Lauric acid C12:0 was converted into a mixture of hydroxy-lauric acids when incubated with microsomes from yeast expressing CYP77A4.. Despite the fact that the implication of a cyto-chro
Trang 1P450 able to catalyze the epoxidation of free fatty
acids in plants
Vincent Sauveplane1, Sylvie Kandel2, Pierre-Edouard Kastner1, Ju¨rgen Ehlting1,
Vincent Compagnon1, Danie`le Werck-Reichhart1and Franck Pinot1
1 Institut de Biologie Mole´culaire des Plantes, University of Strasbourg, France
2 Department of Pharmaceutical Chemistry, University of California, San Francisco, CA, USA
Fatty acid-oxidizing enzymes have been the subject of
an increasing number of studies in all organisms, as
the products of their reactions exhibit fundamental
biological activities [1–3] Among these oxidases,
cyto-chromes P450 play a prominent role For example, in
animals, arachidonic acid (C20:4) is oxidized through
the cytochrome P450 pathway, leading to the
produc-tion of hydroxylated and epoxidized derivatives [4–6] The cytochrome P450 superfamily represents a highly diversified set of heme-containing proteins found in bacteria, fungi, animals and plants [7] In animals, members of the CYP4A gene subfamily mainly cata-lyze the formation of x- and x-1-hydroxyl derivatives
of fatty acids The regulation of some CYP4A enzymes
Keywords
cytochrome P450; defense; epoxide; fatty
acid; plant
Correspondence
F Pinot, IBMP-CNRS UPR 2357, Institut de
Botanique, 28 rue Goethe, F-67083
Strasbourg Cedex, France
Fax: +33 3 90 24 19 21
Tel: +33 3 90 24 18 37
E-mail: franck.pinot@ibmp-ulp.u-strasbg.fr
(Received 4 September 2008, revised 20
November 2008, accepted 26 November
2008)
doi:10.1111/j.1742-4658.2008.06819.x
An approach based on an in silico analysis predicted that CYP77A4, a cytochrome P450 that so far has no identified function, might be a fatty acid-metabolizing enzyme CYP77A4 was heterologously expressed in a Saccharomyces cerevisiae strain (WAT11) engineered for cytochrome P450 expression Lauric acid (C12:0) was converted into a mixture of hydroxy-lauric acids when incubated with microsomes from yeast expressing CYP77A4 A variety of physiological C18fatty acids were tested as poten-tial substrates Oleic acid (cis-D9C18:1) was converted into a mixture of
x-4-to x-7-hydroxyoleic acids (75%) and 9,10-epoxystearic acid (25%) Linoleic acid (cis,cis-D9,D12C18:2) was exclusively converted into 12,13-epoxyocta-deca-9-enoic acid, which was then converted into diepoxide after epoxida-tion of the D9 unsaturation Chiral analysis showed that 9,10-epoxystearic acid was a mixture of 9S⁄ 10R and 9R ⁄ 10S in the ratio 33 : 77, whereas 12,13-epoxyoctadeca-9-enoic acid presented a strong enantiomeric excess in favor of 12S⁄ 13R, which represented 90% of the epoxide Neither stearic acid (C18:0) nor linolelaidic acid (trans,trans-D9,D12C18:2) was metabolized, showing that CYP77A4 requires a double bond, in the cis configuration, to metabolize C18fatty acids CYP77A4 was also able to catalyze the in vitro formation of the three mono-epoxides of a-linolenic acid (cis,cis,cis-D9,
D12,D15C18:3), previously described as antifungal compounds Epoxides gen-erated by CYP77A4 are further metabolized to the corresponding diols by epoxide hydrolases located in microsomal and cytosolic subcellular frac-tions from Arabidopsis thaliana The concerted action of CYP77A4 with epoxide hydrolases and hydroxylases allows the production of compounds involved in plant–pathogen interactions, suggesting a possible role for CYP77A4 in plant defense
Abbreviation
EET, epoxyeicosatrienoic acid.
