Activity and specificity studies of the new thermostable esterase EstDZ2

Article abstract of DOI:10.1016/j.bioorg.2020.104214

In this paper, we study the activity and specificity of EstDZ2, a new thermostable carboxyl esterase of unknown function, which was isolated from a metagenome library from a Russian hot spring. The biocatalytic reaction employing EstDZ2 proved to be an efficient method for the hydrolysis of aryl p-, o- or m-substituted esters of butyric acid and esters of secondary alcohols. Docking studies revealed structural features of the enzyme that led to activity differences among the different substrates.

Full text of DOI:10.1016/j.bioorg.2020.104214

BioorganicChemistry104(2020)104214
Contents lists available at ScienceDirect
Bioorganic Chemistry
journal homepage: www.elsevier.com/locate/bioorg
Activity and specificity studies of the new thermostable esterase EstDZ2
Kamela Myrtollaria, Nikolaos Katsoulakisa, Dimitra Zarafetab, Ioannis V. Pavlidisa,
⁎
Georgios Skretasb, Ioulia Smonoua,
a Department of Chemistry, University of Crete, University Campus-Voutes, 70013 Heraklion, Crete, Greece
b Institute of Chemical Biology, National Hellenic Research Foundation, 11635 Athens, Greece
A R T I C L E I N F O
A B S T R A C T
Keywords:
In this paper, we study the activity and specificity of EstDZ2, a new thermostable carboxyl esterase of unknown
function, which was isolated from a metagenome library from a Russian hot spring. The biocatalytic reaction
employing EstDZ2 proved to be an efficient method for the hydrolysis of aryl p-, o- or m-substituted esters of
butyric acid and esters of secondary alcohols. Docking studies revealed structural features of the enzyme that led
to activity differences among the different substrates.
Thermostable esterase
Hydrolysis
Kinetic resolution
In-silico analysis
1. Introduction
Besides their ability to catalyze hydrolysis, lipolytic enzymes can also
catalyze synthesis of ester bonds or transesterification reactions in non-
Lipolytic enzymes (EC 3.1.1.x) catalyze the hydrolysis of carboxyl
ester bonds and they can be further grouped into several subclasses
based on their substrate specificity and their capacity to hydrolyze es-
ters in solution and emulsion. EC 3.1.1.1 represents carboxylesterases,
enzymes which are active on solutions of short acyl chain esters [1].
The second more widely used subclass includes lipases, which are re-
presented as EC 3.1.1.3 enzymes [2]. It is generally established that
lipases have the unique capacity to cleave carboxylic ester bonds of
water-insoluble long chain triacylglycerols including natural fats.
Moreover, lipases, unlike esterases, are interfacially activated [3].
However, this classical distinction between lipases and esterases
should no longer be considered as the main distinction between these
hydrolytic enzymes, as kinetic and structural studies have revealed that
there are lipases that can hydrolyze water-insoluble-long-chain esters
without bearing a lid domain [4]. Therefore, the distinction between
esterases and lipases should no longer be correlated with the physical
state of the substrate and/or the presence of a lid domain in the enzyme
molecule [5].
aqueous media, following the thermodynamic reversion of the hydro-
lysis reaction route [9]. All these features render them protagonist
biocatalysts in the industry [10].
Their biotechnological significance is reflected by the fact that there
is an enormous effort to find techniques and methods, which will allow
exploration of microbial organisms residing in extreme environments
[11]. Metagenomics is one of the approaches, which aims to address
this issue and bypass the limitations imposed by current culturing
techniques [12]. Such recently developed techniques have provided
access to new esterases or lipases bearing novel sequences and im-
portant features, such as thermostability, which is a crucial prerequisite
for industrial applications.
EstDZ2 is a new lipolytic enzyme, which we have recently isolated
from a metagenomic sample from a hot spring located in the Kamchatka
Peninsula (Russia) as a part of the international project Hotzyme, which
aims at identifying novel thermostable hydrolytic enzymes derived
from hot springs with properties suitable for industrial applications
[13]. The isolated esterase, was cloned, purified from Escherichia coli
and characterized biochemically [14]. According to activity and spe-
cificity studies EstDZ2 can efficiently catalyze the hydrolysis of short to
medium-length acyl chains esters [15]. The biochemical characteriza-
tion proved that this new esterase is a thermostable enzyme with a half-
life of more than six hours and excellent stability at high concentrations
of organic solvents [15]. These properties suggest that EstDZ2 could be
utilized in many different industrial applications. Furthermore, EstDZ2
is an interesting enzyme from a phylogenetic point of view, since its
amino acid sequence did not cluster with that of any previously
Lipolytic enzymes constitute one of the most abundant class of en-
zymes and they have received great attention as they participate in a
variety of biological processes due to their functional diversity [6].
Their functions are pivotal for the human body via digestion, as they
provide building blocks for biosynthesis, help the detoxification and
provide carbon sources for energy [7]. In addition to their fundamental
biological importance, lipolytic enzymes have many potential industrial
and medicinal applications. Increasing interest revolves around their
potential applications in the pulp- and paper-making industries [8].
⁎ Corresponding author at: Department of Chemistry, University of Crete, 70013, Heraklion, Crete, Greece.
