Biocatalytic oxidation of 4-vinylphenol by Nocardia

Article abstract of DOI:10.1139/v02-034

Nocardia species NRRL 5646 stereospecifically hydrates 4-vinylphenol (15) to S-1-(4′-hydroxyphenyl)ethanol (17), and further oxidizes 17 to 4′-hydroxyacetophenone (18). Labeled metabolites 17 and 18 obtained from incubations in D₂O and H₂¹⁸O support initial enzymatic tautomerization of 15 to a reactive quinone methide (16), which adds water in the first reaction. Commitment to catalysis is high in the hydration reaction, while the alcohol dehydrogenation reaction appears to be reversible. The stereochemical features of water addition, alcohol oxidations, and ketone reductions with growing culture biocatalysis were established by chiral HPLC. Alcohol oxidations or ketone reductions in 12 000 × g supernatants preferentially require NADP⁺-NADPH,H⁺ as co-factors. The alcohol dehydrogenase has broad substrate specificity, favoring the oxidation of primary alkanols and 4-hydroxybenzyl alcohols.

Full text of DOI:10.1139/v02-034

582
Biocatalytic oxidation of 4-vinylphenol by Nocardia
Kyung-Seon Lee and John P.N. Rosazza
Abstract: Nocardia species NRRL 5646 stereospecifically hydrates 4-vinylphenol (15) to S-1-(4>-hydroxyphenyl)etha-
nol (17), and further oxidizes 17 to 4>-hydroxyacetophenone (18). Labeled metabolites 17 and 18 obtained from incuba-
tions in D2O and H218O support initial enzymatic tautomerization of 15 to a reactive quinone methide (16), which adds
water in the first reaction. Commitment to catalysis is high in the hydration reaction, while the alcohol
dehydrogenation reaction appears to be reversible. The stereochemical features of water addition, alcohol oxidations,
and ketone reductions with growing culture biocatalysis were established by chiral HPLC. Alcohol oxidations or ketone
reductions in 12 000 × g supernatants preferentially require NADP+–NADPH,H+ as co-factors. The alcohol
dehydrogenase has broad substrate specificity, favoring the oxidation of primary alkanols and 4-hydroxybenzyl alcohols.
Key words : 4-vinylphenol, Nocardia sp., enantiospecific hydration, 1-(4 >-hydroxyphenyl)ethanol, 4 >-hydroxyacetophenone
Résumé : L’espèce Nocardia NRRL 5446 hydrate stéréospécifiquement le 4-vinylphénol (15) en S-1-(4>-hydroxyphé-
nyl)éthanol (17) qu’il oxyde aussi en 4>-hydroxyacétophénone (18). Les métabolites 17 et 18 marqués obtenus par in-
cubation dans D2O et H218O sont en accord avec un mécanisme impliquant une tautomérisation enzymatique initiale de
15 en une méthoquinone réactive (16) qui fixe de l’eau dans la première étape de la réaction. Dans la réaction
d’hydratation, le degré de catalyse est élevé alors que la réaction de déshydrogénation de l’alcool semble réversible.
Faisant appel à la CLHP chirale, on a déterminé les caractéristiques stéréochimiques de l’addition d’eau, des oxyda-
tions de l’alcool et des réductions de cétone par la catalyse de cultures en croissance. Les oxydations d’alcool ou ré-
ductions de cétones dans des liquides surnageant 12,000 x g requièrent le coupe NADP+–NADPH,H+ comme
cofacteurs. La déshydrogénase de l’alcool possède une grande spécificité par rapport au substrat qui favorise
l’oxydation d’alcools primaires et d’alcools 4-hydroxybenzyliques.
Mots clés : 4-vinylphénol, Nocardia sp., hydratation spécifique, 1-(4>-hydroxyphényl)éthanol, 4>-hydroxyacétophénone.
[Traduit par la Rédaction] Lee and Rosazza 588
Introduction
ment for co-factors. They should be robust, stable, and recy-
clable, and ultimately be amenable to large-scale adaptation
to bioreactor use, even under unusual reaction conditions
(3).
