Optimized Reaction Conditions Enable the Hydration of Non-natural Substrates by the Oleate Hydratase from Elizabethkingia meningoseptica

Article abstract of DOI:10.1002/cctc.201601329

The oleate hydratase from Elizabethkingia meningoseptica (Em-OAH) catalyzes the hydration of oleic acid (C18) to (R)-10-hydroxystearic acid. In previous work, low activity of Em-OAH towards chemically synthesized (Z)-undec-9-enoic acid (C11) was observed. Product formation in the hydration of the truncated C11 substrate was improved by optimizing the reaction conditions by applying statistical experiment design. Optimized reaction conditions resulted in a 2.8-fold increase in product formation in just one quarter of the time (64 % conversion in 28 h). The applicability has been assessed in the upscaling of the conversion of (Z)-undec-9-enoic acid to (S)-10-hydroxyundecanoic acid (132 mg product, >95 % purity). Reaction conditions developed for the hydration of C11 facilitated the first hydration of non-natural alkenes. By using a fatty acid dummy substrate, 1-decene was successfully hydrated to (S)-2-decanol with excellent stereoselectivity and 50 % conversion after four days of incubation.

Full text of DOI:10.1002/cctc.201601329

DOI: 10.1002/cctc.201601329
Full Papers
Optimized Reaction Conditions Enable the Hydration of
Non-natural Substrates by the Oleate Hydratase from
Elizabethkingia meningoseptica
Rebecca M. Demming, Konrad B. Otte, Bettina M. Nestl, and Bernhard Hauer*[a]
The oleate hydratase from Elizabethkingia meningoseptica (Em-
OAH) catalyzes the hydration of oleic acid (C18) to (R)-10-hy-
droxystearic acid. In previous work, low activity of Em-OAH to-
wards chemically synthesized (Z)-undec-9-enoic acid (C11) was
observed. Product formation in the hydration of the truncated
C11 substrate was improved by optimizing the reaction condi-
tions by applying statistical experiment design. Optimized re-
action conditions resulted in a 2.8-fold increase in product for-
mation in just one quarter of the time (64% conversion in
28 h). The applicability has been assessed in the upscaling of
the conversion of (Z)-undec-9-enoic acid to (S)-10-hydroxyun-
decanoic acid (132 mg product, >95% purity). Reaction condi-
tions developed for the hydration of C11 facilitated the first
hydration of non-natural alkenes. By using a fatty acid dummy
substrate, 1-decene was successfully hydrated to (S)-2-decanol
with excellent stereoselectivity and 50% conversion after four
days of incubation.
Introduction
Chiral medium-chain hydroxy fatty acids and alcohols are
highly interesting and valuable synthons for the preparation of
enantiomerically pure natural products, pharmaceuticals, and
fine chemicals.[1–5] Moreover, hydroxy fatty acids exhibit great
potential as precursors in polymer chemistry and for the syn-
thesis of cyclic lactones used as antibiotics and flavor com-
pounds in the food industry.[4,6,7] The selective addition of
water to carbon–carbon double bonds is an ongoing challenge
in synthetic organic chemistry. Although acid-based chemical
hydration is poorly selective and can only be applied for mole-
cules that cannot undergo alternative rearrangements, the en-
zymatic addition of water can be achieved with high selectivity
under mild reaction conditions.[8,9] In nature, hydrolyases
(EC 4.2.) catalyze the addition of water at isolated and conju-
gated carbon–carbon double bonds (a,b-unsaturated carbon-
yls) with the first following the rule of Markovnikov
(Scheme 1).[8,10] Recently, oleate hydratases (OAH, EC 4.2.1.53)
catalyzing the cofactor-independent regio- and stereoselective
hydration of oleic acid to (R)-10-hydroxyoctadecanoic acid with
no net redox change involved in catalysis were de-
scribed.[9,11–13] In this context, oleate hydratases are highly inter-
esting biocatalysts for the large-scale production of 10-hydrox-
ystearic acid[14–16] and the synthesis of a,w-dicarboxylic acids
such as 1,10-decanedioic acid in enzyme cascades.[17,18]
The conversion of oleate to (R)-10-hydroxyoctadecanoic acid
was first described in 1962 by Wallen et al., whereas the re-
sponsible enzyme from Elizabethkingia meningoseptica was
only identified and biochemically characterized in 2009.[19] To
date, various bacterial putative hydratases[20,21] have been re-
combinantly expressed, characterized, and identified as oleic
acid hydratases (OAH) or general fatty acid hydratas-
es.[11,13,14,21–23] All tested enzymes only gave 10-hydroxy fatty
acids as products, whereas a hydratase from Macrococcus case-
olyticus introduced a second water molecule on incubation
with linoleic, a-linoleic, and g-linoleic acid.[9,13] Moreover, two
novel regioselective hydratases from Lactobacillus acidophilus
were identified, which convert cis-12 unsaturated fatty acids to
the corresponding (S)-13-hydroxy products[24] and longer poly-
unsaturated fatty acids (PUFAs; C16–C22), resulting in the for-
mation of 12-, 13-, 14-, and 15-hydroxy fatty acids, respective-
ly.[25] The best characterized hydratase is the oleate hydratase
from E. meningoseptica (Em-OAH, GQ144652). The protein ex-
hibits a homodimeric conformational structure with each
73 kDa subunit containing a catalytically non-essential calcium
ion.[12,19] Kinetic studies of recombinant Em-OAH with oleic acid
revealed a maximum activity at pH 6 and an optimal salt con-
centration of 50 mm NaCl.[19] Lately, Engleder et al. solved the
crystal structure of Em-OAH with oleic acid bound in the V-
shaped substrate binding pocket and the carbon–carbon
double bond located at the bend of the predominantly hydro-
phobic cavity near to the active site residues. They further pro-
Scheme 1. Water addition to isolated carbon–carbon double bonds by hy-
drolyases follows Markovnikov’s rule.
[a] R. M. Demming, Dr. K. B. Otte, Dr. B. M. Nestl, Prof. Dr. B. Hauer
Institute of Technical Biochemistry
Universitꢀt Stuttgart
Allmandring 31, 70569 Stuttgart (Germany)
E-mail: bernhard.hauer@itb.uni-stuttgart.de
Supporting information for this article can be found under:
http://dx.doi.org/10.1002/cctc.201601329.
ChemCatChem 2017, 9, 1 – 10
1
ꢀ 2017 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
&
These are not the final page numbers! ÞÞ

Full Papers
posed a reaction mechanism for the hydration of oleic acid
with the formation of a carbocation intermediate at the C10
atom of oleic acid followed by a nucleophilic attack of a gluta-
mic acid-activated water molecule.[12] Moreover, investigations
on various OAHs indicated that the substrate carboxyl group is
important for substrate binding by positively charged amino
acids.[9,12,20] Interestingly, hydratases including Em-OAH do con-
tain a flavin cofactor (FAD/FMN), the function of which is still
unknown.[9,11–14,21–23]
The study of five wildtype oleate hydratases from Chryseo-
bacterium gleum, Desulfomicrobium baculatum, E. meningosepti-
ca, Gemella morbillorum, and Lactobacillus acidophilus showed
that hydratases from E. meningoseptica and C. gleum were able
to hydrate chemically synthesized (Z)-undec-9-enoic acid (C11)
with the highest product yield.[26] However, no conversion was
obtained by employing the smaller C10 fatty acid dec-9-enoic
acid, which possesses a terminal carbon–carbon double bond.
