Biochemical characterization and FAD-binding analysis of oleate hydratase from Macrococcus caseolyticus

Article abstract of DOI:10.1016/j.biochi.2011.12.011

A putative fatty acid hydratase gene from Macrococcus caseolyticus was cloned and expressed in Escherichia coli. The recombinant enzyme was a 68 kDa dimer with a molecular mass of 136 kDa. The enzymatic products formed from fatty acid substrates by the putative enzyme were isolated with high purity (>99%) by solvent fractional crystallization at low temperature. After the identification by GC-MS, the purified hydroxy fatty acids were used as standards to quantitatively determine specific activities and kinetic parameters for fatty acids as substrates. Among the fatty acids evaluated, specific activity and catalytic efficiency (k_cat/K_m) were highest for oleic acid, indicating that the putative fatty acid hydratase was an oleate hydratase. Hydration occurred only for cis-9-double and cis-12-double bonds of unsaturated fatty acids without any trans-configurations. The maximum activity for oleate hydration was observed at pH 6.5 and 25 °C with 2% (v/v) ethanol and 0.2 mM FAD. Without FAD, all catalytic activity was abolished. Thus, the oleate hydratase is an FAD-dependent enzyme. The residues G29, G31, S34, E50, and E56, which are conserved in the FAD-binding motif of fatty acid hydratases (GXGXXG_(A/S)X_(15-21)E_(D)), were selected by alignment, and the spectral properties and kinetic parameters of their alanine-substituted variants were analyzed. Among the five variants, G29A, G31A, and E56A showed no interaction with FAD and exhibited no activity. These results indicate that G29, G31, and E56 are essential for FAD-binding.

Full text of DOI:10.1016/j.biochi.2011.12.011

Biochimie 94 (2012) 907e915
Contents lists available at SciVerse ScienceDirect
Biochimie
journal homepage: www.elsevier.com/locate/biochi
Research paper
Biochemical characterization and FAD-binding analysis of oleate hydratase
from Macrococcus caseolyticus
*
Young-Chul Joo, Ki-Woong Jeong, Soo-Jin Yeom, Yeong-Su Kim, Yangmee Kim, Deok-Kun Oh
Department of Bioscience and Biotechnology, Konkuk University, 1 Hayang-dong Gangjin-gu, Seoul 143-701, Republic of Korea
a r t i c l e i n f o
a b s t r a c t
Article history:
A putative fatty acid hydratase gene from Macrococcus caseolyticus was cloned and expressed in
Escherichia coli. The recombinant enzyme was a 68 kDa dimer with a molecular mass of 136 kDa. The
enzymatic products formed from fatty acid substrates by the putative enzyme were isolated with
high purity (>99%) by solvent fractional crystallization at low temperature. After the identification by
GCeMS, the purified hydroxy fatty acids were used as standards to quantitatively determine specific
activities and kinetic parameters for fatty acids as substrates. Among the fatty acids evaluated, specific
activity and catalytic efficiency (kcat/Km) were highest for oleic acid, indicating that the putative fatty
acid hydratase was an oleate hydratase. Hydration occurred only for cis-9-double and cis-12-double
bonds of unsaturated fatty acids without any trans-configurations. The maximum activity for oleate
hydration was observed at pH 6.5 and 25 ꢀC with 2% (v/v) ethanol and 0.2 mM FAD. Without FAD, all
catalytic activity was abolished. Thus, the oleate hydratase is an FAD-dependent enzyme. The residues
G29, G31, S34, E50, and E56, which are conserved in the FAD-binding motif of fatty acid hydratases
(GXGXXG(A/S)X(15e21)E(D)), were selected by alignment, and the spectral properties and kinetic param-
eters of their alanine-substituted variants were analyzed. Among the five variants, G29A, G31A, and
E56A showed no interaction with FAD and exhibited no activity. These results indicate that G29, G31,
and E56 are essential for FAD-binding.
Received 13 September 2011
Accepted 13 December 2011
Available online 19 December 2011
Keywords:
Oleate hydratase
Characterization
FAD-binding
Hydroxy fatty acid
Substrate specificity
Crown Copyright Ó 2011 Published by Elsevier Masson SAS. All rights reserved.
1. Introduction
hydroxy fatty acids [4,5]. Oleate hydroxylase from the plant
Ricinus communis [6], lipoxygenase from Pseudomonas sp. 42A2L
A hydroxy fatty acid is composed of hydroxyl group(s) and
a long unbranched carbon chain with a carboxyl group at one end.
Hydroxy fatty acids are derived from a variety of natural sources,
including microorganisms, plant seed oils, plant waxes, plant cutin,
tall oil, and cork [1]. They are useful chemical intermediates for
synthesizing fine chemicals and pharmaceuticals. Some may
protect plants against microbial infection, although the mecha-
nisms of these anti-microbial effects are poorly understood [2,3].
The enzymes involved in the hydroxylation of fatty acids have
been reported as P450 monooxygenase, hydroxylase, lip-
oxygenase, and hydratase. P450 monooxygenases from Bacillus
licheniformis, B. subtilis, and Candida tropicalis catalyze the
hydroxylation of several saturated fatty acids into omega-
[7], and fatty acid hydratases [8e10] convert oleic acid into 12-
hydroxyoctadecanoic acid, (E)-10-hydroperoxy-8-octadecenoic
acid, and 10-hydroxyoctadecanoic acid, respectively.
Fatty acid hydratase activity that converts oleic acid into 10-
hydroxyoctadecanoic acid, was first reported in a myosin cross-
reactive antigen (MCRA) protein from Elizabethkingia meningosep-
tica [10]. The MCRA protein from Streptococcus pyogenes is a fla-
voenzyme that catalyzes the hydroxylation of double bonds in
unsaturated fatty acids [8]. Recently, a MCRA protein from Bifido-
bacterium breve has been identified as an FAD-dependent fatty acid
hydratase that catalyzes not only hydration but also isomerization
of linoleic acid to conjugated linoleic acid [9]. However, quantita-
tive analyses and characterization of fatty acid hydratases have not
been attempted because the products of the enzyme reaction are
not commercially available. Therefore, these fatty acid hydratases
have not been fully defined. Moreover, the FAD-binding residues in
these fatty acid hydratases have not been determined and their
roles are not fully understood.