Microbial and enzymatic transformations have been devel-
oped for nearly every class of natural and synthetic chemical
products (1). Microorganisms and their enzymes are bio-
catalysts, which are now widely used as “reagents” in syn-
thetic organic chemistry. Biocatalysts are valued for their
intrinsic abilities to bind organic substrates and to catalyze
highly specific and selective reactions under the mildest of
reaction conditions. Biocatalytic transformations of abundant
renewable natural chemicals afford attractive “green chemis-
try” alternatives to synthetic chemical approaches, which
rely on petrochemically derived starting materials.
p-Coumaric acid (4), ferulic acid (5), and caffeic acid are
abundantly occurring phenolic acids commonly found in
plants in their esterified or free-acid forms. The acids p-
coumaric and ferulic compose nearly 4% of the dry weight
of maize plant parts, rendering them available in the billions
of pounds annually from this source alone (5). The large
quantities of these natural compounds render them useful for
the biocatalytic production of value-added aromatic natural
chemicals. Systematic screening of microorganisms showed
that the major and most common biocatalytic reaction with
hydroxycinnamates is decarboxylation to the corresponding
vinylphenols (hydroxystyrenes) (5, 6). Deuterium-labeling
experiments showed that non-oxidative decarboxylations oc-
curred by enzymatic tautomerization of p-hydroxycinna-
mates to doubly vinylogous b-keto acid intermediates that
spontaneously decarboxylated (5, 6a). Phenolic cinnamate
decarboxylation is capable of providing potentially huge
quantities of oxygenated styrenes, which are difficult to pro-
duce by ordinary chemical means (5, 6a, b). Phenolic
styrenes are valuable starting materials for oxygenated bio-
degradable polymers, fragrances, and flavors, and chiral in-
termediates for organic synthesis.
Green chemistry (2) presents a continuing need for the
discovery of new biocatalysts capable of catalyzing useful
chemical transformations by novel mechanisms. An ideal
biocatalyst has few technical hurdles, such as the require-
Received 23 November 2001. Published on the NRC Research
Press Web site at http://canjchem.nrc.ca on 23 April 2002.
Dedicated to Professor J. Bryan Jones on the occasion of his
65th birthday.
K.-S. Lee and J.P.N. Rosazza.1 Division of Medicinal and
Natural Products Chemistry, and Center for Biocatalysis
and Bioprocessing, College of Pharmacy, University of
Iowa, Iowa City, Iowa 52242, U.S.A.
Biocatalytically unsubstituted styrene (1) is usually con-
verted to styrene epoxides (2) of varying enantiomeric
1Corresponding author: (e-mail: john-rosazza@uiowa.edu).
Can. J. Chem. 80: 582–588 (2002)
DOI: 10.1139/V02-034
© 2002 NRC Canada

Lee and Rosazza
583
Scheme 1. A summary diagram of known styrene bioconversions.
Solid arrows indicate aerobic conditions and dashed arrows indi-
cate anaerobic conditions.
Scheme 2. Oxidation of 4-vinylphenol (15) to 17 and 18 by
Nocardia sp.
-OH(18OH)
Si
18OH
H
H(2H)
O
H(2H)
H2O
D2O/H128
C
H2
C
CH3
CH3
H2
15
O
8
17
6
HO
HO
NADP+
O
16
H-
Re
NADPH
X
OH
O
O
OH
X
18O
H(2H)
3
CH3
19
2'
5'
2
C
1'
HO
3'
4'
20
1
2
HO
H2
6' 18
HO
OH
CHO
COOH
R
1
7
4
5
cohol, b-NAD+ (99%), b-NADP+ (sodium salt, 98%), and b-
NADPH (tetrasodium salt, 95%) were obtained from Sigma
(St. Louis, MO, U.S.A.). Coomassie Plus protein assay re-
agent and Albumin Standard (bovine serum album fraction
V, 2 mg mL–1 in a 0.9% aqueous NaCl solution containing
sodium azide) were obtained from Pierce (Rockford, IL,
U.S.A.). H218O (96 atom% 18O) and NMR solvents were
purchased from Cambridge Isotope Laboratories, Inc.
(Andover, MA, U.S.A.). HPLC solvents were obtained from
Fisher Scientific (Fair Lawn, NJ, U.S.A.), and other reagents
were analytical grade from Fisher Scientific unless other-
wise indicated.
H
OH
OH
OH
OH
COOH
COOH
9 R = CH2OH
10 R = CH=O
11 R = COOH
H
12
13
14
purities by enzymatic activation and insertion of molecular
oxygen into the unsaturated styrene side-chain (7–9). Sty-
rene epoxides (2) are also formed by hydrogen peroxide de-
pendent-peroxidase mediated epoxidations (10). Styrene
epoxides (2) are hydrolytically converted to diols (3) by
epoxide hydrolases (8, 11), to phenylacetaldehyde (4) and
phenylacetic acid (5) (9, 12), and to alkenes in a “reversal”
of the epoxidation reaction (13). With anaerobic consortia,
after water addition to the double bond, styrene (1) is oxi-
dized through the phenylacetate route to yield various prod-
ucts, including ethylbenzene (6), phenylethanol (7), 1-
phenylethanone (8), phenylacetaldehyde (4), phenylacetic
acid (5), benzyl alcohol (9), benzaldehyde (10), and benzoic
acid (11) (14). Styrene (1) is also metabolized via styrene
cis-glycol (12) to 3-vinylcatechol (13) and 2-vinylmuconate
(14), demonstrating double-bond and aromatic-ring oxygen-
ation prior to ring fission (15) (Scheme 1).
