Scalable green approach toward fragrant acetates

Article abstract of DOI:10.3390/molecules25143217

The advantageous properties of ethylene glycol diacetate (EGDA) qualify it as a useful substitute for glycerol triacetate (GTA) for various green applications. We scrutinised the lipase-mediated acetylation of structurally diverse alcohols in neat EGDA furnishing the range of naturally occurring fragrant acetates. We found that such enzymatic system exhibits high reactivity and selectivity towards activated (homo) allylic and non-activated primary/secondary alcohols. This feature was utilised in the scalable multigram synthesis of fragrant (Z)-hex-3-en-1-yl acetate in 70percent yield. In addition, the Lipozyme 435/EGDA system was also found to be applicable for the chemo-selective acetylation of (hydroxyalkyl) phenols as well as for the kinetic resolution of chiral secondary alcohols. Lastly, its discrimination power was demonstrated in competitive experiments of equimolar mixtures of two isomeric alcohols. This enabled the practical synthesis of 1-pentyl acetate isolated as a single product in 68percent yield from the equimolar mixture of 1-pentanol and 3-pentanol.

Full text of DOI:10.3390/molecules25143217

molecules
Article
Scalable Green Approach Toward Fragrant Acetates
Eva Puchl’ová and Peter Szolcsányi *
Department of Organic Chemistry, Slovak University of Technology, Radlinského 9, 81237 Bratislava, Slovakia;
eva.puchlova@stuba.sk
*
Correspondence: peter.szolcsanyi@stuba.sk
ꢀ
ꢁꢂꢀꢃꢄꢅꢆꢇ
ꢀ
ꢁꢂꢃꢄꢅꢆ
Received: 18 June 2020; Accepted: 13 July 2020; Published: 14 July 2020
Abstract: The advantageous properties of ethylene glycol diacetate (EGDA) qualify it as a
useful substitute for glycerol triacetate (GTA) for various green applications. We scrutinised the
lipase-mediated acetylation of structurally diverse alcohols in neat EGDA furnishing the range of
naturally occurring fragrant acetates. We found that such enzymatic system exhibits high reactivity
and selectivity towards activated (homo) allylic and non-activated primary/secondary alcohols. This
feature was utilised in the scalable multigram synthesis of fragrant (Z)-hex-3-en-1-yl acetate in
70% yield. In addition, the Lipozyme 435/EGDA system was also found to be applicable for the
chemo-selective acetylation of (hydroxyalkyl) phenols as well as for the kinetic resolution of chiral
secondary alcohols. Lastly, its discrimination power was demonstrated in competitive experiments of
equimolar mixtures of two isomeric alcohols. This enabled the practical synthesis of 1-pentyl acetate
isolated as a single product in 68% yield from the equimolar mixture of 1-pentanol and 3-pentanol.
Keywords: enzymatic acetylation; green solvents; chemo-selectivity
1
. Introduction
In perfumery and the flavour industry, acetates are the most important aliphatic esters used and
valued for their typical aromas [1]. Although most of them are natural products, their isolation from
sources of origin makes the process economically unfeasible. Therefore, numerous chemical approaches
toward fragrant acetates were successfully developed using homogenous or heterogeneous acid, base,
or transition-metal catalysis [2]. However, perfumery and flavour grade esters need to be free of
inorganic impurities and undesirable colouration. Moreover, increasing the health consciousness and
environmental issues strongly encourages the development of benign yet efficient biocatalytic protocols
as viable green alternatives [
to their high reactivity and remarkable selectivity under diverse reaction conditions [
their stability in a non-aqueous environment, particularly under solvent-free conditions, gives lipases
a comparative advantage over other biocatalysts [ ]. In this context, glycerol-based solvents [
3
]. Among them, the use of lipases gained considerable attention due
4]. In addition,
5
6–8]
and/or reagents represent a green alternative to commonly used reaction media. One of the perspective
glycerol’s derivatives is triacetin (glycerol triacetate, GTA), which was recently used both as a reagent
and solvent for the enzymatic acetylation of some alcohols [9–12]. A similar and commercially available
derivative, ethylene glycol diacetate (EGDA) can be considered an interesting alternative to GTA
for analogous transformations. With the lower boiling point and viscosity, better recyclability, and
regeneration, EGDA has a promising synthetic potential for the scalable preparation of natural acetates
for perfumery and/or flavour purposes.
Therefore, we have decided to scrutinise the practical use of EGDA for the enzymatic acetylation
of selected alcohols that would afford known and naturally occurring fragrant acetates. In addition, we
were interested in the chemo-selectivity of such a reaction system with respect to the structure and/or
reactivity of substrates of commercial interest.
Molecules 2020, 25, 3217; doi:10.3390/molecules25143217
www.mdpi.com/journal/molecules

Molecules 2020, 25, 3217
2 of 11
2
. Results and Discussion
The initial reaction screening was done on (Z)-hex-3-ene-1-ol
1
as a model substrate, which was
acetylated [13
–15] either in neat EGDA or with the cosolvent (MTBE or hexane) using Lipozyme 435
as a biocatalyst to afford (Z)-hex-3-ene-1-yl acetate
2
. The enzyme and EGDA loadings as well as
temperature, concentration, and co-solvent were scrutinised as variable parameters while the progress
of the reaction was monitored by GC-FID (see Table S1 in the Supporting Information). The identified
optimal reaction conditions were then applied for the scale-up preparation of fragrant acetate
2 in 70%
yield. It is noteworthy that the isolation and purification protocol involves environmentally benign
procedures only (filtration, hydrodistillation, and vacuum rectification) and is free of chromatography
and/or any additional organic solvents (Scheme 1).
Scheme 1. Optimised preparation of fragrant (Z)-hex-3-ene-1-ol 1. Reagents and conditions: (a) Ethylene
◦
glycol diacetate (EGDA) (2 equiv), Lipozyme 435 (2% wt), 40 C, 150 rpm, 70% yield (20 g scale).