Abbreviations: FAD, flavin adenine dinucleotide; MCRA, myosin cross-reactive
antigen; GCeMS, gas chromatography-mass spectrometry; LB, Luria-Bertani;
IPTG, isopropyl-b-D-thiogalactopyranoside; EDTA, ethylenediaminetetraacetic acid;
TLC, thin-layer chromatography; PIPES, piperazin-N,N0-ethanesulfonic acid; MES, 4-
morpholine ethanesulfonic acid; GR, glutathione reductase.
* Corresponding author. Tel.: þ82 2 450 4118; fax: þ82 2 444 6176.
E-mail address: deokkun@konkuk.ac.kr (D.-K. Oh).
In the present study, the hydroxylated products formed from
fatty acid substrates by the putative fatty acid hydratase from
0300-9084/$ e see front matter Crown Copyright Ó 2011 Published by Elsevier Masson SAS. All rights reserved.
doi:10.1016/j.biochi.2011.12.011

908
Y.-C. Joo et al. / Biochimie 94 (2012) 907e915
Macrococcus caseolyticus were purified. The purified products were
then used as standards in subsequent analyses. The biochemical
properties of the enzyme were characterized quantitatively using
manufactured standards, and the enzyme was exactly identified
based on its substrate specificity. In addition, the FAD-binding
properties of conserved residues in the FAD-binding motif were
characterized.
2.4. Enzyme purification
Cells were harvested from the culture broth by centrifugation at
6000 ꢁ g for 30 min at 4 ꢀC, washed twice with 0.85% NaCl, and
then resuspended in 50 mM phosphate buffer (pH 8.0) containing
300 mM KCl and 10 mM imidazole. The resuspended cells were
disrupted using Sonic Dismembrator (Fisher Scientific Model 100,
Pittsburg, PA, USA) at 18 W on ice for 2 min. The unbroken cells and
cell debris were removed by centrifugation at 13,000 ꢁ g for 20 min
2. Materials and methods
at 4 ꢀC and the supernatant was filtered through a 0.45
mm filter.
The filtrate was applied to an immobilized metal ion affinity
chromatography cartridge (Bio-Rad, Hercules, CA, USA) equili-
brated with 50 mM phosphate buffer (pH 8.0). The cartridge was
washed extensively with the same buffer and the bound protein
was eluted with a linear gradient of 10e500 mM imidazole at a flow
rate of 1 ml/min. The eluate was collected and loaded immediately
onto a Bio-Gel P-6 desalting cartridge (Bio-Rad) that had been
previously equilibrated with 50 mM PIPES buffer (pH 6.5). The
loaded protein was eluted with 50 mM PIPES buffer (pH 6.5) at
a flow rate of 1 ml/min to obtain a solution of purified enzyme. All
purification steps using the cartridges were carried out in a cold
room at 4 ꢀC with a Profinia protein purification system (Bio-Rad).
The protein expressed from plasmid pET-28a(þ) contained
thrombin restriction site (LeuValProArg/GlySer) at the N terminus
between hexa-histidine tag and putative fatty hydratase. After the
histidine tag in the expressed protein was hydrolyzed by treating
a thrombin kit (Novagen), the added thrombin (35 kDa) was
removed by untrafiltration using a Centricon (50 kDa cutoff, Mil-
lipore, Billerica, MA, USA). The resulting enzyme solution was
applied to a His-bind column (Novagen) and the waste was ob-
tained as a non-tagged enzyme. The column was washed with
50 mM phosphate buffer (pH 8.0) containing 300 mM NaCl and the
bound protein was eluted with 50 mM phosphate buffer (pH 8.0)
containing 87.5 mM imidazole. The eluate was used as a histidine-
tagged enzyme.
2.1. Materials
The expression vector pET-28a(þ) was purchased from Novagen
(San Diego, CA, USA). The expression host, Esherichia coli ER2566,
and all restriction enzymes were purchased from New England
Biolabs (Hertfordshire, UK). LB medium was purchased from BD
Biosciences (San Jose, CA, USA). The substrate fatty acids and the
cofactors FMN, NADþ and NADPþ were purchased from Sigma (St.
Louis, MO, USA). FAD was purchased from Tokyo Chemical Industry
(Tokyo, Japan). The pre-stained maker proteins for the SDS-PAGE
and gel filtration calibration kit were purchased from MBI Fer-
mentas (Hanover, MD, USA) and GE Healthcare (Piscataway, NJ,
USA), respectively.
2.2. Bacterial strains, plasmid, and culture conditions
M. caseolyticus KCTC 3582, E. coli ER2566, and plasmid pET-28a
(þ) were used as sources of genomic DNA encoding a putative fatty
acid hydratase, host cells, and expression vector, respectively.
M. caseolyticus was cultivated on 50 ml of growth medium con-
taining 0.5% glucose, 0.5% yeast extract, 1.0% casein peptone, and
0.5% sodium chloride in a 250 ml flask at 37 ꢀC with shaking at
200 rpm. The recombinant E. coli cells for protein expression were
cultivated in LB medium (1.0% tryptone, 0.5% yeast extract, and 1.0%
sodium chloride) in a 2000 ml flask containing 20 mg/ml kanamycin
at 37 ꢀC with shaking at 200 rpm. When the optical density of
bacteria reached 0.6 at 600 nm, IPTG was added to the culture
medium at a final concentration of 0.1 mM. The culture broth was
then incubated with shaking at 150 rpm at 16 ꢀC for 16 h to express
the enzyme.
2.5. Determination of molecular mass
The subunit molecular mass of the putative fatty hydratase from
M. caseolyticus was examined by SDS-PAGE under denaturing
conditions, using the pre-stained maker proteins as reference
proteins. All protein bands were stained with Coomassie blue for
visualization. The molecular mass of the native enzyme was
determined by gel filtration chromatography using a Sephacryl S-
300 HR 16/60 column (GE Healthcare). The purified enzyme solu-
tion was applied to the column and eluted with 50 mM PIPES buffer
(pH 6.5) containing 150 mM NaCl at a flow rate of 0.3 ml/min. The
2.3. Gene cloning and site-directed mutagenesis
The gene encoding
a putative fatty acid hydratase was
amplified by PCR using M. caseolyticus genomic DNA as the
template. The sequence of primers used for gene cloning
was based on the DNA sequence of the putative fatty acid
hydratase (MCRA) from M. caseolyticus JCSC5402 (GenBank
accession number NC_011999.1). Forward (50-GGGGCTAGCATG-
TACTATAGTAATGGA-30) and reverse primers (50-AGGCTCGAGT-
TATATCAAATTTGCTTC-30) were designed to introduce the
underlined NheI and XhoI restriction sites and were synthesized
by Bioneer (Daejon, Korea). The amplified DNA fragment obtained
by PCR with pfu polymerase (Solgent, Daejon, Korea) was
extracted using the PCR purification kit (Promega, Fitchburg, WI,
USA) and was ligated into the NheI and XhoI sites of pET-28a(þ).
