Introduction

Oxylipins are crucial compounds in plants, playing important developmental roles and being involved in plant defense mechanisms. This family of molecules includes structurally diverse metabolites that are all produced following oxidation of unsaturated fatty acids, comprising hydroperoxides, keto acids, hydroxides, oxoacids, divinyl ethers and aldehydes (Blée 2002; Andreou et al. 2009; Mosblech et al. 2009). While free oxylipins have been studied for years and have well known functions, the roles of plant oxylipins esterified in complex lipids such as galacto- and phospholipids remain unclear (Vu et al. 2012; Griffiths 2015). In 2001, a first oxidized galactolipid was completely characterized in Arabidopsis thaliana (L.) Heynh (Stelmach et al. 2001). Since then, a wide range of esterified oxylipins have been described in the same species (Ibrahim et al. 2011; Vu et al. 2012, 2014). More recently, esterified oxylipins were described outside the Brassicaceae, raising the question as to the extent of their occurrence and their biological functions. Many studies have indicated that these molecules might play important roles in plant defense and developmental processes; however, their precise biological roles remain unclear. This review focuses on new insights concerning esterified oxylipin diversity, biosynthesis and function in plant defense mechanisms and developmental processes.

Esterified oxylipins diversity

Plant esterified oxylipins have been known to exist for many years, but have only been completely structurally characterized in the last two decades. As older analytical methods made full characterization of mixtures composed of oxidized complex lipids very complicated, only the oxylipin parts of the molecules were usually analyzed following methanolysis or otherwise liberating the free oxylipin fatty acid. Generally, the whole structure was not determined (Feussner et al. 1995; Hamberg and Hamberg 1996; Fauconnier et al. 2008). For example, in 1995, Feussner et al. demonstrated that esterified (13S)-hydroxy-(9Z,11E)-octadecadienoic acid is formed during germination of cucumber cotyledons, suggesting a role for this lipoxygenation product in plant developmental processes (Feussner et al. 1995). Esterified (15R)-hydroxylinoleic acid was also found in seeds of Avena sativa L., cv. Freja and Vital (Hamberg and Hamberg 1996). In 2008, it was shown that potato cultivars produced different profiles of esterified oxylipins upon fungal infection, including colneleic and colnelenic acids, 9-hydroxylinolenic acid (9-HOT) and 13-HOT (Fauconnier et al. 2008). Miersch et al. (2004) showed that high quantities of diverse esterified oxylipins containing HOTs, hydroxylinoleic acids (HOD), ketolinolenic acids (KOT), ketolinoleic acids (KOD) and hydroperoxylinoleic acids (HPODs) are found in tomato flower organs (Miersch et al. 2004). Finally, treatment of tomatoes with avirulent rhizobactezrium Pseudomonas putida BTP1 induces accumulation of esterified 13-HPOD, 13-hydroperoxylinolenic acid (13-HPOT), 13-HOT and 13-HOD, which are then released as free compounds upon Botrytis cinerea infection (Mariutto et al. 2014). Interestingly, all those studies highlighted esterified oxylipin ubiquity in plant species and organs.

More recently, advanced analytical methods, such as high-performance liquid chromatography (HPLC)-tandem mass spectrometry (MS/MS) (Stelmach et al. 2001; Böttcher and Weiler 2007; Glauser et al. 2008) and direct infusion MS/MS (Buseman et al. 2006; Vu et al. 2012), have made analyzing complex lipid mixtures less problematic (Göbel and Feussner 2009). Those methods facilitate the structural characterization of complex lipids; however, they cannot separate closely related positional isomers and fully determine their stereochemistry. Even so, many esterified oxylipins have been described, allowing a better understanding of their biosynthetic pathways and functions. While these molecules seem to be highly diverse, their precise biological roles still remain unclear.

Oxidized glycolipids

Arabidopsides

The first esterified oxylipin, sn1-O-(9S,13S-12-oxophytodienoyl)-sn2-O-(7Z,10Z,13Z-hexadecatrienoyl)-sn3-O-(β-D-galactopyranosyl)-glycerol, was completely characterized in A. thaliana by Stelmach et al. (2001), using nuclear magnetic resonance and mass spectrometry. Thereafter, other galactolipids containing 12-oxo-phytodienoic acid (OPDA) and 12-dinor-oxo-phytodienoic acid (dnOPDA) were discovered in the same species and collectively named arabidopsides (from A to G; Fig. 1). Arabidopsides were defined as oxidized monogalactosyldiacylglycerols (MGDG) or digalactosyldiacylglycerols (DGDG) produced by A. thaliana, containing at least one OPDA or dnOPDA residue (Hisamatsu et al. 2003, 2005). Arabidopsides A and C are oxidized MGDGs and DGDGs, respectively, each containing one OPDA residue esterified in the sn-1 position and one dnOPDA residue esterified in the sn-2 position (Hisamatsu et al. 2003, 2005). Arabidopsides B and D are, in turn, oxidized MGDGs and DGDGs, containing OPDA residues esterified to the sn-1 and sn-2 positions, respectively (Hisamatsu et al. 2003, 2005). Arabidopside F is a monogalactosyldiacylglyceride containing esterified dnOPDA in the sn-2 position (Shigemori et al. 2006). Interestingly, two arabidopsides, E and G, contain one additional OPDA esterified on the C6 position of the galactolipid moiety (Andersson et al. 2006; Kourtchenko et al. 2007).