Owing to its reliable heterologous gene expression in E. coli
and structural and biochemical characterization, Em-OAH was
chosen as a promising candidate for the conversion of non-
natural short-chain substrates. The aim of this study was the
improvement of the reaction conditions regarding the hydra-
tion of the non-natural substrate (Z)-undec-9-enoic acid by Em-
OAH. For the optimization of the reaction conditions including
catalyst type, cofactor, and reaction temperature, we applied
the MODDE software for statistical experiment design. We fur-
ther report the first hydration of the smaller short-chain alkene
1-decene by wildtype Em-OAH. We established a novel efficient
methodology for the selective formation of (S)-2-decanol by
using hexanoic acid as a dummy substrate. The successful
usage of dummy substrates (also decoy molecules) has already
been extensively reported in P450-catalyzed hydroxylations of
gaseous alkanes.[27–31]
Figure 1. Conversion of (Z)-undec-9-enoic acid (0.5 mm) by Em-OAH under
standard (stand.) and FAD-reducing (FAD-red.) conditions by applying E. coli
whole cells (50 mgmLÀ1) and cell lysate (4 mgmLÀ1). The results of cell
lysate standard reactions were set as 100%.
observed an enhanced activity of the enzyme towards oleic
acid with FADH2 instead of oxidized cofactor,[12] reducing con-
ditions were applied and compared with the standard reaction
conditions (citrate buffer, glucose was added for whole cells).
Results of the biotransformations using cell lysate under stan-
dard conditions were set as 100% (Figure 1). Product forma-
tion was strongly enhanced by the addition of flavin adenine
dinucleotide (FAD), dithiothreitol (DTT), and NADH (FAD-reduc-
ing conditions) to both cell lysate and whole cell biotransfor-
mations with a 2- and 4-fold increase in product formation, re-
spectively. No background hydration was observed. The best
results were obtained with whole cells under FAD-reducing
conditions with 78% product formation after five days of incu-
bation; a 3.4-fold increase compared with previous results
giving 23%.[26] Moreover, recovery rates were improved by ap-
plying whole cells as long incubation times resulted in an ag-
glomeration of substrate, product, and denatured protein,
forming a white precipitate in cell lysate.
Results and Discussion
Whole cell biotransformations versus lysate assays
The characterization of several wildtype hydratases demon-
strated that oleate hydratase from Elizabethkingia meningosep-
tica (Em-OAH) is a promising candidate for the conversion of
short-chain fatty acids.[26] Although a minimum chain length of
14 carbon atoms is described to be essential for fatty acid con-
version by hydratases,[9] we observed recently the hydration of
C11 (Z)-undec-9-enoic acid (Figure 1).[26] However, the conver-
sion was low with 23% product formation after five days of in-
cubation using 4 mgmLÀ1 E. coli lysate. In contrast to that,
90% of the natural substrate oleic acid was converted within
five minutes with 0.15 mgmLÀ1 cell lysate.
Optimization of reaction conditions by statistical experi-
ment design
Reaction conditions were further improved by using MODDE
as a tool for statistical experiment design.[32–36] MODDE enables
the simultaneous analysis of several factors to identify critical
parameters, synergistic effects, and suitable operating condi-
tions by statistical evaluation.
Owing to the fact that previous studies revealed a low sta-
bility of Em-OAH cell lysate at longer incubation times,[18,26] we
first compared transformations using E. coli whole cells with
cell lysate to improve the conversion of the non-natural short-
chain (Z)-undec-9-enoic acid substrate. In addition to higher
operational stability, whole cell biotransformations are easy to
handle and avoid time-consuming steps such as cell disruption
and protein purification. Owing to the fact that Engleder et al.
In previous studies, parameters influencing hydratase activity
and product yield were identified.[11,13,14,19,21,23–25,37,38] Up to five
conditions including glucose concentration, cofactor addition
(NADH, FAD), reducing agents (DTT, glutathione), salt concen-
tration, enzyme stabilization, and incubation temperature were
analyzed simultaneously in four rounds of optimization with
biotransformation setups designed by MODDE. For more de-
&
ChemCatChem 2017, 9, 1 – 10
www.chemcatchem.org
2
ꢀ 2017 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
ÝÝ These are not the final page numbers!

Full Papers
tailed information, see the Experimental Section. Although glu-
tathione and bovine serum albumin (BSA) showed no effect on
substrate conversion, product formation was decreased by the
addition of NaCl (data not shown). However, improved hydra-
tase activity was obtained by the addition of glucose, FAD,
NADH, or DTT, and an increased incubation temperature (data
not shown). In a final experiment, these parameters were ana-
lyzed consecutively by applying the concentrations optimized
by MODDE (Figure 2). The addition of glucose led to a 7-fold
and S2 in the Supporting Information). This is a 2.8-fold in-
crease in conversion in just one quarter of the time compared
with the starting point of 23% product formation after five
days of incubation.[26]
Upscaling of (Z)-undec-9-enoic acid conversion
The improved conditions were subsequently used for an up-
scaling of 10-hydroxyundecanoic acid production. E. coli BL21
(DE3) pET28a(+)/Em-OAH was grown in a 5 L fed-batch cultiva-
tion for biomass production. Heterologous overexpression was
analyzed by sodium dodecyl sulfate–polyacrylamide gel elec-
trophoresis (SDS-PAGE) and enzyme activity towards oleic acid
and (Z)-undec-9-enoic acid was confirmed. The upscaling was
run in a total volume of about 1 L with 200 mg fatty acid sub-
strate (1% final DMSO concentration), 50 mgmLÀ1 biomass,
100 mm glucose, 0.3 mm FAD, and 3 mm NADH. After three
days of incubation at 308C and 180 rpm, the reaction was
stopped by acidification (about 90% conversion, data not
shown). The product was purified by column chromatography,
the product purity was analyzed by GC-MS and the hydration
was confirmed by NMR analysis (Figures S3–S6 in the Support-
ing Information). The isolated product was subsequently used
to examine the reaction equilibrium in the hydration of (Z)-
undec-9-enoic acid by Em-OAH, which was 99:1 in favor of the
hydrated product. A reaction equilibrium of 85:15 in favor of
the hydrated product was described for oleate hydration by
Em-OAH.[45]
Upscaling of (Z)-undec-9-enoic acid hydration by Em-OAH
under optimized conditions resulted in 132 mg of a light-
brown powder as the product with >95% purity of 10-hydrox-
yundecanoic acid (YP/S=66%, YP/B =0.0024). The stereoselectiv-
ity of (Z)-undec-9-enoic acid hydration by Em-OAH was exam-
ined by Mosher ester analysis (Table S1, Figure S7 in the Sup-
Figure 2. Optimization of reaction conditions for the conversion of 0.5 mm
(Z)-undec-9-enoic acid by Em-OAH (50 mgmLÀ1 biomass). Analysis of the re-
action conditions by MODDE resulted in optimized concentrations of glu-
cose (Glu), FAD, NADH, and DTT for the hydration of (Z)-undec-9-enoic acid
(C11). In a final experiment, each step was built upon the previous optimiza-
tion step (marked by arrows). Finally, the implementation of glucose, FAD,
NADH, and DTT was elucidated at 308C reaction temperature.
