Asymmetric transesterification of secondary alcohols catalyzed by feruloyl esterase from Humicola insolens

Article abstract of DOI:10.1016/j.bioorg.2005.05.002

A new asymmetric transesterification of secondary alcohols catalyzed by feruloyl esterase from Humicola insolens has been found. Although alcohols are not the natural substrates for this enzyme, a high R enantioselectivity was observed. Stereochemical studies showed that variations in substrate structure lead to strong variations in enantioselectivity. The highest enantioselectivities are obtained when the β-carbon of the secondary alcohol is tertiary or quaternary.

Full text of DOI:10.1016/j.bioorg.2005.05.002

Bioorganic Chemistry 33 (2005) 325–337
www.elsevier.com/locate/bioorg
Asymmetric transesterification of secondary
alcohols catalyzed by feruloyl esterase
from Humicola insolens
Nikos S. Hatzakis, Ioulia Smonou *
Department of Chemistry, University of Crete, Iraklion 71409, Crete, Greece
Received 2 March 2005
Available online 20 June 2005
Abstract
A new asymmetric transesterification of secondary alcohols catalyzed by feruloyl esterase
from Humicola insolens has been found. Although alcohols are not the natural substrates
for this enzyme, a high R enantioselectivity was observed. Stereochemical studies showed that
variations in substrate structure lead to strong variations in enantioselectivity. The highest
enantioselectivities are obtained when the b-carbon of the secondary alcohol is tertiary or
quaternary.
Ó 2005 Elsevier Inc. All rights reserved.
Keywords: Ferulic acid; Feruloyl esterase; Humicola insolens; Transesterification; Enantioselectivity;
Secondary alcohols
1. Introduction
Ferulic acid esterases (FAE) [1] are a subclass of the carboxylic ester hydrolases
(EC 3.1.1.1) which hydrolyze the esters of hydroxycinnamic acids [2], such as p-cou-
maric acid, ferulic acid, and sinapic acid, with sugars. Hydroxycinnamates are
*
Corresponding author. Fax: +30 2810 393601.
E-mail address: smonou@chemistry.uoc.gr (I. Smonou).
0045-2068/$ - see front matter Ó 2005 Elsevier Inc. All rights reserved.
doi:10.1016/j.bioorg.2005.05.002

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N.S. Hatzakis, I. Smonou / Bioorganic Chemistry 33 (2005) 325–337
important structural components of plant cell walls where they form cross-links with
various polymers such as hemicellulose and pectin [3,4]. Feruloyl esterases are se-
creted by a number of bacterial [5,6] and fungal [7,8] organisms which exploit plants
either to enter the plant cell or to utilize the cell wall material as nutritional resource
[9]. The complete degradation of plant cell wall polymers requires multienzyme com-
plex systems [10]. Most of the feruloyl esterases have been shown to act synergisti-
cally with xylanases, cellulases, and pectinases [11] to break down complex plant
cell wall carbohydrates [12,13]. In addition to their fundamental biological impor-
tance, feruloyl esterases have many potential industrial and medicinal applications.
Most of the research done to date involves the isolation, purification, and charac-
terization of feruloyl esterases derived from a wide range of microorganisms [5–8], as
well as the enzymatic release of the products from cell-wall degradation [3,4]. In all
cases, the activities of these enzymes have been determined by the release of free feru-
lic acid or other hydroxycinnamic acids. A few studies on substrate specificity and on
the kinetics towards a range of methyl phenylalkanoates have been used to probe the
active site of the enzyme [2,9].
Recently, the crystal structures of feruloyl esterase module of xylanase 10B [14]
xylanase Z [15] from Clostridium thermocellum as well as ferulic acid esterase from
Aspergillus niger were reported [16,17]. The primary structure analysis of FAEs
has shown that they have an a/b hydrolase fold, with a Ser-His-Asp catalytic triad
at their active site [10,14–17]. The overall fold of these proteins is very similar to that
of the fungal lipases. Despite the fact that feruloyl esterases and lipases bear the same
catalytic triad of residues in their active site, they do not show lipase activity. Re-
cently, the synthesis of pentylferulate was achieved by using a water-in-oil micro-
emulsion system containing a feruloyl esterase from Aspergillus niger [18].
However, the potential of feruloyl esterases for the transesterification of alcohols
has never been reported.
We report here for the first time, experimental results of a new and high asymmet-
ric transesterification of secondary alcohols catalyzed by a crude feruloyl esterase
preparation from Humicola insolens [19]. The observed new stereoselectivity of this
enzyme toward non natural substrates, such as secondary alcohols, was unexpected
and may lead to a promising biocatalyst for the enantioselective synthesis of various
alcohols and esters with interesting practical applications.
2. Results and discussion
Selective separation of one enantiomer from a racemic mixture by kinetic resolu-
tion techniques played a central role in the preparation of optically active com-
pounds, despite the disadvantage that only a 50% maximum yield of one
enantiomer can be obtained. In cases where the racemic substrates and the catalyst
are readily available in large quantities, this method can be useful in organic
synthesis.
In this work, a new and highly stereoselective transesterification of secondary
alcohols catalyzed by a crude feruloyl esterase preparation from H. insolens has been

