Oxygenation of monoenoic fatty acids by CYP175A1, an orphan cytochrome P450 from thermus thermophilus HB27

Article abstract of DOI:10.1021/bi300514j

The catalytic activity of CYP175A1 toward monooxygenation of saturated and monounsaturated fatty acids of various chain lengths (C16-C24) has been investigated to assess the enzymatic properties of this orphan thermostable cytochrome P450 enzyme. The resu

Full text of DOI:10.1021/bi300514j

Subscriber access provided by University Libraries, University of Memphis
Article
Oxygenation of monoenoic fatty acids by CYP175A1, an
orphan cytochrome P450 from Thermus thermophilus HB27
Sandeep Goyal, Shibdas Banerjee, and Shyamalava Mazumdar
Biochemistry, Just Accepted Manuscript • DOI: 10.1021/bi300514j • Publication Date (Web): 31 Aug 2012
Downloaded from http://pubs.acs.org on September 6, 2012
Just Accepted
“Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted
online prior to technical editing, formatting for publication and author proofing. The American Chemical
Society provides “Just Accepted” as a free service to the research community to expedite the
dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts
appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been
fully peer reviewed, but should not be considered the official version of record. They are accessible to all
readers and citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered
to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published
in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just
Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor
changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers
and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors
or consequences arising from the use of information contained in these “Just Accepted” manuscripts.
Biochemistry is published by the American Chemical Society. 1155 Sixteenth Street
N.W., Washington, DC 20036
Published by American Chemical Society. Copyright © American Chemical Society.
However, no copyright claim is made to original U.S. Government works, or works
produced by employees of any Commonwealth realm Crown government in the course
of their duties.

Page 1 of 45
Biochemistry
1
2
3
4
5
6
Oxygenation of monoenoic fatty acids by CYP175A1, an orphan cytochrome P450 from
Thermus thermophilus HB27†
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Sandeep Goyal1, Shibdas Banerjee1 and Shyamalava Mazumdar*
Department of Chemical Sciences, Tata Institute of Fundamental Research,
Homi Bhabha Road, Colaba, Mumbai 400005, India.
Running title: Oxygenation of monoenoic fatty acids by CYP175A1
Keywords: P450, CYP175A1, Monoenoic fatty acids, Epoxidation, Allyl-hydroxylation.
*Correspondence: Prof. S Mazumdar, Phone no.: +91 22 2278 2363, FAX: +91 22 2280 4610,
E-mail: shyamal@tifr.res.in, Web: http://www.tifr.res.in/~shyamal/
1 These two authors contributed equally to this work.
† This work was supported by Tata Institute of Fundamental Research.
1
ACS Paragon Plus Environment

Biochemistry
Page 2 of 45
1
2
3
4
Abstract
5
6
7
8
9
The catalytic activity of CYP175A1 towards monooxygenation of saturated and
monounsaturated fatty acids of various chain lengths (C16-C24) has been investigated to assess
the enzymatic properties of this orphan thermostable cytochrome P450 enzyme. The results
showed that the enzyme could catalyze the reaction of monounsaturated fatty acids but not of
saturated fatty acids. The product analyses using ESI-MS and GC-MS revealed an important
regioselectivity in the CYP175A1 catalyzed monooxygenation of the monoenoic fatty acids
depending on the ethylenic double bond (C=C) configuration. When the double bond was in cis-
configuration, an epoxy fatty acid was found to be the major product and two allyl-hydroxy fatty
acids were found to be the minor products. But, when the double bond was in trans-configuration
the product distribution was reversed. The oxygenation efficiency was found to be the highest for
Palmitoleic acid (chain length C16) but there was no direct correlation of the activity with the
chain length or the position of unsaturation of the fatty acid. Molecular docking calculations
showed that the ‘U’-type conformations of the monoenoic fatty acids are particularly responsible
for their binding in the enzyme pocket and that is also consistent with the observed
regioselectivity in the oxygenation reaction. The present results provide evidences that
CYP175A1 can catalyze regioselective oxygenation reaction of several monoenoic fatty acids
though it cannot catalyze the oxygenation of the corresponding saturated analogues. These
studies may provide critical information on the nature of the enzyme pocket and of the possible
the natural substrate of this orphan enzyme.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
2
ACS Paragon Plus Environment

Page 3 of 45
Biochemistry
1
2
3
4
Abbreviations
5
6
7
8
9
Heme: Iron protoporphyrin IX regardless of oxidation and ligation state; WT: Wild type; Tm:
Melting temperature; PDB: Research Collaboratory for Structural Bioinformatics (RCSB)
Protein Data Bank, http://www.rcsb.org/pdb/; PMSF: Phenyl methyl sulphonyl fluoride; IPTG:
Isopropyl-β-D-1-thiogalactopyranoside; TBHP: tert-Butyl hydroperoxide; m-CPBA: m-
chloroperbenzoic acid; EDTA: Ethylenediaminetetraacetic acid dipotassium salt; H2O2:
Hydrogen peroxide; NADH: β-Nicotinamide adenine dinucleotide, reduced dipotassium salt;
ESI-MS: Electrospray ionization mass spectrometry; GC-MS: Gas chromatography mass
spectrometry; CID: Collision induced dissociation; GOLD: Genetic optimization for ligand
docking.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
3
ACS Paragon Plus Environment

Biochemistry
Page 4 of 45
1
2
3
4
5
6
7
8
Cytochrome P450 forms a superfamily of hemoproteins1 that catalyze the monooxygenation of
various organic compounds2 including aromatic and aliphatic hydroxylation, N-oxidation and
also epoxidation of C=C double bonds.
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
These enzymes often catalyze regio- and stereospecific oxygenation of nonactivated
hydrocarbons under mild conditions, which is a challenging task for chemical catalysis in
organic synthesis.3 Thus, cytochrome P450s have great potential as biocatalysts to perform
numerous industrially important reactions such as synthesis of drugs and other fine chemicals.3
However, low tolerance to various solvents and high temperature have generally limited the
usefulness of cytochrome P450 for industrial application. Interestingly, two thermophilic
cytochrome P450s e.g. CYP119 and CYP175A1 have been isolated, purified and characterized4-7
in recent years. Both of them possess extreme stability (Tm = 91oC for CYP1198 and Tm = 88oC
for CYP175A15) and might be used to overcome the above limitations.
The natural substrates and the biological functions of both of these enzymes (CYP119 &
CYP175A1) are unknown9 and therefore they are considered as the so-called orphan P450.10
However, there have been recent advances in using an engineered CYP119 to catalyze the fatty
acid hydroxylation11 and mutant Q67G/Y68I of CYP175A1 to catalyze testosterone
hydroxylation.12 Native CYP175A1 has been used to hydroxylate β-carotene at 3- and 3'-
positions.13
CYP175A1 shows the highest sequence identity of 26% with cytochrome P450BM-3
(CYP102A1)5 from Bacillus megaterium among all the structurally characterized bacterial
cytochrome P450 enzymes. A close inspection of the substrate binding pocket from the crystal
structure of CYP175A1 and CYP102A1 shows that the pockets/active sites of these two enzymes
are similar and mostly surrounded by hydrophobic residues. Hence, both of them should prefer
4
ACS Paragon Plus Environment