Much less is known about biocatalytic transformations of
phenolic styrenes such as 15. The introduction of oxygen
into the styrene side-chain olefin by a hydration mechanism
is a novel and relatively unexplored biocatalytic reaction. In
this study, we examined the mechanism and disposition of
15 by growing cultures, induced resting cells and cell-free
enzyme preparations of the Nocardia species, that afforded
S-1-(4>-hydroxyphenyl)ethanol (17) by vinylphenol hydra-
tase, and subsequently, 4>-hydroxyacetophenone (18) by an
NADP+-dependent alcohol dehydrogenase (Scheme 2).
Racemic 1-(4>-hydroxyphenyl)ethanol (17) was prepared
as described by the NaBH4 reduction of 4>-hydroxyace-
tophenone (18) in CH3OH (16).
S-(–)-17 was prepared by a modification of Reeve et al.
(17), where the lipase mediated enantioselective esterifica-
tion of R-(+)-17 occurs, leaving unreacted S-(–)-17 behind.
Racemic 17 (4 mg) was dissolved in vinyl acetate (4 mL) as
the acyl donor and as the solvent and lipase (4 mg, Pseudo-
monas fluorescens lipase, EC 3.1.1.3, 3097 units mg–1,
Fluka Chemical Co., Milwaukee, WI, U.S.A.) were added.
After incubation for 12 h at 28°C, lipase was filtered off and
the filtrate was evaporated. The residue was dissolved in
CH3OH (2 mL), filtered, and subjected to chiral HPLC anal-
ysis, where the enantiomeric excess of unreacted 17 was
98.8% ee of the S-(–)-isomer (Fig. 1B).
Analytical methods and instrumentation
TLCs were carried out on 0.25 mm layers of silica gel
GF254 (Sigma) precoated on polyester plates (20 × 20 cm).
The developing solvent was
a mixture of CH2Cl2–
CH3OH–HOAc (95:5:0.5, v/v/v). Developed chromatograms
were directly visualized under 254 and 366 nm UV light for
fluorescence quenching or fluorescence. Plates were then
spayed with diazotized sulfanilic acid reagent, which con-
sisted of solution A (0.5% sulfanilic acid in 1 N HCl), solu-
tion B (0.5% NaNO2 in water), and solution C (0.5% NaOH
in 50% (v/v) C2H5OH–H2O). Developed plates were first
sprayed with a fresh mixture of equal volumes of solutions
A and B, warmed with a heat gun, followed by spraying
with solution C to develop the colors. Under these condi-
tions, 15, 17, and 18 gave Rf values of 0.7 (scarlet), 0.3 (yel-
low), and 0.5 (orange), respectively.
Experimental
Chemicals
4-Vinylphenol (15) (10% solution in propylene glycol),
4>-hydroxyacetophenone (18), cinnamyl alcohol (3-phenyl-2-
propen-1-ol), 2-hydroxybenzyl alcohol, 3-hydroxybenzyl al-
cohol, 4-hydroxybenzyl alcohol, oleyl alcohol, and vinyl ac-
etate were obtained from Lancaster Synthesis Inc.
(Windham, NH, U.S.A.). D2O (99.9 atom% D), vanillyl al-
cohol (4-hydroxy-3-methoxybenzyl alcohol), cyclohexanol,
tetralol (1,2,3,4-tetrahydro-1-naphthol), 4-methoxybenzyl al-
cohol, benzyl alcohol, 1-phenyl-1-propyl alcohol, 1-phenyl-
2-propyl alcohol, and 3-phenyl-1-propyl alcohol were pur-
chased from Aldrich (Milwaukee, WI, U.S.A.). Methanol,
ethanol, 1-propanol, 1-butanol, 2-butanol, 2-phenylethyl al-
HPLC was performed with
a
Shimadzu liquid
chromatograph (Shimadzu Corporation, Kyoto, Japan)
equipped with dual pumps (LC-6A) and a variable wave-
length UV detector (SPD-6AV). Eluted peaks were detected
at 280 nm and identified by comparison with standard
compounds. Separations were carried out under isocratic
© 2002 NRC Canada

584
Can. J. Chem. Vol. 80, 2002
Fig. 1. Chiral HPLC resolution of R-(+)- and S-(–)-1-(4 >-
hydroxyphenyl)ethanol (17): (A) racemic 17; (B) S-(–)-17 re-
solved by Pseudomonas fluorescens lipase; (C) 17 isolated from
reaction of Nocardia cultures with 4-vinylphenol (15).