Fig. 1
figure 1

Structure of arabidopsides, linolipins and phosphatidylinositol containing colneleic acid. Arabidopsides are oxidized MGDGs and DGDGs containing OPDA and/or dnOPDA. Linolipins are oxidized MGDGs and DGDGs containing at least one (ω5Z)-etherolenic acid chain. 16:0/colneleic acid-phosphatidylinositol is an oxidized phosphatidylinositol containing one colneleic acid residue. Determined asymmetric carbons absolute configurations are represented on the figure

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Arabidopsides possess several asymmetric centers whose absolute configuration has not been determined for all known arabidopside molecular species. Glycerol asymmetric carbons in arabidopsides A (sn1-O-(12-oxophytodienoyl)-sn2-O-(dinor-oxophytodienoyl)-sn3-O-(β-D-galactopyranosyl)-glycerol) and B (sn1,sn2-di-O-(12-oxophytodienoyl)- sn3-O-(β-D-galactopyranosyl)-glycerol) were determined by subsequent enzymatic hydrolysis and optical rotation comparison and reported to have a S configuration (Hisamatsu et al. 2003). In arabidopside D (sn1,sn2-di-O-(12-oxophytodienoyl)-sn3-O-(6-O-(α-D-galactopyranosyl)-β-D-galactopyranosyl)-glycerol), the same asymmetric carbon was analyzed after derivatization using chiral gas chromatography (GC) and was reported as a racemic mixture of R and S forms (Hisamatsu et al. 2005). In other arabidopsides, the absolute configuration of the glycerol moiety has not been determined yet. Otherwise, esterified OPDA and dnOPDA ((dn)OPDA) were determined to be mainly present as S,S stereoisomers. Chiral GC analysis of (dn)OPDA in arabidopsides isolated from unstressed A. thaliana reported a 10:3 stereoisomer ratio of S,S/R,R (dn)OPDA (Hisamatsu et al. 2003, 2005). It was also shown by subsequent methyl-esterification and chiral HPLC analysis that (dn)OPDAs formed after freeze-thawing in A. thaliana are only present as S,S stereoisomers (Nilsson et al. 2016).

More recently, a diverse range of similar molecules was identified in A. thaliana by direct infusion MS (Ibrahim et al. 2011; Vu et al. 2012, 2014). This, with information detailed below, shows there was much more diversity in those oxidized galactolipids than had been found previously. Furthermore, previously characterized arabidopsides A-D, as well as four novel oxidized galactolipids, were identified in wounded A. thaliana leaves (Glauser et al. 2008). Those metabolites were described as monogalactosylmonoacylglycerols (MGMG) and digalactosylmonoacylglycerols (DGMG) containing (dn)OPDA esterified in the sn-1 or in the sn-2 position, namely sn2-O-(dinor-oxophytodienoyl)-monogalactosyl monoglyceride, sn1-O-(12-oxophytodienoyl)-monogalactosyl monoglyceride, sn2-O-(dinor-oxophytodienoyl)-digalactosyl monoglyceride and sn1-O-(12-oxophytodienoyl)-digalactosyl monoglyceride (Glauser et al. 2008).

While arabidopsides were firstly discovered in A. thaliana leaves, those molecules have also been found in flowers and stems of unstressed plants (Zhao et al. 2013); however, they were not detected in roots, even after wounding (Grebner et al. 2013), or in seeds (Wang et al. 2017). Whereas arabidopsides are constitutively produced in low quantities by A. thaliana, their levels peak under different biotic and abiotic stress conditions, such as mechanical wounding (Buseman et al. 2006; Kourtchenko et al. 2007), exposure to Pseudomonas syringae avirulence proteins AvrRpm1 and AvrRpt2 (Andersson et al. 2006; Kourtchenko et al. 2007), P. syringae infection (Vu et al. 2012) and low temperature treatment (Vu et al. 2012). For example, production of arabidopsides A, B and D increases up to 1000-fold within 15 min of mechanical wounding, with concentrations of more than 2,5 nmol/mg DW for arabidopside A (Buseman et al. 2006). Upon wounding, arabidopsides not only accumulate in A. thaliana stressed leaves but, to a minor extent, also in distal unwounded leaves of the same plant (Glauser et al. 2009). Moreover, in A. thaliana the esterified oxylipin profile depends on the nature of the stress; for example, wounding induces much more oxidized MGDG production than low temperature treatment (Vu et al. 2012).

