Epoxyalcohol synthase of Ectocarpus siliculosus. First CYP74-related enzyme of oxylipin biosynthesis in brown algae

Biochimica et Biophysica Acta 1862 (2017) 167–175

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Epoxyalcohol synthase of Ectocarpus siliculosus. First CYP74-related enzyme of oxylipin biosynthesis in brown algae☆ Yana Y. Toporkova, Valeria S. Fatykhova, Yuri V. Gogolev, Bulat I. Khairutdinov, Lucia S. Mukhtarova, Alexander N. Grechkin ⁎ Kazan Institute of Biochemistry and Biophysics, Russian Academy of Sciences, P.O. Box 30, Kazan 420111, Russia

a r t i c l e

i n f o

Article history: Received 11 June 2016 Received in revised form 16 October 2016 Accepted 14 November 2016 Available online 15 November 2016 Keywords: Oxylipins Cytochrome P450 CYP74 clan Epoxyalcohol synthase Brown alga Ectocarpus siliculosus

a b s t r a c t Enzymes of CYP74 family play the central role in the biosynthesis of physiologically important oxylipins in land plants. Although a broad diversity of oxylipins is known in the algae, no CYP74s or related enzymes have been detected in brown algae yet. Cloning of the first CYP74-related gene CYP5164B1 of brown alga Ectocarpus siliculosus is reported in present work. The recombinant protein was incubated with several fatty acid hydroperoxides. Linoleic acid 9-hydroperoxide (9-HPOD) was the preferred substrate, while linoleate 13-hydroperoxide (13-HPOD) was less efficient. α-Linolenic acid 9- and 13-hydroperoxides, as well as eicosapentaenoic acid 15-hydroperoxide were inefficient substrates. Both 9-HPOD and 13-HPOD were converted into epoxyalcohols. For instance, 9-HPOD was turned primarily into (9S,10S,11S,12Z)-9,10-epoxy11-hydroxy-12-octadecenoic acid. Both epoxide and hydroxyl oxygen atoms of the epoxyalcohol were incorporated mostly from [18O2]9-HPOD. Thus, the enzyme exhibits the activity of epoxyalcohol synthase (EsEAS). The results show that the EsEAS isomerizes the hydroperoxides into epoxyalcohols via epoxyallylic radical, a common intermediate of different CYP74s and related enzymes. EsEAS can be considered as an archaic prototype of CYP74 family enzymes. © 2016 Elsevier B.V. All rights reserved.

1. Introduction Enzymes of CYP74 family play a central role in the plant lipoxygenase pathway [1–3]. They control the conversions (dehydration or isomerization) of fatty acid hydroperoxides to numerous oxylipins, including the bioactive compounds like jasmonates, divinyl ethers, aldehydes and others [1–3]. The diversity of CYP74-related enzymes has recently been extended to some proteobacteria, rhizobacteria and

Abbreviations: EAS, epoxyalcohol synthase; EsEAS, E. siliculosus epoxyalcohol synthase; DES, divinyl ether synthase; AOS, allene oxide synthase; HPL, hydroperoxide lyase; 9-H(P)OD, (9S,10E,12Z)-9-hydro(pero)xy-10,12-octadecadienoic acid; 9-H(P)OT, (9S,10E,12Z,15Z)-9-hydro(pero)xy-10,12,15-octadecatrienoic acid; 13-H(P)OD, (9Z,11 E,13S)-13-hydro(pero)xy-9,11-octadecadienoic acid; 13-H(P)OT, (9Z,11E,13S,15Z)-13hydro(pero)xy-9,11,15-octadecatrienoic acid; 15-H(P)EPE, (5Z,8Z,11Z,13E,15S,17Z)-13hydro(pero)xy-5,8,11,13,17-eicosapentaenoic acid; IMAC, immobilized metal affinity chromatography; Me, methyl; TMS, trimethylsilyl; SIM, selected ion monitoring; COSY, correlation spectroscopy; HSQC, heteronuclear single quantum coherence spectroscopy; HMBC, heteronuclear multiple-bond correlation spectroscopy; NOESY, nuclear Overhauser effect spectroscopy. ☆ This paper is dedicated to the memory of Professor Victor E. Vaskovsky (1935-2016), a distinguished researcher in the fields of lipidology and marine biology, whose friendly care and encouragement were essential for this work. ⁎ Corresponding author. E-mail address: [email protected] (A.N. Grechkin).

http://dx.doi.org/10.1016/j.bbalip.2016.11.007 1388-1981/© 2016 Elsevier B.V. All rights reserved.

Metazoa [4]. These new members expanded the CYP74 diversity from family to the clan (for definitions of P450 family and clan see [5]). Most of these new clan members are waiting for their cloning and studies of specificity of their action. Besides, typical CYP74 products like divinyl ethers have been detected in some living organisms where CYP74-related genes and enzymes have not been encountered yet. Marine algae possess a broad diversity of oxylipins [6–9]. For instance, the brown alga Laminaria sinclairii [10], and red alga Polyneura latissima [11] contain the divinyl ethers. Identical or analogous oxylipins are well known as the products of divinyl ether synthases (CYP74 family), which are widespread in higher plants [12]. The biosynthetic origin of algal divinyl ethers has not been revealed yet. Involvement of CYP74related enzymes in the biosynthesis of algal oxylipins seems to be very likely. Although the CYP74 family enzymes are widespread in land plants, none of CYP74 clan members has been detected in algae until recently. The freshly discovered allene oxide synthase (AOS) of green alga Klebsormidium flaccidum [13] presents the only exception. An unusual gene CYP5164B1 homologous to the CYP74s has been detected recently as a result of the annotation of brown alga Ectocarpus siliculosus (Ectocarpaceae, Phaeophyceae) genome [14]. These considerations prompted us to clone the CYP5164B1 gene and test putative fatty acid hydroperoxide metabolizing activity of the recombinant protein. The results are reported in the present paper.

