A multifunctional lipoxygenase from Pyropia haitanensis— The cloned and functioned complex eukaryotic algae oxylipin pathway enzyme

Algal Research 12 (2015) 316–327

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A multifunctional lipoxygenase from Pyropia haitanensis— The cloned and functioned complex eukaryotic algae oxylipin pathway enzyme Hai-min Chen 1, Zhu-jun Zhu 1, Juan-Juan Chen, Rui Yang, Qi-jun Luo, Ji-lin Xu, He Shan, Xiao-Jun Yan ⁎ School of Marine Science, Ningbo University,Ningbo,Zhejiang315211,China

a r t i c l e

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Article history: Received 20 April 2015 Received in revised form 18 September 2015 Accepted 23 September 2015 Available online xxxx Keywords: Lipoxygenase Pyropia haitanensis Multifunction Oxylipin Eicosanoides

a b s t r a c t Lipoxygenases (LOXs) are critical starting biocatalysts for the synthesis of signaling compounds derived from lipid peroxidation. However, LOXs from prokaryotes or lower eukaryotes always show nonspecific and multifunctional properties. Here, an oxylipin pathway enzyme gene of eukaryotic algae, Pyropia haitanensis of Bangiaceae (Rhodophyta), named PhLOX was cloned and characterized; a phylogenetic analysis was also performed. This unique LOX isoform is a multifunctional enzyme, combined unusually high hydroperoxidelyase (HPL), lipoxygenase, and allene oxide synthase (AOS) three catalytic activities within one catalytic domain of the protein, which may explain the observed diversity of P. haitanensis oxylipins. Phylogenetic analysis indicated that red algae LOXs separated from the LOX clades of the ancestor of higher plant and animal in the early stages of evolution. The substrate flexibility, oxygenation position flexibility, and functional versatility of PhLOX represented typical properties of lower organisms. These results indicated that the origin of the oxylipin biosynthetic gene and the oxylipin biosynthetic pathway of some red algae are unique, which may shed light on the specific defense strategies of these red algae and the evolution of the oxylipin pathway as well as the compact genome of red algae. © 2015 Elsevier B.V. All rights reserved.

1. Introduction Lipid peroxidation is an oxidative process common to all biological systems. Oxylipins derived from the oxidative metabolism of polyunsaturated fatty acids (PUFAs) are known to appear in developmentally regulated processes and in response to environmental changes in both terrestrial plants and animals [1]. Marine algae live in a complex seawater environment, with sessile and intertidal properties of many marine macroalgae continuously challenged by a variety of potentially pathogenic organisms and multivariate environmental changes [2]. These algae obviously had to evolve defensive strategies to survive in such a hostile environment [3]. As a defense against both abiotic and biotic stresses, evolved oxylipin mechanisms have been found to be ubiquitous among eukaryotic algae [4]. In metabolic studies of red and brown algae, a diversity of hydroperoxides, prostaglandins, short Abbreviations: LOX, lipoxygenases; AOS, allene oxide synthase; HPL, hydroperoxide lyase; PUFA, polyunsaturated fatty acid; LA, linoleic; ALA, linolenic acid; ARA, arachidonic acid; EPA, eicosapentaenoic acid; DHA, docosahexaenoic acid; EST, expressed sequence tag; HETE, hydroxyl eicosatetraenoic acid; HEPE, hydroxyl eicosapentaenoic acid; HpODE, hydro(pero)xy octadecadienoic acid; HpOTE, hydro(pero)xy octadecatrienoic acid; HpETE, hydro(pero)xy eicosatetraenoic acid; HpEPE, hydro(pero)xy eicosapentaenoic acid; HpDHE, hydro-(pero)xy docosahexaenoic acid; EETE, epoxy eicosatrienoic acid; GSH-GPx, glutathione–glutathione peroxidase; SPME, solid phase micro-extraction; SRPBCC, START/RHOalphaC/PITP/Bet v1/CoxG/CalC. ⁎ Corresponding author. E-mail address: [email protected] (X.-J. Yan). 1 These authors contributed equally to this work.

http://dx.doi.org/10.1016/j.algal.2015.09.015 2211-9264/© 2015 Elsevier B.V. All rights reserved.

chain aldehydes and alcohols has been found [1,5]. However, until recently, very limited information has been available regarding the enzymes responsible for the biosynthesis of these compounds, with many important questions as yet unaddressed. First, what enzymes produce these complex algal oxylipins? The biosynthesis of oxylipins involves lipoxygenases (LOXs) and several enzyme members of the cytochrome P450 family, designated CYP74, which includes allene oxide synthase (AOS), hydroperoxide lyase (HPL), and divinyl ether synthase [6]. In most plant species, LOXs are encoded by gene families, comprising a number of isozymes, which differ in the substrate oxygenation position and substrate specificity [7]. However, in the lower organism, the oxylipin pathway is strikingly simple and its multifunctional properties have been increasingly reported [8–12]. In previous research, we have found the oxylipins in Pyropia haitanensis were different from that of other algae, with no hydroperoxides detected but an abundance of short chain volatiles found. There is a possibility that the LOX of P. haitanensis and those reported in lower organisms possess common properties, including substrate flexibility and function versatility. Second, what was the role of algae in the evolution of the oxylipin defense pathway? Flowering plants appear to use octadecanoic PUFAs, such as linoleic (18:2, LA) and α-linolenic acid (α-18:3, ALA) as the main precursors for jasmonic acid or other oxylipins. In invertebrate animals and mammals, oxylipins and particularly prostaglandins derive predominantly from eicosanoic PUFAs [13]. Researchers have found that both octadecanoid and eicosanoid metabolism occurred in ancient algal stages, suggesting that they have both animal as well as plant

