Ketols Serve Signaling Roles in Plant–Pathogen Interactions: Comparison
View Latest Version
Please note this is a comparison between Version 1 by Katherine Berg-Falloure and Version 3 by Peter Tang.

Plants produce an array of oxylipins implicated in defense responses against various stresses, with about 600 oxylipins identified in plants to date. Most known oxylipins are the products of lipoxygenase (LOX)-mediated oxygenation of polyunsaturated fatty acids. One of the most well-characterized oxylipins produced by plants is the hormone jasmonic acid (JA); however, the function of the vast majority of oxylipins remains a mystery. One of the lesser-studied groups of oxylipins is comprised of ketols produced by the sequential action of LOX, allene oxide synthase (AOS), followed by non-enzymatic hydrolysis. Ketols were mostly considered mere by-products of JA biosynthesis. Accumulating evidence suggests that ketols exhibit hormone-like signaling activities in the regulation of diverse physiological processes, including flowering, germination, plant–symbiont interactions, and defense against biotic and abiotic stresses. 

  • abiotic stress
  • conjugation to catecholamines
  • CYP74 enzymes
  • induced systemic resistance
  • induction of flowering
  • ketol
  • systemic acquired resistance
  • plant–insect interactions

1. The Lipoxygenase Pathway: CYP74 Enzymes

Plants produce a large number of diverse oxygenated lipids, collectively called oxylipins, that play a role in all aspects of plant physiology, including growth, development, and defense [1][2][3][4]. Lipoxygenases (LOX) oxidize polyunsaturated fatty acid (PUFA) substrates and are the first enzymatic step for the synthesis of most oxylipins [5]. In plants, the major PUFA substrates are linoleic (C18:2), linolenic (C18:3), and hexadecatrienoic acid (C16:3). Depending on the oxidized carbon position of C18:2 or C18:3, LOXs produce either 9- or 13-hydroperoxides of these fatty acids, or 11- or 7-hydroperoxides of the C16:3 substrate. Fatty acid hydroperoxides are fluxed into seven subsequent branches of the LOX pathway, which collectively synthesize an array of structurally and functionally diverse oxylipins [6]. These LOX pathway branches include allene oxide synthase (AOS), hydroperoxide lyase (HPL), divinyl ether synthase (DES), epoxyalcohol synthase (EAS), reductase (RED), peroxidase (POX), and lipoxygenase (LOX) [5]. There are a number of excellent reviews that provide detailed overviews of several of these branches and their oxylipin products [6][7][8][9][10][11][12].
This resviearchw focuses on the synthesis and functions of ketol oxylipins produced by enzymatic actions of the AOS members of the CYP74 subfamily of enzymes. AOS, HPL, EAS, and DES belong to the CYP74 subfamily of cytochrome P450 enzymes, divided into subfamilies designated as CYP74A, B, C, and D based on their specific enzymatic activity [13][14]. CYP74A displays 13-AOS or EAS activity, CYP74B contains enzymes with13-HPL activity, the members of CYP74C contain enzymes with 9/13-HPL and 9/13-AOS activity, and the CYP74D subfamily is composed of enzymes displaying 9/13-DES activity [15].
The 13-HPL pathway leads to the production of C6 volatiles, referred to as green leafy volatiles (GLVs), and C12 compounds, collectively termed traumatins. The 9-HPL pathway produces both volatile and non-volatile C9 compounds. The 9- and 13-DES pathways are responsible for the production of divinyl ethers. The 13-DES activity leads to the production of etheroleic or etherolenic acid, while the 9-DES activity produces colneleic or colnelenic acid [11]. The EAS pathway gives rise to epoxyalcohols [15], with EAS enzymes recently identified in higher plants as part of the CYP74 family [16].