N.S. Hatzakis, I. Smonou / Bioorganic Chemistry 33 (2005) 325–337
327
Table 1
Effect of temperature in the enzymatic resolution of 1-phenyl-1-ethanol (1)
Substrate
Temperature
Time (days)
Conversion (%)
ee (%)
E
1
1
1
25
45
65
3
3
3
12
23
30
94
92
80
36
32
12
studied. This is the first example of the feruloyl esterase reactivity towards substrates
whose structural characteristics are completely different than those of the natural
substrate of this enzyme.
Initially, we investigated the effect of temperature in the enzymeꢀs enantioselectiv-
ity and activity. Transesterification of 1-phenylethanol was used as a model reaction.
In a typical run, 0.2 mmol of the substrate and a 30-fold excess of vinyl acetate were
added in a vial to which 200 mg of the enzyme preparation was placed. Vinyl acetate
was used as the activated acyl donor [20]. The resulting suspension was shaken at
three different temperatures. Samples were collected periodically and were analyzed
by gas chromatography. The reaction was performed in 25, 45, and 65 °C. The re-
sults obtained are summarized in Table 1.
As seen from Table 1, the highest enantioselectivity is observed at 25 °C with a
significant loss of activity (conversion = 12%), whereas the highest activity is at
65 °C with a significant loss of enantioselectivity, may be due to denaturation of
the enzyme. After that, the optimum temperature for this reaction is 45 °C. All
the rest reactions were performed at 45 °C.
The general protocol of the stereoselective transesterification of secondary alco-
hols is as follows: 0.2 mmol of the substrate and a 30-fold excess of vinyl acetate were
added in a vial to which 200 mg of the enzyme preparation was placed. Vinyl acetate
was used as the activated acyl donor [20]. The resulting suspension was shaken at
45 °C. Samples were collected periodically and were analyzed by gas chromatog-
raphy. The enantioselectivity in each reaction was determined by analyzing the enan-
tiomeric excess of the unreacted alcohol and the produced ester by using a 30 m
1
chiral capillary column (HP-5 cross-linked 5% phenyl-methyl silicone), or by H
NMR spectroscopy by using chiral shift reagents or MTPA esters [21]. The results
are summarized in Table 2.
As seen from Table 2, transesterification of a series of aryl alcohols 1–11 shows a
high selectivity for the R enantiomer of the racemic mixture. In most cases, the R
configuration was determined from the measured optical rotations. The high enan-
tiopreference of > 98% with E > 100 is impressively illustrated by substrates 7–9.
Similarly para substituted aryl alcohols with either electron donating, 2 and 3, or
electron withdrawing groups, 4–7, show high enantioselective transesterifications
ranging from 88% to 95% ee. Apparently, the electronic properties of the phenyl sub-
stituent do not significantly affect enzymeꢀs enantioselectivity. However, the substitu-
tion on the meta or para position alter the observed enantioselectivity. FAE shows
higher enantioselectivity for meta than para substituted alcohols, as demonstrated by
the two isomeric pairs 2, 3, and 6, 7. Furthermore, excellent enantioselectivity is
demonstrated again in the 1-(4-pyridinyl)-1-ethanol, 9, where more than 98% ee

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N.S. Hatzakis, I. Smonou / Bioorganic Chemistry 33 (2005) 325–337
Table 2
Substratea
Conversion (%)
23
ee (%)
92
Time (days)
3
Configuration
Eb
1
2
Rc
32
19
18
28
15
18
18
30
17
28
25
52
88
98
3
Rd
–
3
3
>100
75
4
97
2
R
R
R
–
5
94
5
40
6
93
6
34
7
>99
>98
>98
93
7
>100
>100
>224
37
8
2
–
9
0.5
3
Re
Rf
Rg
10
11
91
5
>100
a
Alcohols 1 and 9 were commercially available. Alcohol 6 was prepared by BF3/EtSiH reduction [29] of
the corresponding aryl ketone. Alcohol 10 was prepared by Grignard addition of EtMgI to the
corresponding aldehyde. All the other alcohols were prepared by Grignard addition of CH3MgI to the
corresponding aldehydes.
b
See [22].
c
By GC analysis with a chiral capillary column of the remaining alcohol, compared with standard
S-(ꢀ)-1-phenyl-1-ethanol.
25
d
By the optical rotation of the remaining alcohol after 35% conversion. Measured ½aꢁD ꢀ15 (c 1,
25
CHCl3), ee % 30, reported ½aꢁD +52 (c 1, CHCl3), ee % 87 (R)-alcohol, [23].
e
The configuration of the ester was determined by the optical rotation of the isolated product after 50%
25
25
conversion. Measured ½aꢁD +81 1 (c 1, CHCl3), reported ½aꢁD +81 (c 1.01, CHCl3) [24].
25
f
By the optical rotation of the isolated product after 50% conversion. Measured ½aꢁD +47 1 (c 1,
25
CHCl3), reported ½aꢁD +49 (c 1, CHCl3), [22].
g
By comparison of GC analysis of the produced ester and the enzymatic hydrolysis of racemic ester of
alcohol 7 with PFL, [25].

