Lipase-catalysed chemoselective monoacetylation of hydroxyalkylphenols and chemoselective removal of a single acetyl group from their diacetates

Article abstract of DOI:10.1016/S0957-4166(98)00285-7

It was demonstrated that Pseudomonas cepacia PS lipase adsorbed on Celite, has the ability to catalyse the chemoselective monoacetylation of various hydroxyalkylphenols or the chemoselective removal of a single acetyl group from the corresponding acetate.

Full text of DOI:10.1016/S0957-4166(98)00285-7

TETRAHEDRON:
ASYMMETRY
Pergamon
Tetrahedron: Asymmetry 9 (1998) 2915–2924
Lipase-catalysed chemoselective monoacetylation of
hydroxyalkylphenols and chemoselective removal of a single
acetyl group from their diacetates
Pietro Allevi, Pierangela Ciuffreda, Alessandra Longo and Mario Anastasia ∗
Department of Medical Chemistry and Biochemistry, University of Milan, Via Saldini 50, 20133 Milan, Italy
Received 10 July 1998; accepted 17 July 1998
Abstract
It was demonstrated that Pseudomonas cepacia PS lipase adsorbed on Celite, has the ability to catalyse the
chemoselective monoacetylation of various hydroxyalkylphenols or the chemoselective removal of a single acetyl
group from the corresponding acetate. © 1998 Elsevier Science Ltd. All rights reserved.
1. Introduction
An important topic in organic synthesis is the development of chemical procedures to obtain chemo-
selectively monoacetylated compounds from molecules containing phenolic and aliphatic hydroxyl
groups, as reflected by the number of methods developed recently to effect such transformations.1–3
The literature shows this goal can be achieved by two routes, one involving the selective
monoacylation3 of the dihydroxy compounds, the other the selective removal4–6 of a single acyl
group from the diacetylated compounds. The usual methods (using acyl chlorides or anhydrides) are
generally orientated towards the acylation of phenols,3 while the selective acylation of the alcoholic
hydroxyls is better achieved by relatively recent methods based on transesterification processes that
couple heterogeneous catalysts such as Al2O3 or NaHSO4 supported on silica gel (NaHSO4·SiO2).1
7,8
In the latter case, the acetylation yield is high and the conditions required are milder (6–18 h at
67–80°C) than those reported for selective acylation using tightly twisted active amides (72–120 h
at 80°C).3 The acetyl group is selectively removed from phenols in excellent yield by treating the
phenols with guanidine5 and, in the case of the 3-hydroxybenzylalcohols, by treatment with NaBH4 in
dimethoxyethane (for 18 h at 40°C).6
∗
Corresponding author. E-mail: skinski@imiucca.csi.unimi.it
0957-4166/98/$ - see front matter © 1998 Elsevier Science Ltd. All rights reserved.
PII: S0957-4166(98)00285-7

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Less explored is the possibility of using enzymes in organic solvents to catalyse the chemoselective
acylation of hydroxyalkylphenols or the chemoselective deacetylation of the diacetates, though enzymes
have been used in the enantioselective acylation of unsymmetrical alcohols, diols or polyols,9 in the
regioselective acylation of hydroxyls in polydric phenols10,11a,b and in the regioselective deacylation of
polyacetylated phenols.12–15 In fact, to the best of our knowledge, only three papers have dealt with this
topic. One gave consideration only to the ability of some lipases, from Candida cylindracea (CCL),16 to
mediate the selective acetylation of the alcoholic groups of 2-, 3- and 4-hydroxybenzyl alcohols, together
with the ability of the same enzymes to mediate the chemoselective transesterification of the diacetate.