Trang 2by peroxisome proliferator-activated receptors points
to a role in fatty acid catabolism [8] After
x-hydroxyl-ation, fatty acids can be further oxidized to diacids,
which can then be eliminated by peroxisome
b-oxida-tion [9] However, investigab-oxida-tions describing the effect
of x-hydroxy fatty acids in different physiological
pro-cesses [10–13] have suggested that x-hydroxylation
cannot be considered only as a step leading to
catabo-lism The epoxidation of polyunsaturated fatty acid
double bonds, particularly of arachidonic acid, has
generated much interest because of the biological
activ-ities of the resulting metabolites [14,15] These
epoxi-dation reactions of C20:4are catalyzed by members of
the CYP2C subfamily and by the CYP2J2 isoform
[6,16,17] Human CYP4F8 and CYP4F12 isoforms are
able to epoxidize docosahexaenoic acid (C22:6) [18]
In plants, fatty acids are also metabolized by
cyto-chrome P450-dependent oxygenases [19], and it is
possible to distinguish x-hydroxylases and in-chain
hydroxylases that attack the terminal and
subtermi-nal positions, respectively So far, the majority of
work has addressed x-hydroxylases mainly
repre-sented in CYP86 and CYP94 families [19] Their
involvement in the synthesis of cutin, a protective
biopolymer of fatty acids cross-linked by ester bonds
[20], has been established [21,22] Studies of LCR
(LACERATA) and att1 (aberrant induction of type
three genes), the first Arabidopsis thaliana mutants
with alterations in the coding sequence of CYP86A8
and CYP86A2, respectively, have also shown that
x-hydroxylases have key roles to play in plant
devel-opment [21,22]
Despite the fact that the implication of a
cyto-chrome P450 in the epoxidation of a long-chain fatty
acid was first demonstrated in spinach leaves more
than three decades ago [20,23], a cytochrome P450 able
to epoxidize fatty acids is still poorly documented in
plants Biochemical studies performed with
unsatu-rated analogues of lauric acid (C12:0) clearly
demon-strated the existence in plants of a cytochrome P450
able to epoxidize the double bonds of fatty acids The
terminal olefin 11-dodecenoic acid is converted into
11,12-epoxylauric acid by a cytochrome P450 in
Vicia sativa microsomes [24] The epoxidation of
unsaturated analogues of lauric acid by cytochrome
P450 was also reported in microsomes from Jerusalem
artichoke [25,26], as well as in microsomes from wheat
[27] However, none of the enzymes implicated in these
reactions have been characterized and, to date, no
cytochrome P450 able to epoxidize free fatty acids has
been identified in plants The epoxidation of
physiolog-ical substrates, such as oleic acid (cis-D9C18:1) and
lino-leic acid (cis,cis-D9,D12C18:2), has been reported in
Vicia faba [28] and Glycine max [29] However, these reactions were not catalyzed by cytochrome P450, but rather by peroxygenases, which are hydroperoxide-dependent fatty acid epoxidases Recently, studies of a peroxygenase purified from oat have demonstrated that this enzyme is deeply buried in microsomes or in lipid droplets [30] Lee et al [31] identified a non-heme di-iron enzyme, a ‘desaturase-like’ protein, able to transform linoleic acid into 12,13-epoxyoctadeca-cis-9-enoic acid (vernolic acid) This compound can make
up 50–90% of total fatty acids in seed oil of certain Euphorbiaceae, such as Euphorbia lagascae [32] In this plant, the enzyme involved in its production was described recently [32] This enzyme, classified as CYP726A1, does not epoxidize free fatty acids, but fatty acids bound to phosphatidylcholine [32]
A new approach, based on an in silico analysis of publicly available transcriptome data, has been devel-oped recently to map cytochrome P450 genes onto spe-cific metabolic pathways [33] This analysis identifies metabolic genes that are co-expressed with a given bait P450 during plant development, on stress and hormone treatment, and in mutant wild-type comparisons Based on the functional annotation of co-expressed genes, a metabolic pathway in which the bait P450 may act is predicted This approach suggested that CYP77A4 could be involved in fatty acid metabolism
as it is developmentally co-expressed across hundreds
of biological samples with several characterized enzymes involved in lipid metabolism The most simi-larly expressed genes are CYP86A8 encoding a fatty acid x-hydroxylase, a putative epoxide hydrolase, several genes encoding enzymes involved in the synthesis of fatty acids in plastids, including the stearoyl acyl carrier protein desaturase SSI2, and the plastidic long-chain acyl-CoA synthetase LACS9 (for a complete list of co-expressed genes, see http://www-ibmp.u-strasbg.fr/
~CYPedia/CYP77A4/CoExpr_CYP77A4_Organs.html)
In this work, we report the heterologous expression and functional characterization of CYP77A4 Substrate specificity and catalytic properties were explored using recombinant CYP77A4 expressed in an engineered yeast strain Our study confirms that this enzyme is a fatty acid-metabolizing enzyme We show that CYP77A4 is able to catalyze, in vitro, the epoxidation