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2. Materials and methods 2.1. Materials [18O]Water (95% 18O) and 18O2 gas (88% 18O) were purchased from ZAO Neogaz (Moscow, Russia). [2H6]Benzene (99.5% 2H) was acquired from FSUE RSC “Applied Chemistry” (St. Petersburg, Russia). Linoleic, αlinolenic, and eicosapentaenoic acids, as well as the soybean lipoxygenase type V, were purchased from Sigma. [1-14C]Linoleic acid (2.072 MBq/ μmol) was purchased from Perkin Elmer (former New England Nuclear). NaBH4 and silylating reagents were purchased from Fluka (Buchs, Switzerland). (9S,10E,12Z)-9-Hydroperoxy-10,12-octadecadienoic (9HPOD), and (9S,10E,12Z,15Z)-9-hydroperoxy-10,12,15-octadecatrienoic (9-HPOT) acids, as well as the [1-14C]9-HPOD (5.78 kBq/μmol), were prepared by incubation of linoleic, α-linolenic, and [1-14C]linoleic acids, respectively, with the recombinant maize 9-lipoxygenase (GeneBank: AAG61118.1) [15] at 0 °C, Na-phosphate buffer (100 mM, pH 6.0), under continuous oxygen bubbling. (9Z,11E,13S)-13-Hydroperoxy9,11-octadecadienoic (13-HPOD), (9Z,11E,13S,15Z)-13-hydroperoxy9,11,15-octadecatrienoic (13-HPOT), and (5Z,8Z,11Z,13E,15S,17Z)-13hydroperoxy-5,8,11,13,17-eicosapentaenoic (15-HPEPE) acids were obtained by incubation of linoleic, α-linolenic, and eicosapentaenoic acids, respectively, with the soybean lipoxygenase type V at 23 °C, Tris-HCl buffer (50 mM, pH 9.0), under continuous oxygen bubbling. The extracted hydroperoxides (as free carboxylic acids) were purified by normal phase HPLC (NP-HPLC) on the Kromasil Si columns (7 μm; 4.0 × 250 mm; Elsico, Moscow, Russia) under the isocratic elution with the solvent mixture hexane-isopropanol-acetic acid 98.1:1.8:0.1 (v/v) at a flow rate of 0.4 mL/min. Hydroperoxides were chromatographically pure and at least 98% optically pure, as judged by chiral phase HPLC [16]. Labelled [18O2-hydroperoxy]9-HPOD was obtained similarly by incubations under 18O2 atmosphere. 2.2. Bioinformatics methods The search for CYP74-related proteins predicted from E. siliculosus genome data was performed using the theoretical BLAST analyses in NCBI and Online Resource for Community Annotation of Eukaryotes (ORCAE) databases. Primer construction and multiple sequences alignments were performed using Vector NTI program (Invitrogene, U.S.A.). The BLAST analyses of the CYP74s were performed using the protein NCBI BLAST tool. Multiple protein alignments have been made using the Clustal Omega. The phylogenetic trees of selected CYP74s and P450s were built using the ClustalW2 – Phylogeny and viewed with the TreeView software.

was eluted from the cartridge using 50 mM histidine. The recombinant protein concentration was determined using the Quant-iT™ Protein HS Assay Kit (Invitrogen, USA). The haemoprotein concentration was estimated using the pyridine haemochromogen assay [17]. The relative purity of recombinant protein was estimated by SDS-PAGE and staining of the gel with Coomassie brilliant blue R-250 (Supplementary Fig. 2). Enzymatic activity of the purified recombinant enzyme was determined by monitoring the decrease of the signal at 234 nm in a Perkin Elmer Lambda 25 UV-VIS spectrophotometer with 40 μM substrate concentration. The analyses were performed in 0.6 mL of Na-phosphate buffer (100 mM, pH 7.0) at 25 °C. The initial linear regions of the kinetic curves were used to calculate the rates. The molar extinction coefficient for 9and 13-hydroperoxides of linoleic acid at 234 nm is 25,000 M−1 cm−1. Five independent experiments were performed for each specified variant. 2.4. Incubations of recombinant enzyme with substrates The recombinant enzyme (25 μg) was incubated with 100 μg of 9HPOD, 9-HPOT, 13-HPOD, 13-HPOT, or 15-HPEPE in Na-phosphate buffer (100 mM, 10 mL), pH 7.0, 4 °C, for 15 min. The reaction mixture was acidified to pH 6.0, and the products were extracted with hexane/ethyl acetate (1:1, by volume) mixture, methylated with ethereal diazomethane and trimethylsilylated with pyridine/ hexamethyldisilazane/trimethylchlorosilane (1:1:1, by volume) mixture at 23 °C for 30 min. Then the silylation reagents were evaporated in vacuo. The dry residue was dissolved in 100 μL of hexane and subjected to GC-MS analyses. When specified, the products were reduced with NaBH4, then methylated and trimethylsilylated. Alternatively, the products of NaBH4 reduction were hydrogenated over PtO2, then methylated and trimethylsilylated. Products (without or with the preliminary NaBH4 reduction) were analyzed as Me esters/TMS derivatives (Me/TMS) by GC-MS as described before [18]. For micropreparative isolation of products, the recombinant enzyme (25 μg) was incubated with 18.5 kBq of [1-14C]9-HPOD (5.78 kBq/μmol) in the same way. Products were extracted and methylated with ethereal diazomethane. The resulting methyl esters were separated and purified by NP-HPLC as described in the next section. Incubations of the recombinant enzyme (25 μg) with 100 μg of [18O2-hydroperoxy]9-HPOD were performed in the same way. Alternatively, the enzyme was incubated under identical conditions with the unlabelled 9-HPOD in an 18O2 atmosphere or in the [18O]water medium (1 mL). The products (Me/TMS) were analysed by the selected ion monitoring (SIM) GC-MS. The selected ions are specified in the Results. The data of the selected ion chromatograms were quantified with the Shimadzu data analysis software.

2.3. Expression and purification of recombinant enzyme The CYP5164B1 coding sequence was adapted for expression in Escherichia coli cells (Supplementary Fig. 1) and synthesized in Evrogen Company (Moscow, Russia). The sequence was cloned into the pET-23a vector (Novagen, USA) to yield the target recombinant protein with Histag at C-terminus. The recombinant gene was expressed in BL21(DE3)pLysS strain cells (Novagen, USA) as follows. An overnight culture (10 mL) of bacteria was inoculated into 1 L of Luria-Bertani medium supplemented with one volume of mineral medium M9. Bacteria were grown at 37 °C, 250 rpm to an OD600 of 0.6. Expression of the target gene was induced by addition of 0.5 mM isopropyl-β-D-1thiogalactopyranoside to the medium. Simultaneously, the medium was supplied with 5-aminolevulinic acid to facilitate the haem formation. Induction was controlled by the 12% SDS-PAGE analysis of crude cellular extracts. Bacteria were harvested by centrifugation for 15 min at 4500 rpm at 4 °C and lysed with BugBuster Protein Extraction Reagent (Novagen, USA). Purification of His-tagged recombinant protein was performed using Bio-Scale Mini Profinity IMAC cartridge in BioLogic LP chromatographic system (Bio-Rad, USA). The recombinant enzyme

2.5. Separation and purification of individual oxylipins by normal phase HPLC Products (Me esters) were separated by NP-HPLC on MachereyNagel EC 250/4.6 Nucleodur 100–3 SiOH column under elution with hexane – isopropanol 98:2 (by volume), flow rate 0.4 mL/min. Ultraviolet (UV) detection (190–370 nm) was performed with Shimadzu SPD-M20A diode array detector. Radioactivity was detected by the HPLC radiomonitor model 171 (Beckman Instruments, Fullerton, CA, USA) with a solid scintillator cell (125 μm). Separate products were collected after NP-HPLC separation, redissolved in [2H6]benzene and subjected to NMR spectral records. 2.6. Methods of instrumental analyses The UV spectra of products were scanned during the incubations of the recombinant protein with fatty acid hydroperoxides with Varian Cary 50 spectrophotometer. The UV spectra of reaction products were recorded with the same instrument. Alternatively, the UV spectra of