In all plant studied species, the 13-AOS branch of the LOX pathway is known to produce the well-characterized plant hormones, 12-oxo-phytodienoic acid (12-OPDA) and jasmonic acid (JA), and their derivatives collectively called jasmonates. The 12-OPDA is produced by the sequential action of 13-LOX, 13-AOS, and 13-AOC, which are all localized to plastids. Subsequently, 12-OPDA is transported to peroxisome for further conversion to JA. The 13-AOS pathway also produces 13-ketols, including 9-hydroxy-12-oxo-10(E)-octadecenoic acid (9,12-KOMA), 9-hydroxy-12-oxo-10(E),15(Z)-octadecadienoic acid (9,12-KODA), 13-hydroxy-12-oxo-9(Z)-octadecenoic acid (13,12-KOMA), and 13-hydroxy-12-oxo-9(Z),15(Z)-octadecadienoic acid (13,12-KODA). The 9-AOS pathway produces 10-oxo-11(Z)-phytoenoic acid (10-OPEA), 10-oxo-11(Z),15(Z)-phytodienoic acid (10-OPDA), and their derivatives collectively called “death acids” due to their strong programmed cell death-inducing activity [17]. Recently, an additional AOS branch of oxylipins, collectively named ‘graminoxins’, was identified in wheat, barley, sorghum, and rice roots [18], the function of which is currently unknown. Products from the 9-AOS pathway can undergo spontaneous cyclization into 10-OPEA and 10-OPDA, with speculation that a putative 9-AOC exists in this pathway [17][19][17,19]. The allene oxides produced by the 9-AOS pathway are also converted non-enzymatically to 9-ketols, including 9-hydroxy-10-oxo-12(Z)-octadecenoic acid (9,10-KOMA), 9-hydroxy-10-oxo-12(Z),15(Z)-octadecadienoic acid (9,10-KODA), 13-hydroxy-10-oxo-11(E)-octadecenoic acid (13,10-KOMA), and 13-hydroxy-10-oxo-11(E),15(Z)-octadecadienoic acid (13,10-KODA).

2. Biosynthesis and Occurrence of Ketols

Ketols are C18 compounds that contain a hydroxide group (-OH) present at the 9- or 13-carbon of the fatty acid backbone, a ketone group (C=O) present at the 10- or 12-carbon and ending in a carboxyl functional group. Ketols are designated as ‘α-’ or ‘γ-’ ketols based on the location of the hydroxide functional group in relation to the ketone functional group. If two double-bonds are present, then these ketols are referred to under the abbreviation of ‘KODA’ and are produced from C18:3 substrate, while if a single double-bond is present, then the ketols are referred to as ‘KOMA’ and are produced from the C18:2 substrate [22]. Ketols are grouped into ‘9-AOS-derived ketols’ (9-ketols for short) or ’13-AOS-derived ketols’ (13-ketols) based on the regiospecific 9- or 13-AOS enzymes that produce them. It should be noted that the following four ketols, 9,12-KODA, 9,12-KOMA, 13,10-KODA, and 13,10-KOMA, are classified as reactive electrophilic species (RES) because they contain an α,ß unsaturated carbonyl group. α,ß-unsaturated carbonyls are known to interact with nucleic acids and proteins to initiate adverse biological effects [23]. The proposed mechanism for the synthesis of ketols begins with the conversion of 9- or 13-hydroperoxide fatty acid substrates into epoxide intermediates via AOS activity. In this specific example, hydrogen donated from a hydronium (H3O+) ion attaches to the oxygen in the epoxide ring, leading to the breakage of the epoxide ring and the formation of a hydroxyl functional group on the 10-carbon position. Subsequent hydrolysis reactions occur to add another hydroxyl functional group to the 13-carbon position, leading to the transfer of electrons through double bonds and the formation of an ‘enol’ product that undergoes tautomerization. In the final step of ketol synthesis, a compound acting as a base attracts the hydrogen attached to the carbonyl functional group on the 10-carbon, leading to the ketol product of 13,10-KODA in this specific example. In addition to 13-AOS that all characterized plant species possess, many monocot species encode an additional clade of putatively extraplastidic AOS isoforms that possess dual 9/13-AOS activity [6][24][6,24]. It is likely that these 9/13-AOS isoforms are responsible for the biosynthesis of 9-oxylipins, death acids, and 9-ketols in addition to 13-ketols. Currently, little information is known about the regulation of the synthesis of ketols. It is suggested that ketol production is dependent on JA, as the expression of LOX and AOS genes involved in ketol production is dependent