Biocatalytic study of novel oleate hydratases

Article abstract of DOI:10.1016/j.molcatb.2017.01.010

The direct hydration of C[dbnd]C bonds to yield alcohols or the reverse dehydration is chemically challenging but highly sought after. Recently, oleate hydratases (OAHs) gained attention as biocatalytic alternatives capable of hydrating isolated, non-activated C[dbnd]C bonds. Their natural reaction is the conversion of oleic acid to (R)-10-hydroxystearic acid. In this work, we report the first comparative study of several OAHs. Therefore we established the Hydratase Engineering Database (HyED) comprising 2046 putative OAHs from eleven homologous families and selected nine homologs for cloning in E. coli. The heterologously expressed enzymes were evaluated concerning activity and substrate specificity. The enzymes have a broad substrate scope ranging from oleic acid (C18) to the novel synthetic substrate (Z)-undec-9-enoic acid (C11). The OAHs from Elizabethkingia meningoseptica and Chryseobacterium gleum showed the best expression, highest stability and broadest substrate scope, making them interesting candidates for directed evolution to engineer them for the application as general hydratase catalysts.

Full text of DOI:10.1016/j.molcatb.2017.01.010

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Contents lists available at ScienceDirect
Journal of Molecular Catalysis B: Enzymatic
journal homepage: www.elsevier.com/locate/molcatb
Biocatalytic study of novel oleate hydratases
Jens Schmid1, Lisa Steiner1, Silvia Fademrecht, Jürgen Pleiss, Konrad B. Otte,
Bernhard Hauer∗
Institute of Technical Biochemistry, University of Stuttgart, Allmandring 31, D-70569 Stuttgart, Germany
a r t i c l e i n f o
a b s t r a c t
Article history:
The direct hydration of C C bonds to yield alcohols or the reverse dehydration is chemically challenging
but highly sought after. Recently, oleate hydratases (OAHs) gained attention as biocatalytic alternatives
capable of hydrating isolated, non-activated C C bonds. Their natural reaction is the conversion of oleic
acid to (R)-10-hydroxystearic acid.
Received 18 October 2016
Received in revised form 10 January 2017
Accepted 11 January 2017
Available online xxx
In this work, we report the first comparative study of several OAHs. Therefore we established the
Hydratase Engineering Database (HyED) comprising 2046 putative OAHs from eleven homologous families
and selected nine homologs for cloning in E. coli. The heterologously expressed enzymes were evaluated
concerning activity and substrate specificity. The enzymes have a broad substrate scope ranging from oleic
acid (C18) to the novel synthetic substrate (Z)-undec-9-enoic acid (C11). The OAHs from Elizabethkingia
meningoseptica and Chryseobacterium gleum showed the best expression, highest stability and broadest
substrate scope, making them interesting candidates for directed evolution to engineer them for the
application as general hydratase catalysts.
Keywords:
Alkenes
Biocatalysis
Database analysis
Hydration
Oleate hydratases
© 2017 Elsevier B.V. All rights reserved.
1. Introduction
OAHs (EC 4.2.1.53) are promising biocatalysts for the selec-
tive addition of water to C C bonds. They were first described by
In recent years, hydratases received increasing attention as pos-
sible biocatalysts, as they enable the regio- and stereoselective
addition of water to double bonds or the reverse water elimina-
tion. Chemical hydrations require harsh acidic conditions, which
lead in general to a broad range of side-reactions and a significant
loss of selectivity [1].
Hydratases are classified as hydro-lyases (EC 4.2.1) and can be
divided into two groups regarding their substrates and need of
cofactors [2,3]. Cofactor-dependent hydratases add water to acti-
vated ␣,␤-unsaturated carbonyl compounds, whereas hydratases
acting on isolated carbon–carbon double bonds are described to be
cofactor-independent [3,4]. Rare examples for the latter group are
the carotenoid hydratase [5,6], the linalool dehydratase-isomerase
[7] and the oleate hydratase (OAH) [8].
Wallen et al. in 1962 when they found Elizabethkingia meningosep-
tica (formerly known as Pseudomonas sp. strain 3266) capable of
hydrating oleic acid to 10-hydroxystearic acid [9]. Since then, a
number of other strains were found to possess OAHs and sev-
eral studies have been performed on the selectivity, the kinetics
and the mechanism of the reaction [8]. Isotope labeling exper-
iments confirmed that the alcohol group was introduced at the
C-10 position [10]. Schroepfer and Bloch showed that the hydroxyl
group is formed stereoselectively in (R)-configuration [11]. They
also proved that the reaction does not take place via an initial
epoxidation and subsequent reductive opening by conducting the
reaction under anaerobic conditions. Furthermore, the utilization of
9,10-epoxyoctadecanoic acid as precursor did not lead to the for-
mation of 10-hydroxystearic acid [12,13]. Other studies on novel
fatty acid hydratases also addressed regioselectivity. For instance,
the hydratase from Macrococcus caseolyticus was most active for
oleic acid indicating that it was an OAH, however, hydration also
occurred at cis-12 C C bonds [14]. Only in 2015, a second hydratase
from Lactobacillus acidophilus (La) was discovered, catalyzing the
addition of water selectively to the 12Z-position [15]. In previous
studies, however, it was reported that most hydratases add water
exclusively to the 9Z-double bond, with the hydroxyl group being
introduced at the C-10 atom [8,16–18].
Abbreviations:
C C, carbon–carbon double bond; OAH(s), oleate hydratase(s);
HyED, Hydratase Engineering Database; HFam(s), homologous family/families;
gDNA, genomic DNA; Cg, Chryseobacterium gleum; Db, Desulfomicrobium baculatum;
Em, Elizabethkingia meningoseptica; Gm, Gemella morbillorum; Ht, Haloterri-
gena thermotolerans; La, Lactobacillus acidophilus; Lj, Lactobacillus johnsonii; Mp,
Mucilaginibacter paludis; Tt, Thermoanaerobacterium thermosaccharolyticum.
∗
Corresponding author.
E-mail address: Bernhard.Hauer@itb.uni-stuttgart.de (B. Hauer).
These authors contributed equally to this work
1
http://dx.doi.org/10.1016/j.molcatb.2017.01.010
1381-1177/© 2017 Elsevier B.V. All rights reserved.