of physiological unsaturated fatty acids Our work also shows that the epoxides generated can be further hydrolyzed to the corresponding diols by epoxide hydrolases present in subcellular fractions of A thali-ana Thus, CYP77A4 from A thaliana, described in this work, is the first cytochrome P450 able to catalyze free fatty acid epoxidation, identified in plants Its physiological significance remains to be established
Trang 3and will be assessed by future studies of A thaliana
mutated in the coding sequence of CYP77A4
Results
Selection, cloning and expression of CYP77A4
An approach based on an in silico analysis predicted
that CYP77A4 could be involved in fatty acid
metabo-lism [33] The coding sequence of CYP77A4 was
amplified by PCR from a cDNA library of Arabidopsis
and subsequently cloned into a yeast expression vector
The deduced protein (512 amino acids) has a
calcu-lated mass of 58 134 Da and a pI of 8.71 Enzymatic
characterization of CYP77A4 was carried out
employ-ing microsomes from the yeast strain WAT11
trans-formed with the plasmid pYeDP60 [34] containing the
coding sequence of CYP77A4 WAT11 over-expresses
a plant P450 reductase in order to optimize electron
transfer during catalysis and probably to increase the
stability of the expressed P450 Furthermore, there are
only three cytochromes P450 encoded by the yeast
gen-ome They are either not expressed or expressed at a
negligible level in the growth conditions used here, and
none is able to metabolize fatty acids, ensuring that
the metabolism described here results from enzymatic
reactions catalyzed by CYP77A4 [34] After
micro-somal membrane isolation from the
CYP77A4-trans-formed yeasts, the level of expression of the enzyme
was evaluated on the basis of the differential
absor-bance of reduced CO-bound versus reduced
micro-somes at 450 nm [35] The CYP77A4 content of the
microsomal preparation used in our experiments was
0.1 nmolÆmg)1 protein (Fig S1) No absorbance at
450 nm and no enzymatic activity with the substrates
tested were detected in microsomes from yeast
trans-formed with a void plasmid under the same growth
conditions
Metabolism of lauric acid by CYP77A4
To validate the hypothesis of CYP77A4 being a fatty
acid-metabolizing enzyme, we incubated radiolabeled
lauric acid (C12:0) with microsomes from yeast
express-ing CYP77A4 The resolution of reaction products was
performed by directly loading the incubation medium
onto a TLC plate Figure 1 shows the
radiochromato-grams obtained after incubation in the absence
(Fig 1A) or presence (Fig 1B–D) of NADPH A large
peak of radioactivity was detected after 20 min of
incu-bation (peak 1, Fig 1B) It was not formed in the
absence of NADPH (Fig 1A), with microsomes from
yeast transformed with a void plasmid (Fig 1C) or
with boiled microsomes (Fig 1D) Taken together, these results demonstrate the involvement of CYP77A4
in the formation of this radioactive peak Metabolites from this peak were purified, derivatized and subjected
to GC⁄ MS analysis (Experimental procedures) The mass spectrum of the derivatized metabolite 1 (Fig S2) showed ions at m⁄ z (relative intensity, %) values of 73 (41%) [(CH3)3Si+], 75 (23%) [(CH3)2Si+=O], 117 (100%), 255 (15%) (M-47) [loss of methanol from the (M-15) fragment], 271 (3%) (M-31) (loss of OCH3from the methyl ester), 287 (6%) (M-15) (loss of a methyl from the trimethylsilyl group) This fragmentation pattern is characteristic of the derivative of 11-hydroxy-lauric acid (x-1) (M = 302 gÆmol)1) The mass spec-trum of derivatized metabolite 2 (Fig S2) showed ions
at m⁄ z (relative intensity, %) values of 73 (70%) [(CH3)3Si+], 75 (30%) [(CH3)2Si+=O], 131 (100%),
255 (12%) (M-47) [loss of methanol from the (M-15) fragment], 271 (4%) (M-31) (loss of OCH3 from the methyl ester), 273 (51%), 287 (2%) (M-15) (loss of a methyl from the trimethylsilyl group) This fragmen-tation pattern is characteristic of the derivative of 10-hydroxylauric acid (x-2) (M = 302 gÆmol)1) The mass spectrum of derivatized metabolite 3 (Fig S2) showed ions at m⁄ z (relative intensity, %) values of 73 (75%) [(CH3)3Si+], 75 (31%) [(CH3)2Si+=O], 145 (100%), 255 (11%) (M-47) [loss of methanol from the (M-15) fragment], 259 (59%), 271 (3%) (M-31) (loss of OCH3from the methyl ester), 287 (2%) (M-15) (loss of
a methyl from the trimethylsilyl group) This fragmen-tation pattern is characteristic of the derivative of 9-hydroxylauric acid (x-3) (M = 302 gÆmol)1) The mass spectrum of derivatized metabolite 4 (Fig S2) showed ions at m⁄ z (relative intensity, %) values of 73 (68%) [(CH3)3Si+], 75 (28%) [(CH3)2Si+=O], 159 (100%), 245 (68%), 255 (9%) (M-47) [loss of methanol from the (M-15) fragment], 271 (4%) (M-31) (loss of OCH3from the methyl ester), 287 (2%) (M-15) (loss of
a methyl from the trimethylsilyl group) This fragmen-tation pattern is characteristic of the derivative of 8-hy-droxylauric acid (x-4) (M = 302 gÆmol)1) The mass spectrum of derivatized metabolite 5 (Fig S2) showed ions at m⁄ z (relative intensity, %) values of 73 (97%) [(CH3)3Si+], 75 (39%) [(CH3)2Si+=O], 173 (100%),
231 (71%) 255 (11%) (M-47) [loss of methanol from the (M-15) fragment], 271 (4%) (M-31) (loss of OCH3 from the methyl ester), 287 (5%) (M-15) (loss of a methyl from the trimethylsilyl group) This fragment-ation pattern is characteristic of the derivative of 7-hydroxylauric acid (x-5) (M = 302 gÆmol)1) Their identification revealed that the reaction product is com-posed of a mixture of five different in-chain hydroxyl-ation products of lauric acid, which is predominantly