1
porting Information). By evaluating the shift in H NMR signals
and by applying the Cahn–Ingold–Prelog (CIP) priority rules,
the biotransformation product was identified as (S)-10-hydrox-
yundecanoic acid.
increase in 10-hydroxyundecanoic acid formation and a maxi-
mum recovery rate. Glucose turned out to be essential for the
whole cell system, as without the additional carbon source the
fatty acid substrate was not fully recovered. The loss of sub-
strate can be explained by the metabolic activity of the cells,
which might result in the b-oxidation of the fatty acid.[39–44]
Conversion was further enhanced to 50% by adding FAD and
NADH. Despite the fact that the incorporation of DTT did not
promote product formation, it might be of great significance
for long-time incubations for maintaining reducing conditions
in the presence of oxygen. Enzyme activity was further im-
proved by applying an incubation temperature of 308C. At
higher temperatures, for example, 378C, product formation
and substrate recovery were decreased, most probably due to
enhanced metabolic activity of the whole cells and the instabil-
ity of the oleate hydratase (data not shown). E. coli pET28a(+)
whole cells incubated with glucose, FAD, NADH, DTT, and
0.5 mm substrate served as a negative control. No background
product formation was observed.
Hydration of 1-decene by using the optimized reaction
conditions
The hydration of the truncated C11 fatty acid substrate by Em-
OAH was greatly improved by the optimization of the reaction
conditions. Intrigued by these results, we utilized the opti-
mized conditions in the hydration of smaller C10 substrates.
The non-natural fatty acid dec-9-enoic acid, previously identi-
fied as a non-substrate, and the alkene 1-decene were tested
as substrates in Em-OAH-catalyzed biotransformations.
To enable the enzymatic hydration of alkenes, we compared
the potential binding of fatty acids and alkenes in the active
site of Em-OAH. Figure 3 shows a schematic view of the V-
shaped substrate binding pocket of Em-OAH with bound oleic
acid. The carbon–carbon double bond is located at the bend
of the predominantly hydrophobic cavity near to the active
site residues.[12] Previous studies of various OAHs illustrated
that substrate conversion is dependent on a carboxyl group
The optimization of reaction conditions by MODDE resulted
in 64% product formation after 28 h of incubation (Figures S1
ChemCatChem 2017, 9, 1 – 10
www.chemcatchem.org
3
ꢀ 2017 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
&
These are not the final page numbers! ÞÞ

Full Papers
Figure 3. Strategies for the conversion of short-chain molecules by Em-OAH. Schematic view of the substrate binding pocket with oleic acid bound in the
active site (middle top). While the conversion of (Z)-undec-9-enoic acid was successful (left), no hydration of dec-9-enoic acid was observed (middle). There-
fore, a second strategy for the conversion of non-natural substrates was pursued by using a small short-chain alkene bound in the alkyl substrate binding
pocket (right). A short-chain fatty acid should serve as dummy substrate for substrate recognition bound in the carboxy substrate binding pocket of the
enzyme. The theoretical position of the formed carbocation during the hydration reaction is indicated by red pluses.
for substrate recognition by positively charged amino acids at
the entrance of the substrate binding pocket (indicated as B+
).[9] Based on this assumption and the hydrophobicity of the
cavity, we assumed that non-natural (Z)-undec-9-eoic acid and
dec-9-enoic acid are bound in the carboxy binding pocket in
a way similar to that of the natural substrate oleic acid. More-
over, we assumed that the non-polar molecule 1-decene is
bound in the alkyl substrate binding pocket, filling the space
that is normally occupied by the C9 to C18 atoms of the natu-
ral substrate oleic acid. Consequently, if 1-decene could pene-
trate into the active site of Em-OAH in the presence of
a dummy substrate, the structure of which is similar to fatty
acids, the short-chain alkene could be hydrated.
we showed for the first time the hydration of 1-decene by Em-
OAH. Reports from Engleder et al. indicate that the hydration
of oleic acid proceeds via a secondary carbocation at the C10
atom (the theoretical position of the formed carbocation is in-
dicated by a red plus).[12] The hydration of dec-9-enoic acid
possessing a terminal carbon–carbon double bond implies the
formation of a primary carbocation. The lack of the methyl
group on the omega side of the carbon–carbon double bond
might prevent the reaction,[46] by reason of the low stability of
the formed carbocation lacking inductive effects. As a result of
the proper alkene positioning in the alkyl substrate binding
pocket, the hydration of 1-decene proceeds via a secondary
carbocation as is the case for the natural substrate oleic acid
and the truncated C11 fatty acid. To examine the importance
of the dummy substrate for the hydration of alkenes, 2-dec-
anol formation was analyzed with and without the addition of
hexanoic acid (Figure S8 in the Supporting Information). Prod-
uct formation with the addition of hexanoic acid was set as
100%. The relative conversion was significantly decreased
without the dummy substrate (<7%).
Owing to the apparent dependency of Em-OAH activity on
a carboxylic group for substrate recognition, hexanoic acid was
selected as a dummy substrate for 1-decene hydration. In addi-
tion to the required carboxyl group, it has a suitable chain
length that allows the simultaneous binding of two molecules
within the large cavity. Longer fatty acids might lead to steric
hindrances within the substrate binding pocket and shorter
dummy substrates might be too small to position the alkene
for effective hydration. Furthermore, the utilization of
a dummy substrate might support the organization of the
active site amino acid residues and, thus, activation of the hy-
dratase for catalysis.
Em-OAH-catalyzed conversion of 1-decene was performed in
whole cell biotransformations under the improved conditions
(Figure 2) at 25 and 308C. About 50% of the substrate (0.5 mm
1-decene) was converted to 2-decanol after four days of incu-
bation (Figure 4, Figures S9 and S10 in the Supporting Informa-
tion). 1-Decanol formation was not detected and no product
was found in E. coli pET28a(+) control experiments. Although
Although no hydration was obtained with dec-9-enoic acid
(C10) as a substrate under the optimized reaction conditions,
&
ChemCatChem 2017, 9, 1 – 10
www.chemcatchem.org
4
ꢀ 2017 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
ÝÝ These are not the final page numbers!

Full Papers
Em-OAH whole cells, 2-decanol formation was studied under
the optimized conditions (addition of glucose and FAD) in bio-
transformations using whole cells and cell lysate. Product for-
mation in whole cell biotransformations was set as 100%. In
contrast to whole cell transformations, the hydration activity of
1-decene strongly decreased when cell lysate (10% relative
conversion, data not shown) was used, which is most likely
a result of protein denaturation during the biotransformation.