The resulting plasmid was used to transform E. coli ER2566 as an
expression host. Expression of the gene encoding the putative
fatty acid hydratase was determined by SDS-PAGE. Mutations of
the five conserved FAD-binding residues in the putative fatty acid
hydratase were generated by site-directed mutagenesis using
a QuikChange kit and protocol (Stratagene, Beverly, MA, USA).
DNA sequencing was performed at the Macrogen facility (Seoul,
Korea).
column was calibrated with appoferritin (443 kDa),
b-amylase
(200 kDa), albumin (66 kDa), and carbonic anhydrase (29 kDa) as
reference proteins (GE Healthcare), and the mass of the native
enzyme was calculated by comparing with the migration length of
reference proteins.
2.6. Enzyme assay
Fatty acid substrates were dispersed to a concentration of 4 mM
in 4% (v/v) ethanol and the solution was homogenized at
10,000 rpm for 10 s using a homogenizer (IKA, Kuala Lumpur,
Malaysia). The substrate and enzyme solutions were mixed in 1:1
(v/v) ratio. Unless otherwise stated, the enzymatic reaction was
performed at 25 ꢀC in 50 mM PIPES buffer (pH 6.5) containing 2 mM
fatty acid, 2% (v/v) ethanol, 33 unit/ml (0.01 mg/ml) of enzyme, and
0.2 mM FAD for 10 min. To stop the reaction, the solution was
acidified to pH 2.0 by adding 6 M HCl. One unit of the putative fatty
hydratase activity was defined as the amount of enzyme required to

Y.-C. Joo et al. / Biochimie 94 (2012) 907e915
909
produce 1 nmol of 10-hydroxyoctadecanoic acid per min at 25 ꢀC
and pH 6.5. To measure specific activity, the enzymatic reactions
were performed in 50 mM PIPES buffer (pH 6.5) containing 2 mM
fatty acid at 25 ꢀC for 10 min by adjusting the enzyme concentration
from 33 to 3300 unit/ml.
wild-type and variant enzymes, the enzyme solutions were incu-
bated at 4 ꢀC for 16 h. The unbound FAD in the enzyme solutions
was removed by untrafiltration using the Centricon (50 kDa cutoff),
and the resulting solutions were heated at 100 ꢀC for 15 min. After
heating, the precipitated proteins were removed by centrifugation
at 13,000 ꢁ g for 10 min, and the supernatant was used for the
analysis of UVevisible spectra. The concentration of bound FAD to
enzyme in the supernatant was determined using a linear cali-
bration curve relating optical density at 450 nm and FAD concen-
2.7. Preparation of hydroxy fatty acid standards
The hydroxy fatty acid standards were prepared by solvent
fractional crystallization at low temperature [11]. The enzymatic
reaction was performed at 25 ꢀC in 50 mM PIPES buffer (pH 6.5)
containing 10 mM fatty acid, 2% (v/v) ethanol, and 3300 unit/ml of
enzyme for 16 h. The reaction products thus obtained were extrac-
ted with an equal volume of ethyl acetate. The solvent was removed
from the extract using a rotary evaporator. A mixture of 30%
acetonitrile and 70% acetone was then added to the extracted fatty
acid solution at room temperature. The solution was cooled in an
ultra low temperature freezer for 24 h at ꢂ80 ꢀC. The liquid, which
comprised the unsaturated fatty acid fraction, was removed at room
temperature. The solvent was removed from the solid extract using
a rotary evaporator to obtain the hydroxy fatty acid fraction. This
fractionization procedure was repeated three times. The hydroxy
fatty acid products were obtained with high purity (>99%) and were
used as standard compounds in subsequent analyses.
tration ranging from 0 to 150 mM (Supplementary Fig. 1).
2.10. Analytical methods
The enzymatic reaction solution was extracted with an equal
volume of ethyl acetate. Solvent was removed from the extract
using a rotary evaporator. The obtained fatty acids were silylated
with a 2:1 mixture of pyridine and N-methyl-N-(trimethylsilyl)
trifluoroacetamide [13]. Silylated fatty acids in the organic phase
were analyzed by a GC (Agilent 6890N, Santa Clara, CA, USA)
equipped with a flame ionization detector and a Supelco SPB-1
capillary column. The column temperature was increased from
150 to 210 ꢀC for 15 min and then maintained at 210 ꢀC. The injector
and detector were held at 260 and 250 ꢀC, respectively. The hydroxy
fatty acid products were identified by GCeMS (Agilent 5973N) with
an electron impact ionization source. The ion source was operated
at 70 eV and held at 230 ꢀC.
2.8. Effects of reaction conditions on enzyme activity
The putative fatty hydratase was obtained after incubation at
25 ꢀC with 10 mM EDTA for 1 h followed by overnight dialysis at
4 ꢀC against 50 mM PIPES buffer (pH 6.5). The effect of metal ions on
enzyme activity was investigated using the EDTA-treated enzyme
3. Results
3.1. Gene cloning and molecular mass of the putative fatty acid
hydratase
in the presence of 1 mM Mg2þ, Ca2þ, Co2þ, Mn2þ, Ni2þ, Ba2þ, Zn2þ
,
Fe2þ, or Cu2þ. To examine the effect of pH on enzyme activity, the
pH was varied from 5.5 to 7.5 using 50 mM MES buffer (pH 5.5e6.5)
and 50 mM PIPES buffer (pH 6.5e7.5). The effect of temperature on
enzyme activity was investigated by varying the temperature from
15 to 35 ꢀC. The effect of temperature on enzyme stability was
evaluated over the same temperature range for 120 min. Samples
were withdrawn at specific time intervals and the residual activity
of each sample was measured.
