Plant and Cell Physiology

Goossens,, Alain;Farmer, Edward, E

doi: 10.1093/pcp/pcz197pmid: 31626278

Oxylipins form a steadily growing group of signaling molecules that comprise oxygenated fatty acids and metabolites derived therefrom that can be found in all kingdoms of life. Prokaryotic organisms such as cyanobacteria form only simple oxylipin molecules, typically arising from only one or two metabolic steps, whereas eukaryotic organisms such as plants have evolved complex pathways leading to hundreds of different molecules (Wasternack and Feussner 2018). The most renowned and heavily studied oxylipins are indisputably the animal prostaglandins and the plant jasmonates (JAs). In June 2009, Ted Farmer organized an international symposium on regulatory oxylipins at the University of Lausanne (Switzerland). The main focus of that symposium was on oxylipin discovery and signaling and the biology of nonenzymatic lipid oxidation in whole-organism responses to stress. Those days, the field was greatly marked by the fresh discovery of the JAZ repressor proteins as the direct targets of the JA receptor SCFCOI1 E3 ubiquitin ligase, which revealed the long-sought missing link in JA signaling (Browse 2009). Nonetheless, the oxylipin research presented at the 2009 symposium was not limited to JAs and signaling; on the contrary, it covered research on the identification of structurally diverse oxylipins in a wide variety of plant and nonplant species. Ten years later, the understanding of biosynthesis, metabolism, perception and action of oxylipins has dramatically improved, especially that of land plant JAs. By 2019, this was further reinforced at the international meeting on Regulatory Oxylipins held at the VIB-Ghent University in Ghent, Belgium, and organized by Alain Goossens and Ted Farmer (Fig. 1). Rather unintentionally, it was evidenced that JAs have formed the focal point of plant oxylipin research; however, the diversity in JA research has dramatically increased. Indeed, during the conference, the debate arose as to which and how many JA derivatives may actually be bioactive (see the session on ‘Biochemistry and structural biology’). Similarly, the number of model species for plant oxylipin research is increasing rapidly and is now spanning the entire land plant kingdom—from the ancient liverworts to the classic model Arabidopsis thaliana and many major crops (see the session on ‘Regulatory oxylipins in emerging model systems’). Finally, although the role of the JA pathway in the control of one of the major limbs of the plant defense system is becoming further substantiated, it stands beyond doubt that JA can no longer be considered as ‘only’ a stress hormone. Indeed, more and more unexpected roles for JAs in all kinds of plant cellular processes are being revealed, including metabolism, growth, development and multiple forms of interaction with the environment (see the sessions on ‘Control of growth and defence’ and ‘Control of metabolism, reproduction, and ecology’). Accordingly, our deciphering of the molecular mechanisms and signaling cascades that steer these processes is reaching unprecedented levels of detail (see the session on ‘Jasmonate signalling mechanisms and long-distance signalling’), which can be exemplary for other phytohormones that have been studied longer and more extensively. Because of these collective assets, the JA community is clearly expanding and includes vibrant research from groups that have not necessarily started with JA as the focal point but rather serendipitously ‘bumped’ into this inevitable and crucial phytohormone. We were extremely pleased to note that the group of more established JA researchers, several of whom also attended the 2009 symposium, has been complemented by a versatile and energetic group of new (younger) faces. Fig. 1 Open in new tabDownload slide A group photograph taken in front of the Aula of Ghent University in Belgium. Fig. 1 Open in new tabDownload slide A group photograph taken in front of the Aula of Ghent University in Belgium. Below we provide an overview of the seminars given by our (keynote) speakers. For more details, we invite you to consult the already published papers of the speakers, and obviously keep an eye out for the many others that will hopefully be published soon, from which we were privileged to get a sneak preview. Regulatory Oxylipins in Emerging Plant Model Systems In contrast to the 2009 meeting in Lausanne, nonplant systems were not touched upon in the Regulatory Oxylipins 2019 meeting in Ghent. However, the range of species spanning the plant kingdom has greatly expanded this past decade. The power of including ancestral land plants in the study of oxylipin signaling was brilliantly illustrated by our opening keynote speaker, Roberto Solano (Centro Nacional de Biotecnología, Consejo Superior de Investigaciones Científicas, Spain), who has started including the liverwort Marchantia polymorpha, in addition to A. thaliana, in his research. Compared with other sequenced land plants, M. polymorpha exhibits low genetic redundancy in most regulatory pathways (Fig. 2A), which has allowed the identification of the ancestral bioactive JA ligand and clarification of the phylogenetic history and role of JA signaling proteins in plant defense, growth and development (Fig. 2B, C; Monte et al. 2018, Monte et al. 2019). Similarly, Christiane Gatz (Georg-August-University Göttingen, Germany) presented the work of her group on COI1 proteins from leptosporangiate and eusporangiate ferns (horsetails) and their distinct capacities to mediate gene expression dependent on JA-Ile or other oxylipins. Fig. 2 Open in new tabDownload slide Oxylipin signaling in model systems: the ‘new’ kid on the block, Marchantia polymorpha, vs. the ‘old’ lady, Arabidopsis thaliana. (A) Conservation of the JA signaling pathway with a minimal set of components in the plant kingdom. Shown are gene counts for JA signaling components in A. thaliana, M. polymorpha and other plant model species (courtesy of Roberto Solano). (B) M. polymorpha gemmaling expressing a GFP-tagged MpJAZ protein. The inset shows the ligands of MpCOI1, which are two isomeric forms of the JA-Ile precursor dinor-OPDA, i.e. dinor-cis-OPDA (left) and dinor-iso-OPDA (right) (Monte et al. 2018). (C) Arabidopsis thaliana roots expressing a GFP-tagged AtJAZ1 protein. The inset shows the ligand of AtCOI1, which is (+)-7-iso-jasmonoyl-l-isoleucine (Fonseca et al. 2009). Fig. 2 Open in new tabDownload slide Oxylipin signaling in model systems: the ‘new’ kid on the block, Marchantia polymorpha, vs. the ‘old’ lady, Arabidopsis thaliana. (A) Conservation of the JA signaling pathway with a minimal set of components in the plant kingdom. Shown are gene counts for JA signaling components in A. thaliana, M. polymorpha and other plant model species (courtesy of Roberto Solano). (B) M. polymorpha gemmaling expressing a GFP-tagged MpJAZ protein. The inset shows the ligands of MpCOI1, which are two isomeric forms of the JA-Ile precursor dinor-OPDA, i.e. dinor-cis-OPDA (left) and dinor-iso-OPDA (right) (Monte et al. 2018). (C) Arabidopsis thaliana roots expressing a GFP-tagged AtJAZ1 protein. The inset shows the ligand of AtCOI1, which is (+)-7-iso-jasmonoyl-l-isoleucine (Fonseca et al. 2009). Biochemistry and Structural Biology In a series of invited and selected talks, Abraham Koo (University of Missouri, Columbia, SC, USA), Alexander Grechkin (Kazan Scientific Centre of Russian Academy of Sciences, Russia), Thierry Heitz (University of Strasbourg, France) and Lotte Caarls (Wageningen University, The Netherlands) presented their research on the numerous derivatives of JA and JA-Ile that plants can synthesize, particularly using A. thaliana as a model (e.g. Caarls et al. 2017). The enzymes involved in their biosynthesis and the potential role of these metabolites in different cellular processes, such as wound response and immunity, were discussed. Thierry Heitz also provided a good overview of this topic in his mini-review that features in this special issue, entitled ‘When JA-Ile levels do not dictate jasmonate signalling’ (see Heitz et al. 2019). JA Signaling Mechanisms and Control of Growth, Defense and Metabolism Inevitably, a session of the Regulatory Oxylipins 2019 meeting was devoted to signaling. Katayoon Dehesh (Institute for Integrative Genome Biology, USA) gave an inspiring talk about the interplay between oxylipin and retrograde signaling, which is also covered in the review by Savchenko et al. (2019; in this issue). Chuanyou Li (University of Chinese Academy of Sciences, Beijing, China), who has focused much work on transcriptional regulation (e.g. An et al. 2017), provided an impressive in-depth view on the transcriptional machineries involved in JA signaling in Arabidopsis and tomato (Solanum lycopersicum), comprising among others the well-studied MYC2, MED25 and COI1 proteins as well as some newly identified and less characterized transcription factors (TFs). Several of these factors have also been found to be involved in the touch response in Arabidopsis, as presented by Olivier Van Aken (Lund University, Sweden). Similarly, Andrej Pavlovič (Palacký University Olomouc, Czech Republic) showed that the touch response, as a mimic of prey capture, in the carnivorous Venus Flytrap (Dionaea muscipula) is also dependent on JA signaling (e.g. Pavlovič et al. 2017). Intriguingly, Andrej also demonstrated that the prey capture response could be abolished by anesthetic agents. Signaling was also obviously addressed in the sessions on ‘control of growth and defense’ and ‘control of metabolism’ but then more particularly framed within the control of these cellular processes. Daoxin Xie (Tsinghua University, China) elaborated on the molecular basis of JA-regulated plant defense against insect attack, further elucidating the roles of COI1 and the JAZ8–JAV1–WRKY51 complexes (Yan et al. 2018). Debora Gasperini (Leibniz Institute of Plant Biochemistry, Halle, Germany) presented her work on how damage to cell walls triggers JA production, including a promising suppressor screen of the Arabidopsis korrigan cellulase mutant. This theme is further explored in the review by Mielke and Gasperini (2019; in this issue), in which they provide a detailed insight into cell wall-derived damage signals and discuss how they affect JA biosynthesis as well as future prospects in this fascinating area of research. Gregg Howe (Michigan State University, USA) presented his impressive work on the higher order jaz mutants to demonstrate that the JAZ proteins act at the nexus of growth, defense and metabolism (e.g. Guo et al. 2018). Karsten Melcher (Van Andel Research Institute, USA) showed how well-designed structural biology reveals astonishingly detailed molecular insights into the structural basis of JAZ-mediated gene repression in JA signaling, as evidenced by the crystal structures of (truncated) MYC, JAZ, TOPLESS and NINJA proteins. Knowledge gained by studies of JA perception and signaling in Arabidopsis allowed Minoru Ueda (Tohoku University, Japan) to rationally design JAZ subtype-selective agonists of the COI1-JAZ coreceptor, allowing him to uncouple the JA-modulated defense response from the growth arrest. In-depth RNA-Seq analysis of wild-type Arabidopsis plants exogenously treated with JA either in combination with or without salicylic or abscisic acid by Saskia Van der Wees (Utrecht University, The Netherlands) revealed the involvement of new TFs in hitherto undisclosed aspects of JA signaling. This was further supported by work from Ikram Blilou’s group (KAUST, Saudi Arabia) on RNA-Seq analyses of Arabidopsis mutants for the so-called BIRD proteins, which are plant-specific TFs with multiple roles in plant development and physiology. Lastly, there were two plenary talks on the control of metabolism. The first talk, by Daniel Kliebenstein (UC Davis, USA), with the captivating title ‘Destroying the hierarchy; when the regulated are the regulators’, questioned the concept of ‘the master regulator TF’ by demonstrating that Arabidopsis glucosinolate metabolism is likely controlled by several hundred TFs. The second talk, by Xiao-Ya Chen (Shanghai Institutes for Biological Sciences, China), focused on the transcriptional regulation of gossypol biosynthesis in the trichomes of cotton (Gossypium hirsutum). In this issue, Chen et al. (2019) further discuss how JA signaling impacts on specialized metabolism in plants. Ecology JA signaling in a context that represents the complex environment more closely was addressed in the Ecology session. Ian Baldwin (Max Planck Institute for Chemical Ecology, Jena, Germany) gave a thrilling talk on JA signaling as ‘infochemicals’ and mediators of (insect) resistance in nature, in which he presented some of his field work with the native tobacco species Nicotiana attenuata in the Great Basin Desert of Utah and the creation of his pièce de résistance: the MAGIC collection. Corné Pieterse (Utrecht University, The Netherlands) focused on underground events, by elaborating on his research on induced systemic resistance triggered in Arabidopsis by beneficial microbes. Finally, Carlos Ballaré (University of Buenos Aires & San Martin National University, Argentina) talked about the light regulation of JA metabolism and signaling in Arabidopsis (e.g. Cerrudo et al. 2017), which occurs when plants detect and respond to the proximity of competitors using light signals perceived by photoreceptor proteins. Reproduction The last session of the meeting concentrated on reproduction. Bettina Hause (Leibniz Institute of Plant Biochemistry, Halle, Germany) talked about the role of JAs in tomato flower development (e.g. Schubert et al. 2019), whereas Ivan Acosta (Max Planck Institute for Plant Breeding Research, Cologne, Germany) discussed the spatiotemporal dynamics of JA signaling during stamen maturation in Arabidopsis. Finally, the meeting was closed by John Browse (Washington State University, USA), with a marvelously titled seminar of ‘Sex and the single hormone’, in which he gave a magnificent overview of his research on the role of JAs and the JA receptor COI1 in the regulation of male fertility in Arabidopsis and other plants (e.g. Jewell and Browse 2016). More information on this topic can also be found in this special issue, where Ivan Acosta discusses ‘Jasmonate signalling during Arabidopsis stamen maturation’ (Acosta and Przybyl 2019). Funding Organization of the Regulatory Oxylipins 2019 meeting was sponsored by the Facultaire Commissie voor Wetenschappelijk Onderzoek from Ghent University and the journals Plant and Cell Physiology, The Journal of Experimental Botany, The Plant Journal and The Plant Cell. Acknowledgments The organizers wish to thank all of their group members as well as all other staff for helping to deliver an exciting conference both socially and scientifically. We thank Roberto Solano for kindly providing images for Fig. 2. Disclosures The authors have no conflicts of interest to declare. References Acosta I.F. , Przybyl M. ( 2019 ) Jasmonate signaling during Arabidopsis stamen maturation . Plant Cell Physiol . 60 ; 2648–2659 WorldCat An C. , Li L. , Zhai Q. , You Y. , Deng L. , Wu F. , et al. ( 2017 ) Mediator subunit MED25 links the jasmonate receptor to transcriptionally active chromatin . Proc. Natl. Acad. Sci. USA 114 : E8930 – E8939 . Google Scholar Crossref Search ADS WorldCat Browse J. ( 2009 ) Jasmonate passes muster: a receptor and targets for the defense hormone . Annu. Rev. Plant Biol. 60 : 183 – 205 . Google Scholar Crossref Search ADS PubMed WorldCat Caarls L. , Elberse J. , Awwanah M. , Ludwig N.R. , de Vries M. , Zeilmaker T. , et al. ( 2017 ) Arabidopsis JASMONATE-INDUCED OXYGENASES down-regulate plant immunity by hydroxylation and inactivation of the hormone jasmonic acid . Proc. Natl. Acad. Sci. USA 114 : 6388 – 6393 . 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Google Scholar Crossref Search ADS PubMed WorldCat Wasternack C. , Feussner I. ( 2018 ) The oxylipin pathways: biochemistry and function . Annu. Rev. Plant Biol. 69 : 363 – 386 . Google Scholar Crossref Search ADS PubMed WorldCat Yan C. , Fan M. , Yang M. , Zhao J. , Zhang W. , Su Y. , et al. ( 2018 ) Injury activates Ca2+/Calmodulin-dependent phosphorylation of JAV1-JAZ8-WRKY51 complex for jasmonate biosynthesis . Mol. Cell 70 : 136 – 149 . Google Scholar Crossref Search ADS PubMed WorldCat © The Author(s) 2019. Published by Oxford University Press on behalf of Japanese Society of Plant Physiologists. All rights reserved. For permissions, please email: [email protected] This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Savchenko, Tatyana, V;Rolletschek,, Hardy;Dehesh,, Katayoon

doi: 10.1093/pcp/pcz181pmid: 31529102

Abstract The lipid-derived hormones jasmonates (JAs) play key functions in a wide range of physiological and developmental processes that regulate growth, secondary metabolism and defense against biotic and abiotic stresses. In this connection, biosynthesis, tissue-specific distribution, metabolism, perception, signaling of JAs have been the target of extensive studies. In recent years, the involvement of JAs signaling pathway in the regulation of growth and adaptive responses to environmental challenges has been further examined. However, JAs-mediated mechanisms underlying the transition from ‘growth mode’ to ‘adaptive mode’ remain ambiguous. Combined analysis of transgenic lines deficient in JAs signaling in conjunction with the data from JAs-treated plants revealed the function of these hormones in rewiring of central metabolism. The collective data illustrate JAs-mediated decrease in the levels of metabolites associated with active growth such as sucrose, raffinose, orotate, citrate, malate, and an increase in phosphorylated hexoses, responsible for the suppression of growth and photosynthesis, concurrent with the induction of protective metabolites, such as aromatic and branched-chain amino acids, and aspartate family of metabolites. This finding provides an insight into the function of JAs in shifting the central metabolism from the production of growth-promoting metabolites to protective compounds and expands our understanding of the role of JAs in resource allocation in response to environmental challenges. Introduction Oxylipins are lipid-derived metabolites formed in a multi-branch pathway, characterized by their broad spectrum of regulatory functions. The best-studied branch is the ALLENE OXIDE SYNTHASE (AOS) branch responsible for the production of jasmonates (JAs), the plant hormones with a central role in the regulation of plant adaptive responses to changing environmental conditions. JAs are a group of biologically active compounds comprised jasmonic acid (JA), the JA precursor 12-oxophytodienoic acid, together with JA derivatives that function as regulators of plant reproduction, growth, development and defense responses (Wasternack and Hause 2013). Various regulatory functions of JAs are implemented through CORONATINE INSENSITIVE 1 (COI1), a receptor for JA-Isoleucine conjugate, the most active form of JA, and through JASMONATE ZIM DOMAIN (JAZ) proteins, the negative regulators of the JAs-dependent transcriptional network (Sheard et al. 2010, Howe et al. 2018, Pérez‐Salamó et al. 2019). The role of JAs in the regulation of a broad range of defense responses through transcriptional alteration of selected genes under various environmental conditions has been extensively examined. However, as yet, our knowledge of the JAs-dependent regulation of central metabolism has remained fragmentary (van der Fits and Memelink 2000, Broeckling et al. 2005, Machado et al. 2013, Lu et al. 2015, Machado et al. 2015). This review summarizes our current knowledge of JAs’ role in the reprogramming the central metabolism determined by analyses of transgenic lines modified in JAs biosynthesis and signaling pathways together with the data from JAs-treated plants. The collective analysis reveals JAs-mediated reconfiguration of central metabolism which aids in resource reallocation from growth to adaptive processes in response to stress. JAs-Regulated Growth–Defense Trade-off Overall data provide evidence supporting bimodal regulatory functions of JAs by: (i) induction of defense responses via transcriptional modification of genes involved in the biosynthesis of protective metabolites under a broad range of environmental conditions (Brader et al. 2001, Zhao et al. 2005, Major et al. 2017) and (ii) suppression of plant growth (Staswick et al. 1992, Noir et al. 2013, Attaran et al. 2014, Campos et al. 2016, Guo et al. 2018a) as well as individual cell growth and proliferation (Goossens et al. 2003, Bomer et al. 2018). Analyses of JAs-mediated suppression of plant growth in tandem with the induction of adaptive responses at multi-omics level (transcriptomics, proteomics and metabolomics) confirm the function of JAs as a regulatory hub mediating metabolic reprogramming in response to changing environmental conditions. For example, it was demonstrated that in Arabidopsis methyl jasmonate delays the switch from the mitotic cell cycle to the endoreduplication cycle, thereby arresting cells in G1 phase prior to the S-phase transition in a COI1-dependent manner (Noir et al. 2013). Moreover, exogenous treatment of Arabidopsis cell cultures with methyl jasmonate and overexpression of COI1 modify growth by altering cell proliferation and expansion (Bomer et al. 2018). The accumulation of protective metabolites accompanied by reduced cell growth is also observed in JAs-treated Nicotiana tabacum cell cultures (Goossens et al. 2003). However, as yet, the precise mechanisms underlying the JAs-mediated reconfiguration of ‘growth mode’ to ‘tolerance mode’ have remained ambiguous. Specifically, it is not clear whether JAs-mediated suppression of growth in response to stress is solely the result of redistribution of metabolic resources between growth- and defense-related processes. Recent experiments demonstrate the uncoupling of growth suppression and induction of defense responses in Arabidopsis mutant lines deficient both in the five JAZ genes and in phytochrome B, a key regulator of light signaling (Campos et al. 2016). Plants depleted in the five JAZ proteins display constitutive JA signaling and the consequential growth retardation in conjunction with decreased expression of genes involved in growth-related processes, such as ‘response to light stimulus’, ‘cell wall organization’, ‘carbohydrate biosynthetic process’ and ‘lipid biosynthetic process’ (Guo et al. 2018b), while JAZs and phytochrome B-deficient plants maintain high growth rates and improved resistance to insect invaders. However, the suppression of several genes involved in abiotic stress responses in jazs/phyB mutant suggests a partial loss of tolerance to abiotic stresses. In another study, depletion of JASMONIC ACID OXIDASE 2 enhanced Arabidopsis defense against Botrytis cinerea without negative effect on plant growth (Smirnova et al. 2017). Similarly, treatment of tobacco with semisynthetic macrolactone, an analog of JA-Isoleucine, enhanced tolerance to B. cinerea without suppression of growth (Jimenez-Aleman et al. 2017). The regulatory functions of JAs are intertwined with intricate networks of growth-promoting hormones, such as gibberellins (GAs), auxins and brassinosteroids (Huot et al. 2014, Machado et al. 2017, Howe et al. 2018, Koo 2018), and with photosynthetic machinery as the growth-determining process. Several studies have demonstrated JA-mediated suppression of photosynthesis (Weidhase et al. 1987, Bunker et al. 1995, Attaran et al. 2014, Major et al. 2017, Savchenko et al. 2017) mainly ascribed to altered transcription of photosynthesis-related genes. However, the data concerning JAs-mediated reduction of photosystem II efficiency are thus far contradictory (Jung 2004, Attaran et al. 2014, Savchenko et al. 2017). These contradictions could be due to different experimental conditions and JAs level. Regardless, increased CO2 assimilation per leaf area in mutants depleted of MYC2/3/4 transcription factors, positive regulators of JAs responses, supports the regulatory function of JAs in photosynthetic processes (Major et al. 2017). This conclusion is further supported by carbon starvation symptoms and increased respiration in jaz mutant plants (Guo et al. 2018b). It is also noted that alteration of plant anatomical characteristics, such as an increase in the leaf thickness leading to altered leaf mass per area, could be attributed to JA-mediated changes in photosynthetic capacity (Havko et al. 2016). JAs-Mediated Regulation of Carbohydrate Metabolism Carbohydrate levels in plant cell change dynamically, and the