Page 5 of 45
Biochemistry
1
2
3
4
5
6
7
8
9
hydrophobic substrates to show catalytic activity. Long chain saturated fatty acids (Lauric acid,
myristic acid, palmitic acid and stearic acid) were found to be the natural substrate of P450BM-
3.14 However, no substrate other than β-carotene13 has been reported for the native CYP175A1
so far. It is known that Thermus thermophilus HB27 produces lipolytic enzymes15, 16, and it has
also its own lipolytic activity.17 Therefore, we systematically screened several fatty acids
(saturated and monounsaturated) for their potential as substrates for CYP175A1.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Here we report that CYP175A1 is able to catalyze the monooxygenation reaction of
several monoenoic fatty acids though it cannot catalyze the oxygenation of the corresponding
saturated analogues. Along with the substrate specificity, we also investigated the
regioselectivity of the monooxygenation reactions of those monoenoic fatty acids both
experimentally and theoretically.
Experimental
Materials
The various components of bacteriological culture media were purchased from Himedia, India.
Ampicillin sodium salt, chloramphenicol, DNaseI (from bovine pancrease), PMSF, IPTG,
sodium cholate, anhydrous sodium sulphate, potassium bicarbonate, hexamethyldisilazane,
trimethylchlorosilane, pyridine, 70% aqueous solution of TBHP, (Trimethylsilyl) diazomethane,
NADH, and several fatty acids (saturated and monounsaturated) were purchased from Sigma
Aldrich. The lysozyme (from hen egg white) and m-CPBA was purchased from Fluka. The salt
ammonium sulphate, dipotassium hydrogen ortho phosphate and potassium dihydrogen
orthophosphate, tris-HCl (2-Amino-2-hydroxymethyl-propane-1, 3-diol hydrochloric acid) and
tris-base were purchased from USB Chemicals. The column material Phenyl Sepharose was
5
ACS Paragon Plus Environment

Biochemistry
Page 6 of 45
1
2
3
4
5
6
7
8
9
purchased from GE Healthcare Bio-Sciences AB, Uppasala. Hydroxyapetite material was
purchased from Biorad Laboratories Inc, Hercules, CA. HPLC and/or GC grade methanol,
chloroform, diethyl ether, dichloromethane, toluene, cyclohexane and other required solvents
were purchased from Merck Chemicals. EDTA and was purchased from s. d. Fine Chemicals,
India. Hydrogen peroxide was purchased from Qualigens. All other buffer components were
purchased from Qualigen/s. d. fine chemicals. Millipore water was used for buffer preparation
and for all other experimental work wherever required.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
The concentration of H2O2 was measured spectrophotometrically using the molar
extinction coefficient (ε) 39.4 M-1cm-1 at 240 nm.18 m-CPBA was re-crystallized in a 1:3 diethyl
ether/petroleum ether solution using the method described by Davies et al.19
Protein Expression and Purification
The pKK-223 plasmid, encoding the gene for WT CYP175A1, was a kind gift from Prof. V. B.
Urlacher (University of Stuttgart, Stuttgart, Germany). The wild type CYP175A1 was expressed
and purified by earlier reported method.5, 20, 21 In brief, CYP175A1 was grown by taking a single
colony of the BL21-DE3 codon plus RP cells, harboring the plasmid encoding the gene for WT
protein and inoculating it into the 2×YT media containing ampicillin (100 µg/ml) and
chloramphenicol (50 µg/ml). Expression of protein was induced by addition of IPTG (1mM final
concentration), and the culture was grown at 30oC for 48 hr. After the cell lysis, the protein was
precipitated by (NH4)2SO4 (35 to 50 % saturation), re-dissolved in buffer and purified by a
hydrophobic column (Phenyl Sepharose), and fractions having Rz (Reinheitszahl, A417/A280) ≥
1.4 were collected. The protein was further purified on a hydroxyapatite column and fractions
having Rz ≥ 1.5 were collected.20 The purified fractions were concentrated using centriprep
6
ACS Paragon Plus Environment

Page 7 of 45
Biochemistry
1
2
3
4
5
6
7
8
9
concentrators (10 kD cut-off membrane, Millipore), dialyzed in 50 mM KPi, pH 7.5 buffer and
stored at -25oC in 50 % glycerol. The concentration of CYP175A1 was estimated
spectrophotometrically from the absorbance at 418 nm (ε417 = 104 mM−1 cm−1).5 The purity of the
protein was checked by SDS PAGE, and the ESI-mass spectrum of the purified protein (See
Figure S1 in the Supporting Information) confirms the molecular weight of the protein ~44.87
kD.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
The plasmids encoding wild type putidaredoxin (PdX) and putidaredoxin reductase (PdR)
were kindly supplied by Prof. S. G. Sligar (University of Illinois, USA). The wild
type putidaredoxin (PdX) and putidaredoxin reductase (PdR) were expressed and purified using a
reported procedure22 and the concentration of PdX was determined from the absorbance at 455
nm (ε455 = 5.9 mM−1cm−1)23 and that of PdR was estimated from the absorbance at 454 nm (ε454
10.9 mM−1 cm−1).24
=
Enzymatic Oxygenation of Fatty Acids
Fatty acid oxygenation by CYP175A1 was carried out by incubating 500 µL reaction mixture
containing 20 µM CYP175A1, 600 µM fatty acid and 20 mM H2O2 in 50 mM KPi buffer (pH=
7.5) at 45oC unless otherwise mentioned. The controls for the reactions were carried out that
contained all the reactants except CYP175A1 or except H2O2 in the assay mixture. Each reaction
was allowed to proceed for 60 min unless otherwise stated. The reaction mixture was then
extracted with chloroform and the chloroform layer was collected, dried over anhydrous sodium
sulfate, filtered through 0.2 µm syringe filter and then concentrated for mass spectrometric
analysis.
The kinetic parameters for the enzymatic reaction were determined by initial rate method
at 45oC by mixing 20 mM of H2O2 to a solution containing different concentrations of the
7
ACS Paragon Plus Environment