Next, we have applied the optimal reaction conditions in the acetylation screening of various
alcohols (Scheme 2) that furnish naturally occurring fragrant acetates (entries 1–14, Table 1). In parallel,
we have investigated the chemo-selectivity of such an enzymatic system for discrimination between
structurally different alcohols by adding more substrates (entries 15–20, Table 1)—regardless of aroma
properties of their acetates—to the screening for comparison purposes. The obtained results clearly
show the following reactivity patterns.
(
a) Considering the activated substrates, allylic primary alcohols
–5, 7, 8) were almost quantitatively esterified (GC-FID ratios of alcohol to acetate ranging
from 10:90 to 4:96) to fragrant acetates, namely geranyl acetate 16 34], prenyl acetate
41], phytyl acetate (E/Z = 66:34) 14 [42], and
1R)-nopyl acetate 16, respectively. On the other hand, rac-linalool 11 remained almost intact with
3, 5, 7, 9, 13, and 15 (entries
2
4
[
–
6,
(
(
E)-2-hexenyl acetate 8, cinnamyl acetate 10 [35–
negligible acetylation only (entry 6). Furthermore, the enzyme clearly discriminates between the
primary and tertiary allylic alcohol. Analogously, the primary homoallylic alcohols and 17 also
1
provided corresponding fragrant acetates (entries 1 and 9), namely (Z)-hex-3-ene-1-yl acetate
and (1R)-myrtenyl acetate 18.
2
(
b) Regarding the non-activated substrates, the enzyme clearly distinguishes primary alcohols 27
, 29
(
entries 15, 16), and/or secondary alcohols 19 21 23 33, and 35 (entries 10–13, 18, 19) vs. phenolic
,
,
,
eugenol 31 (entry 17) and/or tertiary alcohol 37 (entry 20). While the former ones were reasonably
acetylated (although not as well as activated alcohols, vide supra) to their corresponding acetates
2
0
,
22
the enzyme
21 (entries 10, 12). In the case of rac-2-heptanol 19 (entry 10), (R)-enantiomer was mostly
acetylated [43 49] as shown by negligible formation of (S)-2-heptenyl acetate (S)-20 (see Figure
,
24
,
28
, 30, 34, and 36, the latter ones were fully reluctant to the transformation. In addition,
´
s enantio-discrimination led to the kinetic resolution of racemic secondary alcohols
19,
–
S1 in the Supporting Information), and, thus, by obeying the Kazlauskas’s rule [50] (entry 11).
Conversely, (1R)-endo-fenchol 25 remained practically intact. This might be due to the significant
steric congestion around the hydroxyl group and/or the result of the enantiomer preference of the
enzyme for the opposite enantiomer (entry 14). In addition, the enzymatic acetylation [51–53]

Molecules 2020, 25, 3217
3 of 11
of bifunctional rac-rhododendrol 21 exhibited both the chemo-selectivity (alcohol vs. phenol)
and the kinetic resolution of racemate (entry 12). Thus, the secondary hydroxyl group of the
(R)-enantiomer (cf. the Kazlauskas’s rule again) was mostly acetylated to furnish the ester (R)-22
along with the enantiomerically enriched (S)-rhododendrol (S)-21 (84% ee, chiral HPLC, see
Figure S2 in the Supporting Information). The clear unreactivity of phenols under the reaction
conditions was also exhibited by zero acetylation of eugenol 31 (entry 17).
(
c) Considering the achiral secondary alcohols, even minor structural differences led to a significant
change in their reactivity. While isopropanol 35 (entry 19) was acetylated in a major extent
(
(
GC-FID ratio 35
GC-FID ratio 33 34 = 45:55). The extent of acetylation of 2-indanol 23 (entry 13), being somewhere
24 = 24:76), might indicate the high sensitivity of the respective
/36 = 15:85), slightly larger pentan-3-ol 33 (entry 18) was much less reactive
/
in between (GC-FID ratio 23
/
lipase to the steric demands of the alcohols in question.
Scheme 2. Reaction screening of the enzymatic acetylation. Reagents and conditions: (a) EGDA (2
◦
equiv), Lipozyme 435 (2% wt), 40 C, 150 rpm, Table 1.
Next, we aimed to practically utilise the observed chemo-selectivity of the enzymatic system,
and, therefore, we tested its discrimination power on two-component mixtures of similar yet different
alcohols. Thus, we have performed competitive acetylation experiments with equimolar mixtures of
four pairs of alcohols: (a) 1-pentanol 39 vs. 3-pentanol 33, (b) 3-pentanol 33 vs. cyclopentanol 41, (c)
prenol 5 vs. divinylcarbinol 43, and (d) 3-pentanol 33 vs. divinylcarbinol 43 (Scheme 3 and Table 2).
The obtained results clearly show that lipase-catalysed acetylation provided excellent discrimination
between primary and secondary alcohols in favour of the former ones (Table 2, entries 1 and 3).
Thus, primary 1-pentanol 39 was almost exclusively acetylated to 1-pentyl acetate 40 (GC-FID ratio
3
9
/
40 = 6:94) at the expense of its secondary isomer 3-pentanol 33 that barely afforded its acetate 34
GC-FID ratio 33 34 = 96.5:3.5). Similarly, primary allylic prenol was overwhelmingly acetylated to
prenyl acetate (GC-FID ratio = 9:91) in the presence of secondary divinylcarbinol 43 that was
concomitantly transformed to its acetate 44 in a minor extent only (GC-FID ratio 43 44 = 81:19). On the
(
/
5
6
5/6
/
other hand, the chemo-selectivity of competitive enzymatic acetylation of cyclic vs. acyclic secondary
alcohols was less pronounced (Table 2, entry 2). Although cyclopentanol 41 was acetylated much faster
to its acetate 42 (GC-FID ratio 41
/42 = 19:81) than acyclic 3-pentanol 33 to 3-pentyl acetate 34 (GC-FID
ratio 33 34 = 80:20) due to the necessary longer reaction time, the discrimination factor was inevitably
/
reduced. On the other hand, the preferential acetylation of 41 over 33 might be attributed to the higher
reactivity of less sterically hindered and more nucleophilic hydroxyl group of cyclopentanol 41 when
compared to conformationally more flexible 3-pentanol 33. Lastly, the reactivity of non-activated
(alkylic) vs. activated (allylic) secondary alcohols was comparable. This was evident from the
competitive acetylation of an equimolar mixture of 3-pentanol 33 and divinylcarbinol 43, which
generated their corresponding acetates 34 and 44 without significant discrimination between each
other (Table 2, entry 4). The time-dependent evolution of the ratio between forming acetates is depicted
in Figure 1. For the corresponding analytical data, see Tables S3–S6 in the Supporting Information.