on JA [6], and disruption of JA biosynthesis in maize results in reduced levels of wound-induced ketols [25]. The 13-AOS enzymatic branch occurs primarily in plastids where it acts upon hydroperoxides of either C18:3 or C18:2 acids produced by 13-LOXs [5]. The products of 13-AOS are short-lived epoxide intermediates that are either non-enzymatically hydrolyzed to form α- and γ- ketols or undergo enzymatic cyclization by 13-AOC, the latter pathway leading to the eventual production of JA. Due to C18:3 being the predominant fatty acid found in the glycerolipids present in plastid membranes, a higher percentage of products derived from C18:3 than C18:2 is expected to be produced in these organelles. Typically, AOSs exhibiting 13-AOS activity are categorized as ‘CYP74A’ enzymes [26]. It is important to note that cyclization of the allene oxide intermediate synthesized under the 13-AOS pathway is possible with the presence of 13-AOC. It has been recently shown that 13-LOXs, 13-AOS, and 13-AOC form a protein complex to channel substrate specifically for the synthesis of JA [27]. However, it is not known if the formation of this complex interferes with ketol synthesis through substrate competition. The majority of tested 9-LOXs and 9-AOSs have been reported to be localized to the cytosol and/or organelles other than the chloroplast [6][28][29][6,28,29]. To date, all characterized cytosolic AOSs possessing 9-AOS activity display a strong affinity for both 9- and 13-hydroperoxide substrates; thus, they are characterized as dual-specific enzymes [6][19][23][28][29][30][31][32][6,19,23,28,29,30,31,32]. Typically, AOSs that have this dual-specific activity are grouped within the CYP74C subfamily [26][28][26,28]. Examples of dual-specific AOS genes include a barley enzyme capable of producing α-ketols from both 9- and 13-hydroperoxides of fatty acids [23]. The 9/13-AOSs were identified in other monocot species, including maize [6][19][6,19], rice [33], and sugarcane [34], based on their phylogenetic relationship and ability to produce both 9- and 13-ketols. Some dicot species also contain a similar mixed-function AOS, including tulips [35], tomatoes [26], and potatoes [28]. The best-characterized JA-producing and plastid-localized 13-AOS enzyme is present in both dicot and monocot plant species. Plant species are known to contain different numbers of AOS genes ranging from a single 13-AOS gene in Arabidopsis to three 13-AOS genes and two 9/13 mixed function AOS genes in maize [6], to twelve mixed function 9/13-AOS genes in sugarcane [34]. The mixed function 9/13-AOSs have been reported in fewer dicot species than monocot species. Flax, barley, maize, rice, tulips, petunia, potato, and tomato are among a few of the plant species identified so far that contain a dual-specific 9/13-AOS enzyme and therefore synthesize both 9- and 13-ketols [6][19][24][28][29][30][31][32][6,19,24,28,29,30,31,32]. Little is known about the evolution of CYP74 enzymes, especially AOS genes. Although Arabidopsis AOS shares structural similarities to other enzymes in mammals [14], it is not known whether mammalian systems possess AOS-like enzymes. Interestingly, 9/13-AOS genes are present in lancelets, which are considered to be an evolutionary intermediate between vertebrates and invertebrates [36]. This recombinant enzyme from the lancelet, Branchiostoma belcheri Gray, was active towards both 9- and 13-hydroperoxides, producing ketols 13,12-KOMA, 13,12-KODA, 9,10-KOMA, and 9,10-KODA [36]. Both soft and stony corals also possess AOS enzymes, though it is not reported if these enzymes produce ketols [37][38][37,38]. Because liverworts, mosses, and green algae contain AOS enzymes [29][30][32][29,30,32], it is suggested that the introduction of AOS genes occurred before the evolution of terrestrial plants, and AOS enzymes were likely present in the last common ancestor of plants and animals [14][34][14,34]. Recently, a metabolic analysis of wounded Physcomitrium patens Mitt knockout mutants of the PpAOS1 gene uncovered that this enzyme is responsible for the synthesis of two α-ketols [39]. Available data suggest that dual-specific AOS enzymes localized either to cytosol or chloroplast likely evolved before the evolution of flowering plant species [14][29][34][14,29,34].