Interestingly, cell disruption in A. thaliana leaves by freeze-thawing leads to the formation of arabidopsides A, B, C, D, E and G (Nilsson et al. 2012), whereas it is known not to generate free jasmonic acid (JA) (Glauser et al. 2009). Among oxidized MGDG species, arabidopsides A and B are produced first and peak approximatively 5 min after freeze-thawing. Those molecules slowly decrease thereafter, while arabidopside E and G production increases and peaks approximately 30 min after freeze-thawing (Nilsson et al. 2012, 2016).

With regard to other species, arabidopsides A and B have been detected upon wounding of Arabidopsis arenosa (L.) Lawalrée (Böttcher and Weiler 2007). The same year, these molecules were also reported in two other Arabidopsis species and three other species of the Brassicaceae (Böttcher 2007), confirming that they are not limited to the genus Arabidopsis. This conclusion was supported by the discovery in 2015 of OPDA-containing lipids in unstressed fresh leaves and freeze-thawed leaves of three other Arabidopsis species and one other species from the Brassicaceae (Nilsson et al. 2015). In 2017, OPDA and arabidopsides A, C and D were reported to be induced in another Brassicaceae plant, Erucastrum canariense Webb & Berthel, upon copper chloride spraying, and by mechanical wounding (Pedras and To 2017). As arabidopsides A and D isolated from E. canariense possessed different optical rotations to those that had been first isolated (Hisamatsu et al. 2003, 2005), it was suggested that they were stereoisomers of A. thaliana arabidopsides (Pedras and To 2017). Arabidopsides A and D were also detected in unstressed leaves of the Brassicaceae plant, Nasturtium officinale R. Br., and in leaves sprayed with copper chloride (Pedras and To 2017).

Whereas (dn)OPDA containing galactolipids where firstly described in Brassicaceae plants, a study found an OPDA-containing MGMG in Ipomoea tricolor Cav. (Convolvulaceae) (Ohashi et al. 2005). In the same way, arabidopsides were also detected in Melissa officinalis L. (Lamiaceae) (Zábranská et al. 2012). Thereafter, it was demonstrated that Cirsium arverse (L.) Scop. (Asteraceae) also produces oxidized glycolipids upon infection by Chaetomium cochlioides, an endophytic fungi, since arabidopsides and other esters of MGDG and DGDG containing OPDA, dihydroOPDA and dihydroJA were detected (Hartley et al. 2015). While arabidopsides could not be detected in several tested plant species (Nilsson et al. 2015), the discovery in other non-Brassicaceae plants indicates they are more ubiquitous than thought upon first discovery (Table 1).

Table 1 Plant species producing (dn)OPDA-containing galactolipids
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Linolipins

In 2009, the discovery of a second family of oxidized galactolipids was reported in leaves of Linum usitatissimum L., cv. Novotorzhski (Fig. 1). Those molecules were named linolipins A and B (Chechetkin et al. 2009). At the present time, four linolipins have been characterized. Linolipins are MGDGs and DGDGs, containing at least one divinyl ether residue and more precisely a (ω5Z)-etherolenic acid chain (Chechetkin et al. 2009, 2013). Linolipin A is an oxidized MGDG, containing one α-linolenic acid (LA) chain in the sn-1 position and one (ω5Z)-etherolenic acid residue esterified in the sn-2 position (Chechetkin et al. 2009). Linolipin C is an oxidized DGDG, containing one (ω5Z)-etherolenic acid esterify in the sn-1 position and one LA chain in the sn-2 position (Chechetkin et al. 2013). Linolipins B and D are respectively an oxidized MGDG and DGDG, both containing two (ω5Z)-etherolenic acid residues esterified in the sn-1 and sn-2 positions (Chechetkin et al. 2009, 2013). Glycerol asymmetric carbon configurations in all described linolipins remain unknown. While linolipin A is constitutively produced by old flax leaves, levels of linolipins B, C and D dramatically increased after bacterial infection (Chechetkin et al. 2009, 2013).

Other oxidized glycolipids

Among other oxidized glycolipids, galactolipids containing esterified hydroperoxides, ketols, hydroxides and phytoprostanes were found in mechanically wounded A. thaliana (Ibrahim et al. 2011; Vu et al. 2012; Zhao et al. 2013). In 2013, MGDGs containing esterified traumatin, dinortraumatin and its derivatives were also found to be produced by wounded leaves of A. thaliana, tobacco, cabbage and common bean (Nakashima et al. 2013).

Other esterified oxylipins

Whereas oxidized glycolipids were increasingly characterized as mentioned above, other esterified oxylipins have been reported in different plant species. In 2003, it was shown that potato tubers contain colneleic acid esterified in phospholipids (Fig. 1) (Fauconnier et al. 2003). In addition, Buseman et al. (2006) reported that wounded A. thaliana leaves not only produce oxidized glycolipids, such as arabidopsides, but also generate OPDA and ketols esterified in phosphatidylglycerol (PG). Vu et al. (2012) highlighted the diversity in oxidized phospholipids produced under stress, with (dn)OPDA, 9-HPOT and 9-HPOD being found esterified in phosphatidylethanolamines, PG and phosphatidylcholines. Nilsson et al. (2014) confirmed those observations and demonstrated that the hypersensitive response in A. thaliana triggers production in leaves of (dn)OPDA containing lipids, such as sulfoquinovosyl diacylglycerol, phosphatidylinositol (PI) and head group acylated PG.

