Lyophilized extracts from vegetable flours as valuable alternatives to purified oxygenases for the synthesis of oxylipins

Article abstract of DOI:10.1016/j.bioorg.2019.103325

In this work, the whole aqueous extracts of soybean flour and oat flour have been used as valuable alternatives to purified oxygenase enzymes for the preparation of oxylipins derived from (5Z,8Z,11Z,14Z,17Z)-eicosapentaenoic acid (EPA). The lipoxygenase activity in the aqueous extracts of soybean (Glycine max. L.) flour was monitored with linoleic acid as substrate and compared with the commercially available purified enzyme (LOX-1). Oat flour extracts (Avena sativa L.) were evaluated for their peroxygenase activity by comparing different enzyme preparations in the epoxidation of methyl oleate. It was found that lyophilization of the aqueous extracts from these vegetable flours offers advantages in terms of enzyme stability, reproducibility and applicability to preparative organic synthesis. The lyophilized enzyme preparations were tested for the oxyfunctionalization of EPA and the formed products were isolated in satisfactory yields. In the presence of lyophilized extract from soybean, EPA gave 15S-hydroxy-(5Z,8Z,11Z,13E,17Z)-eicosapentaenoic acid in enantiopure form as exclusive product. Peroxygenase from oat flour was less selective and catalyzed the formation of different epoxides of EPA. However, the biocatalyzed epoxidation of EPA under controlled conditions allowed to obtain optically active (17R,18S)-epoxy-(5Z,8Z,11Z,14Z)-eicosatetraenoic acid (65% ee) as the main monoepoxide, among the five possible ones.

Full text of DOI:10.1016/j.bioorg.2019.103325

BioorganicChemistry93(2019)103325
Contents lists available at ScienceDirect
Bioorganic Chemistry
journal homepage: www.elsevier.com/locate/bioorg
Lyophilized extracts from vegetable flours as valuable alternatives to
purified oxygenases for the synthesis of oxylipins
Claudia Sanfilippo, Angela Paterna, Daniela M. Biondi, Angela Patti⁎
CNR – Istituto di Chimica Biomolecolare, Via Paolo Gaifami 18, I-95126 Catania, Italy
A R T I C L E I N F O
A B S T R A C T
Keywords:
In this work, the whole aqueous extracts of soybean flour and oat flour have been used as valuable alternatives to
purified oxygenase enzymes for the preparation of oxylipins derived from (5Z,8Z,11Z,14Z,17Z)-eicosapentae-
noic acid (EPA). The lipoxygenase activity in the aqueous extracts of soybean (Glycine max. L.) flour was
monitored with linoleic acid as substrate and compared with the commercially available purified enzyme (LOX-
1). Oat flour extracts (Avena sativa L.) were evaluated for their peroxygenase activity by comparing different
enzyme preparations in the epoxidation of methyl oleate. It was found that lyophilization of the aqueous extracts
from these vegetable flours offers advantages in terms of enzyme stability, reproducibility and applicability to
preparative organic synthesis. The lyophilized enzyme preparations were tested for the oxyfunctionalization of
EPA and the formed products were isolated in satisfactory yields. In the presence of lyophilized extract from
soybean, EPA gave 15S-hydroxy-(5Z,8Z,11Z,13E,17Z)-eicosapentaenoic acid in enantiopure form as exclusive
product. Peroxygenase from oat flour was less selective and catalyzed the formation of different epoxides of EPA.
However, the biocatalyzed epoxidation of EPA under controlled conditions allowed to obtain optically active
(17R,18S)-epoxy-(5Z,8Z,11Z,14Z)-eicosatetraenoic acid (65% ee) as the main monoepoxide, among the five
possible ones.
Oxylipins
Lipoxygenase
Peroxygenase
Lyophilization
Vegetable flours
EPA derivatives
Stereoselective reactions
1. Introduction
whose subsequent conversion through different enzymatic pathways
results in a variety of oxygenated derivatives.
Oxylipins are a large group of oxygenated metabolites of poly-
unsaturated fatty acids (PUFAs), that include fatty acid hydroperoxides,
hydroxy-, epoxy- and keto-derivatives, divinyl ethers, volatile alde-
hydes, and the plant hormone jasmonic acid [1,2]. These molecules,
which are ubiquitously present in eukariots, display important pro-in-
flammatory or anti-inflammatory effects in animals [3,4], whereas in
plants they are involved in a variety of signalling functions as well as in
the resistance to microbial pathogens [5,6].
Due to the increasing interest in the biological effects of ω-3 fatty
acids and related compounds there is the need to synthesize oxylipins in
preparative scale. Considerable synthetic efforts have been devoted to
the development of selective processes for the production of single
isomers of hydroxy- or epoxy- fatty acids, but in most cases multistep
sequences are necessary to control the functionalization of a specific
double bond of the polyenic molecules [12–14].
Two different approaches have been applied (Scheme 1), the first of
which is based on the stereoselective construction of the (Z,Z)-1,4-diene
system by Z-selective Wittig reactions or by CeC coupling of suitably
functionalized fragments followed by Lindlar reduction [15]. Strong
bases and metal catalysts are used and isomerization of the skipped Z-
olefins or over reduction often occur as side reactions. In the second
approach, PUFAs are used as individual starting materials for sub-
sequent modifications [16] and in many cases the selective epoxidation
of the double bond closest or far away to the carboxylic acid group has
been exploited [17] for the access to a variety of products through
oxidative cleavage of the epoxides, isomerization to allylic alcohol or
ring opening.
The in vivo biosynthesis of oxylipins involves a variety of enzymes,
belonging to the class of dioxygenases or monooxygenases, with spe-
cificity for the different positions of the polyene systems of PUFAs. In
this context, most of investigations have been focused on oxylipins
derived from linoleic acid, arachidonic acid and ω-3 fatty acids, eico-
sapentaenoic (EPA) and docosahexaenoic (DHA) acids, due to the im-
portance of their physiological effects as lipid mediators of inflamma-
tion with a potential role in metabolic diseases [7,8] and, more
recently, as cancer preventing agents [9–11].
A common starting step in the biosynthesis of oxylipins is the li-
poxygenase-catalyzed formation of a hydroperoxide of the fatty acid,
⁎ Corresponding author.
E-mail address: angela.patti@cnr.it (A. Patti).
https://doi.org/10.1016/j.bioorg.2019.103325
Received 30 August 2019; Received in revised form 25 September 2019; Accepted 26 September 2019
Availableonline27September2019
0045-2068/©2019ElsevierInc.Allrightsreserved.

C. Sanfilippo, et al.
BioorganicChemistry93(2019)103325
Scheme 1. General methods for the chemical synthesis of functionalized PUFAs.
Enzyme-catalyzed processes, which are usually performed in an
aqueous medium and under mild temperature conditions, offer a va-
luable alternative in terms of selectivity and sustainability, also for their
reduced demand of reagents, required for protection-deprotection of
substrates, and decreased purification steps. In the context of PUFA
modifications, oxygenases are the enzymes of choice but, unfortunately,
their commercial availability in purified form is rather limited and
currently too expensive for their application in synthesis. In many cases
these enzymes have been isolated in laboratory scale from vegetables
[18–20], algae [21,22], fungi [23,24] and bacteria [25,26] for their
characterization in terms of activity and positional specificity in the
formation of fatty acid derivatives.