Please cite this article in press as: J. Schmid, et al., Biocatalytic study of novel oleate hydratases, J. Mol. Catal. B: Enzym. (2017),
http://dx.doi.org/10.1016/j.molcatb.2017.01.010

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The first OAH was isolated in 2009 from Elizabethkingia
meningoseptica (Em-OAH1), subsequently cloned in E. coli and bio-
chemically characterized [8]. Recently, the crystal structures of
La-OAH1 and Em-OAH1 were solved, giving more insight into the
FAD binding and the reaction mechanism [18,19]. It was revealed
that the Em-OAH1 is a dimer and each monomer contains not only
a Ca2+-ion but also FAD as prosthetic group. However, the exact
function of FAD still remains unclear. A structural or mechanistic
purpose has been discussed [18,19].
downloaded from the Protein Data Bank (PDB) and stored in the
respective protein entry [31]. The Hydratase Engineering Database is
publicly available at https://hyed.biocatnet.de and can be browsed
by sequence, structure or source organism. All sequences, multiple
sequence alignments and phylogenetic trees can be accessed online
or be downloaded.
2.3. Conservation analysis
Since the first OAH was described, several hydratases from
different organisms followed, but were in most cases not char-
acterized in detail. Nevertheless, OAHs are successfully applied in
larger-scale biotransformations of unsaturated fatty acids (mainly
oleic acid) [1]. The resulting hydroxy fatty acids and their deriva-
tives are applied in a broad range from detergents and paintings
to biopolymers and flavor compounds [2,20,21]. Beside the pro-
duction of 10-hydroxystearic acid, OAHs have also been embedded
in cascade reactions for the synthesis of plastic monomers (␣,␻-
dicarboxylic acids for polyesters and amino acids for polyamides)
from renewable fatty acids [1,22,23]. Moreover, the enzymatic
dehydration of short chain alcohols (e. g. isobutanol) was described
[2,24].
For identification of conserved positions, a multiple sequence
alignment of all sequences of the database was calculated using
Clustal Omega [32]. For each alignment column, the occurrence
of each amino acid was counted. Positions were considered to be
conserved if one or two amino acids were found in >90 % of all
sequences.
2.4. Cloning of homologous enzymes
The OAH genes were amplified from gDNA via PCR (see Table S2
for primers, Tables S3, S5 and S7 for reaction mixture and Tables S4,
S6 and S8 for temperature program). pET28a(+) was used as vector
backbone. For the cloning of La- and Lj-OAH1, the NcoI restriction
site in pET28a(+) was changed to NdeI by using the ligation-during-
amplification method. Cg-OAH1 was ligated into pET28a(+) using
the XbaI and XhoI restriction sites. For Db-, Em-, Gm-, Mp- and Tt-
OAH1 NcoI and XhoI were used. For Ht-OAH1 XbaI and HindIII, for
La- and Lj-OAH1 the restriction sites NdeI and XhoI were used.
As most of the described enzymes were investigated with diver-
gent substrate panels (ranging from saturated to polyunsaturated
fatty acids with a chain length from C12 to C22) and with different
reaction set-ups (in vitro or in vivo), it is difficult to directly compare
the different enzymes [4].
Herein, we present the compilation of
a comprehensive
database and activity assessment of nine representative hydratases
from three different homologous families with varying similar-
ities on one profound substrate panel. In combination with the
investigation of the enzymes’ stability and their capability to be
overexpressed in E. coli, this study aims for the identification of
an ideal starting candidate for directed mutagenesis as well as the
generation of new insights into the growing OAH family.
2.5. Expression of hydratase genes
A preculture of E. coli BL21 (DE3) containing the corresponding
plasmid was used to inoculate a 2 L baffled flask containing 400 mL
TB medium and kanamycin (final concentration: 30 ␮g mL−1) to an
OD600 of 0.05. The cultures were incubated at 37 ◦C and 180 rpm
until an OD600 of 0.5 was reached. Expression was induced by
adding 0.5 mM IPTG at 20 ◦C. After 16 h at 20 ◦C and 140 rpm the
cells were harvested by centrifugation (30 min, 20,450 × g, 4 ◦C) and
the pellet was stored at −20 ◦C until further use.
2. Experimental
2.1. Chemicals & DNA
The chemicals were purchased from the following suppliers:
Sigma-Aldrich (St. Louis, USA), Serva (Heidelberg, DE), Merck
(Darmstadt, DE), Peqlab (Erlangen, DE), Alfa Aesar (Karlsruhe, DE),
Carl Roth (Karlsruhe, DE) and Macherey-Nagel (Düren, DE).
Genomic DNA (gDNA) as a source of desired OAH genes was
obtained from DSMZ (Leibniz-Institut, Braunschweig, DE). Oligonu-
cleotides were obtained from Metabion (Planegg, DE).
Cell disruption was achieved either by sonication or chemical
lysis. For sonication, a cell suspension (10 mL g−1 cell pellet in the
same buffer as used for reaction) was prepared and kept on ice dur-
ing disruption using a Branson Sonifier 250 (5 cycles: 1 min on/45 s
off, duty cycle 30 %, output 4). Lysate was obtained by centrifu-
gation (20 min, 894 × g, 4 ◦C) as supernatant. For chemical lysis,
harvested cells were resuspended at room temperature in 5 mL g−1
1 x BugBuster Protein Extraction Reagent (Merck, Darmstadt, DE) to
which a spatula tip of DNase was added. The suspension was shaken
(20 min, 25 ◦C, 300 rpm) and subsequently centrifuged (20 min,
16,000 × g, 4 ◦C).The protein concentration was determined by
bicinchoninic acid assay kit (Pierce BCA Protein Assay Kit, Thermo
Scientific, Waltham, USA) according to manufacturer’s microwell
plate protocol and the lysate was analyzed by SDS-PAGE (according
to Laemmli [33]).
Therefore, the obtained lysate was diluted to a concentration of
2 g L−1 and mixed with the same volume of 2 x SDS sample buffer
(2 mL 1 M TRIS/HCl pH 6.8; 190 mg MgCl2; 1 mL glycerol; 0.8 g SDS;
2 mg bromophenol blue; 310 mg DTT, water ad to 20 mL). Whole
cell samples were taken during or after expression normalized to
OD600 = 0.25 and resuspended in 50 ␮L 2 x SDS sample buffer. Both,
whole cell and lysate samples, were denatured at 95 ◦C for 10 min.
Denatured samples were loaded (10 ␮L for lysate, 15 ␮L for pellets)
on the gels along with 5 ␮L PageRuler Prestained Protein Ladder
(Thermo Scientific, product # 26616).
2.2. Database setup
The Hydratase Engineering Database (HyED) was established
within our in-house database system BioCatNet [25] based on 20
sequences (Table S1) derived from a hydratase patent by Mar-
liere describing the dehydration of alcohols for the production
of alkenes [24]. A BLAST search in the non-redundant protein
database of the NCBI was performed for each of these sequences
using an expectation threshold of 10−10 [26,27]. Hits sharing a
sequence identity of >98 % were assigned to a single protein entry.