Oxylipin biosynthesis pathways

As mentioned previously, oxylipins are structurally diverse compounds produced following enzymatic or non-enzymatic oxidation of polyunsaturated fatty acids. While the present section mainly aims to detail plant esterified oxylipin synthesis pathways, biosynthesis of free oxylipins is described below. For more details, plant free oxylipin biosynthesis pathways are reviewed in Creelman and Mulpuri (2002), Mosblech et al. (2009), Acosta and Farmer (2010), Wasternack and Hause (2013), Griffiths (2015), Ahmad et al. (2016) and Wasternack and Song (2016).

Free oxylipins biosynthesis

Free oxylipins biosynthesis (Fig. 2) is initiated by the action of lipases that hydrolyze fatty acids from membrane lipids such as phospholipids, MGDGs and DGDGs. Among them, linoleic acid (LEA), LA and hexadecatrienoic acid (HTA) are plant substrates for lipoxygenases (LOX) that catalyze oxygen insertion into the fatty acid chain. Those unsaturated fatty acids can also be oxidized by α-dioxygenases (DOX) (Hamberg et al. 1999; Stumpe and Feussner 2006; Griffiths 2015; Wang et al. 2018).

Fig. 2
figure 2

Biosynthesis pathways of plant free and esterified oxylipins. Following biotic and abiotic stress, free oxylipins are hydrolyzed from membrane complex lipids by lipases. Free unsaturated fatty acids can then be oxidized by diverse enzymes into a wide range of metabolites. Among those, 13- hydroperoxylinolenic acid (13-HPOT) and 11-hydroperoxyhexadecatrienoic acid (11-HPHT) can be submitted to the subsequent action of allene oxide synthase (AOS) and allene oxide cyclase (AOC) and be transformed into 12-oxo-phytodienoic acid (OPDA) and 12-dinor-oxo-phytodienoic acid (dnOPDA), respectively. Two pathways of jasmonic acid biosynthesis from OPDA/dnOPDA are shown. OPDA/dnOPDA can be reduced by OPDA reductase 3 (OPR3) and then undergo three/two rounds of β-oxidation (in black). An alternative pathway (in red) was recently described by (Chini et al. 2018). In opr3-3 plants, OPDA/dnOPDA directly undergo three/two rounds of β-oxidation and are then reduced to jasmonic acid by OPDA reductase 2 (OPR2). Following biotic and abiotic stress, membrane galactolipids can also be directly oxidized by lipoxygenase 2 (LOX2) into esterified HPOT and HPHT. Subsequent action of AOS and AOC on those products leads to non-acylated arabidopsides formation. Acylated galactolipid associated phospholipase 1 (AGAP1) can then catalyze the transfer of (dn)OPDA from galactolipids to the galactose moiety of other arabidopsides, forming acylated arabidopsides. Other esterified oxylipins such as linolipins have also been described, but their biosynthesis remains unknown. ACX1 (acyl-coenzyme A oxidase 1), ADH (alcohol dehydrogenase), DES (divinyl ether synthase), DOX (α-dioxygenase), EAS (epoxy alcohol synthase), HPL (hydroperoxide lyase), KAT (3-ketoacyl-coenzyme A oxidase), LOX (lipoxygenase), MFP (multifunctional protein), OPC-4 (4-(3-oxo-2-(pent-2-enyl)cyclopentyl)butanoic acid), OPC-6 (6-(3-oxo-2-(pent-2-enyl)cyclopentyl)hexanoic acid), OPC-8 (8-(3-oxo-2-(pent-2-enyl)cyclopentyl)octanoic acid), PLA1 (phospholipase A1), PLIP2 (plastid lipase 2), PLIP3 (plastid lipase 3), POX (peroxygenase), tnOPDA (tetranor-OPDA), 4,5-ddhJA (4,5-didehydro-jasmonic acid). (Color figure online)