In spite of their potential in organic synthesis, the reactions with
soybean flour and oat flour extracts have been carried out on analytical
scale and the products identified by MS-analysis, without any given
information about their isolated yields [37,38,43]. Only a process for
the preparative production of hydroperoxide of linoleic acid with soy-
bean flour extract was reported [37]. We then decided to investigate the
feasibility of preparative reactions in the presence of flour extracts of
soybean or oat using EPA as test substrate. The freeze-drying of flour
extracts allowed us to have stable and active enzyme preparations,
whose use in the synthesis of oxylipins derived from EPA, coupled with
the optimization of reaction and purification conditions, led to good
yields of isolated products and here we report the obtained results.
Lipoxygenase from soybean (Glycine max) is a non-heme iron-con-
taining enzyme [27] able to catalyse the addition of molecular oxygen
across the double bonds of polyunsaturated fatty acids forming their
corresponding hydroperoxides through a well established mechanism
[28]. Different isozymes are known for soybean lipoxygenase, with a
predominant specificity for the 13S-position of linoleic acid [29,30],
and reactions of the purified isozyme LOX-1 (EC 1.13.11.12) with a
variety of fatty acid substrates have been investigated [31–33], in some
cases optimizing the conditions to obtain the dioxygenation products
[34–36]. The use of soybean flour extracts has been proposed as eco-
nomical alternative to purified LOX-1 in the derivatization of linoleic
acid from vegetable oils [37], and more recently has been applied to
DHA [38], but in the latter case the products were not isolated and
diluted solutions of substrates were required in order to avoid inhibi-
tion of the enzyme.
2. Material and methods
2.1. General
(5Z,8Z,11Z,14Z,17Z)-Eicosapentenoic acid (EPA) was obtained by
lipase-catalyzed hydrolysis of the corresponding ethyl ester (from
Solutex, 90% purity). Lipoxygenase from Glycine max (soybean) Type I-
B (Lot. SLBS4722, 226,317 U/mg) and immobilized lipase from Candida
antarctica (Novozym 435) were obtained from Sigma Aldrich. Soybean
from organic crops was purchased by “Negozio leggero” shop network.
Whole oat seeds from organic crops were available from Ki Group.
Lyophilizations were carried out on a Telstar LyoQuest instrument.
Centrifugation was carried out on a Thermo Fisher Scientific SL 40R or
ALC PK130 centrifuge.
Among other legumes and cereals, oat (Avena sativa) has been taken
into consideration for its content in peroxygenase [40] and the whole
oat flour extracts, or the corresponding enzyme-enriched microsomal
fractions, have been used for the preparation of epoxides of some fatty
acids and esters as well as of simple alkenes [41–43], but have not up to
date tested on EPA or DHA. Peroxygenases (EC 1.11.2.1) are a class of
hydroperoxide-dependent enzymes containing heme that have recently
attracted interest for their ability in catalyzing oxyfunctionalization of
double bonds, with the advantage over cytochromes to be independent
by expensive nicotinamide cofactors [44,45].
UV-measurements were carried out on a UV–vis VALUE spectro-
photometer 8453 (Agilent Technologies). GC analyses were performed
on a fast GC Shimadzu 17-A instrument equipped with MS-EI detector
(GCMS-QP5050A) on
a
Supelco SPB-5 capillary column
(15 m × 0.1 mm ID × 0.1 µm film thickness). For the analyses the fol-
lowing parameters were set: He flow rate 32.7 mL/min, column inlet
pressure 350 KPa, linear velocity 31 cm/sec, split ratio 1:92, injector
temperature 250 °C, detector temperature 280 °C. The oven tempera-
ture was held at 100 °C for 1 min, then raised to 280 °C at 10 °C/min.
MS-EI detection was carried out setting 70 eV ionization voltage, 900 V
2

C. Sanfilippo, et al.
BioorganicChemistry93(2019)103325
electron multiplier voltage and 180 °C ion source temperature. Mass
spectra data were acquired in the scan mode in m/z range of 40–400.
Chiral HPLC analyses were carried out on a Dionex instrument
equipped with a Ultimate 3000 high-pressure binary pump, an ASI-100
autosampler, a TCC-100 thermostated column compartment and a
UVD-100 multiple wavelength detector set at 210, 220, 230 and
250 nm. The Chromeleon software (version 6.7) was used for instru-
ment control, data acquisition and data handling.
2.4. Biocatalyzed synthesis of 15S-hydroxy-(5Z,8Z,11Z,13E,17Z)-
eicosapentaenoic acid, 1
To a solution of EPA (100 mg, 0.33 mmol) in 50 mM borate buffer at
pH 9.0 (20 mL) soybean lyophilized enzyme preparation (200 mg) and
sodium borohydride (25 mg, 0.66 mmol) were added. The reaction
mixture was stirred at room temperature under a flow of air (100 mL/
min) until complete disappearance of the substrate (1 h) and then taken
to pH 3 by addition of dil. HCl. The resulting suspension was filtered
through a Celite pad and the clear filtrate was discarded. The Celite pad
was suspended in 60 mL of EtOAc and stirred for 15 min. After filtra-
tion, the organic phase was dried with Na2SO4 and the solvent evapo-
rated under vacuum to give an oily residue (130 mg) which was pur-
ified by column chromatography (Si gel 3.0 g, from n-hexane: HCOOH
99:1 to n-hexane:AcOEt:HCOOH 93:6:1, 150 mL) to give pure 1 (74 mg,
70% yield). [α]D = + 4.4 (c 0.88, MeOH), lit. [47] [α]D = + 4.9 (c
0.11, MeOH). 1H NMR: δ 0.97 (t, 3H, J = 7.6, H-20), 1.71 (br q, 2H, H-
3), 2.09 (m, 2H, H-19), 2.14 (br q, 2H, H-4), 2.38–2.35 (m, 4H, H-2 and
H-16), 2.82 (m, H-7), 2.98 (m, 2H, H-10), 4.28 (br q, 1H, H-15),
5.32–5.48 (m, 6H, H-5, H-6, H-8, H-9, H-11, H-17), 5.57 (m, 1H, H-18),
5.73 (dd, 1H, J = 5.6 and 15.2, H-14), 6.00 (t, 1H, J = 10.8, H-12),
6.60 (dd, 1H, J = 11.2 and 15.2, H-13). 13C NMR: δ 14.2 (C-20), 20.7
(C-19), 24.4 (C-3), 25.7 (C-7), 26.2 (C-10), 26.4 (C-4), 33.0 (C-2), 35.2
(C-16), 72.0 (C-15), 123.5 (CH = ), 125.4 (C-13), 127.4 (CH = ), 127.8
(C-12), 128.6 (CH = ), 128.8 (CH = ), 128.9 (CH = ), 130.3 (CH = ),
135.2 (C-14), 135.4 (C-18), 178.1 (C-1). HR-ESI-MS: 317.21230 [M –
TLC analyses were performed on aluminum plates coated with silica
gel and flourescent indicator F254, revealing the compounds by UV and
cerium sulfate.
Column chromatography was performed on silica gel 60 (Merck,
40–63 μm) or Lichroprep Si 60 (Merck, 25–40 μm) using the specified
eluents. 1H- and 13C NMR spectra were recorded on Bruker Avance™
400 spectrometer at 400.13 and 100.62 MHz, respectively. Chemical
shifts (δ) are given as ppm relative to the residual solvent peak and
coupling constants (J) are in Hz. 2D-experiments were carried out using
an inverse multinuclear probe with pulse-field Z-gradient and standard
Bruker pulse sequence programs. Optical rotations were measured on
Jasco DIP-135 polarimeter using a 10 cm length cell.