The acetylation reaction always occurs quantitatively and with complete and exclusive esterification of
the benzylic hydroxy group. Similarly, deacetylation on the phenolic ester takes place with satisfactory
results except for the 2-acetoxybenzyl alcohol acetate, which shows poor yield and poor selectivity.16
Another of these papers17 discusses the ability of lipase from Aspergillus niger to mediate the acetylation
of the said hydroxybenzylacohols. The reaction was difficult and a considerable amount of the starting
diols was recovered (7–23%; after 48 h). The third paper18 studied the selective enzymatic hydrolysis
of phenolic acetates in the presence of alkyl acetates. Pig liver acetone powder (PLAP) that contains
lipases was used for competitive experiments. Only the hydroxyalkylphenol diacetate was tested, 3-(2-
acetoxypropyl)phenyl acetate, resulting in a partial hydrolysis (12%) of both esters.
From these results it was concluded that the tested enzymes could catalyse, by chemoselective
acetylation, the preparation of acetoxyphenols, but were ineffective when preparing the complementary
isomeric compound (i.e. of the benzylic alcohols with an acetylated phenolic group) by deacetylation.
Thus, there appeared to be a gap in the enzyme-catalysed reactions, compared to reactions mediated by
chemical reagents.
In the course of research concerning the transformation of 40-demethylepipodophyllotoxin (1a) into
the etoposide, a phenolic anticancer O-glycoside,19a,b we searched for an enzymatic route suitable for
the facile acetylation of the alcoholic hydroxyl of 1a. This led us to an extensive study of monoacylation
in organic solvents; we considered various hydroxyalkylphenols and used lipases from Pseudomonas
cepacia PS or PLAP as catalysts in the acetylations.20,21 The next step was the study of the deacetylation
reactions of the corresponding diacetylated compounds in aqueous neutral buffer.
Our results show that Lipase PS can be used to selectively prepare phenolic or alcoholic acetates of
hydroxyalkylphenols containing one or two methoxy groups adjacent to the phenolic hydroxyl. This
selectivity occurs as the alcoholic hydroxyl prevails in enzymatic acetylation in organic solvents, while
in the case of the hydrolysis of the corresponding diacetates what prevails is the ester of the alcoholic
hydroxyl. Moreover, in unsubstituted hydroxyalkylphenols, Lipase PS causes the alcohols (in acetylation)
and the phenol esters (in deacetylation) to be more reactive, always leading to acetylated alcohols
with a free phenol, with complete chemoselectivity. Other lipases, PLAP and lipase from Candida
cylindracea,16 do not show this reversal of chemoselectivity and are, in some cases, ineffectual, while
in others they catalyse both the acetylation and the deacetylation in noticeably longer times than those
required by Lipase PS.
2. Results and discussion
First we evaluated the possibility of obtaining the two monoacetyl esters of the hydroxyalkylphenols
2a–5a, by enzyme-catalysed acetylation and/or deacetylation of the corresponding diacetates. Compound
2a, which contains a phenolic moiety similar to that present in 40-demethylepipodophyllotoxin (1a),19a
is part of a group of well known synthons used for various synthetic purposes in flavonoid or lignan

P. Allevi et al. / Tetrahedron: Asymmetry 9 (1998) 2915–2924
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Table 1
Lipase-catalysed chemoselective acetylation (Ac) of diols (2a–5a) and deacetylation (D) of the
corresponding diacetates (2d–5d)
chemistry.22–24 In addition, some of these congeries/synthons are used as model compounds to evaluate
chemoselective acylations performed by chemical methods.3
In separate experiments, the mono and dimethoxylated hydroxyalkylphenols 2a–5a were subjected
to acetylation with vinyl acetate in diisopropyl ether, catalysed by Lipase PS or PLAP. In all cases,
the alcohol acetylation prevailed, resulting in excellent chemoselectivity regardless of the enzyme used
(Table 1).