H. Chen et al. / Algal Research 12 (2015) 316–327

properties [14]. Thus, as the initial enzyme of oxylipin pathway, how have the LOXs from red algae evolved and what are the evolutionary relationships between LOXs from plants, animals, and prokaryotes? LOXs are the initial and key enzymes in the oxylipin pathway and are ubiquitous in plants, prokaryotes, and animals [15]. Recently, data regarding LOXs from lower organisms, such as cyanobacteria, coral, and moss, have been published [1,8–10]. Nonetheless, there is little information concerning the origin and function of marine algal LOXs and of how and to what extent the oxylipin pathway is present in marine algal kingdoms. In addition, there is the question of whether these pathways evolved independently in the different kingdoms. In this study, a LOX gene from P. haitanensis (Bangiales, Rhodophytes), a typical warm, temperate zone species [16], was identified and its versatile functions and enzyme products described. The results raised interesting questions concerning this enzyme's evolutionary heritage and its relationship to the known diversity of oxylipin structures and functions. Finally, the catalytic character of this LOX variety was elaborated to illuminate its role as a defense strategy in red algae. 2. Materials and methods 2.1. Plant material A gametophyte of P. haitanensis was collected at Hepu, Xiangshan Harbor, Zhejiang Province, China (29°09′18″N, 121°54′05″W) in 2011. 2.2. DNA and RNA isolation P. haitanensis gametophyte genomic DNA and total RNA were isolated using the E.Z.N.A.™ HP Plant DNA kit (Omega Bio-Tek, Inc., Norcross, GA, USA) and Takara RNAiso Plus kit (Takara Bio Inc., Otsu, Japan), respectively. The total RNA was then retranscribed into cDNA using a Takara Primescript RT reagent kit (Takara Bio Inc.). 2.3. Isolation and sequence analysis of P. haitanensis LOX gene Retrieval and alignment of algal LOX associated sequences in NCBI and Pyropia yezoensis EST database (http://est.kazusa.or.jp/en/plant/ porphyra/EST/) were performed to design a pair of primers for amplification of the P. haitanensis LOX gene. The primers were LOXs/LOXa (5′ CTCACCCGCAAGGGAGATGG3′/5′GGACGCTGGGAAGGAGGTGA3′), and a fragment of 624 bp was obtained following a PCR procedure at 94 °C for 3 min; 94 °C for 30 s, 58 °C for 35 s, 72 °C for 40 s, 35 cycles and then 72 °C for 10 min. The technique of 3′ and 5′ RACE (Rapid Amplification of cDNA Ends) was used to obtain its complete ORF. The primers were LOX-os/LOX-is (5′GCCCTCCCGTCCACCCACGTT3′/5′TGCCCCAC TTCGCCGACACC3′) and LOX-oa/LOX-ia (5′CGAGCCCAGGAAGTCCCA CCCTT3′/5′GCCGCCGAGAAGACGTCCATCC3′) for 3′ and 5′-terminal amplification, respectively, by following the nested PCR procedure. The obtained PCR fragment was sequenced and blasted in NCBI data bank and designated as PhLOX. The phylogenetic analysis was monitored by aligning on http://clustalw.ddbj.nig.ac.jp/ with default parameters and then edited in TreeView. An LOX domain structural graphic was constructed according to N- and C-terminal domains, based on all the LOX gene sequences annotated in NCBI “Conserved domains”. The graphic architecture was based on the diagram of Whittaker's five-kingdom system. 2.4. Prokaryotic expression and purification of PhLOX in Escherichiacoli The primers, PhLOXF/PhLOXR (5′GGAATTCCATATGATGGGGAATGCG 3′, NdeI-site underlined, 5′CCCAAGCTTCTAGATGTCGATGGACAG3′, HindIII-site underlined), were combined to clone the target gene from P. haitanensis cDNA. The amplicon was inserted into pET-28a (+) and transformed into E. coli BL21 cells to express a recombinant protein. The transformant protein was induced in the presence of 0.1 mM

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isopropyl thio-β-galactoside and the cells then harvested, lysed, and centrifuged. The resulting crude enzyme supernatant was purified using a Ni-Agarose 6 ×His-Tagged Protein Purification Kit (Accuprep, Bioneer Corp., Alameda, CA, USA). The PhLOX protein purity was evaluated by SDS-PAGE electrophoresis. 2.5. Sample preparation for LC–MS and GC–MS analysis The focus here was on PhLOX catalytic activity for octadecanoids and eicosanoids (LA, α-ALA, arachidonic acid (ARA), eicosapentaenoic acid (EPA), docosahexaenoic acid (DHA), 9-hydroperoxyoctadecatrienoic acid (9-HpOTE), 12-hydroperoxyeicosatetraenoic acid (12-HpETE), and isotopically-labeled [D8]-ARA). For LC–MS sample preparation, enzyme catalytic products were produced by incubating a 100 μM substrate in 0.5 ml 50 mM Tris buffer (pH 8.0) with 0.12 mg ml−1 of pure enzyme at 20 °C and the reaction terminated at defined time points by adding 1 ml of ethyl acetate and shaking under 4 °C for 1 h to extract products. In glutathione–glutathione peroxidase (GSH-GPx) system, 2 U ml−1 GPx and 3 mM GSH were simultaneously added in the reaction solution with substrates [17]. After centrifugation at 12,000 rpm for 10 min, the supernatant organic phase was decanted. The organic solvent was evaporated and redissolved in 500 μl of methanol for LC– MS analysis. The substrate selectivity of PhLOX was determined by adding mixed substrates, at 100 μM each, to a sample of the enzyme, as above. Substrate utilization was quantified using six PUFA standards dissolved at 100 μM in 50 mM Tris buffer (pH 8.0). Calibration curves for the standards were established by peak area integration of different fatty acid standard concentrations. Quantities from utilization analyses were determined using calibration curves. Consumption ratio = [(substrate content before reaction − substrate content after reaction) / substrate content before reaction] × 100%. For GC–MS sample preparation, initial enzyme products were produced by incubation of a 100 μM substrate in 0.5 ml of 3.3 mg ml−1 of pure enzyme at 20 °C and the reaction stopped at defined time points by adjusting the pH to 12.5 with 4 M NaOH. Volatile products were extracted from the resulting mixture by incubation at 40 °C for 50 min using a solid phase micro-extraction device (Supelco Inc., Bellefonte, PA, USA). 2.6. LC–MS analysis LC–MS analysis was carried out on a Finnigan Surveyor and TSQ Quantum Access system (Thermo Fisher Scientific Inc., Pittsburgh, PA, USA). A Hypersil Gold C18 column (2.1 × 100 mm, 3 μm; Thermo Fisher Scientific Inc.) was used at 30 °C with a solvent system of acetonitrile (A)-0.2% acetic acid (B) and a flow rate of 0.2 ml min − 1 . The sample was eluted by a programmed solvent gradient of 30 to 55% A over 4 min, then to 80% A over 32 min, and finally to 100% A over 4 min. Absorption scanning from 234 nm to 280 nm were recorded. High-resolution mass spectrometry was performed on a Q Exactive hybride quadrupole-Orbitrap mass spectrometer operating in the data dependent mode to automatically switch between full scan MS and MS/MS acquisition in negative ion mode. Survey full scan MS spectra with mass range scanning from 50 to 500 was acquired in the Orbitrap with 70,000 resolution (m/z 200) after accumulation of ions to a 1 × 106 target value based on predictive AGC from previous full scan. Dynamic exclusion was set to 70 s. The maximum ion time is 250 ms. The MS/ MS parameters were set as follows: AGC target 2 × 105; maximum ion time 250 ms; isolation width 2 Da. Typical mass spectrometric conditions were: spray voltage, 2 kV; no sheath and auxiliary gas flow; heated capillary temperature, 275; normalized collision energy was set to 25%, and activation time to 20 ms [18]. 9-HpODE, 9-HpOTE, and 13-HpOTE were separated on a Waters Millennium HPLC system (Waters, Milford, MA) with a semi-preparative column (Nova-Pak®HR C18, 6 μm, 60 Å, 7.8 × 300 mm, PrepColumn).