The effect of detergent on enzyme activity was examined using
0.05% (w/v) Span 80, Tween 80, and Triton X-100. Several solvents,
including ethanol, methanol, acetone, isopropyl alcohol, 1-butanol,
and toluene were added to the enzymatic reaction solution at
a concentration of 2% (v/v) to find the optimum solvent for enzyme
activity. The effect of ethanol concentration on enzyme activity was
investigated by varying its concentration from 0 to 4% (v/v). To
remove cofactors, urea was added to the purified enzyme at a final
concentration of 3 M and then the enzyme was dialyzed at 4 ꢀC for
24 h against 50 mM PIPES buffer (pH 6.5) [12]. This procedure has
a negligible impact on enzyme activity. The reconstituted apoen-
zyme was concentrated to 10 mg/ml by ultrafiltration using the
Centricon. After adding 0.2 mM FAD, FMN, NADþ, or NADPþ to the
apoenzyme, the enzyme solution was incubated at 4 ꢀC for 24 h.
The effect of FAD concentration was investigated by varying the
FAD concentration from 0 to 1.0 mM.
The MCRA gene (1770 bp), which was previously proposed as
a fatty acid hydratase gene from M. caseolyticus, with the same
sequence reported in GenBank (accession number NC_011999.1),
was cloned and expressed in E. coli. The expressed enzyme was
purified from the crude extract obtained from harvested cells by
His-Trap HP affinity chromatography. The putative fatty acid
hydratase from M. caseolyticus was purified with a purification of 9-
fold, a yield of 38%, and a specific activity of 3300 unit/mg. The
purified enzyme contained a hexa-histidine tag at the N terminus.
The non-tagged enzyme was obtained from the removal of histi-
dine tag of the histidine-tagged enzyme by treating thrombin. The
specific activity of the non-tagged enzyme was almost the same as
that of the histidine-tagged enzyme (Supplementary Table 1).
The purified protein showed a single band in SDS-PAGE with
a molecular mass of approximately 68 kDa (Fig. 1A), which is
consistent with the calculated value of 68.1 kDa based on the 589-
residue amino acids plus the hexa-histidine tag. Based on the
masses of the reference proteins, the native protein existed as
a dimer with a molecular mass of 136 kDa as determined by gel
filtration chromatography (Fig. 1B). Fatty acid hydratases from
E. meningoseptica, S. pyogenes, and B. breve consisted of 646, 598,
and 625 amino acid residues, respectively, and their subunit
molecular masses were 66e70 kDa. The total molecular mass of
fatty acid hydratase from S. pyogenes is 134 kDa as a dimer [8].
2.9. FAD assay
3.2. Identification of hydroxy fatty acid products formed by the
putative fatty acid hydratase
UVevisible absorption spectra of FAD, apoenzyme, and holo-
enzymes were measured from 300 to 600 nm using a Beckman
Coulter DU 800 UVevisible spectrometer (Brea, CA, USA). Spectral
measurements of the wild-type and variant enzymes were per-
The conversion of unsaturated fatty acid substrates into hydroxy
fatty acid products catalyzed by the putative fatty acid hydratase
from M. caseolyticus were analyzed using GC. The products were
purified by solvent fractional crystallization at low temperature
[11] and the purity of the products was greater than 99% as
formed in 50 mM PIPES buffer (pH 6.5) containing 20
mM FAD and
1 mg/ml enzyme at 25 ꢀC. After adding 1 mM FAD to 10 mg/ml

910
Y.-C. Joo et al. / Biochimie 94 (2012) 907e915
A
B
kDa
1
2
3
1000
800
600
443
400
170
130
95
72
55
200
68 kDa
200
136 kDa
100
80
43
34
26
66.2
60
40
29
20
10
120
130
140
150
160
Retention time (min)
170
180
190
17
Fig. 1. SDS-PAGE analysis and molecular mass determination of the putative fatty acid hydratase from M. caseolyticus. (A) SDS-PAGE analysis. Lane 1, molecular mass marker
proteins; lane 2, crude extract; lane 3, purified putative fatty acid hydratase. (B) Determination of molecular mass of the purified native enzyme by gel filtration chromatography.
The reference proteins were apoferritin (443 kDa), b-amylase (200 kDa), albumin (66 kDa), and carbonic anhydrase (29 kDa).
determined by GC. The purified hydroxy fatty acids were used as
the standard compounds after identification by GCeMS.
The silylated enzymatic product obtained from oleic acid gave
the mass spectra shown in Fig. 2A. Peaks at m/z 229 and 331 were
resulted from the cleavage of the hydroxyl group at the C10 position
because of the loss of C12H27OSi and C8H17, respectively, from the
silylated hydroxy fatty acid, which is represented by peak at m/z
444. A peak at m/z 215 was formed by the loss of C12H25O2Si. These
Fig. 2. Chemical structures and GC/MS spectra of the silylated (A) 10-hydroxyoctadecanoic acid and (B) 10,13-dihydroxyoctadecanoic acid products obtained from oleic acid and
linoleic acid by the putative fatty acid hydratase from M. caseolyticus, respectively.

Y.-C. Joo et al. / Biochimie 94 (2012) 907e915
911
fragment peaks identified that the hydroxy fatty acid was a 10-
A
hydroxyoctadecanoic acid. The mass spectra of the enzymatic
fatty acid products formed from myristoleic acid, palmitoleic acid,
linoleic acid, and a-linolenic acid also contained peaks at m/z 229
and 331, indicating that these compounds were 10-hydroxy fatty
acids.
The mass spectrum of silylated dihydroxy fatty acid obtained
from the enzymatic reaction for linoleic acid (Fig. 2B) contained
peaks at m/z 303 and 331 and at 173 m/z and 461 resulting from the
cleavage of the hydroxyl groups at the C10 and C13 positions,
respectively. These results fragment peaks identified that the
dihydroxy fatty acid was a 10,13-dihydroxyoctadecanoic acid. The
250
200
150
100
50
dihydroxy fatty acid product formed from a-linolenic acid yielded
a mass spectrum with peaks at m/z 301 and 331 and at m/z 171 and
461, indicating two hydroxyl groups at the C10 and C13 positions,
respectively. The dihydroxy fatty acid was identified as a 10,13-
dihydroxy-15(Z)-octadecenoic acid due to these fragment peaks.
