Solvent role in the lipase-catalysed esterification of cinnamic acid and derivatives. Optimisation of the biotransformation conditions

Article abstract of DOI:10.1016/j.tet.2020.131873

The esterification of cinnamic acid has been deeply investigated using ethanol as nucleophile and Candida antarctica lipase type B (CAL-B) as suitable biocatalyst. Special attention has been paid to the role that the solvent plays in the production of ethyl cinnamate. Therefore, volatile organic solvents and deep eutectic mixtures were employed in order to find optimal reaction conditions. Once that hexane was selected as the solvent of choice, other parameters that affect the enzyme activity were investigated in order to produce ethyl cinnamate with excellent yield. The CAL-B loading, nucleophile equivalents, temperature and reaction time have been identified as key parameters in the enzyme efficiency, and the potential of lipase-catalysed esterification has been finally exploited to produce a series of ethyl esters with different pattern substitutions on the aromatic ring.

Full text of DOI:10.1016/j.tet.2020.131873

Tetrahedron 81 (2021) 131873
Contents lists available at ScienceDirect
Tetrahedron
journal homepage: www.elsevier.com/locate/tet
Solvent role in the lipase-catalysed esterification of cinnamic acid and
derivatives. Optimisation of the biotransformation conditions
*
ꢀ
ꢀ
Laura Suarez-Escobedo, Vicente Gotor-Fernandez
ꢀ
Organic and Inorganic Chemistry Department, University of Oviedo. Avenida Julian Clavería 8, 33006, Oviedo, Spain
a r t i c l e i n f o
a b s t r a c t
Article history:
The esterification of cinnamic acid has been deeply investigated using ethanol as nucleophile and
Candida antarctica lipase type B (CAL-B) as suitable biocatalyst. Special attention has been paid to the role
that the solvent plays in the production of ethyl cinnamate. Therefore, volatile organic solvents and deep
eutectic mixtures were employed in order to find optimal reaction conditions. Once that hexane was
selected as the solvent of choice, other parameters that affect the enzyme activity were investigated in
order to produce ethyl cinnamate with excellent yield. The CAL-B loading, nucleophile equivalents,
temperature and reaction time have been identified as key parameters in the enzyme efficiency, and the
potential of lipase-catalysed esterification has been finally exploited to produce a series of ethyl esters
with different pattern substitutions on the aromatic ring.
Received 6 November 2020
Received in revised form
30 November 2020
Accepted 9 December 2020
Available online 16 January 2021
Keywords:
Candida antarctica lipase type B
Cinnamic acid
Deep eutectic solvents
Esterification
© 2020 Elsevier Ltd. All rights reserved.
Phenolic acids
1. Introduction
for the production of valuable organic compounds based on the
exquisite selectivity displayed by enzymes [9e13]. Among the
Cinnamic acid (Fig. 1, 1a) is a key intermediate in the biosyn-
thesis of a wide number of biologically active compounds including
coumarins, flavonoids, isoflavonoids, phenylpropopanoids and
stilbenes among many others, finding the trans-isomer as its most
common form in nature [1,2]. The importance of cinnamic acid and
derivatives is demonstrated by the presence of this structural motif
in numerous antibacterial, antifungal, anti-inflammatory, antioxi-
dant and antitumor agents [3e5], making them attractive targets
for the pharmaceutical industry. Interestingly, the importance of
other cinnamic acid derivatives [6,7], including for instance the
odorous cinnamaldehyde and ethyl cinnamate (2a), has not gone
unnoticed for the production of agrochemicals, polymers cosmetics
and fragrances, expanding the potential of cinnamic acid de-
rivatives to other industrial sectors. In addition, hydroxycinnamic
acids, such as ferulic (1b) and caffeic acid (1c), are currently gaining
great importance because of their presence in numerous food in-
gredients, their esterified forms displaying remarkable bioactivity
values since their solubilisation properties is favoured when
considering different formulations [8].
enzyme classes, hydrolases are when possible the selection of
choice, since they can work in a wide range of experimental con-
ditions (i.e.: temperature, solvent, substrate concentrations …) [14].
This is especially feasible when immobilised forms are employed,
providing higher biocatalyst stabilities and the possibility to reuse
the enzyme [15e18]. In this context, probably Candida antarctica
lipase type B (CAL-B) is the model enzyme to start with a screening
due to its well-known application in aminolysis, ammonolysis,
esterification, perhydrolysis, thiolysis or transesterification re-
actions [19e22].
Focusing on the esterification of cinnamic and phenolic acids, a
few examples have been described in the literature (Scheme 1),
resulting CAL-B, Bacillus lincheniformis lipase or Thermomyces
lanuginosus lipase as efficient enzymes when combined with alkyl
alcohols (methanol, ethanol, n-propanol, isopropanol, butanol,
pentanol, 2-ethylhexanol, octanol, cyclohexanol, benzyl alcohol,
lauryl alcohol, cis-9-octadecen-1-ol …) [23e36]. Remarkably, along
the last three decades the solvent role has been deeply study,
finding hydrophobic ones such as hexane, cyclohexane, cyclo-
octane, 1-octane or isooctane as the best solutions [25,26,29,33,35],
while the design of solvent-free reactions [24,34] and the selection
of DMSO [30], ionic liquids [23,27,28,31] or alcohols [32,36] appear
as suitable alternatives, especially due to their ability to solubilise
the starting carboxylic acid in order to achieve higher conversions.
Biocatalysis is currently considered a sustainable methodology
* Corresponding author.
ꢀ
E-mail address: vicgotfer@uniovi.es (V. Gotor-Fernandez).
https://doi.org/10.1016/j.tet.2020.131873
0040-4020/© 2020 Elsevier Ltd. All rights reserved.

ꢀ
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L. Suarez-Escobedo and V. Gotor-Fernandez
Tetrahedron 81 (2021) 131873
including ethyl ferulate and ethyl caffeate (2b and 2c, entries 2 and
3) and compounds monosubstituted at the C-4 (entries 4e6), C-3
(entries 7e10) or C-2 position (entries 11e14) of the aromatic ring.