Lipase PS showed very high activity, instead PLAP was lower and required noticeably longer reaction
times.25
When the corresponding diacetates 2d–5d were subjected to deacetylation in acetone–aqueous phos-
phate buffer (pH=7), in the presence of free Lipase PS or PLAP (Table 1), the chemoselectivity for the
enzymes was, in two cases, opposing. In fact, Lipase PS catalyses the chemoselective hydrolysis of the
alcoholic acetate esters of 3d and 5d, while PLAP catalyses the hydrolysis of the phenolic esters or, in
the case of 2d and 4d, of both esters.25
On the basis of these results, it can be seen that the same compounds 2b–5b, with acetylated

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alcohols and a free phenol group, can be obtained in good yield by acetylation of the corresponding
diols under Lipase PS or PLAP catalysis, or by the hydrolysis of the corresponding diacetates in the
presence of PLAP. Conversely, the isomeric monoacetylated phenols 2c, 3c and 5c are obtained in
good yield in deacetylation reactions under Lipase PS catalysis. Only in the case of the 4-acetoxy-
3-methoxybenzylalcohol acetate 4d did the deacetylation mediated by Lipase PS afford 4-hydroxy-3-
methoxybenzylalcohol acetate 4b (43% yield), accompanied by the 4-hydroxy-3-methoxybenzyl alcohol
4a (53% yield). The same deacetylation under PLAP catalysis afforded the phenolic benzylalcohol
acetate 4b (in 82% yield), accompanied by a minor amount of the dihydroxycompound 4a (14% yield).
These encouraging results prompted us to evaluate Lipase PS and PLAP behaviour in similar reactions
but involving simpler hydroxyalkylphenols. Thus, a study was made of three classes of compounds,
corresponding to the possible reciprocal positions in a benzene ring (ortho, meta and para) of a phenolic
hydroxy group and an alkyl chain bearing the alcoholic function. The alkyl chains chosen were of three
different lengths (from one to three carbon atoms) in order to also assess the influence of the distance
between the molecules’ hydroxyls on the reaction course.
The results, summarised in Table 2, show that all the tested hydroxy compounds (ortho: 6a, 6e and
6i; meta: 7a, 7e and 7i and para: 8a, 8e and 8i) are chemoselectively acetylated at the alcoholic hydroxy
group affording, exclusively, phenols with an esterified alcoholic group (6b, 6f and 6j; 7b, 7f and 7j;
8b, 8f and 8j); this occurs irrespective of the reciprocal position and distance between the two hydroxyls
in the starting compound. In no case was acetylation of the phenol observed, even on prolonging the
reaction time to several hours after the disappearance of the starting substrate.
PLAP also catalysed the acetylation of the alcoholic hydroxyl, but very long times were needed (the
reactions were incomplete even after 3 days) and are not reported in Table 2.
In the deacetylation of the diacetates (ortho: 6d, 6h and 6l; meta: 7d, 7h and 7l and para: 8d, 8h and 8l),
regardless of the enzyme catalyst used, the regeneration of the phenolic group always prevailed, the only
exception being the case of the hydroxymethylphenol diacetates 6d, 7d and 8d where some dihydroxy
compound (5–30%) accompanied the benzylic alcohol ester. In these cases, prolonging the reaction time
after the disappearance of the starting diacetate results in dihydroxy compounds as exclusive reaction
products.
Thus, with non-hindered phenols, Lipase PS and PLAP show similar behaviour and selectivity, the
selectivity increasing with the length of the alkyl chain, regardless of its position with respect to the
phenol hydroxyl.
The reported results show that the ability of enzymes to discriminate between alcoholic and phenolic
groups in acetylation reactions, and between acetates in deacetylation reactions, has marked potential in
organic syntheses. Indeed enzymatic methods could be used as a good alternative to current chemical
methods for obtaining the same results.