N.S. Hatzakis, I. Smonou / Bioorganic Chemistry 33 (2005) 325–337
329
transesterification product was obtained with E > 100. In this case and in substrate
10, when the conversion was extended to 60%, the unreacted alcohol was practically
enantiopure (> 99% ee) having the S configuration. Also, substitution of an ethyl
(substrate 10) for the methyl group (substrate 1) does not alter the high degree of
stereoselectivity. At this point, it is difficult to rationalize the factors (electronic, ste-
ric) that contribute to this remarkable enantioselectivity with R preference. However,
when the ‘‘small’’ substituent was isopropyl or tert-butyl group no selectivity was ob-
served (data not shown). It must be pointed out, that all these results are consistent
with the Kazlauskas model, [26] which predicts the enantiopreference of lipase to-
wards secondary alcohols. This model is based on the size difference of the substit-
uents at the stereogenic center and suggests that lipases distinguish between
enantiomeric secondary alcohols according to the size of the two substituents.
To examine further the stereochemical factors governing FAEs enantioselectivity
towards secondary alcohols, we studied the secondary alcohols 12–22, where the
small substituent S (methyl) at the stereogenic center is the same through the series
of the examined substrates while the large substituent L is varied. The results are
summarized in Table 3.
As seen from Table 3, FAE exhibits high to excellent enantioselectivity in the transe-
sterification of alcohols 12–15. The common feature among these substrates is the exis-
tence of a tertiary or quaternary moiety next to the carbon which bears the –OH group.
Alcohol 12, with an adamantyl group (a quaternary carbon) next to the stereogenic
alcohol carbon, impressively illustrates this point. However, a significant loss of enan-
tioselectivity is observed when the large group L has an additional methylene group
that is directly bonded to the stereogenic center(enantioselectivities ranging between
35% and 75% ee). Substrates 16–20 illustrate this point. For example, E factor is
> 100 for substrate 12, compared to E factor 9 for alcohol 16. Again, the major product
of these transesterifications is the R enantiomer.
3. Conclusion
In summary, these results show for the first time that feruloyl esterase from
H. insolens can successfully catalyze the stereoselective transesterification of second-
ary alcohols that bear no structural similarity to the natural substrates of this en-
zyme. These reactions proceed with good to excellent enantioselectivity that is
analogous to that of lipases [23–25]. Our results also demonstrate that the enantio-
selectivity of FAE in the transesterification of secondary alcohols depends signifi-
cantly on the substitution of the carbon next to the stereogenic center which bears
the hydroxyl group. Excellent enantiodiscrimination is achieved when the b-carbon
is tertiary or quaternary, whereas moderate enantioselectivity is obtained when this
carbon is secondary. Further increase in the substitution of the b-carbon results in
improvement of the observed enantioselectivity. Additionally, the substitution of
the phenyl ring of secondary benzylic alcohols affects the observed enantioselectivity
of this enzyme. Higher enantioselectivity is observed for meta-substituted compared
to para-substituted substrates.

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N.S. Hatzakis, I. Smonou / Bioorganic Chemistry 33 (2005) 325–337
Table 3
Asymmetric enzymatic transesterification of alcohols 12–22 with FAE
Substratea
Conversion (%)
ee (%)
Time (days)
Configuration
E
12
13
14
15
16
17
18
19
20
21
30
>95
15
R
>100
10
22
23
22
22
21
20
21
25
19
>95
92
85
75
60
40
66
34
62
46
3
24
9
Rb
–
>100
31
16
9
–
18
16
30
9
–
–
6
Rc
Rd
–
3
6
3
2
4
Re
–
5
22
7
3
a
Alcohols 12, 14, 15, 16, 17, 18, 21, and 22 were prepared by MeLi addition to the corresponding
carboxylic acids followed by LiAlH4 reduction of the produced ketones. Alcohols 19 and 20 were prepared
by reduction of the corresponding ketones with LiAlH4. Synthesis of alcohol 13 was accomplished by
Grignard addition of EtMgI to the corresponding aldehyde.
b
See [27].
c
The configuration of the alcohol was determined by the optical rotation of the isolated product after
25
25
35% conversion. Measured ½aꢁD +10.1 (c 1, CHCl3) 38% ee, reported ½aꢁD +12.6 (c 1.01, CHCl3) [23].
d
By chemical esterification of commercially available S-alcohol to the S-ester and comparison of elution
time in GC analysis.
e
See [28].
Further studies are currently under way in our laboratories in order to gain a bet-
ter insight on substrate specificity and to explore the usefulness of this enzyme as a
potent catalyst for the enantioselective resolution of various alcohols and esters with
interesting practical applications.