Trang 4hydroxylated on the x-1 position When oxidizing lauric
acid, CYP77A4 exhibits the following regioselectivity:
x-1 (53%), x-2 (15%), x-3 (8%), x-4 (18%) and x-5
(6%) For substrate oxidation, we determined,
by kinetic studies, Km,app and Vmax,app values of
172 ± 13 lm and 117 ± 5 nmolÆmin)1Ænmol)1 P450,
respectively (Fig S3) The x-1 position of lauric acid
corresponds to carbon 11 of physiological C18 fatty
acids, closely located near the double bonds of oleic,
linoleic and a-linolenic acids We therefore tested these
different unsaturated fatty acids as potential substrates
Metabolism of oleic acid by CYP77A4
We first incubated mono-unsaturated oleic acid
(C18:1) The radiochromatograms obtained after
reso-lution of the reaction products on TLC are presented
in Fig 2 Incubation was carried out in the absence (Fig 2A) or presence (Fig 2B–D) of NADPH Two new peaks of radioactivity (peak 1 and peak 2, Fig 2B) were detected; their formation required the presence of NADPH in the incubation They were also not formed on incubation with microsomes from yeast transformed with a void plasmid (Fig 2C) or with boiled microsomes (Fig 2D) Metabolites from peak 1 were purified, derivatized and subjected to
GC⁄ MS analysis The mass spectrum of derivatized metabolite 1 (Fig S4) showed ions at m⁄ z (relative intensity, %) values of 73 (100%) [(CH3)3Si+], 75 (58%) [(CH3)2Si+=O], 337 (9%) (M-47) [loss of methanol from the (M-15) fragment], 353 (2%) (M-31) (loss of OCH3 from the methyl ester), 369
Fig 1 Radiochromatographic resolution by TLC of metabolites generated in incubations of lauric acid with microsomes from yeast express-ing CYP77A4 Microsomes were incubated with 100 l M [1- 14 C]lauric acid in the absence (A) or presence (B) of NADPH Incubations were performed at 27 C and contained 20 pmol of CYP77A4 They were stopped after 30 min by the addition of 20 lL of acetonitrile (containing 0.2% acetic acid) and directly spotted onto TLC Peak S, lauric acid; peak 1, mixture of 11-, 10-, 9-, 8- and 7-hydroxylauric acids Experiments
in (C) and (D) were performed as in (B), but with microsomes from yeast transformed with a void plasmid (C) or with boiled microsomes (D) The structures of the metabolites are described in (E).
Trang 5(3%) (M-15) (loss of a methyl from the trimethylsilyl
group) and 384 (2%) (M) The mass spectrum also
showed ions at 159 (63%) and 327 (7%), resulting
from cleavage on both sides of the hydroxyl function
carrying the trimethylsilyl group This fragmentation
pattern is characteristic of the derivative of
14-hy-droxyoleic acid (x-4) (M = 384 gÆmol)1) The mass
spectrum of derivatized metabolite 2 (Fig S4) showed
ions at m⁄ z (relative intensity, %) values of 73
(100%) [(CH3)3Si+], 75 (50%) [(CH3)2Si+=O], 337
(11%) (M-47) [loss of methanol from the (M-15)
fragment], 369 (3%) (M-15) (loss of a methyl from
the trimethylsilyl group) The mass spectrum also
showed ions at 173 (34%) and 313 (18%), resulting
from cleavage on both sides of the hydroxyl function
carrying the trimethylsilyl group This fragmentation
pattern is characteristic of the derivative of
13-hy-droxyoleic acid (x-5) (M = 384 gÆmol)1) The mass
spectrum of derivatized metabolite 3 (Fig S4) showed
ions at m⁄ z (relative intensity, %) values of 73 (48%) [(CH3)3Si+], 75 (12%) [(CH3)2Si+=O], 337 (3%) (M-47) [loss of methanol from the (M-15) fragment], 353 (1%) (M-31) (loss of OCH3 from the methyl ester),
369 (0.5%) (M-15) (loss of a methyl from the trim-ethylsilyl group) The mass spectrum also showed ions
at 187 (100%) and 299 (4%), resulting from cleavage
on both sides of the hydroxyl function carrying the trimethylsilyl group This fragmentation pattern is characteristic of the derivative of 12-hydroxyoleic acid (x-6) (M = 384 gÆmol)1) The mass spectrum of derivatized metabolite 4 (Fig S4) showed ions at m⁄ z (relative intensity, %) values of 73 (35%) [(CH3)3Si+],
75 (14%) [(CH3)2Si+=O], 337 (3%) (M-47) [loss
of methanol from the (M-15) fragment], 353 (1%) (M-31) (loss of OCH3 from the methyl ester), 369 (1%) (M-15) (loss of a methyl from the trimethylsilyl group) and 384 (0.5%) (M) The mass spectrum also showed ions at 201 (1%) and 285 (100%), resulting
Fig 2 Radiochromatographic resolution by TLC of metabolites generated in incubations of oleic acid with microsomes from yeast express-ing CYP77A4 Microsomes were incubated with 100 l M [1- 14 C]oleic acid in the absence (A) or presence (B) of NADPH Incubations were performed at 27 C and contained 20 pmol of CYP77A4 They were stopped after 30 min by the addition of 20 lL of acetonitrile (containing 0.2% acetic acid) and directly spotted onto TLC Peak S, oleic acid; peak 1, mixture of 14-, 13-, 12- and 11-hydroxyoleic acids; peak 2, 9, 10-epoxystearic acid Experiments in (C) and (D) were performed as in (B), but with microsomes from yeast transformed with a void plasmid (C) or with boiled microsomes (D) The structures of the metabolites are described in (E).