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products were recorded on-line during the HPLC separations using an SPD-M20A diode array detector (Shimadzu, Japan). Products were analyzed as methyl esters or methyl esters/TMS derivatives by GC-MS as described before [18]. GC-MS analyses were performed using a Shimadzu QP5050A mass spectrometer connected to Shimadzu GC-17A gas chromatograph equipped with an MDN-5S (5% phenyl 95% methylpolysiloxane) fused capillary column (length, 30 m; ID 0.25 mm; film thickness, 0.25 μm). Helium at a flow rate of 30 cm/s was used as the carrier gas. Injections were made in the split mode using an initial column temperature of 120 °C, injector temperature 230 °C. The column temperature was raised at 10 °C/min until 240 °C. The electron impact ionization (70 eV) has been used. The 1H–NMR, 2D–COSY, HSQC, HMBC and NOESY spectra were recorded with a Bruker Avance III 600 instrument (600 MHz, [2H3]acetonitrile, 296 K). 3. Results 3.1. Bioinformatics' analysis of E. siliculosus CYP5164B1 enzyme Currently, there are three partial gene sequences of E. siliculosus encoding the putative P450 proteins (related to the CYP74 clan) in the GenBank database. These are the pseudogene CBN74955.1 (903 bp) and two genes, CBN75517.1 (1113 bp) and CBN74956.1 (861 bp). CBN75517.1 is a partial sequence of the CYP5164B1 gene, lacking the 5′-terminus. The mRNA coding sequence Esi_0111_0095 (1434 bp) encoding the CYP5164B1 protein of 477 AA (Supplementary Fig. 3) is deposited at the ORCAE (Online Resource for Community Annotation of Eukaryotes) database (Ectsi_mRNA_LATEST), see the Supplementary references (Supplementary Material). Furthermore, the expressed sequence tags (ESTs) FP272245, FP278707, and FP286043 (see the Supplementary references), corresponding to the CYP5164B1 gene, have been detected by transcriptomic analyses of E. siliculosus responding to abiotic stresses [19]. Alignment of CYP5164B1 with other P450s suggested the relation between this protein and the CYP74 clan members (Fig. 1). The CYP5164B1 is most similar to the CYP74 clan proteins of methylobacteria. For instance, it possesses 22% identity (at 87% sequence coverage) to the CYP74 clan proteins of Methylobacterium nodulans hydroperoxide lyase (MnHPL, GenBank: ACH43051) and Methylobacterium sp. P450 (GenBank: WP_018263767). The next most homologous proteins are CYP74 family members of plants. The CYP5164B1 is reliably built to the separate subtrees with CYP74 clan members of plants, proteobacteria, and metazoa on phylogenetic trees of P450s. For example, the multiple alignments of CYP5164B1 toward the Arabidopsis P450s, the CYP5164B1 was built to a separate branch of the phylogenetic tree along with CYP74A1 and CYP74B1, separately from monooxygenases, as seen in the Supplementary Fig. 4. The phylogenetic tree of CYP5164B1 and P450s of all phyla is presented in Supplementary Material (Supplementary Fig. 5). The CYP5164B1, as well as the related hypothetical proteins of E. siliculosus, are built to a separate subtree with CYP74 family enzymes of higher plants and CYP74 clan members of methylobacteria, apart from monooxygenases (Supplementary Fig. 5). The CYP5164B1 was aligned with selected CYP74 clan proteins using the Clustal Omega. The resulting multiple alignments were used for generation of a phylogenetic tree with ClustalW2 – Phylogeny. The phylogenetic tree is presented in Fig. 2. The CYP5164B1 was built to a subtree with CYP74 clan proteins of Methylobacterium species and those of Metazoa (Fig. 2). The allene oxide synthase KfAOS of green alga K. flaccidum [13] was built between the mentioned subtree and the CYP74 family proteins of land plants (Fig. 2). BLAST analyses of land plants versus CYP5164B1 as a query revealed its highest identity to proteins of CYP74F (e.g., XP_002451776 of Sorghum bicolor) and CYP74B (e.g., XP_002870238 of Arabidopsis lyrata) subfamilies. Noteworthy, the CYP5164B1 protein possesses a nine amino acid insertion at the haem-binding domain (Fig. 1), which is a distinctive

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feature of all CYP74 clan members [2]. As seen in Fig. 1, the CYP5164B1, as well as all proteins of the CYP74 clan, possess this insertion prior the haem binding cysteine residue while the monooxygenases CYP11A1 and CYP19A1 lack it. Peptide fragment SEPGRVGGM (positions 409–417) at the haem-binding domain of CYP5164B1 corresponds to this insertion. Alignment of CYP5164B1 with monooxygenases and CYP74s of higher plants (Supplementary Fig. 6) revealed one more distinctive feature. Monooxygenases possess a typical A/G-G-X-D/E-T/S-T/ S motif at their oxygen binding domain (I-helix central domain, IHCD, in CYP74s), as seen in Supplementary Fig. 6A. The IHCD of CYP5164B1 has an alteration compared to monooxygenases. This is the glutamine (Q272) instead of the D/E residue (conserved in monooxygenases). Notably, essentially all CYP74 enzymes have a synonymous amino acid residue at this site (Supplementary Fig. 6B), namely the asparagine, e.g. N321 in AtAOS. This conserved N residue is involved in the initial stage of CYP74 catalysis, the homolytic cleavage of the O-O bond of fatty acid hydroperoxides [4]. This similarity indicates that the Q residue of CYP5164B1 might play the same role. Alignment of ORF sequence encoding the CYP5164B1 enzyme with the genomic sequence mapped a gene in a locus 487,736–501,540 and identified its structure. The coding sequence is interrupted into ten exons by eight short introns and one 7396 bp length intron (Supplementary Fig. 7). This sequence entirely corresponds to the full mRNA sequence Esi0111_0095 detected in E. siliculosus transcriptome and deposited at the ORCAE database (Supplementary references, see the Supplementary Material). The CYP5164B1 coding sequence adapted for expression in E. coli cells (Supplementary Fig. 1) has been custom synthesized by the Evrogen Company (Russia) and cloned into the vector pET-23a to yield the target recombinant protein with His-tag at C-terminus. His-tagged recombinant protein was obtained in BL21(DE3)pLysS strain cells (Novagen, USA) and purified by metal affinity chromatography. The enzymatic activity was controlled using the ultraviolet spectroscopy by the decrease of fatty acid hydroperoxide absorbance at 234 nm. 3.2. Preliminary characterization of the recombinant CYP5164B1 Recombinant CYP5164B1 efficiently utilized 9- and 13-HPOD. The estimated turnover rate (kcat) of 9-HPOD was about eight times higher than that of 13-HPOD – 149 s−1 and 18.1 s−1 respectively. At the same time, CYP5164B1 exhibited essentially no activity towards 13-HPOT, 9-HPOT, and 15-HPEPE (Supplementary Fig. 8). Thus, the recombinant enzyme showed a preference for 9-HPOD as a substrate. The GC-MS analyses of NaBH4 reduced products (Me/TMS) of 9-HPOD and 13-HPOD conversions are illustrated in Figs. 3 and 4, respectively. Incubations with 9-HPOD afforded a single predominant product 1 (Fig. 3). In contrast, 9-HPOD remained unutilized after the control incubations of 9-HPOD with boiled CYP5164B1 and the empty vector protein preparation (Supplementary Fig. 9). Incubation of CYP5164B1 with 13-HPOD (Fig. 4) also resulted in one major product 2, but it was accompanied with three isomeric byproducts, eluting a little later than compound 2. Mass spectra of these minor by-products were identical to those of product 2. Identification of compounds 1 and 2 is described below. 3.3. Identification of product 1 The electron impact mass spectrum (Fig. 3A,B) of product 1 (Me/ TMS) exhibited M+ at m/z 398 (0.1%); [M – Me]+ at m/z 383 (0.7%); [M – C1/C8]+ at m/z 241 (3.6%); [M – C1/C9]+ at m/z 212 (3.7%); [M – C1/C10]+ at m/z 199 (97%). As shown in the fragmentation scheme (Fig. 3B inset), the intense fragment at m/z 199 indicates the presence of oxiranyl carbinol function with oxirane at C9/C10 and secondary alcohol (TMS) at C11. Catalytic hydrogenation of product 1 over PtO2 followed by methylation and trimethylsilylation afforded product 3. Its mass spectrum possessed [M – Me]+ at m/z 385 (3.4%); [M – C1/C7]+ at m/z 257 (21.2%); [M – C1/C9]+ at m/z 215 (20.4%); [M – C1/C10]+ at m/z

