Plant-pathogen interactions: underestimated roles of phyto-oxylipins

[en] Plant oxylipins are produced under a wide range of stress conditions, and even though they are well known to activate stress-related signalling pathways, the non-signalling roles of phyto-oxylipins are poorly understood. We describe oxylipins as direct biocidal agents and propose that structure-function relationships play here a pivotal role. Indeed, based on their chemical configuration, plant oxylipins, such as reactive oxygen and electrophile species, activate defence-related genes expression. We also propose that their ability to interact with pathogen membranes is important, but still misunderstood, and that they are involved in cross-kingdom communication. Taken as a whole, the current literature suggests that plant oxylipins have a high potential as biocontrol agents. However, the mechanisms underlying these multi-faceted compounds remain largely unknown.

Disciplines :

Chemistry
Phytobiology (plant sciences, forestry, mycology...)
Biochemistry, biophysics & molecular biology

Author, co-author :

Deboever, Estelle ; Université de Liège - ULiège > Agronomie, Bio-ingénierie et Chimie (AgroBioChem) > Chimie des agro-biosystèmes

Deleu, Magali ; Université de Liège - ULiège > Agronomie, Bio-ingénierie et Chimie (AgroBioChem) > Chimie des agro-biosystèmes

Mongrand, Sébastien

Lins, Laurence ^✱; Université de Liège - ULiège > Agronomie, Bio-ingénierie et Chimie (AgroBioChem) > Chimie des agro-biosystèmes

Fauconnier, Marie-Laure ^✱; Université de Liège - ULiège > Agronomie, Bio-ingénierie et Chimie (AgroBioChem) > Chimie des agro-biosystèmes

^✱ These authors have contributed equally to this work.

Language :

English

Title :

Plant-pathogen interactions: underestimated roles of phyto-oxylipins

Publication date :

2020

Journal title :

Trends in Plant Science

ISSN :

1360-1385

eISSN :

1878-4372

Publisher :

Elsevier, Oxford, United Kingdom

Volume :

Issue :

Pages :

22-34

Peer reviewed :

Peer Reviewed verified by ORBi

Additional URL :

https://www.cell.com/trends/plant-science/fulltext/S1360-1385(19)30246-8

Name of the research project :

Fonds pour la formation à la Recherche dans l’Industrie et dans l’Agriculture’ (FRIA) grant (5100617F)

Funders :

F.R.S.-FNRS - Fonds de la Recherche Scientifique [BE]
FRIA - Fonds pour la Formation à la Recherche dans l'Industrie et dans l'Agriculture [BE]

Available on ORBi :

since 21 November 2019

Statistics

Number of views

298 (39 by ULiège)

Number of downloads

782 (34 by ULiège)