N.S. Hatzakis, I. Smonou / Bioorganic Chemistry 33 (2005) 325–337
331
4. Experimental
4.1. General methods and materials
All commercially available reagents were used as received (Merck and Sigma-Al-
drich) without any further purification. Solvents were distilled and dried using stan-
dard techniques.
The FAE preparation (Pentopan 500 BG) was kindly provided by Novo Nordisk
as a commercial culture supernatant from a strain of H. insolens. Pseudomonas flu-
orescens lipase (PFL) was purchased from Fluka. Diethyl ether and THF were
freshly distilled over sodium. Standard Schlenk techniques were employed by using
argon as the inert atmosphere for all manipulations of air or moisture sensitive
reactions.
The enzymatic reactions were incubated at 45 °C under shaking at 400 rpm. Reac-
tions were quenched on completion by filtering off the enzyme. Solvents were re-
moved under reduced pressure and the products were purified by flash
chromatography using Merck silica gel (230–400 mesh). Control reactions without
enzyme were carried out under the same conditions. All enzymatic reactions were
performed two times.
The progress of the FAE catalyzed reactions and the enantiomeric excesses were
determined by gas chromatography (HP5890II gas chromatograph equipped with an
FID detector; column: 30 m chiral capillary column, HP-5 cross-linked 5% phenyl-
methyl silicone). The estimated error was <2%. In some cases, the enantioselectivity
was determined by 1H NMR analysis of the corresponding MTPA esters or by using
optically active shift reagents [Eu(hfc)3].
MS were taken on a GC–MS (Simatzu GCMS-QP5050 equipped with a SPB-5
1
column and CI mass detector). H NMR and 13C NMR spectra were recorded on
a 300 or a 500 MHz Bruker spectrometers in CDCl3 solutions, by using Me4Si as
internal standard. Chemical shifts are reported in parts per million downfield
from Me4Si. Yields refer to isolated and spectroscopically pure materials. Optical
rotations were measured on a D,P 360 Jasco polarimeter with a Hg 360
lamp. Measurements were made in 1 cm, permanent-window cell at room
temperature.
Feruloyl esterase (FAE) activity was calculated with destarched wheat bran as
substrate. FAEꢀs activity was assayed by the analysis of free ferulic acid (FA)
released from destarched wheat bran. Ferulate release was analyzed by HPLC
(Nucleosil C18, column). The assay was carried out in 100 mM MOPS buffer,
pH 6, and 45 °C. One unit (U) of activity was defined as the amount of enzyme
that catalyzes the release of 1 lmol FA/min. The FAE activity was found to be
0.435 U/gr.
4.2. Preparation of aryl alcohols
Alcohols 2, 3, 4, 5, 7, 8, 9, and 10 were prepared by the addition of the proper
Grignard reagent to the corresponding aldehyde as follows:

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N.S. Hatzakis, I. Smonou / Bioorganic Chemistry 33 (2005) 325–337
To a stirred mixture of 12 mmol of crushed Mg in 20 ml freshly distilled absolute
Et2O, under inert atmosphere, 10 mmol of alkyl iodide in 10 ml of absolute Et2O was
added dropwise, followed by 30 min reflux. In some cases the addition of an iodine
crystal was required. After cooling the reaction mixture at 0 °C, 10 mmol of the
appropriate carbonyl compound in 10 ml of absolute Et2O was added dropwise.
The reaction mixture was stirred under reflux conditions and after cooling at 0 °C,
0.36 mmol of water was added dropwise and the produced alcohol was isolated fol-
lowing standard Grignard work-up conditions.
Alcohol 6 was prepared by reduction of the corresponding p-nitro-acetophenone
with Et3SiH/BF3 system [29].
Alcohol 11 was prepared from the corresponding ketone by reduction with
LiAlH4 [30].
1-(4-Methoxyphenyl)-1-ethanol, 2: yield 1.32 g (90%). 1H NMR (CDCl3
500 MHz, d ppm) 7.32 (d, J = 8.2 Hz, 2H), 6.90 (d, J = 8.2 Hz, 2H), 4.88 (m,
J = 6.3 Hz, 1H), 3.82 (s, 3H), 1.50 (d, J = 6.3 Hz, 3H). MS: m/z 152.
1-(3-Methoxyphenyl)-1-ethanol, 3: yield 2.2 g (96%). 1H NMR (CDCl3
500 MHz, d ppm) 7.16–7.29 (t, J = 7.8 Hz, 1H), 6.95 (d, J = 6.2 Hz, 2H), 6.83
(d, J = 6.2 Hz, 1H) 4.87 (q, J = 6.4 Hz, 1H), 3.82 (s, 3H), 1.5 (d, J = 6.4 Hz,
3H). 13C NMR (CDCl3 500 MHz, d ppm) 159.74, 147.59, 129.49, 117.67, 112.84,
110.89, 70.27, 55.19, 25.10.
1
1-(4-Fluorophenyl)-1-ethanol, 4: yield 1.25 g (90%). H NMR (CDCl3 500 MHz,
d ppm) 7.38–7.41 (m, 2H), 7.06–7.1 (m, 3H), 4.83 (m, 1H), 1.39 (d, J = 4.5 Hz, 3H).
1-(4-Trifluoromethylphenyl)-1-ethanol, 5: yield 1.4 g (90%). 1H NMR (CDCl3
500 MHz, d ppm) 7.63 (d, J = 7.7 Hz, 2H), 7.51 (d, J = 8.0 Hz, 2H), 4.99 (m,
J = 6.4 Hz, 1H), 1.53 (d, J = 6.4 Hz, 3H). MS: m/z 190.
1
1-(4-Nitrophenyl)-1-ethanol, 6: yield 1.6 g (95%). H NMR (CDCl3 500 MHz, d
ppm) 8.22 (d, J = 7.7 Hz, 2H), 7.56 (d, J = 8.3 Hz, 2H), 5.04 (m, J = 6.4 Hz, 1H),
1.54 (d, J = 6.4 Hz, 3H). MS: m/z 152.
1-(3-Nitrophenyl)-1-ethanol, 7: yield 0.84 g (50%). 1H NMR (CDCl3 500 MHz, d
ppm) 8.25 (s, 1H), 8.12 (d, J = 7.6 Hz, 1H), 7.72 (d, J = 6.7 Hz, 1H), 7.53 (m, 1H),
5.03 (m, J = 6.1 Hz, 1H), 1.55 (d, J = 6.1 Hz, 3H).
1-(4-Phenyl-phenyl)-1-ethanol, 8: yield 0.75 g (94%). 1H NMR (CDCl3 500 MHz,
d ppm) 7.61 (d, J = 8.0 Hz, 4H), 7.47 (m, J = 8.0 Hz, 4H), 7.37 (m, 1H), 4.98 (q,
J = 6.4 Hz, 1H), 1.57 (d, J = 6.4 Hz, 3H).
1-(4-Pyridinyl)-1-ethanol, 9; yield 0.5 g (94%). 1H NMR (CDCl3 500 MHz, d
ppm) 8.54 (d, J = 6.6 Hz, 2H), 7.31 (d, J = 3.8 Hz, 2H), 4.92 (m, J = 3.8 Hz, 1H),
1.59 (d, J = 3.8 Hz, 3H).
1
1-Phenyl-1-propanol, 10: yield 1.3 g (96%). H NMR (CDCl3 500 MHz, d ppm)
7.37 (t, J = 3.0 Hz, 4H), 7.28 (m, J = 2.5 Hz, 1H), 4.62 (t, J = 6.6 Hz, 1H), 1.78
(dd, J = 7.1 Hz, 1H), 1.86 (dd, J = 7.3 Hz, 1H), 0.95 (t, J = 7.4 Hz, 2H).
13C NMR (CDCl3 500 MHz, d ppm) 144.59, 128.38, 125.96, 76.006, 31.86,
10.12.
1-(3-Methylphenyl)-1-ethanol, 11: yield 1.17 g (98%). 1H NMR (CDCl3 500 MHz,
d ppm) 7.26–7.29 (t, J = 3.7 Hz, 1H), 7.19 (d, J = 7.5 Hz, 2H), 7.12 (d, J = 7.5 Hz,
1H), 4.87 (q, J = 6.4 Hz, 1H), 2.4 (s, 3H), 1.51 (d, J = 6.4 Hz, 3H).