2.2. Preparation of EPA acid
To a solution of the ethyl ester of EPA (200 mg, 0.6 mmol) in 1.7 mL
of dioxane, 300 µL of distilled water and Novozym 435 (200 mg) were
added. The mixture was stirred at 55 °C and 280 rpm in a shaker for 3 h.
Then, the enzyme was separated by filtration and the solution taken to
dryness under reduced pressure. The residue was purified by column
chromatography (Si gel 3.9 g, from n-hexane to n-hexane/EtOAc 9:1
containing 0.1% of HCOOH, 200 mL) to give pure (TLC analysis) EPA
(180 mg, 0.59 mmol, 98% yield) as a colorless oil.
–
H]–; theor. for C20H29O3 317.21112
2.5. Synthesis of compound 3
According to
a reported procedure [48] compound 1 (30 mg,
0.094 mmol) was converted into the corresponding methyl ester by re-
action with dimethylcarbonate (0.5 mL) in the presence of Novozym 435
lipase (30 mg). The reaction mixture was stirred (280 rpm) in a shaker at
55 °C until complete conversion of the substrate was reached (5 h), then
quenched by filtration of the enzyme. The solution was taken to dryness
under vacuum and the residue was purified by column chromatography
(Si gel, from n-hexane to n-hexane:EtOAc to 9:1) to afford ester 2 (29 mg,
2.3. Test for lipoxygenase activity
Air-dried soybean seeds from organic crops were finely grounded
with a domestic blender and the obtained flour was sifted on a tea sieve.
Defatted flour was prepared by suspending the powder (5 g) in acetone
(10 mL) and stirring the mixture for 10 min at room temperature. The
suspension was then centrifuged at 2930 g and the solution discarded.
The procedure was then repeated another two times and the final solid
was dried overnight in a fume hood to give 4.4 g of defatted flour.
Defatted soybean flour (1 g) was suspended in 20 mL of distilled
water and stirred under magnetic agitation at room temperature for 5,
15 or 30 min. The suspension was then centrifuged at 11,000 g for
20 min at 4 °C and the obtained supernatant was used as “soybean ex-
tract” sample. The same procedure was also applied to untreated soy-
bean flour, which was extracted for 30 min to give the “not defatted”
sample.
0.09 mmol, 96% yield) as
a colorless oil. Compound 2 (10 mg,
0.031 mmol) was dissolved in CH2Cl2 (0.8 mL) and to this solution N,N'-
dicyclohexylcarbodiimmide (DCC, 0.3 mmol), dimethylaminopyridine
(DMAP, 0.15 mmol) and (R)-(–)-methoxyphenylacetic acid (MPA,
0.15 mmol) were added. The mixture was left at room temperature under
stirring for 18 h. After evaporation of the solvent, the residue was purified
by column chromatography (Si gel 1.5 g, from n-hexane to n-hex-
ane:EtOAc 9:1 v/v, 80 mL) to give 12 mg (0.025 mmol, 82% yield) of
diester 3 as a colorless oil. 1H NMR: δ 0.89 (t, 3H, J = 7.6, H-20), 1.72 (m,
2H, H-3), 1.92 (m, 2H, H-19) 2.12 (m, 2H, H-4), 2.25–2.35 (m, 4H, H-2
and H-16), 2.81 (br t, H-7), 2.92 (br t, 2H, H-10), 3.42 (s, 3H, -OMe), 3.67
(s, 3H, -COOMe), 4.76 (s, 1H, -CHOMe), 5.05 (m, 1H, H-17) 5.35–5.46
(m, 7H, H-5, H-6, H-8, H-9, H-11, H-15, H-18), 5.62 (dd, 1H, J = 7.2 and
15.2, H-14), 5.95 (t, 1H, J = 10.8, H-12), 6.52 (dd, 1H, J = 11.2 and
15.2, H-13), 7.35 (m, 3H, Ph), 7.44 (br d, 2H, Ph). 13C NMR: δ 14.0 (C-
20), 20.6 (C-19), 24.8 (C-3), 25.6 (C-7), 26.1 (C-10), 26.5 (C-4), 32.1 (C-
2), 33.4 (C-16), 51.5 (–COOCH3), 57.3 (–OCH3), 75.0 (C-15), 82.7
(-CHOMe), 122.4 (C-17), 127.2 (2 × Ar-C), 127.4 (CH = ), 127.7 (C-12),
128.0 (C-13), 128.5 (2 × Ar-C), 128.6 (CH = ), 128.8 (CH = ), 129.1
(CH = ), 130.5 (C-14), 131.5 (CH = ), 134.7 (C-14), 136.3 (C-1 Ar),
170.0 (MPA-CO), 174.0 (C-1).
Freeze-dried preparation of lipoxygenase was obtained as follows:
soybean flour (10 g) was suspended in 100 mL of distilled water and
stirred under magnetic agitation at room temperature for 30 min at
1250 rpm. The suspension was then centrifuged at 11,000 g for 20 min
at 4 °C and the supernatant frozen at – 4 °C. The frozen supernatant was
freeze-dried to give 2.5 g of lyophilized enzyme preparation as a pale
yellow powder. For the test of enzymatic activity 250 mg of powder
were dissolved in 20 mL of H2O.
The assay of lipoxygenase activity was carried out according to Suda
et al. [46] by mixing 200 µL of 10 mM sodium linoleate, 200 µL of
100 µM methylene blue, 1 mL of 50 mM borate buffer pH 9.0 and dis-
tilled water to a final volume of 2 mL. To this solution, kept under
magnetic agitation, LOX-1 solution (2 mg/mL, 70 μL), soybean fresh
extract (50 mg/mL, 20 µL) or solution of freeze-dried extract (100 μL,
12.5 mg/mL) was added and the absorbance at λ 660 nm was mon-
itored over 10 min.
2.6. Test for peroxygenase activity
Commercial whole seeds (60 g) of air-dried oat (Avena sativa) from
organic crops were ground by an electric coffee mill and the obtained
3

C. Sanfilippo, et al.
BioorganicChemistry93(2019)103325
flour was defatted by suspension in diethyl ether (150 mL) for 10 min
followed by centrifugation at 2930 g for 10 min. The surnatant was
discarded and the procedure repeated on the residue another two times.
The final residue was dried at room temperature overnight to give 56 g
of defatted flour.
The reaction mixture was stirred in a shaker at 55 °C until complete
conversion of the substrate was reached (2 h), then quenched by fil-
tration of the enzyme. The solution was taken to dryness under vacuum
and the residue was purified by column chromatography using
LiChroprep Si 60 (0.8 g) and eluting with n-hexane:EtOAc 95:5 (30 mL)
to afford ester 5 (19 mg, 0.042 mmol, 92% yield) as a colorless oil,
[α]D25 = +3.5 (c = 0.17, CH3OH); lit. [14] [α]D25 = +5.0 (c = 1.0,
CH3OH). 1H NMR: δ 1.05 (t, 3H, J = 7.6, H-20), 1.52–1.65 (m, 2H, H-
19), 1.72 (q, 2H, J = 7.2, H-3), 2.11 (q, 2H, J = 7.2, H-4), 2.20–2.27
(m, 1H, H-16a), 2.33 (t, 3H, J = 7.2, H-2), 2.38–2.43 (m, 1H, H-16b),
2.80–2.85 (m, 6H, H-7, H-10, H-13), 2.88–2.92 (m, 1H, H-18),
2.94–2.98 (m, 1H, H-17), 3.67 (–OCH3), 5.37–5.39 (m, 6H, H-5, H-6, H-
8, H-9, H-11, H-12), 5.47–5.51 (m, 2H, H-14 and H-15). 13C NMR: δ
10.6 (C-20), 21.0 (C-19), 24.8 (C-3), 25.6 (C-7, C-10), 25.8 (C-13), 26.2
(C-16), 26.5 (C-4), 33.4 (C-2), 51.4 (–OCH3), 56.5 (C-17), 58.3 (C-18),
124.5 (C14, C15), 127.8 (CH]), 128.0 (CH]), 128.3 (CH]), 128.4
(CH]), 128.8 (C-5), 130.4 (C14/C15), 174.0 (C-1). HR-ESI-MS:
355.22426 [M + Na]+; theor. for C21H32O3Na+ 355.22437.