content of individual carbohydrates is differentially regulated during growth, development (Rolland et al. 2006), and stress responses (Roitsch 1999). In addition to their role as metabolic currency, soluble sugars also function as signaling molecules regulating gene expression (Jang and Sheen 1994, Xiong et al. 2013, Figueroa and Lunn 2016). As signaling molecules, sugars regulate growth–defense trade-offs through interplay with intrinsic and extrinsic entities such as hormones [abscisic acid (ABA) (Smeekens 2000), ethylene (Zhou et al. 1998), GAs (Perata et al. 1997) and cytokinins (Nemeth et al. 1998)], light signals (Dijkwel et al. 1997, Rolland et al. 2002, Leon and Sheen 2003, Gibson 2004) and energy-sensing pathways (De Vleesschauwer et al. 2018). Several independent studies have established the role of JAs in the regulation of carbohydrates (Hanik et al. 2010, Tytgat et al. 2013, Wang et al. 2014). For example, significantly lower starch concentration was found in tobacco plants impaired in JA signaling (Wang et al. 2014). In addition, exogenous JA application resulted in reduced starch in poplar tree leaves (Babst et al. 2005), decreased soluble sugars in tulip stem (Skrzypek et al. 2005), tobacco (Hanik et al. 2010) and cabbage leaves (Tytgat et al. 2013), and Medicago truncatula cell culture (Broeckling et al. 2005). Recent metabolite profiling of Arabidopsis aos mutant, deficient in the first enzyme of the JAs biosynthesis branch, revealed a significant difference in carbohydrate levels as compared to AOS plants even under standard growth conditions (Savchenko et al. 2019) (Fig. 1). These findings collectively suggest JAs function in the regulation of carbohydrate metabolism and the corresponding physiological ramifications such as affecting plant–insect interaction. This notion is supported by evidence of decreased herbivores’ weight gain when feeding on JAs-deficient plants due to the lower nutritional quality of plants (Machado et al. 2013, Lu et al. 2015), and by altered resistance of Nicotiana attenuata to Manduca sexta through the JAs-mediated regulation of sugars (Machado et al. 2015). Moreover, the control of sugar levels as a key regulatory lever in JAs signaling protects plants against microbial pathogens by inducing pathogenesis-related and antifungal proteins during fruit ripening (Roitsch 1999). Fig. 1 Open in new tabDownload slide Jasmonates-dependent reconfiguration of central metabolism in Arabidopsis. Schematic representation of central metabolism. Metabolites in red boxes are induced by JAs, those noted in green boxes are suppressed by JAs and metabolites, whose levels remain unchanged are noted in back boxes. The data are based on the comparison of metabolic profile of plants possessing functional JA branch of oxylipins and JAs-depleted mutants as the result of the AOS knock out (based on the data from Savchenko et al. 2019). Fig. 1 Open in new tabDownload slide Jasmonates-dependent reconfiguration of central metabolism in Arabidopsis. Schematic representation of central metabolism. Metabolites in red boxes are induced by JAs, those noted in green boxes are suppressed by JAs and metabolites, whose levels remain unchanged are noted in back boxes. The data are based on the comparison of metabolic profile of plants possessing functional JA branch of oxylipins and JAs-depleted mutants as the result of the AOS knock out (based on the data from Savchenko et al. 2019). Sucrose, as a major metabolic currency and as a key signaling compound, controls plant growth, development and stress acclimation. In fact, an increase in sucrose level is one of the common changes in plants during adaptation to environmental stresses. Sucrose-specific sensing and signaling pathways in plants are distinct from those of sucrose degradation products (Smeekens 2000). That is, in general, hexoses promote cell division, while sucrose induces differentiation and maturation (Borisjuk et al. 2002, Borisjuk et al. 2003, Weschke et al. 2003). Sucrose degradation is a central regulatory event in plant carbon metabolism catalyzed by invertase and sucrose synthase. Invertase hydrolyzes sucrose to glucose and fructose, thereby providing monosaccharides to the glycolytic pathway and generating hexose-mediated signals to regulate plant cell metabolism. Subsequently, hexoses are targeted by hexokinase, which forms metabolites that at a low concentration (1–10 mM) repress photosynthetic genes (Jang and Sheen 1994). Phosphorylated hexoses suppress several photosynthetic proteins and reduce chlorophyll content (Jang and Sheen 1994, Pego et al. 1999). Hexokinase is another regulatory hub, functioning as a key sensor and signal transmitter of sugar repression in higher plants. The reduced levels of phosphorylated hexoses observed in JAs-depleted plants may cause enhanced plant growth. In parallel sucrose synthase pathway, sucrose is converted to fructose and UDP-glucose. Sucrose synthase pathway conserves the energy of the sucrose glycoside linkage in the sugar–nucleotide products (UDP-glucose), which in turn is a precursor for the synthesis of starch or cell wall polysaccharides, namely cellulose and callose (Asano et al. 2002, Ruan et al. 2003). Sucrose synthase is ubiquitous in plants and particularly active in sink tissues, such as roots, young leaves, developing seeds or tubers (Koch 2004). Sucrose synthase activity is low in photosynthetic source tissues and high in actively growing sink organs (Rees 1984, Smeekens 2000), in cells with active cell wall deposition, cellulose fibers formation (Ruan et al. 2003) and under anaerobic conditions (Sturm and Tang 1999). The decreased level of UDP-glucose in Arabidopsis aos mutant compared to AOS plants suggests the involvement of JAs in the regulation of sucrose synthase pathway, which is in good agreement with the data supporting the role of JAs in the regulation of sink/source metabolic status (Balcke et al. 2017, Guo et al. 2018b). Moreover, the decreased level of P-gluconate, pentose phosphate pathway-derived metabolites in JAs-depleted plants and induced oxidative pentose phosphate pathway in plants with increased sensitivity to JAs caused by knockout of five JAZ genes in Arabidopsis (Guo et al. 2018b) support the involvement of JAs in the regulation of oxidative pentose phosphate pathway. P-gluconate is the substrate for nucleic acid, lignin, polyphenol and amino acid synthesis (Meyer et al. 2007), and JAs role in regulation of biosynthesis of lignin, a major reinforcer of secondary cell wall formation and a critical structural enabler of long-distance water transport, is demonstrated in Arabidopsis (Denness et al. 2011). It is of note that the oxidative pentose phosphate-dependent sugar-sensing pathway governs the regulation of root nitrogen and sulfur acquisition (Lejay et al. 2008). The JAs-depleted Arabidopsis plants have decreased level of trehalose, a metabolite with compatible solute-like properties that often accumulates in stressed plants, and is perceived by sugar-sensing pathways (Goddijn and van Dun 1999). Instead, the JAs-depleted plants accumulate high level of raffinose and stachyose. Raffinose group oligosaccharides are osmoprotectants, stabilizers of cellular membranes, scavengers of hydroxyl radicals protecting plants from oxidative stress (Nishizawa et al. 2008) and known to accumulate in response to drought, chilling, heat and high-light irradiation, i.e. in response to reactive oxygen species (ROS) generating stresses (Urano et al. 2009). Raffinose accumulation is observed in growing plants (Meyer et al. 2007), and is regulated by the ABA-independent C-repeat binding factors/dehydration-responsive element binding cold-responsive pathway (Cook et al. 2004). The level of myo-inositol, the substrate for raffinose biosynthesis, is also increased in JAs-depleted plants. Besides being a substrate for biosynthesis of oligosaccharides, inositol is an important metabolic and signaling compound (Chen and Xiong 2010) and a substrate for production of cell wall constituents required during plant growth (Roberts and Loewus 1966). Collectively these data establish JAs-mediated modulation of carbohydrate profile involved in the regulation of plant growth and induction of defense responses. JAs-Mediated Alteration of Metabolites of Tricarboxylic Acid Cycle Tricarboxylic acid (TCA) cycle metabolites serve as intermediates in central metabolism and as branching points for the production of secondary metabolites. The JAs-depleted Arabidopsis plants exhibit reduced levels of several glycolytic intermediates, namely 3-phosphoglyceric acid, dihydroxyacetone phosphate, glyceraldehyde 3-phosphate and phosphoenolpyruvate, and simultaneously significantly increased levels of several TCA cycle metabolites, citrate, aconitate, isocitrate, malate and fumarate (Savchenko et al. 2019). Malate through malate shunt could form pyruvate and by extension acetyl-CoA, a key metabolite in de novo fatty acids biosynthesis. Isocitrate and citrate provide carbon skeletons for nitrogen assimilation and reducing equivalents for biosynthetic reactions, supporting the functioning of the glyoxylate cycle and the process of gluconeogenesis (Popova and de Carvalho 1998). The decreased level of isocitrate was observed in plants subjected simultaneously to high light and low temperature, i.e. conditions suppressing photosynthesis and carbon assimilation (Huner et al. 1998). The data support the notion of JAs-mediated suppression of plant growth through reprogramming of central metabolism. Two of the TCA cycle-derived metabolites, oxaloacetate and 2-oxoglutarate, are suppressed in JAs-depleted plants. The level of 2-oxoglutarate, a key regulator in carbon and nitrogen metabolism, is reduced in response to nitrogen starvation (Obata and Fernie 2012). Besides, notable reduction of sulfur-containing metabolites in aos plants corroborates the established connection between the regulatory function of JAs and sulfur metabolism (Park et al. 2002, Jost et al. 2005, Kopriva 2013). The protective functions of sulfur- (Kruse et al. 2007, Mugford et al. 2009, Guo et al. 2018b) and nitrogen-containing secondary metabolites (Ullmann-Zeunert et al. 2013) in plant defense responses have been described. JAs-Mediated Regulation of Amino Acids Amino acids are not only proteins constituents, as they also function as a bridge between primary and secondary metabolism, serve as nitrogen carriers between roots and above-ground plant organs and act as precursors of a number of secondary metabolites with functions in the formation of structural components and defense (Pratelli and Pilot 2014). The levels of most amino acids (except for Gly, Ala, Tyr, Pro and Arg) are reduced in JAs-deficient Arabidopsis plants (Savchenko et al. 2019). Most notable is the reduced levels of aromatic and branched-chain amino acids. Aromatic amino acids, synthesized from phosphoenolpyruvate through the Shikimate pathway, serve as substrates for the formation of secondary metabolites such as lignin, flavonoids, alkaloids, phytoalexins, pigments and the defense hormone salicylic acid, i.e. metabolites with antioxidant property and protective functions (Bailey-Serres et al. 2012). Shikimate pathway as a major consumer of photosynthetically fixed carbon in vascular plants (Vogt 2010) represents an important metabolic hub regulated by JAs. This notion is verified by the accumulation of aromatic amino acids in response to biotic and abiotic stresses (Kim et al. 2007) in conjunction with the transient increase of these amino acids in N. tabacum leaves exogenously treated with JA (Hanik et al. 2010). Similar to aromatic, branched-chain amino acids are substrates for the biosynthesis of a number of secondary metabolites, involved in regulation of GAs and Indole Acetic Acid production (Gao et al. 2009, Parsons et al. 2015) and regulation of gene expression (Kimball and Jefferson 2006, Binder 2010, Obata and Fernie 2012). These amino acids accumulate in drought-stressed (Bowne et al. 2012) and heat-shocked plants (Kaplan et al. 2004). The connection between JAs and biosynthesis of branched-chain and aromatic amino acids has been previously suggested (Balcke et al. 2017, Guo et al. 2018b). In cruciferous, aromatic and branched-chain, amino acids serve as substrates for the biosynthesis of glucosinolates, defensive compounds involved in protection against biotic stresses (Brader et al. 2001), and JA-mediated regulation of glucosinolates accumulation is also established (Guo et al. 2018b). Compared with JAs-producing plants, the JAs-depleted genotypes show a significant decrease in all aspartate family of amino acids, methionine and methionine-related amino acids, together with three amino acids contributing to methionine synthesis (aspartate, serine and cysteine). Also, reduction of 2-oxoglutarate-derived metabolites, including Gln, Glu, ornithine and citrulline in JAs-depleted plants, implicates JAs in regulating the levels of these metabolites. Ornithine and citrulline potentially shuttle nitrate and carbon between mitochondrion and plastids. Levels of these amino acids are increased in response to low temperature (Cook et al. 2004), in concordance with the function of JAs as a regulator of plant responses to low temperature (Hu et al. 2013). Moreover, ornithine is a source of proline synthesis and a ‘gatekeeper’ controlling the biosynthesis of polyamines and gamma-aminobutyric acid (Majumdar et al. 2016). Citrulline is an efficient hydroxyl radical scavenger and a strong antioxidant (Akashi et al. 2001), as such critical to detoxification and elimination of unwanted ammonia within cells (Nelson and Cox 2000). Citrulline accumulation correlates well with plant tolerance to abiotic stress (Yokota et al. 2002). Among amino acids whose levels are positively associated with JAs are Ser, Ile and His (Guo et al. 2018b, Savchenko et al. 2019). Increase in Ser content is a common response to stresses (Rai 2002), and His is an established chelator of toxic ions (Kramer et al. 1996), important for plant reproduction (Stepansky and Leustek 2006). The enhanced His in JA-treated Brassica oleracea confirms the role of JAs in the regulation of His levels (Tytgat et al. 2013). His biosynthesis is a metabolically expensive process, and a decrease in His level in JAs-depleted plants may be the consequence of energy deficiency or altered demands from protein synthesis. Ile plays a special role in JAs signaling, and the majority of JAs-related functions are executed by JA-Ile conjugate (Fonseca et al. 2009). JAs-Mediated Regulation of Antioxidants and Energy Metabolites Ascorbate (Asa) and reduced glutathione (GSH) are major antioxidants present in most plant organelles, mitochondria, chloroplasts and peroxisomes. Asa can react with ROS, such as 1O2, HO·, and act as the substrate for the enzyme Asa peroxidase. GSH is a cellular redox regulator that functions as an ROS scavenger (Foyer and Noctor 2011). The significant decrease of Asa in JAs-depleted Arabidopsis is suggestive of plants experiencing oxidative stress. These data support the previous report, showing upregulation of Asa and GSH metabolic pathways in JAZ-depleted plants displaying constitutive JA responses (Guo et al. 2018b). The other typical features of oxidative stress such as increased levels of sucrose, raffinose, stachyose, myo-inositol and decreased levels of phosphorylated hexoses, fructose-1,6-diphosphate, UDP-glucose, ADP-glucose, p-gluconate, 3-phosphoglyceric acid, altered levels of TCA cycle metabolites, and reduced Asp and Asp-derived amino acids, methionine salvage metabolites, ornithine and citrulline, and low levels of NADP are also observed in JAs-depleted plants (Savchenko et al. 2019). NAD and NADP are two metabolites that play a central role in maintaining plant energy status and redox homeostasis (Hashida et al. 2009). Interestingly, the notable decrease in NADP level in JAs-deficient plants is not accompanied by similar changes in NAD content. NAD is primarily used in respiratory ATP production while NADP is used in reductive biosynthesis and ROS scavenging. It is of note that a decrease in the NAD/NADP ratio is tied directly to photosynthetic activity, at least in cyanobacteria (Tamoi et al. 2005). When higher plants are grown under stressful environments, the reduced form of cytosolic NADP (NADPH) produced in the pentose phosphate pathway plays critical roles in the regulation of ROS (Hashida et al. 2009). The JAs-depleted plants also show reduced levels of other constituents of energy metabolism such as adenosine. In fact, it is reported that adenosine-eliminating enzymes are activated in plants under salt stress conditions (Weretilnyk et al. 2001). Increased level of orotate in aos mutant plants is of special interest because orotate has growth-stimulating activity (Loef et al. 1999, Shopova and Moskova-Simeonova 2000). The ability of JAs to suppress plant growth was documented multiple times, but this observation was not correlated to the modulation of orotate level. Concluding Remarks Plant growth and development are strongly influenced by environmental factors, and as such, in many instances, the adaptation to unfavorable conditions is accompanied by the suppression of growth (Boyer 1982). The two responses to stress, namely growth retardation and enhanced tolerance, stem from complex and intricate adaptive networks aimed at enhancing plant fitness under various conditions, wherein the adaptation process cannot be described as a simple redistribution of resources between growth and protection. From one side, the growth suppression is a common stress avoidance strategy (Maggio et al. 2018), and from the other side, the growth-promoting hormones could aid tailoring protection in response to stress conditions (Smakowska et al. 2016). In general, precise budgeting of the available resources between growth- and defense-related processes is an effective strategy to achieve maximum fitness under conditions of limited resources (Attaran et al. 2014). Here, we portray the role of JAs in modulating the profile of selected central and secondary metabolites with established function in growth regulation and adaptive responses. Metabolites, identified as characteristics of actively growing organs/tissues, including sucrose, raffinose, orotate and TCA cycle metabolites, such as succinate, citrate and malate, are induced in JAs-depleted lines and conversely suppressed in JAs-producing lines or JAs-treated plants. Concurrently, metabolites with protective functions are suppressed in JAs-depleted plants. Collectively, the results provide evidence for JAs-mediated reconfiguration of metabolic profile and lay a foundation for the regulatory role of JAs in the transition from ‘growth mode’ to ‘defense/tolerance mode’ (Fig. 2). Comprehensive profiling of transcripts and metabolic activities will aid us to decipher the molecular and biochemical mechanisms by which JAs mediate regulation of central metabolites. The large-scale analysis of metabolic fluxes is technically still challenging (Williams et al. 2010), but it will provide the most direct link to the observed changes in steady-state metabolite levels in the JA-modulated plants. In general, much remains to be discovered about JAs and their role in cell and organismal physiology, but the rapid progress in this area will define the central role of JAs as the key regulator of physiological processes required for organismal integrity. Fig. 2 Open in new tabDownload slide Simplified model of JAs-mediated shift from ‘growth mode’ to ‘adaptive mode’ (the information is mainly adapted from Huot et al. 2014, Campos et al. 2016, Guo et al. 2018a,b, Noir et al. 2013, Savchenko et al. 2019). Fig. 2 Open in new tabDownload slide Simplified model of JAs-mediated shift from ‘growth mode’ to ‘adaptive mode’ (the information is mainly adapted from Huot et al. 2014, Campos et al. 2016, Guo et al. 2018a,b, Noir et al. 2013, Savchenko et al. 2019). Funding This work was supported by the Russian Science Foundation [grant no 16-14-10155] and Ministry of Education and Science of the Russian Federation [theme AAAA-A17-117030110136-8 to T.V.S.] and National Science Foundation (NSF) IOS-1036491, NSF IOS-1352478 and National Institutes of Health (NIH) [R01GM107311 to K.D]. Disclosures The authors have no conflicts of interest to declare. References Akashi K. , Miyake C. , Yokota A. 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Heitz,, Thierry;Smirnova,, Ekaterina;Marquis,, Valentin;Poirier,, Laure

doi: 10.1093/pcp/pcz172pmid: 31504918

Abstract Regulation of defense and developmental responses by jasmonates (JAs) has been intensively investigated at genetic and transcriptional levels. Plasticity in the jasmonic acid (JA) metabolic pathway as a means to control signal output has received less attention. Although the amplitude of JA responses generally follows the accumulation dynamics of the active hormone jasmonoyl-isoleucine (JA-Ile), emerging evidence has identified cases where this relationship is distorted and that we discuss in this review. JA-Ile is turned over in Arabidopsis by two inducible, intertwined catabolic pathways; one is oxidative and mediated by cytochrome P450 enzymes of the subfamily 94 (CYP94), and the other proceeds via deconjugation by amidohydrolases. Their genetic inactivation has profound effects on JAs homeostasis, including strong JA-Ile overaccumulation, but this correlates with enhanced defense and tolerance to microbial or insect attacks only in the absence of overinduction of negative signaling regulators. By contrast, the impairment of JA oxidation in the jasmonic acid oxidase 2 (jao2) mutant turns on constitutive defense responses without elevating JA-Ile levels in naive leaves and enhances resistance to subsequent biotic stress. This latter and other recent cases of JA signaling are associated with JA-Ile catabolites accumulation rather than more abundant hormone, reflecting increased metabolic flux through the pathway. Therefore, manipulating upstream and downstream JA-Ile homeostatic steps reveals distinct metabolic nodes controlling defense signaling output. Introduction Jasmonates (JAs) regulate major sectors of immune responses and also mediate developmental processes related to growth or fertility (Yuan and Zhang 2015, Campos et al. 2016, Heitz et al. 2016, Wasternack and Song 2017). Induced stress responses constitute useful models to decipher JA metabolism and signaling because they allow to analyze tissues from an initial nonstressed, repressed state through the successive steps leading to full induction of a defended or adapted state (Widemann et al. 2016). JAs belong to the larger family of plant oxygenated fatty acid (FA) derivatives called phytooxylipins that often display signaling activities and/or mediate responses to environmental cues (Wasternack and Feussner 2018). The individual enzymes leading to jasmonic acid (JA) biosynthesis have been extensively and repeatedly reviewed (for recent examples, see Wasternack and Song 2017; Koo 2018; Wasternack and Strnad 2018) and will not be detailed here. JA synthesis is only one of the many possible metabolic routes arising from such lipid-derived precursors and yet JAs play tremendous roles in the adaptation of plants to their ever-changing environment. Their formation is known to be in competition with other metabolic routes branching at several steps in the oxylipin pathway (Wasternack and Feussner 2018). As new enzymatic steps are being elucidated, their functional characterization suggests coordinated and complex control of JA metabolism on hormonal signaling. The present mini-review will focus on the integration of metabolic diversion and elimination mechanisms and their differential impacts on JA signaling. Recent data indicate unexpected positive and negative regulatory loops that will be discussed. Enzymatically generated oxylipins are classified broadly into 9- and 13-derivatives, according to the site of action of lipoxygenase (LOX) on C18 FA. The 13-hydroperoxide of linolenic acid (C18:3) is the main source leading to JA through the allene oxide synthase (AOS)—allene oxide cyclase (AOC) pathway, but this intermediate is also competitively consumed by the hydroperoxide lyase pathway (HPL), and in addition by divinyl ether synthase (DES) in some plants (Fig. 1A). HPL generates C6 or C9 mostly volatile compounds that act in plant–insect interactions (Chehab et al. 2008) and DES produces derivatives that are toxic to microorganisms (Fammartino et al. 2007). All three enzymes belong to the atypical cytochrome P450 subfamily CYP74 (Howe and Schilmiller 2002, Matsui 2006). Genetic studies in rice have shown that HPL-depleted lines overaccumulated JA at the expense of green leaf volatiles and exhibited altered defensive traits along with a lesion mimick phenotype, but the molecular basis of this latter developmental phenotype is unclear (Liu et al. 2012). 