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Enzymes 9- and 13-LOX are regioselective, catalyzing fatty acid oxidation into fatty acids hydroperoxydes, depending on the oxidation site (Griffiths 2015). Hydroperoxides 13-HPOT and 11-hydroperoxyhexadecatrienoic acid (11-HPHT) produced following 13-LOX oxidation of LA and HTA (Griffiths 2015) are substrates for the enzyme allene oxide synthase (AOS) that transforms them into unstable allene oxides. Those molecules are then spontaneously or enzymatically cyclized to form OPDA and dnOPDA, respectively. It is known that spontaneous cyclization gives a blend of OPDA-enantiomers (9R,13R and 9S,13S), while enzymatic cyclization by the allene oxide cyclase (AOC) enzyme only gives the optically pure 9S,13S OPDA stereoisomer (Creelman and Mulpuri 2002; Hofmann et al. 2006; Wasternack and Hause 2013; Nilsson et al. 2016; Bjornson et al. 2017). Two different JA synthesis pathways from (dn)OPDA have currently been described (Chini et al. 2018; Howe 2018; Wasternack and Hause 2018). OPDA and dnOPDA can be reduced by OPDA reductase 3 (OPR3) and the products formed are then transformed into JA by 3 β-oxidations or 2 β-oxidations, respectively (Creelman and Mulpuri 2002; Wasternack and Hause 2013; Ahmad et al. 2016). Surprisingly, an OPR3-independent synthesis pathway for JA has been recently discovered in A. thaliana. OPDA and dnOPDA can indeed enter the peroxisome and follow three or two rounds of β-oxidations, respectively. The product formed then leaves the peroxisome and is reduced in the cytosol by OPDA reductase 2 (OPR2) to form JA (Chini et al. 2018). JA is part of an oxylipin family called jasmonates that includes oxylipins produced enzymatically following LA and HTA acid transformation into JA and its derivatives, such as JA-isoleucine (JA-Ile). Jasmonates have been studied for years and some of them are known to play important developmental and defensive roles (Feussner et al. 1995; Wasternack and Kombrink 2010; Koo 2018). JA derivatives (Ahmad et al. 2016) and some of their precursors (Stintzi et al. 2001; Dave and Graham 2012) are able to modulate gene expression leading to important developmental and defensive changes. For example, they are involved in plant defense against insects (Vijayan et al. 1998), flower development, leaf senescence and seed germination (Ahmad et al. 2016). Biological activities of jasmonates have been extensively studied and were recently reviewed (Wasternack and Strnad 2016, 2018; Huang et al. 2017; Koo 2018; Wasternack and Feussner 2018).

Hydroperoxides 13-HPOT and 11-HPHT are also substrates for hydroperoxide lyase (HPL) that cleaves the C12-C13 bond and produces (3Z)-hexenal and (9Z)-traumatin from 13-HPOT and (3Z)-hexenal and (7Z)-dinortraumatin from 11-HPHT (Vick and Zimmerman 1987; Croft et al. 1993; Nakashima et al. 2013). Recently, it was shown that the initial products of HPL are hemiacetals, unstable compounds that are rapidly transformed into aldehydes and oxoacids (Mukhtarova et al. 2018).

The consecutive action of LOX and divinyl ether synthase (DES) on fatty acids, mainly on LEA and LA, also produces oxidized derivatives called divinyl ethers (Stumpe et al. 2008). Among these compounds, colneleic and colnelenic acids are produced from 9-hydroperoxides of LEA and LA, respectively. Etheroleic and etherolenic acids are, in turn, produced from the 13-hydroperoxides of LEA and LA, respectively (Hamberg 1998). The hydroperoxide fatty acids can also be transformed by other pathways into various other compounds, such as epoxides, epoxy alcohols, ketols, hydroxy fatty acids and aldehydes (Hamberg 1998).

Esterified oxylipins biosynthesis and subcellular localization

Two main hypotheses are currently proposed to explain the biosynthesis of esterified oxylipins. Firstly, it has been suggested that free oxylipins could be produced from free fatty acids and then subsequently be esterified into complex lipids. Alternatively, fatty acids esterified into complex lipids may be directly transformed into oxylipins (Stelmach et al. 2001; Buseman et al. 2006; Chechetkin et al. 2009, 2013). In general, LOX enzymes responsible for the first transformation step of fatty acids into oxylipins are thought to only use free fatty acids and not esterified substrates (Gardner 1991; Babenko et al. 2017). However, a few specific LOX have been reported to use esterified substrates such as phospholipids and galactolipids in vitro (Feussner et al. 1995; Nakashima et al. 2011; Meyer et al. 2013); for example, soybean LOX1 was able to oxidize MGDG (Nakashima et al. 2011).

Different studies have focused on the biosynthesis pathways of (dn)OPDA-containing esterified oxylipins in A. thaliana and these aspects will be discussed here. Unfortunately, there is still no study available on the synthesis of other esterified oxylipins, such as linolipins.