3. Ketols Serve Signaling Roles in Plant–Pathogen Interactions

Several studies implicated ketols in playing a role in defense against biotic stresses. Such a role was first suggested for ketols produced in below-ground organs, as these tissues are characterized by especially high activity of 9/13-AOS enzymes and high levels of ketol production [26][28][35][26,28,35]. Recent research suggests that ketols are involved in plant–pathogen interactions in above-ground tissues. Exogenous application of 9,10-KODA results in strong induction of expression of several SA-inducible PR genes in tobacco (Nicotiana tabacum) leaves to the extent similar to the effect of the application of SA, suggesting that 9,10-KODA may be associated with defenses against pathogen infections and systemic acquired resistance (SAR) [40]. While this study provided strong evidence for the potential relevance of 9,10-KODA to SAR induction, it is unknown if 9,10-KODA acts independently of SA on the induction of SAR marker genes. A further report supported the defensive role of 9,10-KODA as an exogenous application of 9,10-KODA to grape berries suppressed the disease progression by Glomerella cingulata Stonem [41]. Recent genetic and pharmacological evidence obtained by the analysis of maize knockout mutants identified 9,10-KODA as a major xylem-mobile long-distance signal required for the activation of induced systemic resistance (ISR) against leaf pathogens triggered by root colonization with the beneficial fungal symbiont Trichoderma virens [42], suggesting a signaling role for this molecule. Moreover, several γ-ketols were subsequently shown as additional ISR priming agents induced transiently in leaves of maize in response to T. virens-triggered ISR [22]. Importantly, these studies indicated that both 9- and 13-ketols serve as important signals for the induction of ISR in maize [22][42][22,42]. Additionally, several ketols, including 9,10-KODA, 9,12-KODA, 13,12-KODA, and 13,12-KOMA, were upregulated in Colletotrichum graminicola-infected maize plants that were treated with pentyl leaf volatiles (PLVs); suggesting a positive correlation with PLV-mediated pathogen resistance [43]. The 9,10-KODA and other 9-LOX products were highly induced upon infection of maize stems by the hemibiotrophic pathogen Fusarium graminearum Petch, the causal agent of Gibberella Stalk Rot (GSR) [44]. Significantly higher accumulation of this ketol was observed in the maize line resistant to GSR, whereas Zmlox5 mutants disrupted in 9,10-KODA production displayed increased susceptibility to GSR [44].

References

  1. Yan, Y.; Christensen, S.; Isakeit, T.; Engelberth, J.; Meeley, R.; Hayward, A.; Emergy, R.J.N.; Kolomiets, M.V. Disruption of OPR7 and OPR8 Reveals the Versatile Functions of Jasmonic Acid in Maize Development and Defense. Plant Cell 2012, 24, 1420–1436.Yan, Yuanxin; Christensen, Shawn; Isakeit, Tom; Engelberth, Jürgen; Meeley, Robert; Hayward, Allison; Emery, R. J. Neil; Kolomiets, Michael V. Disruption of OPR7 and OPR8 Reveals the Versatile Functions of Jasmonic Acid in Maize Development and Defense. The Plant Cell 2012, 24, 1420, 10.1105/tpc.111.094151.