In our laboratory, we used enzymes to prepare 40-demethylepipodophyllotoxin 4-acetate 1b and α- and
β-estradiol 17-acetates. The acetate 1b was obtained in quantitative yield by hydrolysis of its diacetate

P. Allevi et al. / Tetrahedron: Asymmetry 9 (1998) 2915–2924
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Table 2
Lipase-catalysed chemoselective acetylation (Ac) of hydroxyalkylphenols of type (6–8) and
deacetylation (D) of the corresponding diacetates
1d mediated by PLAP, however lipase PS did not catalyse the deacetylation of compound 1d. The
two epimeric estradiol 17-acetates were obtained by chemoselective deacetylation of the corresponding
diacetates mediated by either PLAP or Lipase PS. These compounds are of interest for synthetic
studies in our laboratory,19a,b,26 and have already been obtained by more tedious chemical procedures
involving the selective protection of the phenolic group, acetylation of the alcohol and deprotection of
the phenol.19a,27,28
In the pioneering work done by Caspi and Njar29 in this area the α-estradiol 17-acetate yield was poor
(25%), while that of the β-estradiol 17 acetate was good (90%). These pioneers obtained the two isomers

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by the enzymatic transesterification of the corresponding diester, mediated by Candida cylindracea in
organic solvents.
In conclusion, though it is still not possible to predict the general course of the enzyme-catalysed reac-
tions of hydroxyalkylphenols, Lipase PS merits some reflection. In the case of the hydroxyalkylphenols
with unhindered alcoholic and phenolic hydroxyls, Lipase PS preferentially catalyses the acetylation
of the alcohol, but in the hydrolysis of the corresponding diacetates it shows chemoselectivity for the
phenolic ester. With an unhindered alcoholic group and a hindered phenolic hydroxyl, Lipase PS still
catalyses the acetylation of the alcoholic group but in the hydrolysis of the corresponding diacetate the
chemoselectivity reverts and hydrolysis of the alcoholic ester is catalysed. Thus, Lipase PS leads to
complementary isomeric products in acetylation–deacetylation steps. On the contrary, when both the
alcoholic and the phenolic hydroxyls are equally hindered (as in 1d), Lipase PS is unable to catalyse
the acetylation or the hydrolysis of the corresponding esters. Both PLAP and CCL are less active than
Lipase PS, however when they are effective, they catalyse the alcoholic group acetylation and phenol
ester hydrolysis or the hydrolysis of both esters.
Also of interest is the behaviour of Lipase PS. It changes its chemopreference not only on the basis of
the steric hindrance but also on the basis of the position of the phenol group. Such knowledge leads to a
better understanding of the characteristics of this enzyme.
3. Experimental section
1H NMR spectra (500.13 MHz) were recorded in CDCl3 at 303 K, the reference being CHCl3 at 7.24
ppm. Mass spectra (MS) were recorded on a Hewlett Packard 5988A mass spectrometer using a direct
inlet at an ionising voltage of 70 eV by electron impact. Mass spectra were reported as m/z (relative
abundance). HPLC analyses were carried out on a Merck superspher 100 RP-18 column (4 mm×15 cm;
eluent H2O:MeOH, 50:50; v/v). The flow rate was 1 mL/min and detection was performed at 270 nm.
GLC analyses were carried out on a SPB-5 Supelco column (30 m×0.32 mm; 0.25 µm film thickness):
column temperature, initially set at 110°C for 1 min, increased by 20°C/min until 280°C where it was
kept for 2 min. TLC was carried out on silica gel 60 F254 microplates eluting with hexane:ethyl acetate
(60:40; v/v). Column chromatography refers to flash chromatography.30 The progress of all the reactions,
column chromatography and compound purity were monitored by TLC, GLC and/or HPLC.
Particular attention was given to the course of the reaction as the starting diacetates and acetoxyphenols
used show very similar Rf in the best eluent used (hexane:ethyl acetate, 60:40; v/v), as do the diols and
alcohols with an acetylated phenol. Experiments with lipase from CCL were performed according to
the general conditions described by Servi and Pedrocchi.16 All organic solvents were dried before use.
Starting substrates were obtained as follows and showed satisfactory elemental analyses and significant
spectroscopic data (IR, 1H NMR and mass spectra).