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To analyse the absolute configuration of 8-HpETE, 2 U ml−1 GPx and 3 mM GSH were simultaneously added into the enzyme reaction system, and the reduced product containing 8-HETE was separated by HPLC. The elution condition is: solvent A, 0.2% acetic acid–water; and solvent B, acetonitrile. The gradient elution profile was as follows, 0– 5 min, from 70% A to 48%; 5–20 min, from 48% A to 20% A; 20–25 min, from 20% A to 0% A, at a flow rate of 1 ml min−1. Enantiomer composition analysis of purified oxylipins was carried out by chiral-phase HPLC on a Chiralcel OB column (25 × 0.46 cm, 5 μm particle size, Daicel Chem. Industries, Osaka, Japan) using a solvent system of hexane:isopropanol:acetic acid (98:2:0.1 v/v/v) at a flow rate of 1 ml min−1, with UV detection at 235 nm. Each isomer was identified by co-injections with the corresponding commercial specimen (9(R,S)HpODE, 9(R)-HpOTE, 9(S)-HpOTE, 13(S)-HpOTE, 8(R)-HETE, 8(S)HETE, Cayman, Ann Arbor, MI).

2.7. Analysis of fatty acids Total lipids were obtained using a modified version of Bligh and Dyer's method [19]. Briefly, 500 mg freeze-dried algae samples were added into 10 ml CHCl3/CH3OH/H2O (1/2/0.8, v/v), which were extracted by grinding at 4 °C for 5 min, followed by sonication for 5 min. The extraction mix was centrifuged at 3500 g for 15 min, and the residue was re-extracted by the described procedure one more time. The liquid solutions were combined, added with 5 ml ice water, and placed on ice for 5 min. After separation by centrifugation, the organic phase was dried on a rotavapor under 35 °C, followed by saponification in MeOH-H2O (4/1, v/v) with 5–6% KOH under an N2 atmosphere at 60 °C for 2 h. After cooling, the pH was adjusted to below 1 and successive triple extractions with hexane–chloroform yielded the total lipids. These lipids were then derivatized to their trimethylsilyl esters by treatment with bis(trimethylsilyl)trifluoroacetamide. The samples were then dried under a N2 stream, redissolved in hexane, and analyzed by QP2010 GC–MS. GC analysis was performed using a SPB-50 fused silica capillary column (30 m × 0.25 mm × 0.25 μm; Supelco Inc.). The temperature of the injector was 250 °C, helium carrier gas at 0.62 ml min− 1, and precolumn pressure at 51.6 kPa. After injection, oven temperature was held at 150 °C for 3.5 min, raised to 200 °C at 20 °C min− 1 , held for 5 min, raised from 250 °C to 280 °C at 5 °C min− 1, and held for 15 min. The mass spectrometer was operated in electron compact mode with 1 kV electron energy. Ion source and interface temperatures were 200 and 250 °C, respectively. The masses were scanned from m/z 50 to 750 [20].

2.8. Analysis of volatile compounds Measurements were carried out using a Shimadzu QP2010 GC system (Shimadzu Corp., Kyoto, Japan) fitted with a VOCOL column (60 m × 0.32 mm; film thickness, 1.8 μm; Supelco Inc.) and coupled with a QP2010 MS (Shimadzu Corp., Kyoto, Japan). After extraction, the SPME device was introduced in a GC splitless injector and maintained at 210 °C for 5 min. The column flow rate of helium carrier gas was at 1.99 ml min− 1 and precolumn pressure at 83.5 kPa. The oven temperature programmed at 35 °C for 3 min, then to 40 °C at 3 °C min− 1, held for 1 min, then to 100 °C at 5 °C min− 1, and finally to 210 °C at 10 °C min− 1, and held for 30 min. Mass spectra were obtained under electron ionization impact at 0.8 kV and data acquisition performed over an m/z range of 45–1000. Ion source and interface temperatures were at 200 and 210 °C, respectively. Analytes were identified by their retention times through comparison of their mass spectra with those recorded in the NIST 147 and Wiley 7 Spectrometry Library and those from commercially available pure standards [21].