More statistics

Scopus citations^®

Scopus citations^®
without self-citations

OpenCitations

Bibliography

Barbosa, M., et al. Biologically active oxylipins from enzymatic and nonenzymatic routes in macroalgae. Mar. Drugs, 14, 2016, 23.
Howe, G.A., Plant hormones: metabolic end run to jasmonate. Nat. Chem. Biol. 14 (2018), 109–110.
Koo, A.J., Metabolism of the plant hormone jasmonate: a sentinel for tissue damage and master regulator of stress response. Phytochem. Rev. 17 (2018), 51–80.
Wasternack, C., Feussner, I., The oxylipin pathways: biochemistry and function. Annu. Rev. Plant Biol. 69 (2018), 363–386.
Garreta-Lara, E., et al. Effect of psychiatric drugs on Daphnia magna oxylipin profiles. Sci. Total Environ. 644 (2018), 1101–1109.
Li, Q., et al. Transporter-mediated nuclear entry of jasmonoyl-isoleucine is essential for jasmonate signaling. Mol. Plant 10 (2017), 695–708.
Farmer, E.E., et al. Jasmonates and related oxylipins in plant responses to pathogenesis and herbivory. Curr. Opin. Plant Biol. 6 (2003), 372–378.
Ponce de León, I., et al. Oxylipins in moss development and defense. Front. Plant Sci., 6, 2015, 483.
Ogorodnikova, A.V., et al. Oxylipins in the spikemoss Selaginella martensii: detection of divinyl ethers, 12-oxophytodienoic acid and related cyclopentenones. Phytochemistry 118 (2015), 42–50.
Lupette, J., et al. Non-enzymatic synthesis of bioactive isoprostanoids in the diatom Phaeodactylum following oxidative stress 1. Plant Physiol. 178 (2018), 1344–1357.
Griffiths, G., Biosynthesis and analysis of plant oxylipins. Free Radic. Res. 49 (2015), 565–582.
Vellosillo, T., et al. Defense activated by 9-lipoxygenase-derived oxylipins requires specific mitochondrial proteins. Plant Physiol 161 (2013), 617–627.
Alméras, E., et al. Reactive electrophile species activate defense gene expression in Arabidopsis. Plant J 34 (2003), 205–216.
Mène-Saffrané, L., et al. Nonenzymatic oxidation of trienoic fatty acids contributes to reactive oxygen species management in Arabidopsis. J. Biol. Chem. 284 (2009), 1702–1708.
Mueller, S., et al. General detoxification and stress responses are mediated by oxidized lipids through TGA transcription factors in Arabidopsis. Plant Cell 20 (2008), 768–785.
Brodhun, F., Feussner, I., Oxylipins in fungi. FEBS J 278 (2011), 1047–1063.
Fischer, G.J., Keller, N.P., Production of cross-kingdom oxylipins by pathogenic fungi: an update on their role in development and pathogenicity. J. Microbiol. 54 (2016), 254–264.
Wasternack, C., Song, S., Jasmonates: biosynthesis, metabolism, and signaling by proteins activating and repressing transcription. J. Exp. Bot. 68 (2017), 1303–1321.
Genva, M., et al. New insights into the biosynthesis of esterified oxylipins and their involvement in plant defense and developmental mechanisms. Phytochem. Rev. 8 (2019), 343–359.
Islam, A.B., et al. Genomic, lipidomic and metabolomic analysis of cyclooxygenase-null cells: eicosanoid storm, cross talk, and compensation by COX-1. Genomics Proteomics Bioinformatics 14 (2016), 81–93.
Behl, T., et al. Role of leukotrienes in diabetic retinopathy. Prostaglandins Other Lipid Mediat 122 (2016), 1–9.
Battilani, P., et al. Oxylipins from both pathogen and host antagonize jasmonic acid-mediated defence via the 9-lipoxygenase pathway in Fusarium verticillioides infection of maize. Mol. Plant Pathol. 19 (2018), 2162–2176.
Allmann, S., et al. Oxylipin channelling in Nicotiana attenuata: lipoxygenase 2 supplies substrates for green leaf volatile production. Plant Cell Environ 33 (2010), 2028–2040.
Chini, A., et al. An OPR3-independent pathway uses 4,5-didehydrojasmonate for jasmonate synthesis. Nat. Chem. Biol. 14 (2018), 171–178.
Böttcher, C., Pollmann, S., Plant oxylipins: plant responses to 12-oxo-phytodienoic acid are governed by its specific structural and functional properties. FEBS J 276 (2009), 4693–4704.
Santino, A., et al. Jasmonate signaling in plant development and defense response to multiple (a)biotic stresses. Plant Cell Rep 32 (2013), 1085–1098.
Sun, Y., et al. The role of wheat jasmonic acid and ethylene pathways in response to Fusarium graminearum infection. Plant Growth Regul 80 (2016), 69–77.