  2. Yan, Y.; Huang, P.-C.; Borrego, E.; Kolomiets, M. New perspectives into jasmonate roles in maize. Plant Signal. Behav. 2014, 9, e970442.Yan, Yuanxin; Huang, Pei-Cheng; Borrego, Eli; Kolomiets, Michael New perspectives into jasmonate roles in maize. Plant Signal. Behav. 2014, 9, e970442, 10.4161/15592316.2014.970442.
  3. Ahmad, R.M.; Cheng, C.; Sheng, J.; Wang, W.; Ren, H.; Aslam, M.; Yan, Y. Interruption of Jasmonic Acid Biosynthesis Causes Differential Responses in the Roots and Shoots of Maize Seedlings against Salt Stress. Int. J. Mol. Sci. 2019, 20, 6202.Feussner, Ivo; Wasternack, Claus The lipoxygenase pathway. Annu Rev Plant Biol 2002, 53, 275, 10.1146/annurev.arplant.53.100301.135248..
  4. Savchenko, T.V.; Zastrijnaja, O.M.; Klimov, V.V. Oxylipins and plant abiotic stress resistance. Biochemistry 2014, 79, 362–375.Borrego, Eli J.; Kolomiets, Michael V. Synthesis and Functions of Jasmonates in Maize. Plants 2016, 5, 41, 10.3390/plants5040041.
  5. Feussner, I.; Wasternack, C. The lipoxygenase pathway. Annu. Rev. Plant Biol. 2002, 53, 275–297.Weichert, H.; Stenzel, I.; Berndt, E.; Wasternack, C.; Feussner, I. Metabolic profiling of oxylipins upon salicylate treatment in barley leaves--preferential induction of the reductase pathway by salicylate(1). FEBS Lett 1999, 464, 133-137, 10.1016/s0014-5793(99)01697-x..
  6. Borrego, E.J.; Kolomiets, M.V. Synthesis and Functions of Jasmonates in Maize. Plants 2016, 5, 41.Feussner, I.; Kühn, H.; Wasternack, C. Lipoxygenase-dependent degradation of storage lipids. Trends Plant Sci 2001, 6, 268-273, 10.1016/s1360-1385(01)01950-1..
  7. Weichert, H.; Stenzel, I.; Berndt, E.; Wasternack, C.; Feussner, I. Metabolic profiling of oxylipins upon salicylate treatment in barley leaves—Preferential induction of the reductase pathway by salicylate. FEBS Lett. 1999, 464, 133–137.Kawano, T. Roles of the reactive oxygen species-generating peroxidase reactions in plant defense and growth induction. Plant Cell Rep 2003, 21, 829-837, 10.1007/s00299-003-0591-z.
  8. Feussner, I.; Kühn, H.; Wasternack, C. Lipoxygenase-dependent degradation of storage lipids. Trends Plant Sci. 2001, 6, 268–273.Howe, Gregg A.; Schilmiller, Anthony L. Oxylipin metabolism in response to stress. Curr Opin Plant Biol 2002, 5, 230-236, 10.1016/s1369-5266(02)00250-9.
  9. Kawano, T. Roles of the reactive oxygen species-generated peroxidase reactions in plant defense and growth induction. Plant Cell Rep. 2003, 21, 829–837.Grechkin, Alexander N. Hydroperoxide lyase and divinyl ether synthase. Prostaglandins and Other Lipid Mediat. 2002, 68, 457-470, 10.1016/s0090-6980(02)00048-5.
  10. Howe, G.A.; Schilmiller, A.L. Oxylipin metabolism in response to stress. Curr. Opin. Plant Biol. 2002, 5, 230–236.Liavonchanka, Alena; Feussner, Ivo Lipoxygenases: occurrence, functions and catalysis. J Plant Physiol 2006, 163, 348-357, 10.1016/j.jplph.2005.11.006.