E-mail address: smonou@uoc.gr (I. Smonou).
https://doi.org/10.1016/j.bioorg.2020.104214
Received 28 February 2020; Received in revised form 23 July 2020; Accepted 30 July 2020
Availableonline29August2020
0045-2068/©2020ElsevierInc.Allrightsreserved.

K. Myrtollari, et al.
BioorganicChemistry104(2020)104214
characterized bacterial esterolytic enzyme and, thus, it was found to
belong to a putative new family of bacterial esterolytic enzymes, for
which we have proposed the index XV [15].
monitoring the EstDZ2-selectivity depending on electronic and/or steric
effects. The progress of the reaction was monitored at various time
intervals (5, 10, 15, 30, 60 min) as well as at 3 h, 5 h and 24 h, by
assessing the conversion of the ester to the corresponding alcohol.
Conversion rate was determined by gas chromatography using nonpolar
column after comparing the retention time of the enzymatic reaction’s
products with the elution time of the corresponding ester and the al-
cohol. Before conducting the screening with EstDZ2, we tested the
stability of each substrate ester under the standard enzymatic reaction
conditions without addition of the enzyme (25 mM Tris-HCl buffer, pH
8 with 0.05% Triton X-100, 5.49 μmol of substrate, 50 °C). All the
substrates are soluble in aqueous buffered solution, under the reaction
conditions and only a small amount of alcohol was detected for most of
the tested esters after 1 h in the buffer solution (2–8%).
In this work, we have examined in more detail the biochemical
profile of the new thermostable esterase EstDZ2 by studying its re-
activity and specificity towards aryl p-substituted esters of butyric acid
and esters of secondary alcohols, and have found that EstDZ2 can
successfully hydrolyze a wide range of esters. In silico analysis reveals
that the activity of this new enzyme is a function of the geometry of the
active site and the structure of the bulky substrate. These are the first
reported reactions catalyzed by EstDZ2 and may eventually lead to
utilization of this enzyme as a potent catalyst in organic synthesis.
2. Results and discussion
As presented in Table 1, among the bromo substituted esters (entries
6–8), the o-bromophenyl ester (entry 8) was hydrolyzed more effi-
ciently than the para and meta isomers. Interestingly, a similar trend
was observed for nitro substituted esters (entries 2–4), among which
better catalytic activity was observed for o-nitrophenyl butyrate (entry
4). These results demonstrate that EstDZ2 activity depends significantly
on steric effects which determine the substrate acceptance. Among es-
ters bearing substituents with altered electronic properties in para po-
sition (entries 2, 5, 6, 9 and 10), enhanced specific activity was ob-
served for esters with halogen substituents (e.g. Br, F) (entries 6 and 9).
However, in the case of a strong electron-withdrawing or electron do-
nating substituent such as nitro and methoxy groups correspondingly,
(p-nitrophenyl- and p-methoxyphenyl butyrates, entries 2 and 5) led to
similar moderately enhanced activity. This result indicates that EstDZ2-
catalyzed hydrolysis is independent on electronic effects. It appears that
steric effects outweigh electronic effects and thus, mainly determine
substrate acceptance. The highest hydrolytic activity among all tested
esters was observed in the case of the larger bromo substituent in
substrate o-bromophenyl butyrate (entry 8). This observation also
confirms the steric dependence of the reaction activity. Highly efficient
hydrolysis for all tested esters was observed at 5 h (conversions 85-
> 99%), after addition of 0.3 μΜ EstDZ2 to 8 μmol of substrate in 4 mL
Tris-HCl buffer 25 mM, pH 8.0, 0.05% Triton X-100, at 50 °C.
Recently published detailed biochemical characterization of EstDZ2
revealed that the three-dimensional modelled structure of the new
hydrolytic enzyme is characterized by the α/β hydrolase fold, forming a
twisted β-sheet [15]. The catalytic triad is constituted by residues of
Ser, Asp and His, which are typical for esterolytic enzymes. Due to its
novel sequence and functional characteristics, EstDZ2 is classified to a
new family of bacterial esterolytic enzymes [15]. To investigate its
esterolytic activity and optimize reaction conditions, p-nitrophenyl
butyrate was used as a substrate since it is a pH-indicator and there is
significant literature for p-nitrophenyl butyrate resolutions catalyzed by
hydrolases [16–18]. The reaction conditions were optimized after
measurements of EstDZ2 activity in a wide range of pH and tempera-
tures, where it was found that EstDZ2 exhibits its maximal activity at
50 °C and pH 8.0 as also reported previously [15].
Initial screening of the new esterase with different p-nitrophenyl
esters derived from fatty acids revealed that EstDZ2 can hydrolyze
short- to medium-chain lengths fatty acid esters [15]. For esters with
long chains, hydrolysis was barely detected. Optimal catalytic effi-
ciency was detected for p-nitrophenyl butyrate [15].