Zhang, L., et al. Jasmonate signaling and manipulation by pathogens and insects. J. Exp. Bot 68 (2017), 1371–1385.
Heitz, T., et al. The rise and fall of jasmonate biological activities. Subcell. Biochem. 86 (2016), 405–426.
Fonseca, S., et al. (+)-7-iso-Jasmonoyl-L-isoleucine is the endogenous bioactive jasmonate. Nat. Chem. Biol. 5 (2009), 344–350.
Delaplace, P., et al. Oxylipin profile and antioxidant status of potato tubers during extended storage at room temperature. Plant Physiol. Biochem. 46 (2008), 1077–1084.
Fauconnier, M.L., et al. Changes in oxylipin synthesis after Phytophthora infestans infection of potato leaves do not correlate with resistance. Plant Physiol. Biochem. 46 (2008), 823–831.
Ghanem, M.E., et al. Organ-dependent oxylipin signature in leaves and roots of salinized tomato plants (Solanum lycopersicum). J. Plant Physiol. 169 (2012), 1090–1101.
Gosset, V., et al. Attacks by a piercing–sucking insect (Myzus persicae Sultzer) or a chewing insect (Leptinotarsa decemlineata Say) on potato plants (Solanum tuberosum L.) induce differential changes in volatile compound release and oxylipin synthesis. J. Exp. Bot. 60 (2009), 1231–1240.
Granér, G., et al. Screening of oxylipins for control of oilseed rape (Brassica napus) fungal pathogens. Phytochemistry 63 (2003), 89–95.
León Morcillo, R.J., et al. Plant 9-lox oxylipin metabolism in response to arbuscular mycorrhiza. Plant Signal. Behav. 7 (2012), 1584–1588.
Tsitsigiannis, D.I., Keller, N.P., Oxylipins as developmental and host-fungal communication signals. Trends Microbiol 15 (2007), 109–118.
Vicente, J., et al. Role of 9-lipoxygenase and α-dioxygenase oxylipin pathways as modulators of local and systemic defense. Mol. Plant 5 (2012), 914–928.
Hamberg, M., et al. Activation of the fatty acid α-dioxygenase pathway during bacterial infection of tobacco leaves: formation of oxylipins protecting against cell death. J. Biol. Chem. 278 (2003), 51796–51805.
Mariutto, M., et al. Reprogramming of fatty acid and oxylipin synthesis in rhizobacteria-induced systemic resistance in tomato. Plant Mol. Biol. 84 (2014), 455–467.
Christensen, S.A., et al. Maize death acids, 9-lipoxygenase-derived, cyclopente(a)nones, display activity as cytotoxic phytoalexins and transcriptional mediators. Proc. Natl Acad. Sci. U. S. A. 112 (2015), 11407–11412.
Andersson, M.X., et al. Oxylipin profiling of the hypersensitive response in Arabidopsis thaliana: formation of a novel oxo-phytodienoic acid-containing galactolipid, arabidopside E. J. Biol. Chem. 281 (2006), 31528–31537.
Jones, J.D.G., Dangl, J.L., The plant immune system. Nature 444 (2006), 323–329.
Zhang, W., et al. Different pathogen defense strategies in Arabidopsis: more than pathogen recognition. Cells, 7, 2018, E252.
Henry, G., et al. PAMPs, MAMPs, DAMPs and others: an update on the diversity of plant immunity elicitors. Biotechnol. Agron. Soc, 16, 2012, 12.
Ramirez-Prado, J.S., et al. Plant immunity: from signaling to epigenetic control of defense. Trends Plant Sci 23 (2018), 833–844.
Chuberre, C., et al. Plant immunity is compartmentalized and specialized in roots. Front. Plant Sci., 9, 2018, 1692.
Tripathi, D., et al. Chemical elicitors of systemic acquired resistance – salicylic acid and its functional analogs. Curr. Plant Biol. 17 (2019), 48–59.
Thaler, J.S., et al. Evolution of jasmonate and salicylate signal crosstalk. Trends Plant Sci 17 (2012), 260–270.
Heyer, M., et al. A holistic approach to analyze systemic jasmonate accumulation in individual leaves of Arabidopsis rosettes upon wounding. Front. Plant Sci., 9, 2018, 1569.
Han, G.Z., Evolution of jasmonate biosynthesis and signalling mechanisms. J. Exp. Bot. 68 (2017), 1323–1331.
Atkinson, N.J., Urwin, P.E., The interaction of plant biotic and abiotic stresses: from genes to the field. J. Exp. Bot. 63 (2012), 3523–3544.
Chung, S.H., et al. Host plant species determines symbiotic bacterial community mediating suppression of plant defenses. Sci. Rep., 7, 2017, 39690.
Hu, Y., et al. Jasmonate regulates leaf senescence and tolerance to cold stress: crosstalk with other phytohormones. J. Exp. Bot. 68 (2017), 1361–1369.
Hoffmann, M., et al. Auxin–oxylipin crosstalk: relationship of antagonists. J. Integr. Plant Biol. 53 (2011), 429–445.
