, H., Eds.; VCH: Weinheim, 1995.
2. Faber, K. Biotransformations in Organic Chemistry, 3rd ed.; Springer-Verlag: Berlin, 1997.
3. Buchholz, K.; Kasche, V. Biokatalysatoren und Enzymtechnologie; VCH: Weinheim, 1997, p. 343.
4. Borgström, B.; Brockmann, H. L. Lipases; Elsevier: Amsterdam, 1984.
5. Brzozowski, A. M.; Derewenda, U.; Derewenda, Z. S.; Dodson, G. G.; Lawson, D. M.; Turkenburg, J. P.; Bjorkling, F.;
Huge-Jensen, B.; Patkar, S. A.; Thim, L. Nature 1991, 351, 491–494.
6. Lawson, D. M.; Brzozowski, A. M.; Dodson, G. G. Current Biology 1992, 2, 473–475.
7. Ikemi, M.; Ishimatsu, Y. J. Biotechnol. 1990, 14, 211–220.
8. Kragl, U.; Kruse, W.; Hummel, W.; Wandrey, C. Biotechnol. Bioeng. 1996, 52, 309–319.
9. Kruse, W.; Hummel, W.; Kragl, U. Recl. Trav. Chim. des Pays-Bas 1996, 115, 239–243.
10. Kruse, W.; Kragl, U.; Wandrey, C. Ger. Pat. Appl. DE P 44.36.149.1, 1994.
11. Liese, A.; Zelinski, T.; Kula, M.-R.; Kierkels, H.; Karutz, M.; Kragl, U.; Wandrey, C. J. Mol. Catal. B: Enzymatic 1998,
4, 91–99.
12. Klibanov, A. M. TIBS 1998, 14, 141–144.
13. Koskinen, A. M. P.; Klibanov, A. M. Enzymatic Reactions in Organic Media; Chapman & Hall: London, 1996, p 314.
14. Biocatalysis in Organic Media, Studies in Organic Chemistry; Laane, C.; Tramper, J.; Lilly, M. D., Eds.; 1987; Vol. 29.
15. Saenger, W. Angew. Chem., Int. Ed. Engl. 1980, 92, 343–361.
16. Clarke, R. J.; Coates, J. H.; Lincoln, S. F. Adv. Carbohydr. Chem. Biochem. 1988, 46, 205–249.
17. Pekic, B.; Lepojevic, Z. Biotechnol. Lett. 1991, 13, 399–404.
18. Otero, C.; Cruzado, C.; Ballasteros, A. Appl. Biochem. Biotechnol. 1991, 27, 185–194.
19. Jadoun, J.; Bar, R. Appl. Microbiol. Biotechnol. 1993, 40, 447–482.
20. Woerdenbag, H. J.; Pras, N.; Frijlink, H. W.; Lerk, C. F.; Malingre, T. M. Phytochemistry 1990, 29, 1551–1554.
21. Zelinski, T.; Kula, M.-R. Biocatalysis Biotransformations 1997, 15, 57–74.
22. Peters, J.; Zelinski, T.; Kula, M.-R. Appl. Microb. Biotech. 1992, 38, 334–340.
23. Peters, J.; Minuth, T.; Kula, M.-R. Enzyme Microb. Technol. 1993, 15, 950–958.
24. Peters, J.; Minuth, T.; Kula, M.-R. Biocatalysis 1993, 8, 31–46.
25. Peters, J. Alcohol dehydrogenases. In Biotechnology, 2nd ed.; Rehm, H. J.; Reed, G., Eds.; Wiley-VCH: Weinheim, 1998;
Vol. 8a, pp. 391–474.
26. Burgess, K.; Jennings, L. D. J. Am. Chem. Soc. 1991, 113, 6129–6139.
27. Ohkuma, T.; Ooka, H.; Ikariya, T.; Noyori, R. J. Am. Chem. Soc. 1995, 117, 10417–10418.
28. Corey, E. J.; Helal, C. J. Tetrahedron Lett. 1995, 50, 9153–9156.
29. Peters, J.; Zelinski, T.; Minuth, T.; Kula, M.-R. Tetrahedron: Asymmetry 1993, 4, 1173–1182.
30. Weuster-Botz, D.; Paschold, H.; Striegel, B.; Gieren, H.; Kula, M.-R.; Wandrey, C. Chem. Ing. Tech. 1994, 17, 131–137.
31. Kula, M.-R.; Wandrey, C. Meth. Enzymol. 1987, 136, 9–21.
32. Brown, H. C.; Chandrasekharan, J.; Ramachandran, P. V. J. Am. Chem. Soc. 1988, 110, 1539–1546