13-HOO-FA is usually generated in or within chloroplasts, which offers multiple possibilities for compartmentation and directed substrate consumption. Recent biochemical, protein–protein interactions and structural modeling uncovered the existence of a LOX2–AOS–AOC2 complex excluding concurrent HPL in Arabidopsis chloroplasts (Pollmann et al. 2019). This indicates a supramolecular enzyme organization that may facilitate enzyme catalysis toward the JA branch (Fig. 1A). Determining whether this results from HOO-FA substrate(s) channeling through this complex will need additional biochemical and genetic analyses. Fig. 1 Open in new tabDownload slide Metabolic bifurcations in the JA biochemical pathway generate complexity and impact jasmonate signaling in flowering plants. (A) Jasmonate biosynthesis from C18:3 FA arises when metabolic flux follows required paths at specific branching points. Some compounds in the pathway are substrates of more than one enzyme activity and, therefore, constitute nodes (red dots) through which flux can be diverted toward distinct end products. For example, JA-Ile homeostasis and accumulation is affected upstream by input into the AOS branch, then by the fate of JA, and finally by its own catabolic pathways. The dotted line leading to X refers to additional JA modification routes that are not depicted in the scheme. Enzymes are shown in blue. Compounds indicated in color possess demonstrated biological activity. (B) Biotic stimuli trigger a burst of JA-Ile accumulation that is cleared by CYP94-mediated oxidation or by amidohydrolase-mediated deconjugation. The stimulus-dependent contribution of each catabolic pathway to JA-Ile elimination has been determined using mutants impaired in either one (3cyp, 2ah) or both (5ko) pathways. In these lines, the stress-induced JA profile is strongly modified, showing the severe metabolic impact of mutations. Enhanced defense and resistance to biotic stress is observed when deficient JA-Ile catabolism is not correlated with overinduction of JAZ or JAM genes encoding negative regulators. (C) Soluble JAO enzymes define a direct JA oxidation route to OH-JA. Impairment of JAO2 results in constitutive expression of JA-Ile dependent defense markers in leaves, and strong resistance to subsequent biotic stress. This occurs with a weak metabolic impact without prior JA-Ile elevation, but detectable JA-Ile catabolites before stress indicate increased flux through the pathway. Fig. 1 Open in new tabDownload slide Metabolic bifurcations in the JA biochemical pathway generate complexity and impact jasmonate signaling in flowering plants. (A) Jasmonate biosynthesis from C18:3 FA arises when metabolic flux follows required paths at specific branching points. Some compounds in the pathway are substrates of more than one enzyme activity and, therefore, constitute nodes (red dots) through which flux can be diverted toward distinct end products. For example, JA-Ile homeostasis and accumulation is affected upstream by input into the AOS branch, then by the fate of JA, and finally by its own catabolic pathways. The dotted line leading to X refers to additional JA modification routes that are not depicted in the scheme. Enzymes are shown in blue. Compounds indicated in color possess demonstrated biological activity. (B) Biotic stimuli trigger a burst of JA-Ile accumulation that is cleared by CYP94-mediated oxidation or by amidohydrolase-mediated deconjugation. The stimulus-dependent contribution of each catabolic pathway to JA-Ile elimination has been determined using mutants impaired in either one (3cyp, 2ah) or both (5ko) pathways. In these lines, the stress-induced JA profile is strongly modified, showing the severe metabolic impact of mutations. Enhanced defense and resistance to biotic stress is observed when deficient JA-Ile catabolism is not correlated with overinduction of JAZ or JAM genes encoding negative regulators. (C) Soluble JAO enzymes define a direct JA oxidation route to OH-JA. Impairment of JAO2 results in constitutive expression of JA-Ile dependent defense markers in leaves, and strong resistance to subsequent biotic stress. This occurs with a weak metabolic impact without prior JA-Ile elevation, but detectable JA-Ile catabolites before stress indicate increased flux through the pathway. Once JA is formed, it can be further metabolized through many enzymatic modifications, including hydroxylation, sulfation, glucosylation, conjugation to amino acids or decarboxylation, and for some compounds specialized biological activities were described (Heitz et al. 2016, Widemann et al. 2016, Wasternack and Song 2017, Koo 2018, Wasternack and Strnad 2018). In early studies, JA was considered as the bioactive form because of its ability to induce powerfully broad spectrum responses, hence a number of JA derivatives were considered as potential JA inactivation forms. However, rigorous evidence for loss of activity was generally lacking (Gidda et al. 2003, Miersch et al. 2008), and these notions need to be reassessed in light of recent findings. In all cases, JA being potentially metabolized by different routes, it constitutes a second important regulatory node controlling flux more downstream in the oxylipin pathway (Fig. 1A). Following initial evidence (Staswick and Tiryaki 2004), more than a decade of research has established in numerous plant species that the conjugation of JA to isoleucine by jasmonate-resistant 1 (JAR1) is the critical activation step that generates the master jasmonate signal jasmonoyl-isoleucine (JA-Ile) (Browse 2009). However, the precise factors that determine when and how much of a given JA pool is converted to JA-Ile are still largely unknown. Since the seminal discoveries that JA-Ile perception relies on coronatine insensitive 1 (COI1) recruitment and degradation of jasmonate-zim domain (JAZ) repressors (Chini et al. 2007, Thines et al. 2007), many other protein players were identified that contribute positively or negatively to modulate target gene transcriptional de-repression (Howe et al. 2018). Among those, jasmonate-associated MYC2-like (JAM) proteins also named MYC2-targeted basic helix loop helix (MTBs) are atypical bHLH proteins that compete with MYC transcription factors and restrain gene expression in a negative feedback loop (Nakata et al. 2013; Sasaki-Sekimoto et al. 2013; Liu et al. 2019). In addition, the recent characterization of several catabolic steps in the JA metabolic pathway has provided a framework to investigate the link between hormone formation, accumulation and modification, and actual defense signaling output. Induced vs. Basal JA Levels The current model of JA biosynthesis regulation assumes that most JA biosynthetic enzymes are present at basal levels (Stenzel et al. 2003; Schaller and Stintzi 2009) in nonstimulated tissues. At least in situations involving physical injury like wounding/herbivory or microbial infections, decompartmentation brings in contact cellular components that are normally separated and activates early signaling events. Complex calcium channel patterns are activated both locally and systemically. cyclic nucleotide gated channel19 (CNGC19) is a Ca2+ channel regulating intravascular, herbivory-induced Ca2+ influxes that are essential for inducing JA-Ile biosynthesis and further defense (Meena et al. 2019). Under resting conditions, JA levels are maintained low by a protein complex termed JJW and involving the negative regulator jasmonate-associated VQ motif gene 1 (JAV1), JAZ8 and WRKY51 that targets JA biosynthetic genes. Upon injury, calmodulin-dependent phosphorylation of JAV1 disintegrates JJW, resulting in JA biosynthesis activation (Yan et al. 2018). Long-distance defense signaling to undamaged leaves is mediated by the translocation of mobile signals, including surface involving membrane depolarizations (Mousavi et al. 2013) and calcium waves mediated by glutamate-gated calcium channels of the glutamate-receptor-like (GLR) family (Mousavi et al. 2013). A whole set of novel highly specific reporter and sensor Arabidopsis lines were reported recently and allowed to visualize the sequence and dynamics of rapid long-distance signals, and JA-Ile action sites at cell-type resolution (Larrieu et al. 2015; Nguyen et al. 2018; Toyota et al. 2018). In addition, JAs themselves can be mobile; e.g. OPDA needs to be translocated over long distances to mediate shoot-triggered responses in Arabidopsis roots (Schulze et al. 2019). This adds on an earlier grafting study in tomato indicating that systemic leaf-to-leaf signaling requires both the biosynthesis of JA at the site of wounding and the ability to perceive a jasmonate signal in remote tissues (Schilmiller and Howe 2005). However, how these events relate to the initial lipolytic activities needed to provide the free FA precursors for JA synthesis is unknown. FA-generating enzymes are complex/multigenic and their mode(s) of activation and possible interplay are still poorly understood (Ellinger et al. 2010, Grienenberger et al. 2010, Scherer et al. 2010, Wang et al. 2018). All these data conceptualize signaling cascades triggered by an external cue and that result in local and frequently systemic elevations of JAs content of tissues. Accordingly, most JA biosynthetic genes are JA-inducible (Sasaki et al. 2001), but the well-admitted concept of an amplification loop where JA induces its own synthesis was refuted by feeding experiments (Miersch and Wasternack 2000, Scholz et al. 2015). Therefore, additional regulatory levels must exist and prevent amplified transcripts to result in increased biosynthetic output. By contrast, under resting conditions, undamaged leaves of mature plants contain barely detectable levels of JA (Glauser et al. 2008, Koo et al. 2009, Heitz et al. 2012). This fits well with the current model assuming that JAs, particularly JA-Ile, needs to accumulate sufficiently and act as a ligand to trigger a COI1-dependent destabilization of JAZ repressors and release subsequent JA responses. We review in the following sections two main advances in our understanding of JA hormone homeostasis, and evaluate how genetic modification in post- and pre-JA-Ile metabolic steps have distinct impacts on JA signaling. Recent evidence highlights considerable control of JA-Ile signaling output in both noninduced and induced states: we emphasize that on the one hand basal JA metabolism is not always nonsignificant, and on the other hand that excessive JA-Ile bursts are powerfully counteracted, possibly to prevent runaway defense. JA-Ile Catabolic Circuitry and Its Impact on Signaling To a large extent, in wild-type (WT) plants, jasmonate responses are proportional to the dynamics (timing and amplitude) of JA-Ile accumulation: the higher the hormone builds-up, the more robust the response(s). This relationship has been challenged in recent years by the detailed characterization of two enzymatic JA-Ile catabolic pathways and their study in Arabidopsis leaf defense, which was initiated by the identification of oxidized forms of JA-Ile in wounded leaves (Glauser et al. 2008). The candidate genes encoding JA-Ile catabolic activities are strongly coregulated with JA biosynthetic and signaling genes, facilitating their discovery. The first pathway is defined by cytochrome P450 of the CYP94 family: CYP94B3 and B1 are JA-Ile ω-hydroxylases generating 12OH-JA-Ile, and CYP94C1 catalyzes a more complete JA-Ile oxidation to 12COOH-JA-Ile (Fig. 1A). Initial in vitro pull-down assays and feeding experiments have shown that 12OH-JA-Ile retains some co-receptor-assembling and gene-inducing activities relative to JA-Ile, and that 12COOH-JA-Ile is fully inactive in both assays (Koo et al. 2011, Koo et al. 2014, Aubert et al. 2015). Ectopic overexpression of CYP94 enzymes in different labs led to the consensus findings that JA-Ile oxidation attenuates JA transcriptional responses, weakens resistance to JA-defended insect or fungus attacks and compromises male fertility, due to increased JA-Ile turnover (Koo et al. 2011, Heitz et al. 2012, Koo et al. 2014, Aubert et al. 2015). Loss-of-function Arabidopsis cyp94 mutants have proven more difficult to phenotype due to gene redundancy. Initial analysis readily evidenced that single or double cyp94 mutant lines sustained more persistent JA-Ile accumulation upon wounding, along with depleted oxidation products (Kitaoka et al. 2011, Koo et al. 2011, Heitz et al. 2012). But here, only marginal and transient increase in JA-Ile-responsive gene expression was found, including enhanced expression of some JAZ repressors. A simple explanation would be that in such mutants, at peak accumulation upon wounding, excessive JA-Ile abundance is saturating signaling capacities, and no stronger response can be delivered. However, higher-order mutants provided more complex patterns and point to the existence of hard-wired safeguards. Surprisingly, despite of blocked oxidation steps and high JA-Ile levels, double cyp94b1b3 and triple cyp94b1b3c1 (henceforth called b1b3 and b1b3c1) mutants were reported counterintuitively to display symptoms of deficient JA-Ile signaling, including reduced wound-induced growth inhibition, reduced anthocyanin and increased susceptibility to insects, at odds with the current signaling model (Poudel et al. 2016). To add on this unexpected behavior, in another study, the b1b3c1 line sustained less multiplication of the necrotrophic fungus Botrytis cinerea than WT, despite of the normal JA-Ile accumulation (Aubert et al. 2015). The second JA-Ile catabolic pathway acts by deconjugation through amidohydrolase (AH) activity, first described in Nicotiana attenuata (Woldemariam et al. 2012) and then characterized as the IAR3 and ILL6 enzymes in Arabidopsis (Widemann et al. 2013). IAR3 was known previously as cleaving auxin conjugates and acts thus as a bifunctional enzyme mediating cross-talk between JA and auxin signaling (Zhang et al. 2016). iar3 but not ill6 mutant displayed enhanced wound-induced JA-Ile accumulation, and both lines had higher 12OH-JA-Ile content (Widemann et al. 2013, Koo et al. 2014). This is consistent with IAR3 and ILL6 enzymes also hydrolyzing CYP94-generated 12OH-JA-Ile, defining an indirect biosynthetic route for 12OH-JA formation (Widemann et al. 2013, Zhang et al. 2016). As for CYP94, AH-overexpressing-lines generated symptoms of JA-Ile deficiency (Zhang et al. 2016), conclusively showing that the primary function of both pathways is in the attenuation of JA responses. IAR3- or ILL6-deficient lines have only partially been analyzed in these initial studies that did not investigate their behavior for induced defenses. In addition to gene redundancy within CYP94 and AH families, compensation mechanisms occur when mutant(s) defective in one pathway overinduce genes/enzymes of the other pathway (Widemann et al. 2013). These dispersed and incomplete data have hampered a global understanding of the functions and impacts of JA-Ile catabolism on downstream responses. This problem was recently overcome in a report by Marquis et al. (2019) where higher-order mutant lines, fully deficient in either one (triple cyp94b1b3c1, named 3cyp or double amidohydrolase, named 2ah) or both (quintuple mutant, named 5ko) JA-Ile catabolic pathways were analyzed (Fig. 1B). Striking differences of impacts were found when these lines were submitted either to mechanical wounding or to infection by B. cinerea. CYP94 and AH pathways were found to contribute similarly to shape JA-Ile dynamics upon wounding. In 5ko, JA-Ile accumulated to unprecedented high levels, indicating that hormone elimination was essentially disrupted in this line. In striking contrast, in B. cinerea infected leaves, 3cyp largely suppressed JA-Ile oxidation but with no change in JA-Ile levels relative to WT. This result and the highly enhanced JA-Ile levels in infected 2ah and 5ko leaves showed that, upon fungal attack, the AH pathway predominates to turnover JA-Ile. Yet, in both biological systems, induced defense responses and pathogen/insect resistance poorly correlated with engineered JA-Ile levels: it was found that impaired JA-Ile catabolism results in prolonged defense and increased pest tolerance only if JAZ and JAM genes are not themselves overinduced, and this occurs in a stress- and pathway-specific manner. JAM constitutes an atypical subclass of bHLH factors that compete with and inhibit the action of MYC transcription factors (Nakata et al. 2013, Sasaki-Sekimoto et al. 2013, Liu et al., 2019). In most cases, exaggerated JA-Ile accumulation leads to stronger or more persistent repressor expression, particularly in 5ko, and no ameliorated defense. Only 2ah after wounding, and 3cyp after infection did not display this enhanced feedback expression of JAZ and JAM repressors, and consistently, these lines are more resilient to insect feeding and fungal infection, respectively (Marquis et al. 2019). These data suggest that signaling is under strong negative feedback control in Arabidopsis and indicate that taking advantage of reduced JA-Ile catabolism requires a prior understanding of the mechanisms of this feedback, particularly what factor(s) determine the transcriptional response of the repressors. This does not exclude that other factors (timing, acclimation) might influence the level of pathway output. It is also remarkable that 2ah, 3cyp or 5ko lines exhibit no constitutively elevated JA-Ile or defenses in the absence of stimulation. It is not yet known how these conclusions can be generalized, because in N. attenuata, the inhibition of CYP94 or AH enzymes extends the half-life of JA-Ile as in Arabidopsis, but this results in stronger direct and indirect defense output, culminating in better performance against insect attacks (Woldemariam et al. 2012, Luo et al. 2016). Another complication of the interpretation is that 12OH-JA-Ile, the product of CYP94B enzymes, was recently shown to display significant COI-dependent signaling activity (Poudel et al. 2019). The drastically modified levels of this compound in the mutants discussed above may also alter signaling but no unified conclusion can be drawn. JA-Ile catabolism has been associated with other traits in Poacea, such as abiotic stress tolerance and reproduction. OsCYP94C2 overexpression results in better survival after salt stress in rice (Kurotani et al. 2015), but an allele conferring lower expression of OsCYP94B4 (HAN1), and thus reduced JA-Ile catabolism, was key to rice adaptation to temperate climate during domestication (Mao et al. 2019). Also, a sexual differentiation defect in maize flowers is due to ectopic overexpression of ZmCYP94B1 that interrupts JA signaling in Tassleseed5 (Ts5) mutant (Lunde et al. 2019). The systematic characterization of CYP94 and AH gene families in crop species is just beginning (Hazman et al. 2019) and should disclose further implications of JA-Ile catabolism in the control of JA-regulated processes. JA Flux Diversion as a Means to Control JA-Ile Signaling A recent breakthrough in JA metabolic biology has identified another node of regulation based on JA oxidation to HO-JA, upstream of JA-Ile formation (Fig. 1C). A new subclade of four JA-coregulated genes within the versatile 2-oxoglutarate/Fe(II) dioxygenase (2ODD) family was characterized and named jasmonate-induced oxygenases (JOX; Caarls et al. 2017) or jasmonic acid oxidases (JAO; Smirnova et al. 2017). Similarly to other plant hormonal pathways (Hagel and Facchini 2018), specific 2ODDs have been recruited in JA metabolism to fulfill important functions. All four recombinant JAO/JOX proteins equally oxidize JA to HO-JA, presumably 12OH-JA, but a specific physiological function could be associated with JAO2, owing to its higher basal expression in leaves. Basal JAO2 expression occurs in leaves even in the coi1 perception mutant, whereas JAO wound- and pathogen-induction is fully COI1-dependent. Opposite to manipulated JA-Ile catabolism, impairing JAO2 has little measurable influence on JA profiles, but turns on signaling in a way that strongly enhances biotic stress tolerance. Even with no stimulation, jao2 mutants exhibit constitutive expression of JA-Ile-dependent defenses in leaves that translate into stronger resistance to B. cinerea infection, and a quadruple jox mutant line also displayed less damage after insect feeding (Caarls et al. 2017, Smirnova et al. 2017). The jao2 phenotype requires JA-Ile biosynthesis and signaling, but interestingly, naive leaves do not show increased basal accumulation of the hormone. Instead, JAO2-deficient lines display signs of increased flux through the JA-Ile metabolic pathway, as indicated by an elevated content in JA-Ile catabolites (Smirnova et al. 2017). Therefore, JAO2 defines a metabolic diversion rather than an inactivation mechanism in WT plants where some basal, inactive JA pool is oxidized to 12HO-JA and this contributes to maintain JA-Ile-regulated defense responses repressed to minimal levels in unstressed leaves (Fig. 1C). This important finding reveals that basal JA catabolism does have an important impact on signaling: some JA is constantly formed at trace levels in unstimulated leaves and its JAO2-mediated oxidation is critical to prevent conjugation to JA-Ile and subsequent signaling. JAO/JOX genes are also stress-inducible along with most of the genes of the JA pathway, but the significance of their inducibility has yet to be determined. It will be also of interest to determine if JAO orthologs in crop species have also regulatory functions. JA Signaling without Increase in JA-Ile The notion that JA-Ile signaling can occur without measurable increase in JA-Ile has received wider support recently. Touch-induced changes in Arabidopsis morphology and growth retardation depend on enzymatic gibberellin breakdown (Lange and Lange 2015), but JA signaling is also clearly activated by repeated touching (Chehab et al. 2012), and was found associated with a JA profile reminiscent of active JA-Ile catabolism while JA-Ile levels remained unchanged (M. and T. Lange and T. Heitz, unpublished data). In a recent report, it was shown that short-term exposure of Arabidopsis to gaseous NO2 triggers JA signaling and pathogen resistance, supported by the upregulation of numerous genes in JA metabolism (Mayer et al. 2018). Consistently, examination of the JA profile indicated that JA and JA-Ile levels were not affected by NO2 fumigation, but the catabolites 12OH-JA, 12OH-JA-Ile and 12COOH-JA-Ile were accumulated upon treatment. These cases illustrate that specific genetic lesions or mild stimuli can efficiently turn on JA signaling, with simultaneous JA-Ile biosynthesis and catabolism permitting hormone perception without a rise in its steady-state levels. JA-Ile accumulates generally in response to stronger stimuli when its biosynthesis temporarily exceeds turnover. Future Prospects The recent metabolic studies reviewed here have shed light on new levels of regulation in the JA/JA-Ile pathway. The whole biochemical pathway, starting from the fate of precursor FA, is highly flexible and many branching points can affect the dynamics of JA-Ile formation/accumulation. Manipulation of upstream or downstream JA-Ile homeostatic steps has revealed distinct constraints on hormonal signaling output that are as many challenges for future research. For example, it is unexplained why contrary to JAO2, CYP94 or AH deficiency does not result in constitutive JA signaling. Is 12OH-JA-Ile just a less effective receptor ligand or does it hold peculiar signaling properties? Do microbial pathogens or other aggressors manipulate further steps in the JA pathway (Gimenez-Ibanez et al. 2016)? Fungus-produced 12OH-JA has been proposed to inhibit JA defense signaling in rice by an unknown mechanism (Patkar et al. 2015), but no gene-regulating properties could yet be assigned to this compound in Arabidopsis (Smirnova et al. 2017). It is not known if the JA metabolic grid is complete, and new conversion steps may confer additional complexity. How does JA metabolism impact growth or specific developmental steps? Broader approaches are certainly needed to capture further dimensions of the regulatory network: metabolic fluxes should be better apprehended because snapshot measurements of compounds can be misleading as to the real activity of a pathway. Mass spectrometry-based imaging could also help achieve a better tissular resolution of the spatial distribution of active and inactive JAs and complement hormone transport studies which also reveal new actors and mechanisms (Li et al. 2017, Guan et al. 2019, Schulze et al. 2019). Finally, (sub)-cellular biology of JA compounds is far from being fully understood: e.g. is JA generated by AH activity handled like neosynthetized JA, or are there subpools with different fates? Or, how can COI1 be activated at seemingly constant (basal) JA-Ile levels in the jao2 mutant or in other situations? The field offers many promising opportunities for development, and important novel insights are expected. Funding Work in the T. Heitz group described in this review was supported by basic funding of CNRS [grant ANR-12-BSV8-005] (Jasmonox) from the Agence Nationale de la Recherche to E.S., and IdEx-2014-208e Interdisciplinary grant to L.P. from Université de Strasbourg and CNRS. V.M. is a recipient of a predoctoral fellowship from the Université de Strasbourg and the Ministère de l’Enseignement Supérieur et de la Recherche. Acknowledgments We apologize to colleagues whose work could not be cited due to space limitations. Disclosures The authors have no conflicts of interest to declare. References Aubert Y. , Widemann E. , Miesch L. , Pinot F. , Heitz T. ( 2015 ) CYP94-mediated jasmonoyl-isoleucine hormone oxidation shapes jasmonate profiles and attenuates defence responses to Botrytis cinerea infection . J. Exp. Bot . 66 : 3879 – 3892 . Google Scholar Crossref Search ADS PubMed WorldCat Browse J. ( 2009 ) Jasmonate passes muster: a receptor and targets for the defense hormone . Annu. Rev. Plant Biol. 60 : 183 – 205 . Google Scholar Crossref Search ADS PubMed WorldCat Caarls L. , Elberse J. , Awwanah M. , Ludwig N.R. , de Vries M. , Zeilmaker T. , et al. 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Mielke,, Stefan;Gasperini,, Debora