For test reactions on flour, the defatted oat flour (2 g) was sus-
pended in 50 mM potassium phosphate buffer pH 7.5 (7 mL) and the
mixture used as a whole. For test reactions on suspensions, the defatted
oat flour (2 g) was suspended in the phosphate buffer, mixed under
magnetic stirrer for 5 min and centrifuged at 2930 g for 2 min. The
bottom fraction was discarded and the suspension was used for the
epoxidation reaction by adding the reagents.
Freeze-dried preparation of peroxygenase was obtained as follows.
Defatted oat flour (110 g) was suspended in water (200 mL) and mixed
under magnetic stirrer for 5 min. The mixture was centrifuged at 2930 g
for 2 min and the suspension was collected. The procedure was repeated
again on the bottom fraction of the centrifugation and the pooled sus-
pension fractions were freeze-dried to give 15.2 g of a white light
powder.
The enantiomeric excess (65% ee) was measured on a Lux Amylose-
1 column (Phenomenex) eluting with a 0.7 mL/min flow of n-hexane/2-
PrOH 99:1 at 23 °C; tR/min: 12.69 [minor, (17S,18R)-enantiomer] and
24.16 [major, (17R,18S)-enantiomer].
In the test for peroxygenase activity, defatted oat flour (2 g) or the
suitable enzyme preparation deriving from 2 g of flour was suspended
in 7 mL of 50 mM potassium phosphate buffer at pH 7.5. To this sus-
pension methyl oleate (13 μL, 11.4 mg, 38 μmol) and t-BuOOH (70 wt%
in H2O, 13 μL, 8.5 mg, 95 μmol) were added and the reaction mixture
was maintained under vigorous stirring at 25 °C. The reaction progress
was monitored by GC analysis of aliquots (0.4 mL) of the reaction taken
at regular intervals and extracted with MeOH:Et2O 1:9 v/v (0.4 mL);
3 μL of the dried organic solution were injected for the GC analyses.
3. Results and discussion
3.1. Lipoxygenase activity in soybean flour extracts
At the onset of our study, the lipoxygenase activity in soybean flour
extracts was tested on linoleic acid by using a simple colorimetric test
based on the bleaching of methylene blue. In the presence of lipox-
ygenase, indeed, linoleic acid is converted into the corresponding hy-
droperoxide which reduces methylene blue to its colourless leucoform
and the enzyme activity can be monitored by following the decrease
over time in the UV absorbance (λ 660 nm) of the reaction mixture
[46].
2.7. Biocatalyzed synthesis of (17R,18S)-epoxy-(5Z,8Z,11Z,14Z)-
eicosatetraenoic acid, 4
To a suspension of freeze-dried oat extract (550 mg) in 50 mM
phosphate buffer at pH 7.5 (28 mL) containing EPA (100 mg,
0.33 mmol), t-BuOOH (70 wt% in H2O) was added in three equal ali-
quots at time 0, 15 and 40 min (total volume 40 µL, 0.29 mmol). The
reaction mixture was maintained for 2 h at 25 °C under vigorous mag-
netic stirring and monitored by TLC analysis (Si gel, n-hexane: EtOAc
7:3 v/v containing 0.8% of HCOOH). The reaction was quenched by
addition of MeOH (0.5 mL) and the mixture was then extracted with
Et2O (3 × 15 mL). The collected organic layers were dried over Na2SO4
and taken to dryness under vacuum. The residue was purified by
column chromatography on LiChroprep Si 60 (6 g) by eluting with n-
hexane:EtOAc 9:1 v:v containing 0.8% of HCOOH (200 mL). Unreacted
EPA (20 mg, 0.066 mmol, 20% yield) was eluted as the first fraction
followed by compound 4 which was obtained as clear oil (46 mg,
0.14 mmol, 44% yield) with 65% ee (from analysis of the corresponding
methyl ester), [α]D25 = + 3.5 (c = 0.84, CH3OH); lit. [14]
[α]D25 = +7.0 (c = 1.0, CHCl3). 1H NMR: δ 1.06 (t, 3H, J = 7.6, H-20),
1.51–1.66 (m, 2H, H-19), 1.72 (q, 2H, J = 7.6, H-3), 2.14 (q, 2H,
J = 7.6, H-4), 2.19–2.26 (m, 1H, H-16a), 2.36 (t, 3H, J = 7.6, H-2),
2.41–2.48 (m, 1H, H-16b), 2.80–2.85 (m, 6H, H-7, H-10, H-13),
2.91–2.96 (m, 1H, H-18), 2.97–3.02 (m, 1H, H-17), 5.33–5.44 (m, 6H,
By this way, it was observed that simple stirring of defatted soybean
flour with water was sufficient to extract lipoxygenase in solution, with
an optimal extraction time of 30 min, and the activity of the extracted
enzyme (3000 U/mg of soybean flour) was determined by comparison
with that of commercial lipoxygenase LOX-1 (Fig. 1).
The preliminary defatting of soybean flour did not affect the enzy-
matic activity, but a significant loss of activity in the extract (as well as
in the solution of LOX-1) was detected as early as 2 h from extraction
Extraction time 5min
Extraction time 15min
Extraction time 30min
LOX-1
1.4
1.2
1.0
0.8
0.6
0.4
0.2
0.0
Not defatted flour
H-5, H-6, H-8, H-9, H-11, H-12), 5.44–5.57 (m, 2H, H-14 and H-15). 13
C
NMR: δ 10.6 (C-20), 21.0 (C-19), 24.5 (C-3), 25.6 (C-7, C-10), 25.8 (C-
13), 26.1 (C-16), 26.4 (C-4), 33.2 (C-2), 56.7 (C-17), 58.6 (C-18), 124.3
(C-15), 127.8 (CH]), 128.0 (CH]), 128.2 (CH]), 128.4 (CH]), 128.8
(C-5), 128.9 (CH]), 130.5 (C-14), 178.6 (C-1). HR-ESI-MS: 317.21231
[M – H]–; theor. for C20H29O3 317.21112. Fractions containing mix-
–
tures of monoepoxides (15.5 mg, 0.049 mmol) and isomeric diepoxides
(11 mg, 0.033 mmol) accounted for 15% and 10% yield, respectively.
0
100
200
300
400
500
600
Time (sec)
2.8. Stereochemical analysis of compound 4
Fig. 1. Lipoxygenase activity in water extracts of defatted soybean flour in
comparison with commercial LOX-1 and extract from non defatted soybean
flour.
Compound
4
(20 mg, 0.062 mmol) was dissolved in di-
methylcarbonate (0.5 mL) and Novozym 435 lipase (20 mg) was added.