The starting diols 4a–8a, 6e–8e, 8i, 17α- and 17β-estradiol were purchased (Aldrich). Diols 2a and 3a
were obtained by NaBH4 reduction of the corresponding aldehydes (Aldrich), compound 6i was obtained
by LiAlH4 reduction of dihydrocoumarin (Aldrich) at 25°C, and the diol 7i was prepared from 3-(3-
methoxyphenyl)propionic acid (Aldrich), described below; 40-demethylepipodophillotoxin was obtained
from commercial podophyllotoxin.19a The acetates 1d31–8d, 6h–8h, 6l–8l and those of 17α- and 17β-
estradiol were obtained by usual acetylation with acetic anhydride in pyridine and were identified by
their physical and/or spectroscopic data. All compounds showed correct elemental analyses.

P. Allevi et al. / Tetrahedron: Asymmetry 9 (1998) 2915–2924
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3.1. Preparation of 3-(3-hydroxypropyl)phenol (7i)
3-(3-Methoxyphenyl)propionic acid (600 mg; 3.33 mmol) was dissolved in CH3CN (5 mL) and added
to NaI (599 mg; 3.99 mmol) and (CH3)3SiCl (0.5 mL; 3.99 mmol). The mixture was heated at 60°C for 4
h, quenched with a saturated solution of NaCl and extracted with CHCl3. The organic layers were washed,
dried and evaporated to afford a crude product (560 mg) which, without purification, was dissolved in
methanol (5 mL) and treated with H2SO4 (0.5 mL; 96%), for 12 h at 25°C. The solution was then
concentrated under reduced pressure, poured into an ice cold aqueous solution of NaOH (pH=12) and
washed with ethyl acetate. The aqueous layer was then acidified with HCl (2 M) and extracted with ethyl
acetate. The obtained organic layers were dried and the solvent evaporated to afford the crude methyl
ester (310 mg). This was dissolved in anhydrous diethyl ether (10 mL) and treated with LiAlH4 (150 mg)
at 25°C for 6 h to give, after usual work-up, the diol 7i (200 mg): a viscous oil, IR (neat) 3380, 1580, 780
cm−1; 1H NMR δ 7.12 (1H, dd, J=8.0, 8.0 Hz), 6.72 (1H, d, J=8.0 Hz), 6.67–6.64 (2H, m), 3.62 (2H, t,
J=7.5 Hz, CH2OH), 2.61 (2H, t, J=7.5 Hz, ArCH2), 1.92 (2H, tt, J=7.5, 7.5 Hz, CH2); MS m/z 152 (M+,
26), 133 (15), 121 (9), 108 (100), 107 (55), 91 (23), 77 (49).
3.2. Enzyme-mediated acetylation of diols. General procedure
Vinyl acetate (0.18 mL; 2 mmol) and enzyme (Lipase PS: 125 mg or PLAP: 125 mg) were added to a
solution of the diol (1 mmol) in diisopropyl ether (20 mL). The resulting suspension was shaken at 25°C
and monitored by TLC, GLC and/or HPLC. After the consumption of the starting diol the reaction was
quenched by filtering off the enzyme and the solvent was then removed in vacuo to give the identified
compound(s).
3.3. Enzyme-mediated deacetylation of diacetates. General procedure
The diacetate (1 mmol) was dissolved in acetone (4 mL) and added to phosphate buffer (20 mL; pH=7)
and the enzyme (free Lipase PS: 65 mg, or PLAP: 125 mg). The resulting suspension was shaken at 25°C
and monitored by TLC, GLC and/or HPLC. After consumption of the starting diacetate the reaction was
quenched by adding ethyl acetate and the resulting mixture was filtered through a pad of Celite. The
organic layer was then separated and dried and the solvent removed in vacuo to give a crude residue
which, if necessary, was chromatographed to afford the identified compound(s).