Trang 6from cleavage on both sides of the hydroxyl function
carrying the trimethylsilyl group This fragmentation
pattern is characteristic of the derivative of
11-hy-droxyoleic acid (x-7) (M = 384 gÆmol)1) The
identifi-cation of metabolites from peak 1 by GC⁄ MS after
purification and derivatization revealed that
CYP77A4 hydroxylates oleic acid with the following
regioselectivity: x-7 (58%), x-6 and x-5 (30%), x-4
(12%) The metabolite from peak 2 displayed the
TLC mobility expected for 9,10-epoxystearic acid,
and was indeed identified as 9,10-epoxystearic acid by
GC⁄ MS analysis (Fig S4) For substrate oxidation,
we determined, by kinetic studies, Km,appand Vmax,app
values of 84 ± 23 lm and 26 ± 5 nmolÆmin)1Ænmol)1
P450, respectively (Fig S3) We determined the
ste-reochemistry of this epoxide after purification and
analysis by HPLC using a chiral column The
radio-chromatogram of Fig 3 shows that it is a mixture of
the two enantiomers, 9S⁄ 10R and 9R ⁄ 10S, in the
ratio 33 : 77, respectively
Metabolism of linoleic acid by CYP77A4
Figure 4 shows the radioactivity profiles obtained after
incubation of linoleic acid (C18:2) with microsomes
from yeast expressing CYP77A4 The addition of
NADPH to the incubation medium led to the
forma-Fig 3 Radiochromatographic resolution by HPLC of the
enantio-mers of 9,10-epoxystearic acid produced by CYP77A4 (A) After
incubation of oleic acid with microsomes from yeast expressing
CYP77A4, the 9,10-epoxystearic produced (peak 2, Fig 2B) was
purified and subjected to chiral HPLC analysis with hexane ⁄
propan-2-ol ⁄ acetic acid (99.7 : 0.2 : 0.1, v ⁄ v ⁄ v) at a flow rate of 0.8
mLÆmin)1 (B) Structures of the enantiomers.
Fig 4 Radiochromatographic resolution by TLC of metabolites gen-erated in incubations of linoleic acid with microsomes from yeast expressing CYP77A4 Microsomes were incubated with 100 l M
[1-14C]linoleic acid in the absence (A) or presence (B) of NADPH Incubations were performed at 27 C and contained 20 pmol of CYP77A4 They were stopped after 30 min by the addition of 20 lL
of acetonitrile (containing 0.2% acetic acid) and directly spotted onto TLC Peak S, linoleic acid; peak 1, 9,10:12,13-diepoxyoctadecanoic acid; peak 2, 12,13-epoxyoctadeca-9-enoic acid Experiments in (C) and (D) were performed as in (B), but with microsomes from yeast transformed with a void plasmid (C) or with boiled microsomes (D) The structures of the metabolites are described in (E).
Trang 7tion of a major radioactive peak (peak 2, Fig 4B)
which was not present in the absence of NADPH
(Fig 4A) It results from a reaction catalyzed by
CYP77A4, as it was not formed when the microsomes
were from yeast transformed with a void plasmid
(Fig 4C) or were boiled (Fig 4D) This peak contains
only one metabolite, which was identified by GC⁄ MS
analysis (Fig S5) after purification reaction in acidic
methanol and derivatization as
12,13-epoxyoctadeca-9-enoic acid (vernolic acid), resulting from the
epoxi-dation of the D12double bond The kinetic parameters
from the reaction of substrate oxidation are Km,app=
61±3 lm and Vmax,app= 13 ± 0.3 nmolÆmin)1Ænmol)1
P450 (Fig S3) Stereochemistry studies presented in
Fig 5 show that CYP77A4 possesses a strong
enantio-specificity: the epoxide formed is a mixture of 12S⁄ 13R
and 12R⁄ 13S in the ratio 90 : 10, thus presenting a
strong enantiomeric excess in favor of the 12S⁄ 13R
conformation The metabolite from the minor peak
(peak 1, Fig 4B) was identified by GC⁄ MS (Fig S5)
as 9,10:12,13-diepoxyoctadecanoic acid after
puri-fication and derivatization CYP77A4 was also able
to catalyze its formation in incubations with purified
12,13-epoxyoctadeca-9-enoic acid (data not shown)
Metabolism of a-linolenic acid by CYP77A4 The incubation of a-linolenic acid (C18:3) with micro-somes from yeast expressing CYP77A4 led to the formation of one major radioactive peak, as shown on the radiochromatogram in Fig 6 (peak 2, Fig 6B) It results from a reaction catalyzed by CYP77A4, as it requires the presence of NADPH and is not formed with microsomes from yeast transformed with a void plasmid (Fig 6C) or on incubation with boiled micro-somes (Fig 6D) The shape of this peak suggests that
it contains more than one metabolite After purifica-tion, acidic treatment and derivatizapurifica-tion, GC⁄ MS analysis showed that it was indeed a mixture of the three epoxide derivatives of a-linolenic acid