Molecules 2020, 25, 3217
4 of 11
Table 1. Screening of the enzymatic acetylation of alcohols according to Scheme 2.
Entry
Alcohol
Acetate
Time (h)
GC-FID Ratio a (%)
OH
OAc
1
19
6:94
5:95
(
1)
(
2)
OH
OAc
(3)
2
(4)
24
3
4
5
23
24
48
4:96
5:95
8:92
(
5)
(7)
OAc
OAc
OAc
(6)
OH
OH
OH
(8)
(9)
(10)
OH
OAc
6
7
(11)
72
24
91:9
4:96
(12)
OH
OAc
(
13) E/Z = 66/34
(14) E/Z = 66/34
OH
OAc
8
9
24
24
10:90
6:94
(
15)
(16)
OH
OAc
(
17)
(18)
OH
OAc
1
1
0
1
(19)
(R)-20
(S)-20
20
72
48:52
91:9
OH
OAc
(S)-19
OH
OAc
1
1
1
2
3
4
(21)
(R)-22
48
72
48
52:48
24:76
98:2
HO
HO
OH
(23)
OAc
(24)
(
25)
(26)
OH
OAc
1
1
5
6
OH
OH
OAc
OAc
72
42
14:86
11:89
(
27)
(28)
(
29)
(
30)
OH
O
OAc
(32)
(
31)
1
1
7
8
48
51
100:0
45:55
O
OH
OAc
(33)
(34)
1
2
9
0
OH
(35)
OAc
(36)
48
48
15:85
100:0
OH
(37)
OAc
(38)
a
The compositions were obtained by GC-FID analysis by comparing peak areas of alcohol vs. acetate.

Molecules 2020, 25, 3217
5 of 11
Scheme 3. Chemo-selective acetylation of equimolar mixtures. Reagents and conditions: (
a
) alcohols
◦
(1:1), ethylene glycol diacetate (EGDA) (1 equiv), Lipozyme 435 (1% wt), 40 C, 150 rpm, Table 2.
Table 2. Screening of competitive enzymatic acetylation of equimolar mixtures of alcohols according to
Scheme 3.
1
1
2
2
R OH/R OAc
R OH/R OAc
1
2
1
2
Entry
Substrates R OH + R OH
Products R OAc + R OAc
Time (h)
(
GC-FID Ratio %) a
(GC-FID Ratio %) a
OH
OAc
1
+
+
8
39/40 (6:94)
33/34 (96.5:3.5)
33/34 (80:20)
OH
OAc
(
39)
(33)
OH
(40)
(34)
OAc
OH
OAc
+
+
2
3
48
41/42 (19:81)
(
41)
(33)
OH
(42)
(6)
(34)
OAc
OH
+
OAc
+
24
56
5/6 (9:91)
43/44 (81:19)
43/44 (51:49)
(5)
(43)
(44)
OH
OH
OAc
OAc
4
a
+
+
33/34 (66:34)
(33)
(43)
(34)
(44)
The compositions were obtained by GC-FID analysis by comparing peak areas of respective alcohol vs. its acetate.
Figure 1. Time-dependent evolution of the ratio between forming acetates during competitive
experiments (cf. Scheme 3, cf. Tables S3–S6 in the Supporting Information).
The observed enzymatic discrimination between interrelated alcohols could be potentially used for
the chemo-selective acetylation of their mixtures and/or for the reactive separation of product-of-choice
from a multi-component (raw) material. To verify such an approach on a model system, the

Molecules 2020, 25, 3217
6 of 11
preparative-scale enzymatic acetylation of an equimolar mixture (5 g each) of 1-pentanol 39 and
-pentanol 33 was carried out. In order to maximise the chemo-selectivity, we have lowered the amount
3
of both the enzyme and EGDA loading by half in comparison to standard conditions (cf. Scheme 2).
Thus, after gentle heating and stirring the reaction mixture for less than 5 h, which was followed by
its hydrodistillation and final extraction of distillate, 1-pentyl acetate 40 was isolated in 68% yield
(Scheme 4).
Scheme 4. Chemo-selective acetylation of equimolar mixture of alcohols. Reagents and conditions: (a)
◦
3
9
+
33 (1:1), EGDA (1 equiv), Lipozyme 435 (1% wt), 40 C, 150 rpm, 4.5 h, 40 (94% GC-FID purity,
6
8% yield from 39).
3
. Materials and Methods
3
.1. Materials and Methods
Chemicals and reagents were purchased from commercial sources (Alfa Aesar, Sigma-Aldrich,
Merck) and were used without further purification. Hexanes refer to a mixture of C-6 alkanes (b.p.