  11. Grechkin, A.N. Hydroperoxide lyase and divinyl ether synthase. Prostaglandins Other Lipid Mediat. 2002, 68, 457–470.Lee, Dong-Sun; Nioche, Pierre; Hamberg, Mats; Raman, C.S. Structural insights into the evolutionary paths of oxylipin biosynthetic enzymes. Nature 2008, 455, 363-368, 10.1038/nature07307.
  12. Liavonchanka, A.; Feussner, I. Lipoxygenases: Occurrence, functions, and catalysis. J. Plant Physiol. 2006, 163, 348–357.Toporkova, Yana Y.; Askarova, Elena K.; Gorina, Svetlana S.; Ogorodnikova, Anna V.; Mukhtarova, Lucia S.; Grechkin, Alexander N. Epoxyalcohol synthase activity of the CYP74B enzymes of higher plants. Biochim Biophys Acta Mol Cell Biol Lipids 2020, 1865, 158743, 10.1016/j.bbalip.2020.158743.
  13. Stumpe, M.; Feussner, I. Formation of oxylipins by CYP74 enzymes. Phytochem. Rev. 2006, 5, 347–357.Stumpe, M.; Feussner, I. Formation of oxylipins by CYP74 enzymes. Phytochem Rev 2006, 5, 347-357, 10.1007/s11101-006-9038-9.
  14. Lee, D.-S.; Nioche, P.; Hamberg, M.; Raman, C.S. Structural insights into the evolutionary paths of oxylipin biosynthetic enzymes. Nature 2008, 455, 363–368.Lee, Dong-Sun; Nioche, Pierre; Hamberg, Mats; Raman, C.S. Structural insights into the evolutionary paths of oxylipin biosynthetic enzymes. Nature 2008, 455, 363-368, 10.1038/nature07307.
  15. Toporkova, Y.Y.; Askarova, E.K.; Gorina, S.S.; Ogorodnikova, A.V.; Mukhtarova, L.S.; Grechkin, A.N. Epoxyalcohol synthase activity of the CYP74B enzymes of higher plants. Biochim. Biophys. Acta Mol. Cell Biol. Lipids 2020, 1865, 158743.Toporkova, Yana Y.; Askarova, Elena K.; Gorina, Svetlana S.; Ogorodnikova, Anna V.; Mukhtarova, Lucia S.; Grechkin, Alexander N. Epoxyalcohol synthase activity of the CYP74B enzymes of higher plants. Biochim Biophys Acta Mol Cell Biol Lipids 2020, 1865, 158743, 10.1016/j.bbalip.2020.158743.
  16. Toporkova, Y.Y.; Smirnova, E.O.; Gorina, S.S.; Mukhtarova, L.S.; Grechkin, A.N. Detection of the first higher plant epoxyalcohol synthase: Molecular cloning and characterisation of the CYP74M2 enzyme of spikemoss Selaginella moellendorffii. Phytochemistry 2018, 156, 73–82.
  17. Christensen, S.A.; Huffaker, A.; Kaplan, F.; Schmelz, E.A. Maize death acids, 9-lipoxygenase-derived cyclopente(a)nones, display activity as cytotoxic phytoalexins and transcriptional mediators. Proc. Natl. Acad. Sci. USA 2015, 112, 11407–11412.
  18. Grechkin, A.N.; Ogorodnikova, A.V.; Egorova, A.M.; Mukhitova, F.K.; Ilyina, T.M.; Khairutdinov, B.I. Allene Oxide Synthase Pathway in Cereal Roots: Detection of Novel Oxylipin Graminoxins. ChemistryOpen 2018, 7, 336–343.
  19. Ogorodnikova, A.V.; Gorina, S.S.; Mukhtarova, L.S.; Mukhitova, F.K.; Toporkova, Y.Y.; Hamberg, M.; Grechkin, A.N. Stereospecific biosynthesis of (9S,13S)-10-oxo-phytoenoic acid in young maize roots. Biochim. Biophys. Acta 2015, 1851, 1262–1270.