3.3.1. 2,6-Dimethoxy-4-(acetoxymethyl)phenol (2b)
1
IR (neat) 3420, 1730 cm−1; H NMR δ 6.58 (2H, s, H-3 and H-5), 5.56 (1H, s, OH), 4.98 (2H, s,
CH2), 3.86 (6H, s, 2×OCH3), 2.06 (3H, s, OCOCH3); MS m/z 226 (M+, 23), 182 (50), 167 (100), 148
(44), 136 (29), 123 (31), 107 (19), 95 (24), 79 (27), 65 (32).
3.3.2. 2,6-Dimethoxy-4-(hydroxymethyl)phenyl acetate (2c)
1
IR (neat) 3430, 1770 cm−1; H NMR δ 6.60 (2H, s, H-3 and H-5), 4.61 (2H, s, CH2), 3.79 (6H, s,
2×OCH3), 2.30 (3H, s, OCOCH3); MS m/z 226 (M+, 3), 184 (100), 167 (24), 155 (15), 141 (8), 123 (25),
109 (21), 95 (17), 81 (15), 53 (17).

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P. Allevi et al. / Tetrahedron: Asymmetry 9 (1998) 2915–2924
3.3.3. 2,3-Dimethoxy-5-(acetoxymethyl)phenol (3b)
IR (neat) 3420, 1725 cm−1; 1H NMR δ 6.59 (1H, d, J=2.1 Hz), 6.45 (1H, d, J=2.1 Hz), 5.82 (1H, s,
OH), 4.97 (2H, s, CH2), 3.87 (3H, s, OCH3), 3.84 (3H, s, OCH3), 2.08 (3H, s, OCOCH3); MS m/z 226
(M+,4), 184 (83), 169 (9), 167 (20), 141 (15), 109 (100), 95 (37), 81 (48), 53 (39).
3.3.4. 2,3-Dimethoxy-5-(hydroxymethyl)phenyl acetate (3c)
IR (neat) 3435, 1770 cm−1; 1H NMR δ 6.80 (1H, d, J=2.1 Hz), 6.63 (1H, d, J=2.1 Hz), 4.57 (2H, s,
CH2), 3.84 (3H, s, OCH3), 3.79 (3H, s, OCH3), 2.29 (3H, s, OCOCH3); MS m/z 226 (M+, 15), 184 (100),
169 (9), 167 (11), 141 (19), 109 (74), 95 (21), 81 (21), 53 (20).
3.3.5. 2-Methoxy-4-(acetoxymethyl)phenol (4b)
IR (neat) 3430, 1735 cm−1; 1H NMR δ 6.88–6.85 (3H, m, H-2, H-3 and H-5), 5.70 (1H, s, OH), 5.00
(2H, s, CH2), 3.87 (3H, s, OCH3), 2.06 (3H, s, OCOCH3); MS m/z 196 (M+, 32), 154 (33), 137 (100),
122 (32), 107 (22), 94 (21), 93 (26), 65 (51).
3.3.6. 2-Methoxy-5-(acetoxymethyl)phenol (5b)3,6
1
IR (neat) 3440, 1730 cm−1; H NMR δ 6.92 (1H, d, J=2.1 Hz, H-6), 6.83 (1H, dd, J=2.1, 7.7 Hz,
H-4), 6.80 (1H, d, J=7.7 Hz, H-3), 5.67 (1H, s, OH), 4.98 (2H, s, CH2), 3.86 (3H, s, OCH3), 2.06 (3H, s,
OCOCH3); MS m/z 196 (M+, 33), 154 (54), 137 (100), 125 (8), 122 (38), 93 (30), 65 (13).
3.3.7. 2-Methoxy-5-(hydroxymethyl)phenyl acetate (5c)3
1
IR (neat) 3400, 1770 cm−1; H NMR δ 7.16 (1H, dd, J=2.1, 7.7 Hz, H-4), 7.06 (1H, d, J=2.1 Hz,
H-6), 6.92 (1H, d, J=7.7 Hz, H-3), 4.57 (2H, s, CH2), 3.80 (3H, s, OCH3), 2.29 (3H, s, OCOCH3); MS
m/z 196 (M+, 7), 154 (100), 137 (27), 125 (19), 122 (18), 93 (36), 65 (25).