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Fig. 1. The P450s sequences alignments. The CYP74 family sequences: Pa, Parthenium argentatum; PaAOS, Q40778; Ca, Capsicum annuum; CaHPL, AAK27266; Lu, Linum usitatissimum; LuDES, ADP03054; Le, Lycopersicon esculentum; LeAOS3, NP_001234833; Mt, Medicago truncatula; MtHPL, CAC86897; Nt, Nicotiana tabacum; NtDES, AAL40900; Ra, Ranunculus acris; RaDES, AJU57209. The CYP74 clan sequences: Msp, Methylobacterium sp. 4–46; MspCYP74, WP_018263767; Mn, M. nodulans; MnHPL, ACH43051. The P450 monooxygenases sequences: Hs, Homo sapiens; HsCYP11A1, NP_000772; HsCYP19A1, NP_000094. The IHCD is numbered by 1–6; ERR-triad, PPV-domain and cysteinyl haem ligand are marked by ♦, ◊ and ● symbols, respectively.

201 (50.3%); [TMS]+ at m/z 73 (100%). The spectrum shows that product 3 has a structure of 9,10-epoxy-11-hydroxyoctadecanoic acid (Me/TMS). Thus, the mass spectral data substantiated the structure of 9,10-epoxy11-hydroxy-12-octadecenoic acid for compound 1. For further structural elucidation compound 1 (Me ester) was purified by radio-NP-HPLC after CYP5164B1 incubation with [1-14C]9-HPOD. NP-HPLC separation

revealed that compound 1 was composed of two epimers 1a and 1b presented at a ratio ca. 4:1. Both epimers were collected, and their NMR spectra were recorded. The NMR spectral data for compounds 1a and 1b are presented in Tables 1 and 2, respectively. Both products possessed one cis-double bond (J 12,13 = 11.0 Hz) and trans-epoxide ring (J9,10 = 2.3 and

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Fig. 2. The unrooted phylogenetic tree of CYP74 clan. Classified CYP74 subfamilies are marked with their letter designations (A, B, C, etc.). Subfamilies consisting of more than one member are outlined with unclosed curves (semi-ellipses). CYP74s of flowering plants: As, Allium sativum; AsDES, GI:83414021; At, Arabidopsis thaliana; AtAOS, CYP74A1, GI:15239032; AtHPL, CYP74B2, GI:3822403; Ca, C. annuum; CaHPL, CYP74B1, GI:1272340; Cm, Cucumis melo; CmHPL, CYP74C2, GI:14134199; Gm, Glycine max; GmCYP74C13, GI:100037482; Hb, Hevea brasiliensis; HbAOS, CYP74A9, GI:84028363; Hv, Hordeum vulgare, HvAOS2, GI:7452981; Le, L. esculentum; LeAOS1, GI:7581989; LeAOS2, CYP74A2, GI:7677376; LeAOS3, CYP74C3, GI:25991603; LeHPL, GI:7677378; LeDES, GI:11991245; Lu, L. usitatissimum; LuAOS, CYP74A1, GI:1352186; LuDES, CYP74B16, GI:310687282; Ms, Medicago sativa; MsHPL, GI:5830465; Mt, M. truncatula; MtHPL1, CYP74C12, GI:33504430; MtHPL3, CYP74B4, GI:63081244; Os, Oryza sativa; OsAOS, CYP74A4, GI:115455571; OsHPL1, CYP74E2, GI:115445057; OsHPL2, CYP74E1, GI:125538638; Pa, P. argentatum; PaAOS, CYP74A1, GI:218511958; Pd, Prunus dulcis; PdHPL, CYP74C5, GI:33300600; Pg, Psidium guajava; PgHPL, CYP74B5, GI:13183137; Pi, Petunia inflata; PiCYP74C9, GI:85720841; Ra, R. acris; RaDES, CYP74Q1, GI:768564485; Sm, Selaginella moellendorffii; CYP74L1, GI: 9645914; CYP74L2, GI: 9651730; CYP74M1, GI:9660714; CYP74M3, GI:9654395; St, Solanum tuberosum; StAOS2, GI:86769479; StAOS3, GI:56605358; StDES, CYP74D2, GI:12667099; Zm, Z. mays; ZmAOS, CYP74A19, GI: 223947589; ZmHPL, CYP74F2, GI:162462890. CYP74s of mosses: Pp, Physcomitrella patens; PpAOS1, CYP74A1, GI:22217985; PpAOS2, CYP74A8, GI:168014176; PpHPL, CYP74G1, GI:76057841. CYP74 clan members of proteobacteria: Mn, M. nodulans; MnHPL, GI:220926268; Msp, Methylobacterium sp. 4–46; MspCYP74, GI:170743950. CYP74 clan members of Metazoa: Ap, Acropora palmata (coral); ApAOS, GI:187948710; Bf, B. floridae (lancelet); BfEAS, GI:189312561. The tree was built with the Clustal Omega and ClustalW2 – Phylogeny and visualized with the TreeView software using the alignments of full amino acid sequences of the above listed CYP74 clan members. The preliminary BLAST analyses of CYP74s were performed using the protein NCBI BLAST tool.

2.2 Hz, respectively). So, the spectra confirmed the structures of oxiranyl carbinols for compounds 1a and 1b. The chemical shifts of H11 in two spectra were markedly different: 4.52 ppm (1a) and 4.25 ppm (1b). This difference suggests that H11 in compound 1a is relatively deshielded due to the spatial proximity of epoxide oxygen. The main Overhauser effects seen from the NOESY data are illustrated in Fig. 5. NOESY spectrum of product 1a possessed a strong cross-peak between H11 and H10 while the correlation between H11 and H9 was not observed. In contrast, the NOESY data for compound 1b showed a strong NOE correlation between H11 and H9, whereas a cross-peak between H11 and H10 was relatively weak. NOESY results (Fig. 5) unambiguously revealed the erythro and threo configurations for the epoxyalcohols 1a and 1b, respectively. These configurations are also fully consistent with the values of spin coupling constants J10,11 for compounds 1a and 1b (3.3 Hz and 5.1 Hz, see Tables 1 and 2) and correspond to the literature data [20,21]. The obtained data allowed us to ascribe the (9S,10S,11S) and (9S,10S,11R) configurations to products 1a and 1b, respectively, assuming the conservation of the original 9(S) configuration, which generally takes place during such conversions.