2.1. Screening of solvents for lipase-catalysed experiments
Fig. 1. Chemical structure of cinnamic acid, ferulic acid, caffeic acid and ethyl cinna-
mate (1a, 1b, 1c and 2a, respectively).
An initial test was performed searching for the most adequate
organic solvent, selecting the esterification of cinnamic acid as the
benchmark reactions. Under enzymatic standard conditions for this
type of lipase-catalysed reactions: the required solvent for a
100 mM concentration of 1a in spite that most of the solvent could
not solubilized the substrate, an excess of EtOH (3 eq) as nucleo-
phile, a substrate:CAL-B 1:0.75 (w/w) ratio, 24 h and 30 ꢀC, the
reaction was monitorised over the time by TLC analyses. The set of
organic solvent tested has been disclosed in Fig. 2 and organised
depending on their logP values. The choice includes mostly hy-
drocarbons (heptane, hexane and pentane) and alkyl ethers such as
cyclopentyl methyl ether (CPME), tert-butyl methyl ether (TBME),
2-methyl tetrahydruran (2-MeTHF) and tetrahydrofuran (THF), but
also other media like ethyl acetate (EtOAc), acetonitrile (MeCN),
Cyrene™ (dihydrolevoglucosenone) or the own EtOH as both
nucleophile and solvent. It must be highlighted that this broad
selection was motivated in some cases for the possibility to develop
more sustainable processes by replacing traditional petroleum
solvents by bio-based ones such as CPME [37,38], 2-MeTHF [39] and
Cyrene™ [40].
As expected, hexane and heptane led to the higher conversions
in these experimental conditions (17e18%), while only TBME (13%)
seems to be a practical alternative although limited by its low
boiling point for further optimisation. None of the bio-based sol-
vents led to good results, so the focus was moved from there to the
use of deep eutectic mixtures as solvents [41e45]. Interestingly, this
class of neoteric solvents have gained recent attention as emerging
environmentally friendly media when exploring the synthetic po-
tential of enzymes [46], mainly hydrolases [47e49] and redox en-
zymes [50,51]. Initially, combinations of choline chloride (ChCl) as
hydrogen bond acceptor (HBA) with either urea (U) or glycerol (Gly)
as hydrogen bond donors (HBDs) were considered in 1:2 mol/mol
ratio. Unfortunately, none of them gave appreciable conversion
values in the esterification of 1a under the reaction conditions
attempted with the volatile organic solvents.
Scheme 1. Hydrolase-catalysed esterification of cinnamic acid and other carboxylic
acid derivatives.
Disappointingly, all these enzymatic processes required high
temperatures (45e80 ꢀC) and sometimes also reduce pressures, so
herein, we have tried to provide a deeper understanding of the
esterification of cinnamic acid and derivatives using CAL-B. For that
reason, a comprehensive study has been performed, paying special
attention to the possibility of using volatile organic solvents
including bio-based solvents, but also novel deep eutectic mixtures,
which could improve the sustainability of the process and lead to a
higher solubilisation and reactivity of these
boxylic acids, respectively.
a,b-unsaturated car-
2. Results and discussion
Prior to the development of the enzymatic study, the synthesis
of ethyl cinnamate (2a) was carried out by refluxing cinnamic acid
in ethanol (EtOH) in the presence of concentrated sulphuric acid.
After monitorisation of the chemical process by TLC analyses,
observing the disappearance of the carboxylic acid, the ester 2a was
recovered in excellent purity and 86% yield after liquid-liquid
extraction (Table 1, entry 1). Similarly, other cinnamic acids
bearing different substitutions in the aromatic ring were converted
into the corresponding ethyl esters (2e24 h, 75e98% yield)
In spite of these discouraging initial results, but considering the
importance of the water presence in enzymatic processes involving
deep eutectic solvents (DES) [52,53], the ChCl:urea mixture was
selected for further investigations since in addition it does not
contain polyalcohols in its structure, avoiding thus the occurrence
of competitive reactions. The DES was prepared again but drying
firstly both hydrophilic components prior to obtain the eutectic
mixture, and then a few ratios of DES:water were attempted in the
esterification of cinnamic acid under previously set-up conditions
(Table 2). Interestingly a maximum lipase activity was found when
using a 10% of water, even higher than the one previously found
with hexane, although long reaction times (72 h) and higher tem-
peratures (60 ꢀC) did not improve the conversion value.
Table 1
Chemical esterification of 1a-n using ethanol and sulphuric acid.
Entry
Acid
R1
R2
R3
Time (h)
Yield 2a-n (%)a
1
2
3
1a
1b
1c
H
OH
OH
H
OCH3
OH
H
H
H
2
24
24
86
94
96
4
5
6
1d
1e
1f
NO2
CF3
CH3
H
H
H
H
H
H
24
22
24
94
98
96
2.2. Optimisation of the reaction conditions
7
8
9
10
1g
1h
1i
H
H
H
H
CH3
OCH3
F
H
H
H
H
24
22
2
87
98
75
93
Once selected hexane as the best solvent for the production of
ethyl cinnamate (2a), other parameters were considered to achieve
higher conversions including the use of different enzyme loadings
(1:0.75 or 1:1, w/w), nucleophile excess (3 or 6 eq of EtOH), tem-
perature (30, 45 or 55 ꢀC) and reaction time (24 or 72 h). A sum-
mary of this research is depicted in Table 3. Starting from the
experimental conditions used in the solvent screening, the reaction
was prolonged from 1 to 3 days, increasing the conversion from 18
1j
NO2
22
11
12
13
14
1k
1l
1m
1n
H
H
H
H
H
H
H
H
NO2
CF3
Br
24
22
24
24
95
79
98
95
OCH3
a
Yields of ethyl esters 2a-n were calculated after isolation by liquid-liquid
extraction.
2

ꢀ
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L. Suarez-Escobedo and V. Gotor-Fernandez
Tetrahedron 81 (2021) 131873
Fig. 2. Influence of the solvent in the esterification of cinnamic acid (1a) catalysed by CAL-B using 3 equivalents of ethanol for 24 h at 45 ꢀC and 250 rpm.
Table 2
The enzyme reusability was finally explored, finding that a
significant loss of the activity was found after the third use of CAL-B
to produce ethyl cinnamate in 66% conversion. This gradual enzyme
deactivation can be explained based on the high temperature
(45 ꢀC) and long reaction times (72 h) required for reaching high
conversion values.