Hydratases characterized so far show a narrow substrate
scope catalyzing the hydration of cis-9 and cis-9,12 unsaturated
fatty acids with chain lengths between 14 and 22 carbon
atoms.[11,13,14,21–23,25,37] Interestingly, in previous work, we ob-
served low activities of oleate hydratases towards (Z)-undec-9-
enoic acid (C11).[26] This motivated us to improve the hydration
of the non-natural C11 fatty acid substrate by Em-OAH by opti-
mizing the reaction conditions. Under reducing conditions, Em-
OAH expressing whole cells showed considerably higher con-
version rates compared with cell lysate. The efficient conver-
sion of hydrophobic substrates in whole cell biotransforma-
tions has already been demonstrated[17,49,50] and led to the suc-
cessful production of 10-hydroxystearic acid and 13-S-hydroxy-
9-(Z)-octadecenoic acid by recombinant oleate hydratase from
Stenotrophomonas maltophilia[14] and linoleate 13-hydratase
from L. acidophilus,[38] respectively. Oleate hydratase from E.
meningoseptica as well as hydratases characterized so far do
contain a flavin cofactor. As no net redox change is involved in
the catalysis, the functional role of FAD remains un-
known.[9,11–14,21–23] However, analysis of hydratases from Macro-
coccus caseolyticus[13] and Streptococcus pyogenes[21] indicates
a stabilizing structural role of the cofactor, which has also been
reported for several other FAD-containing enzymes.[51–53] The
FAD cofactor is non-covalently bound in Em-OAH and previous
studies showed an incomplete cofactor load.[12] Therefore, the
addition of FAD might allow a complete cofactor load, ena-
bling maximum enzymatic activity, whereas the specific activity
is strongly decreased applying the apo-enzyme.[12] The de-
pendency of hydratase activity on FAD was also shown for li-
noleic acid D9 hydratase from L. plantarum.[37] Similar to the
present study, the activity was strongly enhanced by addition
of NAD(P)H. These results suggest an activating role of the nic-
otinamide cofactor in the reduction of FAD.[37] In line with this,
increased specific activity of Em-OAH was also observed after
photo- and chemical reduction of the flavin cofactor.[12]
Figure 4. 1-Decene (0.5 mm) conversion in Em-OAH biotransformations
under improved conditions (50 mgmLÀ1 biomass, 100 mm glucose, 0.3 mm
FAD, 3 mm NADH, and 2 mm DTT at 258C).
GC vials were used as reaction tubes for alkene conversion,
20–30% of 1-decene evaporated during the transformations.
The stereoselectivity of 1-decene hydration was analyzed by
chiral GC (Figure S11 in the Supporting Information). Em-OAH
biotransformations without 1-decene served as control reac-
tions. Although no substrate was added to these reactions,
a small peak co-eluted with R-2-decanol. This background
noise was subtracted from the biotransformation results prior
to the calculation of the enantiomeric excess (ee). An excellent
selectivity for S-(+)-2-decanol with an ee value of >99% was
determined in the biotransformations with and without the
hexanoic acid dummy substrate. The formation of the
branched isomer is favored over the linear one, which corre-
lates with the known favorability of an alkene to undergo Mar-
kovnikov addition. In the natural reaction, the hydration of
oleic acid results in the formation of the R-enantiopure prod-
uct of 10-hydroxyoctadecanoic acid.[12,47,48] The reverse product
configuration in the conversion of 1-decene can be explained
by a change in priority on application of the Cahn–Ingold–
Prelog (CIP) rule as observed for the hydration of 12-oxooleate,
giving (S)-10-hydroxy-12-oxostearic acid.[18]
The chemical differences between fatty acids and alkenes
prompted us to optimize the reactions conditions for the hy-
dration of 1-decene (Figure S12 in the Supporting Information).
Approximately 15% of 2-decanol was obtained after three
days of incubation by applying 50 mgmLÀ1 recombinant cells
without any additive. Product formation was doubled when
glucose was used as an additive and further increased up to
40% upon glucose and FAD addition. Neither NADH and DTT
nor an incubation temperature of 308C showed any effect on
product formation. However, the increased incubation temper-
ature had a slightly negative influence on substrate recovery.
Only up to 70% of the substrate and product were recovered
after three days of incubation. Owing to the enhanced alkene
evaporation at 308C, further experiments were performed at
258C. Based on the results from the hydration of 1-decene by
Analysis of the stereoselectivity showed a S-configuration of
the hydrated product. The natural reaction of Em-OAH is the
regioselective hydration of oleic acid to (R)-10-hydroxyoctade-
canoic acid.[12,47,48] The alteration in stereoselectivity might
result from differences in substrate orientation. The binding of
(Z)-undec-9-enoic acid seems to differ in orientation owing to
the lack of the long alkyl chain on the omega site of the
double bond.
(Z)-Undec-9-enoic acid is the shortest fatty acid converted
by Em-OAH so far. In this light, we pursued the hydration of
smaller non-natural alkenes by Em-OAH in the presence of hex-
anoic acid as a dummy molecule. The general mode of action
for dummy molecules involves the partial filling of the active
site pocket, prompting the substrate to bind in a consistent
ChemCatChem 2017, 9, 1 – 10
www.chemcatchem.org
5
ꢀ 2017 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
&
These are not the final page numbers! ÞÞ

Full Papers
0.5 mm finale isopropyl b-d-1-thiogalactopyranoside (IPTG) concen-
tration. After 20 h of incubation at 208C and 140 rpm, cells were
harvested by centrifugation (12 min, 8000ꢂg, 48C) and washed
twice in citrate buffer (50 mL, 50 mm, pH 6). Pellets were stored at
À208C until usage.
orientation. However, it will not participate in the actual cataly-
sis. The results underline the importance of the dummy sub-
strate as product formation was strongly decreased without
the addition of hexanoic acid. The dependency of OAH activity
on a carboxylic group for substrate recognition by positively
charged amino acids has been demonstrated in previous
work.[9,12,20] The hypothesis is further supported by the inactivi-
ty of hydratases towards substrates such as methyl linoleate,
lacking the corresponding functional group.[23,37]
Cell disruption by sonication
For biotransformations with cell lysate, cell suspensions
(100 mgmLÀ1 cells in 50 mm citrate buffer, pH 6) were constantly
held on ice and disrupted by sonication for 5 min (duty cycle=
35%, output=4; Sonifier 250, Branson, Danbury, USA). The lysed
cells were centrifuged at 16000ꢂg for 15 min at 48C. Protein con-
centrations of the cell lysate soluble supernatant were determined
by using the “Pierce BCA Protein Assay Kit” (Thermo Scientific, Wal-
tham, USA) according to the manufacturer’s microwell plate proto-
col.
The great chemical differences between fatty acids and al-
kenes prompted us to analyze the reaction conditions regard-
ing the hydration of 1-decene. While alkenes are not degraded,
fatty acids are metabolized by E. coli whole cells. The addition
of NADH (and DTT) might have a crucial influence on cell me-
tabolism, saturating the oxidative cell machinery with redox
equivalents. This does not seem to be important for 1-decene
hydration but for (Z)-undec-9-enoic acid hydration, as this
compound is subjected to cell metabolism. Our results show
that the optimization of reaction conditions does not only
depend on the enzyme itself but also on the respective sub-
strate. This work underlines the importance of improved reac-
tion conditions and the applicability of statistical experiment
design for the conversion of non-natural substrates.