3.3.8. 2-(Acetoxymethyl)phenol (6b)16
1
IR (neat) 3380, 1740 cm−1; H NMR δ 7.28–7.22 (2H, m), 6.92 (1H, d, J=7.7 Hz), 6.89 (1H, dd,
J=7.7, 7.7 Hz), 5.10 (2H, s, CH2), 2.09 (3H, s, OCOCH3); MS m/z 166 (M+, 33), 137 (11), 107 (100),
106 (96), 78 (96), 77 (56).
3.3.9. 2-(2-Acetoxyethyl)phenol (6f)1
IR (neat) 3250, 1700 cm−1; 1H NMR δ 7.14–7.06 (2H, m), 6.84 (1H, dd, J=7.7, 7.7 Hz), 6.80 (1H, d,
J=7.7 Hz), 4.27 (2H, t, J=7.0 Hz, CH2OAc), 2.94 (2H, t, J=7.0 Hz, ArCH2), 2.05 (3H, s, OCOCH3); MS
m/z 180 (M+, 2), 137 (3), 120 (100), 107 (42), 91 (30), 77 (25).
3.3.10. 2-(3-Acetoxypropyl)phenol (6j)
IR (neat) 3400, 1720 cm−1; 1H NMR δ 7.08 (1H, dd, J=2.1, 7.7 Hz), 7.05 (1H, ddd, J=2.1, 7.7, 7.7
Hz), 6.84 (1H, dd, J=7.7, 7.7 Hz), 6.75 (1H, d, J=7.7 Hz), 5.08 (1H, s, OH), 4.10 (2H, t, J=7.0 Hz,
CH2OAc), 2.69 (2H, t, J=7.0 Hz, ArCH2), 2.05 (3H, s, OCOCH3), 1.97 (2H, tt, J=7.0, 7.0 Hz, CH2); MS
m/z 194 (M+, 4), 134 (100), 133 (39), 119 (38), 117 (8), 107 (43), 91 (35), 77 (42).
3.3.11. 3-(Acetoxymethyl)phenol (7b)16
IR (neat) 3380, 1730 cm−1; 1H NMR δ 7.19 (1H, dd, J=7.7, 7.7 Hz, H-5), 6.89 (1H, d, J=7.7 Hz, H-4),
6.82 (1H, d, J=2.1 Hz, H-2), 6.78 (1H, dd, J=2.1, 7.7 Hz, H-6), 5.99 (1H, s, OH), 5.04 (2H, s, CH2), 2.09
(3H, s, OCOCH3); MS m/z 166 (M+, 42), 137 (3), 107 (90), 106 (29), 78 (74), 77 (100).

P. Allevi et al. / Tetrahedron: Asymmetry 9 (1998) 2915–2924
2923
3.3.12. 3-(2-Acetoxyethyl)phenol (7f)
IR (neat) 3380, 1700 cm−1; 1H NMR δ 7.14 (1H, dd, J=7.7, 7.7 Hz, H-5), 6.75 (1H, d, J=7.7 Hz, H-4),
6.69 (1H, dd, J=2.1, 7.7 Hz, H-6), 6.67 (1H, d, J=2.1 Hz, H-2), 5.52 (1H, s, OH), 4.26 (2H, t, J=7.0 Hz,
CH2OAc), 2.87 (2H, t, J=7.0 Hz, ArCH2), 2.03 (3H, s, OCOCH3); MS m/z 180 (M+, 1), 137 (2), 120
(100), 107 (16), 91 (14), 77 (18).