Homologous families were automatically generated by cluster-
ing all sequences with USEARCH and an identity cut-off of 50 %
[28]. For further classifications into superfamilies, all sequences
were compared using the Needleman-Wunsch algorithm [29] for
pairwise sequence alignments. Similarities were visualized using
sequence similarity networks generated with Cytoscape version
3.2.1 and the organic layout (Fig. 1) [30]. Available structures were
Please cite this article in press as: J. Schmid, et al., Biocatalytic study of novel oleate hydratases, J. Mol. Catal. B: Enzym. (2017),
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2.6. Biotransformations and analytics
2.8. Freeze-drying and determination of residual activity
For characterization of the enzymes, biotransformations were
performed in triplicates and conducted on 500 ␮L-scale in 1.5 mL
reaction tubes. Protein concentrations and reaction times are listed
in Table 1. Negative controls were performed using lysate of cells
containing empty pET28a(+) vector. All reactions were performed
in 50 mM potassium phosphate buffer pH 6.5 except for biotrans-
formations of (Z)-undec-9-enoic acid and dec-9-enoic acid, which
were performed in 50 mM citrate buffer pH 6. Substrate was added
to a final concentration of 500 ␮M. The reaction mixture was shaken
at 25 ◦C and 600 rpm. The reaction was quenched by the addition of
20 ␮L 1 M HCl, extracted two times with 500 ␮L methyl tert-butyl
ether (MTBE) and 100 ␮M internal standard (heptadecanoic acid
for linoleic acid, oleic acid and palmitoleic acid or pentadecanoic
acid for myristoleic acid, (Z)-undec-9-enoic acid and dec-9-enoic
acid). The organic phase was concentrated in vacuo and the residue
derivatized using 40 ␮L N,O-bis(trimethylsilyl)trifluoroacetamide
containing 1 % trimethylchlorosilane.
Derivatized samples were analyzed on a Shimadzu GC2010
Plus (Shimadzu, Kyo¯to, JP) equipped with an AOC 20 s autosam-
pler, AOC 20i autoinjector (injection volume: 1 ␮L, split ratio: 1:10,
injection temperature: 250 ◦C), carrier gas: hydrogen (linear veloc-
ity: 30 cm s−1), column: DB-5 polyphenylmethylsiloxane (Agilent
technologies, Santa Clara, USA) (30 m, 0.25 mm, 0.25 ␮m), detector
temperature 320 ◦C.
Cells were sonicated in 50 mM ammonium acetate buffer (pH
7.5) as described above. The obtained lysate was frozen in liq-
uid nitrogen and lyophilized for five days using a freeze-dryer
(Christ Alpha 2–4LD plus, Martin Christ Gefriertrocknungsanlagen,
Osterode am Harz, DE). The lyophilized proteins were resolved
in 50 mM potassium phosphate buffer (pH 6.5) and the residual
activity was determined by comparing product formation in bio-
transformations (cf. 2.6) regarding the conversion of 500 ␮M oleic
acid (1 h, 37 ◦C, final protein concentration: 0.5 g L−1) using fresh
lysate or resuspended lyophilisate.
2.9. Synthesis of (Z)-undec-9-enoic acid
In a dry Schlenk tube, 9-undecynoic acid (0.48 g, 2.64 mmol)
was added to Lindlar catalyst (0.06 g) dissolved in 15 mL methanol
(dried over 3 Å molecular sieve) and stirred at room temperature.
The atmosphere was replaced ten times with hydrogen by sub-
sequent evacuation and hydrogen flooding. Afterwards, hydrogen
was allowed to pass through the reaction mixture for 20 h. Finally,
the grey mixture was filtered over celite, washed with methanol
and the filtrate was concentrated in vacuo to yield a yellowish
oil which was purified by column chromatography (cyclohex-
ane/MTBE 5:1, Rf = 0.36, vanillin) to yield (Z)-undec-9-enoic acid
(0.18 g, 0.98 mmol, 38 %) as a colorless oil (>95 % 1H NMR purity).
1H NMR (500 MHz, CDCl3): ␦ = 1.32–1.35 (m, 8H, 4-H, 5-H, 6-H,
7-H), 1.60 (d, J = 6.3 Hz, 3H, 11-H), 1.62–1.65 (m, 2H, 3-H), 2.00–2.04
(m, 2H, 8-H), 2.35 (t, J = 7.5 Hz, 2H, 2-H), 5.36-5.40 (m, 2H, 9-H, 10-
H) ppm.13C NMR (125 MHz, CDCl3): ␦ = 12.8, 24.7, 26.8, 29.0, 29.0,
29.1, 29.5, 33.9, 123.7, 130.8, 179.6 ppm. FT-IR (ATR): v˜ = 3013 (w),
2924 (m), 2855 (m), 1705 (s), 1412 (m), 1284 (m), 1245 (m), 1114
For oleic acid, the oven temperature was set to 180 ◦C for 2 min,
then raised to 240 ◦C at a speed of 15 K min−1, held for 6 min and
finally increased to 300 ◦C at a rate of 20 K min−1 and kept at this
temperature for 2 min. For linoleic acid, palmitoleic acid, myris-
toleic acid, (Z)-undec-9-enoic acid and dec-9-enoic acid, the oven
temperature was set to 50 ◦C for 2 min, then raised to 100 ◦C at a
speed of 10 K min−1, subsequently raised to 200 ◦C with a rate of
(w), 934 (m), 700 (m) cm−1
.
30 K min−1 and raised moreover to 240 ◦C at a speed of 10 K min−1
.
Finally, the temperature was increased to 300 ◦C at 40 K min−1 and
held for 3 min.
3. Results and discussion
GC–MS analysis was performed on
a Shimadzu GC2010
system (Shimadzu, Kyo¯to, JP) equipped with a GCMS-QP2010
mass selective detector (electron impact, 70 eV), AOC 5000
autosampler/autoinjector (injection volume: 1 ␮L, split ratio: 1:20,
injection temperature: 250 ◦C), carrier gas: helium (linear veloc-
ity: 30 cm s−1), column: ZB-5 polyphenylmethylsiloxane (Zebron
− Phenomenex, Torrance, USA) (30 m, 0.25 mm, 0.25 ␮m), detector
temperature: 320 ◦C); MS: ion source temperature 200 ◦C, inter-
face temperature 285 ◦C. The oven temperature was set to 50 ◦C
for 2 min, and then raised to 100 ◦C at a speed of 10 K min−1, sub-
sequently raised to 200 ◦C with a rate of 30 K min−1 and raised
moreover to 240 ◦C at a speed of 10 K min−1. Finally, the tempera-
ture was increased to 300 ◦C at 40 K min−1 and held for 10 min.