Taylor, J.E., et al. Crosstalk between plant responses to pathogens and herbivores: a view from the outside in. J. Exp. Bot. 55 (2004), 159–168.
Di, X., et al. Involvement of salicylic acid, ethylene and jasmonic acid signalling pathways in the susceptibility of tomato to Fusarium oxysporum. Mol. Plant Pathol. 18 (2017), 1024–1035.
Lortzing, T., Steppuhn, A., Jasmonate signalling in plants shapes plant–insect interaction ecology. Curr. Opin. Insect Sci. 14 (2016), 32–39.
Schilmiller, A.L., Howe, G.A., Systemic signaling in the wound response. Curr. Opin. Plant Biol. 8 (2005), 369–377.
Yan, C., Xie, D., Jasmonate in plant defence: sentinel or double agent?. Plant Biotechnol. J. 13 (2015), 1233–1240.
Koo, A.J.K., et al. A rapid wound signal activates the systemic synthesis of bioactive jasmonates in Arabidopsis. Plant J 59 (2009), 974–986.
Choi, W., et al. Rapid, long-distance electrical and calcium signaling in plants. Annu. Rev. Plant Biol. 67 (2016), 287–310.
Choi, W., et al. Orchestrating rapid long-distance signaling in plants with Ca2+, ROS and electrical signals. Plant J 90 (2018), 698–707.
Mousavi, S.A.R., et al. GLUTAMATE RECEPTOR-LIKE genes mediate leaf-to-leaf wound signalling. Nature 500 (2013), 422–426.
Toyota, M., et al. Glutamate triggers long-distance, calcium-based plant defense signaling. Science 6 (2018), 1112–1115.
Nguyen, T.C., et al. Identification of cell populations necessary for leaf-to- leaf electrical signaling in a wounded plant. Proc. Natl Acad. Sci. U. S. A. 115 (2018), 10178–10183.
Schuck, S., et al. The Nicotiana attenuata GLA1 lipase controls the accumulation of Phytophthora parasitica-induced oxylipins and defensive secondary metabolites. Plant Cell Environ 37 (2014), 1703–1715.
Prost, I., et al. Evaluation of the antimicrobial activities of plant oxylipins supports their involvement in defense against pathogens. Plant Physiol 139 (2005), 1902–1913.
Nakamura, S., Hatanaka, A., Green-leaf-derived C6-aroma compounds with potent antibacterial action that act on both Gram-negative and Gram-positive bacteria. J. Agric. Food Chem. 50 (2002), 7639–7644.
Ma, W., et al. Inhibitory effect of (E)-2-hexenal as a potential natural fumigant on Aspergillus flavus in stored peanut seeds. Ind. Crops Prod. 107 (2017), 206–210.
Farmer, E.E., Mueller, M.J., ROS-mediated lipid peroxidation and RES-activated signaling. Annu. Rev. Plant Biol. 64 (2013), 429–450.
Park, S., et al. Cyclophilin 20-3 relays a 12-oxo-phytodienoic acid signal during stress responsive regulation of cellular redox homeostasis. Proc. Natl Acad. Sci. U. S. A. 110 (2013), 9559–9564.
Müller, S.M., et al. The redox-sensitive module of cyclophilin 20-3, 2-cysteine peroxiredoxin and cysteine synthase integrates sulfur metabolism and oxylipin signaling in the high light acclimation response. Plant J 91 (2017), 995–1014.
Farmer, E.E., Davoine, C., Reactive electrophile species. Curr. Opin. Plant Biol. 10 (2007), 380–386.
Desbois, A.P., Smith, V.J., Antibacterial free fatty acids: activities, mechanisms of action and biotechnological potential. Appl. Microbiol. Biotechnol. 85 (2010), 1629–1642.
Kabara, J.J., et al. Fatty acids and derivatives as antimicrobial agents. Antimicrob. Agents Chemother. 2 (1972), 23–28.
Toporkova, Y.Y., et al. Antimicrobial activity of geometric isomers of etherolenic acid – the products of plant lipoxygenase cascade. Biochem. Biophys. Mol. Biol. 480 (2018), 139–142.
Deleu, M., et al. Linoleic and linolenic acid hydroperoxides interact differentially with biomimetic plant membranes in a lipid specific manner. Colloids Surf. B Biointerfaces 175 (2019), 384–391.
Sarwar, A., et al. Biocontrol activity of surfactin A purified from Bacillus NH-100 and NH-217 against rice bakanae disease. Microbiol. Res. 209 (2018), 1–13.
Deravel, J., et al. Mycosubtilin and surfactin are efficient, low ecotoxicity molecules for the biocontrol of lettuce downy mildew. Appl. Microbiol. Biotechnol. 98 (2014), 6255–6264.
Henry, G., et al. The bacterial lipopeptide surfactin targets the lipid fraction of the plant plasma membrane to trigger immune-related defence responses. Cell. Microbiol. 13 (2011), 1824–1837.