The mass spectrum of derivatized metabolite 1 (Fig S6) showed ions at m⁄ z (relative intensity, %) values of 73 (100%) [(CH3)3Si+], 75 (14%) [(CH3)2Si+=O], 439 (4%) (M-31) (loss of OCH3from the methyl ester), 455 (1%) (M-15) (loss of a methyl from the trimethylsilyl group), 470 (0.5%) (M) The mass spectrum also showed ions at 171 (40%) and 299 (44%), resulting from the cleavage between two hydroxyls carrying the trimethylsilyl group generated
by hydrolysis in perchloric acid This fragmentation pattern is characteristic of the derivative after acidic hydrolysis of 12,13-epoxyoctadeca-9,15-dienoic acid (M = 470 gÆmol)1) which represents 87% of the metabolites The mass spectrum of derivatized meta-bolite 2 (Fig S6) showed ions at m⁄ z (relative inten-sity, %) values of 73 (100%) [(CH3)3Si+], 75 (17%) [(CH3)2Si+=O], 439 (2%) (M-31) (loss of OCH3from the methyl ester), 455 (0.5%) (M-15) (loss of a methyl from the trimethylsilyl group), 470 (1%) (M) The mass spectrum also showed ions at 211 (11%) and 259 (81%), resulting from the cleavage between two hydroxyls carrying the trimethylsilyl group generated
by hydrolysis in perchloric acid This fragmentation pattern is characteristic of the derivative after acidic hydrolysis of 9,10-epoxyoctadeca-12,15-dienoic acid (M = 470 gÆmol)1) which represents 7% of the meta-bolites The mass spectrum of derivatized metabolite 3 (Fig S6) showed ions at m⁄ z (relative intensity, %) values of 73 (100%) [(CH3)3Si+], 75 (21%) [(CH3)2Si+=O], 439 (3%) (M-31) (loss of OCH3from the methyl ester), 455 (0.5%) (M-15) (loss of a methyl from the trimethylsilyl group), 470 (4%) (M) The mass spectrum also showed ions at 131 (67%) and 339 (20%), resulting from the cleavage between two hydroxyls carrying the trimethylsilyl group generated
by hydrolysis in perchloric acid This fragmentation pattern is characteristic of the derivative after acidic hydrolysis of 15,16-epoxyoctadeca-9,12-dienoic acid
Fig 5 Radiochromatographic resolution by HPLC of the
enantio-mers of 12,13-epoxyoctadeca-9-enoic acid produced by CYP77A4.
(A) After incubation of linoleic acid with microsomes from yeast
expressing CYP77A4, the 12,13-epoxyoctadeca-9-enoic acid
produced (peak 2, Fig 4B) was purified, methylated and subjected
to chiral HPLC analysis with 100% heptane at a flow rate of
0.5 mLÆmin)1 (B) Structures of the enantiomers.
Trang 8(M = 470 gÆmol)1) which represents 6% of the meta-bolites Kinetic parameters from the reaction of substrate oxidation are Km,app= 29 ± 4 lm and
Vmax,app = 38 ± 2 nmolÆmin)1Ænmol)1 P450 (Fig S3) Metabolites from the minor peak (peak 1, Fig 6B) have not been identified
Fig 7 Radiochromatographic resolution by TLC of metabolite gen-erated in the incubation of 12,13-epoxyoctadeca-9-enoic acid with microsomes or cytosol from A thaliana Microsomes (350 lg pro-tein) or cytosol (600 lg propro-tein) from A thaliana was incubated with 100 l M of 12,13-epoxyoctadeca-9-enoic acid for 20 min at
27 C Incubation was stopped by the addition of 20 lL of acetoni-trile (containing 0.2% acetic acid) and directly spotted onto TLC (A) Experiment performed with microsomes (B) Experiment performed with boiled microsomes (C) Experiment performed with cytosol (D) Experiment performed with boiled cytosol Peak S, 12,13-epoxy-octadeca-9-enoic acid; peak 1, 12,13-dihydroxy12,13-epoxy-octadeca-9-enoic acid The structure of the metabolite is described in (E).
Fig 6 Radiochromatographic resolution by TLC of metabolites
gen-erated in incubations of a-linolenic acid with microsomes from yeast
expressing CYP77A4 Microsomes were incubated with 100 lm
[1- 14 C]a-linolenic acid in the absence (A) or presence (B) of NADPH.
Incubations were performed at 27 C and contained 20 pmol of
CYP77A4 They were stopped after 30 min by the addition of 20 lL
of acetonitrile (containing 0.2% acetic acid) and directly spotted onto
TLC Peak S, a-linolenic acid; peak 1, non-identified; peak 2, mixture
of 12,13-epoxyoctadeca-9,15-dienoic,
9,10-epoxyoctadeca-12,15-die-noic and 15,16-epoxyoctadeca-9,12-die9,10-epoxyoctadeca-12,15-die-noic acids Experiments in
(C) and (D) were performed as in (B), but with microsomes from
yeast transformed with a void plasmid (C) or with boiled
micro-somes (D) The structures of the metabolites are described in (E).