3.1. Database assembly
The Hydratase Engineering Database (https://hyed.biocatnet.de)
was established in the framework of our in-house database system
BioCatNet [25]. The database includes 2046 sequences and thereby
increases the number of previously known putative hydratases
noteably. These sequences were assigned to eleven homologous
families (HFam1-11) sharing a high sequence similarity (average
global sequence identity of 62 % inside the families) (Fig. 1). HFam2
constitutes the largest homologous family (1188 sequences)
and includes the Lactobacillus acidophilus oleate hydratase (La-
OAH1, pdb entries 4IA5 and 4IA6). The next larger families are
HFam1 (436 sequences) and HFam3 (191 sequences). HFam11
(116 sequences) contains the Elizabethkingia meningoseptica oleate
hydratase (Em-OAH1, pdb entry 4UIR). For all other homologous
families (HFam4-10), no structure information is available. These
families contain less than 50 sequences each. The putative OAHs
were mainly derived from bacterial sources, but also 103 fungal
and 24 archaeal sequences are included in the database (Fig. S1).
To identify structurally and functionally important positions,
the homologous families were investigated for similarities and for
systematic differences in their conservation patterns. 80 Positions
were found to be highly conserved among all sequences (identical
amino acids in >90 % of the sequences, Fig. S2, Table S9).
2.7. ThermoFAD
The thermostability of OAHs was measured by ThermoFAD
according to a protocol published elsewhere [34,35]. The protein
lysate samples (25 ␮L) were heated in a microwell plate using
a real-time PCR machine (Eppendorf Mastercycler epgradient S,
Hamburg, DE). The temperature was increased stepwise at 0.5
K from 20 ◦C to 90 ◦C. During the unfolding process of the pro-
teins, FAD is released and the fluorescence signal was measured
at 543 nm. The melting point was determined from the minimum
of the second derivation of the fluorescence signal. ThermoFAD was
performed in triplicates of the same lysate.
Some of these conserved positions have been described
in literature before: The glycine-rich motif of the Rossmann-
fold (usually described as GxGxxG) was described as
GxGxx[GSAN]x(15)[EKD]x(5)[EDGS] for OAHs based on a sequence
alignment using ten enzymes [4]. The conservation analysis
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Fig. 1. Sequence similarity network of the HyED showing the classification into eleven homologous families (HFams). The previously characterized oleate hydratase from
Lactobacillus acidophilus (black) is found in HFam2, while the oleate hydratase from Elizabethkingia meningoseptica (yellow) is found in HFam11. The sequences selected for
characterization are from HFam1, HFam2 and HFam11 and shown as colored triangles.
based on all sequences of the HyED shows that this region is
even more conserved than indicated by the sequence pattern
derived from literature: for 16 of the 28 positions described by
this motif, already two amino acids are sufficient to describe
90 % of the OAHs (Table S10). A more specific and sensitive
sequence motif for the FAD-cofactor binding region of OAHs is
3.2. Selection of homologs
Representative OAHs were selected from literature and our
database (Fig. 1) for the comparative investigation of properties
interesting for biocatalysis including stability, activity and sub-
strate specificity. The two already described oleate hydratases
La-OAH1 [18] (HFam2) and Em-OAH1 [19] (HFam11) for which
also structural information is available, as well as representa-
tive homologs from these families were chosen: The putative
oleate hydratases from Lactobacillus johnsonii (Lj-OAH1), Gemella
morbillorum (Gm-OAH1) and Desulfomicrobium baculatum (Db-
OAH1) [24] from HFam2 were selected for their high sequence
similarity to La-OAH1. Chryseobacterium gleum (Cg-OAH1) from
HFam11 was chosen for its high sequence similarity to Em-OAH1.
Since members of HFam1 show an interesting family-specific con-
servation, three additional putative oleate hydratases from the
thermophilic archaeon Haloterrigena thermotolerans (Ht-OAH1),
from the thermophilic bacterium Mucilaginibacter paludis (Mp-
OAH1) and from Thermoanaerobacterium thermosaccharolyticum
(Tt-OAH1) [24] were selected, especially for their potentially
increased (thermo)stability.
G69xG[LI][AG]x[LM][AS][AG]Ax[FY][LM][IV]R[DE][GA]x(3)Gxx[IV]
x[IFVL][LFY]E96 (positions according to Em-OAH1). Interestingly,
the motif derived from literature describes a charged residue at
position 90 (position underlined in both sequence motifs). Due to
our criteria no residue is conserved at this position, but a tendency
can be seen (35 % E, 16 % K, 15 % S, 14 % D, 5 % Q, 4 % N). For
some of the positions found in the cofactor binding region, specific
functions were reported in previous studies [36,37]. The conserved
glycines at positions 69 and 71 were described to facilitate the
positioning of the FAD. Further, residues at positions 78 and
74 were reported to stabilize the interaction between the first
␤-strand and the following ␣-helix [36,37]. However, although a
tendency towards small amino acids can be seen for position 74,
no residue is found to be conserved.
Of the other regions conserved in OAHs, the loop region
R118GGREM123 (positions according to Em-OAH1) has been
described to have a structural role for cofactor binding and an addi-
tional role in catalysis has been proposed for E122 [19]. Indeed,
this region is highly conserved among all database members (Table
S11). Other highly conserved residues in Em-OAH1 are Y241, which
was also proposed to be involved in the reaction [19] and the
residues suggested to bind the carboxylate of the oleate substrate
(Q265, T436, N438 and H442) [19].
Interestingly, some of these positions showed a family specific
conservation (Table S12). Instead of the proposed catalytic glutamic
acid at position 122, methionine is conserved in HFam1. HFam1 also
forms an exception at position 123 where predominantly leucine
is found, as well as at the position 436 where valine is conserved
and position 438 where predominantly small residues are found.
For none of the other conserved or family-specific conserved
positions, a function has been proposed yet. However, several of
them can be found in the substrate binding pocket and thus, are
supposed to contribute to substrate binding or catalysis.