◦
6
0–80 C). Lipozyme 435 (Candida antarctica lipase B immobilized on a macroporous acrylic resin) with
a reported activity of 8000 PLU (propyl laurate units) per gram was purchased from Novozymes
A/S (Bagsværd, Denmark). All GC-FID analyses were performed on a gas chromatograph Agilent
7890A equipped with FID and a split–splitless injector as follows: (a) standard GC-FID using column
DB-WAX (30 m
gradient 60 C (0 min)
rest of alcohols (except phytol 13): injection 0.01
×
0.25 mm
×
0.15
µ
m), for rhododendrol 21: injection 1.0
µ
L, split 20:1, temperature
◦
◦
◦
→
20 C/min
→
200 C (25 min), carrier gas H , flow 1.2 mL/min, for all the
2
◦
µL, split 50:1, temperature gradient 40 C (0 min)
◦
◦
→
10 C/min
→
200 C (12 min), carrier gas H , flow 1.2 mL/min, (b) standard GC-FID for phytol 13
2
using column DB-5 (30 m
0 C (0 min)
column Lipodex-A (50 m
×
0.25 mm
×
0.25
µ
m), injection 0.01
µL, split 50:1, temperature gradient
◦
◦
◦
4
→
10 C/min
→
200 C (12 min), carrier gas H , flow 1.2 mL/min, (c) chiral GC-FID using
2
◦
×
0.25 mm
×
0.25
µm), injection 0.2
µL, split 50:1, 55 C (50 min) isothermal,
carrier gas H , flow 2.0 mL/min). All GC-MS analyses (except rhododendrol 21) were performed on a
2
gas chromatograph Agilent 7890A coupled with Agilent 5975C inert MSD with Triple-Axis Detector
◦
(
(
column DB-Wax 30 m
×
0.25 mm
×
0.15
µ
m, injection 0.1
µ
L, split 20:1, temperature gradient 40 C
◦
◦
0 min)
→
10 C/min
→
200 C (12 min), carrier gas H , flow 1.0 mL/min), for rhododendrol 21
:
2
◦
◦
◦
220 C (25 min), carrier gas H , flow 1.0 mL/min).
2
temperature gradient 60 C (0 min)
To monitor the reaction progress, the relative amount (reported as a percentage) of each component
alcohol vs. acetate) was determined according to the proportion of each peak area in the total
peak area of the two compositions. The chemical identity of prepared acetates was determined as
→
20 C/min
→
(
follows: (a) GC-FID comparison with the commercial standard for
NIST library) for 10 12 14 16 18, rac-20, (S)-20, rac-22 24 26 28
2
,
4
,
,
6
,
8
, rac-20, (S)-20, (b) GC-MS
1
(
,
,
,
,
,
,
,
,
34
36, (c) H NMR for rac-22
,
30.
NMR spectra were recorded on a Varian INOVA 300 spectrometer and were correctly shifted using
residual non-deuterated solvent (CHCl3:
on Agilent 1260 Infinity LC system equipped with a Lux Amylose-1 column (250
stationary phase: amylose tris-(3,5-dimethylphenyl) carbamate, Phenomenex, Torrance, CA, USA)
and diode-array detector (220 nm). The separation was carried out at 15 C by isocratic elution with
δ
= 7.26 ppm). Chiral HPLC analyses were performed
H
®
×
4.6 mm, 5 µm,
◦
n-hexane/isopropanol (90:10, v/v) with the flow rate of 1 mL/min.
3
.2. Synthetic Procedures and Analytical Data
General experiment for the optimisation screening of acetylation of (Z)-hex-3-en-1-ol (1):

Molecules 2020, 25, 3217
7 of 11
To a solution of an alcohol (10.0 mmol) in either neat EGDA or EGDA/cosolvent mixture was
added Lipozyme 435. The reaction mixture was stirred in an orbital shaker (150 rpm) at two different
◦
temperatures (40 or 45 C, respectively) and the reaction progress was monitored by GC-FID (see
Table S1 in the Supporting Information).
General experiment for the optimised acetylation of alcohols, according to Scheme 2:
Lipozyme 435 (2% wt) was added to a solution of an alcohol (1 equiv) in EGDA (2 equiv).
◦
The reaction mixture was incubated in an orbital shaker (40 C, 150 rpm) and the reaction progress was
monitored by GC-FID (see Table 1).
Scale-up preparation of (Z)-hex-3-en-1-yl acetate (2):
Lipozyme 435 (400 mg, 2% wt) was added to a solution of (Z)-hex-3-en-1-ol
in EGDA (58.3 g, 0.4 mol). The reaction mixture was incubated in an orbital shaker (40 C, 150 rpm)
1
(20.0 g, 0.2 mol)
◦
and the reaction progress was monitored by GC-FID. The enzyme was filtered off and the filtrate was
◦
hydro-distilled. The separated organic phase was purified by vacuum rectification (b.p. 62–63 C/20
mbar) affording (Z)-hex-3-en-1-yl acetate
GC-FID purity, containing 6.48 g of
2.80 g of
.35
Me), 0.96 (t, J = 7.5 Hz, 3H, H-6). C NMR (75 MHz, CDCl ):
2 in two fractions as colourless oils: first fraction (8.11 g, 80%
) and second fraction (13.35 g, 95.7% GC-FID purity, containing
2
1
1
5
2
) in a combined 70% yield. H NMR (300 MHz, CDCl3):
δ
= 5.53
−
5.45 (m, 1H, H-3),
−
5.26 (m, 1H, H-4), 4.05 (t, J = 6.9 Hz, 2H, H-1), 2.39
−
2.32 (m, 2H, H-2), 2.09
−
2.00 (m, 5H, H-5,
1
3
δ
= 171.3 (C = O), 134.7 (C-4), 123.8
3
(
C-3), 64.1 (C-1), 26.8 (C-2), 21.1 (Me), 20.7 (C-5), 14.3 (C-6). The obtained data fully correspond to the
reported spectra [54].