Lipase-catalysed esterification of cinnamic acid (1a) using ethanol (3 eq) as nucle-
ophile and ChCl:urea (1:2 mol/mol) as reaction medium in the presence of water
after 24 h at 45 ꢀC and 250 rpm.
Entry
DES:H2O
Conversion 2a (%)a
1
2
3
4
5
100:0
90:10
75:25
50:50
25:75
<3
27
10
<3
<3
2.3. Reaction scope
Under optimised reaction conditions for 1a, the esterification of
a series of cinnamic acids 1b-n was explored, those bearing a va-
riety of pattern substitutions on the aromatic ring as already dis-
cussed in Table 1. Therefore, employing a 100 mM substrate
concentration in hexane, 3 equivalents of ethanol and using CAL-B
(1:1 w/w substrate/enzyme ratio), the reactions were left for 72 h at
45 ꢀC and 250 rpm (Table 4). On the one hand, in some cases, and
because of the high insolubility of the substrates, even at 45 ꢀC, no
conversion was found for ferulic acid (1b: R1 ¼ OH, R2 ¼ OCH3 and
R3 ¼ H), caffeic acid (1c: R1 ¼ R2 ¼ OH and R3 ¼ H) and 2-
nitrocinnamic acid (1d: R1 ¼ NO2 and R2 ¼ R3 ¼ H), or almost a
negligible one was observed for 4-nitrocinnamic acid (1e, entry 4).
On the other hand, substrates with electron withdrawing (entries 5
and 9e13, or electron donating (entries 6e8 and 14) groups at
different positions of the aromatic ring were tested, finding com-
plete conversions for 1i and 1m, which possess a halogen atom
directed linked to the aromatic ring. In general, moderate to good
isolated yields were obtained after column chromatography puri-
fication, attaining the lower conversions when considering selected
electron donating groups in the phenyl ring (1f and 1n).
a
Conversion values were calculated through 1H NMR analyses of the reaction
crudes after enzyme filtration and liquid-liquid extraction.
Table 3
Esterification of 1a with EtOH using CAL-B and hexane as biocatalyst and solvent,
respectively.
Entry
T (ꢀC)
t (h)
EtOH (eq)
1a: CAL-B (w/w)
Conversion 2a (%)a
1
2
30
30
24
72
3
3
1:0.75
1:0.75
18
40
3
4
45
45
72
72
3
6
1:0.75
1:0.75
82
58
5
6
45
55
72
72
3
3
1:1
1:1
91
92
a
Percentage of product 2a was calculated through 1H NMR analyses of the re-
3. Conclusions
action crudes after enzyme filtration and solvent evaporation.
The esterification of cinnamic acid and other related carboxylic
acids have been explored using ethanol and two independent
strategies, the traditional Fischer esterification using sulphuric acid
as catalyst and the alternative biocatalytic approach with Candida
antarctica lipase type B as biocatalyst. Good to excellent yields were
attained using sulphuric acid and refluxing ethanol, while for the
enzymatic transformation more sustainable conditions were
attempted. Special attention has been paid to the solvent nature in
to 40% (entries 1 and 2). Therefore, the reaction temperature was
increased to 45 ꢀC, finding a 82% conversion after 72 h (entry 3),
while an increase in the amount of EtOH led to a considerable loss
of activity (58% conversion, entry 4). The reactivity was further
improved by increasing the loading of enzyme to a 1:1 CAL-B: 1a
ratio (91% conversion, entry 5), while the use of a higher temper-
ature (55 ꢀC, entry 6) provided similar results.
3

ꢀ
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L. Suarez-Escobedo and V. Gotor-Fernandez
Tetrahedron 81 (2021) 131873
Table 4
silica gel (230e400 mesh). NMR spectra were recorded on a
300 MHz spectrometer for the measurement of conversion values
and characterisation of carboxylic acids and esters. All chemical
CAL-B catalysed esterification of carboxylic acids 1a-n using 3 equivalents of ethanol
and hexane as solvent for 72 h at 45 ꢀC and 250 rpm.
shifts (d) are given in parts per million (ppm) and referenced to the
residual solvent signal as internal standard. IR spectra were recor-
ded on a FT/IR-4700 type A spectrophotometer, and the frequency
(n
) of the most representative absorption bands are listed.
The performance of 1H NMR for carboxylic acids 1a-n and ethyl
esters 2a-n has allowed us the calculation of conversion values for
enzymatic experiments, so experiments in the adequate deuterated
solvent (CDCl3, MeOH-d4 and DMSO‑d6) were performed, assuring
the complete solubility of both class of organic compounds to
achieve reliable results.
Entry
Acid
R1
R2
R3
Conversion 2a-n (%)a
1
2
3
1a
1b
1c
H
OH
OH
H
OCH3
OH
H
H
H
91 (79)
<3
<3
4
5
6
1d
1e
1f
NO2
CF3
CH3
H
H
H
H
H
H
<3
73 (36)
14
4.1. General procedure for the chemical esterification of phenolic
acids (1a-n)
7
8
9
10
1g
1h
1i
H
H
H
H
CH3
OCH3
F
H
H
H
H
97 (90)
78 (65)
100 (73)
76 (69)
A suspension of the corresponding phenolic acid 1a-n (500 mg)
in ethanol (EtOH, 3.8 mL, 65.1 mmol) was dissolved by the addition
of a 98% aqueous solution of sulphuric acid (0.5 mL). The solution
was magnetically stirred at 82 ꢀC and monitorised by TLC analysis
(20e60% EtOAc/hexane). After the disappearance of the starting
carboxylic acid, the reaction was cooled to room temperature, and
water (10 mL) was added. The resulting ethyl ester was extracted
with EtOAc (3 ꢁ 15 mL), combining the organic phases, that were
dried over Na2SO4. In the reactions with phenolic acids 1b and 1c,
an additional washing step of the combined organic phases was
performed, first with an NaHCO3 aqueous saturated solution
(15 mL) and later with brine (20 mL). After filtration and solvent
evaporation under reduced pressure, the corresponding ethyl es-
ters were recovered with high purity (2a-n, 75e98%, see Table 1).