Based on all these previous results, first we performed in silico
analyses to provide insights of substrate structural features that influ-
ence enzyme activity and selectivity. Docking experiments performed
for p-nitrophenyl butyrate and p-nitrophenyl octanoate hydrolysis cat-
alyzed by EstDZ2 agreed with the results from the activity studies. More
specifically, our in silico analyses revealed that binding of the p-ni-
trophenyl butyrate is more favorable as the substrate fits perfectly in
the active site of the enzyme, thus EstDZ2 exhibits higher activity. As
shown in Fig. 1, p-nitrophenyl octanoate binds in a different manner. In
this case, a large part of the substrate does not fit in the active site but it
is oriented towards the solvent and, therefore, this conformation is not
very catalytically productive. These results are in complete agreement
with the activity studies and underline that the activity of EstDZ2 is
largely determined by the geometry of the active site and the structure
of the bulky substrate. To depict the active site in a similar way in both
cases (Fig. 1a and b), the catalytic serine was set to be in the lower
center and the histidine of the catalytic triad in the middle left of the
figures.
In an effort to gain deeper insight into the activity or enantios-
electivity profile of EstDZ2, we investigated the hydrolysis of esters of
secondary alcohols as substrates bearing one stereogenic center. They
were synthesized by acetylation of the corresponding alcohols with
acetic anhydride (Ac2O) in anhydrous ethyl acetate (Fig. 3) [19].
Rac-1-phenylethyl acetate 11 was used for the standard reaction.
Before proceeding to EstDZ2-catalyzed hydrolysis of this ester, in silico
analysis was performed to predict which enantiomer of the ester would
react faster. As seen in Fig. 4, the two enantiomers of rac-11 bind to the
active site of the enzyme in a similar manner. The carboxyl group of
both (R)-11 and (S)-11 faces towards the catalytic Ser141, while both
enantiomers are stabilized via π-π stacking with Phe222. The only
difference between these two conformations is that for (R)-11, distance
between the catalytic Ser and the carbonyl group of the ester is smaller.
This observation indicates that R conformation would be slightly more
catalytically favorable than the S.
With these results at hand, a series of aryl p-, o- or m-substituted
esters of butyric acid was synthesized bearing electron donating or
electron withdrawing groups. The aim was to understand the catalytic
behavior of EstDZ2 by rationally studying two structural elements of
the substrates; (a) the position of the substituent of the aromatic ring
and (b) the electronic character of the ring. The substituents of the
aromatic ring chosen were: -OMe, -NO2, -Br, -F and -propyl. Esters of
butyric acid were readily prepared using a method based on Steglich
esterification, using dicyclohexylcarbodiimide (DCC) and catalytic
amount of 4-dimethylaminopyridine (DMAP) in the presence of dry
dichloromethane (DCM) [19]. All the esters were obtained with good
yields (Fig. 2).
In a typical run, 20 μmol of the substrate rac-11 were diluted in
4 mL of buffer solution (25 mM Tris-HCl pH 8.0 buffer with 0.05%
ο
Triton X-100). The resulting suspension was shaken at 50 C after the
addition of the enzyme. Samples were collected periodically and were
analyzed by GC analysis. Various amounts of enzyme were tested and
the highest activity and enantioselectivity was achieved for 50 μg of
EstDZ2. These were considered to be the optimal reaction conditions for
the stereoselective hydrolysis of esters of secondary alcohols. The ab-
solute configuration of the produced alcohol was determined by GC
analysis using a chiral column. Comparison of the retention time with
the corresponding data for the commercially available optically active
(S)-1-phenylethanol revealed that the R/S enantiomeric ratio of the
produced phenylethanol was 77/23. This faster hydrolysis for the R-
Subsequently, we proceeded to EstDZ2-catalyzed hydrolysis of the
substrates of interest under the optimal reaction conditions, that allow
2

K. Myrtollari, et al.
BioorganicChemistry104(2020)104214
Fig. 1. Docking of p-nitrophenyl octanoate (a) and p-nitrophenyl butyrate (b) in the enzyme intermediate of EstDZ2. The catalytic residues are indicated in red. All
figures were prepared with PYMOL 0.99. The distance from the catalytic Ser141 oxygen to the carbon atom in the carbonyl group is depicted with white dashed line.
(For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)
enantiomer of rac-1-phenylethyl acetate 11 is in total agreement with
the result from the docking studies performed for this substrate. This
also provides extra evidence that the utilized in silico analysis can
predict successfully the preferred enantiomer of substrates.
hydrolysis of esters of secondary alcohols [21]. The enzyme activity
was increased by increasing the size of the larger substituent (L) (entries
13–16). The same trend was observed in the enantioselectivity of the
reaction (entries 13–15). The 1-(4-methoxyphenyl) ethyl acetate 16
was fully hydrolyzed in 24 h and no enantioselectivity was observed
(conv. > 99%, < 1% ee).
In addition to ester 11, analogs with different alkyl, F or -OMe
substituents in para position (esters 13–16) were investigated as well as
the phenylethyl acetylate (ester 12) for the enantioselective hydrolysis
with EstDZ2. All substrates, including the model ester rac-11 (entry 11)
remained stable under the standard reaction conditions for 48 h in the
absence of enzyme. The results obtained are summarized in Table 2.
According to the results shown in Table 2, activity and stereo-
selectivity of EstDZ2 is affected by steric hindrance factors. By in-
creasing the substrate’s chain length from methyl (entry 11) to ethyl
(entry 12) there is a significant increase in the observed conversion at
the same reaction time, but enantioselectivity decreases dramatically
(conversion 17% and 25%, 54% ee and 10% ee respectively). In both
cases EstDZ2 exhibits R enantioselectivity. It is interesting to note here
that these preliminary results indicate that EstDZ2 follows the Ka-
zlauskas rule, which predicts the enantiopreference of lipases towards
3. Conclusions
A detailed activity and specificity analysis of the new thermostable
carboxyl esterase EstDZ2 was performed in this work. EstDZ2 was
found to be highly active in the hydrolysis of aryl-substituted esters of
butyric acid bearing subtituents in the ortho position of the aromatic
ring. While sole steric effects could be explained rationally, the impact
of altered electronic properties did not follow a clear trend in all cases.