Enzymatic acetylation of rhododendrol (21) for the identification purposes:
Lipozyme 435 (10 mg, 2% wt) was added to a suspension of racemic rhododendrol 21 (0.50 g,
3
0.1 mmol) in EGDA (0.88 g, 60.2 mmol, 2 equiv). The reaction mixture was incubated in an orbital
◦
shaker (40 C, 150 rpm) and GC-FID monitored the progress of the reaction. The enzyme was filtered
off, the filtrate was diluted with Et O (10 mL), and it was washed with 2M aq. NaOH (15 mL) to
2
separate the unreacted enantiomer of the substrate [55]. The aqueous layer was washed with Et O
2
(
(
10 mL) and acidified with 9% aq. HCl to pH 6. After extraction of the aqueous phase with Et O
2
10 mL), the separated organic phase was dried over anhydrous Na SO , filtered, and evaporated in
2
4
vacuo to furnish an oil. An aliquot sample of the crude material was subjected to proton NMR [56
]
and GC-MS [57] analyses to identify the acetate 22, and the obtained spectra fully corresponded to
the reported data. In the alternative experiment, after completion of the reaction, the enzyme was
filtered off, the filtrate was diluted with water, and it was extracted with methyl tert-butyl ether
(
MTBE). The separated organic phase was dried over Na SO and concentrated in vacuo to furnish
2 4
an oil. An aliquot sample of the crude material was purified by flash liquid chromatography (FLC)
(
(
silica gel, DCM/MeOH 95:5) to obtain two fractions: the first one containing the less polar acetate 22
R = 0.60) and the second fraction containing the more polar alcohol 21 (R = 0.34). The latter one was
f
f
then analyzed on chiral HPLC and was found to contain enantiomerically enriched (S)-rhododendrol
S)-21 (e.r. = 92/8, see Figure S2 in the Supporting Information, the enantiomeric elution order was
(
adopted from Reference [58]).
Enzymatic acetylation of pent-3-yn-1-ol (29) for identification purposes:
Lipozyme 435 (20 mg, 2% wt) was added to a solution of pent-3-yn-1-ol 29 (1.0 g, 11.9 mmol) in
◦
EGDA (3.47 g, 23.8 mmol, 2 equiv). The reaction mixture was incubated in an orbital shaker (40 C,
1
50 rpm) and the reaction progress was monitored by GC-FID. The enzyme was filtered off and the
filtrate was extracted with Et O (3 15 mL). The separated organic layer was dried over anhydrous
Na SO , filtered, and evaporated in vacuo. An aliquot of the crude material was purified by FLC on
×
2
2
4
1
silica gel (hexanes/AcOEt 1:1) to obtain an analytical sample of pure acetate 30. H NMR (300 MHz,
CDCl3): = 4.12 (t, J = 6.9 Hz, 1H, H-1), 2.46 (ddt, J = 6.9, 4.4, 2.6 Hz, 1H, H-2), 2.07 (s, 1H, Me), 1.78 (t,
δ
J = 2.6 Hz, 1H, H-5).
General experiment for the competitive acetylations, according to Scheme 3.

Molecules 2020, 25, 3217
8 of 11
EGDA (1 equiv) and Lipozyme 435 (1% wt) was added to an equimolar mixture of two alcohols
◦
(
0.5 + 0.5 equiv). The reaction mixture was stirred in an orbital shaker (40 C, 150 rpm) and the reaction
progress was monitored by GC-FID. (see Table 2).
Chemo-selective acetylation of pentan-1-ol (39):
EGDA (16.6 g, 113.4 mmol) and Lipozyme 435 (100 mg, 1% wt) was added to an equimolar
mixture of pentan-1-ol 39 (5.0 g, 56.7 mmol) and pentan-3-ol 33 (5.0 g, 56.7 mmol). The reaction mixture
◦
was stirred in an orbital shaker (40 C, 150 rpm) and the reaction progress was monitored by GC-FID.
The enzyme was filtered off and the filtrate was hydro-distilled. The separated organic phase was
diluted with hexane (75 mL), and washed with aq. NaHCO soln. (3
×
75 mL, pH 9) and distilled water
3
◦
(
150 mL). The separated organic phase was dried over Na SO and concentrated in vacuo (40 C, 500
2
4
→
200 mbar) to yield pent-1-yl acetate 40 as colourless oil (5.3 g, 94% GC-FID purity, 68% yield from
1
3
9
). H NMR (300 MHz, CDCl ):
δ
= 4.03 (t, J = 6.8 Hz, 2H, H-1), 2.02 (s, 3H, Me), 1.64
−
1.56 (m, 2H,
3
H-2), 1.31 (m, 4H, H-3, H-4), 0.90
−
0.87 (m, 3H, H-5). 13C NMR (75 MHz, CDCl3):
δ
= 171.3 (C = O),
64.7 (C-1), 28.4 (C-2), 28.2 (C-3), 22.4 (C-4), 21.1 (Me), 14.0 (C-5). The obtained data fully correspond to
the reported spectra [59].
4
. Conclusions
In combination with Lipozyme 435, we have successfully employed EGDA as a useful acetylation
reagent and solvent for chemo-selective and/or stereoselective enzymatic preparation of natural acetates
for perfumery and/or flavour purposes. We found that such a green system exhibits high reactivity and
selectivity toward activated (homo) allylic and non-activated primary/secondary alcohols. This feature
was utilised in the scalable multigram synthesis of fragrant (Z)-hex-3-en-1-yl acetate 2 in 70% yield after
vacuum rectification. In addition, the Lipozyme 435/EGDA system was also found to be applicable to
the chemo-selective acetylation of (hydroxyalkyl) phenols as well as to the kinetic resolution of chiral
secondary alcohols. Lastly, its remarkable discrimination power enabled the preferential acetylation
of respective alcohol from the equimolar mixture of two isomeric substrates. Thus, pure 1-pentyl
acetate 40 was isolated as a single product in 68% yield in a multigram quantity from the 1:1 mixture of
1
-pentanol 39 and 3-pentanol 33.
Supplementary Materials: The following are available online: optimisation screening conditions, GC-FID
retention times of all compounds, copies of chiral GC-FID and HPLC spectra, copies of NMR spectra of
isolated compounds.
Author Contributions: Conceptualization and methodology, E.P. and P.S. Synthesis and analytics, E.P.
Writing—original draft preparation, review and editing, E.P. and P.S. Supervision and project administration, P.S.
Funding acquisition, P.S. All authors have read and agreed to the published version of the manuscript.
Funding: The Grant Agency of the Slovak Republic (VEGA 1/0162/20) funded this research.