Biotransformations—standard/FAD-reducing conditions
Biotransformations were performed in citrate buffer (50 mm, pH 6)
on a 500 mL scale using 1.5 mL reaction tubes. E. coli BL21 (DE3)
pET28a(+) whole cells and cell lysate, respectively, served as nega-
tive controls. Expression cultures were thawed on ice, resuspended
in citrate buffer, and disrupted by sonication for lysate assays. The
cell suspension (50 mgmLÀ1) and protein (4 mgmLÀ1) served for
whole cell and cell lysate biotransformations, respectively. Glucose
(50 mm) was added to whole cell biotransformations and FAD
(0.2 mm), DTT (5 mm), and NADH (5 mm) were applied for the
FAD-reducing conditions. Reactions were started by the addition of
substrate (0.5 mm (Z)-undec-9-enoic acid, 1% final DMSO concen-
tration) and incubated at 258C and 800 rpm. After five days, the
biotransformations (in triplicate) were quenched by addition of HCl
(20 mL, 1m) and extracted twice with methyl tert-butyl ether
(MTBE, 500 mL) and dodecanoic acid (100 mm) as internal standard.
After each extraction step, the organic phase (300 mL) was collect-
ed and concentrated in vacuo (EZ-2 Plus, GeneVac, Ipswich, UK).
Fatty acids were derivatized for 30 min at 708C by using N,O-bis(-
trimethylsilyl)trifluoroacetamide (40 mL) containing 1% trimethyl-
chlorosilane. Product formation was analyzed by GC-FID.
Conclusions
In this study, we demonstrated the hydration of two non-natu-
ral short-chain fatty acid and alkene substrates by the oleate
hydratase from E. meningoseptica. Based on the optimization
of reaction conditions by statistical experiment design (S)-10-
hydroxyundecanoic acid was produced from (Z)-undec-9-enoic
acid on a preparative scale by using recombinant E. coli whole
cells. Moreover, optimized conditions enabled the first stereo-
selective hydration of 1-decene to S-(+)-2-decanol by using
a fatty acid dummy substrate.
The conversion of alkenes represents the next step towards
the hydration of non-natural short-chain molecules. Activity to-
wards these substrates might be further enhanced by methods
of protein engineering applying random mutagenesis or ra-
tional protein design. Future efforts will also concentrate on in-
vestigating the role of the fatty acid dummy substrate.
Design of experiments (DoE)
Reaction conditions were optimized by using MODDE for statistical
experiment design (Version 11; MKS Umetrics AB, Sweden). All bio-
transformations were performed by using whole cells (50 mgmLÀ1
)
Experimental Section
at the 500 mL scale in 1.5 mL reaction tubes. Expression cultures
were thawed on ice and resuspended in citrate buffer (50 mm,
pH 6). Compositions were designed by MODDE according to the
selected interaction model (FracFac Res V+, Full Fac (2 levels)). Up
to five conditions were analyzed simultaneously in four rounds of
optimization. The following components and concentrations were
tested: glucose (0–100 mm), NaCl (0–100 mm), FAD (0–0.5 mm),
DTT (0–5 mm), NADH (0–5 mm), glutathione (0–5 mm), and BSA (0–
0.5 mgmLÀ1). In the first round, glucose, NADH, FAD, DTT, and glu-
tathione concentrations were analyzed. Based on the obtained re-
sults, glutathione was excluded from further analysis and the glu-
cose concentration was set to 100 mm in the second round of op-
timization, varying once again the concentrations of NADH, FAD,
and DTT. In the third round, glucose, NADH, and FAD concentra-
tions were fixed, while the amount of DTT was adjusted. The four
parameters were fixed in the last round of optimization, analyzing
Unless otherwise stated, all solvents, buffer components, and
chemicals were purchased from Acros Organics (Geel, Belgium),
Alfa Aesar (Karlsruhe, Germany), Carl Roth GmbH (Karlsruhe, Ger-
many), Macherey–Nagel (Dꢁren, Germany), Merck (Darmstadt, Ger-
many), Peqlab (Erlangen, Germany), Serva (Heidelberg, Germany),
Sigma–Aldrich (St. Louis, USA), and VWR (Darmstadt, Germany).
Expression of recombinant Em-OAH
Terrific broth (TB) medium (400 mL) in a 2 L baffled flask with
30 mgmLÀ1 final kanamycin concentration was inoculated with an
overnight preculture of E. coli BL21 (DE3) pET28a(+)/Em-OAH to an
OD600 of 0.02. Cultures were incubated at 378C and 180 rpm and
gene expression was induced at an OD600 of 0.5–0.7 by addition of
&
ChemCatChem 2017, 9, 1 – 10
www.chemcatchem.org
6
ꢀ 2017 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
ÝÝ These are not the final page numbers!

Full Papers
different NaCl and BSA concentrations at 25, 30, and 378C. Bio-
transformations were started with the addition of substrate
(0.5 mm (Z)-undec-9-enoic acid, 1% finale DMSO concentration).
After 28 h of incubation at 800 rpm, the reactions were quenched
by the addition of HCl (20 mL, 1m) and extracted as described
above. Substrate conversion and product formation was analyzed
by GC-FID. The optimized conditions for (Z)-undec-9-enoic acid
conversion were evaluated by MODDE, applying tools for analysis
and prediction.
H), 1.60–1.66 (m, 2H, 3-H), 2.34 (t, J=7.5 Hz, 2H, 2-H), 3.77–
3.83 ppm (sext, J=6 Hz, 1H, 10-H).
Stereoselectivity of (Z)-undec-9-eoic acid hydration
Purified 10-hydroxyundecanoic acid (25 mg, 0.125 mmol) was dis-
solved in trimethylsulfonium hydroxide solution (TMSH, 1 mL,
0.25m in methanol) for the derivatization of the free carboxyl
group. After 1 h of incubation at 408C and 800 rpm, formic acid (>
98%, 5 mL, 46 gmolÀ1, ꢀ0.125 mmol) was added to remove the
surplus TMSH. After 30 min of incubation at room temperature and
300 rpm, the solution was transferred to a 5 mL pear-shaped flask
and the volatile compounds were removed by using a rotary evap-
orator.
Upscaling of (Z)-undec-9-enoic acid conversion
TB medium (5 L) with kanamycin (30 mgmLÀ1 final kanamycin con-
centration) was inoculated with preculture of E. coli BL21 DE3
pET28a(+)/Em-OAH (200 mL) to an OD600 of 0.05. Cells were culti-
vated at 378C and 800 rpm with 8 LminÀ1 aeration rate. The pH
value was kept at 7.2 by addition of ammonium hydroxide (28%)
and phosphoric acid (10%). For induction of heterologous protein
expression, the temperature was reduced to 308C and IPTG was
added to a final concentration of 0.5 mm at an OD600 of 5.5.
MgSO4·7H2O (25 mL, 1m), trace solution (2.5 mL) containing
CaCl2·2H2O (0.38 gLÀ1), ZnSO4·7H2O (0.18 gLÀ1), MnSO4·H2O
The stereoselectivity of 10-OH-undecanoic acid hydration by Em-
OAH was examined by Mosher ester analysis by applying a modi-
fied protocol by Hoye et al.[54] TMSH-derivatized product (5 mg,
ꢀ0.025 mmol) was dissolved in deuterated chloroform (1 mL, dried
with Na2SO4) in a 2 mL glass vial. Anhydrous pyridine (40 mL,
ꢀ0.5 mmol, 20 equiv) and (S)-(+)-a-methoxy-a-trifluoromethylphe-
nylacetyl chloride (S-(+)-MTPA-Cl, 74 mL, ꢀ0.4 mmol, 16 equiv) was
added and the reaction was stirred for 3 h at room temperature.