Ethyl cinnamate (2a). Colourless liquid (86% isolated yield). Rf
1j
NO2
11
12
13
14
1k
1l
1m
1n
H
H
H
H
H
H
H
H
NO2
CF3
Br
6
78 (67)
100 (90)
19
OCH3
a
Conversion values were calculated through 1H NMR analysis of the reaction
crudes after enzyme filtration and solvent evaporation. Isolated yields after column
chromatography purification appear in parentheses.
order to investigate new sustainable media solutions for the
enzymatic production of alkyl esters, although the higher conver-
sions were found with petroleum-derived solvents such as heptane
and hexane. The use of bio-based solvents such as cyclopentyl
methyl ether, 2-methyltetrahydrofurane or Cyrene™ does not seem
to be a practical alternative, although when exploring the use of
deep eutectic solvents in combination with water, significant con-
versions were achieved, which open the window for the esterifi-
cation of highly insoluble compounds.
Optimisation of the reaction conditions in terms of CAL-B
loading, ethanol equivalents, temperature and reaction time have
allowed the production of a series of ethyl esters, some of them
with complete conversion, and many of them with good isolated
yields after column chromatography purification on silica gel.
Overall, an alternative strategy to the traditional Fischer esterifi-
cation was deeply studied, presenting the use of CAL-B in esterifi-
cation reactions as a complementary approach to traditional
chemical methods or the use of lipase-catalysed transesterification
reactions starting from another ester instead of a carboxylic acid as
starting material [54,55].
(60% EtOAc/hexane): 0.90. IR (
NMR
(300.13 MHz, CDCl3): 7.69 (d, 1H, J ¼ 16.0 Hz), 7.57e7.48 (m,
n .
): 1737, 1577, 1254, 1072 cmꢂ1 1H
d
2H), 7.42e7.35 (m, 3H), 6.44 (d, 1H, J ¼ 16.3 Hz), 4.27 (q, 2H,
J ¼ 7.1 Hz), 1.34 (t, 3H, J ¼ 7.1 Hz) ppm. 13C RMN
d (75.5 MHz, CDCl3):
167.1 (C),144.7 (CH),134.6 (C),130.3 (CH),129.0 (2 CH),128.2 (2 CH),
118.4 (CH), 60.6 (CH2), 14.4 (CH3) ppm.
Ethyl ferulate (2b). Dark yellow colour solid (94% isolated
yield). Melting point 64e65 ꢀC. Rf (50% EtOAc/hexane): 0.74. IR (
n
):
3528, 1737, 1510, 1252, 1031 cmꢂ1.1H NMR
d (300.13 MHz, MeOD-
d4): 7.60 (d, 1H, J ¼ 16.0 Hz), 7.18 (d, 1H, J ¼ 2.0 Hz), 7.06 (dd, 1H,
J ¼ 8.3, 2.0 Hz), 6.80 (d, 1H, J ¼ 8.2 Hz), 6.35 (d, 1H, J ¼ 16.0 Hz), 4.22
(q, 2H, J ¼ 7.1 Hz), 3.89 (s, 3H), 1.31 (t, 3H, J ¼ 7.1 Hz) ppm. 13C NMR
d
(75.5 MHz, MeOD-d4): 169.3 (C), 150.6 (C), 149.3 (C), 146.6 (CH),
127.7 (C), 124.0 (CH), 116.4 (CH), 115.6 (CH), 111.6 (CH), 61.4 (CH2),
56.4 (CH3), 14.6 (CH3) ppm.
Ethyl caffeate (2c). Pale yellow colour solid (96% isolated yield).
Melting point 144e148 ꢀC. Rf (40% EtOAc/hexane): 0.37. IR (
n
): 3450,
4. Experimental section
1658, 1278, 1041 cmꢂ1. 1H NMR
d (300.13 MHz, MeOD-d4): 7.53 (d,
1H, J ¼ 15.9 Hz), 7.03 (d, 1H, J ¼ 2.00 Hz), 6.93 (dd, 1H, J ¼ 8.2,
2.0 Hz), 6.77 (d, 1H, J ¼ 8.2 Hz), 6.24 (d, 1H, J ¼ 15.9 Hz), 4.21 (q, 2H,
All chemical reagents were obtained from Sigma-Aldrich and
used as received. The corresponding DES were prepared by heating
the corresponding hydrogen bond donor (HBD, urea or glycerol)
and acceptor (ABD, ChCl) overnight (ChCl:Gly 1:2 mol/mol at 60 ꢀC;
ChCl:urea 1:2 mol/mol at 100 ꢀC). Initial experiments were per-
formed with the chemicals as received but since especially ChCl is
highly hygroscopic, urea and ChCl were dried in a high vacuum
pump for a few days prior to be mixed under nitrogen atmosphere
(ChCl at 60 ꢀC and urea at room temperature). Candida antarctica
lipase type B (CAL-B, Novozym® 435, 7300 PLU/g) was obtained
from Novozymes.
J ¼ 7.1 Hz), 1.30 (t, 3H, J ¼ 7.1 Hz) ppm. 13C NMR
d (75.5 MHz, MeOD-
d4): 169.3 (C), 149.5 (2 C), 146.7 (CH), 127.7 (C), 122.9 (CH), 116.4
(CH), 115.2 (CH), 115.0 (CH), 61.4 (CH2), 14.6 (CH3) ppm.
Ethyl 4-nitrocinnamate (2d). Scaly yellow solid (94% isolated
yield). Melting point 137e140 ꢀC. Rf (30% EtOAc/hexane): 0.82. IR
(n d (300.13 MHz, DMSO‑d6):
): 1708, 1644, 1390, 1110 cmꢂ1. 1H NMR
8.23 (d, 2H, J ¼ 8.8 Hz), 8.01 (d, 2H, J ¼ 8.8 Hz), 7.75 (d, 1H,
J ¼ 16.1 Hz), 6.85 (d, 1H, J ¼ 16.1 Hz), 4.21 (q, 2H, J ¼ 7.1 Hz), 1.27 (t,
3H, J ¼ 7.1 Hz) ppm. 13C NMR
d (75.5 MHz, DMSO‑d6): 165.7 (C),
148.1 (C),141.8 (CH),140.5 (C),129.5 (2 CH),123.9 (2 CH),122.5 (CH),
60.5 (CH2), 14.2 (CH3).