N.S. Hatzakis, I. Smonou / Bioorganic Chemistry 33 (2005) 325–337
333
13C NMR (CDCl3 500 MHz, d ppm) 145.81, 138.10, 128.38, 128.17, 126.10,
122.42, 70.36, 25.09, 21.43.
4.3. Preparation of alcohols 12, 14, 16, 17, 18, 21, and 22
The above alcohols were prepared from their corresponding ketones by reduction
with LiAlH4. These ketones were prepared from their carboxylic acids by addition of
MeLi [31] as follows:
To a stirred solution of 5 mmol of carboxylic acid dissolved in 50 ml freshly dis-
tilled absolute Et2O, under inert atmosphere, which was cooled to ꢀ78 °C,
11 mmol of MeLi was added dropwise. The reaction mixture was stirred at this
temperature for 15 min and then it was left to reach room temperature. It was
then stirred for 3 h. After completion of the reaction, the reaction mixture was
cooled at 0 °C and 1 ml of water was added dropwise. The organic layer with
washed with distilled water and saturated aqueous NaCl and then it was dried over
MgSO4. The corresponding ketone was isolated after removal of the ether under
vacuum.
The overall yield of isolated alcohols was in the range of 50–60%.
1
1-Adamantyl-1-ethanol, 12: yield 0.50 g (58%). H NMR (CDCl3 500 MHz, d
ppm) 3.3 (m, 1H), 2.0 (s, 3H), 1.74 (d, 3H), 1.65 (m, 7H), 1.50 (d, 3H), 1.2 (d,
J = 6.4 Hz, 3H). 13C NMR (CDCl3 500 MHz, d ppm) NMR (CDCl3 500 MHz, d
ppm) 75.75, 37.69, 37.23, 36.53, 28.31, 16.41.
1
3,3-Dimethyl-2-butanol, 14: yield 0.50 g (58%). H NMR (CDCl3 500 MHz, d
ppm) 3.49 (q, J = 6.3 Hz, 1H), 1.13 (d, J = 6.3 Hz, 3H), 0.91 (s, 9H).
1
3-Methyl-2-butanol, 15: yield 0.42 g (57%). H NMR (CDCl3 500 MHz, d ppm),
3.49 (q, 1H, J = 6.3 Hz), 1.13 (d, 3H, J = 6.3 Hz), 0.91 (s, 9H).
1
1-Adamantyl-2-propanol, 16: yield 0.66 g (60%). H NMR (CDCl3 500 MHz, d
ppm) 4.05 (m, 1H), 1.96 (s, 3H), 1.67 (dd, 6H, J1 = 12.1 Hz, J2 = 33.3 Hz), 1.57
(s, 6H), 1.25 (m, 2H), 1.19 (d, J = 6.2 Hz, 3H). 13C NMR (CDCl3 500 MHz, d
ppm) 64.35, 54.18, 42.98, 37.04, 32.21, 28.66, 26.04.
1
4,4-Diphenyl-2-butanol, 17: yield 0.66 g (68.5%). H NMR (CDCl3 500 MHz, d
ppm) 7.36 (m, 8H), 7.27 (m, 2H), 4.28 (m, 1H), 3.73 (m, 1H), 2.26 (t, J = 7.1 Hz,
2H), 1.28 (d, 6.1 Hz, 3H). 13C NMR (CDCl3 500 MHz, d ppm) 145.11, 144.44,
128.60, 128.58, 128.13, 127.82, 126.33, 126.25, 65.85, 47.74, 45.02, 24.06.
1
1-Phenyl-2-propanol, 18: yield 0.63 g (72%). H NMR (CDCl3 500 MHz, d ppm)
7.34 (m, 2H), 7.26 (m, 3H), 4.04 (m, 1H), 1.27 (d, J = 6.2 Hz, 3H).
4-Phenyl-3-buten-2-ol, 21: yield 0.91 g (63%). 1H NMR (CDCl3 500 MHz, d ppm)
7.40 (d, J = 6.9 Hz, 2H), 7.34 (d, J = 7.3 Hz, 2H), 7.27 (m, 1H), 6.59 (d, J = 16.2 Hz,
1H), 6.29 (dd, J1 = 6.2 Hz, J2 = 15.8 Hz, 1H), 4.51 (m, J = 6.2 Hz, 1H), 1.40 (t,
J = 6.6 Hz, 3H).
4-Methyl-3-buten-2-ol, 22: yield 0.50 g (60%). 1H NMR (CDCl3 500 MHz, d
ppm) 5.22 (m, 1H), 4.56 (m, 1H), 1.73 (s, 3H), 1.71 (s, 3H), 1.23 (d, J = 7.7 Hz,
3H).
Alcohols 15, 19, and 20 were prepared from their corresponding ketones by reduc-
tion with LiAlH4.