Mass spectra were obtained by direct inlet probe mass
spectrometry (DIPMS) performed on a Voyager mass spec-
trometer (ThermoQuest, Manchester, England). One dimen-
sional 1H and 13C NMR spectra were obtained with a Bruker
1
DMX-400 (operating at 400 MHz for H and 100 MHz for
13C) or an AMX-600 spectrometer (operating at 600 MHz
1
for H, Karlsruhe, Germany), equipped with an IBM Aspect
2000 processor and with VNMR version 4.1. Impurities in
deuterated NMR solvents were used as internal standards.
Chemical shift values (d) are reported in parts per million
(ppm) and coupling constants (J values) are given in Hz.
Abbreviations for NMR are s, singlet; d, doublet; and q,
quartet.
Fermentation and biocatalysis
Nocardia sp. NRRL 5646 was grown and maintained on
slants of Sabouraud-dextrose agar or sporulation agar
(ATCC No. 5 medium) (18). For analytical scale reactions,
cultures were grown by a two-stage protocol (19) in 25 mL
of sterile soybean meal-glucose medium held in stainless
steel-capped 125-mL DeLong culture flasks. The medium
contained 2% glucose, 0.5% yeast extract, 0.5% soybean
meal, 0.5% NaCl, and 0.5% K2HPO4 in doubly distilled wa-
ter, and was adjusted to pH 7.0 with 6 N HCl before being
autoclaved at 121°C for 15 min. Cultures were incubated
with shaking at 250 rpm at 28°C on an Innova 5000
Gyrotory tier Shaker (New Brunswick Scientific Co., Edi-
son, NJ, U.S.A.). A 10% inoculum derived from a 72-hour-
old first-stage culture was used to initiate the second-stage
cultures, which were incubated as above for 24 h. For
bioconversions, 15 and 17 were added directly, usually at
concentrations of 1 mg mL–1, and 18 was added at
0.4 mg mL–1 of media.
Induced Nocardia resting cells were prepared by adding
1 mg mL–1 15 to 24-hour-old stage-two cultures. After an
additional 24 h, incubations were filtered through a cheese
cloth to remove soybean meal solids, and the cells were con-
centrated by centrifugation (Sorvall RC 26 Plus centrifuge
with a SLA-600TC rotor, Kendro Laboratory Products,
Newton, CT, U.S.A.) at 10 000 × g at 4°C for 20 min. Cell
pellets were suspended and washed in cold phosphate buffer
(pH 7.0, 0.1 M) and centrifuged again. Typical wet cell
yields ranged from 20 to 24 g L–1.
Cell-free extracts were prepared from induced cells.
Typically, 15 g wet weight of cells were suspended in cold
phosphate buffer (45 mL, pH 7.0, 0.1 M) containing
dithiothreitol (1 mM) (Aldrich) and disrupted by a Sonifier
Cell Disrupter 350 (Branson Sonic Power Co., Danbury, CT,
U.S.A.) operating at 10 W with a 60% intermittent duty cy-
cle for 5 min. Cell debris was removed by centrifugation at
12 000 × g for 30 min at 4°C to provide a supernatant as the
cell-free extract. The cell-free extract was assayed for total-
protein content by the Bradford protein microassay (20) with
bovine serum albumin as the standard.
conditions over a Hypersil C18 column (250 × 4.6 mm ID,
5
m particle size, Alltech Associates, Inc., Deerfield, IL,
U.S.A.) protected with a guard column (10 × 4.6 mm ID,
particle size, Alltech) eluted with
5
m
CH3OH–CH3CN–H2O (7:3:10, v/v/v) at a flow rate of
0.8 mL min–1 to give retention volumes (Rv) of 3.6 mL for
17, 4.5 mL for 18, and 10.1 mL for 15. Standard curves
were established over the range of 1–10 g for 15, 1–8
for 17, and 0.2–0.8 g for 18.
g
Chiral HPLC was carried out with a Chiralcel OJ column
(250 × 4.6 mm ID, 10 m particle size, Chiral Technoloies
Inc., Exton, PA, U.S.A.) linked to a Chiralcel OJ guard col-
umn (50 × 4.6 mm ID, 10 m particle size) at a flow rate of
0.8 mL min–1 with n-hexane–2-propanol (9:1, v/v), which
gave retention volumes (Rv, mL) of 16.6 for 15, 16.8 for 18,
33.6 for S-(–)-17, and 37.5 for R-(+)-17.