doi: 10.1093/pcp/pcz119pmid: 31241137

Abstract Plant cell walls are sophisticated carbohydrate-rich structures representing the immediate contact surface with the extracellular environment, often serving as the first barrier against biotic and abiotic stresses. Notably, a variety of perturbations in plant cell walls result in upregulated jasmonate (JA) production, a phytohormone with essential roles in defense and growth responses. Hence, cell wall-derived signals can initiate intracellular JA-mediated responses and the elucidation of the underlying signaling pathways could provide novel insights into cell wall maintenance and remodeling, as well as advance our understanding on how is JA biosynthesis initiated. This Mini Review will describe current knowledge about cell wall-derived damage signals and their effects on JA biosynthesis, as well as provide future perspectives. Introduction Plant cells are surrounded by complex plant cell walls, consisting predominantly of polysaccharides, phenolic compounds and a lower amount of structural proteins (Polko and Kieber 2019). Plant cell walls constitute a decisive evolutionary feature that enabled plants to fulfill a wide range of biological functions that are central to plant life (Popper et al. 2011). They provide mechanical support for growth, enable the adoption of a remarkable variety of cell shapes while securing cellular integrity and tissue cohesiveness, facilitate cellular exchange by increasing contact areas among cells, and can serve as major carbon reservoirs (Polko and Kieber 2019). In addition, their intricated structure protects plant cells and represents the immediate contact interface with the extracellular environment. Contrary to the suggestive concept of a permanent construction invoked by the term ‘wall’, the composition, architecture, properties and functions of plant cell walls are highly dynamic and responsive to both developmental and environmental cues (Lampugnani et al. 2018). Polymer structures in growing plant cells are continuously modified by the synthesis and secretion of new polysaccharides which are then further modified in muro by cell wall residing proteins (Carpita 2011). Moreover, cell wall modification is one of the first signs of pathogen attack and symbiont colonization, and the perception of plant cell wall breakdown products and consequent intracellular signaling is essential for the induction of plant defense responses, reviewed in Bacete et al. (2018). To therefore ensure cell wall integrity (CWI) during expansion and enable modifications following environmental signals, the status of the cell wall is constantly monitored by cell wall surveillance systems that can trigger intracellular signaling in response to cell wall-derived cues (Wolf 2017). However, due to the astonishing structural and biochemical complexity of plant cell walls, the signaling pathways arising from cell wall-derived cues have started to emerge only in the last decade and few molecular components have been identified to date (Wolf 2017, Polko and Kieber 2019). Notably, a variety of perturbations in plant cell walls often lead to altered hormonal responses (Sanchez-Rodriguez et al. 2010, Wolf 2017, Bacete et al. 2018), indicating that cell wall-derived signals are critical to initiate intracellular signaling pathways to coordinate cellular, tissue and whole plant responses. The phytohormone jasmonate (JA), an essential regulator of plant defense and growth responses, can be induced by chemical or genetic perturbations of plant cell walls (Ellis et al. 2002, Sanchez-Rodriguez et al. 2010). However, knowledge on how are cell wall-derived damage signals perceived and transmitted intracellularly to initiate JA production is still completely missing. Furthermore, the roles of JA production and signaling resulting from cell wall damage are still far from being understood. Here, after outlining relevant aspects of plant cell wall composition and its surveillance system, we will highlight recent findings on the interplay between altered plant cell walls and JA production, formulate impending questions and propose future research directions. Plant cell walls Based on structural and functional differences, plant cell walls can be broadly classified into primary and secondary cell walls. Primary cell walls are synthesized during cell division at the cell plate, surround newly formed cells and increase their area when cells expand due to the forces of internal turgor pressure (Lampugnani et al. 2018). They are mainly composed of cellulose, hemicelluloses and pectins. Some plant tissues that have ceased growing may initiate the deposition of secondary walls impregnated with lignin, that give further compressive and tensile strength, and are a hallmark of cell differentiation (Cosgrove and Jarvis 2012). Cellulose is an unbranched, linear polymer of (1→4)-β-D-glucose synthesized by cellulose synthase (CesA) complexes (CSC) at the plasma membrane which use cytosolic UDP-glucose as a substrate (Carpita 2011). For primary cell wall biosynthesis in Arabidopsis, CesA1, CesA3 and CesA6 are necessary to form a functional CSC, and their viable mutant alleles result in stunted growth phenotypes due to impaired cellulose production (Polko and Kieber 2019). Nascent glucan chains organize into para-crystalline cellulose microfibrils that provide high tensile strength necessary to shape cells and reinforce tissues. During cell expansion, the alignment and orientation of cellulose microfibrils is guided by the arrangement of plasma membrane-associated cortical microtubules through their interaction with the CSCs (Polko and Kieber 2019). Cellulose microfibrils are embedded in a hydrated network of hemicelluloses and pectins, which are both synthesized in the Golgi and reach the cell wall through secretory vesicles (Gendre et al. 2013). Unlike cellulose, hemicelluloses are branched polymers, having a cellulose-like backbone decorated with different monosaccharides, such as glucose, xylose, arabinose and mannose, through (1→4)-β-D-links. The most abundant hemicellulose in Arabidopsis is xyloglucan. Other hemicelluloses include glucomannan, galactomannan or arabinoxylan (Pauly and Keegstra 2016). The intricated network of primary cell wall cellulose microfibrils and hemicelluloses is embedded in a gel-like pectin matrix. Pectins are galacturonic acid-containing multiblock polymers consisting of homogalacturonans (HG) and rhamnogalacturonans carrying complex side chains (Atmodjo et al. 2013). The major type of pectin, HG, is delivered into the cell wall in a highly methylesterified, neutral state. After delivery into the extracellular space, HG is de-methyl esterified by pectin methylesterase (PME) with the consequent exposure of negative charges that can form strong cross-links with calcium (Ca2+) (Senechal et al. 2014). These three main polysaccharide classes (cellulose, hemicelluloses and pectins) are interconnected through a variety of different linkages, whose type and function are still emerging (Cosgrove 2016). More than 10% of the Arabidopsis genome, about 2,500 genes, is predicted to be involved in cell wall biosynthesis and its remodeling, highlighting the enormous complexity and challenges in deciphering various aspects of this pivotal plant structure (Carpita 2011). Moreover, plant cell walls have diversified not just between species, but also among cell types, and cell wall microdomains within the same cell (Pattathil et al. 2010). This large heterogeneity of plant tissues and cell types, further hampers the interpretation of biochemical and physical studies of plant cell walls, illustrating the need to study cell wall structures by using a variety of multidisciplinary approaches. Plant cell wall surveillance signaling The existence of a pathway, constantly monitoring the status of plant cell walls was hypothesized based on the well-characterized CWI maintenance system in yeast (Levin 2011). Yeast CWI maintenance involves the perception of the cell wall status by specific osmo-, mechano- and epitope-sensing receptors at the plasma membrane that transduce the signal intracellularly through ionic [Ca2+] changes, Rho1 GTPases and mitogen-activated protein kinases (MAPK). The triggered signaling cascades then initiate appropriate physiological responses such as reorganization of the cytoskeleton, adjustment of cell wall metabolism, and cell cycle progression (Levin 2011). The first genetic evidence for the existence of a plant cell wall sensing system was found when a mutant in THESEUS, a member of the Catharanthus roseus receptor-like kinase1-like protein (CrRLK1L), suppressed the stunted growth phenotype and ectopic lignification of the CesA6 mutant prc1 without restoring cellulose production (Hematy et al. 2007). Ectopic lignification is commonly induced in response to reduced cellulose synthesis (Hamann 2015), supporting the notion that plant cell wall remodeling requires receptors that sense changes in the cell wall and trigger adequate cellular responses that in turn reestablish a new cell wall equilibrium (Wolf 2017). THE1 and other CrRLK1L family members, including FERONIA (FER), CURVY1, HERCULES1/2, ANXUR1/2, BUDDHA’s PAPER SEAL1/2 and ERULUS are plasma membrane proteins with a cytoplasmic serine/threonine kinase domain and an ectodomain exposed towards the cell wall that have been implicated in sensing and transducing cell wall-derived signals (Wolf 2017). Several CrRLK1L are able to bind RAPID ALKALINIZATION FACTOR peptides and thereby regulate immune and growth responses to developmental and environmental cues (Stegmann et al. 2017, Ge et al. 2019). Furthermore, the malectin-like ectodomains in FER can directly bind pectins in vitro, suggesting a role in sensing wall-derived cues and regulating downstream cellular responses (Feng et al. 2018). Recently, a mechanism involving the direct interaction between FER and Extracellular Leucine‐Rich Repeat Extensin (LRX) proteins was found to sense mechanical constraints in the cell wall and consequently drive vacuolar size and cell elongation (Dunser et al. 2019). In addition to CrRLK1Ls, increasing evidence indicates that other receptor-like kinases (RLKs) such as leucine-rich repeat (LRR)-RLK and wall-associated kinases (WAKs), stretch-activated ion channels, highly glycosylated hydroxyproline-rich glycoproteins like arabinogalactans and LRR-extensins play important roles in sensing and transducing cell wall cues across the plasma membrane (Wolf 2017). These different receptors may recognize and respond to a number of potential signals, including binding specific cell wall epitopes or breakdown products, interact with peptides, or sense membrane tension, mechanical stress and strain. Downstream signaling from membrane receptors perceiving cell wall alterations is currently less understood but may involve elements of responses against pathogens such as protein kinases, Ca2+-dependent protein kinases (CDPKs), MAPK and small GTPases to regulate post-translational and transcriptional processes to coordinate cell wall with intracellular responses. It is likely that these cell wall sensing pathways provide the basis for the substantial plasticity of plant cell wall composition (Wolf 2017). Initiation of JA biosynthesis Insect herbivory, necrotrophic pathogen infections and mechanical wounding are very effective in inducing the accumulation of lipid-derived mediators including the hormone precursor JA and the biologically active jasmonoyl-l-isoleucine (JA-Ile), which activates defense responses and inhibits plant growth to promote plant fitness (Howe et al. 2018). JA-Ile is perceived by a co-receptor complex composed of JASMONATE ZIM-DOMAIN (JAZ) repressors and the substrate receptor for an E3 Ubiquitin ligase CORONATINE INSENSITIVE 1 (COI1). As a consequence of JA-Ile perception, JAZ repressors become ubiquitylated and degraded, liberating JA-dependent transcription factors (TFs), such as MYC2, to activate JA-dependent responses (Chini et al. 2007, Thines et al. 2007). Interestingly, all the above-mentioned triggers of JA-Ile biosynthesis involve the disruption of cell walls and release of cellular contents and signaling molecules which may activate directly or indirectly JA biosynthesis. JA-Ile biosynthesis requires a concerted synergism among three subcellular compartments: plastids, peroxisomes and cytosol (Fig. 1), reviewed in Wasternack and Feussner (2018). It initiates from the enzymatic oxygenation of poly unsaturated fatty acids (α-linolenic acid, LeA) present in plastids and further conversion into a cyclic 12-oxo-phytodienoic acid (OPDA) intermediate. OPDA then translocates to peroxisomes for subsequent reduction and β-oxidation to produce JA. Once in the cytosol, JA is conjugated to the bioactive JA-Ile triggering nuclear JA-Ile signaling (Wasternack and Feussner 2018). During male fertility development, a process that is also under JA regulation in Arabidopsis, specific lipases such DEFECTIVE IN ANTHER DEHISCENCE1 (DAD1) release LeA from membrane galactolipids to initiate JA production (Ishiguro et al. 2001). Recently, two abscisic acid (ABA)-regulated PLASTID LIPASEs (PLIP2 and PLIP3) induced JA production and integrated ABA-JA crosstalk (Wang et al. 2018). However, it is still unknown which lipases control LeA release during JA-inducing stimuli such as mechanical wounding. Fig. 1 Open in new tabDownload slide Genetic mutations in specific cell wall biosynthesis and sensing components activate the JA pathway. Mutations in specific components of plant cell walls, which are composed of complex polysaccharides including cellulose (schematically represented by orange rods), hemicelluloses (by purple lines) and pectins (by green ribbons), result in the activation of jasmonoyl-l-isoleucine (JA-Ile) biosynthesis and signaling. JA-Ile biosynthesis initiates in plastids from the enzymatic oxygenation of α-linolenic acid and further conversion into 12-oxophytodienic acid (OPDA). The OPDA precursor then translocates to peroxisomes for several rounds of β-oxidation and conversion into JA. In the cytosol, JA is conjugated to Ile, resulting in the production of bioactive hormone JA-Ile which triggers JA-dependent transcriptional reprogramming in the nucleus. Cell wall biosynthesis and sensing components whose loss of function results in the activation of the JA pathway are highlighted in magenta and include: CELLULASE SYNTHASE 1 and 3 (cesa1, cesa3), KORRIGAN1 (kor1), COBRA (cob), MURUS1 (mur1), UDP-D-GLUCURONATE 4-EPIMERASE 1 and 6 (gae1 gae6), and FERONIA (fer). Nevertheless, intracellular components linking alterations in cell wall biosynthesis to JA-Ile production are still unknown. Those putative signaling pathways could either directly trigger plastidial enzymes to initiate JA production or involve a nuclear transcriptional reprogramming step that then induces JA biosynthesis. Fig. 1 Open in new tabDownload slide Genetic mutations in specific cell wall biosynthesis and sensing components activate the JA pathway. Mutations in specific components of plant cell walls, which are composed of complex polysaccharides including cellulose (schematically represented by orange rods), hemicelluloses (by purple lines) and pectins (by green ribbons), result in the activation of jasmonoyl-l-isoleucine (JA-Ile) biosynthesis and signaling. JA-Ile biosynthesis initiates in plastids from the enzymatic oxygenation of α-linolenic acid and further conversion into 12-oxophytodienic acid (OPDA). The OPDA precursor then translocates to peroxisomes for several rounds of β-oxidation and conversion into JA. In the cytosol, JA is conjugated to Ile, resulting in the production of bioactive hormone JA-Ile which triggers JA-dependent transcriptional reprogramming in the nucleus. Cell wall biosynthesis and sensing components whose loss of function results in the activation of the JA pathway are highlighted in magenta and include: CELLULASE SYNTHASE 1 and 3 (cesa1, cesa3), KORRIGAN1 (kor1), COBRA (cob), MURUS1 (mur1), UDP-D-GLUCURONATE 4-EPIMERASE 1 and 6 (gae1 gae6), and FERONIA (fer). Nevertheless, intracellular components linking alterations in cell wall biosynthesis to JA-Ile production are still unknown. Those putative signaling pathways could either directly trigger plastidial enzymes to initiate JA production or involve a nuclear transcriptional reprogramming step that then induces JA biosynthesis. Because JA biosynthetic enzymes are present under basal conditions and their overexpression does not lead to major increases in basal hormone levels (Bachmann et al. 2002, Sharma et al. 2006) it is presumed that JA-Ile biosynthesis is triggered by the post-transcriptional activation of pre-existing plastidial enzymes. Alternatively, JA-Ile synthesis may initiate from the conversion of existing pools of hormone precursors such as OPDA (Koo et al. 2009). Several intracellular signaling events involved in enzyme activation have been proposed based on parallels with well-studied ligand elicitors from the immunity field, reviewed in Campos et al. (2014). Those include plasma membrane receptors that upon elicitor perception, induce the increase of cytosolic [Ca2+] and apoplastic reactive oxygen species (ROS), leading to the activation of MAPK cascades and CDPKs that finally induce transcriptional reprogramming. Nevertheless, the precise activation mechanisms remain elusive. The involvement of Ca2+ in triggering JA-Ile biosynthesis was hypothesized in several studies (e.g. Scholz et al. 2014, Matschi et al. 2015, Lenglet et al. 2017), and has recently been reinforced by the identification of clade 3 GLUTAMATE RECEPTOR-LIKE (GLR) proteins as regulators of Ca2+ fluxes stimulating distal hormone production (Nguyen et al. 2018, Toyota et al. 2018). Nevertheless, how do [Ca2+] changes induce JA-Ile biosynthesis upon wounding or insect herbivory remains an open question. Furthermore, herbivory-induced [Ca2+] changes activate JA signaling independently of JA-Ile biosynthesis and COI1 perception, through a direct Calmodulin-dependent phosphorylation of JASMONATE-ASSOCIATED VQ MOTIF GENE 1 (JAV1) leading to the disassembly of a JAV1-JAZ8-WRKY51 nuclear repressor complex (Yan et al. 2018). Hence, the interpretation of [Ca2+] signals involved in JA biosynthesis and signaling will require careful evaluation. Plant Cell Wall Alterations Can Trigger the JA Pathway Interestingly, several lines of evidence indicate that cell wall perturbation alone is able to induce JA biosynthesis. Specifically, exogenous applications of cell wall fragments and cellulose inhibitors, cell wall degrading enzymes (CWDEs) and genetic perturbations in plant cell walls result in the activation of the JA pathway (e.g. Ellis et al. 2002, Denness et al. 2011, Engelsdorf et al. 2018). This has led to the hypothesis that cell wall disruption may result in intracellular stress signaling leading to the initiation of JA biosynthesis (Ellis et al. 2002). Moreover, the signaling pathways involved in sensing cell wall alterations and transducing the information intracellularly to initiate JA production in plastids are currently unknown. Unambiguous evidence for the upregulation of JA biosynthesis is ideally provided through quantitative hormone measurements of JA and JA-Ile expressed as molar amount per unit of fresh weight from flash-frozen tissues and/or through the induction of JA-dependent marker genes (Acosta and Farmer 2010). Among these genes are JAZ repressors, the TF MYC2, as well as the JA biosynthesis genes ALLENE OXIDE SYNTHASE, LIPOXYGENASE 2 (LOX2), LOX3, LOX4 and OPDA REDUCTASE 3 (OPR3) (Hickman et al. 2017). Defense-related genes such as VEGETATIVE STORAGE PROTEIN 2 (VSP2) and PLANT DEFENSIN 1.2 (PDF1.2) may be also used, but in combination to hormone measurements and other JA-dependent markers, as in addition to JA their transcripts are regulated by abscisic acid and ethylene, respectively (Penninckx et al. 1998, Anderson et al. 2004). By using these criteria, we will next summarize which plant cell wall alterations result in upregulated JA biosynthesis or signaling (Table 1). Table 1 List of chemically and genetically induced plant cell wall alterations in Arabidopsis that induce the JA pathway Cell wall alteration Evidence of JA pathway activation References Chemical treatments Driselase Increased JA levels Engelsdorf et al. (2018) Pectinase Increased JA levels Engelsdorf et al. (2018) OGs (DP10-DP15) Upregulated AOS and LOX2 expression Moscatiello et al. (2006) OGs (DP12-DP25) Upregulated AOS, LOX3 and LOX4 expression de Azevedo Souza et al. (2017) Cellobiose Upregulated of AOS, LOX3 and LOX4 expression de Azevedo Souza et al. (2017) Isoxaben Increased JA levels Denness et al. (2011),Engelsdorf et al. (2018) Degradation of JA biosensor Jas9-VENUS Larrieu et al. (2015) Upregulated VSP2 expression Ellis et al. (2002) 2, 6-dichlorbenzonitrile (DCB) Upregulated VSP2 expression Ellis et al. (2002) Thaxtomin A Upregulated VSP2 expression Bischoff et al. (2009) Upregulated expression of JAZ1 Duval and Beaudoin (2009) Cell wall mutants (gene) cev1 (CesA3) Ectopic JA levels and upregulated VSP2 levels Ellis et al. (2002) ixr1-1 (CesA3) Ectopic JA levels and upregulated JAZ6, JAZ7, JAZ10, LOX3 and LOX4 expression Engelsdorf et al. (2018) rsw1-1 (CesA1) Upregulated VSP2 expression Ellis et al. (2002) kor1-1 (KOR1) Ectopic JA levels and upregulated LOX3 expression Lopez-Cruz et al. (2014) cob-5 (COB) Upregulated AOS, LOX2 and OPR3 expression Ko et al. (2006) gae1 gae6 (GAE1 and GAE6) Upregulated JAZ5 expression Bethke et al. (2016) mur1 (MUR1) Upregulated JAZ1, JAZ2, JAZ5, JAZ7, JAZ8, JAZ9, JAZ10, LOX2 and OPR3 expression Voxeur et al. (2017) fer1-4 (FER) Upregulated JA-dependent transcriptome Guo et al. (2018) Cell wall alteration Evidence of JA pathway activation References Chemical treatments Driselase Increased JA levels Engelsdorf et al. (2018) Pectinase Increased JA levels Engelsdorf et al. (2018) OGs (DP10-DP15) Upregulated AOS and LOX2 expression Moscatiello et al. (2006) OGs (DP12-DP25) Upregulated AOS, LOX3 and LOX4 expression de Azevedo Souza et al. (2017) Cellobiose Upregulated of AOS, LOX3 and LOX4 expression de Azevedo Souza et al. (2017) Isoxaben Increased JA levels Denness et al. (2011),Engelsdorf et al. (2018) Degradation of JA biosensor Jas9-VENUS Larrieu et al. (2015) Upregulated VSP2 expression Ellis et al. (2002) 2, 6-dichlorbenzonitrile (DCB) Upregulated VSP2 expression Ellis et al. (2002) Thaxtomin A Upregulated VSP2 expression Bischoff et al. (2009) Upregulated expression of JAZ1 Duval and Beaudoin (2009) Cell wall mutants (gene) cev1 (CesA3) Ectopic JA levels and upregulated VSP2 levels Ellis et al. (2002) ixr1-1 (CesA3) Ectopic JA levels and upregulated JAZ6, JAZ7, JAZ10, LOX3 and LOX4 expression Engelsdorf et al. (2018) rsw1-1 (CesA1) Upregulated VSP2 expression Ellis et al. (2002) kor1-1 (KOR1) Ectopic JA levels and upregulated LOX3 expression Lopez-Cruz et al. (2014) cob-5 (COB) Upregulated AOS, LOX2 and OPR3 expression Ko et al. (2006) gae1 gae6 (GAE1 and GAE6) Upregulated JAZ5 expression Bethke et al. (2016) mur1 (MUR1) Upregulated JAZ1, JAZ2, JAZ5, JAZ7, JAZ8, JAZ9, JAZ10, LOX2 and OPR3 expression Voxeur et al. (2017) fer1-4 (FER) Upregulated JA-dependent transcriptome Guo et al. (2018) OGs, oligogalacturonides (pectic breakdown products); DP, degree of polymerization. Open in new tab Table 1 List of chemically and genetically induced plant cell wall alterations in Arabidopsis that induce the JA pathway Cell wall alteration Evidence of JA pathway activation References Chemical treatments Driselase Increased JA levels Engelsdorf et al. (2018) Pectinase Increased JA levels Engelsdorf et al. (2018) OGs (DP10-DP15) Upregulated AOS and LOX2 expression Moscatiello et al. (2006) OGs (DP12-DP25) Upregulated AOS, LOX3 and LOX4 expression de Azevedo Souza et al. (2017) Cellobiose Upregulated of AOS, LOX3 and LOX4 expression de Azevedo Souza et al. (2017) Isoxaben Increased JA levels Denness et al. (2011),Engelsdorf et al. (2018) Degradation of JA biosensor Jas9-VENUS Larrieu et al. (2015) Upregulated VSP2 expression Ellis et al. (2002) 2, 6-dichlorbenzonitrile (DCB) Upregulated VSP2 expression Ellis et al. (2002) Thaxtomin A Upregulated VSP2 expression Bischoff et al. (2009) Upregulated expression of JAZ1 Duval and Beaudoin (2009) Cell wall mutants (gene) cev1 (CesA3) Ectopic JA levels and upregulated VSP2 levels Ellis et al. (2002) ixr1-1 (CesA3) Ectopic JA levels and upregulated JAZ6, JAZ7, JAZ10, LOX3 and LOX4 expression Engelsdorf et al. (2018) rsw1-1 (CesA1) Upregulated VSP2 expression Ellis et al. (2002) kor1-1 (KOR1) Ectopic JA levels and upregulated LOX3 expression Lopez-Cruz et al. (2014) cob-5 (COB) Upregulated AOS, LOX2 and OPR3 expression Ko et al. (2006) gae1 gae6 (GAE1 and GAE6) Upregulated JAZ5 expression Bethke et al. (2016) mur1 (MUR1) Upregulated JAZ1, JAZ2, JAZ5, JAZ7, JAZ8, JAZ9, JAZ10, LOX2 and OPR3 expression Voxeur et al. (2017) fer1-4 (FER) Upregulated JA-dependent transcriptome Guo et al. (2018) Cell wall alteration Evidence of JA pathway activation References Chemical treatments Driselase Increased JA levels Engelsdorf et al. (2018) Pectinase Increased JA levels Engelsdorf et al. (2018) OGs (DP10-DP15) Upregulated AOS and LOX2 expression Moscatiello et al. (2006) OGs (DP12-DP25) Upregulated AOS, LOX3 and LOX4 expression de Azevedo Souza et al. (2017) Cellobiose Upregulated of AOS, LOX3 and LOX4 expression de Azevedo Souza et al. (2017) Isoxaben Increased JA levels Denness et al. (2011),Engelsdorf et al. (2018) Degradation of JA biosensor Jas9-VENUS Larrieu et al. (2015) Upregulated VSP2 expression Ellis et al. (2002) 2, 6-dichlorbenzonitrile (DCB) Upregulated VSP2 expression Ellis et al. (2002) Thaxtomin A Upregulated VSP2 expression Bischoff et al. (2009) Upregulated expression of JAZ1 Duval and Beaudoin (2009) Cell wall mutants (gene) cev1 (CesA3) Ectopic JA levels and upregulated VSP2 levels Ellis et al. (2002) ixr1-1 (CesA3) Ectopic JA levels and upregulated JAZ6, JAZ7, JAZ10, LOX3 and LOX4 expression Engelsdorf et al. (2018) rsw1-1 (CesA1) Upregulated VSP2 expression Ellis et al. (2002) kor1-1 (KOR1) Ectopic JA levels and upregulated LOX3 expression Lopez-Cruz et al. (2014) cob-5 (COB) Upregulated AOS, LOX2 and OPR3 expression Ko et al. (2006) gae1 gae6 (GAE1 and GAE6) Upregulated JAZ5 expression Bethke et al. (2016) mur1 (MUR1) Upregulated JAZ1, JAZ2, JAZ5, JAZ7, JAZ8, JAZ9, JAZ10, LOX2 and OPR3 expression Voxeur et al. (2017) fer1-4 (FER) Upregulated JA-dependent transcriptome Guo et al. (2018) OGs, oligogalacturonides (pectic breakdown products); DP, degree of polymerization. Open in new tab Cell wall degrading enzymes and cell wall fragments as triggers of the JA pathway Phytopathogenic fungi, bacteria and nematodes require breaking the integrity of the host cell wall to infect and complete their life cycle in plant tissues. To breach the cell wall and reach the plasma membrane to initiate infection, they often secrete a variety of CWDEs, including cellulases, hemicellulases (e.g. Xylanases), polygalacturonases (PGs), PMEs and acetyl esterases acting on acetylated pectins or xylans (Bacete et al. 2018). Concurrently, necrotroph infection triggers the JA pathway and thereby the induction of defense responses (Howe et al. 2018). Treatment of Arabidopsis plants with cell culture filtrates of the necrotrophic bacteria Pectobacterium carotovorum, containing secreted CWDEs, induced the expression of JA-dependent genes AOS and VSP2 (Norman-Setterblad et al. 2000). This suggests that CWDEs could be sufficient to activate the JA pathway. However, cell culture filtrates might contain additional pathogen-derived molecules that could also trigger host defense responses (e.g. Zhang et al. 2013). Recently, an isolated mixture of CWDEs from Basidomycetes sp. called Driselase induced the production of JA in treated Arabidopsis seedlings. Further fractionation of CWDEs in Driselase revealed that homogenous preparations of pectinase, but not of cellulase or xylanase induced JA production, suggesting that enzymatic pectin degradation activates JA biosynthesis (Engelsdorf et al. 2018). The enzymatic degradation of the carbohydrate-rich cell wall generates cell wall breakdown fragments. Some of those extracellular, endogenous plant molecules are defined as damage-associated molecular patterns (DAMPs), and can be perceived by membrane receptors and trigger intracellular defense responses (Choi and Klessig 2016). Among the pectic breakdown fragments, α-(1→4)-linked oligogalacturonides (OGs) are potent defense response elicitors and are likely perceived by WAK1 and WAK2 receptors in Arabidopsis (Brutus et al. 2010). However, not all OGs induce JA-mediated defense responses, and OG oligomers with a degree of polymerization (DP) between 9 and 15 showed the most potent intracellular activities (Kohorn 2015). Indeed, treatment with OGs (DP ∼20) triggered JA production in tomato (Doares et al. 1995), and OGs (DP10-15 or DP12-25) elevated the expression of JA-responsive genes AOS, LOX2, LOX3 and LOX4 in both Arabidopsis seedlings and cell cultures (Moscatiello et al. 2006, Souza et al. 2017). However under different experimental conditions, OG treatment did not induce elevated JA levels in Arabidopsis even though pectin degradation triggered JA biosynthesis (Engelsdorf et al. 2018), further highlighting that the DP of OGs is critical for their activity. In addition to pectin oligomers, the cellulose-derived fragment cellobiose also induced JA-responsive genes AOS, LOX3 and LOX4, although its receptor is still unknown (Souza et al. 2017). Cell wall biosynthesis inhibitors inducing JA signaling The inhibitory effect of various herbicides on cell wall biosynthesis has been effectively employed in plant cell wall research. Treatment of Arabidopsis seedlings with the herbicide isoxaben leads to the inhibition of cellulose biosynthesis (Heim et al. 1990) and is therefore regularly used to trigger cellulose deficiency in growing tissues. In addition of depleting CesA3 and CesA6 from the plasma membrane and leading to the disorganization of cortical microtubules, it is unclear if isoxaben has other cellular targets (Paredez et al. 2006). Interestingly, the first evidence indicating that defects in cellulose biosynthesis may lead to the induction of the JA pathway was provided by isoxaben treatment and its consequent upregulation of VSP2 (Ellis et al. 2002). This observation was later corroborated by several studies showing that isoxaben triggers the degradation of the negative JA biosensor Jas9-VENUS (J9V) (Larrieu et al. 2015) and induces high JA levels (Denness et al. 2011, Engelsdorf et al. 2018). Similarly to isoxaben, treatments with other cellulose biosynthesis inhibitors such as 2,6-dichlorbenzonitrile (DCB) and Thaxtomin A trigger the expression of JA-dependent genes VSP2 and JAZ1 (Ellis et al. 2002, Bischoff et al. 2009, Duval and Beaudoin 2009), suggesting that they can also trigger JA biosynthesis (Table 1). Importantly, isoxaben treatment leads to considerable remodeling of cell wall composition, including alterations in hemicelluloses, pectins and ectopic lignification (Denness et al. 2011, Manfield et al. 2004), possibly as compensatory mechanisms to counteract cellulose loss. Furthermore, isoxaben treatment leads to higher salicylic acid levels (Engelsdorf et al. 2018). Hence, it remains unclear if these chemical treatments impacting cellulose production trigger the JA pathway as a direct consequence of cellulose loss, or they do so by secondary effects involving the remodeling of other cell wall components or even through cross-talk with other hormonal pathways. Genetic alterations in plant cell walls resulting in upregulated JA biosynthesis Mutations in cell wall biosynthesis genes involved in the three major cell wall constituents cellulose, pectin and hemicellulose, often lead to impaired production of the respective carbohydrate polymers and severely stunted growth phenotypes (Sanchez-Rodriguez et al. 2010). Additionally, cell wall alterations in one of the cell wall components result in wall alterations in the other two classes (Cano-Delgado et al. 2003), emphasizing the dynamic and tight regulation of cell wall sensing and consequent remodeling. Interestingly, several loss of function mutants in cellulose biosynthesis genes lead to constitutive JA responses (Table 1). The constitutive expression of vsp1 (cev1) mutant allele of CesA3 exhibits high levels of JA, ethylene and defense-related genes like VSP1, VSP2 and PDF1.2 (Ellis et al. 2002). Constitutive expression of these particular defense-related genes was also observed in other mutant alleles of CesA3, ectopic lignification 1 (eli1) (Cano-Delgado et al. 2000) and isoxaben resistant 1 (ixr1) (Hernandez-Blanco et al. 2007). Furthermore, ixr1-1 plants show upregulated basal expression of JA-responsive genes JAZ6, JAZ7, JAZ10, LOX3 and LOX4 as well as elevated JA levels (Engelsdorf et al. 2018). The CesA1 mutant radially swollen 1 (rsw1-1) also exhibits higher basal VSP2 levels (Ellis et al. 2002). In addition to CesAs, mutations in KORRIGAN1 (KOR1), a CesA-interacting membrane protein involved in cellulose biosynthesis (Vain et al. 2014), also showed slightly elevated JA levels at basal conditions, although this elevation was not consistent among all assays (Lopez-Cruz et al. 2014). Nevertheless, upon infection with Botrytis cinerea or Pseudomonas syringae pv tomato, kor1-1 mutants exhibited a higher inducibility of JA and JA-Ile production, as well as a potentiated expression of defense-related genes in comparison to wild-type plants (Finiti et al. 2013, Lopez-Cruz et al. 2014). The glycosylphosphatidylinositol (GPI)-anchored protein COBRA modulates cellulose deposition and oriented cell expansion in roots (Roudier et al. 2005). A loss-of-function allele cobra-5 constitutively expressed JA biosynthesis genes AOS, LOX2 and OPR3 as well as higher basal JA levels (Ko et al. 2006). A double mutant in other GPI-anchored proteins, shaven3 shaven3-like1 (shv3-2 svl1-1), and a mutant in KOBITO 1 (abi8), a glycosyltransferase located in the Golgi, also showed higher basal levels of PDF1.2 and VSP1 (Brocard-Gifford et al. 2004, Hayashi et al. 2008), although hormone levels were not measured. Taken together, the data show that genetic defects in cellulose production impact the JA pathway. In addition to mutants in cellulose biosynthesis, a double mutant in GLUCURONATE 4-EPIMERASE 1 and 6 (gae1 gae6), involved in pectin biosynthesis, exhibited higher basal expression of JAZ5 and a higher inducibility of JA signaling (JAZ5 and JAZ10 expression) upon OG (DP 9–15) and macerozyme treatment (Bethke et al. 2016). Likewise, a mutant in MURUS1 (mur1-1), which is deficient in GDP-L-fucose production and has reduced rhamnogalacturonan-II cross linking, constitutively expressed a battery of JA-dependent genes JAZ1,2,5,7,8,9,10, LOX2 and OPR3 (Voxeur et al. 2017). In agreement with the effects of OGs and pectinase treatment, the available data thus indicate that in addition to cellulose defects, alterations in pectins are able to induce the JA pathway. Nevertheless, as the JA pathway was not systematically assayed in all available cell wall mutants, it is probable that numerous other genes implicated in cell wall biosynthesis and its remodeling regulate JA biosynthesis. A startling example is the finding that fer loss of function mutants accumulate high transcript levels of JA-regulated genes and JA precursors both basally and after isoxaben treatment (Engelsdorf et al. 2018, Guo et al. 2018). Specifically, under basal conditions FER inhibited JA signaling by phosphorylating and destabilizing MYC2, while upon RALF23 elicitation FER stabilized MYC2 and promoted JA-dependent responses (Guo et al. 2018). Thus FER is a negative regulator of JA biosynthesis. Furthermore, while herk1 and herk2 mutations resulted in isoxaben-induced JA overaccumulation as in fer, a the1 loss of function mutant restricted it (Engelsdorf et al. 2018), suggesting that cell wall-derived cues and signaling steps regulating the JA pathway might be multiple. In fact, not all cell wall mutants impact JA-Ile production (Bacete et al. 2018). For example, mutants in CesAs involved in secondary cell wall formation, irx5 and irx1, do not display elevated PDF1.2 expression, and the transcriptome of the growth-impaired friable1 mutant, bearing several cell wall alterations including pectin methylesterification, did not identify the upregulation of JA markers (Hernandez-Blanco et al. 2007, Neumetzler et al. 2012). Hence, induced JA-Ile biosynthesis may be a consequence of specific rather than general alterations of plant cell walls. Role of Upregulated JA Signaling Following Cell Wall Alterations The initiation of JA signaling in response to wounding or herbivory triggers transcriptional changes necessary to activate defense responses at the expense of plant growth (Howe et al. 2018). Furthermore, elevated JA levels inhibit root growth by reducing cell division and cell elongation in the root meristem (Chen et al. 2011). Many cell wall mutations as well as isoxaben treatments lead to growth impairments and concomitant ectopic JA biosynthesis (Table 1). Hence, it is possible that the activation of JA responses resulting from cell wall alterations contribute to hamper plant growth. So far, this has been confirmed in a double mutant between the CesA3 allele cev1 and the JA-insensitive mutant coi1-1 (Ellis et al. 2002). The cev1 coi1-1 double mutant partially restored the cev1 short root phenotype, indicating that ectopic JA signaling can contribute to the growth arrest resulting from cell wall alterations. However, it is not clear if the impact JA on growth can be viewed as a general compensatory mechanism observed after isoxaben treatment and in cell wall mutants or if this a cev1-specific effect. In addition to inducing JA biosynthesis, cell wall alterations often lead to increased levels of other stress-related hormones, reviewed in Sanchez-Rodriguez et al. (2010) and Bacete et al. (2018). For example, treatment with isoxaben, OGs, cellobiose or genetic alterations in CesA3 result in higher salicylic acid or ethylene accumulation (Ellis et al. 2002, Moscatiello et al. 2006, Souza et al. 2017, Engelsdorf et al. 2018). Furthermore, isoxaben treatment increased the production of ROS (Denness et al. 2011). Although partially JA-dependent defense response genes VSP1, VSP2 and PDF1.2 are upregulated in mutant alleles of CesA3 and in wild-type Arabidopsis seedlings treated with isoxaben (Ellis et al. 2002, Cano-Delgado et al. 2003), it is still unclear what is the role of JA (and other hormonal pathways) in the described elevated defense responses of some cell wall mutants. Future efforts should unveil a deeper understanding of the contribution of JA in cell wall-mediated defense responses. Elevated JA levels following cell wall damage may also impact cell wall composition. For example, the observed isoxaben-induced ectopic lignification is potentiated in JA-deficient mutant backgrounds (Denness et al. 2011), suggesting that the activation of the JA pathway contributes to regulate cell wall composition. Furthermore, induced JA signaling resulted in increased expression of cell wall modifying PMEs upon pathogen infection (Bethke et al. 2016). Exogenous JA treatment as well as overexpression of COI1 were also shown to increase the expression of several cell wall modifying enzymes, strengthening the hypothesis that JA signaling may be involved in cell wall remodeling (Bomer et al. 2018). Interestingly, transcriptomic data from ixr1-1 revealed upregulation of only a specific subset of the canonical JA-responsive genes, specifically JAZ6, JAZ7 and JAZ10 (Engelsdorf et al. 2018). This expression pattern is remarkably different when compared with the transcriptional changes occurring after wounding, which leads to the upregulation of almost all JAZ transcripts as well as, e.g. MYC2, AOS, LOX2 and OPR3 (Hickman et al. 2017). Therefore, the characterization of cell wall mutants with respect to the JA pathways might provide the opportunity to uncover specific and more subtle effects of the JA responses that were thus far masked under strong JA-inducing stimuli such as wounding and herbivory. Future Perspectives and Research Directions Deciphering the complex signaling networks arising from cell wall alterations and leading to the upregulation of JA biosynthesis will not only provide crucial insights into how cell wall-derived cues signal intracellularly to activate hormonal responses, but also advance our understanding on how is JA biosynthesis initiated. Although downstream biosynthetic and signaling steps in the JA pathway are well characterized, precise mechanisms of stress perception at the plasma membrane and inter-compartmental transduction events leading to activation of plastidial enzymes are still unknown (Wasternack and Feussner 2018). Putative signaling pathways could involve plasma membrane receptors that sense the status of the cell wall through ligand and epitope binding of either cell wall fragments such as DAMPs or other structures, or membrane tension sensors that could induce ionic fluxes across the plasma membrane of cells with altered cell walls (Campos et al. 2014). Those receptors could transduce the signal intracellularly directly to plastids or first to the nucleus that would then control the activation of plastidial enzymes through transcriptional reprogramming (Fig. 1). Importantly, the initiation of JA biosynthesis following isoxaben treatment is significantly slower (>2 h) compared with the rapid (30 s—few min) JA-Ile induction triggered by wounding (Glauser et al. 2009, Larrieu et al. 2015), suggesting that elicitors and signaling mechanisms might differ between wounding and cell wall damage. Nevertheless, cell wall damage is an inevitable consequence of tissue injury and might be involved in potentiating wound-induced responses over time. Therefore, it will be important to clarify if the activation of JA-Ile biosynthesis following cell wall damage is the result of a direct post-translational activation of JA biosynthesis enzymes or if it requires a nuclear transcriptional reprogramming step. The identification of mutations in MEDIATOR16 (MED16), a tail mediator subunit, as suppressors of growth phenotypes in cob-6 (Sorek et al. 2015), favors a transcriptional activation of JA-Ile biosynthesis which will require further investigation. In addition to ligand- and epitope-based cues from the cell wall, osmotic and mechanic stress at the plasma membrane could be involved in initiating stress responses from altered cell walls. In fact, the addition of an osmoticum (eg. sorbitol or mannitol) putatively balancing the osmotic pressure across the plasma membrane, neutralizes several effects of cellulose biosynthesis inhibitors. For example, combined treatments with isoxaben or driselase with osmotic support, nullified the increased JA levels observed for isoxaben treatment alone (Engelsdorf et al. 2018). An important function of plant cell walls is to counteract the high intracellular turgor pressure during growth and development (Polko and Kieber 2019). Cell wall alterations caused by cellulose biosynthesis inhibitors or cell wall mutants might result in weaker walls that are inefficient in counterbalancing intracellular turgor pressure, leading to mechanical stress at the plasma membrane. Radial turgor pressure changes have already been proposed to be important for wound-induced activation of JA biosynthesis in xylem contact cells (Farmer et al. 2014). Furthermore, (Engelsdorf et al. 2018) found that osmosensitive alterations in the cell wall provoked by isoxaben require the kinases THE1 and FEI2 as well as the ion channel MATING PHEROMONE INDUCED DEATH 1 (MID1)-COMPLEMENTING ACTIVITY 1 (MCA1) in order to activate JA biosynthesis. Hence, elucidating the relationships between osmoregulation and the JA pathways will be another promising research direction. Acknowledgments We apologize to colleagues whose work could not be cited due to space limitations. Funding This work was supported by the Deutsche Forschungsgemeinschaft [grant number GA 2419/2–1]; and by the IPB core funding from the Leibniz Association to D.G. Disclosures The authors have no conflicts of interest to declare. References Acosta I.F. , Farmer E.E. ( 2010 ) Jasmonates . Arabidopsis Book 8 : e0129 . 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Chen,, Xueying;Wang,, Dan-Dan;Fang,, Xin;Chen,, Xiao-Ya;Mao,, Ying-Bo

doi: 10.1093/pcp/pcz161pmid: 31418777