Arabidopside synthesis

As arabidopsides are oxidized MGDG and DGDG, it was firstly suggested that these compounds were produced following oxidation of plant chloroplast membrane galactolipids (Stelmach et al. 2001). Free (dn)OPDA could then be rapidly esterified into chloroplastic MGDG and DGDG, or unsaturated fatty acids in MGDG and DGDG could be directly transformed into (dn)OPDA by LOX, AOS and AOC action (Stelmach et al. 2001; Buseman et al. 2006). Notably, the second proposal was supported by the discovery in 2012 that galactolipid fatty acid chains can be directly oxidized. A. thaliana leaves incubated in 18O-labelled water before being wounded, did not lead to the formation of 18O-labelled arabidopsides. This experiment proved that esterified fatty acids were not hydrolyzed, oxidized and re-esterified. In addition, arabidopsides are quickly formed in completely disrupted cells without free fatty acid formation, which favors a production mechanism from esterified fatty acids (Nilsson et al. 2012). As described above, OPDA production starts with LOX oxygenation of fatty acids. A. thaliana possesses six LOX isoforms, four of them are 13-LOX (LOX2, LOX3, LOX4, LOX6) and two are 9-LOX (LOX1, LOX5) (Wasternack and Hause 2013). Interestingly, LOX2 is largely implicated in (dn)OPDA-containing esterified oxylipin synthesis, as production of arabidopsides A, B and C in wounded plants decreases dramatically in Arabidopsis lox2-1 mutants compared to wild type plants (Glauser et al. 2009; Zoeller et al. 2012; Nilsson et al. 2016). As the increase in HPL expression leads to less arabidopside accumulation, it was suggested than this enzyme competes with AOS for esterified fatty acid hydroperoxides (Nilsson et al. 2016). These results are consistent with the fact that A. thaliana Columbia 0 wild type, which does not express any functional HPL, produces high quantities of arabidopsides (Duan et al. 2005; Chehab et al. 2008; Nilsson et al. 2016). Finally, since (dn)OPDA esterified in galactolipids and formed after freeze-thawing in A. thaliana are only present as S,S stereoisomers, it was proposed that allene oxide cyclization does not occur spontaneously but enzymatically by AOC action, suggesting that freeze-thawing triggers the activation of LOX2, AOS and AOC that are likely already synthesized. However, A. thaliana possesses four AOC isoforms and it is not known if one of them or all of them are involved in this transformation (Nilsson et al. 2016).

MGDGs containing one oxidized chain acylated on the galactose 6′-hydroxyl group, such as arabidopsides E and G, were also reported (Andersson et al. 2006; Kourtchenko et al. 2007; Vu et al. 2014). In A. thaliana, non-acylated (dn)OPDA containing MGDG and DGDG are rapidly produced upon stress and peak after approximatively 5 min. The presence of those molecules then decreases and oxidized acyl MGDGs are produced and reach a maximum approximatively 30 min after stress (Vu et al. 2014; Nilsson et al. 2016). Actually, (dn)OPDA esterified in galactolipids is transferred to the galactose moiety of other MGDGs (Vu et al. 2014; Nilsson et al. 2015) by acylated galactolipid associated phospholipase 1 (AGAP1), forming OPDA-acylated MGDG (Nilsson et al. 2015). Mutants of A. thaliana with an AGAP1 knockout are unable to produce oxidized and non-oxidized acyl MGDGs (Nilsson et al. 2015).

Localization of arabidopsides

It has been suggested that arabidopsides are mainly localized in chloroplast membranes (Böttcher and Weiler 2007), since galactolipids are a major constituent of these membranes (Dörmann and Benning 2002). Böttcher and Weiler (2007) found high quantities of esterified (dn)OPDA in thylakoid membranes but low quantities in the chloroplast envelope membranes. AOS and AOC, the two enzymes responsible for fatty acid hydroperoxide transformation into (dn)OPDA are localized in chloroplasts, supporting this hypothesized location (Wasternack and Hause 2013). However, it has not been established if the OPDA located in chloroplasts is esterified in galactolipids. It is also not known if those molecules can be found in other plant organelles, or if they are transferred from chloroplast membranes to other plant cell organelle. In this context, it has been shown that under specific conditions, such as phosphate starvation, non-oxidized DGDG can be transferred from chloroplast membranes to non-plastidial membranes such as mitochondrial membranes (Jouhet et al. 2004) and plasma membrane (Andersson et al. 2005; Tjellström et al. 2008). As it is known that phosphate deprivation induces LOX2 expression in A. thaliana leaves (Khan et al. 2016) and that this enzyme is implicated in arabidopside synthesis (Glauser et al. 2009), this same trafficking process could occur for arabidopsides. It is known that any change in plant plasma membrane composition can affect its integrity and its wide range of functions (Deleu et al. 2014). Therefore, arabidopside trafficking to the plasma membrane could have many effects on plant physiology. Due to their particular structure, arabidopsides should have different interfacial properties compared to plant plasma membrane lipids. Arabidopside presence in this membrane might modify lipid organization and plasma membrane properties such as fluidity and permeability. A wide range of proteins are present in the plasma membrane, including some involved in signaling functions. As a result, modifications in the lipid environment of these proteins might alter their activities and so influence signaling pathways, possibly those involved in plant defense mechanisms.

Function of esterified oxylipins

Although specific roles have not been assigned to esterified oxylipins, they are thought to be involved in plant defense and developmental processes (Fig. 3).