Thin layer chromatographies (TLCs) were conducted with silica
gel precoated plates and visualised with UV and potassium per-
manganate stain. Column chromatographies were performed using
Ethyl 4-trifluoromethylcinnamate (2e). Colourless liquid (98%
isolated yield). Rf (30% EtOAc/hexane): 0.79. IR (n): 1705, 1631, 1397,
4

ꢀ
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L. Suarez-Escobedo and V. Gotor-Fernandez
Tetrahedron 81 (2021) 131873
1067 cm ꢂ1. 1H NMR
d
(300.13 MHz, MeOD-d4): 7.72e7.56 (m, 5H),
J ¼ 5.8 Hz), 123.5 (CH), 61.9 (CH2), 14.5 (CH3) ppm. 19F NMR
6.56 (d, 1H, J ¼ 16.0 Hz), 4.22 (q, 2H, J ¼ 7.1 Hz), 1.29 (t, 3H,
d
(282 MHz, MeOD-d4): 60.2 ppm.
J ¼ 7.1 Hz). 13C NMR
d
(75.5 MHz, MeOD-d4): 168.0 (C), 144.0 (CH),
Ethyl 2-bromocinnamate (2m). Colourless liquid (98% isolated
139.5 (C), 129.7 (2 CH), 127.2 (C), 126.8 (q, 2 CH, J ¼ 3.8 Hz), 123.6 (q,
yield). Rf (30% EtOAc/hexane): 0.9. IR (
n): 1708, 1634, 1094,
C, J ¼ 271.3 Hz), 121.9 (CH), 61.9 (CH2), 14.6 (CH3) ppm. 19F NMR
593 cmꢂ1. 1H NMR
d
(300.13 MHz, CDCl3): 8.04 (d, 1H, J ¼ 15.9 Hz),
d
(282 MHz, MeOD-d4): 64.4 ppm.
7.59 (dt, 2H, J ¼ 7.8,1.8 Hz), 7.31 (tdd,1H, J ¼ 7.9,1.3, 0.6 Hz), 7.21 (td,
1H, J ¼ 7.7, 1.7 Hz), 6.38 (d, 1H, J ¼ 15.9 Hz), 4.27 (q, 2H, J ¼ 7.1 Hz),
Ethyl 4-methylcinnamate (2f). Colourless liquid (96% isolated
yield). Rf (20% EtOAc/hexane): 0.70. IR (
n
): 2872, 1705, 1634,
1.34 (t, 3H, J ¼ 7.1 Hz). 13C NMR
d (75.5 MHz, CDCl3): 166.5 (C), 143.0
1095 cmꢂ1 1H NMR
.
d
(300.13 MHz, MeOD-d4): 7.64 (d, 1H,
(CH), 134.6 (C), 133.5 (CH), 131.3 (CH), 127.9 (CH), 127.8 (CH), 125.4
(C), 121.2 (CH) 60.8 (CH2), 14.4 (CH3) ppm.
J ¼ 16.0 Hz), 7.47 (d, 2H, J ¼ 8.1 Hz), 7.21 (d, 2H, J ¼ 8.00 Hz), 6.45 (d,
1H, J ¼ 16.0 Hz), 4.23 (q, 2H, J ¼ 7.1 Hz), 2.35 (s, 3H), 1.32 (t, 3H,
Ethyl 2-methoxycinnamate (2n). Colourless liquid (95% iso-
J ¼ 7.1 Hz) ppm. 13C NMR
d
(75.5 MHz, MeOD-d4): 168.8 (C), 146.1
lated yield). Rf (20% EtOAc/hexane): 0.51. IR (
n): 1703, 1629,
(CH), 142.1 (C), 133.0 (C), 130.6 (CH), 129.2 (CH), 117.8 (CH), 61.5
(CH2), 21.4 (CH3), 14.6 (CH3) ppm.
1095 cmꢂ1 1H NMR
.
d
(300.13 MHz, MeOD-d4): 7.96 (d, 1H,
J ¼ 16.2 Hz), 7.55 (dd, 1H, J ¼ 7.7, 1.6 Hz), 7.41e7.33 (m, 1H),
Ethyl 3-methylcinnamate (2g). Colourless liquid (87% isolated
7.07e6.91 (m, 2H), 6.53 (d, 1H, J ¼ 16.2 Hz), 4.22 (q, 2H, J ¼ 7.1 Hz),
yield). Rf (30% EtOAc/hexane): 0.90. IR (
n
): 2870, 1705, 1635,
3.89 (s, 3H), 1.31 (t, 3H, J ¼ 7.1 Hz) ppm. 13C NMR
d (75.5 MHz,
1093 cmꢂ1. 1H NMR
d
(300.13 MHz, CDCl3): 7.69 (d, 1H, J ¼ 16.0 Hz),
MeOD-d4): 169.2 (C), 159.8 (C), 141.3 (CH), 132.9 (CH), 129.7 (CH),
124.2 (C), 121.8 (CH), 119.1 (CH), 112.4 (CH), 61.5 (CH2), 56.0 (CH3),
14.6 (CH3) ppm.
7.35e7.19 (m, 4H), 6.45 (d, 1H, J ¼ 16.0 Hz), 4.29 (q, 2H, J ¼ 7.1 Hz),
2.38 (s, 3H), 1.36 (t, 3H, J ¼ 7.1 Hz) ppm. 13C NMR
d (75.5 MHz,
CDCl3): 167.1 (C), 144.8 (CH), 138.5 (C), 134.3 (C), 131.1 (CH), 128.8
(CH), 128.7 (CH), 125.3 (CH), 118.0 (CH), 60.5 (CH2), 21.3 (CH3), 14.4
(CH3) ppm.