Biochemistry
Page 8 of 45
1
2
3
4
5
6
7
8
9
substrate fatty acid (varied between 100 - 500 µM), 20 µM CYP175A1 in 50 mM buffer (pH
7.5). The reaction was stopped by acetic acid (1%) after 1 min, and the unreacted fatty acid along
with the products of the enzymatic reaction were extracted in CHCl3 and subjected to ESI-mass
spectrometric identification. The concentrations of the reactants and products were determined
from the ratio of peak currents, as well as by comparing with the ion currents from standard
solutions of the reactants and the products. The initial rates of formation of monooxygenated
fatty acid obtained from the mass spectrum were estimated at different concentrations of the
substrate and analysed by Lineweaver-Burk method to evaluate the kinetic parameters Km and
kcat of the reaction.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Synthesis of Standard Samples of the Oxygenation Products
Epoxidation of monoenoic fatty acids
The epoxidation of the ethylenic double bond present in the monoenoic fatty acid was performed
following a slight modification of the literature procedure.25 5 mg monoenoic fatty acid (16C9,
18C9, 18T9, 18C6, and 18C11, see Table 1) was reacted with 4 mg m-CPBA in 500 µL
chloroform at room temperature overnight. The epoxy fatty acid derivative was solvent extracted
with diethyl ether after addition of 1 ml 5% aqueous potassium bicarbonate solution to the
reaction mixture. These epoxy derivatives of the monoenoic fatty acids were used as standards in
the tandem mass spectrometry for identification of the enzymatic reaction products of these
substrates.
Allylic hydroxylation of monoenoic fatty acids
Allyl-hydroxy fatty acids were synthesized following a slight modification of the literature
procedure.26 100 µL 70% aqueous solution of tert-butyl hydroperoxide (TBHP) mixed with ~1.5
8
ACS Paragon Plus Environment

Page 9 of 45
Biochemistry
1
2
3
4
5
6
7
8
9
mg selenium dioxide in 200 µL dichloromethane (DCM) was stirred for 30 min This reaction
mixture was added to 200 µL of DCM solution of the monoenoic fatty acid (~25 mM, 16C9,
18C9, 18T9, 18C6, and 18C11, see Table 1) and the solution was kept at shaker for 24 hr. The
solution was subsequently mixed with 1 ml water and solvent extracted with DCM/brine
solution. The unreacted fatty acids and the reaction products (allyl hydroxy- derivatives) were
separated in the DCM phase and identified by ESI-MS, which showed formation of allylic
mono-hydroxylation products from each of the monoenoic fatty acids (16C9, 18C9, 18T9, 18C6,
and 18C11, see Table 1). There were two isomers of the allylic hydroxy- derivatives identified
by tandem mass spectrometry, and these allyl hydroxy derivatives were used as standards for
identification of the enzymatic reaction products of the monoenoic fatty acids in the present
experiment.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Esterification of the fatty acids and their derivatives for GC-MS analysis
Esterification of the fatty acids was carried out following a slight modification of the literature
procedure.27 ~1 mg fatty acid/epoxy fatty acid/hydroxy (allylic) fatty acid was dissolved in 150
µL toluene and 100 µL methanol followed by the addition of 2 M trimethylsilyl diazomethane
(~5 µL in ether). The reaction mixture was kept in a shaker for 30 min at room temperature and
subsequently concentrated to give the corresponding methyl ester of the fatty acid/epoxy fatty
acid/hydroxyl fatty acid, that was used for GC-MS analysis.
Trimethylsilylation of the hydroxyl functional group28, 29 for GC-MS analysis
The ester of the allyl hydroxy product of the monoenoic fatty acid (as described above) was
further converted to trimethylsilyl derivative of the hydroxy group for GC-MS analyses. The
hydroxy fatty acid ester was reacted with 15 µL hexamethyldisilazane and 5 µL
9
ACS Paragon Plus Environment

Biochemistry
Page 10 of 45
1
2
3
4
5
6
trimethylchlorosilane in presence of 50 µL pyridine, and subsequently the trimethylsilyl ether
derivative was extracted in cyclohexane and used for GC-MS analysis.
7
8
9
UV-Visible Absorption Study
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
The UV-Visible absorption spectra were recorded from 220-700 nm (as per needed) in quartz
cuvette having path length of 1 cm on a double beam Perkin Elmer (Lambda 750-UV/Vis/NIR)
spectrophotometer. The wavelength accuracy for the spectrophotometer was ± 0.15 nm and it
was equipped with a Peltier cell temperature controller with an accuracy of ± 0.3-0.5 oC.
Spectral titration of binding of the substrates:
The dissociation constants for various fatty acids were determined by titrating the substrate free
enzyme (~5 µM) with fatty acids at room temperature. A decrease in absorbance at Soret (417
nm) and increase in absorbance at 390 nm with the increasing concentration of fatty acids are
indicative of binding of the substrate to CYP175A1 and of the conversion of heme iron from
low-spin to high-spin state. Data were analysed by reported method30 to obtain the dissociation
constant (Kd) values from the intercept and the slope of the plot of 1/ΔA417 against 1/[Substrate]
according to the following equation:
1/ΔA417 = (Kd/(PT × Δε417)) × (1/[Substrate]) + (1/(PT × Δε417))
where,
ΔA417 is change in absorbance at 417nm.
Kd (Slope/Intercept) = dissociation constant of enzyme-substrate complex.
PT is the total concentration of the holo enzyme.
Δε417 is difference in extinction coefficient of substrate-free and substrate-bound form at 417 nm.
[Substrate] is total concentration of substrate.
Electrospray Ionization Mass Spectrometry (ESI-MS)
10
ACS Paragon Plus Environment

Page 11 of 45
Biochemistry
1
2
3
4
5
6
7
8
9
ESI-MS and collision induced dissociation (ESI-MS/CID) studies were carried out using a
Thermo Finnigan LCQ Deca Electrospray quadrupole ion trap mass spectrometer.31, 32 All the
experiments were done under identical conditions unless otherwise stated. The flow rate of the
analyte solution (in methanol) was maintained at 5 µL/min, and the solution was directly injected
through the ion source kept in negative ion mode. Nitrogen was used as sheath and auxiliary gas.
The ion source conditions were: sheath gas flow rate 25 (arbitrary unit), with no auxiliary gas
flow. Capillary temperature (for desolvation) was maintained at approximately 200oC and
capillary voltage was kept at -15 V. The ion-spray voltage and tube lens offset were maintained
at -5 kV and -30 V respectively. CID product ion spectra were acquired using an isolation width
of 2 m/z under the normalized collision energy ranged from 40-50% with wide band activation
turned on. The activation Q (related to the parameter qz in the Mathieu equation for the precursor
ions)33 of 0.250 with an activation time 30 ms was used for CID-MS/MS (tandem mass
spectrometry) studies. Helium was used as buffer as well as collision gas in the collision cell (ion
trap). Data acquisition was performed for 1 min using XCalibur software (Thermo Fisher
Scientific).
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Gas chromatography-mass spectrometry (GC-MS)
The fatty acids and their oxidized products were derivatized (esterified and/or silylated) to make
them amenable to GC-MS analysis. The GC-MS analysis was performed using a gas
chromatograph (Trace GC ultra, Thermo Finnigan Electron Corporation) coupled with an ion
trap mass spectrometer (Polaris Q, Thermo Finnigan).34 Agilent J & W GC capillary column (30
m length, 0.32 mm i.d., 0.50 µm film thickness) was utilized. Helium was used as a carrier gas
(99.99% purity) maintained at a constant flow rate of 1.6 ml/min. The GC column temperature
program was as follows: initial temperature 150°C for 2 min, from 150°C to 300°C at a rate of
11
ACS Paragon Plus Environment