For preparative-scale conversions, stage-two incubations
were carried out in 200 mL of medium held in 1-L DeLong
flasks. After 24 h of incubation in the second stage, 15 was
added as the substrate and incubated with shaking at
Flash column chromatography was conducted over silica
gel, 40 m for flash columns (J.T. Baker, Phillipsburg, NJ,
U.S.A.).
-
250 rpm at 28°C for 24 h. The reaction mixture was har
vested, acidified to pH 2 with 6 N HCl, followed by
centrifugation at 10 000 × g at 4°C for 20 min to remove
© 2002 NRC Canada

Lee and Rosazza
585
1
Table 1a. H NMR and mass spectral analyses for 1-(4>-hydroxyphenyl)ethanol (17) isolated from reactions of 4-vinylphenol (15) with
Nocardia conducted in H2O- or D2O-containing buffers.
Relative intensity at m/z (%)
1H NMR relative
intensity – C-2H
at d 1.35
D2O
123
[M – CH3]+
124
138
[M]+
139
[M + 1]+
140
[M + 2]+
amount
(%)
[M + 1 – CH3]+
0
50
100
100
100
100
9
18
69
1
2
11
100
100
100
14
9
8
3.0
2.7
1.6
1
Table 1b. H NMR and mass spectral analyses for 4 >-Hydroxyacetophenone (18) isolated from reactions of 4-vinylphenol (15) with
Nocardia conducted in H2O- or D2O-containing buffers.
Relative intensity at m/z (%)
1H NMR relative
intensity – C-2H
at d 2.51
D2O
121
[M – CH3]+
122
136
[M]+
137
[M + 1]+
138
[M + 2]+
amount
(%)
[M + 1 – CH3]+
0
50
100
100
100
100
24
23
83
7
6
18
100
100
100
9
8
10
3.0
2.8
1.6
cells. The supernatant was extracted wiht EtOAc (3 ×
200 mL), the combined EtOAc extracts were dried over an-
hydrous Na2SO4 and concentrated by rotary evaporation to
give 431 mg of crude extract. The extract was dissolved in
CH3OH (5 mL), adsorbed onto 2 g of silica gel, and loaded
onto a flash column (2 × 25 cm). The column was eluted
stepwise with CHCl3 and CHCl3–CH3OH (99:1 to 97:3) to
give fraction A (39 mg) and fraction B (173 mg). These
fractions were separately resolved by flash column chroma-
tography over silica gel using mixtures of n-hexane–EtOAc
(95:5) for fraction A, and n-hexane–EtOAc (8:2 to 7:3) for
fraction B. The resulting pure products were subjected to
chromatographic and spectral analyses.
from plates, extracted with CH3OH (3 × 20 mL), and the
extracts were evaporated to dryness for H NMR and MS
analysis.
1
Labeled-oxygen incorporation experiments also used in-
duced Nocardia resting cells that were suspended in a phos-
phate buffer (10 mL, pH 7.0, 0.1 M) containing glucose
(0.5%, prepared with 20% H218O) and contained in 50-mL
DeLong flasks. After addition of 15 (30 mg), the incubation
was carried out at 28°C with spinning at 250 rpm. The con-
trol consisted of an identical reaction mixture, except the
buffer was prepared with unlabeled H2O. The 24-hour-old
culture solutions were extracted with EtOAc (10 mL), the
extracts were concentrated and purified by preparative TLC,
as previously mentioned, to give products that were sub-
jected to spectral analysis.
Induced resting cell conversions of 4-vinylphenol
Induced resting-cell reactions were performed by sus-
pending 0.6 g wet weight of Nocardia cells in phosphate
buffer (25 mL, pH 7.0, 0.1 M) containing glucose (0.5%)
and 15 (25 mg). Samples of 2.0 mL were diluted with
CH3OH (2.0 mL), filtered through 0.2- m nylon membrane
filters, and 10 L of the filtrates were injected for HPLC
analysis. The same procedure was used in induced cell con-
versions of 17 at 1 mg mL–1 and 18 at 0.4 mg mL–1.
Enantiospecificity of 4-vinylphenol hydration
The enantiospecificity of the hydration reaction was eval-
uated by adding 15 (25 mg) to 25 mL volumes of 24-hour-
old stage-two cultures. Incubations were conducted in 125-
mL DeLong flasks. Samples of 3 mL were taken after 24,
48, 72, and 144 h and each was extracted with EtOAc
(3 mL). After evaporation, crude extracts were redissolved in
CH3OH to concentrations of 2 mg mL–1, passed through sy-
ringe filters (Acrodisc 13 mm with 0.45 m nylon mem-
brane filter, Gelman Science, Ann Arbor, MI, U.S.A.), and
the filtrate samples (20 L) were injected for chiral HPLC
analysis.