Peaks at m/z 227 and 329 in the mass spectrum of the hydroxy fatty
0
0
200
400
600
800
1000
acid formed from
the C10 position. The mass spectrum of the silylated dihydroxy fatty
acid obtained from -linolenic acid exhibits peaks at m/z 303 and
g-linolenic acid also indicate a hydroxyl group at
Time (min)
g
329 and at m/z 173 and 459 resulting from the cleavage of the
hydroxyl group at C10 and C13 positions, respectively. The double
B
bond between C6 and C7 of g-linolenic acid caused a difference of
2 m/z units between the product peaks (Table 1). The mass spec-
trum indicated that the dihydroxy fatty acid was a 10,13-dihydroxy-
6(Z)-octadecenoic acid.
3.3. Conversion of unsaturated fatty acids into hydroxy fatty acids
by the putative fatty acid hydratase
The enzymatic conversion of oleic acid was examined under
the standard conditions described above. After 840 min, the
conversion of oleic acid to 10-hydroxyoctadecanoic acid occurred
with a molar yield of 98% (Fig. 3A). The enzymatic reaction did not
occur with 2 mM 10-hydroxyoctadecanoic acid as a substrate.
Thus, the enzymatic reaction was irreversible (Fig. 3B). The
enzyme also converted myristoleic acid, palmitoleic acid, and oleic
acid into 10-hydroxytetradecanoic acid, 10-hydroxyhexadecanoic
acid, and 10-hydroxyoctadecanoic acid, respectively, with no
other byproducts (Supplementary Fig. 2A, B, and C). Linoleic acid,
Fig. 3. Irreversible catalytic reaction of oleic acid into 10-hydroxyoctadecanoic acid by
oleate hydratase from M. caseolyticus. (A) Time-course reaction. The conversion reac-
tions of oleic acid (B) into 10-hydroxyoctadecanoic acid (C). The reactions were
performed by varying the reaction time in 50 mM PIPES buffer (pH 6.5) containing
2 mM oleic acid and 33 unit/ml enzyme at 25 ꢀC for 840 min. Data represent the means
of three experiments and error bars represent standard deviation. (B) Reaction
mechanism. The oleate hydratase hydrated oleic acid at its cis-9-double bond to yield
10-hydroxyoctadecanoic acid.
3.4. Identification of the putative fatty acid hydratase
a
-linolenic acid, and g-linolenic acid were each converted into the
two products 10-hydroxy-12(Z)-octadecenoic acid and 10,13-
dihydroxyoctadecanoic acid, 10-hydroxy-12(Z),15(Z)-octadecadie-
noic acid and 10,13-dihydroxy-15(Z)-octadecenoic acid, and 10-
hydroxy-6(Z),12(Z)-octadecadienoic acid and 10,13-dihydroxy-
6(Z)-octadecenoic acid, respectively (Supplementary Fig. 2D, E,
and F).
The control experiment for the enzymatic reaction was per-
formed with heat-inactivated enzyme or without the enzyme
under the same reaction conditions. In the experiment, no hydroxy
fatty acids were detected. The substrate specificity of the putative
fatty hydratase was evaluated by determining the concentrations of
Table 1
Specific activity and kinetic parameters of the putative fatty acid hydratase from M. caseolyticus for fatty acids as substrates.
Substrate
Product(s)
GC retention
time (min)
GC/MS fragments (m/z)
Specific
activity
Km
(
mM
)
kcat
kcat/Km
(minꢂ1
)
(mmꢂ1 minꢂ1
)
(unit/mg)
Myristoleic acid
Palmitoleic acid
Oleic acid
10-Hydroxytetradecanoic acid
10-Hydroxyhexadecanoic acid
10-Hydroxyoctadecanoic acid
10-Hydroxy-12(Z)-octadecenoic acid
10,13-Dihydroxyoctadecanoic acid
7.15
10.26
13.76
13.39
13.60
388(M), 331, 229, 159
416(M), 331, 229, 187
444(M), 331, 229, 215
442(M), 331, 229, 213
532(M), 461, 331, 303, 173
440(M), 331, 229, 211
530(M), 461, 331, 301, 171
440(M), 329, 227, 213
530(M), 459, 329, 303, 173
18 ꢃ 0.2 300 ꢃ 7
2 ꢃ 0.0
230 ꢃ 4
470 ꢃ 1
8 ꢃ 0.2
1 ꢃ 0.1
8 ꢃ 0.2
2 ꢃ 0.6
160 ꢃ 1
12 ꢃ 0.2
8 ꢃ 0.2
390 ꢃ 6
1360 ꢃ 5
20 ꢃ 0.4
3 ꢃ 0.6
1680 ꢃ 8
570 ꢃ 9
340 ꢃ 2
3300 ꢃ 12
Linoleic acid
58 ꢃ 0.9 390 ꢃ 1
5 ꢃ 0.8 340 ꢃ 2
a
-Linolenic acid
-Linolenic acid
10-Hydroxy-12(Z),15(Z)-octadecadienoic acid 13.45
59 ꢃ 2.9 320 ꢃ 2
17 ꢃ 0.9 330 ꢃ 4
1120 ꢃ 1.2 590 ꢃ 2
90 ꢃ 0.9 600 ꢃ 4
25 ꢃ 0.6
8 ꢃ 0.2
10,13-Dihydroxy-15(Z)-octadecenoic acid
10-Hydroxy-6(Z),12(Z)-octadecadienoic acid
10,13-Dihydroxy-6(Z)-octadecenoic acid
13.65
12.72
12.84
g
270 ꢃ 0.1
21 ꢃ 0.5
ND, kinetic parameters are not detected by the analytical methods used in this study.
Data represent the mean of three experiments and ꢃ values represent standard deviations.

912
Y.-C. Joo et al. / Biochimie 94 (2012) 907e915
the reaction products by GC and then identifying the products
by GCeMS. No enzymatic activity was observed for lauric acid
(C12), myristic acid (C14), palmitic acid (C16), stearic acid
(C18), arachidic acid (C20), petroselinic acid (C18:166Z), elaidic
acid (C18:169E), vaccenic acid (C18:1611Z), conjugated linoleic
acids (C18:269E,11E, C18:269Z,11E, and C18:2610E,12Z), arachidonic
acid (C20:465Z,8Z,11Z,14Z), erucic acid (C22:1613Z), or nervonic
acid (C22:1615Z). However, the enzyme exhibited some activity
for myristoleic acid (C14:169Z), palmitoleic acid (C16:169Z), oleic
The enzyme exhibited the highest catalytic efficiency (kcat/Km) for
oleic acid among the substrates tested. The kcat/Km for oleic acid
(1359 mMꢂ1 minꢂ1) was 3.5-fold higher than that for the second
favored substrate palmitoleic acid (387 mMꢂ1 minꢂ1). These results
indicate that the putative fatty acid hydratase was an oleate
hydratase.