Biochemistry
Page 12 of 45
1
2
3
4
5
6
7
8
9
10°C/min, finally kept at 300°C for 2 min. Analyte (in chloroform) injection (1 μl) was
performed in a split less mode and the temperature at the injector was 280°C. The MS conditions
were as follows: electron ionization (EI) mode at ionization energy of 70 eV, emission current of
250 μA, ion source and transfer line temperature at 230°C and 280°C, respectively. Standard
samples were used to identify the retention times and the products were further verified by
matching the fragmentation pattern with a database in the software provided with the instrument.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Molecular docking calculations
Docking of various fatty acids with CYP175A1 (PDB ID: 1N97) was performed using GOLD
software (version 5.1).35, 36 GOLD is an automated ligand docking program used to calculate the
possible conformations of the substrate that binds to the active site of the protein. It uses a
genetic algorithm (GA) to check the ligand conformational flexibility and partial flexibility of
the protein.37 It satisfies the fundamental requirement that the ligand must displace loosely bound
water on binding.37 The genetic algorithm parameters used are: population size 100, number of
islands 5, niche size 2, selection pressure 1.1, migration frequency 10, number of operations
100000, mutation frequency 95, and crossover frequency 95. The default speed selection was
used to avoid a potential reduction in docking accuracy. One hundred genetic algorithm (GA)
runs were performed without early termination with diverse solution generation. The substrate
conformer with the highest binding score was taken into account for prediction of possible mode
of binding of the substrate to the enzyme.
12
ACS Paragon Plus Environment

Page 13 of 45
Biochemistry
1
2
3
4
Results and Discussion
5
6
7
Binding of monoenoic fatty acids with CYP175A1
8
9
The crystal structure analysis suggested that the active site (enzyme pocket) of CYP175A1
consists predominantly of hydrophobic residues analogous to that in case of cytochrome
P450BM-3, which has high structural similarity with CYP175A1 . Since fatty acids are known
to be the substrates of P450 BM-314, we screened the binding of several saturated and
unsaturated (monoenoic) fatty acids in the enzyme pocket of CYP175A1 spectrophotometrically.
However, among all the fatty acids tested only some unsaturated monoenoic fatty acids induced
changes in the absorption spectra in the Soret region. Titration of CYP175A1 with different
monoenoic fatty acids showed an increase in absorbance at 390 nm and decrease in absorbance
at 417 nm (see Figure S2 in the Supporting Information). A typical isosbestic point at 405 nm
was observed in the difference spectra of substrate-bound versus substrate free CYP175A1 (see
Figure S2 in the Supporting Information). This result suggests a typical type I substrate binding.2
The calculated Kd values are reported in the Table 1. The observation of large dissociation
constant (Kd) values (Table 1) suggests that the substrate may be weakly bound or binding of the
substrate does not efficiently displace the axial water molecule from the heme center of the
enzyme.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Oxygenation of monoenoic fatty acids by CYP175A1
The catalytic activity of CYP175A1 was assayed using H2O2 as the source of oxidizing
equivalents.2, 20, 38-44 The high-valent redox intermediate (cpdI) of the cytochrome P450 could be
generated by the peroxide shunt pathway using H2O2 or analogues (m-CPBA,
cumenehydroperoxide, peroxyacid etc.) to catalyze the oxygenation reaction of the substrate.2, 20
13
ACS Paragon Plus Environment

Biochemistry
Page 14 of 45
1
2
3
4
5
6
A reaction mixture containing CYP175A1, fatty acids and H2O2 (see experimental section for
details) at 45oC showed the oxygenation of the monoenoic fatty acids.
7
8
9
The product and the unreacted substrate extracted in organic phase were analyzed in ESI-
MS by direct injection in the negative ion mode (see Figure 1). In the negative ion mode of ESI,
ionization of the fatty acid or oxygenated fatty acid occurs via deprotonation of the carboxylic
acid functional groups.33, 45 The ionization efficiency45 of both substrate (fatty acid) and product
(oxygenated fatty acid) were found to remains essentially the same. Control experiments with
known concentrations of the fatty acids and their oxygenated derivatives showed almost the
same peak current per M of the sample (e.g., a control mixture of 500 M of 16C9 and 500 M
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
of its standard oxygenated derivative showed an average ion count of 4.92105 at m/z=253.4 for
the fatty acid, and an average ion count of 4.3105 at m/z=269.4 for the oxygenated derivative,
under identical instrument conditions). Thus, the ratio of the concentration of the unreacted
substrate to the concentration of product in the enzymatic reaction could be calculated from the
ratio of the peak current (ion signal) of the substrate (unreacted fatty acid) to that of the product
(oxygenated fatty acid) recorded in ESI-MS (Figure 1).
It is evident from Figure 1 that the monooxygenation has taken place (an increase by 16
Da in the molecular weight of the products relative to that of the corresponding substrates) in all
the substrates under investigation (see Table 1). Moreover, no ion signals corresponding to
addition of two or more oxygen atoms to the fatty acid were detected in the present studies,
indicating that only single monooxygenation of the fatty acids is catalyzed by CYP175A1. The
percentage yields for monooxygenation of each fatty acid are shown in the Figure 1. Figure 1
shows that the catalytic activity of CYP175A1 is maximum for 16C9 (64%) & 18C6 (60%), and
minimum for 20C11 (12%), 22C13 (12%) and 24C15 (7%). No significant product formation
14
ACS Paragon Plus Environment

Page 15 of 45
Biochemistry
1
2
3
4
5
6
7
8
9
was observed in the control experiments containing the fatty acid with H2O2 but without
CYP175A1, or containing the fatty acid with the enzyme but without H2O2 in the reaction
mixtures (see Figure S3 in the Supporting Information for a typical substrate 16C9). Thus the
oxygenation of the above fatty acids is essentially carried out by the enzymatic action of
CYP175A1. These results thus indicate that the substrate binding cavity of the enzyme might be
such that it can accommodate the monoenoic fatty acid 16C9 more favorably compared to the
larger fatty acids.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
We also studied the fatty acid oxygenation in the reductive pathway using putidaredoxin
(PdX)-putidaredoxin reductase (PdR) as the redox partners for the class-I type of CYP enzyme
system and NADH as the electron donor. PdX-PdR redox partners were earlier13 shown to drive
the reductive pathway for hydroxylation of β-carotene by CYP175A1. The reductive pathway
also resulted in the oxygenation of the monoenoic fatty acids. However, the reaction efficiency
(e.g., 21% for 16C9) was lower compared to that in the peroxide shunt pathway (64% for 16C9),
indicating poor redox coupling of PdX-PdR system with CYP175A1 (see Figure S4 in the
Supporting Information).
The effects of the experimental conditions on the enzymatic efficiency were assessed by
varying the time course, temperature and H2O2 concentration in the reaction mixture without
changing the initial concentrations of the enzyme and of the substrate. The percentage yield of
the product increased with incubation time and reached a plateau at ~30 min (Figure 2a).
Variation of the reaction temperature from 30oC to 70oC also showed distinct temperature
dependence of the percentage yield of the product formation and the maximal activity
temperature for the monoenoic fatty acid oxygenation was found to be 45oC (Figure 2b).
15
ACS Paragon Plus Environment