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1
3-Methyl-2-butanol, 15: yield 0.31 g (70%). H NMR (CDCl3 500 MHz, d ppm)
3.58 (m, 1H), 1.63 (m, 1H), 1.15 (d, J = 6 Hz, 3H), 0.93 (d, J = 6.8 Hz, 3H), 0.91
(d, , J = 6.8 Hz, 3H).
1
2-Octanol, 19: yield 0.54 g (85%). H NMR (CDCl3 500 MHz, d ppm), 3.80 (m,
1H), 1.45 (m, 2H), 1.26–1.36 (m, 8H), 1.21 (d, J = 6.2 Hz, 3H), 0.90 (t,
J = 6.9 Hz, 3H).
1
2-Butanol, 20: yield 0.26 g (70%). H NMR (CDCl3 500 MHz, d ppm), 3.58 (m,
1H), 1.40 (m, 2H), 1.10 (d, J = 6.2 Hz, 3H), 0.89 (t, J = 6.3 Hz, 3H).
4.4. Acyl esterification of alcohols [32]
To a stirred mixture of 0.5 mmol of the appropriate alcohol in 30 ml of ethyl ace-
tate were added 15 mmol of distilled acetic anhydride, 15 mmol of anhydrous K2CO3
and a catalytic amount of N,N-dimethyl-amino pyridine (DMAP).The reaction mix-
ture was stirred at room temperature for 24 h. Following the addition of diethyl
ether, the mixture was extracted with saturated CuSO4Æ5H2O, saturated aqueous so-
dium bicarbonate NaHCO3 and finally washed with brine. The organic layer was
dried over MgSO4 and the corresponding acetyl ester was isolated after removal
of the solvent under vacuum in good yields.
All the prepared substrates are known compounds and were identified by their
NMR, and MS spectra.
Acetyl ester of alcohol 2: yield 0.082 g (85%). 1H NMR (CDCl3 500 MHz, d ppm)
7.31 (d, J = 8.5 Hz, 2H), 6.90 (d, J = 8.5 Hz, 2H), 5.87 (m, J = 6.5 Hz, 1H), 3.82 (s,
3H), 2.06 (s, 3H), 1.54 (d, J = 6.6 Hz, 2H). MS: m/z 152.
Acetyl ester of alcohol 3: yield 0.084 g (87%). 1H NMR (CDCl3 500 MHz, d ppm)
7.26–7.29 (t, J = 7.8 Hz, 1H), 6.95 (d, J = 6.2 Hz, 2H), 6.83 (d, J = 6.2 Hz, 1H), 5.77
(q, J = 6.4 Hz, 1H), 3.82 (s, 3H), 1.50 (d, J = 6.4 Hz, 3H).
Acetyl ester of alcohol 4: yield 0.082 g (91%). 1H NMR (CDCl3 500 MHz, d ppm)
7.33–7.36 (m, 2H), 7.03–7.06 (m, 2H), 5.8 (q, J = 6.5 Hz, 1H), 2.08 (s, 3H), 1.5 (d,
J = 6.5 Hz, 3H).
Acetyl ester of alcohol 5: yield 0.096 g (81%). 1H NMR (CDCl3 500 MHz, d ppm)
7.63 (d, J = 8.0 Hz, 2H), 7.48 (d, J = 8.0 Hz, 2H), 5.92 (m, J = 6.7 Hz, 1H), 2.11 (s,
3H), 1.56 (d, J = 6.7 Hz, 3H).
Acetyl ester of alcohol 6: yield 0.083 g (80%). 1H NMR (CDCl3 500 MHz, d ppm)
7.23 (d, J = 8.6 Hz, 2H), 7.53 (d, J = 8.6 Hz, 2H), 5.94 (m, J = 6.7 Hz, 1H), 2.13 (s,
3H), 1.57 (d, J = 6.7 Hz, 3H).
Acetyl ester of alcohol 7: yield 0.077 g (75%). 1H NMR (CDCl3 500 MHz, d ppm)
7.61 (d, J = 7.0 Hz, 2H), 7.43 (m, 1H), 7.31 (m, 1H), 5. 72 (m, 1H), 2.1 (s, 3H), 1.57
(d, J = 6.5 Hz, 3H).
Acetyl ester of alcohol 8: yield 0.101 g (90%). 1H NMR (CDCl3 500 MHz, d ppm)
7.60 (d, J = 7.9 Hz, 4H), 7.46 (t, J = 7.0 Hz, 4H), 7.37 (m, 1H), 5.96 (q, J = 6.5 Hz,
1H), 1.60 (d, J = 6.4 Hz, 3H).
Acetyl ester of alcohol 9: yield 0.070 g (85%). 1H NMR (CDCl3 500 MHz, d ppm)
8.61 (d, J = 6.0 Hz, 2H), 7.26 (d, J = 6.0 Hz, 2H), 5.84 (m, J = 6.5 Hz, 1H), 2.1 (s,
3H), 1.54 (d, J = 6.5 Hz, 3H).