Acknowledgments: We thank Veronika Výbohová and Katarína Hrobo nˇ ová for chiral HPLC analyses. We very
much appreciate the technical support from Axxence Slovakia Ltd.
Conflicts of Interest: The authors declare no conflict of interest.
References
1.
2.
3.
Surburg, H.; Panten, J. Common Fragrance and Flavor Materials, 5th ed.; Wiley-VCH: Weinheim, Germany,
006.
2
Anbu, N.; Nagarjun, N.; Jacob, M.; Kalaiarasi, J.M.V.K.; Dhakshinamoorthy, A. Acetylation of Alcohols,
Amines, Phenols, Thiols under Catalyst and Solvent-Free Conditions. Chemistry 2019, 1, 69–79. [CrossRef]
Stergiou, P.-Y.; Foukis, A.; Filippou, M.; Koukouritaki, M.; Parapouli, M.; Theodorou, L.G.; Hatziloukas, E.;
Afendra, A.; Pandey, A.; Papamichael, E.M.; et al. Advances in lipase-catalyzed esterification reactions.
Biotechnol. Adv. 2013, 31, 1846–1859. [CrossRef] [PubMed]
4
.
S
á
, A.G.A.; de Meneses, A.C.; de Ara
esters used as flavor ingredients for food, cosmetics and pharmaceuticals industries. Trends Food Sci. Technol.
017, 69, 95–105. [CrossRef]
újo, P.H.H.; de Oliveira, D. A review on enzymatic synthesis of aromatic
2

Molecules 2020, 25, 3217
9 of 11
5
6
7
8
9
1
.
.
.
.
.
Dhake, K.P.; Thakare, D.D.; Bhanage, B.M. Lipase: A potential biocatalyst for the synthesis of valuable
flavour and fragrance ester compounds. Flavour Fragr. J. 2013, 28, 71–83. [CrossRef]
Gu, Y.; J
CrossRef]
az- lvarez, A.E.; Francos, J.; Lastra-Barreira, B.; Crocheta, P.; Cadierno, V. Glycerol and derived solvents:
new sustainable reaction media for organic synthesis. Chem. Commun. 2011, 47, 6208–6227. [CrossRef]
Garc a, J.I.; Garc a-Mar na, H.; Piresa, E. Glycerol based solvents: synthesis, properties and applications.
érôme, F. Glycerol as a sustainable solvent for green chemistry. Green Chem. 2010, 12, 1127–1138.
[
D
í
Á
í
í
í
Green Chem. 2014, 16, 1007–1033. [CrossRef]
Wolfson, A.; Atyya, A.; Dlugy, C.; Tavor, D. Glycerol triacetate as solvent and acyl donor in the production of
isoamyl acetate with Candida antarctica lipase B. Bioprocess Biosyst. Eng. 2010, 33, 363–366. [CrossRef]
´
0. Milivojevi c´ , A.; Corovi c´ , M.; Carevi c´ , M.; Banjanac, K.; Vujisi c´ , L.; Veli cˇ kovi c´ , D.; Bezbradica, D. Highly
efficient enzymatic acetylation of flavonoids: Development of solvent-free process and kinetic evaluation.
Biochem. Eng. J. 2017, 128, 106–115. [CrossRef]
1
1. Varga, Z.; Kmecz, I.; Sz
-phenylethanol and a straightforward modelling of the reaction. Biocatal. Biotransformation 2017, 35,
27–433. [CrossRef]
écsényi, A.; Székely, E. Neat lipase-catalysed kinetic resolution of racemic
1
4
1
2. Bayout, I.; Bouzemi, N.; Guo, N.; Mao, X.; Serra, S.; Riva, S.; Secundo, F. Natural flavor ester synthesis
catalyzed by lipases. Flavour Fragr. J. 2020, 35, 209–218. [CrossRef]
1
3. Bourg-Garros, S.; Razafindramboa, N.; Pavia, A.A. Optimization of lipase-catalyzed synthesis of
(z)-3-hexen-1-yl acetate by direct esterification in hexane and a solvent-free medium. Enzym. Microb.
Technol. 1998, 22, 240–245. [CrossRef]
1
1
1
1
1
1
2
2
2
4. Bourg-Garros, S.; Razafindramboa, N.; Pavia, A.A. Large-scale preparation of (Z)-3-hexen-1-yl acetate using
Candida antarctica-immobilized lipase in hexane. Biotechnol. Biochem. 1998, 59, 495–500.
5. Chiang, W.D.; Chang, S.W.; Shieh, C.J. Studies on the optimized lipase-catalyzed biosynthesis of
cis-3-hexen-1-yl acetate in n-hexane. Process Biochem. 2003, 38, 1193–1199. [CrossRef]
6. Langrand, G.; Triantaphylides, C.; Baratti, J. Lipase catalyzed formation of flavour esters. Biotechnol. Lett.
1
988, 10, 549–554. [CrossRef]
7. Chulalaknanukul, W.; Condoret, J.; Combes, D. Kinetics of geranyl acetate synthesis by lipase-catalysed
transesterification in n-hexane. Enzyme Microb. Technol. 1992, 14, 293–298. [CrossRef]