4

C. Sanfilippo, et al.
BioorganicChemistry93(2019)103325
1.2
1
1.2
1
(A)
(B)
0.8
0.6
0.4
0.2
0
0.8
0.6
0.4
0.2
0
0
100
200
300
400
500
600
0
100
200
300
400
500
600
Time (sec)
Time (sec)
LOX-1
Fresh extract
Freeze-dried extract
1h
2h
4h
Freeze-dried extract (after 6 months)
Freezed extract (after 24h)
Fig. 2. (A) Lipoxygenase activity in soybean flour extracts at different times after extraction; (B) lipoxygenase activity in freeze-dried extracts (100 μL) compared
with fresh extract (20 μL) and purified enzyme. (Also see the experimental section).
(Fig. 2A), so that the enzymatic conversion of less reactive substrates
could pose some concerns. The freezing of the extract was not sufficient
to maintain the enzymatic activity so that the extract should be freshly
prepared for its use in biocatalyzed reactions.
In this context, we envisaged that the freeze-drying of soybean flour
extract could be advantageous in giving the availability of active en-
zyme as if it was freshly extracted and the possibility to portionwise add
the enzyme in case of slow-reacting substrates without volume increase.
Furthermore, the reproducibility between different reactions could be
ensured by using the same lot of lyophilized enzyme preparation.
Thus, soybean flour was stirred in water for 30 min and the solid
separated by centrifugation; the solution was then freeze-dried and
Scheme 2. Biocatalyzed synthesis of 15-OH EPA with soybean flour extract.
about 20% of the activity of the fresh extract was recovered in the
obtained enzyme preparation, probably due to the aforementioned
time-dependent deactivation, inherent in the enzyme, during the pre-
paration and lyophilization of the sample. However, it is noteworthy
that the recovered lipoxygenase activity was still present after 6 months
of storage at –20 °C and the ultimate reaction rate was the same as the
freshly freeze-dried enzyme preparation (Fig. 2B).
context, the simultaneous presence of a water-soluble triphenylpho-
sphine as reducing agent during the enzymatic oxidation has been
proposed to overcome the problem [33].
Among the different reducing agents we have tested, NaBH4 in a
2:1 M ratio with respect to the substrate was the best and allowed the
concentration of EPA to be increased up to 24 mM maintaining a full
conversion within 1 h. When a preparative reaction was carried out we
faced with extensive loss in the recovered product, due to the formation
of a gelatinous emulsion during the extraction of the acidified reaction
mixture with an organic solvent so that the reported work-up protocol
was modified. In our optimized conditions, HCl was added to the re-
action mixture up to pH 3 and the resulting cloudy suspension was
filtered over a Celite pad in a sintered funnel; the clear filtrate was then
discarded and the Celite washed with EtOAc to give an organic fraction
containing the alcohol 1. In the 1H NMR spectra of 1, in addition to the
diagnostic broad quartet of the –CHOH proton at 4.3 ppm, three well-
resolved resonances in the region of ethylenic protons were observed
and associated with the two conjugated double bonds, in which two
hydrogens are in a trans configuration as deduced from their large
coupling constant (J = 15.2 Hz). The presence of just two methylene
groups located between two double bonds, recognizable for their pe-
culiar chemical shift around 2.8–3.0 ppm is compatible with the func-
tionalization of C8-C9 or C14-C15 double bond and the discrimination
between the two possibilities was obtained by 2D experiments. Indeed,
3JHC correlation of the C20 triplet with an ethylenic carbon in the 2D-
HMBC spectrum allowed to unequivocally assign the C-18 carbon re-
sonance, from which the H-16 proton was identified and found scalarly
coupled (2D-COSY) with the eCHOH proton.
3.2. Biocatalyzed hydroperoxidation of EPA
Although the hydroperoxidation of fatty acids by soybean lipox-
ygenase has been widely investigated, an optimal experimental proce-
dure is not still established and a variety of conditions have been
adopted in terms of temperature, pressure of oxygen, substrate con-
centration and amount of enzyme.
In most cases the enzyme displays optimal activity and selectivity at
pH 9.0 [31] giving an hydroperoxide which is subsequently chemically
reduced to the corresponding alcohol.
In our experiments, we chose to dissolve the lyophilized preparation
of lipoxygenase in 50 mM borate buffer at pH 9.0, and magnetically stir
the reaction mixture at room temperature under a vigorous bubbling of
air. In a first reaction, full conversion of 1.65 mM EPA substrate in the
presence of 1.8 mg/ml of soybean freeze-dried extract was reached
within 1 h and the subsequent chemical reduction of the crude reaction
mixture with NaBH4 gave the expected monohydroxy derivative 1 as
almost exclusive product (Scheme 2).
Increasing the substrate concentration led to unreacted EPA even
after 2 h and complete conversion of the substrate was achieved only
after the addition of fresh enzyme. This behaviour has been reported for
other PUFAs and attributed to inhibition/deactivation of lipoxygenase
by substrate and/or product or other reactive oxygen species. In this
5

C. Sanfilippo, et al.
BioorganicChemistry93(2019)103325
The structure of
1
was then established as 15-hydroxy-
preparation converted 0.32 μmol of methyl oleate/mg of lyophilized
(5Z,8Z,11Z,13E,17Z)-eicosapentaenoic acid and the S-configuration of
the molecule was assigned by comparison of its optical rotation with
literature data [47], so confirming the stereoselectivity of soybean li-
poxygenase. Further support came from the 1H NMR spectrum of the
diester 3, obtained by reaction of the ethyl ester 2 with (R)-α-meth-
oxyphenylacetic acid (MPA), that was in agreement with the (R)-MPA-
15S-diastereoisomer [46]. The resonances reported for the (R)-MPA-
15R-diastereoisomer,1 were not visible in the spectrum of 3 (see sup-
porting information) thus the enantiomerically pure nature of 1 was
established. For alcohol 1, which has been recently isolated from some
microalgae [47] and marine diatoms [49,50], interesting antibacterial
[51] and anti-inflammatory [47] activities have been reported.
enzyme preparation in 1 h.
3.4. Biocatalyzed epoxidation of EPA
The epoxidation of PUFAs by peroxygenase from oat has been
previously applied only to linoleic and linolenic acids (or their methyl
esters), that were converted in mixtures of the corresponding mono-
epoxides and diepoxides [42]. Stereoselective epoxidation of PUFAs
with higher degree of unsaturation by different human cytochromes has
been reported [54–56], but these studies are mostly directed to un-
derstanding the in vivo metabolism of PUFAs and other oxygenases have
not been investigated up to date.
The lack of data on a biocatalytic route to EPA-derived epoxides in
preparative scale prompted us to test our lyophilized extract from oat
flour as catalyst for the epoxidation reaction. The enzyme preparation
was then suspended in phosphate buffer (pH 7.5) and EPA was left to
react in the presence of t-BuOOH as oxidant (t-BuOOH:EPA 5:1 M ratio,
three equal aliquots) at room temperature. The substrate was quickly
converted into a mixture of mono- and poly- epoxide derivatives, so
evidencing that oat peroxygenase is not selective toward the mono-
functionalization of EPA and partially epoxidized derivatives of EPA are
also good substrates for the enzyme. However, the time-course mon-
itoring of the enzymatic reaction, in comparison with the chemical
epoxidation of EPA in the presence of m-chloroperbenzoic acid, re-
vealed that one monoepoxide among the five possible ones was mainly
formed as transient product in the peroxygenase-catalyzed epoxidation.
Thus, we focused our attention on the possibility to maximize the for-
mation of a single monoepoxide derivative of EPA and a biocatalyzed
reaction was carried out in the presence of 0.9 eqv. of t-BuOOH. In these
conditions, 80% conversion of the substrate was reached in 2 h and
epoxide 4 was obtained in 44% isolated yield and its structure was
established by NMR-analysis (Scheme 3).