These results indicate that substrate acceptance is mainly determined
by steric effects.
Docking experiments revealed structural features of the enzyme that
led to activity differences. By increasing the substrate’s alkyl chain
Fig. 2. Esterification of substituted phenols with butyric acid.
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K. Myrtollari, et al.
BioorganicChemistry104(2020)104214
Table 1
Substrate specificity for the enzymatic hydrolysis of aryl butyrate catalyzed by EstDZ2.
Entry
Substrate
Conversiona
5 (min)
%
10 (min)
15 (min)
30 (min)
60 (min)
1
2
3
4
5
6
7
8
9
10
phenyl-butyrate
26
5
35
9
41
10
14
45
10
70
67
82
25
14
40
27
21
52
13
72
75
88
28
29
43
28
25
72
35
79
71
83
31
56
p-nitrophenyl butyrate
m-nitrophenyl butyrate
o-nitrophenyl butyrate
p-methoxyphenyl butyrate
p-bromophenyl butyrate
m-bromophenyl butyrate
o-bromophenyl butyrate
p-fluorophenyl butyrate
p-propylphenyl butyrate
10
37
8
12
42
13
52
58
56
22
17
36
34
39
20
9
a
Conversions were derived from Gas Chromatography using nonpolar column and are in good agreement with the data obtained from 1H NMR, (error
1%).
Reaction conditions: 4 mL Tris-HCl buffer (25 mM, pH 8.0, 0.05% Triton X-100), substrate 1–10 (5.49 μmol), EstDZ2 (0.04 μΜ), 50 °C.
length from butyl to octyl, the activity of the enzyme decreased sig-
nificantly since larger alkyl-groups do not appear to fit in the active site
of the enzyme. EstDZ2 was also tested for the first time as a catalyst for
the kinetic resolution of secondary alcohols. Preliminary studies re-
vealed that this enzyme exhibits (R)-enantioselectivity. The bulkier the
ester of secondary alcohol is, the higher activity and enantioselectivity
was observed. The stereoselectivity of EstDZ2 was successfully pre-
dicted also by in-silico analysis. Further studies for the kinetic resolution
of secondary alcohols are under investigation in our group.
4.2. Materials
Unless otherwise noted, all solvents and reagents were purchased
from Sigma–Aldrich, Merck, Riedel or Fluka in the highest purity
available and were used without any further purification. EstDZ2 was
prepared as described previously by D. Zarafeta et al. [15].
4.3. General procedure for enzymatic hydrolysis of aryl substituted esters
The enzymatic hydrolysis was performed as follows: In a Tris-HCl
buffer solution (4 mL, 25 mM, pH 8.0), the substrate (5.49 μmol) and
the EstDZ2 (0.04 μM) were added. The reaction was incubated at 50 °C.
Aliquots were withdrawn at predetermined time intervals and analyzed
using GC. After completion of the reaction, the products were isolated
by extracting the crude reaction mixture with EtOAc (3 × 1.5 mL). The
combined organic layers were dried over MgSO4 and evaporated to
dryness.
4. Experimental section
4.1. General
NMR spectra were recorded on Bruker Avance series 500 and 300
spectrometers at room temperature. Chemical shifts (δ) are reported in
ppm relative to the residual solvent peak (CDCl3, δ: 7.26, 13CDCl3, δ:
77.16) and the multiplicity of each signal is designated by the following
abbreviations: s, singlet; d, doublet; t, triplet; q, quartet; m, multiplet;
br, broad. Coupling constants (J) are quoted in Hz. Column chroma-
tographic separations were carried out using a flash chromatography
system with silica gel and hexane/ethyl acetate or petroleum ether/
ethyl acetate solvent mixtures. Thin Layer Chromatography (TLC) was
conducted using Merck silica gel (grade 60, F245). The progress of the
enzymatic reactions was followed by gas chromatography using a
SHIMADZU GC-2014 gas chromatograph equipped with an FID detector
and non-polar column (HP-5 capillary, 30 m × 0.32 mm × 0.25 μm,
5% diphenyl and 95% dimethylpolysiloxane - Nonpolar). The progress
of the EstDZ2-catalyzed hydrolysis of esters of secondary alcohols was
4.4. General procedure for enzymatic hydrolysis of esters of secondary
alcohols
In a vial of 22 mL, 4 mL of buffer solution Tris-HCl pH 8.0, 0.05%
Triton-X 100 (v/v) were added. The reaction was incubated at 50 °C
followed by the addition of 0.4 μM of EstDZ2 and 20 μmol of ester. After
completion of the reaction, the products were isolated by extracting the
crude reaction mixture with EtOAc (3 × 1.5 mL). The combined or-
ganic layers were dried over MgSO4 and evaporated to dryness.