3.3. Activity, regioselectivity and stability
The nine selected genes were cloned into pET28a(+) vector for
expression in E. coli BL21 (DE3). A small expression study was
performed to optimize several parameters including incubation
temperature, time and induction method. The improved expression
procedure was then utilized to investigate the overexpression capa-
bility and the individual expression levels were compared (Figure
S3). Overexpression was detectable for seven out of nine homol-
ogous genes (∼65 kDa). Two hydratase genes from Haloterrigena
thermotolerans and Thermoanaerobacterium thermosaccharolyticum
showed no expression. The expression of the oleate hydratase
genes from Mucilaginibacter paludis and Lactobacillus johnsonii led
to the formation of inclusion bodies. The five oleate hydratases
from Chryseobacterium gleum, Desulfomicrobium baculatum, Eliza-
bethkingia meningoseptica, Gemella morbillorum and Lactobacillus
acidophilus showed good overexpression.
Please cite this article in press as: J. Schmid, et al., Biocatalytic study of novel oleate hydratases, J. Mol. Catal. B: Enzym. (2017),
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Activity tests were performed with oleic acid (C18:1) as sub-
strate. Four hydratases showed moderate to high conversion
(Cg-OAH1: 87 %, Db-OAH1: 26 %, Em-OAH1 89 % and La-OAH1: 33
%), one hydratase (Gm-OAH1) showed only minor conversion (2 %).
Lj- and Mp-OAH1 formed inclusion bodies during expression. No
activity was detected for lysates of cells showing no overexpres-
sion in SDS-PAGE. Negative controls were performed using lysate
of induced cells containing empty pET28a(+) vector.
In order to characterize the enzyme panel regarding regiose-
lectivity, the active homologs were incubated with linoleic acid
(C18:2) and the fragmentation patterns of the hydrated products
were analyzed via GC–MS. The five enzymes were also active with
linoleic acid, forming the 10-hydroxy product (Table 1).
To analyze robustness, we investigated thermostability and
residual activity after freeze-drying (Table S13). All homologs
showed comparable melting points Tm ranging from 56.0 0.2 ◦C
for La-OAH1 to 65.6 0.2 ◦C for Em-OAH1. This is in accordance with
a study of the Lysinibacillus fusiformis oleate hydratase demonstrat-
ing that the enzyme was stable at 40 ◦C for 1 h showing increasingly
faster denaturation for temperatures between 40 and 70 ◦C [38].
Clear differences in their behavior were measurable by compar-
ing their activity for the hydration of oleic acid under the same
conditions before and after freeze-drying. Db- and Gm-OAH1 lost
a majority of their initial activity. After freeze-drying only 17 %
or 15 %, respectively, were detectable. La-OAH1 retained 57 %,
whereas Cg- and Em-OAH1 out of HFam11 where clearly not as
much affected as the other enzymes from HFam2. Cg-OAH1 lost 9
% activity and Em-OAH1 only 7 %.
from whole cells over cell-free extract to purified enzymes are
utilized [19,39,41,42]. In the case of fatty acids, this can have a
huge impact on the performance of biotransformations as the long-
chain substrates have to be actively imported in the case of whole
cells, whereas cell-free protein extract does not face this limitation
[43,44]. Therefore we preferred cell-free protein extract over whole
cells for studying the substrate scope. In order to obtain comparable
results we focused on the substrate panel shown in Table 1. This set
comprises six different substrates. Starting from the C18 substrates
oleic acid and linoleic acid, the latter used for the determination of
the regioselectivity, we continued with successively shorter sub-
strates. Biotransformations were performed for these substrates
with all cloned enzymes regardless of their expression behavior.
Consistent with the SDS-PAGE (Fig. S3), which showed no over-
expression for the hydratases from the thermophilic organisms,
no activity could be detected for the enzymes from Haloterrigena
thermotolerans (Ht-OAH1) and Thermoanaerobacterium thermosac-
charolyticum (Tt-OAH1) towards any of the substrates. This was also
the case for Mp-OAH1 from Mucilaginibacter paludis and Lj-OAH1
from Lactobacillus johnsonii because of the formation of inclusion
bodies. Negative controls were performed using lysate of induced
cells containing an empty pET28a(+) vector.
The activity of OAHs for the natural substrate oleic acid (C18:1)
was high, leading to a conversion of up to 90 % after 5 min. Never-
theless, due to the different kinetics for each substrate, the reaction
conditions had to be adjusted accordingly in terms of reaction time
and enzyme concentration. The different parameters and results of
the biotransformations are listed in Table 1.
Oleic acid (C18:1) was the best substrate and high conversions
were reached with Cg-OAH1 and Em-OAH1, moderate product
formation was observed with Db-OAH1 and La-OAH1 whereas Gm-
OAH1 was almost inactive at protein concentrations of 0.15 g L−1
and reaction times of 5 min. With shorter chain lengths of the
substrates, the reaction times were prolonged and the enzyme con-
centrations were increased to reach a substantial and comparable
amount of product. For the short-chain substrates (Z)-undec-
9-enoic acid (C11:1) and dec-9-enoic acid (C10:1) the protein
concentration was increased to 4 g L−1 and the reaction time pro-
longed to 5 d.
Interestingly, Gm-OAH1 (Gemella morbillorum) was more active
on the shorter chain substrates relative to the homologous enzymes
compared with the biotransformation of oleic acid (Table 1).
The remaining active homologs behaved relatively similar when
compared amongst each other. However, some remarkable dif-
ferences are recognizable. The two hydratases from Lactobacillus
acidophilus (La-OAH1) and Desulfomicrobium baculatum (Db-OAH1)
exhibited a lower activity towards the long-chain substrates C18:1
and C18:2 but showed comparable high activity to the medium
chain substrates C16:1 and C14:1. Gm-OAH1, also found in HFam2,
displayed even lower activity on linoleic acid (C18:2) and oleic
acid (C18:1), but had a relative high or increased activity towards
substrates of shorter chain length. The two homologous enzymes
from Chryseobacterium gleum and Elizabethkingia meningoseptica
demonstrateda good overall performance throughout the substrate
panel ranging from C18:2 to C14:1 and displayed together with
Gm-OAH1 the highest conversion for the short-chain substrate (Z)-
undec-9-enoic acid (C11:1). Whereas Gm-OAH1 was able to form
16 % of the corresponding 10-hydroxyundecanoic acid, Em-OAH1
and Cg-OAH1 were able to produce almost the double amount (23
% and 30 %).
3.4. Optimization of reaction conditions
The reported pH-optima for most of the oleate hydratases is in
the range from pH 5.5–6.8[38–40]. Therefore we used potassium
phosphate buffer pH 6.5 for our initial tests. However, in addi-
tion to the pH, enzyme stability becomes increasingly important
for the biotransformation of non-natural substrates with prolonged
reaction time and higher protein concentration. It was not possi-
ble under standard conditions with potassium phosphate buffer
to monitor any oleate hydratase reaction longer than two days.