Fig. 3
figure 3

Potential functions of esterified oxylipins in plants. Esterified oxylipins are largely produced under biotic and abiotic stress such as wounding, pathogen infection and low temperature treatment. Among them, oxidized galactolipids such as arabidopsides are likely formed in chloroplast membranes. Esterified oxylipins may have direct (red arrows) and indirect (blue arrows) functions. Potential direct functions of those molecules include modifications of chloroplast function, plant senescence, stomatal opening, inhibition of root growth, and antifungal and antibacterial activities. Esterified oxylipins may also be transferred to plasma and mitochondrial membranes, and thus modify membrane properties. Indirect functions have also been suggested for esterified oxylipins, including modulation of gene expression and inhibition of pathogen growth (pests, bacteria and fungi). Drawing by Carolina Levicek. (Color figure online)

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Esterified oxylipin involvement in plant defense and developmental processes

Many insights about esterified oxylipins, such as arabidopsides and linolipins, highlight their probable involvement in plant defense mechanisms. Firstly, these molecules are largely produced under stress conditions. For example, A. thaliana esterified oxylipins peak under a wide range of different biotic and abiotic stresses, such as wounding, freezing and pathogen infection (Buseman et al. 2006; Böttcher and Weiler 2007; Kourtchenko et al. 2007; Glauser et al. 2008, 2009; Chechetkin et al. 2009, 2013; Koo et al. 2009; Vu et al. 2012, 2015). Moreover, upon wounding, arabidopsides not only accumulate in wounded leaves but also in distal unwounded leaves of the same plant (Glauser et al. 2009). In addition, the A. thaliana esterified oxylipin profile is dependent on the nature of the stress. For example, wounding induces much more oxidized MGDG production than freezing (Vu et al. 2012). Moreover, the A. thaliana lox2-1 mutant, which is defective in arabidopside production, is more susceptible to herbivory by larvae of the insect Spodoptera littoralis than wild type plants (Glauser et al. 2009). In the case of linolipins, a recent study showed that their production in flax leaves can be induced by 24-epibrassinolide (EPB) (Fedina et al. 2017), a plant steroid hormone implicated in plant stress responses (Lee et al. 2018). Linolipin production following Pectobacterium atrosepticum infection of leaves pretreated with EPB was greatly increased compared to non-treated leaves, suggesting an hormonal regulation of esterified oxylipin production during plant bacterial infection (Fedina et al. 2017).

It has also been suggested that esterified oxylipins might play a role in plant development. As an example, linolipin profiles are different depending on the plant developmental stage. These molecules were not detected in young flax leaves, at day 23 of culture, with production starting at a later stage of development, from day 35 of culture, and reaching a concentration of 71 nmol/g FW (Chechetkin et al. 2009, 2013).

Indirect functions of esterified oxylipins

It has been suggested that esterified oxylipins might act as a pool of free oxylipins that could be released when necessary (Stelmach et al. 2001; Ohashi et al. 2005; Dave and Graham 2012). This hypothesis was supported by the discovery of two A. thaliana phospholipases, pPLAIIα and AtPLAI, able to release free OPDA from oxidized glycolipids (Yang et al. 2007, 2012). In the same way, arabidopside levels are increased in some A. thaliana mutants with reduced pPLAIIα expression (Davoine et al. 2017). (dn)OPDA containing MGMG and DGMG found in, for example, A. thaliana were proposed as by-products formed after free oxylipins hydrolysis from oxidized MGDG and DGDG (Glauser et al. 2008). As such, released (dn)OPDA could be used as a substrate for JA production, an essential compound in the regulation of plant developmental, physiological and defense mechanisms (Wasternack and Strnad 2016, 2018; Huang et al. 2017; Koo 2018; Wasternack and Feussner 2018). Free OPDA might also directly modulate gene expression, as this compound has been shown to induce JA-independent responses. OPDA is able to activate gene expression, leading to various responses independent of JA-Ile (Wasternack and Strnad 2016). OPDA is notably involved in plant fungal resistance (Stintzi et al. 2001), in drought stress responses (Savchenko et al. 2014) and in defense processes during plant–insect interactions (Schafer et al. 2011). Moreover, OPDA and abscisic acid were shown to cause inhibition of seed germination, though this effect was previously attributed to JA (Dave et al. 2011; Dave and Graham 2012). However, little is known about OPDA signaling as this molecule is not able to bind the co-receptor complex allowing JA-Ile perception (Arnold et al. 2016). Recently, OPDA conjugated with isoleucine (OPDA-Ile) was described in A. thaliana (Floková et al. 2016) and its ability to induce OPDA-specific gene expression was confirmed (Arnold et al. 2016). In vitro studies recently showed that an isoleucine conjugate of LA can be used as a substrate for OPDA-Ile formation (Uchiyama et al. 2018); however, the mechanisms behind the perception of this molecule have not been determined.

Oxidized MGDG and DGDG containing dihydroOPDA and dihydroJA were also detected in Cirsium arvense infected with a foliar endophyte, C. cochlioides. As dihydroOPDA is an intermediate of OPDA transformation into JA, it was suggested that with esterified OPDA further metabolism is possible. Moreover, as dihydroJA was detected but not free jasmonates, the authors suggested that the latter ones may have been synthesized and that endophyte infection may have cause their further sequestration (Hartley et al. 2015).