Biochemistry
Page 16 of 45
1
2
3
4
5
6
7
8
9
Figure 3 shows that with the increase in the concentration of H2O2 from 1 mM to 20 mM
in the reaction mixture (containing 16C9), the amount of product (after 1h reaction) increases
and reaches a maximum at 20 mM H2O2 concentration. When the peroxide concentration was
200 µM and the reaction time was 24 h, the total yield of the oxygenated product for 16C9 was
found to be 30% (data not shown). This yield is almost half of that obtained in the presence of
100 times more concentration of H2O2 (20 mM H2O2 ) in 1 h reaction time (64% yield for 16C9).
It is however important to note that presence of H2O2 causes degradation of the enzyme, which
becomes predominant at higher concentrations of the peroxide. Increase of H2O2 concentration
above 20 mM causes decrease in the amount of the product formation (Figure 3) possibly due to
the H2O2-dependent bleaching of the heme chromophore and degradation of the enzyme.38, 40
We also assessed the heme degradation of CYP175A1 in presence of the substrate
(16C9) at different concentrations of H2O2 by monitoring the Soret absorption at 417 nm with
time (see Figure S5 in the Supporting Information). The results indicated that even with 20 mM
H2O2, there were ~38% degradation of the enzyme in 1 hr, though the overall yield of the
enzymatic product at this concentration of peroxide were the highest (64%, Figure 1). Moreover,
the degradation of the enzyme by peroxide was not linear with time, and there was only <10%
degradation of the enzyme in 1 min in presence of 20 mM H2O2 (see later).
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Identification of the products
CID-MS/MS analysis
Electrospray ionization in the negative ion mode resulted in the production of carboxylate anion
of both the substrate (fatty acids) and product (oxygenated fatty acids) in the gas phase. In order
to identify the site of monooxygenation in the product, the product ion was mass selected and
16
ACS Paragon Plus Environment

Page 17 of 45
Biochemistry
1
2
3
4
5
6
7
8
9
isolated in the ion trap for collision induced dissociation (CID) fragmentation. The CID-MS/MS
spectra of two typical monooxygenated fatty acids of 18C9 and of 18T9 are shown in Figure 4
and Figure 5 respectively (see Figure S6 for 16C9, Figure S7 for 18C6 and Figure S8 for 18C11
in the Supporting Information). Both 18C9 and 18T9 are C18-monounsaturated fatty acid with
the double bond at the same position (C9-C10). They only differ in the configuration around the
double bond. 18C9 has a cis-double bond and 18T9 has a trans-double bond (see Table 1).
CID-MS/MS spectrum (see Figure 4a) of the enzymatically monooxygenated 18C9 (m/z
297, labeled as [M-H]−) shows an intense fragment ion signal at m/z 171 (base peak) along with
the fragment ion signals at m/z 155 as well as at m/z 141, m/z 157, m/z 169 and m/z 183. The
monooxygenation of the fatty acid may lead to the formation of epoxide (e.g., cis-9,10-
epoxystearic acid in the case of 18C9) and/or the allyl-hydroxy derivative with hydroxylation
either at C8 or at C11 of 18C9. In order to identify the monooxygenation product(s) of the fatty
acid, we compared the CID-MS/MS spectra of the monooxygenated fatty acid with those of the
standard epoxy derivative (Figures 4b) and of the standard hydroxy derivative (Figures 4c) of
the fatty acid.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Figure 4b shows that the CID-MS/MS spectrum of cis-9,10-epoxystearic acid (epoxy
derivative of 18C9) gives two fragment ions at m/z 171 and m/z 155 with relative ion counts
(relative abundance) 5:1. Figure 4c shows that allylhydroxy oleic acids (hydroxy derivatives of
18C9) give CID-MS/MS ion fragments at m/z 157, m/z 183, and m/z 169, m/z 141, which were
assigned to fragments of 8-hydroxyoleic acid and 11-hydroxyoleic acid respectively. Thus the
CID-MS/MS spectrum (Figure 4a) of the monooxygenated product of 18C9 consists of
fragments from the epoxy derivative (Figure 4b) as well as those from the hydroxy derivative
(Figure 4c) of the fatty acid. Analyses of the relative ion counts for each of these derivatives
17
ACS Paragon Plus Environment

Biochemistry
Page 18 of 45
1
2
3
4
5
6
7
8
9
suggested that the enzymatic monooxygenation of 18C9 gives 7:3 ratio of the epoxy (~70%) and
hydroxy (~30%) products. Moreover, the results indicate that the 8-hydroxy derivative was
higher (~24%) compared to the 11-hydroxy derivative (~6%) of the fatty acid.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Analogous to the cis-isomer (18C9), the CID-MS/MS spectrum of the enzymatic
monooxygenation product of 18T9 shows six ion fragments (Figure 5a) with the m/z values
identical to those of the cis-isomer. However, the relative ion counts of the ion fragments from
the trans-isomer (18T9) were distinctly different from those obtained from the cis-isomer
(18C9). Analogous to the cis-derivative, the CID-MS/MS spectrum of the standard trans-9,10-
epoxystearic acid (epoxy derivative of 18T9) gives two fragment ions at m/z 171 and m/z 155
(Figure 5b) with slightly different relative ion counts (relative abundance) of 10:1. Similarly,
Figure 5c shows that chemically synthesized allylhydroxyelaidic acids (hydroxy derivatives of
18T9) give the same CID-MS/MS ion fragments as those observed in case of the cis-isomer
(Figure 4c) but the relative ion counts of the ion fragments are different than those observed for
the allylhydroxyoleic acid (Figure 4c). Analyses of the relative ion counts for each of the
fragment ions obtained from the enzymatic monooxygenation of 18T9 (Figure 5a) with the help
of the standard epoxy- and allylhydroxy- derivatives suggested that the enzymatic
monooxygenation of 18T9 gives 1:4 ratio of the epoxy (20%) and hydroxy (80%) products. The
8-hydroxy derivative was lower (~35%) compared to the 11-hydroxy derivative (~45%) of the
fatty acid. The formation of the different fragment ions by collision induced dissociation of the
cis- and trans-isomers of the monooxygenated fatty acids are shown in Scheme S1 in the
Supporting Information.
These results thus suggest that monooxygenation of 18C9 catalysed by CYP175A1 yields
cis-9,10-epoxystearic acid as major product (70%) and 8-hydroxyoleic acid (24%) and 11-
18
ACS Paragon Plus Environment