4.2. General procedure for the esterification of carboxylic acids 1a-
n using CAL-B
Ethyl 3-methoxycinnamate (2h). Colourless liquid (98% iso-
lated yield). Rf (30% EtOAc/hexane): 0.87. IR (
n): 1704, 1635,
Ethanol (87.6 mL, 1.5 mmol) was added to a suspension of the
1094 cmꢂ1. 1H NMR
d
(300.13 MHz, CDCl3): 7.65 (d, 1H, J ¼ 16.0 Hz),
corresponding carboxylic acid 1a-n (0.5 mmol: 74.1 mg for 1a,
97.1 mg for 1b, 90.1 mg for 1c, 96.6 mg for 1d, 1j and 1k, 108.1 mg
for 1e and 1l, 81.1 mg for 1f and 1g, 89.1 mg for 1h and 1n, 83.1 for
1i, and 113.5 mg for 1m) and CAL-B (1:1 substrate:enzyme w/w
ratio) in the desired solvent (5 mL, 100 mM of 1a-n, see Fig. 2). The
mixture was shaken in an orbital shaker for 72 h at 45 ꢀC and
250 rpm. After this time, the enzyme was filtered-off and washed
with ethyl acetate (2 ꢁ 5 mL). The solvent was distilled under
reduced pressure, affording a reaction crude that was analysed by
1H NMR to calculate the conversion of the lipase-catalysed esteri-
fication reaction. For those reactions leading to high conversion
values, chromatography purification on silica gel were performed
to isolated the corresponding ethyl esters and compared with the
standards produced by chemical esterification (see Tables 1 and 4).
Column chromatographies on silica gel were made using 10.0 g SiO2
and mixtures of ethyl acetate and hexane as eluents (10e30%
EtOAc/hexane).
7.30 (d, 1H, J ¼ 7.9 Hz), 7.11 (dt, 1H, J ¼ 7.6, 1.3 Hz), 7.03 (t, 1H,
J ¼ 2.1 Hz), 6.92 (ddd, 1H, J ¼ 8.2, 2.6, 1.0 Hz), 6.42 (d, 1H,
J ¼ 16.0 Hz), 4.26 (q, 2H, J ¼ 7.1 Hz), 3.81 (s, 3H), 1.33 (t, 3H,
J ¼ 7.1 Hz) ppm. 13C NMR
d (75.5 MHz, CDCl3): 167.1 (C), 160.0 (C),
144.6 (CH), 135.9 (C), 130.0 (CH), 120.8 (CH), 118.6 (CH), 116.2 (CH),
113.0 (CH), 60.6 (CH2), 55.4 (CH3), 14.4 (CH3) ppm.
Ethyl 3-fluorocinnamate (2i). Colourless liquid (75% isolated
yield). Rf (30% EtOAc/hexane): 0.90. IR (
n): 1706, 1640, 1392,
1076 cmꢂ1 1H NMR
.
d
(300.13 MHz, MeOD-d4): 7.62 (d, 1H,
J ¼ 16.0 Hz), 7.44e7.30 (m, 3H), 7.16e7.07 (m, 1H), 6.51 (d, 1H,
J ¼ 16.0 Hz), 4.23 (q, 2H, J ¼ 7.1 Hz), 1.31 (t, 3H, J ¼ 7.1 Hz). 13C NMR
d
(75.5 MHz, MeOD-d4): 166.0 (C), 163.0 (d, C, J ¼ 413.3 Hz),144.5 (d,
CH, J ¼ 2.5 Hz), 138.1 (d, C, J ¼ 7.8 Hz), 131.8 (CH, d, J ¼ 7.9 Hz), 125.4
(d, CH, J ¼ 3.2 Hz), 120.6 (CH), 118.1 (d, CH, J ¼ 21.6 Hz), 115.2 (d, CH,
J ¼ 23.0 Hz), 61.8 (CH2), 14.6 (CH3) ppm. 19F NMR
MeOD-d4): 114.8 ppm.
d (282 MHz,
Ethyl 3-nitrocinnamate (2j). Scaly white solid (98% isolated
yield). Melting point 73e76 ꢀC Rf (30% EtOAc/hexane): 0.84. IR (
):
1712, 1643, 1392, 1100 cmꢂ1. 1H NMR
(300.13 MHz, CDCl3): 8.37 (t,
n
4.3. General procedure for the esterification of cinnamic acid (1a)
using CAL-B in DES as reaction media
d
1H, J ¼ 2.0 Hz), 8.22 (ddd, 1H, J ¼ 8.2, 2.3, 1.1 Hz), 7.82 (d, 1H,
J ¼ 8.0 Hz), 7.70 (d, 1H, J ¼ 16.0 Hz), 7.58 (t, 1H, J ¼ 8.0 Hz), 6.55 (d,
1H, J ¼ 16.0 Hz) 4.28 (q, 2H, J ¼ 7.1 Hz),1.34 (t, 3H, J ¼ 7.1 Hz) ppm.
Ethanol (35.0
mL, 0.6 mmol) and CAL-B (29.6 mg, 1:1 sub-
13C NMR
d (75.5 MHz, CDCl3): 166.2 (C), 148.8 (C), 141.8 (CH), 136.3
strate:enzyme w/w ratio) was added to a mixture of cinnamic acid
(1a, 29.6 mg, 0.2 mmol) and the corresponding eutectic mixture
(2 mL). For these experiments involving the influence of water, the
total volume of the (DES:H2O) mixture was 2 mL for a final 100 mM
cinnamic acid concentration (see Table 2). The mixture was shaken
in an orbital shaker for 24 h at 45 ꢀC and 250 rpm. After this time,
the enzyme was filtered-off and washed with ethyl acetate
(5 ꢁ 5 mL), finding two phases that were shaken in a vortex stirrer
(3 ꢁ 10 s). The upper organic phase with the organic compounds 1a
and 2a dissolved in EtOAc were collected, and the solvent distilled
under reduced pressure. The resulting reaction crude was analysed
by 1H NMR to measure the reaction conversion.
(C), 133.7 (CH), 130.1 (CH), 124.6 (CH), 122.5 (CH), 121.6 (CH), 61.0
(CH2), 14.4 (CH3) ppm.
Ethyl 2-nitrocinnamate (2k). Brown liquid (93% isolated yield).