8. Akoh, C.C.; Claon, P.A. Enzymatic synthesis of geranyl acetate in n-hexane with Candida antarctica lipases.
J. Am. Chem. Soc. 1994, 71, 575–578.
9. Claon, P.A.; Akoh, C.C. Lipase-catalyzed synthesis of primary terpenyl acetates by transesterification: study
of reaction parameters. J. Agric. Food Chem. 1994, 42, 2349–2352. [CrossRef]
0. Yee, L.N.; Akoh, C.C. Enzymatic synthesis of geranyl acetate by transesterification with acetic anhydride as
acyl donor. J. Am. Oil Chem. Soc. 1996, 73, 1379–1384. [CrossRef]
1. Huang, S.; Chang, H. Kinetic study on the esterification of geraniol and acetic acid in organic solvents using
surfactant-coated lipase. J. Chem. Technol. Biotechnol. 1999, 74, 183–187. [CrossRef]
2. Bartling, K.; Thompson, J.U.; Pfromm, P.H.; Czermak, P.; Rezac, M.E. Lipase-catalyzed synthesis of geranyl
acetate in n-hexane with membrane-mediated water removal. Biotechnol. Bioeng. 2001, 75, 676–681. [CrossRef]
[PubMed]
2
2
2
2
2
3. Peres, C.; Harper, N.; da Silva, M.D.R.G.; Barreiros, S. Effect of zeolites on lipase catalyzed esterification in
nonaqueous media. Enzyme Microb. Technol. 2005, 37, 145–149. [CrossRef]
4. Barahona, D.; Pfromm, P.H.; Rezac, M.E. Effect of water activity on the lipase catalyzed esterification of
geraniol in ionic liquid [bmim]PF . Biotechnol. Bioeng. 2006, 93, 318–324. [CrossRef] [PubMed]
6
5. De la Casa, R.M.; Sinisterra, J.V.; Sánchez-Montero, J.M. Characterization and catalytic properties of a new
crude lipase from C. Rugosa. Enzyme Microb. Technol. 2006, 38, 599–609. [CrossRef]
6. Antoniotti, S.; Fernandez, X.; Duñach, E. Reaction design for evaluation of the substrate range of hydrolases.
Biocatal. Biotransformation 2008, 26, 228–234. [CrossRef]
7. Couto, R.; Vidinha, P.; Peres, C.; Ribeiro, A.S.; Ferreira, O.; Oliveira, M.V.; Macedo, E.A.; Loureiro, J.M.;
Barreiros, S. Geranyl Acetate Synthesis in a Packed-Bed Reactor Catalyzed by Novozym in Supercritical
Carbon Dioxide and in Supercritical Ethane. Ind. Eng. Chem. Res. 2011, 50, 1938–1946. [CrossRef]
8. Lozano, P.; Bernal, J.M.; Navarro, A. A clean enzymatic process for producing flavour esters by direct
esterification in switchable ionic liquid/solid phases. Green Chem. 2012, 14, 3026–3033. [CrossRef]
2

Molecules 2020, 25, 3217
10 of 11
2
9. Gupta, A.; Dhakate, S.R.; Pahwa, M.; Sinha, S.; Chand, S.; Mathur, R.B. Geranyl acetate synthesis catalyzed by
Thermomyces lanuginosus lipase immobilized on electrospun polyacrylonitrile nanofiber membrane. Process
Biochem. 2013, 48, 124–132. [CrossRef]
3
0. Badgujar, K.C.; Bhanage, B.M. Synthesis of geranyl acetate in non-aqueous media using immobilized
Pseudomonas cepacia lipase on biodegradable polymer film: Kinetic modelling and chain length effect study.
Process Biochem. 2014, 49, 1304–1313. [CrossRef]
3
1. Lozano, P.; Bernal, J.M.; G
Green bioprocesses in sponge-like ionic liquids. Catal. Today 2015, 255, 54–59. [CrossRef]
2. Adarme, C.A.A.; Le o, R.A.C.; de Souza, S.P.; Itabaiana, I., Jr.; de Souza, R.O.M.A.; Rezende, C.M.
ómez, C.; García-Verdugo, E.; Burguete, M.I.; Sánchez, G.; Vaultier, M.; Luis, S.V.
3
ã
Continuous-Flow Chemo and Enzymatic Synthesis of Monoterpenic Esters with Integrated Purification. Mol.
Catal. 2018, 453, 39–46. [CrossRef]
3
3. Murcia, M.D.; G
and kinetic parameters calculation in the lipase-catalysed synthesis of geranyl acetate. Chem. Eng. Res. Des.
018, 138, 135–143. [CrossRef]
ómez, M.; Gómez, E.; Gómez, J.L.; Hidalgo, A.M.; Sánchez, A.; Vergara, P. Kinetic modelling
2
3
3
3
3
3
3
4
4. Perdomo, I.C.; Gianolio, S.; Pinto, A.; Romano, D.; Contente, M.L.; Paradisi, F.; Molinari, F. Efficient Enzymatic
Preparation of Flavor Esters in Water. J. Agric. Food Chem. 2019, 67, 6517–6522. [CrossRef] [PubMed]
5. Yadav, G.D.; Devendran, S. Lipase catalyzed synthesis of cinnamyl acetate via transesterification in
non-aqueous medium. Process Biochem. 2012, 47, 496–502. [CrossRef]
6. Wolfson, A.; Dlugy, C.; Karanet, A.; Tavor, D. A sustainable one-pot synthesis of cinnamyl acetate in triacetin.
Tetrahedron Lett. 2012, 53, 4565–4567. [CrossRef]
7. Geng, B.; Wang, M.; Qi, W.; Su, R.; He, Y. Cinnamyl acetate synthesis by lipase-catalyzed transesterification
in a solvent-free system. Biotech. Appl. Biochem. 2012, 59, 270–275. [CrossRef] [PubMed]
8. Wu, Z.; Qi, W.; Wang, M.; Su, R.; He, Z. Lipase immobilized on novel ceramic supporter with Ni activation
for efficient cinnamyl acetate synthesis. J. Mol. Catal. B Enzym. 2014, 110, 32–38. [CrossRef]
9. Tomke, D.P.; Rathod, V.K. Ultrasound assisted lipase catalyzed synthesis of cinnamyl acetate via
transesterification reaction in a solvent free medium. Ultrason. Sonochemistry 2015, 27, 241–246. [CrossRef]
0. Badgujar, K.C.; Sasaki, T.; Bhanage, B.M. Synthesis of lipase nano-bio-conjugates as an efficient biocatalyst:
characterization and activity–stability studies with potential biocatalytic applications. RSC Adv. 2015, 5,
5
5238–55251. [CrossRef]
4
4
4
1. Perdomo, I.C.; Contente, M.L.; Pinto, A.; Romano, D.; Fernandes, P.; Molinari, F. Continuous preparation of
flavour-active acetate esters by direct biocatalytic esterification. Flavour Fragr. J. 2020, 35, 190–196. [CrossRef]
2. Nishio, T.; Takahashi, K.; Yoshimoto, T.; Kodera, Y.; Saito, Y.; Inada, Y. Terpene alcohol ester synthesis by
polyethylene glycol-modified lipase in benzene. Biotechnol. Lett. 1987, 9, 187–190. [CrossRef]
3. Yanagishita, H.; Sakaki, K.; Hirata, H. Optical Resolution of 2-Alkanol by Lipase-Catalyzed Acetylation
with Vinyl Acetate in Packed-Bed Reactor with Recycling System. J. Oleo Sci. 2007, 56, 137–148. [CrossRef]
[
PubMed]