Blurring of the reaction mixture after 1 d indicated the begin-
ning denaturation of the enzymes from the cell lysate leading to
the exposure of hydrophobic areas and the subsequent interaction
with substrate and product. As a result, neither substrate nor prod-
uct were detectable in the extract of the reaction mixture. Hence,
we compared the product formation for the biotransformation of
myristoleic acid (C14:1) over 1 d and 2 d in four different buffer sys-
tems at pH 6.0 (potassium phosphate, citrate, BIS-TRIS, MES) using
Em-OAH1. The experiments showed that the product recovery in
the 2 d sample was less than in the 1 d sample for potassium phos-
phate and BIS-TRIS, while more product was isolated after 2 d for
citrate and MES buffer. Particularly, the obtained product decreased
for potassium phosphate from 100 % to 58 % and for BIS-TRIS from
105 % to 78 %, whereas increased from 122 % to 134 % and 114
% to 148 % for citrate and MES buffer, respectively. The percent-
ages are all normalized to the product formation after 1 d in 50 mM
potassium phosphate buffer pH 6.0.
From the tested buffer systems, we chose 50 mM citrate buffer
pH 6.0 for higher stability of oleate hydratases at prolonged reaction
times and better reproducibility of long-term experiments.
3.5. Substrate scope analysis
Despite their activity on 9Z-double bonds, the homologous
enzymes were also characterized on their capability to hydrate
a terminal double bond as in dec-9-enoic acid (C10:1). However,
none of the homologs was active towards this substrate.
Cg-OAH1 and Em-OAH1, both found in HFam11, could be iden-
tified as enzymes with the highest activity on all substrates. The
The so far investigated substrate scope described in litera-
ture and the applied reaction conditions often differ for each
individual homolog, which makes it difficult to compare the differ-
ent enzymes. Furthermore, different enzyme preparations ranging
Please cite this article in press as: J. Schmid, et al., Biocatalytic study of novel oleate hydratases, J. Mol. Catal. B: Enzym. (2017),
http://dx.doi.org/10.1016/j.molcatb.2017.01.010

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6
Table 1
Substrate scope analysis of putative oleate hydratases. Product formation [%], reaction time and protein concentration of biotransformations.
Product formation is indicated as follows: no highlighting (< 20 %), light grey (20–34 %), grey (35–74 %), dark grey (> 75 %). Reaction times (treaction) and protein concentrations
(b)
(c)
(d)
(e/f)
(cprotein): (a) 4.5 h, 0.5 g L−1
,
5 min, 0.15 g L−1
,
30 min, 0.5 g L−1
,
21 h, 2 g L−1
,
5 d, 4 g L−1
.
enzymes of HFam2 on the other hand showed only high yields
for fatty acids of medium chain length (C14 and C16) under the
tested conditions. Therefore, enzymes of HFam11 are particularly
interesting for further investigations and for the applications as
hydration catalyst for short-chain fatty acids and alkenes.
tremendous support and discussions throughout the writing pro-
cess of this publication.
Appendix A. Supplementary data
Supplementary data associated with this article can be found, in
the online version, at http://dx.doi.org/10.1016/j.molcatb.2017.01.
010.
4. Conclusion
In this study, we established the publicly available Hydratase
Engineering Database (HyED) to serve as navigation tool and to col-
lect information about this interesting protein family. Particularly,
we examined the oleate hydratase family around the enzymes from
Elizabethkingia meningoseptica and Lactobacillus acidophilus in fur-
ther detail and used the HyED for the investigation of conserved
sequence motifs to gain further insights into the molecular base of
these enzymes.
References
[1] V. Resch, U. Hanefeld, The selective addition of water, Catal. Sci. Technol. 5
(2015) 1385–1399, http://dx.doi.org/10.1039/C4CY00692E.
[2] B.N.M. van Leeuwen, A.M. van der Wulp, I. Duijnstee, A.J.A. van Maris, A.J.J.
Straathof, Fermentative production of isobutene, Appl. Microbiol. Biotechnol.
93 (2012) 1377–1387, http://dx.doi.org/10.1007/s00253-011-3853-7.
[3] J. Jin, U. Hanefeld, The selective addition of water to CC bonds; enzymes are
the best chemists, Chem. Commun. (Camb.) 47 (2011) 2502–2510, http://dx.
doi.org/10.1039/c0cc04153j.
Furthermore, we have chosen nine homologous oleate
hydratases for detailed investigations and heterologously
expressed them in E. coli. Five homologous genes were suitable
for clear and stable overexpression. The enzymes were analyzed
with a set of substrates, ranging from long-chain to medium-chain
fatty acids. For the first time, we have tested two novel substrates
(C11:1 and C10:1) and activity could be demonstrated towards
(Z)-undec-9-enoic acid (C11:1). All actively expressed hydratases
can be classified for their comparable substrate scope as oleate
hydratases.
The oleate hydratases from HFam11, Em-OAH1 and Cg-OAH1,
showed the highest stability as well as activity towards the novel
non-natural shorter chained substrates. This makes them ideal can-
didates for further mutagenesis experiments with the goal to evolve
a hydration biocatalyst with improved properties. We believe that
using the inherent hydration potential of oleate hydratases can
lead to new chemistry and, beyond, can provide an alternative to
chemical synthesis, not only for hydroxy fatty acids but also for the
formation of chiral alcohols starting from alkenes/double bonds.
[4] A. Hiseni, I.W.C.E. Arends, L.G. Otten, New cofactor-independent hydration
biocatalysts: structural, biochemical, and biocatalytic characteristics of
carotenoid and oleate hydratases, ChemCatChem 7 (2015) 29–37, http://dx.
doi.org/10.1002/cctc.201402511.
[5] Z. Sun, S. Shen, C. Wang, H. Wang, Y. Hu, J. Jiao, T. Ma, B. Tian, Y. Hua, A novel
carotenoid 1 2-hydratase (CruF) from two species of the non-photosynthetic
bacterium Deinococcus, Microbiology 155 (2009) 2775–2783, http://dx.doi.
org/10.1099/mic.0.027623-0.
[6] S. Steiger, A. Mazet, G. Sandmann, Heterologous expression, purification, and
enzymatic characterization of the acyclic carotenoid 1,2-hydratase from
Rubrivivax gelatinosus, Arch. Biochem. Biophys. 414 (2003) 51–58, http://dx.
doi.org/10.1016/S0003-9861(03)00099-7.