3.3. Peroxygenase activity in oat flour extracts
In oat seeds the peroxygenase is localized in the microsomial
fraction, which can be separated from the other cellular components
by ultracentrifugation to give an enriched enzyme preparation
[39,41,42]. As an alternative, the whole oat seed flour has been di-
rectly used for some biocatalyzed transformations [43]. A procedure
for the immobilization of the enzyme activity on a protein-binding
synthetic membrane has been also developed and the resultant pre-
paration was tested either in aqueous medium or organic solvents
[52,53].
Encouraged by the results obtained with the freeze-dried extract of
soybean flour, we were interested the possibility to apply the same
procedure also to oat flour extracts. Since the peroxygenase from oat is
not available in purified form, the different enzyme preparations were
compared for their activity in a standard reaction of methyl oleate in
the presence of t-butyl hydroperoxide (t-BuOOH) as oxidant, by mon-
itoring the reaction course by GC-MS analysis. When H2O2 was tested as
alternative oxidant, the enzymatic activity was fully inhibited and only
the unreacted substrate was detected.
In the 1H NMR spectrum, the signals in the region 5.3–5.6 ppm ac-
counted for four ethylenic bonds and the two methinic protons on the
oxirane ring were identified by their CeH correlation in the 2D-HSQC
spectrum with signals at 56.7 and 58.6 ppm. Diagnostic correlation be-
tween the oxirane carbon at 58.6 and methyl protons on C-20 was ob-
served in the 2D-HMBC spectrum, so that structure of 17,18-epoxy-
(5Z,8Z,11Z,14Z)-eicosatetraenoic acid was unequivocally assigned to
compound 4. Compound 4 was isolated in 65% enantiomeric excess, as
determined by chiral HPLC analysis of the corresponding methyl ester 5,
and the 17R,18S absolute configuration was assigned by comparison of
optical rotation of 5 with data reported in literature [14]. However, we
found an opposite elution order of the two enantiomers of 5 on the
Amylose-1 (Phenomenex) HPLC column with respect the reported data
for the same chiral stationary phase [14]. When our sample was analysed
on a Chiralcel OB column the 17R,18S-enantiomer was eluted as the first
peak, in agreement with previous studies [54], so confirming the assigned
configuration of 5 and giving evidence that the elution order of en-
antiomers is opposite on Chiralcel OB and Amylose-1 columns.
In the first reactions the substrate was added to the whole slurry
obtained by homogenization of the oat flour in phosphate buffer at pH
7.5 [43] and preliminary defatting of the flour was found to be essential
for the peroxygenase activity (Table 1, entries 1 and 2), which was
maintained also after 2 h from extraction (entry 3).
Centrifugation of the slurry at 2930 g allowed to discard a solid
fraction and recover the enzyme activity in the microsome-containing
surnatant, which appeared as a white milky suspension. When methyl
oleate was added to this suspension it was fully converted into the
corresponding epoxide in 1 h and the increase in the reaction rate could
be related with a better contact between the enzyme and reactants,
favoured by the lower density of reaction mixture compared with that
containing the whole homogenate (Table 1, compare entries 2 and 4).
Centrifugation of the slurry at 11,000 g gave a clear solution with
about halved activity with respect to the above cited suspension
(Table 1, compare entries 4 and 5). Homogenization of the oat flour
with water instead the phosphate buffer and subsequent centrifugation
gave a suspension that was freeze-dried to afford a white light powder
(5.5% weight with respect to flour) with slightly lower activity with
respect to the parent enzyme preparation (Table 1, compare entries 6
and 4). The activity of the freeze-dried suspension was monitored up to
6 months (storage at –20 °C) without giving significant differences, thus
ensuring reproducibility within the same batch of enzyme preparation.
Differences in the enzymatic activity could be expected depending
on the biological variability of the oat source, the efficiency of cen-
trifugation and the yield of lyophilized powder. Our enzyme
Although epoxidation reactions catalysed by oat peroxygenase have
been reported for a variety of ethylenic substrates, no information has
been given regarding the stereochemistry and the optical purity of the
obtained products. Thus, our data provide the first report of the ste-
reochemical selectivity of oat peroxygenase.
Positive effects in decreasing cell proliferation have been reported
for compound 4, that has been shown to be the only active EPA-derived
monoepoxide compared to the other regioisomers [57], and its ability
to regulate the contraction of cardiomyocytes [58] makes it attractive
as antiarrythmic drug.
1 For the (S)-MPA-15S-diastereoisomer (which is magnetically equivalent to
(R)-MPA-15R-diastereoisomer) the following resonances (selected resonances)
were reported: δ 0.95 (H-20), 2.76 (H-10), 5.25 (H-17), 5.51 (H-14), 5.85 (H-
12), 6.28 (H-13).
4. Conclusions
Simple and effective bioprocesses for the stereo- and regioselective
preparation of two oxygenated derivatives of EPA were developed by
6

C. Sanfilippo, et al.
BioorganicChemistry93(2019)103325
Table 1
Activity of different peroxygenase preparation from oat floura.
Entry
Enzyme preparation
% Epoxide (0.5 h)b
% Epoxide (1 h)b
1
2
3
4
5
6
7
8
Slurryc
32
43
44
80
42
74
71
72
50
82
75
99
65
92
88
88
Slurry
Slurryd
Microsomes suspension
Microsomes suspensione
Freeze-dried suspension
Freeze-dried suspensiond
Freeze-dried suspensionf
a
Experimental conditions: defatted oat flour (2 g) or enzyme preparation from 2 g of defatted flour; 50 mM phosphate buffer (7 mL), methyl oleate (0.013 mL,
0.038 mmol), 70% v/v of TBHP (0.013 mL, 0.1 mmol), 25 °C.
b
c
d
e
By GC analysis of the reaction mixture.
Flour was not defatted.
Reagents were added after 2 h of stirring in phosphate buffer.
From centrifugation at 11000 g.
f
After 6 months from freeze-drying
Lipids 1801 (2010) 1260–1273.
[4] C.N. Serhan, Pro-resolving lipid mediators are leads for resolution physiology,
Nature 510 (2014) 92–101.
[5] E. Blée, Impact of phyto-oxylipins in plant defense, Trends Plant Sci. 7 (2002)
315–322.
[6] H.C. Pohl, L.F.J. Kock, Oxidized fatty acids as inter-kingdom signaling molecules,
Molecules 19 (2014) 1273–1285.
[7] C.N. Serhan, J. Dalli, R.A. Colas, J.W. Winkler, N. Chiang, Protectins and maresins:
new pro-resolving families of mediators in acute inflammation and resolution
bioactive metabolome, Biochim. Biophys. Acta (BBA)-Mol. Cell Biol. Lipids. 1851
(2015) 397–413.
[8] M. Spite, J. Clària, C.N. Serhan, Resolvins, specialized proresolving lipid mediators,
and their potential roles in metabolic diseases, Cell Metab. 19 (2014) 21–36.
[9] N.B. Janakiram, A. Mohammed, C.V. Rao, Role of lipoxins, resolvins, and other
bioactive lipids in colon and pancreatic cancer, Cancer Metastasis Rev. 30 (2011)
507–523.
Scheme 3. Biocatalyzed synthesis of 17,18-epoxy-EPA with oat flour extract.
exploiting the lipoxygenase and peroxygenase content of soybean flour
and oat flour, respectively. The enzymatic activity was recovered from
these flours in the aqueous extracts, which could be used as such in
place of the purified enzymes. The freeze-drying of the aqueous extracts
gave enzyme preparations stable up to 6 months and allowed re-
producibility of the reactions and simplification of the workup. Further
studies are in progress to extend the procedure to polyunsaturated fatty
acids other than EPA, also targeting at the preparation of specific de-
rivatives with relevant biological activities.