3.3.13. 3-(3-Acetoxypropyl)phenol (7j)
IR (neat) 3380, 1710 cm−1; 1H NMR δ 7.12 (1H, dd, J=7.7, 7.7 Hz, H-5), 6.72 (1H, d, J=7.7 Hz, H-4),
6.66 (1H, dd, J=2.1, 7.7 Hz, H-6), 6.64 (1H, d, J=2.1 Hz, H-2), 5.60 (1H, s, OH), 4.07 (2H, t, J=7.0 Hz,
CH2OAc), 2.61 (2H, t, J=7.0 Hz, ArCH2), 2.04 (3H, s, OCOCH3), 1.92 (2H, tt, J=7.0, 7.0 Hz, CH2); MS
m/z 194 (M+, 6), 134 (100), 133 (97), 119 (16), 117 (77), 107 (71), 91 (40), 77 (44).
3.3.14. 4-(Acetoxymethyl)phenol (8b)16
IR (neat) 3380, 1725 cm−1; 1H NMR δ 7.22 (2H, d, J=7.7 Hz), 6.80 (2H, d, J=7.7 Hz), 5.71 (1H, s,
OH), 5.02 (2H, s, CH2), 2.07 (3H, s, OCOCH3); MS m/z 166 (M+, 15), 137 (2), 124 (17), 107 (100), 78
(30), 77 (26).
3.3.15. 4-(2-Acetoxyethyl)phenol (8f)1,16
IR (neat) 3380, 1700 cm−1; 1H NMR δ 7.04 (2H, d, J=8.4 Hz), 6.75 (2H, d, J=8.4 Hz), 5.72 (1H, s,
OH), 4.22 (2H, t, J=7.0, CH2OAc), 2.84 (2H, t, J=7.0 Hz, ArCH2), 2.02 (3H, s, OCOCH3); MS m/z 180
(M+, 0.2), 137 (4), 120 (100), 107 (63), 91 (10), 77 (21).
3.3.16. 4-(3-Acetoxypropyl)phenol (8j)
IR (neat) 3380, 1710 cm−1; 1H NMR δ 7.00 (2H, dd, J=7.7 Hz), 6.74 (2H, d, J=7.7 Hz), 5.70 (1H, s,
OH), 4.06 (2H, t, J=7.0 Hz, CH2OAc), 2.59 (2H, t, J=7.0 Hz, ArCH2), 2.04 (3H, s, OCOCH3), 1.89 (2H,
tt, J=7.0, 7.0 Hz, CH2); MS m/z 194 (M+, 6), 134 (80), 133 (76), 119 (8), 117 (6), 107 (100), 91 (14), 77
(33).
3.3.17. 40-Demethylepipodophyllotoxin 4-acetate (1b)
M.p. 172–174°C; was identical with that reported.28 1H NMR δ 6.84 (1H, s, H-5), 6.53 (1H, s, H-8),
6.26 (2H, s, H-20 and H-60), 6.12 (1H, d, J=3.5 Hz, H-4), 5.97, 5.94 (2H, 2×d, J=1.0 Hz, OCH2O), 4.63
(1H, d, J=5.0 Hz, H-1), 4.31 (1H, dd, J 9.5 and J=7.0 Hz, H-11a), 3.87 (1H, dd, J=11.0, 9.5 Hz, H-11b),
3.75 (6H, s, 30-OCH3 and 50-OCH3), 3.19 (1H, dd, J=14.0, 5.0 Hz, H-2), 2.95 (1H, dddd, J=14.0, 11.0,
7.0, 3.5 Hz, H-3), 2.08 (3H, s, OCOCH3).
3.3.18. 17β-Acetoxy-1,3,5(10)-estratrien-3-ol
M.p. 217–220°C; all chemicophysical properties were identical with an authentic sample.1,3b
3.3.19. 17α-Acetoxy-1,3,5(10)-estratrien-3-ol
M.p. 178–180°C; all chemicophysical properties were identical with an authentic sample.32

2924
P. Allevi et al. / Tetrahedron: Asymmetry 9 (1998) 2915–2924
Acknowledgements
We would like to thank G. Gritti for the technical assistance and A. Lorenzi for the mass spectra, and
gratefully acknowledge the Italian Ministero dell’Università e della Ricerca Scientifica e Tecnologica
(MURST) for the financial support of this work.
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