Trang 9Requirements for CYP77A4 activity
To check the importance of the double bond for
CYP77A4 activity, we tested as potential substrates
stearic acid (C18:0), which is saturated, and linolelaidic
acid, which is linoleic acid containing trans double
bonds No metabolites were detected on TLC after
incubation of radiolabeled C18:0with microsomes from
yeast expressing CYP77A4 (data not shown) To test
linolelaidic acid, which is not available radiolabeled,
we performed two experiments In the first experiment,
we incubated radiolabeled linoleic acid with
micro-somes in the presence of an increasing concentration
of unlabeled linolelaidic acid, and did not detect any
inhibition of epoxidation of linoleic acid (data not
shown) In a second experiment, we ran GC⁄ MS
anal-ysis after the incubation of linolelaidic acid with yeast
microsomes expressing CYP77A4, and did not detect
any metabolite (data not shown) Together, this shows
that CYP77A4 requires the presence of unsaturation
to metabolize C18 fatty acids; furthermore,
unsatura-tion must be in the cis configuraunsatura-tion
Hydrolysis of vernolic acid in microsomes and
cytosol from A thaliana
In order to test whether the metabolites generated by
CYP77A4 were end products or could be substrates of
other enzymatic systems (i.e epoxide hydrolase) from
A thaliana, we purified vernolic acid that was produced
by CYP77A4 (peak 2, Fig 4B) This epoxide was
sub-sequently incubated with microsomes isolated from
A thalianaseedlings The results are presented in Fig 7
A peak of radioactivity (peak 1, Fig 7A) was detected
after resolving the products of reaction on TLC No
metabolite was formed if the microsomes were boiled
before incubation (Fig 7B) This demonstrates the
enzy-matic origin of the metabolite from peak 1 The mass
spectrum of this derivatized metabolite (Fig S7) showed
ions at m⁄ z (relative intensity, %) values of 73 (100%)
[(CH3)3Si+], 75 (17%) [(CH3)2Si+=O], 457 (2%)
(M-15) (loss of a methyl from the trimethylsilyl group)
The mass spectrum also showed ions at 173 (40%) and
299 (8%), resulting from the cleavage between two
hydroxyls carrying the trimethylsilyl group This
frag-mentation pattern is characteristic of the derivative of
12,13-dihydroxyoctadeca-9-enoic acid (M = 472 gÆ
mol)1) The same results were obtained when incubation
was carried out with the cytosolic fraction of A thaliana
(Fig 7C) Together, these experiments show that
vernol-ic acid produced by CYP77A4 can be converted to the
corresponding diol by microsomal and cytosolic epoxide
hydrolase (Fig 8) These epoxide hydrolases can also
convert epoxides from C18:3into the corresponding diols (data not shown)
Discussion
A new approach, based on an in silico analysis of pub-licly available transcriptome data (http://www-ibmp u-strasbg.fr/~CYPedia/), has been developed recently for the mapping of cytochromes P450 onto specific metabolic pathways based on large-scale co-expression analysis [33] This approach showed that CYP77A4 was co-regulated across 167 developmental samples (cover-ing more than 400 publicly available Affymetrix micro-array data sets) with a set of enzymes implicated in fatty acid metabolism Although co-expression correlations were relatively low compared with other co-expressed genes acting in a common pathway [33], with Pearson correlation coefficients not exceeding 0.75, it was strik-ing that the top eight co-expressed genes with CYP77A4 have been functionally characterized as being involved
in fatty acid metabolism (http://www-ibmp.u-strasbg.fr/
~CYPedia/CYP77A4/CoExpr_CYP77A4_Organs.html)
We thus found it worthwhile to test experimentally the hypothesis generated by this bioinformatic approach and to elucidate the physiological role of CYP77A4, also because no function has been reported for members belonging to this cytochrome P450 family to date Heterologous expression of CYP77A4 in an engineered strain of yeast, and incubations of a diverse set of fatty acids with yeast microsomes, allowed us to confirm the capacity of this newly characterized P450 to metabolize fatty acids, highlighting the predictive power of the
in silico co-expression analysis Based on phylogenetic reconstructions [36], CYP77A4 belongs to the CYP71
Fig 8 Conversion of linoleic acid to 12,13-dihydroxyoctadeca-9-enoic acid by CYP77A4 and epoxide hydrolases from A thaliana (A) Linoleic acid (B) Epoxyoctadeca-9-enoic acid (C) 12,13-Dihydroxyoctadeca-9-enoic acid.