[10] M. Murray, A. Hraiki, M. Bebawy, C. Pazderka, T. Rawling, Anti-tumor activities of
lipids and lipid analogues and their development as potential anticancer drugs,
Pharmacol. Ther. 150 (2015) 109–128.
[11] K. Moro, M. Nagahashi, R. Ramanathan, K. Takabe, T. Wakai, Resolvins and omega
three polyunsaturated fatty acids: Clinical implications in inflammatory diseases
and cancer, World J. Clin. Cases 4 (2016) 155–164.
[12] L. Balas, T. Durand, Dihydroxylated E, E, Z-docosatrienes. An overview of their
synthesis and biological significance, Prog. Lipid Res. 61 (2016) 1–18.
[13] T. Goto, D. Urabe, K. Masuda, Y. Isobe, M. Arita, M. Inoue, Total synthesis of four
stereoisomers of (4 Z, 7 Z, 10 Z, 12 E, 16 Z, 18 E)-14, 20-dihydroxy-4, 7, 10, 12, 16,
18-docosahexaenoic acid and their anti-inflammatory activities, J. Org. Chem. 80
(2015) 7713–7726.
[14] Y. Nanba, R. Shinohara, M. Morita, Y. Kobayashi, Stereoselective synthesis of 17,
18-epoxy derivative of EPA and stereoisomers of isoleukotoxin diol by ring opening
of TMS-substituted epoxide with dimsyl sodium, Org. Biomol. Chem. 15 (2017)
8614–8626.
Declaration of Competing Interest
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.
[15] S. Durand, J.-L. Parrain, M. Santelli, Construction of (Z, Z) skipped 1, 4-dienes.
Application to the synthesis of polyunsaturated fatty acids and derivatives, J. Chem.
Soc. Perkin Trans. 1 (2000) 253–273.
[16] A. Vik, T.V. Hansen, Synthetic manipulations of polyunsaturated fatty acids as a
convenient strategy for the synthesis of bioactive compounds, Org. Biomol. Chem.
16 (2018) 9319–9333.
Acknowledgements
[17] E.J. Corey, H. Niwa, J.R. Falck, Selective epoxidation of eicosa-cis-5, 8, 11, 14-
tetraenoic (arachidonic) acid and eicosa-cis-8, 11, 14-trienoic acid, J. Am. Chem.
Soc. 101 (1979) 1586–1587.
Training grant (Paterna A.) from FSE PO 2014-2020 (avviso 11/
2017 Regione Sicilia) is gratefully acknowledged. Thanks are also due
to Dr. Sandro Dattilo, Institute for Polymers, Composites and
Biomaterials of CNR, for HR-ESI-MS spectra.
[18] T. Galliard, D.R. Phillips, Lipoxygenase from potato tubers. Partial purification and
properties of an enzyme that specifically oxygenates the 9-position of linoleic acid,
Biochem. J. 124 (1971) 431–438.
[19] P.R.Q. Chang, A.R. McCurdy, Lipoxygenase activity in fourteen legumes, Can. Inst.
Food Sci. Technol. J. 18 (1985) 94–96.
References
[20] J.-M. Kuo, A. Hwang, D.-B. Yeh, M.-H. Pan, M.-L. Tsai, B.S. Pan, Lipoxygenase from
banana leaf: purification and characterization of an enzyme that catalyzes linoleic
acid oxygenation at the 9-position, J. Agric. Food Chem. 54 (2006) 3151–3156.
[21] C.J. Tsai, W.F. Li, B.S. Pan, Characterization and immobilization of marine algal 11-
lipoxygenase from Ulva fasciata, J. Am. Oil Chem. Soc. 85 (2008) 731–737.
[22] Z. Zhu, F. Qian, R. Yang, J. Chen, Q. Luo, H. Chen, X. Yan, A lipoxygenase from red
alga Pyropia haitanensis, a unique enzyme catalyzing the free radical reactions of
polyunsaturated fatty acids with triple ethylenic bonds, PLoS ONE 10 (2015)
e0117351.
[1] A. Mosblech, I. Feussner, I. Heilmann, Oxylipins: structurally diverse metabolites
from fatty acid oxidation, Plant Physiol. Biochem. 47 (2009) 511–517.
[2] M. Gabbs, S. Leng, J.G. Devassy, M. Monirujjaman, H.M. Aukema, Advances in our
understanding of oxylipins derived from dietary PUFAs, Adv. Nutr. 6 (2015)
513–540.
[3] G. Bannenberg, C.N. Serhan, Specialized pro-resolving lipid mediators in the in-
flammatory response: an update, Biochim. Biophys. Acta (BBA)-Mol. Cell Biol.
7

C. Sanfilippo, et al.
BioorganicChemistry93(2019)103325
[23] M. Hofrichter, R. Ullrich, Oxidations catalyzed by fungal peroxygenases, Curr. Opin.
Chem. Biol. 19 (2014) 116–125.
conjugated linoleic acid with peroxygenase, Lipids 38 (2003) 255–261.
[42] G.J. Piazza, A. Nuñez, T.A. Foglia, Epoxidation of fatty acids, fatty methyl esters,
and alkenes by immobilized oat seed peroxygenase, J. Mol. Catal. B Enzym. 21
(2003) 143–151.
[24] I. Plagemann, U. Krings, R.G. Berger, Isolation and characterization of wild-type
lipoxygenase LOXPsa1 from Pleurotus sapidus, Zeitsch. Naturforsch. C. 69 (2014)
149–154.
[43] G.J. Piazza, T.A. Foglia, One-pot synthesis of fatty acid epoxides from triacylgly-
cerols using enzymes present in oat seeds, J. Am. Oil Chem. Soc. 83 (2006)
1021–1025.
[25] H.M. Girvan, A.W. Munro, Applications of microbial cytochrome P450 enzymes in
biotechnology and synthetic biology, Curr. Opin. Chem. Biol. 31 (2016) 136–145.
[26] J.-U. An, S.-H. Hong, D.-K. Oh, Regiospecificity of a novel bacterial lipoxygenase
from Myxococcus xanthus for polyunsaturated fatty acids, Biochim. Biophys. Acta
(BBA)-Mol. Cell Biol. Lipids 1863 (2018) 823–833.
[44] E. Fernández-Fueyo, Y. Ni, A.G. Baraibar, M. Alcalde, L.M. van Langen,
F. Hollmann, Towards preparative peroxygenase-catalyzed oxyfunctionalization
reactions in organic media, J. Mol. Catal. B Enzym. 134 (2016) 347–352.
[45] Y. Wang, D. Lan, R. Durrani, F. Hollmann, Peroxygenases en route to becoming
dream catalysts. What are the opportunities and challenges? Curr. Opin. Chem. Biol.
37 (2017) 1–9.
[27] W. Minor, J. Steczko, B. Stec, Z. Otwinowski, J.T. Bolin, R. Walter, B. Axelrod,
Crystal structure of soybean lipoxygenase L-1 at 1.4 Å resolution, Biochemistry 35
(1996) 10687–10701.
[28] M.H. Glickman, J.P. Klinman, Lipoxygenase reaction mechanism: demonstration
that hydrogen abstraction from substrate precedes dioxygen binding during cata-
lytic turnover, Biochemistry 35 (1996) 12882–12892.