3. Results 3.1. Protein sequences and alignments The obtained open reading frame sequence of PhLOX (GenBank Accession No. AFQ59981) was 2697 bp in length, encoding a protein of 898 amino acids with a molecular weight of 98 kDa (Figs. S1 and S2). The whole sequence showed the highest identity to PyLOX2 from P. yezoensis (Contig No. 31643, 90% identical). In addition, four similar LOXs were located, including Porphyra purpurea LOX (AAA61791), Gracilaria chilensis (G. chilensis) LOX (AEH16747, segment), Chondrus crispus (C. crispus) LOX (CHC_T00008739001), and P. yezoensis PyLOX1 (Contig. No. 22618), whose sequence identity to PhLOX was 54%, 41%, 34%, and 31%, respectively (Fig. S3a). A low sequence similarity was found in the N-terminal portion of the protein, which was directed to the SRPBCC (START/RHOalphaC/PITP/Bet v1/CoxG/CalC) superfamily (Fig. S3b). The other known LOXs possess N-terminal PLAT domains, which were absent in PhLOX. However, the C-terminal domain, consisting primarily of α-helices and harboring the enzyme's catalytic site, exhibits sequence similarity to that of other LOXs. In particular, it possesses a highly conserved, central, histidine-rich region around His-583, His-588, His-770, Asn-774, and Ile-898, which is involved in iron binding within the active site [22]. 3.2. Phylogenetic analysis of PhLOX In this study, a phylogenetic analysis of the LOX family using Cterminal conserved LOX domains was conducted to predict the evolutionary origin as well as some biochemical features of PhLOX (Fig. 1a). The phylogenetic tree separated plant, red algae, and animal enzymes and formed several subgroups within each kingdom, indicating that the three branches should have the same ancestral LOX. It was clear that LOXs from red algae formed individual groups in separate branches. The LOX from a marine bacterium, Shewanella violacea DSS12, is closely related to the red algae branch, and LOXs of prokaryotes are closely related to animals, except for this marine bacterium. However, interestingly, LOXs of brown algae appear more closely related to prokaryotes than to red algae and plants. The evolutionary origin of PhLOX was elaborated here by performing a conserved-domain search for its sequence in the known LOX protein sequences, and a LOX structural-evolution diagram was produced based on LOX domain structures (Fig. 1b), which revealed that according to N-terminal domains, LOXs could be divided into several groups. Most of the known LOXs from flowering plants and animals were characterized by an N-terminal polycystin/lipoxygenase/α-toxin (PLAT) domain, except for the AOS-LOX fusion protein of Triticum urartu (EMS58940). As shown in the diagram, the AOS–LOX fusion proteins, with an AOS-like domain at the N-terminus, pervaded the cyanobacteria, coral, green algae, wheat, and Proteus. An SRPBCC domain, belonging to the SRPBCC superfamily, was found in the Rhodophyta, in particular, Pyropia and Porphyra, and in the marine bacterium S. violacea DSS12 (YP_003557190). But two other red algae LOX genes from C. crispus and G. conferta did not show this domain. 3.3. Fatty acid composition of P. haitanensis and catalytic activity of PhLOX on fatty acid The natural content of PUFAs in P. haitanensis was determined by GC–MS. Chromatogram analysis showed the presence of a mixture of C20 unsaturated FAs, with C20:5 acids as the most prominent types, along with general levels of C20:4, C20:3, and C20:1. The percentage of two saturated FAs C16:0 and C18:0 was also high, but there was a low level of C18 PUFAs. The content of C20:5 acid was 26.85-fold higher than that of C18:3 acid (Fig. 2a). C22 and higher FAs were absent. The activity of purified recombinant full-length PhLOX was tested with several free PUFA substrates, including LA, α-ALA, ARA, EPA, and

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Fig. 1. Evolutionary history of PhLOX. (a) Phylogenetic tree of amino acid sequences of lipoxygenases (LOXs) from algae, prokaryotes, fungi, plants, and animals. Moss-mt shows that this LOX located in Physcomitrella patens mitochondrion. Superscript-numbers indicate the positional specificity on corresponding substrates. The alignment was performed without the N-terminal domain of LOXs. (b) Structural evolution diagram of lipoxygenases (LOXs) according to N- and C-terminal domains. An evolution diagram of N- and C-terminal domain constructed using all LOX protein sequences annotated in NCBI “Conserved domains.” Graphic architecture was according to diagram of Whittaker's five-kingdom system.

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3.4. Flexibility and diversity of substrate catalyzes

Fig. 2. The catalytic activity of PhLOX on different polyunsaturated fatty acids (PUFAs) substrates. (a) Identified fatty acids in P. haitanensis in this study. (b) Different substrates (linoleic acid (LA), α-linolenic acid (ALA), arachidonic acid (ARA), eicosapentaenoic acid (EPA), and docosahexaenoic acid (DHA)) catalyzed by enzyme for different times and detected by LC–MS. (c) Substrate selectivity of PhLOX on polyunsaturated fatty acid substrates.

DHA. The enzyme was active with all these substrates; among them, EPA and DHA were most rapidly converted by PhLOX (87.4 and 86.5% conversion of EPA and DHA respectively in 15 min). ARA was also rapidly consumed (83.9% in 15 min). In contrast, LA and α-ALA were converted to a lesser extent (43.8 and 39.4% respectively; Fig. 2b). At pH 8.0, the enzyme activities of PhLOX to five substrates LA, ALA, ARA, EPA, and DHA were 9.51, 19.43, 666.03, 718.34, and 378.50 μmol (substrate) s− 1 mg−1 (enzyme protein), respectively. Incubation of this recombinant LOX with a mixture of substrates indicated that ARA and EPA were the preferred substrates. In 15 min, ARA and EPA were decreased by 98.48% and 100% respectively, while only 40.97% of α-ALA were consumed (Fig. 2c).

The study on substrate showed that PhLOX was able to catalyze a broad range of substrates to produce structurally diverse oxylipins. The structure types of products were mainly divided into three kinds as mono-hydroperoxide, di-hydroperoxide, and ketol. In mono-hydroperoxide, for C18 FA substrates, for example, 9hydroperoxyoctadecadienoic acid (HpODE) was produced from LA, and 9- and 13-HpOTE were from ALA. Meanwhile, tiny amounts of monohydroperoxides detected with ARA, EPA, and DHA substrates, e.g., 8-HpETE, 8-hydroperoxyeicosapentaenoic acids (HpEPE), or 7and 10-hydroperoxydocosahexaenoic acids (HpDHE) were found. In dihydroperoxide products, such as 9,12-di-HpOTE from ALA, 9,12-diHpETE from ARA, 9,12-di-HpEPE from EPA, and 11,14-di-HpDHE and 14,17-di-HpDHE from DHA were found, with an exception that no dihydroperoxides were produced from LA. Besides, small amounts of α and γ-ketols were observed. For example, 9-hydroxy-10-oxo-αketol was derived from LA and ALA; 8-hydroxy-9-oxo-α-ketol was from ARA; 12-hydroxy-9-oxo-γ-ketol was from EPA; and 14-hydroxy11-oxo-γ-ketol was from DHA (Fig. 3). Our results revealed that PhLOX exhibited no strict positional specificity. For example, among the octadecanoid acids, the enzyme released C9 oxidized products from LA and C9, C12, and C13 products from ALA. Similarly, C8 , C 9, and C12 products were released from eicosanoid acids, and C7, C10, C11, C14, and C17 products were from DHA (Fig. 3). The accurate structure of metabolites obtained was analyzed by several methods. The exact mass of MS/MS fragments of each product was identified by high-resolution mass spectrometry (Table S1). Fig. 4 lists the LC–MS spectra and the corresponding fragmentation scheme of four detected products of α-C18:3 acid catalyzed by PhLOX, including 9-HpOTE, 13-HpOTE, 9,10-α-ketol, and 9,12-di-HpOTE. The complete structural analysis was then confirmed by the addition of stable isotopically labeled FAs. Fig. 5 presents the reaction on [D8]-ARA and subsequent demonstration that the fragments of the labeled precursor were in the deuterium-labeled products. The MS spectra data of some common oxylipins, e.g., 13-HpOTE and 9,10-ketol, were confirmed by comparison with the MS/MS spectra of commercially available standards (Figs. S4 and S5). The UV characteristics of the three kinds of products were recorded simultaneously, and UV maxima were observed at 235.9, 267.7, and 280 nm for mono-hydroperoxide, di-hydroperoxide, and ketol, respectively. The absolute configuration of several oxylipins was determined by chiral HPLC. Here, we could only determine the stereochemistry in few cases where commercial racemic mixture and pure enantiomers were available. By these means, the absolute stereochemistry of 9-HpODE, 9-HpOTE, 13-HpOTE, and 8-HpETE was established as S (Fig. 6).