3.4. Identification of product 2 As mentioned above, incubations of CYP5164B1 with 13-HPOD resulted in one major product 2, which was accompanied with three byproducts, having slightly bigger retention times than compound 2 (Fig. 4A). Mass spectrum of product 2 (Fig. 4B) possessed M+ at m/z 398 (0.1%); [M – Me]+ at m/z 383 (2.2%); [M – n-pentyl]+ at m/z 327 (3.0%); [M – C12/C18]+ at m/z 285 (89.2%); [TMS]+ at m/z 73 (100%). The fragmentation patterns (Fig. 4B, inset) confirm the structure of 11hydroxy-12,13-epoxy-9-octadecenoic acid (Me/TMS) for compound 2. Catalytic hydrogenation of product 2 over PtO2 followed by methylation and trimethylsilylation yielded product 4, possessing [M – Me]+ at m/z 385 (9.7%); [M – Me – MeOH]+ at m/z 353 (1.9%); [M – CH3(CH2)4CHO]+ at m/z 301 (10.1%); [M – C12/C18]+ at m/z 287 (48.8%); 271 (17.2%); [M – (CH2)9COOMe]+ at m/z 215 (7.6%); [TMS]+ at m/z 73 (100%). The spectrum corresponds to the structure of 11-hydroxy-12,13epoxyoctadecanoic acid (Me/TMS), thus confirming the identification of compound 2. Minor peaks eluting closely after the product 2 (Fig. 4A) exhibited the identical mass spectral patterns. So, these are the stereoisomers of

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Fig. 4. GC-MS analyses of NaBH4-reduced products (Me/TMS) of 13-HPOD incubations with the recombinant EsEAS. A), the total ion current (TIC) GC–MS chromatogram of incubation products (Me esters TMS derivatives). Peak 2, 11-hydroxy-12,13-epoxy-9octadecenoic acid (Me/TMS). B), mass spectrum and fragmentation scheme for product 2 (Me/TMS). Conditions of incubation, extraction, derivatization and analysis are described in Materials and Methods.

[18O]water solution have shown that the rest of hydroxyl oxygen comes either from the atmosphere or from water (Table 3). 4. Discussion The obtained data demonstrated that CYP5164B1 is an epoxyalcohol synthase (EAS). No AOS, hydroperoxide lyase (HPL), or divinyl ether synthase (DES) products have been detected along with the epoxyalcohol. We ascribed the trivial name EsEAS (E. siliculosus epoxyalcohol synthase) Fig. 3. GC-MS analyses of NaBH4-reduced products (Me/TMS) of 9-HPOD incubations with the recombinant CYP5164B1 (EsEAS). A), the total ion current (TIC) GC–MS chromatogram of incubation products (Me esters TMS derivatives). Peak 1, 9.10-epoxy-11-hydroxy-12octadecenoic acid (Me/TMS). B), mass spectrum and fragmentation scheme for product 1 (Me/TMS), C), mass spectrum and fragmentation scheme for product 1 (Me/TMS) of the incubation of the EsEAS with [18O2-hydroperoxy]9-HPOD. Conditions of incubation, extraction, derivatization and analysis are described in Materials and Methods.

compound 2, presumably having a distinct stereo configuration at C11 or C12. The NMR spectra for products of 13-HPOD conversion were not recorded and their stereochemistry was not studied. 3.5. The 18O labelling experiments The 18O labelling experiments were performed to spread more light on the mechanism of the enzymatic reaction. First of all, the EsEAS was incubated with [18O2-hydroperoxy]9-HPOD. The mass spectrum of product 1 (Me/TMS) is depicted in Fig. 3C. The labelled compound 1 exhibited M+ at m/z 402 (which was 398 in the unlabelled product) and [M – Me]+ at m/z 387 (which was 383 in the unlabelled product). Selected ion monitoring (SIM) GC-MS analyses of compound 1 species possessing 2, 1 or 0 atoms of 18O were performed using [M – Me]+ at m/z 387.3, 385.3 and 383.3, respectively. Quantifications were conducted by estimation of areas of compound 1 peaks in the corresponding ion chromatograms. As seen from the results presented in Table 3, the predominant part of epoxide oxygen and ca. 70% of hydroxyl oxygen were incorporated from [18O2-hydroperoxy]9-HPOD. Incubations of the EsEAS with the unlabelled 9-HPOD under the 18O atmosphere or in

Table 1 The 1H–NMR data (1H–NMR, 2D–COSY, HSQC, HMBC) for compound 1a (Me ester). 600 MHz, [2H6]benzene, 303 K. Position number

13 C chemical shifts (ppm); functional group

1 H chemical shifts (ppm); multiplicity; coupling constant (Hz)

Heteronuclear multiple bond correlation

1 2 3 4 5 6 7 8a 8b 9

173.37; COOMe 34.22; CH2 25.35; CH2 29.48; CH2 29.48; CH2 29.47–29.85; CH2 29.83; CH2 31.97; CH2 31.97; CH2 54.87; CH

COOMe, H2 H3, COOMe H2, H4 H3

10 11

60.47; CH 65.91; CH

12

128.35; CH

13

133.60; CH

14 15 16 17 18 (1) (11)

28.45; CH2 26.36; CH2 31.82; CH2 23.05; CH2 14.29; CH3 51.03; COOMe OH

n 2.11; t; 7.5 (H3) 1.53; m; 7.5 (H2) 1.14; m 1.07–1.31; m 1.07–1.31; m 1.20–1.31; m 1.43; m 1.35; m 2.92; ddd; 6.3 (H8a); 5.7 (H8b); 2.3 (H10) 2.70; dd; 3.3 (H11); 2.3 (H9) 4.52 ddd (dt-like); 8.0 (H12); 3.3 (H10); 2.7 (OH) 5.47; ddt (AM); 11.0 (H13); 8.0 (H11); 1.4 (H14) 5.51; ddt (AM); 11.0 (H12); 7.3 (H14); 1.0 (H11) 1.98; m 1.27; m 1.17–1.31; m 1.23; m 0.87; t; 7.1 (H17) 3.37; s 1.56; d; 2.7 (H11)

H8a, H8b H7, H9 H7, H9 H8a, H8b, H10 H9, H11 H10, H12, OH H11, H13 H12, H14 H13, H15 H14, H16 H15 H18 H17 H2 H11

Y.Y. Toporkova et al. / Biochimica et Biophysica Acta 1862 (2017) 167–175 Table 2 The 1H–NMR data (1H–NMR, 2D–COSY, HSQC, HMBC) for compound 1b (Me ester). 600 MHz, [2H6]benzene, 303 K. Position number

13 C chemical shifts (ppm); functional group

1 2 3 4 5 6 7 8a 8b 9

173.37; COOMe 34.25; CH2 25.35; CH2 29.51; CH2 29.51; CH2 29.87–29.50; CH2 30.32–29.80; CH2 32.03; CH2 32.03; CH2 56.63; CH

10 11

61.70; CH 68.27; CH

12

133.69; CH

13

133.55; CH

14 15 16 17 18 (1) (11)

28.41; CH2 26.39; CH 31.83; CH 23.07; CH2 14.35; CH3 51.05; COOMe OH

1

H chemical shifts (ppm); multiplicity; coupling constant (Hz)

2.11; t; 7.5 (H3) 1.53; m; 1.14; m 1.07–1.20; m 1.07–1.30; m 1.19–1.38; m 1.40; m 1.34; m 2.82; ddd; 6.1 (H8a); 5.1 (H8b); 2.2 (H10) 2.71; dd; 5.1 (H11); 2.2 (H9) 4.25 dddd; 8.5 (H12); 5.3 (OH); 5.1 (H10); 1.1 (H13) 5.53; ddt; 11.0 (H13); 8.5 (H11); 1.4 (H14) 5.45; ddt; 11.0 (H12); 7.4 (H14); 1.1 (H11) 1.97; m 1.25; m 1.15–1.24; m 1.23; m 0.87; t; 7.2 (H17) 3.37; s 1.54; d; 5.3 (H11)