3.5. Effects of reaction conditions on the activity of the oleate
hydratase
acid (C18:169Z), linoleic acid (C18:269Z,12Z),
a-linolenic acid
(C18:369Z,12Z,15Z), and -linolenic acid (C18:366Z,9Z,12Z). These
g
The purified enzyme obtained after the removal of metal ions
by treating 10 mM EDTA exhibited 90% activity of the native enzyme
activity. The addition of EDTA to oleate hydratase from
E. meningoseptica had no effect on enzyme activity [10]. After adding
divalent metal ions to the EDTA-treated oleate hydratase, the activity
was reduced to 25ꢂ44% of the native enzyme activity, following the
order Mg2þ > Ca2þ > Co2þ > Mn2þ > Ni2þ > Ba2þ > Zn2þ > Fe2þ
(Supplementary Fig. 3A). The addition of Cu2þ resulted in no
activity. Maximum enzyme activity was observed at pH 6.5 and 25 ꢀC
(Fig. 4A and B). The effect of temperature on enzyme stability was
investigated. After 120 min, the enzyme activity nearly unchanged
at 25 ꢀC, but was reduced to only 25% at 35 ꢀC (Supplementary
Fig. 3B). Enzyme activity was decreased with the addition of any
detergent (Supplementary Fig. 3C). Among the solvents tested,
ethanol was the most beneficial for enzyme activity (Supplementary
Fig. 3D), with an optimal concentration of 2% (v/v) (Fig. 4C). The
addition of the cofactor FMN, NADþ, or NADPþ did not affect enzyme
results indicate that the putative fatty acid hydratase had no
specificity for saturated fatty acids and trans-, cis-5-, cis-6-, cis-8-,
cis-11-, cis-13-, cis-14-, and cis-15-double bond unsaturated fatty
acids, but had for cis-9- and/or cis-12-double bond unsaturated
fatty acids without trans-configurations. Thus, cis-9 unsaturated
fatty acids were converted into 10-hydroxy fatty acids as a single
product, cis-9 and cis-12 unsaturated fatty acids were converted
into the two products 10-hydroxy fatty acids and 10,13-dihydroxy
fatty acids.
The specific activity, kcat, and kcat/Km of the enzyme followed the
order oleic acid > palmitoleic acid >
product) > -linolenic acid (the slower product) >
(the faster product) > linoleic acid (the faster product) > myristoleic
acid > -linolenic acid (the slower product) > linoleic acid (the
g-linolenic acid (the faster
g
a-linolenic acid
a
slower product) (Table 1). The faster and slower products were 10-
hydroxy fatty acids and 10,13-dihydroxy fatty acids, respectively.
A
B
120
120
100
80
60
40
20
0
100
80
60
40
20
0
5.0
5.5
6.0
6.5
7.0
7.5
8.0
15
20
25
30
35
o
Temperature ( C)
pH
C
D
140
2.0
1.5
1.0
0.5
0.0
120
100
80
60
40
5
0
1
2
3
4
0.0
0.2
0.4
0.6
0.8
1.0
Ethanol (%, v/v)
FAD (mM)
Fig. 4. Effects of pH, temperature, and concentrations of ethanol and FAD on the activity of oleate hydratase from M. caseolyticus. (A) Effect of pH. The reactions were performed in
50 mM MES buffer for pH 5.5 to 6.5 (C) and 50 mM PIPES buffer for pH 6.5 to 7.5 (B) with 2 mM oleic acid and 33 unit/ml enzyme at 25 ꢀC for 10 min. (B) Effect of temperature. The
reactions were performed in 50 mM PIPES buffer (pH 6.5) containing 2 mM oleic acid and 33 unit/ml enzyme for 10 min in temperature range of from 15 to 35 ꢀC. (C) Effect of
ethanol concentration. The reactions were performed by varying the concentration of ethanol in 50 mM PIPES buffer (pH 6.5) containing 2 mM oleic acid and 33 unit/ml enzyme at
25 ꢀC for 10 min. (D) Effect of FAD concentration. The reactions were performed by varying the concentration of FAD in 50 mM PIPES buffer (pH 6.5) containing 2 mM oleic acid and
33 unit/ml enzyme at 25 ꢀC for 10 min. 0 mM FAD indicates the apoenzyme without the addition of FAD. Data represent the means of three experiments and error bars represent
standard deviation.

Y.-C. Joo et al. / Biochimie 94 (2012) 907e915
913
activity. But, when FAD was not added to the apoenzyme, all catalytic
activity was abolished (Fig. 4D), and enzyme activity increased with
increasing FAD concentration, reaching a plateau above 0.2 mM.
The maximum activity of oleate hydratase from M. caseolyticus
was observed at pH 6.5 and 25 ꢀC with 2% (v/v) ethanol and 0.2 mM
FAD. The maximum activity of E. meningoseptica oleate hydratase
occurred at pH 6.0 with 5% isopropyl alcohol and 50 mM NaCl [10].
However, the effects of temperature and solvent were not reported.
The reaction conditions of S. pyogenes fatty acid hydratase were
reported to be pH 6.1, 37 ꢀC, 2% ethanol, and 0.02 mM FAD [8]. The
reaction of B. breve fatty acid hydratase was performed at pH 7.5
and 25 ꢀC [9].
enzymes after heat precipitation were similar to those of the
enzymes before heat precipitation (Supplementary Fig. 6). These
results mean that a non-covalently bound FAD is fully released in
the solution upon protein denaturation by heat. After heat precip-
itation, the FAD concentration of the supernatant of the wild-type
was 141
mM in 145 mM wild-type enzyme (Fig. 6), indicating that
the coenzyme FAD was bound to apoenzyme with a ratio of 1:1. The
bound FAD concentrations followed the order wild-type enzyme
(141
m
M) > E50A (98
M) > G31A (8
The specific activities and kcat/Km values of the S34A and E50A
m
mM).