3.5. HPL activity The generation of several kinds of short carbon-chain, volatile products were addressed using a GC–MS method. Fig S6 shows that PhLOXproduced FA hydroperoxides can be further cleaved to form C-5, -6, -8, and -9 aldehydes, with C-5 and -6 aldehydes from C18 PUFAs and C-8 and -9 aldehydes from C20 and C22 PUFAs. ARA gave most kinds of lyase products, including 2Z-octen-1-ol, 3Z-nonenal, 1-octen-3-ol, 2Enonenal, and 1-octen-3-one as secondary fragments, while 2Z-pentenal, and 3Z -hexenal were formed from ALA; EPA and DHA were transformed into 2E,6Z-nonadienal and 3,5Z-octadien-2-one. However, no

Fig. 3. PhLOX-catalyzed pathways for five substrates. Linoleic acid (LA), α-linolenic acid (ALA), arachidonic acid (ARA), eicosapentaenoic acid (EPA), and docosahexaenoic acid (DHA) were used as substrates in enzyme reaction and reaction products detected by LC–MS and GC–MS. Deduced reaction pathways shown in diagram. Molecules with red frame were not detected but deduced as existing intermediates based on their downstream product. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

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Fig. 4. LC–MS detection of α-linolenic acid catalysis by PhLOX. I, II, III, and IV represent four detected compound types; I and II, monohydroperoxide products, 9hydroperoxyoctadecatrienoic acid (9-HpOTE) and 13-HpOTE; III, product of AOS-like activity of PhLOX, 9,10-ketol; and IV, a dihydroperoxide product, 9,13-di-HpOTE.

hydroperoxides derived from LA were transformed to detectable amounts of lyase products. Furthermore, addition of ARA into the reaction solution remarkably decreased the substrate peak. No 12-HpETE was detected even as early as 6 s (Fig. 7a), while short chain aldehydes were observed by 6 s (Fig. 7b), indicating that intermediate 12-HpETE was transformed within a few seconds or instantly after substrate addition to yield products, such as 2-octen-1-ol and 1-octen-3-ol. These results suggested that PhLOX possesses an unusually high HPL activity for C20 substrates.

3.6. Hydroperoxy FAs as the intermediates According to current knowledge of the double dioxygenation and the forming pathway of aldehyde and ketol products in higher plants, the compounds shown above might be formed from the hydroperoxide. It can be assumed that PhLOX generates hydroperoxides from PUFAs; the former are then further catalyzed by other activities of PhLOX. In order to examine whether hydroperoxides are formed as intermediates, and to confirm that the derivatives of ketols or short chain volatiles are formed from the hydroperoxides intermediates, we used GSH–GPx system, which is known to reduce lipid hydroperoxides to the corresponding hydroxides in mammalian cells [23] and thereby block the formation of ketol and aldehydes, because that FA hydroxides are not substrates for AOS-like or lyase-like activities of PhLOX. As a result, the AOS activity was successfully halted, such that the peaks of ketol derived from ARA disappeared, and the reduction form of 8-HpETE, 8hydroxy eicosatetraenoic acid (8-HETE), appeared (Fig. 8). Strangely, under these conditions, the HPL activity was not affected; 12-HpETE was not detected, but C8 volatiles were still produced. When 12HpETE was used as the substrate, these C8 volatiles were also detected, but no 2-nonenal appeared. Furthermore, the formation pattern of dihydroperoxides was also examined using 12-HpETE or 9-HpOTE as substrate, and 9,12-diHpETE and 9,12-di-HpOTE were detected in the HPLC spectra (Fig. 9). Meanwhile, when 9-HpOTE was used as a substrate, 9,10-α-ketol production was observed, further demonstrating that the formation of ketol also needs hydroperoxide as intermediate (Fig. 9).

4. Discussion Lipid peroxidation is common to biological systems. Especially, a large variety of unique oxylipin classes have been found in marine algae [1,5]. Although several lipid peroxidation enzymes have been better characterized in flowering plants, animal, and cyanobacteria, there are few reports on the sequences and protein functions of LOX genes from eukaryotic algae. This makes the source of these abundant oxylipins in algae a puzzle. Here we cloned and characterized a complex oxylipin pathway enzyme in eukaryotic algae. Sequence alignment analysis of PhLOX revealed a highly conserved central histidine-rich region, indicating that the enzyme retained its function over time. However, the N-terminal sequence of PhLOX has a low sequence similarity to that of others. According to the diagram produced based on LOX conserved domain structure. PhLOX does not possess N-terminal PLAT domains, which function in some cases to direct proteins to membranes [22]. However, PhLOX and LOXs from Pyropia and Porphyra have an annotated “SRPBCC domain” in the corresponding region. SRPBCC superfamily usually has a deep hydrophobic ligandbinding pocket and can bind diverse ligands. Till now, the domains found in this superfamily include the steroidogenic acute regulatory protein (StAR)-related lipid transfer (START) domains of mammalian STARD1–STARD15, and the C-terminal catalytic domains of the α oxygenase subunit of Rieske-type non-heme iron aromatic ringhydroxylating oxygenases (RHOs_α_C), as well as the SRPBCC domains of phosphatidylinositol transfer proteins (PITPs). Interestingly, a marine bacterium S. violacea DSS12 also has this domain. It was thus presumable that LOXs from Pyropia and Porphyra and the marine bacterium S. violacea DSS12 may have the same ancestor. Then we constructed a phylogenetic tree using only LOX domains (excluding N-terminal domains), but the LOXs of Pyropia and this marine bacterium still clustered into one group (Fig. 1a). As Pyropia and S. violacea DSS12 might coexist in the same environments, we thus speculated that the SRPBCC domain and the Pyropia LOX portion might have been acquired from this marine bacterium by horizontal gene transfer; it is thus that the LOX acquisition in Rhodophytes might have occurred after the primary endosymbiosis, but it does not necessarily have a cyanobacterial origin. However, the other two red algae LOX genes from C. crispus and G. conferta do not have this domain. The phylogenetic tree showed that these LOXs were