Heteronuclear multiple bond correlation COOMe, H2 H3, COOMe H2, H4 H3

H8a, H8b H7, H9 H7, H9 H8a, H8b, H10 H9, H11 H10, H12 H11, H13

Table 3 GC-MS (SIM) quantification of 18O incorporation from [18O2-hydroperoxy]9-HPOD, 18O2 atmosphere and [18O2]water into compound 1 in the presence of EsEAS (CYP5164B1). A. 18O incorporation from [18O2-hydroperoxy]9-HPOD into compound 1 species possessing 2, 1 or 0 atoms of 18O Ion, m/z

Relative abundances⁎ of compound 1 species possessing 2, 1 or 0 atoms of 18O, %

387.3 385.3 383.3

54.60 ± 0.40⁎⁎ 37.34 ± 0.13⁎⁎ 8.06 ± 0.53

B. The total isotopic content of 18O in compound 1 and the precursory [18O2]9-HPOD Total estimated 18O content in product 1 (100% corresponds to two 18O atoms incorporated)

Control, 18O content in [18O2-hydroperoxy]9-HPOD

73.27 ± 0.46

84.95 ± 0.52%⁎⁎

C. Incorporation of 18O into the hydroxyl group (C11) of compound 1 after EsEAS (CYP5164B1) incubations with [18O2]9-HPOD, unlabelled 9-HPOD under the 18O2 atmosphere, or unlabelled 9-HPOD in [18O2]water. 18

O labelled precursor

H12, H14 H13, H15 H14, H16 H15 H18 H17 H2 H11

to the novel enzyme and the name EsEAS to the corresponding gene. The described CYP5164B1 is a first CYP74-related enzyme from brown algae and a second P450 possessing the EAS activity. The only example of EAS described before is the CYP74 clan enzyme CYP440A1 (BfEAS) of the lancelet Branchiostoma floridae [4]. The catalytic mechanisms of EsEAS and the lancelet enzyme are similar. Both produce epoxyalcohols. EsEAS synthesizes mainly the (9S,10S,11S)-epimer of the epoxyalcohol (trans-epoxide) while the BfEAS yields the (S,R,S)-stereoisomers with cis-epoxide [4]. The specificity of 18O incorporation into epoxyalcohol by BfEAS was not reported yet. As for EsEAS, it largely incorporates both 18 O atoms from [18O2-hydroperoxy]9-HPOD into the epoxyalcohol. Thus, the EsEAS synthesizes the epoxyalcohol mainly through the isomerization of hydroperoxide. Incubations of EsEAS with the unlabelled hydroperoxide under the 18O2 gas or in [18O]water medium have shown that the rest of hydroxyl group oxygen comes from the atmosphere, and a little part is incorporated from the water (Supplementary Fig. 10). The proposed major mechanism of EsEAS catalysis is presented in Fig. 6. The 18O labelling data indicate that the enzymatic reaction goes through the (1) homolysis of hydroperoxyl function; (2) rearrangement of the resulting oxy radical into the epoxyallylic radical; (3) recombination of the epoxyallylic radical with hydroxyl radical, resulting in the epoxyalcohol formation (Fig. 6). The mechanism is consistent with those proposed before for CYP74 enzymes [4,18,23].

173

[18O2]9-HPOD 18 O2 atmosphere [18O2]water

Isotopic content of 18O in hydroxyl group at C11, compound 1, % 62.10 ± 0.40 24.23 ± 0.66 7.45 ± 0.39

Relative abundances of compound 1 species possessing 1 and 0 atoms of 18O have been estimated from integral intensities of compound 1 peak in SIM chromatograms at m/z 201.2 and 199.2, respectively. All data were corrected in accordance with the relative abundance of peak at m/z 201 in the spectra of unlabelled product 1. All experiments were performed in triplicate. Mean values and standard deviations are presented. ⁎ Relative abundances of compound 1 species possessing 2, 1 or 0 atoms of 18O have been estimated from integral intensities of compound 1 peaks in SIM chromatograms (corresponding to ions at m/z 387.3, 385.3 and 383.3) of products of [18O2-hydroperoxy]9-HPOD incubations with the EsEAS (CYP5164B1). ⁎⁎ Presented quantification data for ions (m/z 387.3 and 385.3) were corrected in accordance with the relative abundances of the same ions in the spectrum of unlabelled product. ⁎⁎ [18O2]9-HPOD was successively reduced with sodium borohydride, methylated with diazomethane and trimethylsilylated. The resulting [18O]9-HOD was subjected to SIM GC-MS analyses. 18O content was estimated from the relative abundance of ion pairs 384.3 and 382.3, [M]+; 313.2 and 311.2 [M-C5H11]+; 227.2 and 225.2 [M-(CH2)8COOMe]+.

One more kind of CYP74 enzyme besides the EAS catalyzes the isomerization of fatty acid hydroperoxides. This is the HPL, which is widespread in higher plants [1,2]. Both atoms of 18O from 18O2hydroperoxides are incorporated into the hemiacetal, the short-lived primary product of HPLs [18,23]. Two more CYP74 enzymes, namely the AOS and DES, are dehydrases [1,2]. Thus, unlike in EAS and HPLs, the fate of the oxygens cannot be followed in AOS and DES. Oxiranyl carbinols like compound 1 are known to be formed along with other products through the non-enzymatic conversions of fatty acid hydroperoxides, occurring in the presence of acids [24,25], Fe3+ ions [26], hematin [27]. Epoxyalcohol production from hydroperoxides in the presence of monooxygenases [28] have been observed, but at

Fig. 5. The nuclear Overhauser effect (NOE) correlations in NOESY spectra of products 1a and 1b. R = n-pentyl; R’ = MeOOC(CH2)7–.

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Fig. 6. The proposed reaction mechanism of the EsEAS (CYP5164B1). R = n-pentyl; R’ = MeOOC(CH2)7–.

low yield and much slower rate compared to the EsEAS. Epoxyalcohols have also been detected as the minor side products of some CYP74 enzymes, in particular, the AOSs [29,30], as well as some nonclassical P450 AOSs of fungi [31,32]. The CYP5164B1 is evidently phylogenetically related to the CYP74 clan, most of all to the genes of methylobacteria and Metazoa (see the phylogenetic tree in Fig. 2). Nine amino acid insertion at the haembinding domain (Fig. 1), as well as the replacements in the IHCD, are also the distinctive common peculiarities of EsEAS and CYP74 clan enzymes [2]. The main mechanistic features (the involvement of the epoxyallylic radical intermediate) are common as well. Altogether, this allowed us to classify the new protein as a CYP74-related enzyme. To our knowledge, this is the first characterized recombinant enzyme of oxylipin biosynthesis from brown algae. Brown algae species possess various polyenoic C18 and C20 fatty acids [33,34]. Major constituents of E. siliculosus are linoleic, α-linolenic, arachidonic, and eicosapentaenoic acids [35]. At least three different lipoxygenase genes are present in E. siliculosus genome. However, the specificity of action of E. siliculosus lipoxygenases has not been studied yet. Metabolomic analysis of E. siliculosus revealed 9- and 13hydro(pero)xides of linoleic and α-linolenic acids [36]. These hydroperoxides, as well as 15-HPEPE, were studied in the present work. Our data revealed that 9-HPOD is the preferred EsEAS substrate among studied hydroperoxides. Thus, it is quite likely that the pathway mediated by 9-lipoxygenase and EsEAS might occur in E. siliculosus thallus. Various C18 and C20 epoxyalcohols have been detected in red [11,37, 38] and green [39] algae, as well as in the diatoms [40]. To our knowledge, no epoxyalcohols have been identified in brown algae yet. On the other hand, our unpublished observation revealed the prominent 11hydroxy-12,13-epoxy-9-octadecenoic acid in Sargassum pallidum. Nonetheless, brown algae possess a great diversity of oxylipins including the divinyl ethers (in some species) [10], as well as the different 1,2epoxycyclopentane and cyclopropane derivatives [41–43]. All these oxylipins are biosynthesized through the intramolecular conversions of the epoxyallylic radical. Thus, their biosynthesis is related to the epoxyalcohol formation. Ritter et al. [36] reported the detection of diverse oxylipins produced via the 9-lipoxygenase and 13-lipoxygenase pathways in the copper-stressed E. siliculosus tissues. Detection of the CYP74-related enzyme in brown alga raises new questions on the genesis of CYP74 clan genes. EsEAS is phylogenetically distant from CYP74s of other taxa. Thus, the filling of many existing gaps in the genomic knowledge is required for better understanding the relations between the EsEAS and CYP74s of bacteria, metazoan, and green plants. 5. Conclusions 1. The full-length coding sequence of E. siliculosus CYP5164B1 enzyme has been expressed in Escherichia coli cells.