Linolipins consisting of etherolenic acid esterified into MGDG or DGDG can generate free etherolenic acid and its degradation product, (E)-hexenal, known to inhibit flax seed germination and root development (Chechetkin et al. 2009, 2013). Moreover, some divinyl ethers possess antifungal and/or antibacterial activities against plant pathogens (Prost et al. 2005), supporting the hypothesis that linolipins could act as a storage of free colnelenic acid that could be released rapidly when needed for biological activities.

Direct functions of esterified oxylipins

It has been suggested that plant esterified oxylipins are not only used as a pool of free oxylipins, but that the whole molecule per se could have a direct function. Notably, in vitro studies showed that some of them could decrease pathogen growth, such as arabidopside E with P. syringae, while MGDG and free OPDA had no significant effect at the same concentration (Andersson et al. 2006). Additionally, arabidopsides E and G both decrease growth of B. cinerea in vitro (Kourtchenko et al. 2007), and arabidopside A possesses direct antifungal properties against Alternaria brassicicola, Leptosphaeria maculans and Sclerotinia sclerotiorum (Pedras and To 2017).

Plant esterified oxylipins may also have developmental roles, as arabidopsides A, B, D and F are able to inhibit cress root growth (Hisamatsu et al. 2005; Shigemori et al. 2006). Arabidopsides may have a role in senescence as those molecules accumulate in A. thaliana mutants showing early leaf senescence (Xiao et al. 2010; Hu et al. 2018). In the same way, arabidopsides A, B, C, D, and F have a senescence promoting effect on leaves of Avena sativa L., cv. zenshin. At the same concentration, the senescence promoting effect of oxidized DGDGs is stronger than oxidized MGDGs. Oxidized DGDGs are in fact able to induce senescence in a similar way to JA, OPDA and methyl jasmonate, a well-known senescence promotor (Hisamatsu et al. 2006; Shigemori et al. 2011). In addition, an OPDA-containing MGMG isolated from I. tricolor was shown to possess stomatal opening properties in Commelina communis L. (Ohashi et al. 2005).

All these studies highlight a possible direct involvement of esterified oxylipins in plant stress responses and developmental processes. However, it should be noted that biological functions described here were investigated using purified molecules obtained from plant extracts whose chirality was not fully determined. As arabidopside D was reported to be a racemic blend of two enantiomers (Hisamatsu et al. 2005), it is possible that some arabidopsides exist in stereoisomer mixtures and that distinct enantiomers possess different biological activities. Total synthesis of arabidopsides has not been done yet, and esterified oxylipin extraction and purification remain challenging as low molecule yields are obtained. This presents an obstacle to certain biological studies such as inhibition of plant pathogen growth, study of arabidopsides and plant membrane lipid interactions and the study of biological activities of different stereoisomers.

As mentioned above, oxidized galactolipids in A. thaliana are likely formed in thylakoid membranes upon MGDG and DGDG oxidation (Böttcher and Weiler 2007). Lipids of the latter are part of photosystems I and II of the light harvesting complex II and are hence essential for plant photosynthesis and growth (Boudière et al. 2014; Kelly et al. 2016). A. thaliana mutants unable to synthesize DGDG show a reduced photosynthetic ability and an altered chloroplast morphology, suggesting that lipid integrity is essential for its correct function (Lin et al. 2016). Arabidopside production in these membranes, notably upon stress, might modify membrane properties. Since photosynthesis ability of chloroplasts containing arabidopsides has not been studied yet, it represents an exciting avenue for future investigation.

Conclusion

Plant esterified oxylipins were only reported in a restricted number of plant species. Those molecules have been mainly described in Arabidopsis species, where many galactolipids and phospholipids containing esterified (dn)OPDA have been reported. However, the number of recent publications describing esterified oxylipins in plant species and different plant organs is increasing. Notably, esterified oxylipins have not only been found in other species of the Brassicaceae but also in other families such as Lamiaceae, Asteraceae, Convolvulaceae and Linaceae. The recent literature suggests that esterified oxylipins may be widespread metabolites in plants and that those molecules may have numerous functions that have not been identified yet. Those molecules seem to play important roles in plant stress responses and developmental processes. We propose that esterified oxylipins are considered as rare metabolites because analytical methods did not allow routine analysis of these molecules up until recently. New methods, including direct infusion mass spectrometry and HPLC–MS/MS, have allowed an easy identification and quantification of a large number of these esterified oxylipins; however, these methods do not provide information about their stereochemistry. Esterified oxylipins are produced under specific conditions such as following biotic and abiotic stresses, at precise plant developmental stages and in particular plant organs. Besides these conditions, it is possible that esterified oxylipins are not detected in plants because their levels are lower than analytical detection limits. Future investigation on their biological functions should open the way to identify the importance of these metabolites in plant developmental processes and defense mechanisms.