Reactions of 4-vinylphenol with Nocardia sp. in D2O
and 20% H218O
Deuterium-incorporation experiments used induced rest-
ing cells of Nocardia sp. as described, except that flasks
held phosphate buffer (25 mL, pH 7.0, 0.1 M) containing
glucose (0.5%, prepared with 50 and 100% D2O) and 15
(1 mg mL–1). For each reaction, incubations were stopped
after 24 h, acidified and pelleted as before, and supernatants
were extracted with EtOAc (3 × 25 mL). The combined ex-
tracts were dried over anhydrous Na2SO4 and evaporated to
dryness. The extracts were purified on glass preparative-
TLC plates coated with 1-mm layers of silica gel GF254
(20 × 20 cm, Sigma). UV-quenching bands were scraped
Substrate specificity of alcohol dehydrogenase in
Nocardia cell-free extracts
Incubations in quartz cuvettes were conducted in phos-
phate buffer (1.0 mL, pH 7.0, 0.1 M) containing the sub-
strates (3 mM), NADP+ or NAD+ (0.25 mM), and cell-free
extract (500 L, 1.70 mg protein). Substrates were dissolved
in dimethylformamide (DMF), and 10 L of the DMF solu-
tions were added to incubation mixtures. Reactions were
© 2002 NRC Canada

586
Can. J. Chem. Vol. 80, 2002
Table 2a. Mass spectral data for 1-(4>-hydroxyphenyl)ethanol (17) isolated from reactions of 4-vinylphenol (15) with Nocardia con-
ducted in H2O- and 20% H218O-containing buffers.
138
[M]+
139
[M + 1]+
140
[M + 2]+
123
[M – CH3]+
125
[M + 2 – CH3]+
Control H2O
20% H218O
100
100
10
23
0
18
100
100
2
23
Table 2b. Mass spectral data for 4 >-hydroxyacetophenone (18) isolated from reactions of 4-vinylphenol (15) with Nocardia conducted
in H2O- and 20% H218O-containing buffers.
136
[M]+
137
[M + 1]+
138
[M + 2]+
121
[M – CH3]+
123
[M + 2 – CH3]+
Control H2O
20% H218O
100
100
11
16
3
17
100
100
3
20
monitored under initial-rate conditions by measuring
changes in absorbance at 340 nm for 5 min at 20°C
(Shimadzu 2101-PC equipped with UVPC Optional Kinetics
Software v 2.5). The molar extinction coefficient used for
NADPH at pH 7.0 was ꢀ340 = 6.22 mM–1 cm–1.
buffer reaction, and 60% and 10%, respectively, from the
100% D2O-containing buffer reaction. Fragment ions at m/z
123 and 124, where methyl groups were lost from the mo-
lecular ions, were essentially identical to those from
unlabeled 17, showing that all the deuterium in labeled 17
1
resided in the methyl groups. H NMR of relative peak in-
tensities for C-2H showed approximately 10 and 47% deute-
rium incorporations vs. signals for Ar-2> and 6>-H,
respectively, similar to the mass spectral findings. Similar
analyses of 18 showed that all deuterium labeling resided in
the C-2 methyl group. Deuterium incorporations of 59 and
11%, respectively, for [M + 1]+ and [M + 2]+ from 100%
D2O-containing reactions matched very well with the results
Results and discussion
Conversion of 4-vinylphenol by Nocardia sp.
The reaction of Nocardia sp. with 4-vinylphenol (15) gave
two major products identified as 17 and 18. Characterization
1
2
data for 17: Rf = 0.3. H NMR (CD3OD) d: 7.19 (d, J =
8.4 Hz, 2H, Ar-2>, 6>H), 6.76 (d, 2J = 8.4 Hz, 2H, Ar-3>, 5>H),
2
2
1
4.73 (q, J = 6.5 Hz, 1H, C-1H), 1.35 (d, J = 6.5 Hz, 3H,
C-2H); identical to the results of Everhart and Craig (21).
MS m/z (%): 138 ([M]+, 15.4), 123 ([M – CH3]+, 63.5).
for 17 from 100% D2O. H NMR results for 18 again gave
6.6% incorporation from the 50% D2O reaction and 47%
from the 100% D2O buffer reaction. Results of deuterium in-
corporation from 50% D2O buffers were inexplicably lower
than expected.
1
Characterization data for 18: Rf = 0.5. H NMR (CD3OD) d:
2
2
7.87 (d, J = 8.8 Hz, 2H, Ar-2>, 6>H), 6.82 (d, J = 8.8 Hz,
2H, Ar-3>, 5>H), 2.51 (s, 3H, C-1H); spectrum was identical
to that for authentic 18. MS m/z (%): 136 ([M]+, 35.2), 121
([M – CH3]+, 100).