4. Dlugy, C.; Wolfson, A. Lipase catalyse glycerolysis for kinetic resolution of racemates. Bioprocess Biosyst. Eng.
007, 30, 327–330. [CrossRef]
4
4
2
5. Zarcula, C.; Corîci, L.; Croitoru, R.; Ursoiu, A.; Peter, F. Preparation and properties of xerogels obtained by
ionic liquid incorporation during the immobilization of lipase by the sol–gel method. J. Mol. Catal. B. Enzym.
2
010, 65, 79–86. [CrossRef]
4
4
6. Strohalm, H.; Dold, S.; Pendzialek, K.; Weiher, M.; Engel, K.H. Preparation of Passion Fruit-Typical 2-Alkyl
Ester Enantiomers via Lipase-Catalyzed Kinetic Resolution. J. Agric. Food Chem. 2010, 58, 6328–6333.
[
CrossRef] [PubMed]
7. Tomin, A.; Weiser, D.; Hellner, G.; Bata, Z.; Corici, L.; P
generation sol–gel lipase immobilization with ternary alkoxysilane precursor systems. Process Biochem. 2011
6, 52–58. [CrossRef]
é
ter, F.; Koczka, B.; Poppe, L. Fine-tuning the second
,
4
4
8. Hellner, G.; Boros, Z.; Tomin, A.; Poppe, L. Novel Sol-Gel Lipases by Designed Bioimprinting for
Continuous-Flow Kinetic Resolutions. Adv. Synth. Catal. 2011, 353, 2481–2491. [CrossRef]
4
9. Schober, M.; Gadler, P.; Knaus, T.; Kayer, H.; Birner-Grünberger, R.; Gülly, C.; Macheroux, P.; Wagner, U.;
Faber, K. A Stereoselective Inverting sec-Alkylsulfatase for the Deracemization of sec-Alcohols. Org. Lett.
2
011, 13, 4296–4299. [CrossRef]

Molecules 2020, 25, 3217
11 of 11
5
0. Kazlauskas, R.J.; Weissfloch, A.N.E.; Rappaport, A.T.; Cuccia, L.A. A rule to predict which enantiomer of
a secondary alcohol reacts faster in reactions catalyzed by cholesterol esterase, lipase from Pseudomonas
cepacia, and lipase from Candida rugosa. J. Org. Chem. 1991, 56, 2656–2665. [CrossRef]
5
5
1. Yuasa, Y.; Shibuya, S.; Yuasa, Y. Resolution of Racemic Rhododendrol by Lipase-Catalyzed Enantioselective
Acetylation. Synth. Commun. 2003, 33, 1469–1475. [CrossRef]
2. Kitayama, T.; Isomori, S.; Nakamura, K. Asymmetric synthesis of enantiomerically pure zingerols by
lipase-catalyzed transesterification and efficient synthesis of their analogues. Tetrahedrom Asymmetry 2013
4, 621–627. [CrossRef]
,
2
5
5
5
5
3. Karume, I.; Takahashi, M.; Hamdan, S.M.; Musa, M.M. Deracemization of Secondary Alcohols by using a
Single Alcohol Dehydrogenase. ChemCatChem 2016, 8, 1459–1463. [CrossRef]
4. Velusamy, S.; Borpuzari, S.; Punniyamurthy, T. Cobalt (II)-catalyzed direct acetylation of alcohols with acetic
acid. Tetrahedron 2005, 61, 2011–2015. [CrossRef]
5. Miyazawa, T.; Yamamoto, M.; Danjo, H. Chemoselective acylation of (hydroxyalkyl) phenols catalyzed by
Candida antarctica lipase B. Biotechnol. Lett. 2013, 35, 625–630. [CrossRef]
6. Song, L.; Liu, Y.; Tong, R. Cephalosporolide B Serving as a Versatile Synthetic Precursor: Asymmetric
Biomimetic Total Syntheses of Cephalosporolides C, E, F, G, and (4-OMe-)G. Org. Lett. 2013, 15, 5850–5853.
[CrossRef]
5
7. Kulangiappar, K.; Anbukulandainathan, M.; Raju, T. Nuclear Versus Side-Chain Bromination of 4-Methoxy
Toluene by an Electrochemical Method. Synth. Commun. 2014, 44, 2494–2502. [CrossRef]
5
8. Boffi, A.; Cacchi, S.; Ceci, P.; Cirilli, R.; Fabrizi, G.; Prastaro, A.; Niembro, S.; Shafir, A.; Vallribera, A. The Heck
Reaction of Allylic Alcohols Catalyzed by Palladium Nanoparticles in Water: Chemoenzymatic Synthesis of
(R)-(–)-Rhododendrol. ChemCatChem 2011, 3, 347–353. [CrossRef]
5
9. Kumar, H.M.; Joyasawal, S.; Reddy, B.V.S.; Chakravarthy, P.P.; Krishna, A.D.; Yadav, J.S. Reaction of orthoesters
with alcohols in the presence of acidic catalysts: a study. Indian J. Chem. 2005, 44, 1686–1692. [CrossRef]
Sample Availability: Samples of the compounds are not available from the authors.
©
2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access
article distributed under the terms and conditions of the Creative Commons Attribution
(CC BY) license (http://creativecommons.org/licenses/by/4.0/).