Gronnier, J., et al. Divide and rule: plant plasma membrane organization. Trends Plant Sci 23 (2018), 899–917.
Siebers, M., et al. Lipids in plant–microbe interactions. Biochim. Biophys. Acta 1861 (2016), 1379–1395.
Brodhagen, M., et al. Reciprocal oxylipin-mediated cross-talk in the Aspergillus–seed pathosystem. Mol. Microbiol. 67 (2008), 378–391.
Okazaki, Y., Saito, K., Roles of lipids as signaling molecules and mitigators during stress response in plants. Plant J 79 (2014), 584–596.
Singh, A., Del Poeta, M., Lipid signalling in pathogenic fungi. Cell. Microbiol. 13 (2011), 177–185.
Sagini, K., et al. Extracellular vesicles as conveyors of membrane-derived bioactive lipids in immune system. Int. J. Mol. Sci., 19, 2018, E1227.
Kachroo, A., Kachroo, P., Fatty acid–derived signals in plant defense. Annu. Rev. Phytopathol. 47 (2009), 153–176.
Walley, J.W., et al. Fatty acids and early detection of pathogens. Curr. Opin. Plant Biol. 16 (2013), 520–526.
Ishimaru, Y., et al. GTR1 is a jasmonic acid and jasmonoyl-l-isoleucine transporter in Arabidopsis thaliana. Biosci. Biotechnol. Biochem. 8451 (2017), 249–255.
Saito, H., et al. The jasmonate-responsive GTR1 transporter is required for gibberellin-mediated stamen development in Arabidopsis. Nat. Commun., 6, 2015, 6095.
Yang, C.C., Chang, K.W., Eicosanoids and HB-EGF/EGFR in cancer. Cancer Metastasis Rev 37 (2018), 385–395.
Wang, D., Dubois, R.N., Eicosanoids and cancer. Nat. Rev. Cancer 10 (2010), 181–193.
Cornejo-García, J.A., et al. Pharmacogenomics of prostaglandin and leukotriene receptors. Front. Pharmacol., 7, 2016, 316.
Noverr, M.C., et al. Production of eicosanoids and other oxylipins by pathogenic eukaryotic microbes. Clin. Microbiol. Rev. 16 (2003), 517–533.
Affeldt, K.J., et al. Aspergillus oxylipin signaling and quorum sensing pathways depend on G protein-coupled receptors. Toxins (Basel) 4 (2012), 695–717.
Affeldt, K.J., et al. Global survey of canonical Aspergillus flavus GPCRs. MBio 5 (2014), e01501–e01514.
Raviv, Z., et al. The anti-cancer activities of jasmonates. Cancer Chemother. Pharmacol. 71 (2013), 275–285.
Savchenko, T., et al. Arachidonic acid: an evolutionarily conserved signaling molecule modulates plant stress signaling networks. Plant Cell 22 (2010), 3193–3205.
Christensen, S.A., Kolomiets, M.V., The lipid language of plant–fungal interactions. Fungal Genet. Biol. 48 (2011), 4–14.
Gao, X., Kolomiets, M.V., Host-derived lipids and oxylipins are crucial signals in modulating mycotoxin production by fungi. Toxin Rev 28 (2009), 79–88.
Tsitsigiannis, D.I., et al. Aspergillus infection inhibits the expression of peanut 13S-HPODE-forming seed lipoxygenases. Mol. Plant Microbe Interact. 18 (2005), 1081–1089.
Maschietto, V., et al. Resistance to Fusarium verticillioides and fumonisin accumulation in maize inbred lines involves an earlier and enhanced expression of lipoxygenase (LOX) genes. J. Plant Physiol. 188 (2015), 9–18.
Pandey, S., et al. Two novel GPCR-type G proteins are abscisic acid receptors in Arabidopsis. Cell 136 (2009), 136–148.
Risk, J.M., et al. Reevaluation of abscisic acid-binding assays shows that G-protein-coupled receptor2 does not bind abscisic acid. Plant Physiol 150 (2009), 6–11.
Chini, A., et al. The fungal phytotoxin lasiojasmonate A activates the plant jasmonic acid pathway. J. Exp. Bot. 69 (2018), 3095–3102.
Chanclud, E., Morel, J., Plant hormones : a fungal point of view. Mol. Plant Pathol. 17 (2016), 1289–1297.
Patkar, R.N., et al. A fungal monooxygenase-derived jasmonate attenuates host innate immunity. Nat. Chem. Biol. 11 (2015), 733–740.
Tran Thanh, H., et al. Toxicity of fatty acid hydroperoxides towards Yarrowia lipolytica: implication of their membrane fluidizing action. Biochim. Biophys. Acta 1768 (2007), 2256–2262.
Wang, M., et al. Bidirectional cross-kingdom RNAi and fungal uptake of external RNAs confer plant protection. Nat. Plants, 2, 2017, 16151.
Weiberg, A., et al. Fungal small RNAs suppress plant immunity by hijacking host RNA interference pathways. Science 342 (2014), 118–123.
Ongena, M., et al. Surfactin and fengycin lipopeptides of Bacillus subtilis as elicitors of induced systemic resistance in plants. Environ. Microbiol. 9 (2007), 1084–1090.