Page 19 of 45
Biochemistry
1
2
3
4
5
6
7
8
9
hydroxyoleic acid (6%) as minor products. Analogously, CYP175A1 catalysis gives 8-
hydroxyelaidic acid (35%) and 11-hydroxyelaidic acid (45%) as major products and trans-9,10-
epoxystearic acid (20%) as minor product on enzymatic monooxygenation of 18T9 indicating
high regioselectivity of the enzymatic reaction of CYP175A1.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
In a similar manner, by comparing the CID-MS/MS spectra of the CYP175A1 catalyzed
oxygenation products of the other fatty acids (16C9, 18C6, and 18C11) with the standard
samples, we observed that 16C9, 18C6, and 18C11 (cis-configuration of the double bond) yield
epoxy fatty acid as major product and allylhydroxy fatty acid as minor product upon enzymatic
action of CYP175A1 (Figures S6, S7, S8 in supporting information).
The enzymatic reaction products obtained by the fatty acid oxygenation in the reductive
pathway using putidaredoxin (PdX)-putidaredoxin reductase (PdR) as the redox partners were
also analyzed by ESI-MS and CID-MS/MS to identify the products. The results showed that the
products of the enzymatic monooxygenation by the reductive pathway were the same as those
obtained by the oxidative (driven by H2O2) shunt pathway (data not shown).
GC-MS analysis
The products of the CYP175A1 catalyzed monooxygenation of the fatty acids were further
characterized by GC-MS. Fatty acids require esterification and/or other derivatization to make
them sufficiently volatile for detection by GC technique.46 The carboxylic acid group of the fatty
acids and their derivatives were esterified (methyl ester) to detect them in the GC. The unreacted
fatty acid as well as the corresponding epoxy-product could be detected by esterification (Figure
S9a & b in supporting information). The total-ion current (TIC) gas chromatogram of the
enzymatic oxygenation (Figure S9a in supporting information for a typical fatty acid 16C9)
19
ACS Paragon Plus Environment

Biochemistry
Page 20 of 45
1
2
3
4
5
6
7
8
9
shows distinct GC peaks for unreacted fatty acid ester (e.g., at retention time 9.9 min. for 16C9)
and for the epoxy derivative of the fatty acid ester (e.g., at retention time 11.92 min. for methyl
9,10-epoxypalmitoleate). The TIC of the peaks in the gas chromatogram of the methyl esters of a
known mixture of the fatty acid and the corresponding standard epoxy derivative (Figure S9b in
supporting information) match with that of the chromatogram of the methyl esters of the
enzymatic reaction solution, supporting the formation of the epoxy product by the enzyme. It is
important to note that unlike in case of the soft ionisation method of ESI-MS, the total ion count
(TIC) in the GCMS analyses cannot be used to get the quantitative estimates of the relative
concentrations of various products of the fatty acid in the present case. Moreover, simple
esterification of the sample could not detect the allyl-hydroxy products of the fatty acids.
The formation of the allyl-hydroxy products could be detected by silylation of the
hydroxyl group (Figure S9c & d in supporting information). When the esterified
monooxygenated product (produced enzymatically from 16C9) was further derivatized by
silylation reaction, two additional peaks at retention time 11.47 min. and 11.59 min. were
appeared along with the previous peak of epoxy-16C9 at the retention time 11.92 min. (see
Figure S9c in supporting information). The esterification followed by silylation of the chemically
synthesized standard allyl-hydroxy derivatives of 16C9 also gave two peaks in GC at retention
time 11.47 min. and 11.59 min. (see Figure S9d in supporting information). This result indeed
confirmed the formation of two allyl-hydroxy products of 16C9 along with epoxy-16C9 in the
CYP1751 catalyzed oxygenation of 16C9. The enzymatic products were further confirmed by
comparing the GC-mass spectral fragmentation data (see Figure S10 in supporting information)
of each species with a specific retention time in the TIC chromatogram with that of the standard
samples having the same retention time (Figure S11 in the Supporting Information). An intense
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
20
ACS Paragon Plus Environment

Page 21 of 45
Biochemistry
1
2
3
4
5
6
7
8
9
ion signal at m/z 213 (retention time 11.47 min.) corresponding to [CH3-(CH2)5-CH=CH-CH-O-
SiMe3]+ (see Figure S10a and Figure S11a in supporting information) was observed in the
enzymatic reaction solution as well as in the standard allyl-hydroxy derivative of 16C9
suggesting the presence of the hydroxyl group at C8 position in the monooxygenated 16C9.47
Similarly an intense ion signal at m/z 285 (retention time 11.59 min.) corresponding to [Me3Si-
O-CH-CH=CH-(CH2)7-COOMe]+ (see Figure S10b and Figure S11b in supporting information)
was observed both in the enzymatic product and in standard allyl-hydroxy derivative of 16C9
suggesting the hydroxyl group at C11 position in the monooxygenated 16C9.47 Analogously, the
species with retention time 11.92 min was confirmed as the epoxy derivative of 16C9 (see Figure
S10c and Figure S11c in supporting information).
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Molecular docking of the substrates
Molecular docking of the monoenoic fatty acids on the CYP175A1 (PDB: 1N97) was carried out
to assess the structural basis of the interaction of the fatty acids with the enzyme. The docking
results showed that fatty acids adopt ‘U’-shaped conformation inside the active site of the protein
(see Figures 6 & 7 for two typical fatty acids 18C9 and 18T9 respectively). This ‘U’ shaped
conformation of the substrate possibly arises due the restricted rotation of the fatty acid around
the double bond inside the active site. The docking results with several fatty acids (16C9, 18C9,
18T9, 18C6, and 18C11) showed that there are two possible types of interactions between the
substrate and the enzyme in the present case, viz., 1) the hydrophobic interaction of the alkyl
chain with the hydrophobic amino acid residues (e.g. Leu, Ile, Val, and Ala) at the substrate
binding site (see Figure 6 and 7 for two typical fatty acids 18C9 and 18T9), and 2) the ionic
interaction (hydrogen bonding) of the carboxylate group of the fatty acid with the amide
21
ACS Paragon Plus Environment

Biochemistry
Page 22 of 45
1
2
3
4
5
6
hydrogens of nearby amino acids (e.g. Glu 224 & Thr 225 in Figure 6 and Val 371 & Thr 372 in
Figure 7) and with water inside the active site.
7
8
9
Since no binding of the saturated fatty acids (e.g. palmitic acid, stearic acid etc.) to the
enzyme was observed, the double bond in the monoenoic fatty acid possibly helps in taking the
'U' shape and thus facilitates binding of the substrate at the enzyme pocket. The docking results
also indicated that depending on the chain length, double bond position and stereochemistry of
the substrate, different monoenoic fatty acids adopted different spatial orientation in the enzyme
pocket (data not shown). Thus the polar residue(s) of the enzyme, that are involved in the
interaction with the carboxylic acid group of the substrate, may be different for different fatty
acids of varied length and geometry. The docking result of 16C9 also exhibited a possibility of
polar interaction involving Thr168 and the carboxylate group of 16C9 (see Figure S12 in the
Supporting Information) in the active site of the enzyme pocket, while the same residue is not
involved in interaction with other fatty acids. Furthermore, the docking results indicated that the
double bond of the cis-monoenoic fatty acid (18C9, see Figure 6) and allylic-methylene group of
the trans-monoenoic fatty acid (18T9, see Figure 7) reside nearer to the heme iron (at ~4 Å for
18C9, and at ~3.4 Å for 18T9) making them as the potential site for oxygenation, which agrees
with the experimental results (Figures 4 and 5). The binding of the fatty acid to the enzyme is
however weak (large Kd values, Table 1) and thus the fluctuations in the conformation of the
fatty acid inside the enzyme cavity could account for observation of both the epoxy- as well as
allyl-hydroxy products on enzymatic monooxygenation by CYP175A1.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
It is however important to note that the present in-silico analyses of the structures of the
enzyme and the fatty acids only provide the possible conformations of the substrates at the active
22
ACS Paragon Plus Environment