Rf (30% EtOAc/hexane): 0.80. IR (
NMR
n
): 1711, 1638, 1392, 1095 cmꢂ1. 1H
(300.13 MHz, DMSO‑d6): 8.08 (dd, 1H, J ¼ 8.1, 1.4 Hz),
d
8.00e7.86 (m, 2H), 7.78 (td, 1H, J ¼ 7.7, 1.4 Hz), 7.68 (td, 1H, J ¼ 7.7,
1.5 Hz), 6.63 (d, 1H, J ¼ 15.8 Hz), 4.22 (q, 2H, J ¼ 7.1 Hz), 1.26 (t, 3H,
J ¼ 7.1 Hz) ppm. 13C NMR
d (75.5 MHz, DMSO‑d6): 165.5 (C), 148.3
(C), 139.5 (CH), 133.9 (CH), 131.0 (CH), 129.3 (CH), 129.2 (C), 124.7
(CH), 122.6 (CH), 60.5 (CH2), 14.1 (CH3) ppm.
Ethyl 2-trifluoromethylcinnamate (2l). Pale yellow liquid (79%
isolated yield). Rf (30% EtOAc/hexane): 0.81. IR (
n): 1714, 1632, 1393,
1061 cmꢂ1. 1H NMR
d
(300.13 MHz, MeOD-d4): 8.00 (dq,1H, J ¼ 15.8,
Declaration of competing interest
2.3 Hz), 7.89e7.81 (m, 1H), 7.75e7.48 (m, 3H), 6.52 (d, 1H,
J ¼ 15.8 Hz), 4.25 (q, 2H, J ¼ 7.1 Hz), 1.31 (t, 3H, J ¼ 7.1 Hz) ppm. 13
C
The authors declare that they have no known competing
financial interests or personal relationships that could have
appeared to influence the work reported in this paper.
NMR
d (75.5 MHz, MeOD-d4): 167.6 (C), 140.9 (CH), 134.3 (C), 133.7
(CH), 131.1 (CH), 129.8 (C), 129.4 (C), 129.2 (CH), 127.2 (q, CH,
5

ꢀ
ꢀ
L. Suarez-Escobedo and V. Gotor-Fernandez
Tetrahedron 81 (2021) 131873
ꢀ
[21] V. Gotor-Fernandez, E. Busto, V. Gotor, Adv. Synth. Catal. 348 (2006) 797e812.
[22] E.M. Anderson, K.M. Larsson, O. Kirk, Biocatal. Biotransform. 16 (1998)
181e204.
Acknowledgments
Financial support of this work by the Spanish Ministry of Science
and Innovation (PID2019-109253RB-I00) and the Asturian Regional
[23] Y-g. Shi, Y. Wu, X.-y. Lu, Y.-p. Ren, Q. Wang, C.-m. Zhu, D. Yu, H. Wang, Food
Chem. 220 (2017) 249e256.
[24] K.-C. Huang, Y. Li, C.-H. Kuo, Y.-K. Twu, C.-J. Shieh, Molecules 21 (2016) 478.
[25] Y. Wang, D.-H. Zhang, N. Chen, G.-Y. Zhi, Bioresour. Technol. 198 (2015)
256e261.
[26] V. Kumar, F. Jahan, K. Kameswaran, R.V. Mahajan, R.K. Saxena, J. Ind. Micro-
biol. Biotechnol. 41 (2014) 907e912.
[27] J. Wang, S.-S. Gu, H.-S. Cui, L.-Q. Yang, X.-Y. Wu, Bioresour. Technol. 149
(2013) 367e374.
Government
(FC-GRUPIN-IDI/2018/000181)
are
gratefully
acknowledged. We thank Novozymes for the generous gift of CAL-B
and the technical support offered from the Scientific-Technical
Services of the University of Oviedo.
Appendix A. Supplementary data
[28] N. Pang, S.-S. Gu, J. Wang, H.-S. Cui, F.-Q. Wang, X. Liu, X.-Y. Zhao, F.-A. Wu,
Bioresour. Technol. 139 (2013) 337e342.
ꢀ
ꢀ
ꢀ
[29] S.M. Jakovetic, B.Z. Jugovic, M.M. Gvozdenovic, D.I. Bezbradica, M.G. Antov,
Supplementary data associated with this article can be found in
the online version, at https://doi.org/10.1016/j.tet.2020.131873.
These data include NMR spectra, MOL files and InChiKeys of the
most important compounds described in this article.
ꢁ
ꢁ
ꢀ
ꢀ
D.Z. Mijin, Z.D. Knezevic-Jugovic, Appl. Biochem. Biotechnol. 170 (2013)
1560e1573.
[30] C.K. Sharma, S.S. Kanwar, Res. J. Recent Sci. 1 (2012) 68e71.
[31] M.H. Katsoura, A.C. Polydera, L.D. Tsironis, M.P. Petraki, S.K. Rajacic,
ꢁ ꢀ
A.D. Tselepis, H. Stamatis, N. Biotech. 26 (2009) 83e91.
[32] D.E. Stevenson, S.G. Parkar, J. Zhang, R.A. Stanley, D.J. Jensen, J.M. Cooney,
Enzym. Microb. Technol. 40 (2007) 1078e1086.
[33] J. Cassani, H. Luna, A. Navarro, E. Castillo, Electron. J. Biotechnol. 10 (2007)
509e513.
References
[1] L. Chen, L. Zhang, G. Yan, D. Huang, Asian J. Org. Chem. 9 (2020) 842e862.
[2] B. Vanholme, I. El Houari, W. Boerjan, Curr. Opin. Biotechnol. 56 (2019)
156e162.
[34] K. Vosmann, P. Witkamp, N. Weber, J. Agric. Food Chem. 54 (2006)
2969e2976.
[35] G.-S. Lee, A. Widjaja, Y.-H. Ju, Biotechnol. Lett. 28 (2006) 581e585.
[36] B. Guyot, B. Bosquette, M. Pina, J. Graille, Biotechnol. Lett. 19 (1997) 529e532.
^
[3] A.T. Silva, C.M. Bento, A.C. Pena, L.M. Figueiredo, C. Prudencio, L. Aguiar,
T. Silva, R. Ferraz, M.S. Gomes, C. Teixeira, P. Gomes, Molecules 25 (2020) 66.
[4] E. Pontiki, A. Peperidou, I. Fotopoulos, D. Hadjipavlou-Litina, Curr. Pharmaceut.