[46] I. Suda, M. Hajika, Y. Nishiba, S. Furuta, K. Igita, Simple and rapid method for the
selective detection of individual lipoxygenase isoenzymes in soybean seeds, J.
Agric. Food Chem. 43 (1995) 742–747.
[29] E. Skrzypczak-Jankun, L.M. Amzel, B.A. Kroa, M.O. Funk Jr, Structure of soybean
lipoxygenase L3 and a comparison with its L1 isoenzyme, Prot. Struct. Funct. Bioinf.
29 (1997) 15–31.
[47] C. de los Reyes, J. Ávila-Román, M.J. Ortega, A. de la Jara, S. García-Mauriño,
V. Motilva, E. Zubía, Oxylipins from the microalgae chlamydomonas debaryana and
Nannochloropsis gaditana and their activity as TNF-α inhibitors, Phytochemistry
102 (2014) 152–161.
[30] C.P.A. Van Os, G.P.M. Rijke-Schilder, J.F.G. Vliegenthart, 9-lR-linoleyl hydroper-
oxide, a novel product from the oxygenation of linoleic acid by type-2 lbpox-
ygenases from soybeans and peas, Biochim. Biophys. Acta (BBA)-Lipids Lipid
Metab. 575 (1979) 479–484.
[48] R. Morrone, N. D’Antona, D. Biondi, D. Lambusta, G. Nicolosi, Preparation of n-3
PUFAs ethyl esters by an efficient biocatalyzed solvent-free process, J. Mol. Catal. B
Enzym. 84 (2012) 173–176.
[31] S. Nanda, J.S. Yadav, Lipoxygenase biocatalysis: a survey of asymmetric oxygena-
tion, J. Mol. Catal. B Enzym. 26 (2003) 3–28.
[49] D. Nanjappa, G. D’Ippolito, C. Gallo, A. Zingone, A. Fontana, Oxylipin diversity in
the diatom family leptocylindraceae reveals DHA derivatives in marine diatoms,
Mar. Drugs 12 (2014) 368–384.
[32] B.-J. Kim, K.-C. Shin, D.-K. Oh, Enzymatic production of 15-hydro-
xyeicosatetraenoic acid from arachidonic acid by using soybean lipoxygenase, J.
Microbiol. Biotechnol. 24 (2014) 359–362.
[50] J. Rettner, M. Werner, N. Meyer, O. Werz, G. Pohnert, Survey of the C20 and C22
oxylipin family in marine diatoms, Tetrahedron Lett. 59 (2018) 828–831.
[51] P.A. Desbois, C.K. Lawlor, Antibacterial activity of long-chain polyunsaturated fatty
acids against propionibacterium acnes and staphylococcus aureus, Mar. Drugs 11
(2013) 4544–4577.
[33] T. Itoh, T. Saito, Y. Yamamoto, H. Ishida, K. Yamamoto, Gram scale synthesis of
specialized pro-resolving mediator 17 (S)-HDHA using lipoxygenase enhanced by
water-soluble reducing agent TCEP, Bioorg. Med. Chem. Lett. 26 (2016) 343–345.
[34] I.A. Butovich, A one-step method of 10, 17-dihydro (pero) xydocosahexa-4Z, 7Z,
11E, 13Z, 15E, 19Z-enoic acid synthesis by soybean lipoxygenase, J. Lipid Res. 47
(2006) 854–863.
[52] G.J. Piazza, T.A. Foglia, A. Nuñez, Epoxidation of fatty acids with membrane-sup-
ported peroxygenase, Biotechnol. Lett. 22 (2000) 217–221.
[53] G.J. Piazza, A. Nuñez, T. A. Foglia, US Patent 6,485,949 B1, 26 Nov. 2002.
[54] J.H. Capdevila, S. Wei, C. Helvig, J.R. Falck, Y. Belosludtsev, G. Truan,
S.E. Graham-Lorence, J.A. Peterson, The highly stereoselective oxidation of poly-
unsaturated fatty acids by cytochrome P450BM-3, J. Biol. Chem. 271 (1996)
22663–22671.
[35] E.P. Dobson, C.J. Barrow, J.L. Adcock, Solvent-induced 7R-dioxygenase activity of
soybean 15-lipoxygenase-1 in the formation of omega-3 DPA-derived resolvin
analogs, J. Mol. Catal. B Enzym. 108 (2014) 96–102.
[36] E.P. Dobson, C.J. Barrow, J.A. Kralovec, J.L. Adcock, Controlled formation of mono-
and dihydroxy-resolvins from EPA and DHA using soybean 15-lipoxygenase, J.
Lipid Res. 54 (2013) 1439–1447.
[55] D. Schwarz, P. Kisselev, S.S. Ericksen, G.D. Szklarz, A. Chernogolov, H. Honeck, W.-
H. Schunck, I. Roots, Arachidonic and eicosapentaenoic acid metabolism by human
CYP1A1: highly stereoselective formation of 17(R),18(S)-epoxyeicosatetraenoic
acid, Biochem. Pharmacol. 67 (2004) 1445–1457.
[37] M.B.W. Elshof, M. Janssen, G.A. Veldink, J.F.G. Vliegenthart, Biocatalytic large-
scale production of 13 (S)-hydroperoxy-9 (Z), 11 (E) octadecadienoic acid from
hydrolysed safflower oil by a crude soybean-flour extract as lipoxygenase source,
Recl. Des Trav. Chim. Des Pays-Bas. 115 (1996) 499–504.
[56] D. Lucas, S. Goulitquer, J. Marienhagen, M. Fer, Y. Dreano, U. Schwaneberg,
Y. Amet, L. Corcos, Stereoselective epoxidation of the last double bond of poly-
unsaturated fatty acids by human cytochromes P450, J. Lipid Res. 51 (2010)
1125–1133.
[38] H.-A.T. Tu, E.P. Dobson, L.C. Henderson, C.J. Barrow, J.L. Adcock, Soy flour as an
alternative to purified lipoxygenase for the enzymatic synthesis of resolvin analo-
gues, N. Biotechnol. 41 (2018) 25–33.
[57] P.H. Cui, N. Petrovic, M. Murray, The ω-3 epoxide of eicosapentaenoic acid inhibits
endothelial cell proliferation by p38 MAP kinase activation and cyclin D1/CDK4
down-regulation, Br. J. Pharmacol. 162 (2011) 1143–1155.
[39] M. Hamberg, G. Hamberg, Peroxygenase-catalyzed fatty acid epoxidation in cereal
seeds (sequential oxidation of linoleic acid into 9 (S), 12 (S), 13 (S)-trihydroxy-10
(E)-octadecenoic acid), Plant Physiol. 110 (1996) 807–815.
[58] J.R. Falck, G. Wallukat, N. Puli, M. Goli, C. Arnold, A. Konkel, M. Rothe, R. Fischer,
D.N. Müller, W.-H. Schunck, 17(R),18(S)-epoxyeicosatetraenoic acid, a potent ei-
cosapentaenoic acid (EPA) derived regulator of cardiomyocyte contraction: struc-
ture-activity relationships and stable analogues, J. Med. Chem. 54 (2011)
4109–4118.
[40] A. Hanano, M. Burcklen, M. Flenet, A. Ivancich, M. Louwagie, J. Garin, E. Blée,
Plant seed peroxygenase is an original heme-oxygenase with an EF-hand calcium
binding motif, J. Biol. Chem. 281 (2006) 33140–33151.
[41] G.J. Piazza, A. Nuñez, T.A. Foglia, Isolation of unsaturated diols after oxidation of
8