[7] D. Brodkorb, M. Gottschall, R. Marmulla, F. Lüddeke, J. Harder, Linalool
dehydratase-isomerase, a bifunctional enzyme in the anaerobic degradation
of monoterpenes, J. Biol. Chem. 285 (2010) 30436–30442, http://dx.doi.org/
10.1074/jbc.M109.084244.
[8] L.E. Bevers, M.W.H. Pinkse, P.D.E.M. Verhaert, W.R. Hagen, Oleate hydratase
catalyzes the hydration of a nonactivated carbon–carbon bond, J. Bacteriol.
191 (2009) 5010–5012, http://dx.doi.org/10.1128/J.B.00306-09.
[9] L.L. Wallen, R.G. Benedict, R.W. Jackson, The microbiological production of
10-Hydroxystearic acid from oleic acid, Arch. Biochem. Biophys. 99 (1962)
249–253, http://dx.doi.org/10.1016/0003-9861(62)90006-1.
[10] S. Koritala, C.T. Hou, C.W. Hesseltine, M.O. Bagby, Microbial conversion of
oleic acid to 10-hydroxystearic acid, Appl. Microbiol. Biotechnol. 32 (1989)
299–304, http://dx.doi.org/10.1007/BF00184978.
[11] G.J. Schroepfer, K. Bloch, The stereospecific conversion of stearic acid to oleic
acid, J. Biol. Chem. 240 (1965) 54–63.
[12] W.G. Niehaus, A. Torkelson, A. Kisic, D.J. Bednarczyk, G.J. Schroepfer,
Stereospecific hydration of the delta-9 double bond of oleic acid, J. Biol. Chem.
245 (1970) 3790–3797.
Acknowledgements
[13] W.G. Niehaus, G.J. Schroepfer, The reversible hydration of oleic acid to
10D-hydroxystearic acid, Biochem. Biophys. Res. Commun. 21 (1965)
271–275, http://dx.doi.org/10.1016/0006-291X(65)90282-2.
[14] Y.-C. Joo, K.-W. Jeong, S.-J. Yeom, Y.-S. Kim, Y. Kim, D.-K. Oh, Biochemical
characterization and FAD-binding analysis of oleate hydratase from
Macrococcus caseolyticus, Biochimie 94 (2012) 907–915, http://dx.doi.org/10.
1016/j.biochi.2011.12.011.
The research leading to these results has received funding
from the Ministry for Science, Research and the Arts of Baden-
Württemberg, Germany [MWK; Baden-Württemberg Biosynthesis
netWork (BW2)]. Furthermore we want to thank Constantin Vogel
for the initial work on the hydratase engineering database and
Linda Otten (TU Delft) for helpful discussions. Moreover we
would like to give our special thanks to Bettina Nestl for her
[15] J.-Y. Park, S.-H. Lee, K.-R. Kim, J.-B. Park, D.-K. Oh, Production of
13S-hydroxy-9(Z)-octadecenoic acid from linoleic acid by whole recombinant
Please cite this article in press as: J. Schmid, et al., Biocatalytic study of novel oleate hydratases, J. Mol. Catal. B: Enzym. (2017),
http://dx.doi.org/10.1016/j.molcatb.2017.01.010

G Model
MOLCAB-3509; No. of Pages7
ARTICLE IN PRESS
J. Schmid et al. / Journal of Molecular Catalysis B: Enzymatic xxx (2017) xxx–xxx
7
cells expressing linoleate 13-hydratase from Lactobacillus acidophilus, J.
Biotechnol. 208 (2015) 1–10, http://dx.doi.org/10.1016/j.jbiotec.2015.05.006.
[16] A. Kisic, Y. Miura, G.J. Schroepfer, Oleate hydratase: studies of substrate
specificity, Lipids 6 (1971) 541–545, http://dx.doi.org/10.1007/BF02531133.
[17] T. Kaneshiro, J.-K. Huang, D. Weisleder, M. Bagby, 10(R)-Hydroxystearic acid
production by a novel microbe, NRRL B-14797, isolated from compost, J. Ind.
Microbiol. 13 (1994) 351–355, http://dx.doi.org/10.1007/BF01577218.
[18] A. Volkov, S. Khoshnevis, P. Neumann, C. Herrfurth, D. Wohlwend, R. Ficner, I.
Feussner, Crystal structure analysis of a fatty acid double-bond hydratase
from Lactobacillus acidophilus, Acta Crystallogr. D. Biol. Crystallogr. 69 (2013)
648–657, http://dx.doi.org/10.1107/S0907444913000991.
models of biomolecular interaction networks, Genome Res. 13 (2003)
2498–2504, http://dx.doi.org/10.1101/gr.1239303.
[31] H.M. Berman, The protein data bank, Nucleic Acids Res. 28 (2000) 235–242,
http://dx.doi.org/10.1093/nar/28.1.235.
[32] F. Sievers, A. Wilm, D. Dineen, T.J. Gibson, K. Karplus, W. Li, R. Lopez, H.
McWilliam, M. Remmert, J. Soding, J.D. Thompson, D.G. Higgins, Fast, scalable
generation of high-quality protein multiple sequence alignments using Clustal
Omega, Mol Syst. Biol. 7 (2014) 539, http://dx.doi.org/10.1038/msb.2011.75.
[33] U.K. Laemmli, Cleavage of structural proteins during the assembly of the head
of bacteriophage T4, Nature 227 (1970) 680–685, http://dx.doi.org/10.1038/
227680a0.
[19] M. Engleder, T. Pavkov-Keller, A. Emmerstorfer, A. Hromic, S. Schrempf, G.
Steinkellner, T. Wriessnegger, E. Leitner, G.A. Strohmeier, I. Kaluzna, D. Mink,
M. Schürmann, S. Wallner, P. Macheroux, K. Gruber, H. Pichler,
Structure-based mechanism of oleate hydratase from Elizabethkingia
meningoseptica, Chembiochem 16 (2015) 1730–1734, http://dx.doi.org/10.
1002/cbic.201500269.
[20] F.C. Naughton, Production, chemistry, and commercial applications of various
chemicals from castor oil, J. Am. Oil Chem. Soc. 51 (1974) 65–71, http://dx.doi.
org/10.1007/BF00000015.
[21] Y. Wache, M. Aguedo, A. Choquet, I.L. Gatfield, J.-M. Nicaud, J.-M. Belin, Role of
beta-Oxidation Enzymes in gamma-Decalactone Production by the Yeast
Yarrowia lipolytica, Appl. Environ. Microbiol. 67 (2001) 5700–5704, http://dx.
doi.org/10.1128/AEM.67.12.5700-5704.2001.