4.5. Steglich esterification
followed by using
a chiral column (J&W CP-Chirasil-Dex CB
In a 50 mL round flask equipped with magnetic stirring, butyric acid
(1 mmol), DCC (1.1 mmol) and DMAP (5%) were added with 10 mL of
(25 × 0.25 × 0.25 μm) with total lengh 25 m).
Fig. 3. Synthesis of acyl esters of secondary alcohols. The isolated yield is given in bracket.
4

K. Myrtollari, et al.
BioorganicChemistry104(2020)104214
Fig. 4. Docking of (R)-1-phenylethyl acetate (green a) and (S)-1-phenylethyl acetate (green b) in the acetyl-enzyme intermediate of EstDZ2 active site. The catalytic
residues are indicated in red. The distance between the oxygen of the hydroxyl group of the catalytic Ser141 and the carbonyl carbon of (R)- and (S)-phenyl ethyl
acetate is 3.349 Å and 3.414 Å respectively. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this
article.)
6.88 (d, J = 9.12 Hz, 2H), 3.79 (s, 3H), 2.52 (t, J = 7.32 Hz, 2H),
1.84–1.71 (m, 2H), 1.03 (t, J = 7.41 Hz, 3H); 13C NMR (CDCl3,
500 MHz): δ 172.7, 157.3, 144.4, 122.5, 114.6, 55.7, 36.3, 18.6, 13.8.
Table 2
Kinetic resolution of esters of secondary alcohols using EstDZ2.
Entry
Substrate
Conversiona % (24 h)
Enantiomeric excessb
%
Ec
11
17
54
3.7
4.5.3. o-nitrophenyl butyrate (3)
1
Yield: 69%. H NMR (CDCl3 500 MHz): δ 8.09 (dd, J1 = 8.25 Hz,
J2 = 1.6 Hz, 1H), 7.65 (ddd, J1 = 15.6 Hz, J2 = 8.1 Hz, J3 = 1.6 Hz,
1H), 7.39 (ddd, J1 = 15.75 Hz, J2 = 8.2 Hz, J3 = 1.35 Hz, 1H), 7.23
(dd, J1 = 8.1 Hz, J2 = 1.25 Hz, 1H), 2.63 (t, J = 7.4 Hz, =2H),
1.85–1.77 (m, 2H), 1.06 (t, J = 7.4 Hz, 3H); 13C NMR (CDCl3,
500 MHz): δ 171.2, 144.2, 141.9, 134.6, 126.5, 125.7, 125.2, 35.8,
18.0, 13.6.
12
13
25
10
10
54
1.3
3.4
14
12
72
6.8
4.5.4. m-nitrophenyl Butyrate (4)
Yield: 64%. 1H NMR (CDCl3 300 MHz): δ 8.11 (ddd, J1 = 7.75 Hz,
J2 = 2.17 Hz, , J3 = 1.00 Hz, 1H), 7.99 (t (dd), J1 = 2.20 Hz, 1H), 7.56
(dd, J1 = 8.20 Hz, 1H), 7.45 (ddd, J1 = 8.15 Hz, J2 = 2.25 Hz,
J3 = 1.00 Hz, 1H), 2.59 (t, J = 7.35 Hz, =2H), 1.84–1.77 (m, 2H),
1.06 (t, J = 7.35 Hz, 3H); 13C NMR (CDCl3, 300 MHz): δ 171.6, 151.2,
148.9, 130.1, 128.3, 120.8, 117.5, 36.2, 18.4, 13.7.
15
16
20
74
8.1
–
> 99
< 1
4.5.5. p-nitrophenyl butyrate (5)
a,
b
Conversion and enantiomeric excess were derived from Gas
Yield: 70%. 1H NMR (CDCl3 300 MHz): δ 8.27 (d, J = 9.21 Hz, 2H),
7.28 (d, J = 9.27 Hz, 2H), 2.59 (t, J = 7.35 Hz, 2H), 1.84–1.76 (m,
2H), 1.05 (t, J = 7.38 Hz, 3H); 13C NMR (CDCl3, 300 MHz): δ 171.3,
155.6, 145.5, 125.3, 122.6, 36.3, 18.4, 13.7.
Chromatography by using a chiral capillary column.
c
E value was calculated as defined by Chen et al. [20].
dry CH2Cl2 at 0 °C. The mixture was kept under vigorous stirring for
30 min. Then, 3 mmol of alcohol dissolved in 4 mL of dry CH2Cl2 was
added dropwise and the reaction was stirred for 6 h. After completion of
the reaction the produced urea was filtered and the filtrate was eva-
porated. The product was purified by column chromatography with
pet.ether/EtOAc (20:1) as eluent. Isolated yield of the pure corre-
sponding ester ~ 66%.
4.5.6. p-propylphenyl butyrate (6)
Yield: 64%. 1H NMR (CDCl3 300 MHz): δ 7.16 (d, J = 8.45 Hz, 2H),
6.97 (d, J = 8.45 Hz, 2H), 2.57 (t, J = 7.50 Hz, 2H), 2.52 (t,
J = 7.35 Hz, 2H), 1.82–1.74 (m, 2H), 1.66–1.59 (m, 2H), 1.04 (t,
J = 7.40 Hz, 3H), 0.93 (t, J = 7.35 Hz, 3H); 13C NMR (CDCl3,
300 MHz): δ 172.5, 148.8, 140.3, 129.4, 121.3, 37.6, 36.4, 24.7, 18.6,
13.9, 13.8.