Abstract As sessile and autotrophic organisms, plants have evolved sophisticated pathways to produce a rich array of specialized metabolites, many of which are biologically active and function as defense substances in protecting plants from herbivores and pathogens. Upon stimuli, these structurally diverse small molecules may be synthesized or constitutively accumulated. Jasmonate acids (JAs) are the major defense phytohormone involved in transducing external signals (such as wounding) to activate defense reactions, including, in particular, the reprogramming of metabolic pathways that initiate and enhance the production of defense compounds against insect herbivores and pathogens. In this review, we summarize the progress of recent research on the control of specialized metabolic pathways in plants by JA signaling, with an emphasis on the molecular regulation of terpene and alkaloid biosynthesis. We also discuss the interplay between JA signaling and various signaling pathways during plant defense responses. These studies provide valuable data for breeding insect-proof crops and pave the way to engineering the production of valuable metabolites in future. Introduction As sessile organisms, plants do not have the ability to escape and are restricted to their environment, which may include insect herbivores. In the long evolutionary process, plants have evolved multilayered and efficient systems that influence fitness, which can be divided into constitutive and inductive defenses (Howe and Jander 2008). Once attacked by herbivorous insects or infected by pathogens, plant defense signaling pathways are triggered to reprogram gene expression for the biosynthesis of diverse defense compounds (Browse 2009). Jasmonic acid (JA) and related precursors and derivatives, which are collectively called JAs, play an essential role in plant inductive defense in response to biotic or abiotic stress. As a oxylipin-derived phytohormone, JAs lay the foundation of plant signaling networks as well as immune systems and have been intensively studied during the last decade (Campos et al. 2014, Goossens et al. 2016). Pattern recognition receptors (PRRs) located at the plasma membrane enable plants to recognize attackers-associated patterns of herbivores or pathogens, including microbe/pathogen/herbivore/damage-associated molecular patterns (MAMPs/PAMPs/HAMPs/DAMPs), many of which induce the rapid accumulation of JAs directly or indirectly through mitogen-activated protein kinase cascades, calcium-dependent protein kinases (Romeis and Herde 2014, Zhang et al. 2017), calcium ions and the burst of reactive oxygen species (Fig. 1) (Xia et al. 2015, Choi et al. 2017, Toyota et al. 2018). JA signaling has been shown to mediate plant defense against multiple attackers, especially herbivorous insects, in a variety of ways (Zhang et al. 2017). The biosynthetic pathways that lead to specialized metabolites, especially secondary compounds, such as terpenoids, alkaloids and glucosinolates (GLSs), have been proven to be induced by the JA signaling pathway (De Geyter et al. 2012, Goossens et al. 2017). Additionally, JAs are also required for plant development and reproduction process (Maes et al. 2011, Zhou et al. 2019). JAs directly, or through the interaction with other hormones, modulate the accumulation of specialized metabolites in order to coordinate plant defense and development (Colinas and Goossens 2018, Wasternack and Feussner 2018). Here, we focus on the formation of specialized metabolites induced by JA signaling and how JAs play a role as the coordinator among multiple signaling pathways involved in defense and development. In this mini review, we summarize recent research on certain JA-regulated specialized metabolites by describing the biosynthesis process and the TFs involved (Table 1). We also discuss important strategic functions of the interactions between JAs and other signaling pathways under multistress conditions. Fig. 1 Open in new tabDownload slide Elicitors and effectors influence the core regulation module of JA-mediated defense response. Inductive signals, including DAMPs, HAMPs and MAMPs, are perceived by PRRs at the cell surface to trigger de novo JA biosynthesis. At the resting stage, JAZ proteins block JA signaling by binding to transcription factors including MYCs. JAZs directly recruit corepressors, such as TPL, partially through NINJA. TPL then recruits HDAs that repress JA-response genes through chromatin remodeling. In the presence of JA-Ile, JAZs interact with COI1 to form a coreceptor, which leads to ubiquitination and proteasome-dependent degradation of JAZs, resulting in de-repression of MYCs. By binding with MED25 and recruiting additional coactivators (e.g. HAC1), MYCs recruit RNA polymerase II to form a transcription preinitiation complex that transcripts JA-responsive genes. Since the degradation of JAZ is crucial for activating JA response, it is frequently targeted by effectors of diverse attackers. Pathogen effectors COR, HopZ1a and HopBB1 promote the degradation of JAZs, resulting in enhanced JA signaling and depressed SA signaling. Additionally, viral effector (2b protein and βC1) and whitefly-secreted salivary protein Bt56 repress JA singing pathway in favor of insect performance. Fig. 1 Open in new tabDownload slide Elicitors and effectors influence the core regulation module of JA-mediated defense response. Inductive signals, including DAMPs, HAMPs and MAMPs, are perceived by PRRs at the cell surface to trigger de novo JA biosynthesis. At the resting stage, JAZ proteins block JA signaling by binding to transcription factors including MYCs. JAZs directly recruit corepressors, such as TPL, partially through NINJA. TPL then recruits HDAs that repress JA-response genes through chromatin remodeling. In the presence of JA-Ile, JAZs interact with COI1 to form a coreceptor, which leads to ubiquitination and proteasome-dependent degradation of JAZs, resulting in de-repression of MYCs. By binding with MED25 and recruiting additional coactivators (e.g. HAC1), MYCs recruit RNA polymerase II to form a transcription preinitiation complex that transcripts JA-responsive genes. Since the degradation of JAZ is crucial for activating JA response, it is frequently targeted by effectors of diverse attackers. Pathogen effectors COR, HopZ1a and HopBB1 promote the degradation of JAZs, resulting in enhanced JA signaling and depressed SA signaling. Additionally, viral effector (2b protein and βC1) and whitefly-secreted salivary protein Bt56 repress JA singing pathway in favor of insect performance. The Core Module of JA Signaling JA and related products are synthesized from α-linolenic acid, which is released from plastidial membrane lipids by lipases (Wasternack and Hause 2013). Along with coupled dehydration–cyclization reactions by allene oxide synthase and allene oxide cyclase, the first cyclopentenone oxylipin 12-oxo-phytodienoic acid (OPDA) is generated. After being transported from chloroplasts into the peroxisome, it is reduced to 3-oxo-2–(2′-pentenyl)-cyclopentane-1-octanoic acid (OPC8) by OPDA reductase 3 (OPR3) (Wasternack and Hause 2013). Subsequently, OPC8 is shortened in the carboxylic acid side chain through three cycles of β-oxidation to produce JA, which is then transported into the cytosol. On the other hand, in the OPR3-independent pathway, OPDA directly enters into the β-oxidation pathway to produce 4, 5-ddh-JA, which can be converted into JA through reduction catalyzed by OPR2 (Chini et al. 2018). However, JA itself is inactive, and can be converted into the active form through conjugation with isoleucine (Ile) to form JA-Ile [(+)–7-iso-JA-Ile] by JA-amido synthetase JAR1 (Li et al. 2017). Other modifications of JA include methylation to methyl jasmonate (MeJA) by jasmonic acid carboxyl methyltransferase, leading to increased volatility and capacity to cross membranes (Seo et al. 2001). The release of MeJA is believed to trigger plant–plant communication for defense responses (Wasternack and Hause 2013). In order to balance plant defense and growth, JA signaling is elaborately regulated to avoid overreaction. Recent research proves that after oxidation or deconjugation, JA-Ile forms the derivative 12-hydroxyjasmonic acid (12OH-JA), leading to the suppression of JA defense responses (Widemann et al. 2013, Smirnova et al. 2017). Additionally, other existing JA compounds and their possible functions have been discussed in detail in previous reviews (Wasternack and Hause 2013, Wasternack et al. 2013). Coronatine insensitive1 (COI1), a member of the ubiquitin E3 ligase family, is the core component of the SKP1–CUL1–F-box protein (SCF) E3 ubiquitin ligase complex (SCFCOI1), which is responsible for JA perception and signal transduction (Xie et al. 1998, Sheard et al. 2010). Jasmonate ZIM-domain (JAZ) proteins act as the main repressors in JA signaling pathway. In response to stimuli, surged JA-Ile directly promote the interaction between JAZ proteins and COI1, leading to ubiquitination of JAZ proteins and their degradation via 26S proteasome (Sheard et al. 2010), and subsequently, the degradation of JAZs leads to a de-repression of transcription factors that in turn activates defensive gene expression (Fig. 1). Of the TFs targeted by JAZ proteins, the basic helix–loop–helix (bHLH) transcription factor (TF) MYCs (MYC2, MYC3 and MYC4) are the best studied and are considered primary response regulators of the JA signaling pathway (Kazan and Manners 2013). MYC2 is involved in two branches of the JA-mediated defense process. In wound/insect damage, MYC2 is the transcription factor that synergizes JA and abscisic acid (ABA) pathways by activating the transcription of common downstream genes, such as vegetative storage protein (VSP) (Lorenzo et al. 2004). On the other hand, MYC2 also acts as a linker in JA-ethylene crosstalk in plant–microbe interactions, and regulates the expression of relevant defense genes, such as plant defensin 1.2 (PDF1.2) (Ndamukong et al. 2007). Additionally, MYC2 may directly regulate specialized metabolism-related genes or indirectly (or in synergy) with other downstream transcription factors between JA and other phytohormone signaling pathways during plant defense (Kazan and Manners 2013, Goossens et al. 2016). In contrast to positive regulators such as MYC2, subgroup IIId bHLH factors, including bHLH03/JA-associated MYC2-LIKE 3 (JAM3), bHLH13/JAM2, bHLH14 and bHLH17/JAM1, function as transcriptional repressors. By competitively binding to the target genes of positive TFs, IIId bHLHs attenuate the JA-mediated developmental, defensive processes in case of hypersensitive JA responses (Nakata et al. 2013). On the other hand, a recent study has described the function of MYC2 in regulating the termination of JA signaling, which proceeds by activating the MYC2-targeted bHLH proteins to inhibit the formation of MYC2-MED25 complex (Liu et al. 2019). JA Regulates the Biosynthesis of Specialized Metabolites Terpenoids Plants produce a wide spectrum of specialized metabolites against environmental challenges, and JA signaling plays an important role in the regulation of the biosynthesis of defense substances (Fig. 2). Terpenoids form the largest group of plant metabolites and exert a range of bioactivities in many aspects of plant growth, development, adaptation and defense (Tian et al. 2016). Although our understanding of the ecological function of terpenoids is still primitive, defense roles for some of these compounds, including (E)-β-caryophyllene, (E,E)-farnesol, caulerpenyne, avenacoside B, perforalactone A and gossypol, have been reported. Most of these compounds are sesquiterpens or their derivatives (Fang et al. 2015). Through the 2-C-methyl-d-erythritol 4-phosphate and mevalonate pathway, isopentenyl diphosphate and its isomer, dimethylallyl diphosphate, are produced and function as the precursors of terpenoids, as well as many other isoprenoid-derived primary and specialized metabolites in higher plants (Bohlmann et al. 1998, Burlat et al. 2004). Fig. 2 Open in new tabDownload slide The JA-mediated regulation of biosynthetic pathways of specialized metabolites in plants. Schematic diagram of the regulation of specialized metabolite biosynthesis by JA. The modulated pathways include the biosynthesis of GLSs, sesquiterpenes and anthocyanins in Arabidopsis thaliana, artemisinin in A. annua, terpenoid indole alkaloid (TIA) in C. roseus, nicotine in Nicotiana species and tanshinones and phenolic acids in S. miltiorrhiza. Unbroken lines indicate proven links, whereas broken lines represent hypothetical links. Black arrows indicate synergistic interactions and brown T-bars indicate counteractions. Fig. 2 Open in new tabDownload slide The JA-mediated regulation of biosynthetic pathways of specialized metabolites in plants. Schematic diagram of the regulation of specialized metabolite biosynthesis by JA. The modulated pathways include the biosynthesis of GLSs, sesquiterpenes and anthocyanins in Arabidopsis thaliana, artemisinin in A. annua, terpenoid indole alkaloid (TIA) in C. roseus, nicotine in Nicotiana species and tanshinones and phenolic acids in S. miltiorrhiza. Unbroken lines indicate proven links, whereas broken lines represent hypothetical links. Black arrows indicate synergistic interactions and brown T-bars indicate counteractions. Sesquiterpenes are a subclass of terpenoids that contain three isopentenyl C5-units in their carbon skeleton (Fang et al. 2011). In Arabidopsis, the sesquiterpene synthases, AtTPS21 and AtTPS11, are involved in the production of the vast majority of floral sesquiterpenes (Tholl et al. 2005). AtTPS21 converts FPP into (E)-β-caryophyllene and α-humulene, the predominant component of sesquiterpene volatiles released from flowers, and AtTPS11 is responsible for the formation of essentially all other remaining floral sesquiterpenes, including (+)-thujopsene (Chen et al. 2003). It has been shown that AtMYC2 can directly bind to the promoters of both AtTPS21 and AtTPS11 and activate their expressions. DELLAs, the repressors of the gibberellin (GA) signaling pathway, appear to negatively affect sesquiterpene biosynthesis by directly interacting with MYC2; thus, the transcription factor perceives both JA and GA signals for the formation of floral sesquiterpenes (Hong et al. 2012). Artemisinin is a sesquiterpene lactone that accumulates in glandular trichomes of Artemisia annua. Several gene-encoding enzymes of the artemisinin biosynthetic pathway, including AaADS, AaCYP71AV1, AaDBR2 and AaALDH1, are expressed in glandular trichomes (Zhang et al. 2008). It has been reported that jasmonate signaling can stimulate the biosynthesis of artemisinin and promote the formation of glandular trichomes (Maes et al. 2011). Early investigations have identified that the WRKY transcription factor, AaWRKY1, regulates the artemisinin biosynthetic genes AaADS, AaCYP71AV1 and AaDBR2 through directly binding to the W-box motifs (Ma et al. 2009, Han et al. 2014), and AaWRYK1 can rapidly be induced by MeJA treatment (Ma et al. 2009). More recently, another trichome-specific WRKY transcription factor, AaGSW1, has been shown to positively regulate AaCYP71AV1 expression (Chen et al. 2017). The transcription of AaGSW1 is directly regulated by the JA-responsive AaMYC2, and overexpression of AaMYC2 in A. annua significantly increases the transcription levels of AaCYP71AV1 and AaDBR2, as well as artemisinin content (Shen et al. 2016, Chen et al. 2017). AaERF1 and AaERF2 in A. annua belong to the JA-responsive apetala2/ethylene response factor (AP2/ERF) transcription factor family. Both these proteins can bind to the promoters of AaADS and AaCYP71AV1 to activate gene transcription (Yu et al. 2012b). Furthermore, many other JA-mediated transcription factors, such as those of the ERF (AaORA, AaTAR1), HD-ZIP (AaHD1), LSD and E2F/DP families are also involved in the regulation of artemisinin synthesis (Hao et al. 2017). Gossypol is a dimer of sesquiterpene aldehydes, which specifically accumulate in cotton plants where they are involved in defense mechanisms against pests and pathogens (Tian et al. 2016, Tian et al. 2018). GaWRKY1, a WRKY transcription factor, regulates the expression of the gene encoding (+)-δ-cadinene synthase (CAD or CDN), a sesquiterpene synthase and the first specialized enzyme of the gossypol biosynthetic pathway (Xu et al. 2004). In suspended G. arboretum cells, the expressions of both the GaWRKY1 and CAD1-A genes and the biosynthesis of sesquiterpene aldehydes are strongly induced by a fungal elicitor preparation and by MeJA (Xu et al. 2004). Gossypol and sesquiterpene aldehydes are also constitutively synthesized and stored in pigmented glands of aerial organs and in epidermal and subepidermal layers of roots. A bHLH transcription factor, GoPGF, has been identified to control gland formation (Ma et al. 2016), though further investigation is required to find out whether the regulatory mechanism of PGF is related to JA signaling. The Traditional Chinese Medicinal plant, Salvia miltiorrhiza, contains two types of active components: the diterpenoid tanshinones and the phenolic salvianolic acids (Fang et al. 2017). It has been demonstrated that application of exogenous MeJA enhances the accumulation of tanshinone IIA and salvianolic acid B (Sal B) in S. miltiorrhiza hairy root cultures (Luo et al. 2014). Based on gene expression patterns in response to MeJA treatment, seven bHLH factor genes, especially SmbHLH37, SmbHLH74 and SmbHLH92, were found to be potentially involved in the regulation of tanshinone biosynthesis via JA signaling (Zhang et al. 2015). SmMYC2a and SmMYC2b, which are JA-inducible genes, affect multiple genes in the tanshinone and phenolic acid biosynthesis pathway. Specifically, SmMYC2a is able to bind to an E-box motif within promoters of hydroxy-cinnamoyl transferase 6 (SmHCT6) and cytochrome P450 monooxygenase 98A14 (SmCYP98A14) genes, and SmMYC2b binds to an E-box motif of the SmCYP98A14 promoter. SmHCT6 and SmCYP98A14 are enzymes of the biosynthetic pathway of salvianolic acids, and their gene expressions are also induced by JA (Zhou et al. 2016). Another report showed that the overexpression of either SmJAZ3 or SmJAZ9 represses the JA signaling pathway in S. miltiorrhiza hairy roots and leads to decreased accumulation of tanshinones (Shi et al. 2016). Alkaloids The medical plant Catharanthus roseus produces bioactive terpenoid indole alkaloids (TIA), including the anticancer drugs, vinblastine and vincristine (Courdavault et al. 2014). In general, TIA compounds are generated through the formation of the common alkaloid precursor, strictosidine, which is a condensation product of the indole compound, tryptamine and the monoterpene/iridoid compound, secologanin (Van Moerkercke et al. 2013, Facchini and De Luca 2008). In previous studies, cell suspensions exposed to 50 uM MeJA for 6 h were found to largely induce the expression of most TIA biosynthesis genes in C. roseus (van der Fits and Memelink 2000), which has been confirmed as crucial function of JA elicitation in the TIA biosynthesis pathway (Goossens et al. 2017). The transcription factors octadecanoid responsive catharanthus alkaloid3 (ORCA3) and its homolog, ORCA2, which belong to the AP2/ERFs family, have been identified as positive regulators that activate gene transcriptions which encode enzymes in TIA biosynthesis, including strictosidine synthase (STR) genes (Memelink et al. 2001). Furthermore, the CrORCA gene cluster CrORCA, 3, 4 and 5 are mainly regulated by the bHLH transcription factor, CrMYC2, the main transcription factor in JA signaling. Both CrORCAs and CrMYC2 are activated by phosphorylation, which is mediated by a MAP kinase cascade (Paul et al. 2017). Nevertheless, overexpression of CrORCA3 does not lead to a large accumulation of TIA, because iridoid genes upstream of loganic acid methyltransferase (LAMT) contain eight tightly coexpressed JA-inducible genes, such as geraniol-8-oxidase (G8O), which are not under the control of CrORCA3 (van der Fits and Memelink 2000). JA-regulated bHLH TFs of the clade IVa, iridoid synthesis 1 (BIS1) and BIS2, have been identified to function in a complementary manner to CrORCAs. BIS1 and BIS2 both activate the above-mentioned iridoid genes. Overexpression of either BIS1 or BIS2 specifically activates genes of the iridoid pathway, and results in the accumulation of TIA (Van Moerkercke et al. 2015, Van Moerkercke et al. 2016). Therefore, these JA-regulated TFs can function more accurately and efficiently in controlling TIA biosynthesis through joint regulation, and activate species-specific terpenoid synthesis pathways by recruiting defined cis-regulatory elements, in order to form conserved regulatory modules (Mertens et al. 2016). Nicotine is an alkaloid that specifically accumulates in Nicotiana species. In general, the biosynthesis of nicotine occurs exclusively in roots and is subsequently transported to the aerial parts. As the predominant defense compound in Nicotiana, the accumulation of nicotine largely increases in response to herbivore attacks (Steppuhn et al. 2004). Nicotine is composed of a pyrrolidine ring and a pyridine ring. The structural genes which encode ornithine decarboxylase, putrescine N-methyltransferase (PMT) and arginine decarboxylase are involved in the synthesis of the pyrroline ring, whereas the structural genes which encode aspartate oxidase, quinolinic acid synthase and quinolinate phosphoribosyl transferase are involved in the synthesis of the pyridine ring (Xu et al. 2017). The biosynthesis of nicotine has been studied extensively through genetic analysis and omics research, and JA is considered to play a central role in damage-induced nicotine formation (Kajikawa et al. 2017, Xu et al. 2017). In tobacco, nicotine levels are specifically controlled by two distinct genetic loci: nicotine1 (NIC1) and NIC2. A small group of JA-inducible AP2/ERF TFs, which form a clade within the group IXa subfamily, are clustered at NIC2 locus (Todd et al. 2010). These NIC2-locus ERFs are homologs of C. roseus ORCA3, with functional redundancy and divergence between each other. By recognizing a GCC-box-like element in the promoter of a nicotine biosynthesis gene, the NIC2-locus ERFs specifically activates the most known structural genes in the nicotine pathway (Shoji et al. 2010). ORC1, also known as ERF221, is one of the NIC2-locus ERFs. Under the stimulation of JA, ORC1 positively regulates the biosynthesis of pyridine alkaloid by inducing the expression of genes in all nicotine pathways, and in turn is enhanced by JA-modulated post-translational modifications (De Boer et al. 2011). NtMYC2s in Nicotiana tabacum have been considered to be indispensable to nicotine production, either directly, by activating various nicotine biosynthesis genes or indirectly, by regulating the NIC2-locus ERF TFs (De Boer et al. 2011). In addition, JA-induced bHLH TFs NbbHLH1 and NbbHLH2, which are homologs of MYC2, positively regulate nicotine production in Nicotiana benthamiana (Todd et al. 2010); the synergistic activation mechanism is also controlled in a JAZ-repressive manner. In the absence of JA-Ile production, NtMYC2s are repressed by JAZ proteins, which simultaneously block downstream nicotine biosynthesis in tobacco (De Boer et al. 2011). Glucosinolates GLSs are known as the most important defense compounds in Brassicaceae (Manzaneda et al. 2010). GLSs can be induced by mechanical damage or herbivorous attack via JA signaling. Once attacked by herbivores, the damaged tissue causes the contact between GLSs and myrosinases, which causes the release of toxic products such as nitriles, thiocyanates, isothiocyanates, oxazolidine-2-thiones and epithionitriles. Thus, the GLS–myrosinase system makes plant defense mechanisms more instant and flexible (Bones and Rossiter 2006). The biosynthetic genes of specific groups of GLSs are regulated by a series of R2R3-MYB transcription factors, including MYB28, MYB29 and MYB76 in aliphatic GLSs biosynthesis, whereas MYB34, MYB51 and MYB122 in indole GLSs biosynthesis (Gigolashvili et al. 2007, Sonderby et al. 2010). These R2R3-MYB TFs are regulated by MYC TFs, and the content of GLS is decreased in an extreme manner in the triple mutant myc2myc3myc4 (Schweizer et al. 2013). In addition, evidence shows that MYC2 directly binds to the promoter of several GLSs biosynthesis genes, such as BCAT4, CYP79B3 and SOT16, and induces the de novo synthesis of GLSs, when treated with MeJA (Schweizer et al. 2013). JA Plays a Central Role in the Defense Regulatory Network Plants tend to rely on the coordination between different hormone signaling pathways in order to maintain and adjust efficient defense responses against insects and pathogens. JA is the central regulator in defense response and controls the biosynthesis of specialized metabolites required in this process by interacting with other signaling pathways, such as ABA, salicylic acid (SA), GAs and auxin (IAA) (De Geyter et al. 2012, Verma et al. 2016). SA is the main hormone involved in the regulation of plant resistance to biotrophic and hemibiotrophic pathogens, while JA signaling is triggered upon wounding caused by herbivore attacks or necrotrophic pathogenic infections (Thomma et al. 1998). Usually, SA and JA act antagonistically, based on the type of pathogens or herbivores encountered. When infected by the hemibiotrophic pathogens Pseudomonas syringae. pv. tomato (Pst) DC3000, enhanced SA production leads to reduced resistance to the necrotrophic pathogens Alternaria brassicicola in neighboring tissues (Van der Does et al. 2013, Zhang et al. 2017). On the other hand, exogenous SA treatment (1 mM SA for 24 h) decreased JA production and the JA-mediated defense response against insects. Nevertheless, recent investigations have indicated that during effector-triggered immunity (ETI), SA and JA accumulate at high levels to resist both biotrophs and necrotrophs (Liu et al. 2016). The SA receptors, NPR3 and NPR4, help activate JA response, which leads to positive regulation of RPS2-mediated ETI (Liu et al. 2016). Based on the antagonism between JA and SA, pathogens and insects take advantage of this mechanism to overcome plant resistance during infection. For example, the pathogen P. syringae suppresses SA-dependent defense by inducing JA signaling for successful infection (Campos et al. 2014). It does so by producing Coronatine (COR), which acts as a molecular mimic of JA-Ile (Melotto et al. 2006), and secretes effectors including T3SE HopZ1a and HopBB1 to promote JA signaling by directly interacting with JAZ proteins for degradation (Jiang et al. 2013). Recent reports have revealed that Bt56, a whitefly-secreted salivary protein, can elicit the SA signaling pathway by suppressing JA-dependent defense for better fitness (Xu et al. 2019). The JA Response Is Modulated by Plant Growth and Development Increased plant defense against insect herbivore or pathogen infection is usually accompanied by decreased growth (Guo et al. 2018). Recently, JA signaling has been verified to regulate the equilibrium between immunity and growth, and help reprogram primary metabolism (Colinas and Goossens 2018). The generation of a jaz decuple (jazD) mutant defective in JAZ1-7, -9, -10 and -13 displays high resistance to caterpillar feeding and necrotrophic fungal pathogenic infections, but has a seriously suppressed growth phenotype. Through omics analyses, the data indicated that metabolic pathways, such as the biosynthesis of indole GLSs, are largely induced by jazD (Guo et al. 2018). As the accumulation of excessive metabolites is potentially harmful to plant cells, JAZ-repressive transcriptional hierarchy is important for successful plant growth. In plants, the miR156-SPL module regulates the phase transition from the juvenile to adult stage through targeting a