2. The recombinant CYP5164B1 converted 9- and 13-hydroperoxides of linoleic acid into the epoxyalcohols, e.g. (9S,10S,11S,12Z)-9,10epoxy-11-hydroxy-12-octadecenoic acid. Thus, the enzyme possessed the activity of epoxyalcohol synthase (EAS). 3. Both 18O atoms largely incorporated into epoxyalcohol after the incubation of CYP5164B1 (EsEAS) with [18O2-hydroperoxy]9-HPOD, thus pointing the rearrangement as a primary mechanism of conversion. 4. The described EsEAS (CYP5164B1) is a first CYP74-related enzyme of oxylipin biosynthesis in brown algae. Transparency document The Transparency document associated with this article can be found, in the online version. Acknowledgements Experiments on the expression and purification of recombinant protein were supported by grant 16-34-00648-mol_a from the Russian Foundation for Basic Research (RFBR). Experiments on purification of reaction products by HPLC were supported by grant 14-04-01532-a from the RFBR. Experiments on GC-MS analyses of reaction products of the CYP5164B1 were supported by grant MK-6529.2015.4. Experiments on NMR analyses of reaction products of CYP5164B1 were supported by grant 15-04-04108-а from the RFBR. Studies of catalytic mechanism of the CYP5164B1 enzyme were carried out under financing from the Russian Science Foundation (Project No. 16-14-10286). The authors thank Dr. Fakhima Mukhitova for GC-MS records.

Appendix A. Supplementary data Supplementary data to this article can be found online at http://dx. doi.org/10.1016/j.bbalip.2016.11.007. References [1] A.N. Grechkin, Recent developments in biochemistry of the plant lipoxygenase pathway, Prog. Lipid Res. 37 (1998) 317–352. [2] A.R. Brash, Mechanistic aspects of CYP74 allene oxide synthases and related cytochrome P450 enzymes, Phytochemistry 70 (2009) 1522–1531. [3] R.K. Hughes, S. De Domenico, A. Santino, Plant cytochrome CYP74 family: biochemical features, endocellular localisation, activation mechanism in plant defence and improvements for industrial applications, Chembiochem 10 (2009) 1122–1133. [4] D.-S. Lee, P. Nioche, M. Hamberg, C.S. Raman, Structural insights into the evolutionary paths of oxylipin biosynthesis enzymes, Nature 455 (2008) 363–370. [5] D.R. Nelson, Cytochrome P450 nomenclature, 2004, Methods Mol. Biol. 320 (2006) 1–10. [6] W.H. Gerwick, M.A. Roberts, A. Vulpanovici, D.L. Ballantine, Biogenesis and biological function of marine algal oxylipins, Adv. Exp. Med. Biol. 447 (1999) 211–218. [7] W.H. Gerwick, I.P. Singh, Structural diversity of marine oxylipins, in: T.M. Kuo, H. Gardner (Eds.), Lipid Biotechnology, CRC Press 2002, pp. 277–308.

Y.Y. Toporkova et al. / Biochimica et Biophysica Acta 1862 (2017) 167–175 [8] A. Andreou, F. Brodhun, I. Feussner, Biosynthesis of oxylipins in non-mammals, Prog. Lipid Res. 48 (2009) 148–170. [9] M. Barbosa, P. Valentão, P.B. Andrade, Biologically active oxylipins from enzymatic and nonenzymatic routes in macroalgae, Mar. Drugs 14 (1) (2016) 23. [10] P.J. Proteau, W.H. Gerwick, Divinyl ethers and hydroxy fatty acids from three species of Laminaria (brown algae), Lipids 28 (1993) 783–787. [11] Z.D. Jiang, W.H. Gerwick, Novel oxylipins from the temperate red alga Polyneura latissima: evidence for an arachidonate 9(S)-lipoxygenase, Lipids 32 (1997) 231–235. [12] A.N. Grechkin, Hydroperoxide lyase and divinyl ether synthase, Prostaglandins Other Lipid Mediat. 68-69 (2002) 457–470. [13] T. Koeduka, K. Ishizaki, C.M. Mwenda, K. Hori, Y. Sasaki-Sekimoto, H. Ohta, T. Kohchi, K. Matsui, Biochemical characterization of allene oxide synthases from the liverwort Marchantia polymorpha and green microalgae Klebsormidium flaccidum provides insight into the evolutionary divergence of the plant CYP74 family, Planta 242 (2015) 1175–1186. [14] J.M. Cock, L. Sterck, P. Rouze, D. Scornet, A.E. Allen, G. Amoutzias, V. Anthouard, F. Artiguenave, J.M. Aury, J.H. Badger, B. Beszteri, K. Billiau, E. Bonnet, J.H. Bothwell, C. Bowler, C. Boyen, C. Brownlee, C.J. Carrano, B. Charrier, G.Y. Cho, S.M. Coelho, J. Collen, E. Corre, C. Da Silva, L. Delage, N. Delaroque, S.M. Dittami, S. Doulbeau, M. Elias, G. Farnham, C.M. Gachon, B. Gschloessl, S. Heesch, K. Jabbari, C. Jubin, H. Kawai, K. Kimura, B. Kloareg, F.C. Kupper, D. Lang, A. Le Bail, C. Leblanc, P. Lerouge, M. Lohr, P.J. Lopez, C. Martens, F. Maumus, G. Michel, D. Miranda-Saavedra, J. Morales, H. Moreau, T. Motomura, C. Nagasato, C.A. Napoli, D.R. Nelson, P. NyvallCollen, A.F. Peters, C. Pommier, P. Potin, J. Poulain, H. Quesneville, B. Read, S.A. Rensing, A. Ritter, S. Rousvoal, M. Samanta, G. Samson, D.C. Schroeder, B. Segurens, M. Strittmatter, T. Tonon, J.W. Tregear, K. Valentin, P. von Dassow, T. Yamagishi, Y. van de Peer, P. Wincker, The Ectocarpus genome and the independent evolution of multicellularity in brown algae, Nature 465 (2010) 617–621. [15] R.A. Wilson, H.W. Gardner, N.P. Keller, Cultivar-dependent expression of a maize lipoxygenase responsive to seed infesting fungi, Mol. Plant Microbe Interact. 14 (2001) 980–987. [16] I.R. Chechetkin, E.V. Osipova, N.B. Tarasova, F.K. Mukhitova, M. Hamberg, Y.V. Gogolev, A.N. Grechkin, Specificity of oxidation of linoleic acid homologs by plant lipoxygenases, Biochemistry (Moscow) 74 (2009) 855–861. [17] J.B. Schenkman, I. Jansson, Spectral analyses of cytochromes P450, Methods Mol. Biol. 320 (2006) 11–18. [18] A.N. Grechkin, F. Bruhlmann, L.S. Mukhtarova, Y.V. Gogolev, M. Hamberg, Hydroperoxide lyases (CYP74C and CYP74B) catalyze the homolytic isomerization of fatty acid hydroperoxides into hemiacetals, Biochim. Biophys. Acta 1761 (2006) 1419–1428. [19] S.M. Dittami, D. Scornet, J.L. Petit, B. Ségurens, C. Da Silva, E. Corre, M. Dondrup, K.H. Glatting, R. König, L. Sterck, P. Rouzé, Global expression analysis of the brown alga Ectocarpus siliculosus (Phaeophyceae) reveals large-scale reprogramming of the transcriptome in response to abiotic stress, Genome Biol. 10 (2009) R66. [20] J. Mercier, B. Agoh, Behavior of long-chain allylic hydroperoxides in presence of complexes of certain transition-metals. 1. Structure of epoxy alcohols formed from hydroxyperoxides of methyl-10-undeceneoate in presence of vanadyl acetylacetonate, Chem. Phys. Lipids 12 (1974) 232–238. [21] C.P. Thomas, W.E. Boeglin, Y. Garcia-Diaz, V.B. O'Donnell, A.R. Brash, Steric analysis of epoxyalcohol and trihydroxy derivatives of 9-hydroperoxy-linoleic acid from hematin and enzymatic synthesis, Chem. Phys. Lipids 167 (2013) 21–32. [23] A.N. Grechkin, M. Hamberg, The “heterolytic hydroperoxide lyase” is an isomerase producing a short-lived fatty acid hemiacetal, Biochim. Biophys. Acta 1636 (2004) 47–58.