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Fig. 5. Examination of reactant products from PhLOX using isotopically-labeled [D8]-arachidonic acid substrate. (a) Negative ionization mass spectra for products after incubation with recombinant PhLOX. (b) ESI mass spectrum of 1-octen-3-ol produced by PhLOX using isotopically-labeled [D8]-arachidonic acid substrate.

still assembled into one group, indicating that they have the same ancestor; however, during the LOX gene evolution from the Protoflorideae to Florideae, the SRPBCC domain might be lost in C. crispus and G. conferta. Nonetheless, as only a restricted search of the published genomes of bacteria and algae was currently possible, considerations of their origin remained speculative.

The PhLOX protein identified in the present study exerted several special catalytic properties. Most notably, PhLOX showed no substrate specificity. PhLOX was able to catalyze all the five substrates tested in this study, though with certain preference towards ARA and EPA, indicating that PhLOX possessed animal-like LOX characteristics, preferring to use C20 FA substrates. Comparison of LOX

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Fig. 6. Enantiomeric analysis of the reaction products. Insets show the respective enantiomeric separation of each product.

substrate preferences in different organisms indicated that LOXs appeared to have evolved by adapting to the PUFA composition of the corresponding organisms, and thus the preferred LOX substrates were mainly decided by an organism's PUFA composition. For example, flowering plants are predominantly composed of C 18 PUFAs, whereas ARA is widely distributed in terrestrial animal and corals [24]. Thus, in flowering plants, C18 FAs are the main precursors of oxylipins, while in animals, oxylipins are derived predominantly from C20 FAs [13]. Marine red algae are rich in various classes of eicosanoids [4]. Analysis of the FA composition of P. haitanensis revealed high concentrations of C20:4 and C20:5 acids. The presence of large amounts of ARA and EPA in this organism might have led to this observed substrate preference. The second notable property of PhLOX is that it exhibited no strict positional specificity. Multiple catalytic sites are available, such as 9-/12-/13lipoxygenase against ALA; 7-/10-/11-/14-/17-lipoxygenase against DHA. This positional nonspecificity of PhLOX is not unique, as many LOXs have been reported to convert PUFAs at several positions. Fungal and bacterial 15-LOXs convert ARA to 5-, 8-, 12-, and 15-HpETEs, while NpLOX1 converts ARA to 8-, 11-, 12-, 15-HETE [25]; LOXs from liverwort,

e.g., MpLOX1, generate 11- and 15-hydroxyeicosapentaenoic acid (HEPE) as products from EPA [26]. Thus, the positional nonspecificity of LOX appears ubiquitous in nature. Additionally, PhLOX was a unique multifunctional enzyme, based on product analysis. It has LOX-, AOS- and HPL-like activities. Primarily, it had lipoxygenase activity. Some single-position hydroperoxides were detected. Surprisingly, some dihydroperoxide were also found. Similar properties have only been reported in a few LOXs from soybean, wheat seed, rice seed, and all of them can convert ARA into a mixture of 5,15- and 8,15-diHpETE [27]. The formation mechanism of doubledioxygenation products has been described by Bild et al. [28] who studied the enzymatic processes of soybean LOX-1, which is able to catalyze 15-, 5-, and 8-LOX reactions in two steps. The initial oxygenation to form 15-HpETE is followed by its reverse orientation (“flip”) and then by 5 or 8 oxygenation to form 5,15- or 8,15-di-HpETE respectively. In this case, the catalytic reaction of PhLOX may also be performed in two steps. Here, the formation pattern of dihydroperoxides was examined using 12-HpETE or 9-HpOTE as substrate, and 9,12-di-HpETE and 9,12-di-HpOTE were detected in the HPLC spectra, which partially confirmed the present hypothesis. As LA has no double-bond between C-15 and C-16, and the hydrogen atom cannot be abstracted from the C-14 position, there is no possible dihydroperoxide formation during LA catalysis by PhLOX. Interestingly, besides lipoxygenase activity, PhLOX also has AOS-like activity, because small amounts of α and γ-ketols were observed. The presence of ketols, formed from unstable allene oxides by nonenzymatic hydrolysis, indicated that PhLOX possessed AOS activity, as allene oxide products are converted from hydroperoxides by AOS [29]. In this study, when 9-HpOTE was used as a substrate, 9,10-α-ketol production was observed, suggesting that the formation of ketol by PhLOX needs hydroperoxide as intermediate. In addition, PhLOX had HPL-like function. The generation of C-5, -6, -8, and -9 aldehydes or alcohols by PhLOX could be attributed to its HPL activity. Among these products, 1-penten-3-one, 1-octen-3-ol (a C8 volatile), and 3,5-octadien-2-one were mainly produced respectively from 13-HpOTE, 12-HpETE, and 14-HpDHE (Fig. 4). Because PhLOX preferred to catalyze C20:4 and C22:6 FAs, short chain volatiles generated by PhLOX were predominantly C8. These results are consistent with studies on other red algae too, suggesting that the most prominent theme in red algal oxylipin biosynthesis is the metabolism of C20 PUFAs via a 12-LOX activity [1], which was also observed in diatom Stephanophxis turris[30]. In the present analysis of volatile product, 12LOX activity might be the main activity of PhLOX as well, and we also noticed that PhLOX possessed an unusually high HPL activity for C20 substrates, because the volatiles were detected in 6 s of reaction, suggesting that the catalytic mechanism of this HPL-like activity of PhLOX may be specific and unique. Our previous research on the responses of P. haitanensis to agaro-oligosaccharides found that hydroperoxide generation was not detected in the presence of agaro-oligosaccharides over 0.5–3 h of incubation, but C8 volatiles significantly increased in the first 0.5 h [31]. We also observed the increased PhLOX expression following agaro-oligosaccharides stimulation, which suggesting that PhLOX may be a part of the P. haitanensis defense system. Wichard et al. reported a rapid depletion of polyunsaturated fatty acids (PUFAs) in diatoms and some macroalgae within the first minutes after cell disruption. This fatty acid depletion is directly linked with the production of PUFA-derived polyunsaturated aldehydes (PUA), which are thought to be involved in the chemical defense of algae [32,33]. Therefore, the observed high HPL activity in PhLOX may be used to defense external stresses rapidly and effectively in P. haitanensis. In addition, we found that PhLOX can convert 12-HpETE directly to C8 volatiles. Barofsky et al. also reported that diatom Thalassiosira rotula releases short chain polyunsaturated aldehydes by a unique transformation of polyunsaturated hydroperoxy fatty acids [34]. Another interesting phenomenon was observed in this study. In plant LOXs, a hydrogen atom is abstracted from a FA C-11 carbon atom. The