M) > S34A (75
mM) > E56A (25 mM) > G29A
(19
m
variants for oleic acid were 60e85% those of the wild-type enzyme
(Table 2). The affinity and kcat of these variants for oleic acid were
also lower than those of the wild-type enzyme. Alanine-
substitution of G29, G31, and E56 resulted in loss of enzyme
activity.
3.6. Spectral and kinetic analyses of FAD-binding residues in the
oleate hydratase
The wild-type holoenzyme exhibited two peaks at 375 and
445 nm as a typical UVevisible absorption spectrum of an FAD-
dependent enzyme. These peaks were not present in the absorp-
tion spectrum of the apoenzyme obtained after the removal of FAD
(Supplementary Fig. 4). The five conserved residues of G29, G31,
S34, E50, and E56 in the FAD-binding motif of fatty acid hydratases
(Supplementary Fig. 5) were individually replaced with alanine.
The UVevisible absorption spectra of S34A and E50A displayed
reduced peaks at 375 and 445 nm relative to those of the wild-type
enzyme (Fig. 5A, D, and E), and the FAD-dependent peaks of G29A,
G31A, and E56A disappeared (Fig. 5B, C, and F). The heat precipi-
tation was performed to release FAD in the enzyme. The UVevisible
absorption spectra of the supernatant of the wild-type and variant
4. Discussion
The MCRA protein family is widely distributed in bacteria,
especially pathogenic and intestinal bacteria. MCRA was found first
in a strain of S. pyogenes that causes human pathogenesis, including
acute rheumatic fever and heart tissue damage [14]. The MCRA of
pathogenic bacteria can survive in blood with the aid of human
keratinocytes for adherence and internalization [8] and are less
sensitive to heat and solvent stresses [9,15]. The MCRA of intestinal
bacteria is involved in anti-microbial [3,16] and anti-stress effects
[17]. Many bacteria are sensitive to fatty acids, and unsaturated
fatty acids are much more toxic than saturated fatty acids [18,19].
Fig. 5. UVevisible absorption spectra of FAD-containing wild-type, and variant enzymes. (A) Wild-type. (B) G29A. (C) G31A. (D) S34A. (E) E50A. (F) E56A. The concentrations of the
wild-type and variant enzymes were approximately 1 mg/ml in 50 mM PIPES buffer (pH 6.5).

914
Y.-C. Joo et al. / Biochimie 94 (2012) 907e915
(Fig. 2, Table 1) and used as standards in subsequence analyses.
Based on the quantitative determination of the substrate specificity,
the putative fatty acid hydratase was identified as an oleate
hydratase. The enzyme exhibited hydration activity for myristoleic
acid, palmitoleic acid, oleic acid, linoleic acid, and
acids. Fatty acid hydratase from S. pyogenes recognized specifically
palmitoleic acid, oleic acid, linoleic acid, -linolenic acid but not
myristoleic acid and -linolenic acid [8]. Fatty acid hydratase from
B. breve showed activity for palmitoleic acid, oleic acid, linoleic acid,
but not for myristoleic acid and -linolenic acid [9]. The hydration
activity of this enzyme for -linolenic acid was not investigated.
a- and g-linolenic
a
g
a
g
The enzyme from Flavobacterium sp. DS5 involved in the hydration
of unsaturated fatty acids was suggested to be a hydratase [23,24].
Flavobacterium sp. DS5 [24] and Stenotrophomonas nitritireducens
[25] had hydration activities for myristoleic acid, palmitoleic acid,
oleic acid, linoleic acid, and a- and g-linolenic acids, indicating that
the substrate specificity of these bacteria is the same as that of
oleate hydratase from M. caseolyticus. Unfortunately, the different
specificity of the fatty acid hydratases and bacteria cannot be
elucidated because the crystal structure and active site of hydratase
have not been determined.
Fig. 6. Concentrations of bound FAD to the wild-type and variant enzymes. The
concentrations of the wild-type and variant enzymes were approximately 10 mg/ml in
50 mM PIPES buffer (pH 6.5). After heat precipitation, the FAD concentration of the
supernatant was measured at 450 nm of optical density.
It has been suggested that the conversion of oleic acid into 10-
hydroxyoctadecanoic acid using crude extract from Pseudomonas
sp. strain 3266 is a reversible reaction [10,26]. This putative enzyme
involved in the study was referred to as an oleate hydratase,
although the enzyme was not purified and characterized. The strain
converted oleic acid into not only 10-hydroxyoctadecanoic acid but
also 10-oxo-octadecanoic acid, suggesting that other fatty acid-
converting enzymes must be present in the crude extract. In the
present study, the purified oleate hydratase did not convert 10-
hydroxyoctadecanoic acid into oleic acid. The conversion of oleic
acid with D2O and/or H128O by Nocardia cholesterolicum [27] and
Pseudomonas sp [28]. showed the incorporation of D at the C9
position and 18O at the C10 position of 10-hydroxyoctadecanoic
acid. Therefore, oleate hydratase catalyzes the irreversible addi-
tion of a hydrogen atom and a hydroxy group from water to the
carbonecarbon double bond at the C9 and C10 positions, respec-
tively, to make 10-hydroxyoctadecanoic acid (Fig. 3B).
Spectral analyses [29] and kinetic analyses of variants [12] have
been widely used for the characterization of dinucleotide-
dependent enzymes. When FAD was not bound to the enzyme, its
catalytic activity was abolished (Fig. 4D). These results indicate that
the oleate hydratase from M. caseolyticus is FAD-dependent. All
members of the glutathione reductase (GR) family, including fatty
acid hydratases, contain a FAD-binding motif such as a GXGXXS(A/
G)X15E(D)X5E (where X denotes any amino acid) sequence at the N
terminus [30,31]. The roles in FAD-binding for the five conserved
residues of G29, G31, S34, E50, and E56 in the FAD-binding motif
were investigated. The absorption peaks at 375 and 445 nm, bound
FAD amounts, specific activities, and kcat values of S34A and E50A
were all reduced by alanine-substitution (Figs. 5 and 6, and Table 2).