Trang 10clan and, within this clan, forms a basal clade with the
CYP89, CYP753 and CYP752 families, none of which
has been functionally characterized In contrast, most
functionally characterized plant fatty acid hydroxylases
belong to the divergent CYP86 clan (including CYP86
and CYP94 families) Both the CYP71 and CYP86 clans
appear to have evolved independently within the green
plant lineage, as they are evolutionary separated by
fam-ilies that pre-date land plant evolution [36] Thus, a
function of CYP77A4 as a fatty acid-metabolizing
enzyme would not have been predicted based on
phylo-genetic reconstructions, again highlighting the power of
the co-expression analysis approach, which is
indepen-dent of sequence or structural similarities
On the model substrate lauric acid, CYP77A4
hydroxylated predominantly the x-1 carbon, which
cor-responds to a carbon in the environment of unsaturation
in oleic (C18:1), linoleic (C18:2) and a-linolenic (C18:3)
acids, the common physiological C18 fatty acids in
plants We therefore assayed these compounds as
poten-tial substrates and demonstrated that CYP77A4 was
able to produce, in vitro, epoxide derivatives of these
fatty acids Investigations on the members of the CYP2
family in animals have previously demonstrated that the
regioselectivity and enantioselectivity of epoxidation are
cytochrome P450 dependent [37,38] For CYP77A4, the
requirement of unsaturation, in the cis configuration,
together with the regioselectivity and enantioselectivity
observed, probably reflect steric constraints on the
sub-strate in the active site The fact that C18:2is epoxidized
first exclusively on the D12 unsaturated position, with
strong enantiomeric excess (the epoxide formed is a
mix-ture of 12S⁄ 13R 12R ⁄ 13S in the ratio 90 : 10), shows
that it is probably hindered in the active site, suggesting
that it could be a physiological substrate
In animals, epoxides of arachidonic acid (C20:4),
formed by epoxidases (mainly belonging to the CYP2
family), are well documented This is mainly a result
of the large array of biological effects attributed to
epoxyeicosatrienoic acids (EETs) For example,
activa-tion of CYP epoxidases in endothelial cells is a key
step in vasodilatation events [14] EETs also play a
major role in cell proliferation and angiogenesis via
the activation of an epidermal growth factor [39–41]
Over-expression of CYP2C9 and exogenous
applica-tion of EETs to cultured endothelial cells are
associ-ated with angiogenesis [41,42] CYP2C and CYP2J2
have also been shown to be expressed in different
tumor tissues [43,44] Epoxides of fatty acids are less
described in plants, and only a few biological activities
have been attributed to them The discovery of such
activities in plants might help to understand the
physi-ological role of CYP77A4 This lack of data could
explain the small amount of information available today concerning the ability of plant enzymes to gener-ate epoxides of fatty acids, despite the fact that this type of reaction was described for the first time more than three decades ago [23] Thus, the discovery of CYP77A4 carrying such activity opens the door not only for detailed biochemical characterizations, but also for an understanding of the physiological role of epoxides of fatty acids in plants
In addition to cytochromes P450, two distinct types
of plant enzyme, unrelated to cytochrome P450, with epoxidase activity, have been described The first, a peroxygenase, was reported in Vicia faba [28] and Gly-cine max [29] This type of enzyme uses hydroperox-ides as cofactors to catalyze the epoxidation of fatty acids The second, described by Lee et al [31], is a non-heme di-iron enzyme, also named ‘desaturase-like’ enzyme It thus appears that fatty acid epoxidation in plants can be facilitated by evolutionarily divergent sets of enzymes, further suggesting a pivotal role of these epoxides or derivatives thereof
CYP77A4, described in this work, catalyzed the oxy-gen incorporation into double bonds of oleic (C18:1), linoleic (C18:2) and a-linolenic (C18:3) acids, but did not metabolize saturated stearic acid (C18:0) Furthermore,
it did not metabolize linolelaidic acid, which is the homolog of linoleic acid possessing two trans double bonds, not commonly found in natural fatty acids These observations suggest that the physiological func-tion of CYP77A4 could be epoxidafunc-tion of unsaturated
C18fatty acids This hypothesis is supported by in silico co-expression analysis, showing that CYP77A4 is co-regulated with a stearoyl acyl carrier protein desat-urase and a putative epoxide hydrolase [33] Cahoon
et al [32], in E lagascae seed, identified a cytochrome P450, classified as CYP726A1, able to convert linoleic acid into 12,13-epoxyoctadeca-9-enoic acid (vernolic acid) CYP77A4 differs from this enzyme; indeed, it metabolizes free fatty acids, whereas CYP726A1 meta-bolizes fatty acids incorporated into phosphatidylcho-line [32,45] The physiological role of CYP77A4 is unlikely to be the production of fatty acid epoxides for accumulation in seeds as, unlike E lagascae and plants belonging to the Aesteraceae genera, such as Crepis palaestina, A thaliana does not store fatty acid epox-ides Cytochrome P450-dependent fatty acid oxidases in plants have been mainly investigated with regard to cutin synthesis [19] Cutin consists of a biopolymer of fatty acids belonging to the protective envelope of plants: the cuticle [20] Epoxides of fatty acids may rep-resent up to 60% of cutin monomers [46,47] Cutin anal-ysis of A thaliana has been performed recently [48], and 18-hydroxy-9,10-epoxystearic acid was shown to be