4.5.1. Phenyl butyrate (1)
Yield: 60%. 1H NMR (CDCl3 500 MHz): δ 7.38 (t (dd), J = 8.4 Hz,
2H), 7.22 (t, J = 7.5 Hz, 1H), 7.08 (d, J = 7.8 Hz, 2H), 2.55 (t,
J = 7.4 Hz, 2H), 1.84–1.76 (m, 2H), 1.05 (t, J = 7.4 Hz, 3H); 13C NMR
(500 MHz, CDCl3): δ 172.3, 150.9, 129.5, 125.9, 121.7, 36.4, 18.6,
13.8.
4.5.7. p-fluorophenyl butyrate (7)
Yield: 69%. 1H NMR (CDCl3 300 MHz): δ 7.07–7.04 (m, 4H), 2.53 (t,
J = 7.35 Hz, 2H), 1.82–1.74 (m, 2H), 1.04 (t, J = 7.40 Hz, 3H); 13C
NMR (CDCl3, 500 MHz): δ 172.3, 161.3, 159.3, 146.70, 146.68,
123.13, 123.06, 116.2, 116.1, 36.3, 18.6, 13.8.
4.5.2. p-methoxyphenyl butyrate (2)
4.5.8. o-bromophenyl butyrate (8)
1
Yield: 65%. 1H NMR (CDCl3 300 MHz): δ 7.00 (d, J = 9.12 Hz, 2H),
Yield: 68%. H NMR (CDCl3 300 MHz): δ 7.60 (dd, J1 = 8.31 Hz,
5

K. Myrtollari, et al.
BioorganicChemistry104(2020)104214
J2 = 1.71 Hz, 1H), 7.35–7.30 (m, 1H), 7.20–7.09 (m, 2H), 2.60 (t,
J = 7.32 Hz, 2H), 1.89–1.77 (m, 2H), 1.07 (t, J = 7.38 Hz, 3H); 13C
NMR (CDCl3, 300 MHz): δ 171.4, 148.4, 133.5, 128.6, 127.4, 124.0,
116.4, 36.1, 18.5, 13.9.
6.88 (d, J = 8.65 Hz, 2H), 5.85 (q, J = 6.6 Hz, 1H), 3.80 (s, 3H), 2.05
(s, 3H), 1.52 (d, J = 6.6 Hz, 3H); 13C NMR (500 MHz, CDCl3): δ 170.3,
159.2, 133.7, 127.6, 113.8, 72.0, 55.2, 21.9, 21.4.
4.7. In silico analysis
4.5.9. m-bromophenyl Butyrate (9)
Yield: 63%. 1H NMR (CDCl3 300 MHz): δ 7.36 (ddd, J1 = 8.05 Hz,
J2 = 1.8 Hz, , J3 = 0.95 Hz, 1H), 7.28 (t , J1 = 2.05 Hz, 1H), 7.24 (t,
J1 = 8.15 Hz, 1H), 7.04 (ddd, J1 = 8.15 Hz, J2 = 2.20 Hz,
J3 = 0.95 Hz, 1H), 2.53 (t, J = 7.50 Hz, =2H), 1.82–1.74 (m, 2H),
1.04 (t, J = 7.35 Hz, 3H); 13C NMR (CDCl3, 500 MHz): δ 171.8, 151.4,
130.6, 129.0, 125.3, 122.5, 120.6, 36.3, 18.5, 13.8.
In silico analyses were performed using the YASARA 18.11.21 soft-
ware. All esters used as substrates were drawn in YASARA and refined
under the AMPER03 force field. The docking experiments were per-
formed in a water cell (pH 8.0, 50 °C). The protein (“receptor”) was
flexible during docking simulations. Four parameters were used to
identify productive clusters from analysis of the substrate–protein
complexes: 1) protein distortion, 2) stability of the hydrogen bond in-
teractions with the oxyanion hole residues, 3) localization of the hy-
droxyl group with regard to the region included between the catalytic
histidine and serine residues, and 4) a distance of less than 4 Å between
the catalytic serine and the substrate. All figures were prepared with
PyMOL 0.99.
4.5.10. p-bromophenyl butyrate (10)
Yield: 63%. 1H NMR (CDCl3 300 MHz): δ 7.48 (d, J = 8.79 Hz, 2H),
6.97 (d, J = 8.73 Hz, 2H), 2.50 (t, J = 7.35 Hz, 2H), 1.83–1.71 (m,
2H), 1.03 (t, J = 7.41 Hz, 3H), 13C NMR (CDCl3, 500 MHz): δ 171.9,
149.9, 132.6, 123.5, 118.9, 36.3, 18.5, 13.8.
Declaration of Competing Interest
4.6. Acetylation of secondary alcohols
In a 100 mL round flask equipped with magnetic stirring, the sec-
ondary alcohol (1 mmol) was added with 7 mL of EtOAc. Next, acetic
anhydride (2 mmol), K2CO3 (4 mmol) and catalytic amount of DMAP
(5%) were added and the mixture was stirred at room temperature for
the appropriate time (24 h, monitored by TLC). After completion, the
reactant was filtered and water was added to the filtrate. The mixture
was extracted with ethyl acetate (10 mL × 3) and the combined organic
layer was washed with saturated NaHCO3 (10 mL × 2), brine (10 mL),
driedover MgSO4 and concentrated to give the pure product. Isolated
yield ~ 87%.