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Fig. 7. The time course of arachidonic acid (ARA) decrease by PhLOX catalysis. (a) The time course of decreased integrated ARA peak area analyzed by HPLC. (b) GC–MS analysis of short chain volatiles produced after ARA catalysis by PhLOX for 6 s.

resulting carbon-centered, radical intermediate undergoes rearrangement in the (n−2) and (n + 2) positions, and oxygen is inserted at C9 and C-13 in the final step [15]. Subsequently, ALA 13-HpOTE is converted into an allene oxide by AOS, whereas HPL can act on 9- or 13-HpOTE to form volatile aldehydes [35]. But in the present case, the AOS activity of PhLOX specifically acted on the (n−2) position, i.e., the 9-HpOTE, 8HpETE, and 10-HpDHE from ALA, ARA, and DHA, respectively. Meanwhile, the HPL activity of PhLOX was found to specifically catalyze the (n + 2) position, i.e., the 13-HpOTE, 12-HpETE, and 14-HpDHE from ALA, ARA, and DHA, respectively (Fig. 4). These specific products might be related to the substrate orientation in the active site pocket. Thus, PhLOX is a multifunctional enzyme. Besides hydroperoxide formation, it exhibits pronounced HPL and AOS activities. This property of PhLOX is not unique; recently, some enzymes involved in oxylipin formation have been reported to carry out similar, if not overlapping functions [8–11,36]. A typical example is a catalase-like-AOS–LOXs fusion protein from cyanobacteria and corals. In these cases, the catalase-like-AOS and LOX were encoded by two separate genes that form an operon, comprising a monomeric LOX fused with a hemecontaining AOS [8,11]. However, in the present case, there was only

one catalytic domain, one active site, belonging to a conserved nonheme LOX family, suggesting that although PhLOX catalyzed equivalent reactions, its AOS function had no relationship with the actual heme AOS protein of the CYP74 family. This is a very unique property, because, until now, all reported functional AOS enzymes belong to the hemoproteins, which thus makes PhLOX quite exceptional. In other cases, PpLOX1 from moss Physcomitrella patens[10] and a LOX from diatom T. rotula show additional HPL activity [37], and LOXs from soybean and the alga Chlorella pyrenoidosa show HPL activity under oxygen deprivation [9]. In the former enzymes, product 12-HpETE is cleaved to (2Z)-octen-1-ol and 1-octen-3-ol. Prerequisites for this activity are elevated substrate concentrations and a double bond either at the ω-6 position in the case of C18 PUFAs or at ω-5 for C20 PUFAs. In the present case, there was no substrate concentrations required, from 5 to 125 μM we tested, the products were same, but the prerequisite instead appeared to be the position of the substrates'–OOH group, such as ω-9 for C20 and C22 and ω-6 for C18:3 FAs (Fig. 4). In the other enzymes above, anaerobic LOX activity only further metabolizes 13hydroperoxides. Here, PhLOX catalyzed both the 13-hydroperoxide (produced from C18:3 acid as starting substrate) and the 12-

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Fig. 8. The PhLOX reaction products were altered by reduction of hydroperoxide intermediates with glutathione–glutathione peroxidase (GSH–GPx) system. Arachidonic acid is used as substrate. After GSH–GPx addition, hydroperoxide product 8-hydroperoxyeicosatetraenoic acid (8-HpETE) reduced to 8-hydroxy eicosatetraenoic acid (8-HETE), which then prevented conversion to 9,12-ketol.

hydroperoxide (produced from C20:4 and C20:5 acid substrates) under aerobic conditions. However, whether functional aerobic and anaerobic HPL–LOXs are evolutionarily related remained unclear. Lee et al. [38] suggested that oxylipin biosynthetic genes might have been present in the last common ancestor of plants and animals. LOXs are ancient genes, probably more ancient than CYP74. To date, only a few studies of cloned and expressed LOXs from algae have been reported [1]. Here we hypothesized that under the requirements of functional or environmental stress, Rhodophyta might have evolved LOX to execute

the functions of CYP74 family. Thus, in the present situation, the CYP74 enzymes' heme portion was substituted by the nonheme portion of LOX, to achieve the similar catalytic activity. Therefore, our hypothesis here was that the ancestral LOX might have functioned in a manner similar to some CYP74 enzymes before gene duplication and neofunctionalization occurred. Many primary red algae have compact genomes. Extremophilic red alga Galdieria sulphuraria has a 13.7-Mbp genome [39]; Florideophyte C. crispus and P. yezoensis have a nuclear genome of 105-Mbp [40] and ~ 43-Mbp [41] respectively. In their study on the genome of C. crispus, Collén et al. surprisingly found that there were only two genes encoding LOX but no candidates for AOS, HPL, and DES. The present study also searched the P. haitanensis transcriptomic information published by Xie et al. [16] and the genomic data of P. yezoensis but did not find candidate genes for these enzymes. To date, no evidence has indicated that CYP74 family existed at the evolutionary stage when Rhodophyta emerged. If these primary organisms were to function normally in some situations, they might need certain genes to involve in multiple tasks. Many primary organisms evolve a similar strategy; for example, cyanobacteria and corals choose to use a cAOS–LOX fusion protein to satisfy their demand for oxylipins. Red algae, however, might have selected a more economical and easy means for achieving this purpose, using a multifunctional monoenzyme to produce the diversity of C 18 and C 20 oxylipins and related compounds that are needed to respond to biotic stresses. 5. Conclusion

Fig. 9. Product specificity of PhLOX on hydroperoxide intermediates. (a) 9hydroperoxyoctadecatrienoic acid (9-HpOTE) as substrate. (b) 12hydroperoxyeicosatetraenoic acid (12-HpETE) as substrate.