[22] J.W. Song, E.Y. Jeon, D.H. Song, H.Y. Jang, U.T. Bornscheuer, D.K. Oh, J.B. Park,
Multistep enzymatic synthesis of long-chain ␣, ␻-dicarboxylic and
␻-hydroxycarboxylic acids from renewable fatty acids and plant oils, Angew.
Chem. Int. Ed. 52 (2013) 2534–2537, http://dx.doi.org/10.1002/anie.
201209187.
[34] F. Forneris, R. Orru, D. Bonivento, L.R. Chiarelli, A. Mattevi, Thermo FAD, a
Thermofluor ® −adapted flavin ad hoc detection system for protein folding
and ligand binding, FEBS J. 276 (2009) 2833–2840, http://dx.doi.org/10.1111/
j.1742-4658.2009.07006.x.
[35] S. Reich, N. Kress, B.M. Nestl, B. Hauer, Variations in the stability of NCR ene
reductase by rational enzyme loop modulation, J. Struct. Biol. 185 (2014)
228–233, http://dx.doi.org/10.1016/j.jsb.2013.04.004.
[36] G. Kleiger, D. Eisenberg, GXXXG and GXXXA Motifs Stabilize FAD and
NAD(P)-binding Rossmann Folds Through C␣-H-O Hydrogen Bonds and van
der Waals Interactions, J. Mol. Biol. 323 (2002) 69–76, http://dx.doi.org/10.
1016/S0022-2836(02)00885-9.
[37] R.K. Wierenga, P. Terpstra, W.G. Hol, Prediction of the occurrence of the
ADP-binding beta alpha beta-fold in proteins, using an amino acid sequence
fingerprint, J. Mol. Biol. 187 (1986) 101–107, http://dx.doi.org/10.1016/0022-
2836(86)90409-2.
[38] M.-H. Seo, K.-R. Kim, D.-K. Oh, Production of a novel compound
10,12-dihydroxystearic acid from ricinoleic acid by an oleate hydratase from
Lysinibacillus fusiformis, Appl. Microbiol. Biotechnol. 97 (2013) 8987–8995,
http://dx.doi.org/10.1007/s00253-013-4728-x.
[39] M. Takeuchi, S. Kishino, A. Hirata, S.-B. Park, N. Kitamura, J. Ogawa,
Characterization of the linoleic acid ꢀ9 hydratase catalyzing the first step of
polyunsaturated fatty acid saturation metabolism in Lactobacillus plantarum
AKU, J. Biosci. Bioeng. 119 (2015) (2015) 636–641, http://dx.doi.org/10.1016/j.
jbiosc.2014.10.022.
[23] J.W. Song, J.H. Lee, U.T. Bornscheuer, J.B. Park, Microbial synthesis of
medium-chain ␣, ␻-dicarboxylic acids and ␻-aminocarboxylic acids from
renewable long-chain fatty acids, Adv. Synth. Catal. 356 (2014) 1782–1788,
http://dx.doi.org/10.1002/adsc.201300784.
[24] P. Marliere, Method for Producing an Alkene Comprising the Step of
Converting an Alcohol by an Enzymatic Dehydration Step,
WO2011/076691A1/EP2336341A1, (2011).
[40] C.T. Hou, Production of hydroxy fatty acids from unsaturated fatty acids by
flavobacterium sp. DS5 hydratase, a C-10 positional- and cis
unsaturation-specific enzyme, J. Am. Oil Chem. Soc. 72 (1995) 1265–1270,
http://dx.doi.org/10.1007/BF02546197.
[41] M.H. Kim, M.S. Park, C.H. Chung, C.T. Kim, Y.S. Kim, K.H. Kyung, Conversion of
unsaturated food fatty acids into hydroxy fatty acids by lactic acid bacteria, J.
Microbiol. Biotechnol. 13 (2003) 360–365.
[42] I.-S. Yu, H.-J. Kim, D.-K. Oh, Conversion of linoleic acid into
10-Hydroxy-12(Z)-octadecenoic acid by whole cells of stenotrophomonas
nitritireducens, Biotechnol. Prog. 24 (2008) 182–186, http://dx.doi.org/10.
1021/bp0703073.
[43] W.D. Nunn, R.W. Simons, Transport of long-chain fatty acids by Escherichia
coli: mapping and characterization of mutants in the fadL gene, Proc. Natl.
Acad. Sci. 75 (1978) 3377–3381.
[25] P. Buchholz, C. Vogel, W. Reusch, M. Pohl, D. Rother, A. Spieß, J. Pleiss,
BioCatNet: a database system for the integration of enzyme sequences and
biocatalytic experiments, Chembiochem (2016), http://dx.doi.org/10.1002/
cbic.201600462.
[26] S.F. Altschul, T.L. Madden, A.A. Schäffer, J. Zhang, Z. Zhang, W. Miller, D.J.
Lipman, Gapped BLAST and PSI-BLAST: a new generation of protein database
search programs, Nucleic Acids Res. 25 (1997) 3389–3402, http://dx.doi.org/
10.1093/nar/25.17.3389.
[27] D.A. Benson, M. Cavanaugh, K. Clark, I. Karsch-Mizrachi, D.J. Lipman, J. Ostell,
E.W. Sayers, GenBank. Nucleic Acids Res. 41 (2013) D36–D42, http://dx.doi.
org/10.1093/nar/gks1195.
[28] R.C. Edgar, Search and clustering orders of magnitude faster than BLAST,
Bioinformatics 26 (2010) 2460–2461, http://dx.doi.org/10.1093/
bioinformatics/btq461.
[44] J.H. Bae, B.G. Park, E. Jung, P.-G. Lee, B.-G. Kim, fadD deletion and fadL
overexpression in Escherichia coli increase hydroxy long-chain fatty acid
productivity, Appl. Microbiol. Biotechnol. 98 (2014) 8917–8925, http://dx.doi.
org/10.1007/s00253-014-5974-2.
[29] S.B. Needleman, C.D. Wunsch, A general method applicable to the search for
similarities in the amino acid sequence of two proteins, J. Mol. Biol. 48 (1970)
443–453, http://dx.doi.org/10.1016/0022-2836(70)90057-4.
[30] P. Shannon, A. Markiel, O. Ozier, N.S. Baliga, J.T. Wang, D. Ramage, N. Amin, B.
Schwikowski, T. Ideker, Cytoscape: a software environment for integrated
Please cite this article in press as: J. Schmid, et al., Biocatalytic study of novel oleate hydratases, J. Mol. Catal. B: Enzym. (2017),
http://dx.doi.org/10.1016/j.molcatb.2017.01.010