The authors declare that they have no known competing financial
interests or personal relationships that could have appeared to influ-
ence the work reported in this paper.
Appendix A. Supplementary data
Supplementary data to this article can be found online at https://
doi.org/10.1016/j.bioorg.2020.104214.
References
4.6.1. 1-phenylethyl acetate (11)
[1] M. Levisson, J. Van der Oost, S.W.M. Kengen, Carboxylic ester hydrolases from
hyperthermophiles, Extremophiles 13 (2009) 567, https://doi.org/10.1007/
s00792-009-0260-4.
1
Yield: 89%. H NMR (CDCl3 500 MHz): δ 7.36–7.27 (m, 5H), 5.88
(q, J = 6.6 Hz, 1H), 2.07 (s, 3H), 1.52 (d, J = 6.6 Hz, 3H); 13C NMR
(500 MHz, CDCl3): δ 170.5, 141.8, 128.6, 128.0, 126.2, 72.5, 22.3,
21.5.
[2] M. Fischer, J. Pleiss, The Lipase Engineering Database: a navigation and analysis
tool for protein families, Nucleic Acids Res. 31 (1) (2003) 319–321, https://doi.org/
10.1093/nar/gkg015.
[3] R. Verger, ‘Interfacial activation’ of lipases: facts and artifacts, Trends Biotechn. 15
(1997) 32–38, https://doi.org/10.1016/S0167-7799(96)10064-0.
[4] A. Hjorth, F. Carrière, C. Cudrey, H. Woldike, E. Boel, D.M. Lawson, F. Ferrato,
C. Cambillau, G.G. Dodson, L. Thim, R. Verger, A structural domain (the lid) found
in pancreatic lipases is absent in the guinea pig (phospho) lipase, Biochemistry 32
(1993) 4702–4707, https://doi.org/10.1021/bi00069a003.
4.6.2. 1-phenylpropyl acetate (12)
Yield: 85%. 1H NMR (CDCl3 500 MHz): δ 7.36–7.27 (m, 5H), 5.66 (t,
J = 7.3 Hz, 1H), 2.08 (s, 3H), 1.99–1.88 (m, 1H), 1.86–1.77 (m, 1H),
0.88 (t, J = 7.4 Hz, 3H); 13C NMR (500 MHz, CDCl3): δ 170.6, 140.7,
128.5, 128.0, 126.7, 77.5, 29.4, 21.4, 10.1.
[5] H.W. Anthonsen, A. Baptista, F. Drabløs, P. Martel, S.B. Petersen, M. Sebastião,
L. Vaz, Lipases and esterases: a review of their sequences, structure and evolution,
Biotechnol. Annu. Rev. 1 (1995) 315–371, https://doi.org/10.1016/S1387-
2656(08)70056-5.
4.6.3. 1-(4-ethylphenyl)ethyl acetate (13)
[6] P.C. Kienesberger, M. Oberer, A. Lass, R. Zechner, Mammalian patatin domain
containing proteins: a family with diverse lipolytic activities involved in multiple
biological functions, J. Lipid Res. 50 (2008) 63–68, https://doi.org/10.1194/jlr.
R800082-JLR200.
Yield: 82%. 1H NMR (CDCl3 500 MHz): δ 7.28 (d, J = 8.05 Hz, 2H),
7.18 (d, J = 8.25 Hz, 2H), 5.87 (q, J = 6.6 Hz, 1H), 2.64 (q,
J = 7.35 Hz, 2H), 2.06 (s, 3H), 1.53 (d, J = 6.6 Hz, 3H), 1.23 (t,
J = 7.6 Hz, 3H); 13C NMR (300 MHz, CDCl3): δ 170.6, 144.1, 139.0,
128.1, 126.3, 72.4, 28.7, 22.2, 21.6, 15.7.
[7] M.S. Jensen, S.K. Jensen, K. Jakobsen, Development of digestive enzymes in pigs
with emphasis on lipolytic activity in the stomach and pancreas, J. Anim. Sci. 75 (2)
(1997) 437, https://doi.org/10.2527/1997.752437x.
[8] a) S. Sharma, S.S. Kanwar, Organic solvent tolerant lipases and applications, Sci.
World J. 625258 (2014), https://doi.org/10.1155/2014/625258;
b) K.R. Jegannathan, P.H. Nielsen, Environmental assessment of enzyme use in
industrial production – a literature review, J. Cleaner Prod. 42 (2013) 228–240,
https://doi.org/10.1016/j.jclepro.2012.11.005.
4.6.4. 1-(4-isopropylphenyl)ethyl acetate (14)
Yield: 89%. 1H NMR (CDCl3 500 MHz): δ 7.28 (d, J = 8.15 Hz, 2H),
7.21 (d, J = 8.10 Hz, 2H), 5.87 (q, J = 6.6 Hz, 1H), 2.94–2.86 (m, 1H),
[9] A.M. Klibanov, Improving enzymes by using them in organic solvents, Nature 409
(2001) 241–246, https://doi.org/10.1038/35051719.