Comparisons of [M + 2]+ ions for 17 and 18 from reac-
tions conducted in buffers prepared with 20% H218O (Ta-
bles 2a and 2b) revealed oxygen-incorporation levels of 18
and 14%, respectively. Relative intensities of [M + 2 – CH3]+
ions revealed 21 and 17% increases in similar ions vs.
unlabeled 17. Both results are consistent with complete in-
corporation of oxygen from water and not from molecular
oxygen.
With induced resting cells, 15 was transformed to 17
(25%) and 18 (1%) within 48 h. Trace amounts of 4-
ethylphenol (19) were detected as a peak at Rv 12.1 mL, but
only after 144 h, indicating this to be a very minor
biotransformation pathway. With racemic 17, induced rest-
ing cells gave 18 in 10% yield, while 15 was undetectable.
In the reverse direction, 4>-hydroxyacetophenone (18) gave
17 (33%) in 48 h, again producing trace amounts of 4-
ethylphenol (19). Neither 17 nor 18 gave 15 as a product, in-
dicating the irreversibility of the initial hydration reaction.
Resting cell incubations in 50% D2O, 100% D2O, and
20% H218O containing buffers were conducted to examine
the source of oxygen and the mechanism by which Nocardia
oxidized 15 to 17 and 18. Reaction products isolated from
these incubations were analyzed by mass spectrometry and
1H NMR spectroscopy to determine the extent and positions
of isotope incorporations. Tables 1a and 1b show the results
of deuterium incorporation. For 17, when compared with
control reactions conducted in H2O-containing buffer, ions at
m/z 139 and 140 for [M + 1]+ and [M + 2]+ were enhanced
by 9% and 1%, respectively, from 50% D2O-containing
Labeling results permit a description of the reaction pro-
cess as shown in Scheme 2. Nocardia contains a hydratase
that binds and tautomerizes 4-vinylphenol (15) to a quinoid
intermediate (16), which adds water to afford 17. The
tautomerization reaction is apparently partially reversible,
because both 17 and its oxidation product 18 contain 1.4
deuterium atoms on the C-2 methyl group. Addition of oxy-
gen from water and not air was clearly confirmed by the
H218O labeling experiment, which gave nearly theoretical
incorporations of 18O into 17 and 18. Slightly lower values
than expected in deuterium and H218O incorporations can be
explained by dilutions of isotopically labeled water by
intracellular stores of water during the hydration reaction.
Reactions conducted under an argon atmosphere with in-
duced resting cells and in cell free extracts without co-
factors gave similar products (data not shown), supporting
© 2002 NRC Canada

Lee and Rosazza
587
Table 3. Influence of time on ee% of 17 from hydration of 4-vinylphenol (15), reduction of 4>-hydroxyacetophenone (18), and oxida-
tion of racemic 17 by Nocardia cultures.
From hydration of 15
S-(–)-17 ee%
From reduction of 18
S-(–)-17 ee%
From oxidation of R,S-17
Time (h)
Yield (%)
Yield (%)
R-(+)-17 ee%
Yield of 18 (%)
24
48
72
90
90
88
84
4
10
13
24.6
98
90
90
86
0.5
1.6
2.4
5.7
1.4
2.2
4.9
7.7
16
29
36
52
144
Table 4. Substrate specificity of Nocardia sp. alcohol dehydrogenase.
Enantiospecificity of 4-vinylphenol hydration
Relative activity (%)a
The stereospecificity of Nocardia 4-vinylphenol hydration
was established by chiral HPLC comparison of 17 formed
by Nocardia cultures to S-(–)-17 prepared by Pseudomonas
fluorescens lipase resolution of racemic 17 (17, 26).
Chromatograms (Fig. 1) for racemic 17 (Fig. 1A), lipase re-
solved S-(–)-17 (Fig. 1B), and 17 from incubation reactions
(Fig. 1C.) clearly show that 17 produced by Nocardia is pre-
dominantly S-(–)-17. We then compared the enantiomeric
purity of 17 formed by hydration of 15 vs. time (Table 3).
From 24 to 144 h, yields of 17 ranged from 4% to about
25%, while the ee of S-(–)-17 decreased from 90% at 24 h to
84% at 144 h. Over time, the reduction of 18 to 17, a less fa-
vorable reaction over all, progressed from 0.5% at 24 h to
about 6% at 144 h, while the ee of S-(–)-17 ranged from 98
to 86% (Table 3). These results indicate that hydration oc-
curs preferentially by si-face delivery of water to 16, and
that the reduction of 18 occurs preferentially by re-face hy-
dride delivery. As both reactions progressed, the ee% de-
creased, suggesting the enhancement of R-(+)-17 by reaction
reversibility, or the involvement of more than one alcohol
dehydrogenase with varying enantiospecificity or selectivity.