  20. Blée, E. Phytooxylipins and plant defense reactions. Prog. Lipid Res. 1998, 37, 33–72.
  21. Deboever, E.; Deleu, M.; Mongrand, S.; Lins, L.; Fauconnier, M.-L. Plant–pathogen Interactions: Underestimated Roles of Phyto-oxylipins. Trends Plant Sci. 2020, 25, 22–34.
  22. Wang, K.-D.; Gorman, Z.; Huang, P.-C.; Kenerley, C.M.; Kolomiets, M.V. Trichoderma virens colonization of maize roots triggers rapid accumulation of 12-oxophytodienoate and two γ-ketols in leaves as priming agents of induced systemic resistance. Plant Signal. Behav. 2020, 15, 1792187.
  23. Mueller, M.J.; Berger, S. Reactive electrophilic oxylipins: Pattern recognition and signaling. Phytochemistry 2009, 70, 1511–1521.
  24. Maucher, H.; Hause, B.; Feussner, I.; Ziegler, J.; Wasternack, C. Allene oxide synthases of barley (Hordeum vulgare cv. Salome): Tissue specific regulation in seedling development. Plant J. 2000, 21, 199–213.
  25. He, Y.; Borrego, E.J.; Gorman, Z.; Huang, P.-C.; Kolomiets, M.V. Relative contribution of LOX10, green leaf volatiles and JA to wound-induced local and systemic oxylipin and hormone signature in Zea mays (maize). Phytochemistry 2020, 174, 112334.
  26. Itoh, A.; Schilmiller, A.L.; McCaig, B.C.; Howe, G.A. Identification of a Jasmonate-regulated Allene Oxide Synthase That Metabolizes 9-Hydroperoxides of Linoleic and Linolenic Acids. J. Biol. Chem. 2002, 277, 46051–46058.
  27. Pollmann, S.; Springer, A.; Rustgi, S.; Wettstein, D.V.; Kang, C.; Reinbothe, C.; Reinbothe, S. Substrate channeling in oxylipin biosynthesis through a protein complex in the plastid envelope of Arabidopsis thaliana. J. Exp. Bot. 2019, 70, 1483–1495.
  28. Stumpe, M.; Göbel, C.; Demchenko, K.; Hoffmann, M.; Klösgen, R.B.; Pawlowski, K.; Feussner, I. Identification of an allene oxide synthase (CYP74C) that leads to formation of α-ketols from 9-hydroperoxides of linoleic and linolenic acid in below-groundvorgans of potato. Plant J. Cell Mol. Biol. 2006, 47, 883–896.
  29. Scholz, J.; Brodhun, F.; Hornung, E.; Herrfurth, C.; Stumpe, M.; Beike, A.K.; Faltin, B.; Frank, W.; Reski, R.; Feussner, I. Biosynthesis of allene oxides in Physcomitrella patens. BMC Plant Biol. 2012, 12, 228.
  30. Koeduka, T.; Ishizaki, K.; Mwenda, C.M.; Hori, K.; Sasaki-Sekimoto, Y.; Ohta, H.; Kohchi, T.; Matsui, K. Biochemical characteriza- tion 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 2015, 242, 1175–1186.
  31. Toporkova, Y.Y.; Smirnova, E.O.; Mukhtarova, L.S.; Gorina, S.S.; Grechkin, A.N. Catalysis by allene oxide synthases (CYP74A and CYP74C): Alterations by the Phe/Leu mutation at the SRS-1 region. Phytochemistry 2020, 169, 112152.
  32. Toporkova, Y.Y.; Askarova, E.K.; Gorina, S.S.; Mukhtarova, L.S.; Grechkin, A.N. Oxylipin biosynthesis in spikemoss Selaginella moellendorffi: Identification of allene oxide synthase (CYP74L2) and hydroperoxide lyase (CYP74L1). Phytochemistry 2022, 195, 113051.