2.06 (s, 3H), 1.53 (d, J = 6.6 Hz, 3H), 1.24 (d, J = 6.95 Hz, 6H); 13
C
NMR (500 MHz, CDCl3): δ 170.4, 148.5, 138.9, 126.5, 126.1, 72.2,
[10] N. Gurung, S. Ray, S. Bose, V. Rai, A broader view: microbial enzymes and their
relevance in industries, medicine, and beyond, BioMed Res. Int. 2013 (2013) 1–18,
https://doi.org/10.1155/2013/329121.
33.8, 23.9, 22.0, 21.4.
[11] J.L. Adrio, A.L. Demain, Microbial enzymes: tools for biotechnological processes,
Biomolecules 4 (2014) 117–139, https://doi.org/10.3390/biom4010117.
[12] P. Hugenholtz, G.W. Tyson, Metagenomics, Nature 455 (7212) (2008) 481–483,
https://doi.org/10.1038/455481a.
4.6.5. 1-(4-fluorophenyl)ethyl acetate (15)
Yield: 91%. 1H NMR (CDCl3 500 MHz): δ 7.3.4–7.31 (m, 2H),
7.05–7.01 (m, 2H), 5.85 (q, J = 6.6 Hz, 1H), 2.06 (s, 3H), 1.52 (d,
J = 6.6 Hz, 3H) ; 13C NMR (500 MHz, CDCl3): δ 170.2, 163.3, 161.3,
137.5, 137.4, 127.9, 127.8, 115.4, 115.2, 71.6, 22.1, 21.3.
[13] P. Menzel, R. Gudbergsd, G. Rike, L. Lin, Q. Zhang, P. Contursi, M. Moracci,
J.K. Kristjansson, B. Bolduc, S. Gavrilov, N. Ravin, A. Mardanov, E. Bonch-
Osmolovskaya, M. Young, A. Krogh, X. Peng, Comparative metagenomics of eight
geographically remote terrestrial hot springs, Microb. Ecol. 70 (2) (2015) 411–424,
https://doi.org/10.1007/s00248-015-0576-9.
4.6.6. 1-(4-methoxyphenyl)ethyl acetate (16)
[14] R. Wohlgemuth, J. Littlechild, D. Monti, K. Schnorr, B. Siebers, P. Menzel, T. van
Rossum, I. Kublanov, A.G. Rike, G. Skretas, Z. Szabo, X. Peng, M.J. Young,
Yield: 85%. 1H NMR (CDCl3 500 MHz): δ 7.29 (d, J = 8.60 Hz, 2H),
6

K. Myrtollari, et al.
BioorganicChemistry104(2020)104214
Discovering of novel hydrolases from hot environments, Biotechnol. Adv. 36 (8)
00072-6.
(2018) 2077–2100, https://doi.org/10.1016/j.biotechadv.2018.09.004.
[15] D. Zarafeta, D. Moschidi, E. Ladoukakis, S. Gavrilov, E.D. Chrysina,
A. Chatziioannou, I. Kublanov, G. Skretas, F.N. Kolisis, Metagenomic mining for
thermostable esterolytic enzymes uncovers a new family of bacterial esterases, Sci.
Rep. 6 (1) (2016), https://doi.org/10.1038/srep38886.
[18] J. Pliego, J. Mateos, J. Rodriguez, F. Valero, M. Baeza, R. Femat, E. Herrera- López,
Monitoring lipase/esterase activity by stopped flow in a sequential injection ana-
lysis system using p-nitrophenyl butyrate, Sensors 15 (2) (2015) 2798–2811,
https://doi.org/10.3390/s150202798.
[19] W. Steglich, B. Neises, Simple method for the esterification of carboxylic acids,
Angew. Chem. Int. Ed. Engl. 7 (1978) 17, https://doi.org/10.1002/anie.
197805221.
[16] L.E. Janes, A.C. Löwendahl, R.J. Kazlauskas, Quantitative screening of hydrolase
libraries using ph indicators: identifying active and enantioselective hydrolases,
Chem. Eur. J. 4 (11) (1998) 2324–2331, https://doi.org/10.1002/(SICI)1521-
3765(19981102)4:11<2324::AID-CHEM2324>3.0.CO;2-I.
[20] C.S. Chen, Y. Fujimoto, G. Girdaukas, C.J. Sih, Quantitative analyses of biochemical
kinetic resolutions of enantiomers, J. Am. Chem. Soc. 104 (1982) 7294–7299,
https://doi.org/10.1021/ja00389a064.
[17] A.M.F. Liu, N.A. Somers, R.J. Kazlauskas, T.S. Brush, F. Zocher, M.M. Enzelberger,
U.T. Bornscheuer, G.P. Horsman, A. Mezzeti, C. Schmidt-Dannert, R.D. Schmid,
Mapping the substrate selectivity of new hydrolases using colorimetric screening:
lipases from Bacillus thermocatenulatus and Ophiostoma piliferum, esterases from
Pseudomonas fluorescens and Streptomyces diastatochromogenes, Tetrahedron:
Asymmetry 12 (4) (2001) 545–556, https://doi.org/10.1016/S0957-4166(01)
[21] R.J. Kazlauskas, A.N.E. Weissfloch, A.T. Rappaport, L.A. Cuccia, 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. 56 (8) (1991) 2656–2665, https://doi.org/10.1021/
jo00008a016.
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