In this regard, when racemic 17 was oxidized to 18, small
excesses of R-(+)-17 (7.7% ee) accumulated as the reaction
time and yield progressed (52% yield at 144 h) (Table 3).
Substrate (3 mM)
Primary alcohols
Vanillyl alcohol
Benzyl alcohol
100
10
15
20
65
11
4
2-Hydroxybenzyl alcohol
3-Hydroxybenzyl alcohol
4-Hydroxybenzyl alcohol
4-Methoxybenzyl alcohol
2-Phenylethyl alcohol
Cinnamyl alcohol
3-Phenyl-1-propyl alcohol
Methanol
Ethanol
1-Propanol
1-Butanol
Oleyl alcohol
23
2
59
25
39
39
0
Secondary alcohols
1-(4>-Hydroxyphenyl)ethanol
2-Butanol
12
9
9
2
3
Cyclohexanol
1-Phenyl-1-propyl alcohol
1-Phenyl-2-propyl alcohol
Tetralol
3
aVanillyl alcohol was oxidized by cell free extracts containing NADP+
at a rate of 101.2 M sec–1 with a DmAbs sec–1 of 0.630.
Substrate specificity of alcohol dehydrogenase in cell-
free Nocardia extracts
the proposed hydration mechanism. The addition of water to
a styrene double bond through a hydration mechanism and
not by activation and insertion of molecular oxygen through
an epoxide intermediate has not been observed before, ex-
cept with a mixed anaerobic microbial consortium (14).
Hopper and co-workers (22) reported 17 and 18 as the
products of Pseudomonas sp. derived, flavoprotein-catalyzed
oxidations of 4-ethylphenol (19). 4-Ethylphenol methy-
lenehydroxylase, p-cresol methylhydroxylase, and vanillyl
alcohol oxidase (17, 22, 23) are all flavoproteins that oxidize
4-ethylphenol (19) to a proposed quinoid intermediate
(16) similar to that proposed here for Nocardia sp. with 15.
The R vs. S stereochemistry of 17 produced in flavoprotein-
mediated reactions varies with the enzyme used, and
may be affected by the addition of electron acceptors
(17). Heresztyn (24) and Eldin et al. (25) showed that
4-vinylphenol (15) could be microbiologically reduced to
4-ethylphenol (19). To rule out a combination of the
4-vinylphenol reduction and coupled flavoprotein oxidation of
4-ethylphenol (19) to 17, Nocardia sp. was incubated with
19. No metabolites were obtained, thus ruling out this oxida-
tive pathway and the benzylic hydroxylation of 19 to 17.
Both NADP+ and NAD+ were co-factors in the oxidation
of 17 to 18 in Nocardia cell free extracts. The alcohol oxida-
tion reaction, however, was nearly 17 times more favorable
with NADP+. The range of substrates used by the crude en-
zyme preparation was broad, as shown in Table 4. Among
the primary alcohols, phenolic benzyl alcohols were favor-
able substrates with vanillyl alcohol giving the highest rela-
tive oxidation rates. Among the mono-phenolic benzyl
alcohols, 4-hydroxybenzyl alcohol was most readily oxi-
dized, indicating a preference for and a possible participa-
tion of the 4-phenolic moiety in the oxidation reaction. As a
group of compounds, aliphatic alcohols including methanol,
ethanol, 1-propanol, and 1-butanol were oxidized more
readily than secondary alcohols. We compared the relative
rates of oxidation vs. reduction of 4-hydroxybenzalde-
hyde–4-hydroxybenzyl alcohol and benzaldehyde–benzyl
alcohol using crude cell free Nocardia extracts prepared
from cells induced with 4-vinylphenol (15). In these prepa-
rations, the rate of alcohol oxidation was 13 times greater
than aldehyde reduction, while the benzaldehyde reduction
was greater by 14 times than the alcohol oxidation of benzyl
© 2002 NRC Canada

588
Can. J. Chem. Vol. 80, 2002
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Witholt, A. Schmid, and M.G. Wubbolts. Appl. Environ.
Microbiol. 64, 2032 (1998).
alcohol. We have initiated the purification and characteriza-
tion of vinylphenol hydratase.
Acknowledgments
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We are grateful for the inspirational and pioneering efforts
of Professor J. Bryan Jones in applying enzymes in synthetic
organic chemistry. His infectious enthusiasm and chemical
excellence has influenced a generation of chemists and has
fostered the widespread application of enzymes in green
chemistry approaches in the field. This work was supported
by the USDA through the Iowa Biotechnology Byproducts
Consortium.
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© 2002 NRC Canada