subset of transcription factors named SQUAMOSA PROMOTER BINDING PROTEINLIKE (SPL) (Wu and Poethig 2006). During plant growth, miR156 decays and the level of SPL increases, which regulates multiple aspects of plant growth, development and adaptation (Yu et al. 2015). SPL9 has been found to directly bind to the AtTPS21 promoter and activates its expression, and this regulation is independent of MYC2. Similar regulation exists in the perennial fragrant herb Pogostemon cabilin. By overexpressing SPL10 in patchouli plant, the accumulation of patchouli oil, in which the sesquiterpene alcohol (-)-patchoulol is a predominant component, substantially increases. Consistently, the expression of corresponding patchoulol synthase (PatPTS) is also upregulated by miR156-targeted SPLs in patchouli (Yu et al. 2015). In addition, a recent investigation uncovered that defense responses against insect herbivores are temporally regulated by miR156-targeted SPL (Mao et al. 2017). In plant, SPLs interact with a defined group of JAZ proteins and impair COI1-dependent JAZ degradation. JA response progressively decays with the miR156-SPL-JAZ module. During the young stages, plant has low concentrations of defensive compounds; therefore, active JA response is crucial for defense. During plant growth, defense compounds, such as GLSs in Arabidopsis, are constitutively accumulated and enable adult plants to exert higher resistance against insect herbivores. The age-dependent decay of JA signaling is one strategy that plants can use to balance defense with growth. On the other hand, the phytohormone GA, an important regulator of plant growth and development, has been shown to cross-react with both JA signaling and miR156-SPL-mediated aging pathways by DELLA proteins (Hong et al. 2012, Yu et al. 2012a). GA signaling also functions in the regulation of anthocyanin accumulation by DELLA proteins. Previous data indicate that the MYB/bHLH/WD40 (MBW) complex, a core regulator of anthocyanin biosynthesis, can be a direct target of DELLAs (Qi et al. 2011). In addition, DELLA proteins repress the expression of sesquiterpene synthase genes by directly interacting with MYC2 (Hong et al. 2012). Conclusion and Perspectives Plants synthesize a large number of specialized metabolites, with a large fraction being of great value to humans, such as artemisinin. Generally, the content of these metabolites is low at the stress-free stage. Due to biological activity, their de novo biosynthesis is often strictly regulated. Jasmonate is one of the most important phytohormones in defense signaling network. The most important repressors of the JA signaling pathway, the JAZ proteins, benefit from their structural features and interact with many transcription factors to expand the range of JA-regulated processes, including the biosynthesis of specialized metabolites. The JAZ proteins also act as linkers in crosstalk with other signaling pathways. In the course of co-evolution, pathogens, insects and even viruses have tended to use effectors that target defense hormone signaling for better adaptation to their host plant and JA signaling components appear to be the most common targets of virulence factors from a diverse range of attackers. Further investigation into JA signaling in plant–biotic interactions will provide valuable data for breeding insect-proof crops and will allow the production of valuable metabolites to be engineered. The regulation of the biosynthesis of specialized metabolites does not merely depend on a simple on/off switch; rather, it depends on multiplex signals that are integrated into the defense network. This requires the discovery of new key regulators of the defense network and novel technologies and processes for engineering plant metabolites. Along with increasing omics data, further research on the crosstalk between JA and other signaling pathways will greatly enrich our knowledge of the complex systems that coordinate plant growth, metabolism and defense, and will also help to develop specific plants or plant systems to synthesize targeted metabolites that are of a high value. Funding The Strategic Priority Research Program of Chinese Academy of Sciences [XDB11030000], National Natural Sciences of China [31772177, 31788103], Chinese Academy of Sciences [QYZDY-SSW-SMC026] and The Ministry of Agriculture of China [2016ZX08009001-009]. Table 1 Transcription factors involved in JA-mediated specialized metabolites biosynthesis pathways JA-induced metabolites JA-induced TFs Regulated genes or pathways Plant species Refs. Terpenoids and related derivatives     Volatile terpenes WRKY3 and WRKY6 Upstream of JA biosynthesis Nicotiana attenuata Skibbe e al. (2008)     Gossypol WRKY1 CAD1-A Gossypium arboreum Xu e al. (2004)  Artemisinin WRKY1 CYP71AV1, ADS and DBR2 A. annua Han e al. (2014) and Ma e al. (2009) MYC2 CYP71AV1 and DBR2 A. annua Shen e al. (2016) ERF1 and ERF2 CYP71AV1 and ADS A. annua Yu e al. (2012b) GSW1 CYP71AV1 A. annua Chen e al. (2017)     Ginsenoside WRKY1 Ginsenoside biosynthesis Panax quinquefolius Sun e al. (2013)  Taxol WRKY1 DBAT Taxus chinensis Li e al. (2013) JAMYC1, JAMYC2 and JAMYC4 Paclitaxel biosynthetic pathway Taxus cuspidata Lenka e al. (2015)  Tanshinones and phenolic salvianolic acids MYC2a HCT6 and CYP98A14 S. miltiorrhiza Zhou e al. (2016) MYC2b CYP98A14 S. miltiorrhiza Zhou e al. (2016) Alkaloids  Nicotine NbbHLH1 and NbbHLH2 PMT N. benthamiana Todd e al. (2010) MYC2a/MYC2b PMT1a N. tabacum Zhang e al. (2012) ORC1/ERF221 Nicotine biosynthesis N. tabacum De Boer e al. (2011) MYC2 and ERF189 PMT2 and QPT2 N. tabacum Shoji and Hashimoto (2011)  TIAs ORCA3,4,5 and ORCA2 STR and TDC C. roseus Menke e al. (1999) and Paul e al. (2017) MYC2 TDC C. roseus Paul e al. (2017) MYC1 STR C. roseus Chatel e al. (2003) BIS1 and BIS2 Iridoid biosynthesis C. roseus Van Moerkercke e al. (2016) and Van Moerkercke e al. (2015) ZCT1, ZCT2 and ZCT3 STR and TDC C. roseus Pauw e al. (2004) Flavonoids  Flavonoids MYC2 MYB75/PAP1 and EGL3 A. thaliana Dombrecht e al. (2007) DOF4;2 Produce hydroxycinnamic acid A. thaliana (Skirycz e al. (2007)  Anthocyanins GL3, EGL3, TT8, GL1 and MYB75 Anthocyanins synthesis A. thaliana Qi e al. (2011) MYBL2 TT8 and DFR A. thaliana Xie e al. (2016) MYB115 and MYB134 Proanthocyanidin synthesis Populus James e al. (2017) Phenolic compounds  Phenylpropanoids MYBJS1 PAL and 4CL N. tabacum Galis e al. (2006) MYB8 AT1 and DH29 N. attenuata Onkokesung e al. (2012) Others  GLSs MYC2, MYC3 and MYC4 BCAT4, CYP79B3 and SUR1 A. thaliana Schweizer e al. (2013) MYB34, MYB51 and MYB122 CYP79B2/3, CYP83B1 and SOT16 A. thaliana Frerigmann and Gigolashvili (2014) MYB28, MYB29 and MYB76 Aliphatic GLSs biosynthesis A. thaliana Sonderby e al. (2010)     Camalexin WRKY33 PAD3(CYP71B15) A. thaliana Mao e al. (2011) JA-induced metabolites JA-induced TFs Regulated genes or pathways Plant species Refs. Terpenoids and related derivatives     Volatile terpenes WRKY3 and WRKY6 Upstream of JA biosynthesis Nicotiana attenuata Skibbe e al. (2008)     Gossypol WRKY1 CAD1-A Gossypium arboreum Xu e al. (2004)  Artemisinin WRKY1 CYP71AV1, ADS and DBR2 A. annua Han e al. (2014) and Ma e al. (2009) MYC2 CYP71AV1 and DBR2 A. annua Shen e al. (2016) ERF1 and ERF2 CYP71AV1 and ADS A. annua Yu e al. (2012b) GSW1 CYP71AV1 A. annua Chen e al. (2017)     Ginsenoside WRKY1 Ginsenoside biosynthesis Panax quinquefolius Sun e al. (2013)  Taxol WRKY1 DBAT Taxus chinensis Li e al. (2013) JAMYC1, JAMYC2 and JAMYC4 Paclitaxel biosynthetic pathway Taxus cuspidata Lenka e al. (2015)  Tanshinones and phenolic salvianolic acids MYC2a HCT6 and CYP98A14 S. miltiorrhiza Zhou e al. (2016) MYC2b CYP98A14 S. miltiorrhiza Zhou e al. (2016) Alkaloids  Nicotine NbbHLH1 and NbbHLH2 PMT N. benthamiana Todd e al. (2010) MYC2a/MYC2b PMT1a N. tabacum Zhang e al. (2012) ORC1/ERF221 Nicotine biosynthesis N. tabacum De Boer e al. (2011) MYC2 and ERF189 PMT2 and QPT2 N. tabacum Shoji and Hashimoto (2011)  TIAs ORCA3,4,5 and ORCA2 STR and TDC C. roseus Menke e al. (1999) and Paul e al. (2017) MYC2 TDC C. roseus Paul e al. (2017) MYC1 STR C. roseus Chatel e al. (2003) BIS1 and BIS2 Iridoid biosynthesis C. roseus Van Moerkercke e al. (2016) and Van Moerkercke e al. (2015) ZCT1, ZCT2 and ZCT3 STR and TDC C. roseus Pauw e al. (2004) Flavonoids  Flavonoids MYC2 MYB75/PAP1 and EGL3 A. thaliana Dombrecht e al. (2007) DOF4;2 Produce hydroxycinnamic acid A. thaliana (Skirycz e al. (2007)  Anthocyanins GL3, EGL3, TT8, GL1 and MYB75 Anthocyanins synthesis A. thaliana Qi e al. (2011) MYBL2 TT8 and DFR A. thaliana Xie e al. (2016) MYB115 and MYB134 Proanthocyanidin synthesis Populus James e al. (2017) Phenolic compounds  Phenylpropanoids MYBJS1 PAL and 4CL N. tabacum Galis e al. (2006) MYB8 AT1 and DH29 N. attenuata Onkokesung e al. (2012) Others  GLSs MYC2, MYC3 and MYC4 BCAT4, CYP79B3 and SUR1 A. thaliana Schweizer e al. (2013) MYB34, MYB51 and MYB122 CYP79B2/3, CYP83B1 and SOT16 A. thaliana Frerigmann and Gigolashvili (2014) MYB28, MYB29 and MYB76 Aliphatic GLSs biosynthesis A. thaliana Sonderby e al. (2010)     Camalexin WRKY33 PAD3(CYP71B15) A. thaliana Mao e al. (2011) Open in new tab Table 1 Transcription factors involved in JA-mediated specialized metabolites biosynthesis pathways JA-induced metabolites JA-induced TFs Regulated genes or pathways Plant species Refs. Terpenoids and related derivatives     Volatile terpenes WRKY3 and WRKY6 Upstream of JA biosynthesis Nicotiana attenuata Skibbe e al. (2008)     Gossypol WRKY1 CAD1-A Gossypium arboreum Xu e al. (2004)  Artemisinin WRKY1 CYP71AV1, ADS and DBR2 A. annua Han e al. (2014) and Ma e al. (2009) MYC2 CYP71AV1 and DBR2 A. annua Shen e al. (2016) ERF1 and ERF2 CYP71AV1 and ADS A. annua Yu e al. (2012b) GSW1 CYP71AV1 A. annua Chen e al. (2017)     Ginsenoside WRKY1 Ginsenoside biosynthesis Panax quinquefolius Sun e al. (2013)  Taxol WRKY1 DBAT Taxus chinensis Li e al. (2013) JAMYC1, JAMYC2 and JAMYC4 Paclitaxel biosynthetic pathway Taxus cuspidata Lenka e al. (2015)  Tanshinones and phenolic salvianolic acids MYC2a HCT6 and CYP98A14 S. miltiorrhiza Zhou e al. (2016) MYC2b CYP98A14 S. miltiorrhiza Zhou e al. (2016) Alkaloids  Nicotine NbbHLH1 and NbbHLH2 PMT N. benthamiana Todd e al. (2010) MYC2a/MYC2b PMT1a N. tabacum Zhang e al. (2012) ORC1/ERF221 Nicotine biosynthesis N. tabacum De Boer e al. (2011) MYC2 and ERF189 PMT2 and QPT2 N. tabacum Shoji and Hashimoto (2011)  TIAs ORCA3,4,5 and ORCA2 STR and TDC C. roseus Menke e al. (1999) and Paul e al. (2017) MYC2 TDC C. roseus Paul e al. (2017) MYC1 STR C. roseus Chatel e al. (2003) BIS1 and BIS2 Iridoid biosynthesis C. roseus Van Moerkercke e al. (2016) and Van Moerkercke e al. (2015) ZCT1, ZCT2 and ZCT3 STR and TDC C. roseus Pauw e al. (2004) Flavonoids  Flavonoids MYC2 MYB75/PAP1 and EGL3 A. thaliana Dombrecht e al. (2007) DOF4;2 Produce hydroxycinnamic acid A. thaliana (Skirycz e al. (2007)  Anthocyanins GL3, EGL3, TT8, GL1 and MYB75 Anthocyanins synthesis A. thaliana Qi e al. (2011) MYBL2 TT8 and DFR A. thaliana Xie e al. (2016) MYB115 and MYB134 Proanthocyanidin synthesis Populus James e al. (2017) Phenolic compounds  Phenylpropanoids MYBJS1 PAL and 4CL N. tabacum Galis e al. (2006) MYB8 AT1 and DH29 N. attenuata Onkokesung e al. (2012) Others  GLSs MYC2, MYC3 and MYC4 BCAT4, CYP79B3 and SUR1 A. thaliana Schweizer e al. (2013) MYB34, MYB51 and MYB122 CYP79B2/3, CYP83B1 and SOT16 A. thaliana Frerigmann and Gigolashvili (2014) MYB28, MYB29 and MYB76 Aliphatic GLSs biosynthesis A. thaliana Sonderby e al. (2010)     Camalexin WRKY33 PAD3(CYP71B15) A. thaliana Mao e al. (2011) JA-induced metabolites JA-induced TFs Regulated genes or pathways Plant species Refs. Terpenoids and related derivatives     Volatile terpenes WRKY3 and WRKY6 Upstream of JA biosynthesis Nicotiana attenuata Skibbe e al. (2008)     Gossypol WRKY1 CAD1-A Gossypium arboreum Xu e al. (2004)  Artemisinin WRKY1 CYP71AV1, ADS and DBR2 A. annua Han e al. (2014) and Ma e al. (2009) MYC2 CYP71AV1 and DBR2 A. annua Shen e al. (2016) ERF1 and ERF2 CYP71AV1 and ADS A. annua Yu e al. (2012b) GSW1 CYP71AV1 A. annua Chen e al. (2017)     Ginsenoside WRKY1 Ginsenoside biosynthesis Panax quinquefolius Sun e al. (2013)  Taxol WRKY1 DBAT Taxus chinensis Li e al. (2013) JAMYC1, JAMYC2 and JAMYC4 Paclitaxel biosynthetic pathway Taxus cuspidata Lenka e al. (2015)  Tanshinones and phenolic salvianolic acids MYC2a HCT6 and CYP98A14 S. miltiorrhiza Zhou e al. (2016) MYC2b CYP98A14 S. miltiorrhiza Zhou e al. (2016) Alkaloids  Nicotine NbbHLH1 and NbbHLH2 PMT N. benthamiana Todd e al. (2010) MYC2a/MYC2b PMT1a N. tabacum Zhang e al. (2012) ORC1/ERF221 Nicotine biosynthesis N. tabacum De Boer e al. (2011) MYC2 and ERF189 PMT2 and QPT2 N. tabacum Shoji and Hashimoto (2011)  TIAs ORCA3,4,5 and ORCA2 STR and TDC C. roseus Menke e al. (1999) and Paul e al. (2017) MYC2 TDC C. roseus Paul e al. (2017) MYC1 STR C. roseus Chatel e al. (2003) BIS1 and BIS2 Iridoid biosynthesis C. roseus Van Moerkercke e al. (2016) and Van Moerkercke e al. (2015) ZCT1, ZCT2 and ZCT3 STR and TDC C. roseus Pauw e al. (2004) Flavonoids  Flavonoids MYC2 MYB75/PAP1 and EGL3 A. thaliana Dombrecht e al. (2007) DOF4;2 Produce hydroxycinnamic acid A. thaliana (Skirycz e al. (2007)  Anthocyanins GL3, EGL3, TT8, GL1 and MYB75 Anthocyanins synthesis A. thaliana Qi e al. (2011) MYBL2 TT8 and DFR A. thaliana Xie e al. (2016) MYB115 and MYB134 Proanthocyanidin synthesis Populus James e al. (2017) Phenolic compounds  Phenylpropanoids MYBJS1 PAL and 4CL N. tabacum Galis e al. (2006) MYB8 AT1 and DH29 N. attenuata Onkokesung e al. (2012) Others  GLSs MYC2, MYC3 and MYC4 BCAT4, CYP79B3 and SUR1 A. thaliana Schweizer e al. (2013) MYB34, MYB51 and MYB122 CYP79B2/3, CYP83B1 and SOT16 A. thaliana Frerigmann and Gigolashvili (2014) MYB28, MYB29 and MYB76 Aliphatic GLSs biosynthesis A. thaliana Sonderby e al. (2010)     Camalexin WRKY33 PAD3(CYP71B15) A. thaliana Mao e al. (2011) Open in new tab Acknowledgments We thank the many researchers devoted to plant specialized metabolites and jasmonate signaling. Disclosures The authors have no conflicts of interest to declare. References Bohlmann J. , Meyer-Gauen G. , Croteau R. ( 1998 ) Plant terpenoid synthases: molecular biology and phylogenetic analysis . Proc. Natl. Acad. Sci. USA 95 : 4126 – 4133 . Google Scholar Crossref Search ADS WorldCat Bones A.M. , Rossiter J.T. 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For permissions, please email: [email protected] This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Acosta, Ivan F; Przybyl, Marine

doi: 10.1093/pcp/pcz201pmid: 31651948

Abstract The last stages of stamen development, collectively called stamen maturation, encompass pollen viability, filament elongation and anther dehiscence or opening. These processes are essential for male fertility in Arabidopsis and require the function of jasmonate signaling. There is a good understanding of jasmonate synthesis, perception and transcriptional outputs in Arabidopsis stamens. In addition, the spatiotemporal localization of jasmonate signaling components at the tissue and cellular levels has started to emerge in recent years. However, the ultimate cellular functions activated by jasmonate to promote stamen maturation remain unknown. The hormones auxin and gibberellin have been proposed to control the activation of jasmonate synthesis to promote stamen maturation, although we hypothesize that this action is rather indirect. In this review, we examine these different areas, attempt to clarify some confusing aspects found in the literature and raise testable hypothesis that may help to further understand how jasmonate controls male fertility in Arabidopsis. Introduction In angiosperms, the development of stamens and pistils, the flower organs bearing the male and female reproductive material, is exquisitely controlled to guarantee the success of offspring generation (Ma 2005, Gomez et al. 2015, Erbasol Serbes et al. 2019). Mutations in genes important for any stage of stamen or pistil development, from organ formation to maturation, through cell-specific differentiation and function, can lead to plant sterility (Sanders et al. 1999). For example, in the model plant Arabidopsis, the jasmonate family of phytohormones is indispensable for the final stages of stamen development, as shown by mutants impaired in jasmonate biosynthesis or perception, which are male sterile without compromised female fertility (Feys et al. 1994, Sanders et al. 2000, Stintzi and Browse 2000). In contrast to most Arabidopsis male sterile mutants, jasmonate mutants display remarkably normal-looking stamens until late in development (Fig. 1A). Accordingly, pollen grains seem to develop normally after meiosis, undergoing two rounds of mitosis to produce the expected tricellular gametophyte composed of one vegetative cell and two sperm cells (McConn and Browse 1996). However, pollen grains lose viability after this point and are unable to germinate. Furthermore, the following two other processes fail to occur: the elongation of stamen filaments, which ensures that anthers reach the pistil stigmata for fertilization, and the opening of anthers (dehiscence), which is essential for pollen release (Fig. 1A;Sanders et al. 2000, Stintzi and Browse 2000). We refer to these three aspects of stamen development as maturation. In this review, we examine our current understanding of jasmonate signaling during this process in Arabidopsis, including synthesis, perception, transcriptional changes, possible cellular functions and the influence of auxin and gibberellin. In addition, we identify open questions and potentially interesting research avenues on these topics. Fig. 1 Open in new tabDownload slide Jasmonate signaling during Arabidopsis stamen maturation. See main text for details. (A) Filament (F) elongation starts in wild-type (WT) Arabidopsis stamens at flower stage 12, and it finishes at stage 13 along with anther (A) opening. These processes fail in the jasmonate synthesis mutant aos. Notice that two of the six stamens in Arabidopsis flowers are always shorter and delayed. (B) Jasmonate synthesis pathway in stamens. The genes encoding the corresponding enzymes at each step are abbreviated in bold and italics. Question marks indicate that it is not yet clear which specific AOC and JAR1-type enzymes are required in stamens. 13-HPOT, 13(S)-hydroperoxy-octadecatrienoic acid; 12,13-EOT, (13S)-12,13-epoxy-octadecatrienoic acid. Fig. 1 Open in new tabDownload slide Jasmonate signaling during Arabidopsis stamen maturation. See main text for details. (A) Filament (F) elongation starts in wild-type (WT) Arabidopsis stamens at flower stage 12, and it finishes at stage 13 along with anther (A) opening. These processes fail in the jasmonate synthesis mutant aos. Notice that two of the six stamens in Arabidopsis flowers are always shorter and delayed. (B) Jasmonate synthesis pathway in stamens. The genes encoding the corresponding enzymes at each step are abbreviated in bold and italics. Question marks indicate that it is not yet clear which specific AOC and JAR1-type enzymes are required in stamens. 13-HPOT, 13(S)-hydroperoxy-octadecatrienoic acid; 12,13-EOT, (13S)-12,13-epoxy-octadecatrienoic acid. Jasmonate Synthesis Mutant analysis of jasmonate synthesis and perception genes in Arabidopsis has provided the basis to understand several biological functions of this hormone family. The failure of stamen maturation in mutants devoid of jasmonates in Arabidopsis flowers can be rescued easily by spraying with a concentrated solution of volatile methyl jasmonate. This restores male fertility and self-pollination, allowing the propagation of pure mutant populations (Acosta and Farmer 2010). A summary of the jasmonate synthesis pathway and corresponding enzymes during Arabidopsis stamen maturation is presented in Fig. 1B. Jasmonates are one type of oxylipins, molecules derived from the oxygenation of polyunsaturated fatty acids (Hamberg and Gardner 1992). Jasmonates in particular are made from trienoic α-linonenic acid, the most abundant polyunsaturated fatty acid in plants. One of the first hints that jasmonate is essential for Arabidopsis stamen maturation was provided by the fad3 fad7 fad8 triple mutant. This mutant lacks α-linonenic acid due to a loss of function in all the desaturases catalyzing the last step in the synthesis of trienoic fatty acids (McConn and Browse 1996). α-Linonenic acid is mainly found within plastid membranes as part of glycerolipids, from which it is specifically released in Arabidopsis stamens by the lipase DEFECTIVE IN ANTHER DEHISCENCE1 (DAD1) to initiate jasmonate synthesis (Fig. 1B;Ishiguro et al. 2001). Plastid-localized 13-lipoxygenases (13-LOXs) oxygenate α-linonenic acid in carbon 13 to generate a lipid hydroperoxyde (Bannenberg et al. 2009). The Arabidopsis genome encodes four 13-LOXs (LOX2, LOX3, LOX4 and LOX6). However, only LOX3 or LOX4 are indispensable and sufficient for stamen maturation, as demonstrated by the male sterility of the lox3 lox4 double mutant (Caldelari et al. 2011) and the full fertility of lox2 lox3 lox6 and lox2 lox4 lox6 triple mutants (Chauvin et al. 2013). It is not yet known what determines this specific function of LOX3 and LOX4 in Arabidopsis stamens, but we can propose at least two testable explanations. First, they may be the only 13-LOXs specifically present in the relevant stamen cells or sub-plastidial compartments. Alternatively, they may be better suited than LOX2 and LOX6 to use as substrate the ‘free’ α-linonenic acid released by the DAD1 lipase. Two subsequent enzymes, ALLENE OXIDE SYNTHASE (AOS) and ALLENE OXIDE CYCLASE (AOC), process the lipid hydroperoxyde to produce 12-oxo-phytodienoic acid (OPDA). The AOS enzyme is encoded by a single copy gene, whose loss of function abolishes all jasmonate production in the plant (Park et al. 2002). Conversely, the AOC step is encoded by four genes, three of which are located in tandem in the Arabidopsis genome, which has so far prevented the genetic dissection of their role in stamen maturation (Stenzel et al. 2012). OPDA is transported to peroxisomes where the synthesis of jasmonic acid is completed by an OPDA reductase (OPR3) and three rounds of β-oxidation. The vast majority of jasmonate synthesis occurs through OPR3. However, it was shown recently that OPR2, a paralog localized in the cytosol, is able to partly provide jasmonates in the opr3 mutant through an alternative route during defense responses (Chini et al. 2018). This route is certainly not functional in stamens, where the full sterility of the opr3 mutant clearly indicates that OPR3 is the sole contributor to jasmonate production (Sanders et al. 2000, Stintzi and Browse 2000). Jasmonic acid (JA) eventually accumulates in the cytosol, where the JA-amido synthetase JASMONATE RESISTANT1 (JAR1) conjugates it to one of several amino acids (Staswick and Tiryaki 2004). By far, the most abundant bioactive jasmonate is jasmonoyl-isoleucine (JA-Ile) (Katsir et al. 2008, Yan et al. 2016). No JA-amido synthetase mutant devoid of JA-Ile has been identified yet, although the jar1 mutant shows a reduction of >70% in JA-Ile levels both in flowers and after leaf wounding (Suza and Staswick 2008). Thus, another enzyme or enzymes not yet identified produce the remaining JA-Ile in these tissues. Interestingly, the jar1 mutant is fully fertile but shows reduced jasmonate defense responses upon insect attack (Acosta et al. 2013), suggesting that the low levels of JA-Ile in jar1 flowers sustain stamen maturation more robustly than defense responses. Alternatively, a different jasmonate derivative or conjugate not yet known may be the bioactive molecule activating stamen maturation. Interestingly, recent work has shown that the ABCG-type JASMONATE TRANSPORTER1 (JAT1) is important to translocate JA-Ile from the cytoplasm to the nucleus, the site of jasmonate signaling activation (Li et al. 2017). Similar to jar1, the single mutant jat1 is fertile. However, haploinsufficiency or higher-order loss of both JAR1 and JAT1 causes male sterility that can be rescued by methyl jasmonate application. Therefore, Li et al. (2017) attributed this phenotype to stamen maturation defects, although a detailed description of anther, filament and pollen features is missing. These results emphasize that JA-Ile levels in jar1 stamens are at the limit so that further reducing its transport to the nucleus abolishes jasmonate signaling. Moreover, the rescue of jar1 jat1 fertility with methyl jasmonate supports the existence of additional unknown enzyme(s) capable of converting JA into JA-Ile in the absence of JAR1. Jasmonate Perception and the Activation of Transcription Jasmonate-regulated responses are effected through large changes in