Page 23 of 45
Biochemistry
1
2
3
4
5
6
site of the enzyme, and the exact disposition of the fatty acid inside the enzyme cannot be
predicted without the crystal structure of the enzyme-substrate complex.
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Activity assay of the enzyme
The enzymatic activity of CYP175A1 for monooxygenation of the monoenoic fatty acids by
H2O2 following the oxidative shunt pathway of the enzymatic cycle was determined at 45oC by
Lineweaver-Burk method from the linear fit to the plot of 1/rate (V) vs. 1/[substrate] (see Figure
S13 in supporting information). The values of Km and kcat for enzymatic reaction of various
monoenoic fatty acids with H2O2 catalysed by CYP175A1 are given in Table 2. The results
indicate that CYP175A1 show highest enzymatic efficiency (kcat/Km) for fatty acids 16C9 and
18C9, while that for the trans- isomer 18T9 is the lowest. The enzymatic activity (kcat) was
highest for 16C9 with the turn over number ~16 min-1. Earlier studies on enzymatic reaction of
-carotene by CYP175A1 following the reductive pathway (i.e., with NADH, PdX, PdR and
oxygen) reported the turnover rate of 0.23 min-1, which is much lower than that observed for
reaction of the monoenoic fatty acid by the enzyme following oxidative shunt pathway (i.e., with
H2O2 ) in the present study. The reductive pathway of the enzymatic reaction using PdX and
PdR, was also found to be less efficient in the present case possibly because of weaker redox
coupling between PdX and CYP175A1. However, it is important to note that unlike in the
reductive pathway, the peroxide induced oxidative shunt pathway of the enzymatic reaction was
associated with partial degradation of the enzyme (see Figure S5 in supporting information).
Nevertheless, there was <10 % degradation of the enzyme by 20 mM H2O2 in the initial 1 min of
the reaction (see Figure S5 in supporting information), hence the initial rates (Table 2)
determined at this time period, may be assumed to have almost no effect of degradation of the
23
ACS Paragon Plus Environment

Biochemistry
Page 24 of 45
1
2
3
4
5
6
7
8
9
enzyme. The apparent rates of the reaction over 1 hr (e.g., from the percentage yield data shown
in Figure 1) would however be much less than that estimated in short time period (Table 2),
because of partial degradation of the enzyme (see Figure S5 in supporting information) as well as
because of saturation of the reaction (Figure 2) at longer time scales.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Conclusions
The thermostable enzyme, CYP175A1 catalyzes monooxygenation reaction of various
monoenoic fatty acids by H2O2 in the oxidative shunt pathway as well as by oxygen and NADH
in the reductive pathway (in presence of PdX and PdR). This monooxygenation reaction exhibits
regioselectivity and the product distribution depends on the configuration of the ethylenic double
bond in the substrate (monoenoic fatty acid). Mass spectrometric studies of the enzyme
(CYP175A1) catalyzed oxygenation reaction showed that the epoxy fatty acid (double bond
oxygenation) is the major product for the cis-monoenoic fatty acid substrate, while the allyl-
hydroxy fatty acids are the major products for the trans-monoenoic fatty acid substrate. This
regioselectivity is consistent with the substrate docking analyses. The specificity of the enzyme
also varies with the chain length and position of the double bond (C=C) in the monoenoic fatty
acids. Among all the monoenoic fatty acid substrates, palmitoleic acid (16C9) was found to be
the better substrate than others. The results further support that CYP175A1 may have an
important role in lipid metabolism in Thermus thermophilus HB27, as it can efficiently
metabolise various fatty acids. The present work further suggests that the natural substrate of this
orphan enzyme might have a 'U' shape structure for efficient binding to the active site of the
enzyme.
24
ACS Paragon Plus Environment

Page 25 of 45
Biochemistry
1
2
3
4
Supporting Information
5
6
7
8
9
ESI-MS of CYP175A1, binding titration of various fatty acids with CYP175A1, control reaction
of 16C9 with CYP175A1, enzymatic reaction of 16C9 with CYP175A1 using reductive
pathway, H2O2 dependent heme degradation, CID-MS/MS of enzymatic and standard products
of 16C9, 18C6 and 18C11, TIC gas chromatogram of enzymatic and standard products of 16C9,
GC-MS of enzymatic and standard products of 16C9, docking result of palmitoleic acid (16C9),
Lineweaver-Burk plot for various fatty acid kinetics, and possible mechanism of fragmentation
of epoxy- and different allyl-hydroxy fatty acids during CID/MS/MS study. This material is
available free of charge via the Internet at http://pubs.acs.org.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Acknowledgements
The authors thank Mr. Bharat T. Kansara and Mr. Abhishek Acharya for their help.
Note:
The authors declare no competing financial interest.
25
ACS Paragon Plus Environment

Biochemistry
Page 26 of 45
1
2
3
4
References
5
6
7
8
9
(1)
(2)
Guengerich, F. P. (1993) Cytochrome P450 enzymes, Am. Sci. 81, 440-447.
Ortiz de Montellano, P. R. (2004) Cytochrome P450: structure, mechanism, and
biochemistry, 3rd Edition ed., Springer, New York.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
(3)
(4)
Urlacher, V. B., and Girhard, M. (2011) Cytochrome P450 monooxygenases: an update
on perspectives for synthetic application, Trends Biotechnol. 30, 26-36.
Wright, R. L., Harris, K., Solow, B., White, R. H., and Kennelly, P. J. (1996) Cloning of
a potential cytochrome P450 from the archaeon Sulfolobus solfataricus, FEBS Lett. 384,
235-239.
(5)
(6)
Yano, J. K., Blasco, F., Li, H., Schmid, R. D., Henne, A., and Poulos, T. L. (2003)
Preliminary characterization and crystal structure of a thermostable cytochrome P450
from Thermus thermophilus, J. Biol. Chem. 278, 608-616.
Yano, J. K., Koo, L. S., Schuller, D. J., Li, H., Ortiz de Montellano, P. R., and Poulos, T.
L. (2000) Crystal structure of a thermophilic cytochrome P450 from the archaeon
Sulfolobus solfataricus, J. Biol. Chem. 275, 31086-31092.
(7)
(8)
Behera, R. K., and Mazumdar, S. (2010) Thermodynamic basis of the thermostability of
CYP175A1 from Thermus thermophilus, Int. J. Biol. Macromol. 46, 412-418.
McLean, M. A., Maves, S. A., Weiss, K. E., Krepich, S., and Sligar, S. G. (1998)
Characterization of a cytochrome P450 from the acidothermophilic archaea Sulfolobus
solfataricus, Biochem. Biophys. Res. Commun. 252, 166-172.
(9)
Nishida, C. R., and Ortiz de Montellano, P. R. (2005) Thermophilic cytochrome P450
enzymes, Biochem. Biophys. Res. Commun. 338, 437-445.
26
ACS Paragon Plus Environment