This study discovered a unique multifunctional LOX with unusual catalytic properties from a marine red alga P. haitanensis. Pyropia LOX gene groups, along with those of other red algae, were concluded to have separated from the ancestor of higher plant and animal LOX clades in the early stages of evolution. These genes might be evolved after horizontal gene transfer from a marine bacterium. In terms of substrate and position flexibility and functional versatility, PhLOX possessed properties typical for some lower organisms, suggesting that the oxylipin pathway is an ancient pathway in organisms. Although PhLOX represented only a minor LOX isoform, its properties, along with its unique characteristic of producing abundant downstream volatiles, have implications for possible defense strategies of marine red algae in the marine ecosystem. However, the multifunctional properties of PhLOX raised several

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new questions regarding lipid-derived signals as follows: 1) How the trifunctional ability of PhLOX was evolved? Whether the multifunctional property of PhLOX was evolved by mutation of some gene positions or the PhLOX ancestor already possessed such property? 2) Whether the multifunctional property of LOX in this study is typical and specific for P. haitanensis only or it occurs ubiquitously in algae? These important questions warrant future crystallographic analyses and systemic researches on algal LOXs. Acknowledgments This project was funded by the NSFC project (No. 81370532); Ningbo Marine Algae Biotechnology Innovative Team (No. 2011B81007); National Key Technology Research and Development Program of the Ministry of Science and Technology of China (No. 2011BAD13B08); Science and Technology Major Project of the Ministry of Science and Technology of Zhejiang, China (No. 2012C12907-6); K.C. Wong Magna Fund of Ningbo University, and Zhejiang 151 Talents Project. Appendix A. Supplementary data Supplementary data to this article can be found online at http://dx. doi.org/10.1016/j.algal.2015.09.015. References [1] A. Andreou, F. Brodhun, I. Feussner, Biosynthesis of oxylipins in non-mammals, Prog. Lipid Res. 48 (2009) 148–170. [2] P. Potin, K. Bouarab, J.P. Salaün, G. Pohnert, B. Kloareg, Biotic interactions of marine algae, Curr. Opin. Plant Biol. 5 (2002) 308–317. [3] A. Cosse, C. Leblanc, P. Potin, Dynamic defense of marine macroalgae against pathogens: from early activated to gene regulated responses, Adv. Bot. Res. 46 (2007) 221–266. [4] K. Bouarab, F. Adas, E. Gaquerel, B. Kloareg, J.P. Salaün, P. Potin, The innate immunity of a marine red alga involves oxylipins from both the eicosanoid and octadecanoid pathways, Plant Physiol. 135 (2004) 1838–1848. [5] 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. [6] M. Stumpe, I. Feussner, Formation of oxylipins by CYP74 enzymes, Phytochem. Rev. 5 (2006) 347–357. [7] A.Z. Andreou, E. Hornung, S. Kunze, S. Rosahl, I. Feussner, On the substrate binding of linoleate 9-lipoxygenases, Lipids 44 (2009) 207–215. [8] M.L. Oldham, A.R. Brash, M.E. Newcomer, The structure of coral allene oxide synthase reveals a catalase adapted for metabolism of a fatty acid hydroperoxide, Proc. Natl. Acad. Sci. U. S. A. 102 (2005) 297–302. [9] A. Nuñez, T.A. Foglia, B.J. Savary, G.J. Piazza, Anaerobic lipoxygenase activity from Chlorella pyrenoidosa responsible for the cleavage of the 13-hydroperoxides of linoleic and linolenic acids, Eur. J. Lipid Sci. Technol. 102 (2000) 181–188. [10] T. Senger, T. Wichard, S. Kunze, C. Göbel, J. Lerchl, G. Pohnert, I. Feussner, A multifunctional lipoxygenase with fatty acid hydroperoxide cleaving activity from the moss physcomitrella patens, J. Biol. Chem. 280 (2005) 7588–7596. [11] B.L. Gao, W.E. Boeglin, Y.X. Zheng, C. Schneider, A.R. Brash, Evidence for an ionic intermediate in the transformation of fatty acid hydroperoxide by a catalase-related allene oxide synthase from the cyanobacterium Acaryochloris marina, J. Biol. Chem. 284 (2009) 22087–22098. [12] C. Wasternack, I. Feussner, Multifunctional enzymes in oxylipin metabolism, Chembiochem 9 (2008) 2373–2375. [13] A.R. Brash, Lipoxygenases: occurrence, functions, catalysis, and acquisition of substrate, J. Biol. Chem. 274 (1999) 23679–23682. [14] K. Bouarab, F. Adas, E. Gaquerel, B. Kloareg, J.P. Salaün, P. Potin, The innate immunity of a marine red alga involves oxylipins from both the eicosanoid and octadecanoid pathways, Plant Physiol. 135 (2004) 1838–1848. [15] A. Andreou, I. Feussner, Lipoxygenases—structure and reaction mechanism, Phytochemistry 70 (2009) 1504–1510. [16] C.T. Xie, B. Li, Y. Xu, D.H. Ji, C.S. Chen, Characterization of the global transcriptome for Pyropia haitanensis (Bangiales, Rhodophyta) and development of cSSR markers, BMC Genomics 14 (2013) 107. [17] Y. Akakabe, K. Matsui, T. Kajiwara, Stereochemical correlation between 10hydroperoxyoctadecadienoic acid and 1-octen-3-ol in Lentinula edodes and Tricholoma matsutake mushrooms, Biosci. Biotechnol. Biochem. 69 (2005) 1539–1544.

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