  33. Yoeun, S.; Sukhanov, A.; Han, O. Separation of enzymatic functions and variation of spin state of rice allene oxide synthase-1 mutation of Phe-92 and Pro-430. Bioorg. Chem. 2016, 68, 9–14.
  34. Sun, T.; Chen, Y.; Feng, A.; Zou, W.; Wang, D.; Lin, P.; Chen, Y.; You, C.; Que, Y.; Su, Y. The allene oxide synthase gene family in sugarcane and its involvement in disease resistance. Ind. Crops Prod. 2023, 192, 116136.
  35. Grechkin, A.N.; Mukhtarova, L.S.; Hamberg, M. The lipoxygenase pathway in tulip (Tulipa gesneriana): Detection of the ketol route. Biochem. J. 2000, 352, 501–509.
  36. Toporkova, Y.Y.; Smirnova, E.O.; Lantsova, N.V.; Mukhtarova, L.S.; Grechkin, A.N. Detection of the First Epoxyalcohol Syn- thase/Allene Oxide Synthase (CYP74 clan) in the Lancelet (Branchiostoma belcheri, Chordata). Int. J. Mol. Sci. 2021, 22, 4737.
  37. Brash, A.R. Catalase-Related Allene Oxide Synthase, on a Biosynthetic Route to Fatty Acid Cyclopentenones: Expression and Assay of the Enzyme and Preparation of the 8R-HPETE Substrate. Methods Enzymol. 2018, 605, 51–68.
  38. Lõhelaid, H.; Samel, N. Eicosanoid Diversity of Stony Corals. Mar. Drugs 2018, 16, 10.
  39. Resemann, H.C.; Feussner, K.; Hornung, E.; Feussner, I. A non-targeted metabolomics analysis identifies wound-induced oxylipins in Physcomitrium patens. Front. Plant Sci. 2023, 13, 1085915.
  40. Endo, J.-I.; Takahashi, W.; Yokoyama, M.; Tanaka, O. Induction of gene expression for systemic acquired resistance in tobacco by 9-hydroxy-10-oxo-12(Z),15(Z)-octadecadienoic acid (KODA). Can. J. Plant Sci. 2013, 93, 827–830.
  41. Wang, S.; Saito, T.; Ohkawa, K.; Ohara, H.; Shishido, M.; Ikeura, H.; Takagi, K.; Ogawa, S.; Yokoyama, M.; Kondo, S. α-Ketol linolenic acid (KODA) application affects endogenous abscisic acid, jasmonic acid and aromantic volatiles in grapes infected by a pathogen (Glomerella cingulata). J. Plant Physiol. 2016, 192, 90–97.
  42. Wang, K.-D.; Borrego, E.J.; Kenerley, C.M.; Kolomiets, M.V. Oxylipins Other Than Jasmonic Acid Are Xylem-Resident Signals Regulating Systemic Resistance Induced by Trichoderma virens in Maize. Plant Cell 2020, 32, 166–185.
  43. Gorman, Z.; Tolley, J.P.; Koiwa, H.; Kolomiets, M.V. The Synthesis of Pentyl Leaf Volatiles and Their Role in Resistance to Anthracnose Leaf Blight. Front. Plant Sci. 2021, 12, 719587.
  44. Wang, Q.; Sun, Y.; Wang, F.; Huang, P.-C.; Wang, Y.; Ruan, X.; Ma, L.; Li, X.; Kolomiets, M.V.; Gao, X. Transcriptome and Oxylipin Profiling Joint Analysis Reveals Opposite Roles of 9-Oxylipins and Jasmonic Acid in Maize Resistance to Gibberella Stalk Rot. Front. Plant Sci. 2021, 12, 699146.
More
© Text is available under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/)
Video Production Service
Feedback