The same protocol was applied for the derivatization with R-(À)-
MTPA-Cl. Aliquots (600 mL) of each solution were analyzed by NMR
spectroscopy (500 Hz, 1H).
(0.1 gLÀ1),
Na2·ETDA
(20.1 gLÀ1),
FeCl3·6H2O
(16.7 gLÀ1),
CuSO4·5H2O (0.15 gLÀ1), and CoCl2·6H2O (0.18 gLÀ1), as well as
(NH4)2HPO4 (150 mL, 2.5m) were added to the culture. Glycerol
(86%) served as the carbon source with a feeding rate of 15 ghÀ1
,
which was increased to 22 ghÀ1 10 h after induction. Cells were
harvested after 23 h of incubation at a final OD600 of 44 and pellets
were stored at À208C.
Conversion of 1-decene
1-Decene biotransformations were performed in citrate buffer
(50 mm, pH 6) on the 500 mL scale in 2 mL glass vials as reaction
tubes. E. coli BL21 (DE3) pET28a(+) served as the negative control.
Expression cultures were thawed on ice, resuspended in citrate
buffer, and disrupted by sonication for lysate assays. Cell suspen-
sion (50 mgmLÀ1) and protein (3.8 mgmLÀ1) served for whole cell
and lysate biotransformations, respectively. Glucose (100 mm), FAD
(0.3 mm), NADH (3 mm), and DTT (2 mm) were applied for the de-
tailed analysis of MODDE-optimized conditions. Glucose (100 mm)
and FAD (0.3 mm) were applied for the direct comparison of whole
cells and lysate under optimized and non-optimized conditions.
Reactions were started by the addition of substrate and dummy
substrate (0.5 mm 1-decene and 0.5 mm hexanoic acid, respective-
ly) with 1% final DMSO concentration and incubated at 258C and
800 rpm. Biotransformation samples (in triplicate) were quenched
by addition of NaOH (20 mL, 1m) and extracted twice with MTBE
(500 mL) and 1-octanol (500 mm) as internal standard. After each ex-
traction step, 300 mL of the organic phase was collected and direct-
ly analyzed by GC/GC-MS.
Upscaling of (Z)-undec-9-enoic acid conversion was performed
with 200 mg fatty acid substrate (1 mm final concentration, 1%
final DMSO concentration) in a 2 L Schott flask under the opti-
mized conditions (50 mgmLÀ1 biomass, 100 mm glucose, 0.3 mm
FAD, and 3 mm NADH). The biotransformation was started with the
addition of substrate and incubated at 308C and 180 rpm. After
two and three days of incubation, samples (500 mL, in triplicate)
were taken and prepared as described above. Substrate conversion
and product formation was analyzed by GC-FID. After three days of
incubation, the biotransformation was stopped with HCl (40 mL,
5m) and fatty acids were extracted by adding the equivalent
volume of MTBE. The organic phase was collected and the aque-
ous phase was extracted three times with MTBE (600 mL). The or-
ganic phases were combined, concentrated in vacuo, and stored at
48C.
The extracted biotransformation product was dissolved in the
mobile phase (98% dichloromethane with 2% methanol and 1%
acetic acid as additive) and purified by column chromatography
(40 cm length, 5 cm diameter, 300 g silica gel 60). Purification frac-
tions were collected and analyzed by thin layer chromatography
(pre-coated TLC sheets ALUGRAMꢃXtra SIL G/UV254; 0.2 mm, silica
gel 60). TLC sheets were developed in phosphomolybdic acid hy-
drate (5 g in 50 mL ethanol) and by subsequent heating. Samples
of each fraction were prepared for GC-MS analysis as described
above. Product fractions were pooled and concentrated in vacuo.
A light-brown powder (132 mg) was obtained as the product with
>95% purity (YP/S=66%, YP/B =0.0024). Extracted and purified 10-
hydroxyundecanoic acid (10 mg) was mixed with deuterated
chloroform (600 mL CDCl3) for NMR analysis (1H, 13C). 13C NMR
(125 MHz, CDCl3): d=23.4, 24.7, 25.7, 29.0, 29.1, 29.3, 29.5, 34.0,
For chiral GC analysis, acetic anhydride (250 mL) and a catalytic
amount of 4-dimethylaminopyridine (DMAP, 2 mg) were added to
the organic phase (600 mL). After 1 h of incubation at 258C and
800 rpm, water (300 mL) was added. The organic phase was trans-
ferred to a new reaction tube, dried over Na2SO4, and analyzed by
GC-FID (CP-Chirasil-Dex CB).
Analytics—GC and GC-MS
Derivatized fatty acid samples were analyzed by GC by using a DB-
5 polyphenylmethylsiloxane column (30 m, 0.25 mm, 0.25 mm; Agi-
lent technologies, Santa Clara, USA) on a Shimadzu GC2010 Plus
(Shimadzu, Kyo¯to, Japan) equipped with an AOC 20s autosampler
and AOC 20i autoinjector (injection volume: 1 mL, split ratio: 1:10,
1
39.0, 68.0, 179.0 ppm; H NMR (500 MHz, CDCl3): d=1.18 (s, 1H, 11-
H), 1.19 (s, 2H, 11-H), 1.30–1.47 (m, 12H, 4-H, 5-H, 6-H, 7-H, 8-H, 9-
ChemCatChem 2017, 9, 1 – 10
www.chemcatchem.org
7
ꢀ 2017 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
&
These are not the final page numbers! ÞÞ

Full Papers
injection temperature: 2508C). Hydrogen served as the carrier gas
with a linear velocity of 30 cmsÀ1 and the detector temperature
was 3208C. The initial oven temperature was set to 1608C, raised
to 2208C at a rate of 10 KminÀ1, and subsequently increased to
2608C (20 KminÀ1). Finally, the temperature was raised to 3208C
with a rate of 40 KminÀ1 and held for 1 min. Quantification was
done with dodecanoic acid as internal standard.
influx into the solution. The grey mixture was filtered over Celite,
washed with methanol, and the filtrate was concentrated in vacuo.
The yellowish oil was purified by column chromatography (cyclo-
hexane/MTBE 5:1, Rf =0.36, vanillin) to yield 394 mg of (Z)-undec-9-
enoic acid (2.14 mmol, 81% yield) as a colorless oil (>95% H NMR
purity).
1
1H NMR (500 MHz, CDCl3): d=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 ppm (m, 2H, 9-H,
10-H); 13C NMR (125 MHz, CDCl3): d=12.8, 24.7, 26.8, 29.0, 29.1,
29.5, 33.9, 123.7, 130.8, 179.6 ppm; FTIR (ATR): n˜ =3013 (w), 2924
(m), 2855 (m), 1705 (s), 1412 (m), 1284 (m), 1245 (m), 1114 (w), 934
(m), 700 cmÀ1 (m).