Page 27 of 45
Biochemistry
1
2
3
4
5
6
7
8
9
(10) Guengerich, F. P., Tang, Z., Salamanca-Pinzón, S. G., and Cheng, Q. (2010)
Characterizing Proteins of Unknown Function: Orphan Cytochrome P450 Enzymes as a
Paradigm, Mol. Interv. 10, 153-163.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
(11) Koo, L. S., Immoos, C. E., Cohen, M. S., Farmer, P. J., and Ortiz de Montellano, P. R.
(2002) Enhanced electron transfer and lauric acid hydroxylation by site-directed
mutagenesis of CYP119, J. Am. Chem. Soc. 124, 5684-5691.
(12) Mandai, T., Fujiwara, S., and Imaoka, S. (2009) Construction and engineering of a
thermostable self-sufficient cytochrome P450, Biochem. Biophys. Res. Commun. 384, 61-
65.
(13) Momoi, K., Hofmann, U., Schmid, R. D., and Urlacher, V. B. (2006) Reconstitution of -
carotene hydroxylase activity of thermostable CYP175A1 monooxygenase, Biochem.
Biophys. Res. Commun. 339, 331-336.
(14) Boddupalli, S. S., Estabrook, R. W., and Peterson, J. A. (1990) Fatty acid
monooxygenation by cytochrome P450BM-3, J. Biol. Chem. 265, 4233-4239.
(15) Dominguez, A., Pastrana, L., Longo, M. A., Rúa, M. L., and Sanroman, M. A. (2005)
Lipolytic enzyme production by Thermus thermophilus HB27 in a stirred tank bioreactor,
Biochem. Eng. J. 26, 95-99.
(16) Domínguez, A., Deive, F., Pastrana, L., Rúa, M., Longo, M., and Sanroman, M. (2010)
Thermostable lipolytic enzymes production in batch and continuous cultures of Thermus
thermophilus HB27, Bioproc. Biosyst. Eng. 33, 347-354.
(17) Fucinos, P., Rua, M. L., Longo, M. A., Sanroman, M. A., and Pastrana, L. (2008)
Thermal spring water enhances lipolytic activity in Thermus thermophilus HB27, Process
Biochemistry 43, 1383-1390.
27
ACS Paragon Plus Environment

Biochemistry
Page 28 of 45
1
2
3
4
5
6
7
8
9
(18) Nelson, D. P., and Kiesow, L. A. (1972) Enthalpy of decomposition of hydrogen
peroxide by catalase at 25°C (with molar extinction coefficients of H2O2 solutions in the
UV), Anal. Biochem. 49, 474-478.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
(19) Davies, D. M., Jones, P., and Mantle, D. (1976) The kinetics of formation of horseradish
peroxidase compound I by reaction with peroxobenzoic acids. pH and peroxo acid
substituent effects, Biochem. J. 157, 247-253.
(20) Goyal, S., Deshpande, M. S., and Mazumdar, S. (2011) Structural design of the active
site for covalent attachment of the heme to the protein matrix: studies on a thermostable
cytochrome P450, Biochemistry 50, 1042-1052.
(21) Behera, R. K., Goyal, S., and Mazumdar, S. (2010) Modification of the heme active site
to increase the peroxidase activity of thermophilic cytochrome P450: A rational
approach, J. Inorg. Biochem. 104, 1185-1194.
(22) Sevrioukova, I. F., Garcia, C., Li, H., Bhaskar, B., and Poulos, T. L. (2003) Crystal
Structure of Putidaredoxin, the [2Fe-2S] Component of the P450cam Monooxygenase
System from Pseudomonas putida, J. Mol. Biol. 333, 377-392.
(23) Kuznetsov, V. Y., Blair, E., Farmer, P. J., Poulos, T. L., Pifferitti, A., and Sevrioukova, I.
F. (2005) The Putidaredoxin Reductase-Putidaredoxin Electron Transfer Complex, J.
Biol. Chem. 280, 16135-16142.
(24) Sevrioukova, I. F., and Poulos, T. L. (2002) Putidaredoxin Reductase, a New Function
for an Old Protein, J. Biol. Chem. 277, 25831-25839.
(25) Klaas, M. R. g., and Warwel, S. (1999) New oxidation methods for unsaturated fatty
acids, esters, and triglycerides, In Recent developments in the synthesis of fatty acid
derivatives, pp 157-181, AOCS Publishing, Urbana, USA.
28
ACS Paragon Plus Environment

Page 29 of 45
Biochemistry
1
2
3
4
5
6
7
8
9
(26) Knothe, G., Weisleder, D., Bagby, M., and Peterson, R. (1993) Hydroxy fatty acids
through hydroxylation of oleic acid with selenium dioxide/tert-butylhydroperoxide, J.
Am. Oil Chem. Soc. 70, 401-404.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
(27) Hashimoto, N., Aoyama, T., and Shioiri, T. (1981) New Methods and Reagents in
Organic Synthesis. 14. A Simple Efficient Preparation of Methyl Esters with
Trimethylsilyldiazomethane (TMSCHN2) and Its Application to Gas Chromatographic
Analysis of Fatty Acids, Chem. Pharm. Bull. 29, 1475-1478.
(28) Poole, C. F., and Schuette, S. A. (1984) Contemporary practice of chromatography,
Elsevier Science, Amsterdam, New York.
(29) Blau, K., and Halket, J. M. (1978) Handbook of Derivatives for Chromatography,
Heyden & Sons, London.
(30) Griffin, B. W., and Peterson, J. A. (1972) Camphor binding of Pseudomonas putida
cytochrome P-450. Kinetics and thermodynamics of the reaction, Biochemistry 11, 4740-
4746.
(31) Banerjee, S., Prakash, H., and Mazumdar, S. (2011) Evidence of molecular fragmentation
inside the charged droplets produced by electrospray process, J. Am. Soc. Mass Spectrom.
22, 1707-1717.
(32) Banerjee, S., and Mazumdar, S. (2010) Non-covalent dimers of the lysine containing
protonated peptide ions in gaseous state: electrospray ionization mass spectrometric
study, J. Mass Spectrom. 45, 1212-1219.
(33) Banerjee, S., and Mazumdar, S. (2012) Electrospray ionization mass spectrometry: A
technique to access the information beyond the molecular weight of the analyte, Int. J.
Anal. Chem. 2012, DOI:10.1155/2012/282574.
29
ACS Paragon Plus Environment