G29A, G31A, and E56A exhibited no adsorption peaks and no
enzyme activity. The bound FAD concentrations of S34A and E50A
were more than 50% of the wild-type enzyme FAD, whereas those
of G29A, G31A, and E56A were less than 20%. Mutations at G29,
G31, and E56 resulted in significantly reduced molecular interac-
tions with FAD and resulted in no activity. Thus, G29, G31, and E56
are essential for FAD-binding and S34 and E50 are involved in FAD-
binding. UVevisible absorption spectra of the holoenzyme of
M. caseolyticus oleate hydratase were measured at 30 s intervals for
10 min (Supplementary Fig. 7A). The spectra changed very little,
indicating that the oxidationereduction reaction of FAD did not
occur. To investigate the effect of the addition of FAD on enzyme
stability, the reactions were performed after incubating at 25 ꢀC for
24 h with 1 mg/ml purified enzyme in the presence and absence of
Unsaturated fatty acids inhibit the development of cellular
membranes because their kinked structure disrupts the lipid
bilayer formation [20] and inhibits enoyl-ACP reductase, which
involved in bacterial fatty acid synthesis [21]. The hydration of
unsaturated fatty acids is suggested to be a detoxification mecha-
nism in bacteria harboring MCRA proteins and a survival strategy
for living in fatty acid-rich environments [8]. Thus, MCRA proteins
are fatty acid hydratases.
The amino acid sequence of oleate hydratase from
M. caseolyticus exhibited 57, 55, and 41% identity with the charac-
terized fatty acid hydratases from S. pyogenes [8], B. breve [9], and
E. meningoseptica [10], respectively, and showed 11, 10, and 10%
identity with the other oleate-converting enzymes P450 mono-
oxygenase from B. licheniformis [22], oleate hydroxylase from the
plant R. communis L [6], and lipoxygenase from Pseudomonas 42A2
[7], respectively. These results indicate that the oleate hydratase is
much more closely related to fatty acid hydratases than to other
oleate-converting enzymes.
The substrate specificity of fatty acid hydratases for fatty acids
was not investigated quantitatively and the name of the specific
hydratase was not defined because the hydroxy fatty acid products
of the previously reported fatty acid hydratases are not commer-
cially available and have not been isolated [8e10]. In the present
study, all of the hydroxy fatty acids produced by the putative fatty
acid hydratase from M. caseolyticus were purified, and the purified
products (>99%) (Supplementary Fig. 2) were identified by GCeMS
Table 2
Kinetic parameters of the wild-type and variant enzymes in the FAD-binding motif
of oleate hydratase from M. caseolyticus.
Enzyme
Specific activity
(unit/mg)
Km
(
mM)
kcat (minꢂ1
)
kcat/Km
(mMꢂ1 minꢂ1
)
Wild-type
G29A
3300 ꢃ 54
ND
340 ꢃ 2
ND
470 ꢃ 1
ND
1360 ꢃ 5
ND
G31A
ND
ND
ND
ND
S34A
E50A
E56A
2540 ꢃ 25
2820 ꢃ 36
ND
380 ꢃ 5
390 ꢃ 7
ND
300 ꢃ 4
440 ꢃ 5
ND
800 ꢃ 1
1140 ꢃ 2
ND
ND, kinetic parameters are not detected by the analytical methods used in this
study.
Data represent the mean of three experiments and ꢃ values represent standard
deviations.

Y.-C. Joo et al. / Biochimie 94 (2012) 907e915
915
[10] L.E. Bevers, M.W. Pinkse, P.D. Verhaert, W.R. Hagen, Oleate hydratase cata-
lyzes the hydration of a nonactivated carbon-carbon bond, J. Bacteriol. 191
(2009) 5010e5012.
1 mM FAD (Supplementary Fig. 7B). The production of 10-
hydroxystearic acid from oleic acid by the enzyme in the absence
of FAD was 7-fold lower than that in the presence of 1 mM FAD.
Thus, FAD in the oleate hydratase does not seem to be involved in
the hydration reaction itself but rather in the structural stabiliza-
tion of the protein.
In the present study, the putative fatty acid hydratase from
M. caseolyticus was identified as an oleate hydratase by character-
izing the quantitative biochemical properties of the enzyme. The
enzyme exhibited hydration activity only for cis-9- and/or cis-12-
double bond unsaturated fatty acids without trans-configurations,
irreversibly producing 10-hydroxy fatty acids and 10,13-dihydroxy
fatty acids. The oleate hydratase was FAD-dependent, and G29,
G31, and E56 were essential for FAD-binding.
[11] T.C. Chen, Y.H. Ju, An improved factional crystallization method for the
enrichment of
g-linolenic acid in borage oil fatty acid, Ind. Eng. Chem. Res. 40
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[13] J.A. Hudson, C.A. MacKenzie, K.N. Joblin, Conversion of oleic acid to 10-
hydroxystearic acid by two species of ruminal bacteria, Appl. Microbiol. Bio-
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[14] K.S. Kil, M.W. Cunningham, L.A. Barnett, Cloning and sequence analysis of
a gene encoding a 67-kilodalton myosin-cross-reactive antigen of Strepto-
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[15] A. Liavonchanka, E. Hornung, I. Feussner, M.G. Rudolph, Structure and
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Acknowledgment
[16] E.F. O’Shea, P.D. Cotter, C. Stanton, R.P. Ross, C. Hill, Production of bioactive
substances by intestinal bacteria as a basis for explaining probiotic mecha-
nisms: Bacteriocins and conjugated linoleic acid, Int. J. Food Microbiol. (2011).
doi:10.1016/j.ijfoodmicro.2011.1005.1025.
[17] S.J. O’Flaherty, T.R. Klaenhammer, Functional and phenotypic characterization
of a protein from Lactobacillus acidophilus involved in cell morphology, stress
tolerance and adherence to intestinal cells, Microbiology 156 (2010)
3360e3367.
This study was supported by a grant (Code# 000407980110)
from the Small and Medium Business Administration, Republic of
Korea.
[18] H. Galbraith, T.B. Miller, A.M. Paton, J.K. Thompson, Antibacterial activity of
long chain fatty acids and the reversal with calcium, magnesium, ergo-
calciferol and cholesterol, J. Appl. Bacteriol. 34 (1971) 803e813.
[19] C. Henderson, The effects of fatty acids on pure cultures of rumen bacteria,
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Appendix. Supplementary material
Supplementary material associated with this article can be
found, in the online version, at doi:10.1016/j.biochi.2011.12.011.
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