Biotechnol. 19 (2018) 1019e1048.
[5] P. De, F. Bedos-Belval, C. Vanucci-Bacque, M. Baltas, Curr. Org. Chem. 16 (2012)
747e768.
ꢀ
[37] G. de Gonzalo, A.R. Alcantara, P. Domínguez de María, ChemSusChem 12
(2019) 2083e2097.
[38] U. Azzena, M. Carraro, L. Pisano, S. Monticelli, R. Bartolotta, V. Pace, Chem-
SusChem 12 (2019) 40e70.
ꢀ
[39] V. Pace, P. Hoyos, L. Castoldi, P. Domínguez de María, A.R. Alcantara, Chem-
[6] A.C. Fonseca, M.S. Lima, A.F. Sousa, A.J. Silvestre, J.F.J. Coelho, A.C. Serra, Polym.
Chem. 10 (2019) 1696e1723.
SusChem 5 (2012) 1369e1379.
[40] N. Guajardo, P. Domínguez de María, Mol. Catal. 485 (2020) 110813.
[41] Y. Liu, J.B. Friesen, J.B. McAlpine, D.C. Lankin, S.-N. Chen, G.F. Pauli, J. Nat. Prod.
81 (2018) 679e690.
ꢀ
ꢀ
[7] O. Taofiq, A.M. Gonzalez-Paramas, M.F. Barreiro, I.C. Ferreira, Molecules 22
(2017) 281.
[8] G. Murtaza, S. Karim, M.R. Akram, S.A. Khan, S. Azhar, A. Mumtaz,
M.H.H.A. Asad, BioMed Res. Int. (2014) 145342.
[9] B. Hauer, ACS Catal. 10 (2020) 8418e8427.
[10] R. Mu, Z. Wang, M.C. Wamsley, C.N. Duke, P.H. Lii, S.E. Epley, L.C. Todd,
P.J. Roberts, Catalysts 10 (2020) 832.
[11] R.A. Sheldon, D. Brady, M.L. Bode, Chem. Sci. 11 (2020) 2587e2605.
[12] R.A. Sheldon, D. Brady, ChemSusChem 12 (2019) 2859e2881.
[42] D.A. Alonso, A. Baeza, R. Chinchilla, G. Guillena, I.M. Pastor, D.J. Ramon, Eur. J.
Org Chem. (2016) 612e632.
ꢀ
[43] J. García-Alvarez, E. Hevia, V. Capriati, Chem. Eur J. 24 (2018) 14854e14863.
[44] A. Paiva, R. Craveiro, I. Aroso, M. Martins, R.L. Reis, A.R.C. Duarte, ACS Sustain.
Chem. Eng. 2 (2014) 1063e1071.
[45] A.P. Abbott, D. Boothby, G. Capper, D.L. Davies, R.K. Rasheed, J. Am. Chem. Soc.
126 (2004) 9142e9147.
ꢀ
ꢀ
ꢀ
[13] J. Albarran-Velo, D. Gonzalez-Martínez, V. Gotor-Fernandez, Biocatal. Bio-
transform. 36 (2018) 102e130.
[46] C. Aranda, G. de Gonzalo, Molecules 25 (2020) 3016.
[47] J.-N. Tan, Y. Dou, Appl. Microbiol. Biotechnol. 104 (2020) 1481e1496.
[48] I. Juneidi, M. Hayyan, M.A. Hashim, Process Biochem. 66 (2018) 33e60.
[49] P. Xu, G.-W. Zheng, M.-H. Zong, N. Li, W.-Y. Lou, Bioresour. Bioprocess. 4
(2017) 34.
[14] U.T. Bornscheuer, R.J. Kazlauskas, Hydrolases in Organic Synthesis: Regio- and
Stereoselective Transformations, second ed., Wiley-VCH Verlag GmbH & Co.
KGaA, Weinheim, 2005.
[15] J. Mulinari, J.V. Oliveira, D. Hotza, Biotechnol. Adv. 42 (2020) 107581.
[16] R.C. Rodrigues, J.J. Virgen-Ortíz, J.C.S. dos Santos, A. Berenguer-Murcia,
ꢀ
[50] V. Gotor-Fernandez, C.E. Paul, J. Biotechnol. 293 (2019) 24e35.
ꢀ
[51] F.Z.I.M. Hassani, S. Amzazi, I. Lavandera, Molecules 24 (2019) 2190.
[52] N. Guajardo, P. Domínguez de María, K. Ahumada, R.A. Schrebler, R. Ramírez-
Tagle, F.A. Crespo, C. Carlesi, ChemCatChem 9 (2017) 1393e1396.
ꢀ
A.R. Alcantara, O. Barbosa, C. Ortiz, R. Fernandez-Lafuente, Biotechnol. Adv. 37
(2019) 746e770.
[17] Z. Zhao, M.-G. Zhou, R.-L. Liu, Catalysts 9 (2019) 647.
[18] J. Chapman, A.E. Ismail, C.Z. Dinu, Catalysts 8 (2018) 238.
[19] C. Ortiz, M.L. Ferreira, O. Barbosa, J.C.S. dos Santos, R.C. Rodrigues,
€
[53] M. Patzold, S. Siebenhaller, S. Kara, S. Liese, C. Syldatk, D. Holtmann, Trends
Biotechnol. 37 (2019) 943e959.
[54] A. Kurata, Y. Kitamura, S. Irie, S. Takemoto, Y. Akai, Y. Hirota, T. Fujita, K. Iwai,
M. Furusawa, N. Kishimoto, J. Biotechnol. 148 (2010) 133e138.
[55] N. Nakajima, K. Ishihara, S. Matsumura, H. Hamada, K. Nakamura, T. Furuya,
Biosci. Biotechnol. Biochem. 61 (1997) 1926e1928.
ꢀ
ꢀ
A. Berenguer-Murcia, L.E. Briand, R. Fernandez-Lafuente, Catal. Sci. Technol. 9
(2019) 2380e2420.
[20] R.N. Lima, C.S. dos Anjos, E.V.M. Orozco, A.L.M. Porto, Mol. Catal. 466 (2019)
75e105.
6