GC-MS analysis of derivatized fatty acids was performed by using
a ZB-5 polyphenylmethylsiloxane column (30 m, 0.25 mm, 0.25 mm;
Zebron-Phenomenex, Torrance, USA) on
a Shimadzu GC2010
system (Shimadzu, Kyo¯to, Japan) equipped with a GCMS-QP2010
mass selective detector (electron impact, 70 eV) and AOC 5000 au-
tosampler/autoinjector (injection volume: 1 mL, split ratio: 1:10, in-
jection temperature: 2508C). Hydrogen served as the carrier gas
with a linear velocity of 30 cmsÀ1 and the detection temperature
was 3208C. The MS ion source temperature was set to 2508C and
the interface temperature was 2708C. The temperature program
was the same as for GC-FID analysis.
Acknowledgments
This project has received funding from the European Union’s Ho-
rizon 2020 Research and Innovation Programme under Grant
agreement No 635536. The production of biomass by fed-batch
cultivation was kindly supervised by Sven Richter. Many thanks
to Silvia Fademrecht for the creative and skilled support regard-
ing the preparation of the schematic substrate binding pocket.
Special thanks go to Max Fischer and Jens Schmid for interesting
and highly motivating discussions.
1-Decene biotransformation samples were analyzed by GC without
derivatization by using a HP-1 dimethylpolysiloxane column (30 m,
0.25 mm, 0.25 mm; Agilent technologies, Santa Clara, USA) on a Shi-
madzu GC2010 (Shimadzu, Kyo¯to, Japan) equipped with an AOC
20s autosampler and AOC 20i autoinjector (injection volume: 1 mL,
split ratio: 1:5, injection temperature: 2508C). Hydrogen served as
the carrier gas with a linear velocity of 30 cmsÀ1 and the detector
temperature was 3308C. The initial oven temperature was set to
808C, raised to 1608C at a rate of 10 KminÀ1, subsequently in-
creased to 3208C with a rate of 50 KminÀ1, and held for 4 min.
Quantification was done with 1-octanol as internal standard.
Keywords: enzyme catalysis · oleate hydratase · reaction
optimization · stereoselectivity · upscaling
GC-MS analysis of 1-decene biotransformation samples was per-
formed by using a DB-5 column ((5% phenyl)–methylpolysiloxane,
30 m, 0.25 mm, 0.25 mm; Agilent technologies, Santa Clara, USA)
with an Agilent 7890A GC/MS system (Santa Clara, US) equipped
with a flame ionization detector as well as mass selective detector
(Agilent 5975C inert MSD, Santa Clara, US). Helium served as the
carrier gas (constant pressure of 16.521 psi) and the detection tem-
perature was 3208C. The sample (1 mL) was injected with a split
ratio of 1:10 at an injection temperature of 2508C. The initial oven
temperature was set to 808C, raised to 1408C at a rate of
[1] K. Faber, M. C. R. Franssen, Trends Biotechnol. 1993, 11, 461–470.
[2] N. ꢅhrner, C. Orrenius, A. Mattson, T. Norin, K. Hult, Enzyme Microb.
Technol. 1996, 19, 328–331.
[3] S. Panke, M. Wubbolts, Curr. Opin. Chem. Biol. 2005, 9, 188–194.
[4] S. Schneider, M. G. Wubbolts, D. Sanglard, B. Witholt, Tetrahedron: Asym-
metry 1998, 9, 2832–2844.
[5] K. B. Otte, B. Hauer, Curr. Opin. Biotechnol. 2015, 35, 16–22.
[6] Y. Pagot, A. Endrizzi, J. M. Nicaud, J. M. Belin, Lett. Appl. Microbiol. 1997,
25, 113–116.
[7] M. Dietrich, T. A. Do, R. D. Schmid, J. Pleiss, V. B. Urlacher, J. Biotechnol.
2009, 139, 115–117.
15 KminÀ1
40 KminÀ1, and held for 5 min.
, subsequently increased to 3108C with a rate of
[8] J. Jin, U. Hanefeld, Chem. Commun. 2011, 47, 2502–2510.
[9] A. Hiseni, I. W. C. E. Arends, L. G. Otten, ChemCatChem 2015, 7, 29–37.
[10] B. N. M. van Leeuwen, A. M. van Der Wulp, I. Duijnstee, A. J. A. van Maris,
A. J. J. Straathof, Appl. Microbiol. Biotechnol. 2012, 93, 1377–1387.
[11] B. N. Kim, Y. C. Joo, Y. S. Kim, K. R. Kim, D. K. Oh, Appl. Microbiol. Biotech-
nol. 2012, 95, 929–937.
[12] 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. Schꢁrmann, S. Wallner, P. Macheroux, K. Gruber, H. Pichler,
ChemBioChem 2015, 16, 1730–1734.
The stereoselectivity of 1-decene hydration was analyzed by chiral
GC by using a CP-Chirasil-Dex CB column (25 m, 0.25 mm, 0.25 mm;
Agilent technologies, Santa Clara, USA) with a Shimadzu GC2010
(Shimadzu, Kyo¯to, Japan) equipped with an AOC 20s autosampler
and AOC 20i autoinjector (injection volume: 1 mL, split ratio: 1:20,
injection temperature: 2208C). Hydrogen served as the carrier gas
with a linear velocity of 40 cmsÀ1 and the detector temperature
was 2308C. The initial oven temperature was set to 1008C and
held for 1 min, raised to 1208C at a rate of 10 KminÀ1 and held for
[13] Y. C. Joo, K. W. Jeong, S. J. Yeom, Y. S. Kim, Y. Kim, D. K. Oh, Biochimie
2012, 94, 907–915.
[14] Y. C. Joo, E. S. Seo, Y. S. Kim, K. R. Kim, J. B. Park, D. K. Oh, J. Biotechnol.
2012, 158, 17–23.
2.5 min, subsequently increased to 1508C with a rate of 10 KminÀ1
and finally raised to 2008C (50 KminÀ1) and held for 3 min.
,
[15] B. N. Kim, S. J. Yeom, D. K. Oh, Biotechnol. Lett. 2011, 33, 993–997.
[16] E. Y. Jeon, J. H. Lee, K. M. Yang, Y. C. Joo, D. K. Oh, J. B. Park, Process Bio-
chem. 2012, 47, 941–947.
Synthesis of (Z)-undec-9-enoic acid
[17] J. W. Song, E. Y. Jeon, D. H. Song, H. Y. Jang, U. T. Bornscheuer, D. K. Oh,
J. B. Park, Angew. Chem. Int. Ed. 2013, 52, 2534–2537; Angew. Chem.
2013, 125, 2594–2597.
[18] K. B. Otte, PhD Thesis, University of Stuttgart (Germany) 2014, DOI
10.18419/opus-1424.
(Z)-Undec-9-enoic acid (0.48 g, 2.64 mmol) was added to Lindlar
catalyst (0.06 g) in a dry Schlenk tube and dissolved in methanol
(15 mL, dried over 3 ꢄ molecular sieves). The mixture was stirred at
room temperature and the atmosphere was replaced ten times
with hydrogen by subsequent evacuation and hydrogen flooding.