175

[24] H.W. Gardner, D. Weisleder, E.C. Nelson, Acid catalysis of a linoleic acid hydroperoxide: Formation of epoxides by an intramolecular cyclization of the hydroperoxide group, J. Org. Chem. 49 (1984) 508–515. [25] H.W. Gardner, E.C. Nelson, L.W. Tjarks, R.E. England, Acid-catalyzed transformation of 13(S)-hydroperoxylinoleic acid into epoxyhydroxyoctadecenoic and trihydroxyoctadecenoic acids, Chem. Phys. Lipids 35 (1984) 87–101. [26] H.W. Gardner, Decomposition of linoleic acid hydroperoxides. Enzymic reactions compared with nonenzymic, J. Agric. Food Chem. 23 (1975) 129–136. [27] T.A. Dix, L.J. Marnett, Conversion of linoleic acid hydroperoxide to hydroxy, keto, epoxyhydroxy, and trihydroxy fatty acids by hematin, J. Biol. Chem. 260 (1985) 5351–5357. [28] M.S. Chang, W.E. Boeglin, F.P. Guengerich, A.R. Brash, Cytochrome P450-dependent transformations of 15R-and 15S-hydroperoxyeicosatetraenoic acids: Stereoselective formation of epoxy alcohol products, Biochemistry 35 (1996) 464–471. [29] W.C. Song, S.W. Baertschi, W.E. Boeglin, T.M. Harris, A.R. Brash, Formation of epoxyalcohols by a purified allene oxide synthase. Implications for the mechanism of allene oxide synthesis, J. Biol. Chem. 268 (1993) 6293–6298. [30] R.K. Hughes, F.K. Yousafzai, R. Ashton, I.R. Chechetkin, S.A. Fairhurst, M. Hamberg, R. Casey, Evidence for communality in the primary determinants of CYP74 catalysis and of structural similarities between CYP74 and classical mammalian P450 enzymes, Proteins 72 (2008) 1199–1211. [31] I. Hoffmann, E.H. Oliw, Discovery of a linoleate 9S-dioxygenase and an allene oxide synthase in a fusion protein of Fusarium oxysporum, J. Lipid Res. 54 (2013) 3471–3480. [32] I. Hoffmann, F. Jernerén, E.H. Oliw, Expression of fusion proteins of Aspergillus terreus reveals a novel allene oxide synthase, J. Biol. Chem. 288 (2013) 11459–11469. [33] G.R. Jamieson, E.H. Reid, The component fatty acids of some marine algal lipids, Phytochemistry 11 (1972) 1423–1432. [34] S.V. Khotimchenko, V.E. Vaskovsky, T.V. Titlyanova, Fatty acids of marine algae from the Pacific coast of North California, Botanica Marina 45 (2002) 17–22. [35] S.M. Dittami, A. Gravot, D. Renault, S. Goulitquer, A. Eggert, A. Bouchereau, C. Boyen, T. Tonon, Integrative analysis of metabolite and transcript abundance during the short-term response to saline and oxidative stress in the brown alga Ectocarpus siliculosus, Plant Cell Environ. 34 (2011) 629–642. [36] A. Ritter, S.M. Dittami, S. Goulitquer, J.A. Correa, C. Boyen, P. Potin, T. Tonon, Transcriptomic and metabolomic analysis of copper stress acclimation in Ectocarpus siliculosus highlights signaling and tolerance mechanisms in brown algae, BMC Plant Biol. 14 (2014) 116. [37] M.W. Bernart, W.H. Gerwick, Eicosanoids from the tropical red alga Murrayella periclados, Phytochemistry 36 (1994) 1233–1240. [38] W.H. Gerwick, Epoxy allylic carbocations as conceptual intermediates in the biogenesis of diverse marine oxylipins, Lipids 31 (1996) 1215–1231. [39] M.W. Bernart, G.G. Whatley, W.H. Gerwick, Unprecedented oxylipins from the marine green alga Acrosiphonia coalita, J. Nat. Prod. 56 (1993) 245–259. [40] N. Lamari, M.V. Ruggiero, G. d'Ippolito, W.H. Kooistra, A. Fontana, M. Montresor, Specificity of lipoxygenase pathways supports species delineation in the marine diatom genus Pseudo-nitzschia, PLoS One 8 (8) (2013) e73281. [41] P.J. Proteau, J.V. Rossi, W.H. Gerwick, Absolute stereochemistry of neohalicholactone from the brown alga Laminaria sinclairii, J. Nat. Prod. 57 (1994) 1717–1719. [42] K. Kousaka, N. Ogi, Y. Akazawa, M. Fujieda, Y. Yamamoto, Y. Takada, J. Kimura, Novel oxylipin metabolites from the brown alga Eisenia bicyclis, J. Nat. Prod. 66 (2003) 1318–1323. [43] H. Choi, P.J. Proteau, T. Byrum, W.H. Gerwick, Cymatherelactone and cymatherols A−C, polycyclic oxylipins from the marine brown alga Cymathere triplicate, Phytochemistry 73 (2012) 134–141.