N.S. Hatzakis, I. Smonou / Bioorganic Chemistry 33 (2005) 325–337
335
1
Acetyl ester of alcohol 10: yield 0.087 g (94%). H NMR (CDCl3 500 MHz, d
ppm) 7.35 (d, J = 6.5 Hz, 4H), 7.29 (m, J = 3.4 Hz, 1H), 5.69 (t, J = 6.9 Hz, 1H),
2.1 (s, 3H), 1.56 (m, J = 7.4 Hz, 1H), 1.85 (m, J = 7.1 Hz, 1H), 0.91 (t,
J = 7.4 Hz, 3H). 13C NMR (CDCl3 500 MHz, d ppm) 170.39, 140.54, 128.357,
127.79, 126.56, 29.27, 21.24, 9.88.
1
Acetyl ester of alcohol 11: yield 0.080 g (97%). H NMR (CDCl3 500 MHz, d
ppm) 7.27 (t, J = 7.4 Hz, 1H), 7.18 (d, J = 7.4 Hz, 2H), 7.13 (d, J = 7.4 Hz, 1H),
5.88 (q, J = 6.6 Hz, 1H), 2.4 (s, 3H), 2.1 (s, 3H), 1.55 (d, J = 6.6 Hz, 3H). 13C
NMR (CDCl3 500 MHz, d ppm) 170.30, 141.60, 138.10, 128.60, 128.38, 126.81,
123.09, 72.33, 22.17, 21.41.
Acetyl ester of alcohol 12: yield 0.10 g (91%). 1H NMR (CDCl3 500 MHz, d ppm)
4.52 (q, J = 6.5 Hz, 1H), 2.08 (s, 3H), 1.98 (s, 3H), 1.67 (dd, J1 = 12.1 Hz,
J2 = 38.63 Hz, 6H), 1.54 (dd, J1 = 12.1 Hz, J2 = 26.1 Hz, 6H), 1.01 (d, J = 6.5 Hz,
3H). 13C NMR (CDCl3 500 MHz, d ppm) 170.86, 77.69, 37.82, 37.06, 35.67,
28.14, 21.16, 13.34.
Acetyl ester of alcohol 14: yield 0.64 g (89%). 1H NMR (CDCl3 500 MHz, d ppm)
4.70 (q, J = 6.7 Hz, 1H), 2.1 (s, 3H), 1.15 (d, J = 6.7 Hz, 3H), 0.92 (s, 9H).
Acetyl ester of alcohol 15: yield 0.55 g (90%). 1H NMR (CDCl3 500 MHz, d ppm)
4.7 (m, J = 6.3 Hz, 1H), 2.04 (s, 3H), 1.77 (m, 1H), 1.16 (d, J = 6.3 Hz, 3H), 0.91 (d,
J = 6.7 Hz, 6H).
Acetyl ester of alcohol 16: yield 0.11 g (92%). 1H NMR (CDCl3 500 MHz, d ppm)
5.09 (m, 1H), 2.02 (s, 3H), 1.95 (s, 3H), 1.06 (dd, J1 = 11.7 Hz, J2 = 37.5, 6H), 1.52
(m, 9H), 1.20 (d, J = 6.2 Hz, 3H).
Acetyl ester of alcohol 17: yield 0.91 g (88%). 1H NMR (CDCl3 500 MHz, d ppm)
7.30–7.35 (m, 6H), 7.20–7.25 (m, 4H), 4.84 (m, 1H), 4.08 (t, J = 7.6 Hz, 1H), 2.47 (m,
1H), 2.25 (m, 1H), 1.98 (s, 3H), 1.28 (d, J = 6.2 Hz, 3H). 13C NMR (CDCl3
500 MHz, d ppm) 170.53, 128.61, 128.51, 127.79, 127.75, 126.40, 126.29, 65.59,
47.86, 41.69, 21.17, 20.32.
1
Acetyl ester of alcohol 18: yield 0.77 g (88%). H NMR (CDCl3 500 MHz, d
ppm) 7.30 (m, 2H), 7.22 (m, 3H), 5.14 (m, 1H), 2.94 (dd, J1 = 6.5 Hz,
J2 = 13.5 Hz, 1H), 2.77 (dd, J1 = 6.5 Hz, J2 = 13.5 Hz, 1H), 2.02 (s, 1H), 1.24 (d,
J = 6.5 Hz, 3H).
Acetyl ester of alcohol 19: yield 0.60 g (92%). 1H NMR (CDCl3 500 MHz, d ppm)
4.88 (m, 1H), 2.04 (s, 3H), 1.58 (m, 1H), 1.48 (m, 1H), 1.3 (m, 8H), 1.22 (d,
J = 6.2 Hz, 3H), 0.90 (t, J = 6.9 Hz, 3H).
Acetyl ester of alcohol 20: yield 0.40 g (82%). 1H NMR (CDCl3 500 MHz, d ppm)
4.35 (m, 1H), 2.01 (s, 3H), 1.68 (m, 1H), 1.55 (m, 1H), 1.23 (d, J = 6.4 Hz, 3H), 0.88
(t, J = 7.3 Hz, 3H).
Acetyl ester of alcohol 21: yield 0.77 g (82%). 1H NMR (CDCl3 500 MHz, d ppm)
7.40 (d, J = 7.3 Hz, 2H), 7.34 (d, J = 7.7 Hz, 2H), 7.27 (m, 1H), 6.62 (d, J = 15.8 Hz,
1H), 6.22 (dd, J1 = 6.6 Hz , J2 = 15.8 Hz, 1H), 5.56 (m, J = 6.6 Hz, 1H), 2.10 (s,
3H), 1.42 (t, J = 6.2 Hz, 3H).
Acetyl ester of alcohol 22: yield 0.62 g (88%). 1H NMR (CDCl3 500 MHz, d ppm)
5.6 (m, 1H), 5.18 (d, J = 7.3 Hz, 1H), 2.04 (s, 3H), 1.74 (d, J = 5.5 Hz, 6H), 1.27 (d,
J = 6.2 Hz, 3H).

336
N.S. Hatzakis, I. Smonou / Bioorganic Chemistry 33 (2005) 325–337
4.5. General procedure for the enzymatic transesterification of alcohols catalyzed by
feruloyl esterase
All enzymatic transesterification reactions were performed as follows:
100 mg (0.0435 U) of the FAE preparation and 0.277 ml (3 mmol) of dry vinyl
acetate were placed in a vial and 0.1 mmol of the (R,S)-alcohol was added. Molec-
˚
ular sieves 3 A were also added to the reaction mixture. The resulting suspension
was shaken at 400 rpm, 45 °C. Samples were collected periodically and were ana-
lyzed by gas chromatography. When the desired conversion was reached the mixture
was filtered through a paper funnel and the mixture of the residual substrate and the
resulting ester was washed three times with ethyl acetate. The solvent was removed
under vacuum and the products were separated from the remaining substrate by
flash column chromatography with 5–30% diethyl ether in hexane as eluent.
Acknowledgments
We thank the Greek Secretariat of Research and Technology (PENED 2001) and
the Ministry of Education (B-EPEAEK graduate program) for financial support and
graduate fellowship to N.S.H.
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