The reactions were incubated for 20 h under continuous hydrogen
[19] L. E. Bevers, M. W. H. Pinkse, P. D. E. M. Verhaert, W. R. Hagen, J. Bacteriol.
2009, 191, 5010–5012.
&
ChemCatChem 2017, 9, 1 – 10
www.chemcatchem.org
8
ꢀ 2017 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
ÝÝ These are not the final page numbers!

Full Papers
[20] A. Volkov, S. Khoshnevis, P. Neumann, C. Herrfurth, D. Wohlwend, R.
Ficner, I. Feussner, Acta Crystallogr. Sect. D 2013, 69, 648–657.
[21] A. Volkov, A. Liavonchanka, O. Kamneva, T. Fiedler, C. Goebel, B. Kreike-
meyer, I. Feussner, J. Biol. Chem. 2010, 285, 10353–10361.
[22] B. Yang, H. Chen, Y. Song, Y. Q. Chen, H. Zhang, W. Chen, Biotechnol.
Lett. 2013, 35, 75–81.
[23] E. Rosberg-Cody, A. Liavonchanka, C. Gçbel, R. P. Ross, O. O’Sullivan,
G. F. Fitzgerald, I. Feussner, C. Stanton, BMC Biochem. 2011, 12, 9.
[24] K.-R. Kim, H.-J. Oh, C.-S. Park, S.-H. Hong, J.-Y. Park, D.-K. Oh, Biotechnol.
Bioeng. 2015, 112, 2206–2213.
[38] J.-Y. Park, S.-H. Lee, K.-R. Kim, J.-B. Park, D.-K. Oh, J. Biotechnol. 2015,
208, 1–10.
[39] P. N. Black, C. C. Dirusso, Microbiol. Mol. Biol. Rev. 2003, 67, 454–472.
[40] K. T. Hughes, R. W. Simons, W. D. Nunn, J. Bacteriol. 1988, 170, 1666–
1671.
[41] W. D. Nunn, R. W. Simons, Proc. Natl. Acad. Sci. USA 1978, 75, 3377–
3381.
[42] D. Clark, J. Bacteriol. 1981, 148, 521–526.
[43] F. Kamp, D. Zakim, F. Zhang, N. Noy, J. A. Hamilton, ACS Biochem. 1995,
34, 11928–11937.
[44] F. Kamp, J. A. Hamilton, Prostaglandins Leukotrienes Essent. Fatty Acids
2006, 75, 149–159.
[25] A. Hirata, S. Kishino, S.-B. Park, M. Takeuchi, N. Kitamura, J. Ogawa, J.
Lipid Res. 2015, 56, 1340–1350.
[45] W. Niehaus, G. Schroepfer, Biochem. Biophys. Res. Commun. 1965, 21,
271–275.
[46] L. L. Wallen, E. N. Davis, Y. V. Wu, W. K. Rohwedder, Lipids 1971, 6, 745–
750.
[47] W. G. Niehaus, A. Kisic, A. Torkelson, D. J. Bednarczyk, G. J. Schroepfer, J.
Biol. Chem. 1970, 245, 3790–3797.
[48] G. J. Schroepfer, K. Bloch, J. Am. Chem. Soc. 1963, 85, 3310–3311.
[49] K. B. Otte, J. Kittelberger, M. Kirtz, B. M. Nestl, B. Hauer, ChemCatChem
2014, 6, 1003–1009.
[50] N. Kishimoto, I. Yamamoto, K. Toraishi, S. Yoshioka, K. Saito, H. Masuda,
T. Fujita, Lipids 2003, 38, 1269–1274.
[51] D. Vargo, A. Pokora, S. W. Wang, M. S. Jorns, J. Biol. Chem. 1981, 256,
6027–6033.
[52] T. H. Cromartie, C. T. Walsh, J. Biol. Chem. 1976, 251, 329–333.
[53] R. G. Duggleby, S. S. Pang, J. Biochem. Mol. Biol. 2000, 33, 1–36.
[54] T. R. Hoye, C. S. Jeffrey, F. Shao, Nat. Protoc. 2007, 2, 2451–2458.
[26] J. Schmid, L. Steiner, S. Fademrecht, J. Pleiss, K. B. Otte, B. Hauer, J. Mol.
Catal. B: Enzym. 2017, DOI: 10.1016/j.molcatb.2017.01.010.
[27] N. Kawakami, O. Shoji, Y. Watanabe, Chem. Sci. 2013, 4, 2344–2348.
[28] N. Kawakami, O. Shoji, Y. Watanabe, Angew. Chem. Int. Ed. 2011, 50,
5315–5318; Angew. Chem. 2011, 123, 5427–5430.
[29] O. Shoji, T. Kunimatsu, N. Kawakami, Y. Watanabe, Angew. Chem. Int. Ed.
2013, 52, 6606–6610; Angew. Chem. 2013, 125, 6738–6742.
[30] O. Shoji, Y. Watanabe, Isr. J. Chem. 2015, 55, 32–39.
[31] F. E. Zilly, J. P. Acevedo, W. Augustyniak, A. Deege, H. Uw, M. T. Reetz,
Angew. Chem. Int. Ed. 2011, 50, 2720–2724; Angew. Chem. 2011, 123,
2772–2776.
[32] http://umetrics.com/product/modde-go.
[33] J. Pezzini, G. Joucla, R. Gantier, M. Toueille, A. M. Lomenech, C. Le Sꢆnꢆ-
chal, B. Garbay, X. Santarelli, C. Cabanne, J. Chromatogr. A 2011, 1218,
8197–8208.
[34] J. Fricke, K. Pohlmann, N. A. Jonescheit, A. Ellert, B. Joksch, R. Luttmann,
Biotechnol. J. 2013, 8, 738–747.
Manuscript received: October 19, 2016
Revised: December 21, 2016
[35] H. Tye, M. Whittaker, Org. Biomol. Chem. 2004, 2, 813–815.
[36] M. Stꢁtzle, A. Moll, R. Handrick, K. Schindowski, BMC Proc. 2013, 7, P74.
[37] M. Takeuchi, S. Kishino, A. Hirata, S.-B. Park, N. Kitamura, J. Ogawa, J.
Biosci. Bioeng. 2015, 119, 636–641.
Accepted Article published: December 21, 2016
Final Article published: && &&, 0000
ChemCatChem 2017, 9, 1 – 10
www.chemcatchem.org
9
ꢀ 2017 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
&
These are not the final page numbers! ÞÞ

FULL PAPERS
R. M. Demming, K. B. Otte, B. M. Nestl,
B. Hauer*
Selective addition of water: The forma-
tion of (S)-10-hydroxyundecanoic acid
and (S)-2-decanol with high yield and
excellent stereoselectivity by oleate hy-
dratase from E. meningoseptica is report-
ed as a first step towards the hydration
of short-chain non-natural substrates.
&& – &&
Optimized Reaction Conditions Enable
the Hydration of Non-natural
Substrates by the Oleate Hydratase
from Elizabethkingia meningoseptica
&
ChemCatChem 2017, 9, 1 – 10
www.chemcatchem.org
10
ꢀ 2017 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
ÝÝ These are not the final page numbers!