gene expression that in most cases require the transcription factor MYC2, which belongs to subclade IIIe of the basic helix–loop–helix (bHLH) family (Lorenzo et al. 2004, Fernandez-Calvo et al. 2011). During stamen maturation, MYC2 works redundantly with its close paralogs MYC3, MYC4 and MYC5 (Qi et al. 2015). Triple myc mutant combinations show delayed anther dehiscence and low pollen viability. These defects worsen in the quadruple myc2 myc3 myc4 myc5 (myc2/3/4/5) mutant, which carries limited viable pollen and additionally displays slower filament elongation. Ultimately, seed set is reduced by 50% in the quadruple mutant, but this partial fertility suggests that additional jasmonate-dependent transcription factors can support the stamen maturation program. MYC2 is believed to recruit the RNA polymerase II transcriptional machinery through its interaction with MED25, a subunit of the Mediator coactivator complex (Chen et al. 2012). However, the biological relevance of this interaction has been mainly reported for jasmonate-dependent defense responses (Kidd et al. 2009), with no documentation of stamen maturation defects in med25 mutants. Thus, which subunits of the Mediator complex may be required for MYC function in this process remains an open question. The current model of jasmonate perception (Fig. 2; reviewed by Howe et al. 2018) proposes that under low JA-Ile concentrations, a multiprotein complex coordinated by JASMONATE-ZIM-DOMAIN (JAZ) proteins represses MYC activity through several simultaneous mechanisms (Chini et al. 2007, Thines et al. 2007, Sheard et al. 2010). These may include directly blocking interactions with transcriptional coactivators, or recruiting TOPLESS-type corepressors either directly or through the adaptor protein NOVEL INTERACTOR OF JAZ (NINJA; Pauwels et al. 2010, Howe et al. 2018). When JA-Ile accumulates, it acts as a molecular glue that brings together a coreceptor formed by JAZ proteins and CORONATINE INSENSITIVE1 (COI1). COI1 is an F-box protein that is part of an E3 ubiquitin ligase complex of the SKP1-CUL1-F-box type, which ubiquitylates JAZs, effectively sending them for degradation by the 26S proteasome (Fig. 2; Xie et al. 1998, Thines et al. 2007, Katsir et al. 2008). This disassembles the entire corepressor complex, liberating MYCs to recruit the transcription machinery and activate the expression of target genes. Since a single copy gene encodes COI1 in the Arabidopsis genome, coi1 mutants are impaired in all jasmonate-mediated responses known to date, including stamen maturation (Xie et al. 1998). Thus, they are indistinguishable from the male sterile jasmonate synthesis mutants, except that jasmonate application obviously does not rescue coi1 fertility. Overexpression of nondegradable JAZ proteins also causes a failure of stamen maturation, nicely supporting their role as repressors of jasmonate signaling (Thines et al. 2007, Chung and Howe 2009). Fig. 2 Open in new tabDownload slide Proposed mechanism of jasmonate perception and transcriptional control of jasmonate-responsive genes with factors known to work during Arabidopsis stamen maturation. See main text for details. Fig. 2 Open in new tabDownload slide Proposed mechanism of jasmonate perception and transcriptional control of jasmonate-responsive genes with factors known to work during Arabidopsis stamen maturation. See main text for details. The core JAZ-MYC jasmonate signaling module is a robust off/on switch that is activated by a wide range of developmental or environmental cues capable of stimulating JA-Ile accumulation (Howe et al. 2018). On the other hand, each cue prompts specific transcriptional changes suited to the required response. Such specificity may be provided by additional transcriptional activators that interact in a combinatorial manner with MYCs (Goossens et al. 2017, Howe et al. 2018). For jasmonate-mediated stamen maturation, it has been proposed that the MYC partners may be at least two related R2R3-type MYB transcription factors, MYB21 and MYB24. The genes encoding these proteins were first identified as induced by jasmonate signaling at the onset of stamen maturation, along with MYB57 and MYB108, two other related genes (Mandaokar et al. 2006). Null myb21 mutants display delayed anther opening and complete failure of filament elongation, which causes full sterility because anthers are unable to reach the pistil stigmata; however, the pollen is viable and the mutants can be propagated by manual ‘self’ pollination (Mandaokar et al. 2006, Reeves et al. 2012). Moreover, myb24 mutants are completely fertile, but double myb21 myb24 mutants fail in all three aspects of stamen maturation and are fully sterile (Mandaokar et al. 2006, Reeves et al. 2012). This indicates that MYB21 is alone essential for filament elongation, while it acts redundantly with MYB24 to promote pollen viability and anther dehiscence. The earlier discovery of the MYB21 and MYB24 genes during stamen maturation suggested a simple scenario where the outcome of jasmonate synthesis and perception was the induced expression of these transcription factors, which would then act as ‘master regulators’ of the stamen maturation program. However, in addition to the importance of MYCs in this process, recent work uncovered that MYB21/24 physically interact with both MYC and JAZ proteins and that JAZs can inhibit their transcriptional activation function (Song et al. 2011, Qi et al. 2015). This suggested the slightly more complex picture of the general JAZ-MYC signaling module cooperating with MYB21/24 to specifically trigger stamen maturation after jasmonate activation (Fig. 2; Qi et al. 2015, Goossens et al. 2017, Howe et al. 2018). This model supposes a pre-existing MYC-JAZ-MYB complex, but it remains an open question what controls the ‘basal’ expression of the different components. This is unclear not only for stamen maturation but also for all other jasmonate-activated responses that use similar modules throughout the plant. One possibility is that the basal transcriptional machinery constantly drives the expression of core JAZ-MYC components at low levels in all tissues, while stamen-specific factors additionally determine the basal expression of MYB21 and MYB24. A remarkable outcome of jasmonate signaling activation is the rapid and transient accumulation of transcripts encoding jasmonate biosynthesis enzymes (e.g. LOX2, LOX3 and LOX4), MYC transcription factors, JAZs and other repressors, and jasmonate catabolic enzymes (Chung and Howe 2009, Koo et al. 2011). Accordingly, the expression of many of these genes is found in the transcriptome of maturing stamens (e.g. Reeves et al. 2012). This ‘jasmonate transcriptional signature’ is believed to create positive and negative feedback loops that, on the one hand, increase the capacity to synthesize and respond to jasmonates and, on the other hand, attenuate the transcriptional signaling output (Wasternack 2007, Howe et al. 2018). It has been suggested that positive feedback upregulation of jasmonate biosynthetic genes does not necessarily result in further jasmonate accumulation (Scholz et al. 2015). However, mathematical modeling suggests that the two opposing loops generate a transient pulse of jasmonate biosynthesis and response (Banerjee and Bose 2011). This agrees well with the expression data of jasmonate-responsive genes in maturing stamens, where they reach a transitory expression peak at flower stage 12 that declines at stage 13 (Nagpal et al. 2005, Reeves et al. 2012). It is conceivable that transient pulses of jasmonate signaling are essential to correctly pattern and limit the expression and function of MYB21 and MYB24 because uncontrolled accumulation of these factors is detrimental not only to fertility but also to plant vegetative growth (Shin et al. 2002, C. Yang et al. 2007, Song et al. 2011). Not surprisingly, loss of jasmonate synthesis/perception or MYC function strongly reduces the expression of the ‘jasmonate transcriptional signature’. Interestingly, however, jasmonate-responsive genes remain upregulated in the myb21 myb24 double mutant past flower stage 13, suggesting that MYB21 and MYB24 also contribute to the negative feedback loop (Reeves et al. 2012). A practical aspect of the feedback upregulation of jasmonate signaling genes is that they constitute excellent markers to diagnose jasmonate signaling activation. However, it is imperative to remember that induction of a gene’s expression does not always imply an underlying function. A clear example is the jasmonate synthesis gene LOX2, which is highly expressed in stamens and follows the kinetics of other jasmonate-responsive genes (Reeves et al. 2012), but it is obviously not required for the process of stamen maturation. Localization and Regulation of Jasmonate Synthesis and Perception in Maturing Stamens There is limited and fragmentary information on the spatiotemporal dynamics of jasmonate synthesis factors during stamen maturation. A DAD1 transcriptional reporter indicates that, in flowers, this gene is exclusively expressed in stamen filaments, starting shortly before the onset of stamen maturation (Ishiguro et al. 2001). Low-resolution in situ hybridization of OPR3 transcripts suggests a similar localization in stamen filaments and in the vascular region at the junction of anthers and filaments; however, OPR3 expression does not seem restricted to the stamen maturation phase, being already present at early developmental stages and in additional flower organs, such as petals and pistils (Sanders et al. 2000). Of the four AOC genes, only AOC1 and AOC4 seem active in mature stamens (filaments and pollen) according to β-glucuronidase transcriptional reporters, but an aoc1 aoc4 double mutant did not have fertility defects (Stenzel et al. 2012). Nevertheless, it should be noted that the T-DNA line that these authors used for aoc4 (SALK_124897) may not be a true loss-of-function allele because their data show only marginal reductions in AOC4 expression and the most recent update of the SALK T-DNA index indicates that this insertion lies within the 5’ untranslated region of AOC4. Alternatively, higher-order mutants including the other AOC genes may be required to unravel which of the four copies are required for stamen maturation. Lastly, immunolocalization of AOC proteins resembles the expression of OPR3 transcripts: widespread in stamens of early flowers but restricted to filaments in maturing flowers (Hause et al. 2003b). These authors also attempted immunocytology of AOS protein and claimed that it was found in pollen, but they did not describe the situation in filaments. In sum, filaments seem the single most shared location of jasmonate synthesis factors during stamen maturation. Although clearly more work is needed to accurately describe their spatiotemporal dynamics, an exclusive localization of jasmonate synthesis to the filaments, if confirmed, raises the interesting possibility that jasmonate exerts non-cell-autonomous effects in pollen and anther tissues. Constitutive levels of jasmonates are normally very low in vegetative tissues but increase within seconds after mechanical wounding (Glauser et al. 2009). In contrast, jasmonates are found at relatively high levels in flowers at the stages of stamen maturation (Reeves et al. 2012); thus, it is expected that this accumulation is developmentally regulated. In both cases, it is not known exactly how stress or developmental signals trigger jasmonate synthesis and, for stamen maturation, there are some limited hints. Lack of the homeotic factor AGAMOUS late in stamen development causes maturation defects that can be rescued by the application of jasmonate or α-linonenic acid. Importantly, AGAMOUS seems able to bind putative cis-regulatory elements of the jasmonate synthesis gene DAD1 and to ectopically activate its expression in petals (Ito et al. 2007). Although no evidence was presented to support that AGAMOUS can induce DAD1 in stamens, the work of Ito et al. (2007) suggests the intriguing model that this transcription factor activates timely DAD1 expression before the onset of stamen maturation to initiate jasmonate synthesis. In addition to this putative direct regulatory function of AGAMOUS, there are several reports of additional factors that seem to indirectly impact the initiation of jasmonate synthesis, including other hormones, such as auxin and gibberellin, that will be discussed below (Nagpal et al. 2005, Cheng et al. 2009, Tabata et al. 2010, Cecchetti et al. 2013, Peng et al. 2013). Interestingly, most of these reports suggest DAD1 expression as the ‘limiting’ step for jasmonate synthesis in stamens. Recent work has shed light on the sites of jasmonate perception in maturing stamens by expressing a COI1-YFP reporter under the control of tissue- or organ-specific promoters in a coi1-1 mutant background (Jewell and Browse 2016). First, the promoter of COI1 rescues all three aspects of stamen maturation and confers expression in most stamen cells except pollen. This expression pattern suggests that (i) jasmonate perception is not required in pollen and is sufficient in sporophytic tissues to drive pollen viability and (ii) most stamen cells are poised for jasmonate perception and COI1 transcription is not a limiting factor. Second, expressing COI1 only in the filament using the DAD1 promoter partly rescues filament elongation but not anther opening or pollen viability; conversely, expressing COI1 only in anther tissues partly rescues anther opening and pollen viability but not filament elongation. These results indicate that jasmonate perception within each tissue type (anther or filament) is necessary and (only) sufficient to activate maturation within such tissue. This also emphasizes that if jasmonate production does occur only in filaments, some jasmonate should be transported to anthers to activate responses there. Third, expressing COI1 in all stamen epidermal cells suffices to partly rescue all three aspects of stamen maturation, suggesting the intriguing idea that the epidermis is the sole site of jasmonate perception in stamens. However, since the rescue is only partial, it is possible that additional cell layers need to activate jasmonate signaling for normal stamen maturation. This may be particularly expected for filament growth, where a coordinated expansion of the epidermis, cortex and vascular cell layers is likely. In situ hybridization of MYB21 around stage 12 finds it expressed in filaments, most strongly in the apical region and in the junction with the anthers, including the anther vasculature tissue (Cheng et al. 2009, Reeves et al. 2012). This pattern agrees with the expression of OPR3 described above and fits with the role of MYB21 in filament elongation. It also suggests that jasmonate synthesis and perception in filaments suffice to activate MYB21 function there. MYB24 shows a similar expression pattern in the filaments but seems absent from anthers (Cheng et al. 2009, Reeves et al. 2012). However, this absence and the restricted expression of MYB21 in the anther vasculature are difficult to reconcile with the role of MYB21 and MYB24 in anther opening. As discussed below, one of the proposed effects of jasmonate signaling to promote anther opening is the breakage of stomium epidermal cells, which are several cell layers beyond the vasculature. Moreover, the work of Jewell and Browse (2016) suggests that jasmonate perception occurs not only in the vasculature but also in other anther cells, such as the epidermis, where it may suffice for anther opening. Thus, similar to jasmonate synthesis factors, the spatiotemporal dynamics of MYB21 and MYB24 requires more detailed analysis to clarify where exactly it occurs to promote filament elongation and anther opening. Possible Cellular Events Activated by Jasmonate Signaling to Drive Stamen Maturation In contrast to the mostly clear understanding of the jasmonate signaling components required for stamen maturation, there has been only limited research on how the transcriptional reprogramming activated by jasmonate redirects cell functions to drive pollen viability, filament elongation and anther opening. The two published transcriptomes of jasmonate signaling mutants may provide a starting point to hypothesize potential executors of the stamen maturation program: the time-course transcriptome of jasmonate-deficient opr3 mutant stamens after jasmonate treatment (Mandaokar et al. 2006) and the differentially expressed genes in myb21 myb24 mutant flowers at stages 12 and 13 (Reeves et al. 2012). For example, Mandaokar et al. (2006) proposed that jasmonate signaling induces the synthesis of waxes that may be important for pollenkitt formation and, therefore, pollen viability. Moreover, stamen filament elongation occurs through increases in cell length (Mandaokar et al. 2006, Cheng et al. 2009, Reeves et al. 2012) and the transcriptomes induced by jasmonate or MYB21/24 include cell wall-modifying enzymes that may play a role in the expansion of filament cells. However, all these hypotheses remain to be tested. What seems clear is that jasmonate promotes stamen maturation by instructing different cell types to perform particular and disparate functions, such as expansion and degeneration (see below). Thus, another unsolved question is how each cell-specific response is achieved. It is possible that MYB21/24 activate the expression of a second level of transcription factors in different cell types. For example, one of the primary targets of MYB21/MYB24 may be MYB108, which seems involved in anther dehiscence only (Mandaokar and Browse 2009, Reeves et al. 2012). Pollen is produced and enclosed within two pairs of anther locules or chambers (Fig. 3). Adjacent locules within a pair are kept apart by a group of cells forming a septum. In addition, adjacent locules converge at the stomium, the epidermal region where the anther actually opens for pollen release. Several processes are required for anther dehiscence (represented in Fig. 3; reviewed by Wilson et al. 2011): (i) deposition of ligno-cellulosic thickenings at the cell walls of subepidermal endothecium cells; (ii) separation or break down of septum cells, leading to locule pair fusion; (iii) separation or break down of the specialized epidermal cells forming the stomium; and (iv) dehydration of the anther surface. It is believed that this dehydration along with the tension created by the secondary thickenings of the endothecium bends the locule outward, widening the stomium opening to allow pollen release (Keijzer 1987, Nelson et al. 2012). Fig. 3 Open in new tabDownload slide Schematic representation of Arabidopsis anther cross-sections at stages 11 and 12. Distinct cell types are shown with different colors. The tapetum layer is not visible anymore at stage 12. Red bars on the endothecium layer represent secondary thickenings. At stage 12 in this depiction, septum rupture is complete in both locule pairs, while stomium breakage to allow pollen release has only occurred in the right locule pair. Fig. 3 Open in new tabDownload slide Schematic representation of Arabidopsis anther cross-sections at stages 11 and 12. Distinct cell types are shown with different colors. The tapetum layer is not visible anymore at stage 12. Red bars on the endothecium layer represent secondary thickenings. At stage 12 in this depiction, septum rupture is complete in both locule pairs, while stomium breakage to allow pollen release has only occurred in the right locule pair. In Arabidopsis, endothecium secondary thickening does not require jasmonate signaling (Ishiguro et al. 2001, Steiner-Lange et al. 2003, Cecchetti et al. 2013). Instead, it is solely dependent on the transcription factor MYB26, which partly acts by activating the expression of two other essential transcription factors, NST1 and NST2, presumed activators of genes encoding cellulose and lignin biosynthetic enzymes (Mitsuda et al. 2005, X. Y. Yang et al. 2007, Yang et al. 2017). Jasmonate signaling seems only necessary for stomium breakage because anther histology shows that only this process is absent or delayed in the jasmonate-deficient mutants dad1 and opr3; in contrast, they display normal septum rupture (Sanders et al. 2000, Ishiguro et al. 2001). Based on several model species, it has been proposed that cell separation events in the septum and the stomium involve pectin-degrading enzymes that facilitate cell wall loosening (Keijzer 1987, Keijzer et al. 1996, Wilson et al. 2011). Loss-of-function mutations in a specific clade of pectin-degrading polygalacturonases of tomato (PS-2) and Arabidopsis (ADPG1, ADPG2, QRT2) block or delay anther dehiscence, supporting the importance of cell wall enzymatic lysis in this process (Gorguet et al. 2009, Ogawa et al. 2009). Based only on histological observations, the authors attributed this mutant defect to a failed rupture of the stomium, not the septum. The transcriptomic data suggest that jasmonate and MYB21/24 may control the expression of genes encoding these or similar pectin-degrading enzymes (Mandaokar et al. 2006, Reeves et al. 2012), but deeper analyses are required. Stomium breakage has also been variously attributed to simple mechanical rupture (Keijzer 1987, Wilson et al. 2011) or to active programmed cell death (Sanders et al. 2000, Sanders et al. 2005). In particular, Sanders et al. (2000) claim that the opr3 mutant shows a delayed anther dehiscence due to a lag in the degeneration of stomium cells, raising the interesting possibility that jasmonate activates developmentally programmed cell death. However, a clearer, unequivocal definition of the cellular and molecular events leading to the separation or break down of septum and stomium cells is still necessary. For example, it is important to detect and follow the kinetics of pectin degradation or cell death in those cells to investigate if these events are truly necessary for anther opening in Arabidopsis. Ishiguro et al. (2001) hypothesized that jasmonate signaling is important for anther dehydration, promoting water movement out of the anthers into the filaments, which would also cause filament cell expansion. Such mechanism would achieve an elegant synchronization of anther dehiscence and filament elongation. However, the tissue-specific rescue of jasmonate perception reviewed earlier indicates that these two processes occur mainly independent of each other (Jewell and Browse 2016). This is further supported by the phenotype of the myb21 mutant stamens, where filaments fail to elongate but anthers open successfully although with some delay. Still, it is formally possible that water movements that follow an osmotic potential in filaments contribute to elongation (Keijzer 1987, Bonner and Dickinson 1990). Ishiguro et al. (2001) suggested that jasmonate produced in filaments could stimulate this osmotic potential by activating the expression of the sugar transporter AtSUC1 at the vascular interface of anther and filament tissues. Nevertheless, the transcriptome data does not support that jasmonate or MYB21/24 control the expression of AtSUC1. Instead, myb21/24 mutant stamens seem to show lower expression of several ion channels that may facilitate the transport of ions, such as potassium, which could be a faster and more sensitive source of osmotic potential in filaments (Heslop-Harrison et al. 1987, Heslop-Harrison and Heslop-Harrison 1996). Recent work has provided genetic and physiological evidence for the requirement of dehydration in Arabidopsis anther opening and pollen viability. INDUCER OF CBF EXPRESSION 1 (ICE1), another MYC-type transcription factor, is essential for the differentiation of abaxially localized anther stomata (Wei et al. 2018). Although stomium breakage seems to occur in an ice1 mutant, the anther epidermis remains hydrated, preventing the widening of the stomium and, therefore, pollen release. This supports that water evaporation via stomata is essential for anther dehydration and full dehiscence and suggests that it is another event putatively activated by jasmonate signaling. Still, this does not rule out that osmotic potentials are also needed to drive additional water movements toward the filament, petals or other organs. Interestingly, jasmonate and ICE1 promote cold stress tolerance; instead, JAZ proteins repress it because they bind and block ICE1 function; thus, jasmonate likely activates ICE1 under cold stress (Hu et al. 2013). This raises the interesting possibility that jasmonate also activates ICE1 in anthers to allow stomata differentiation. This hypothesis has not yet been tested and would imply that jasmonate signaling mutants carry undifferentiated stomata precursor cells similar to the ice1 mutant. Alternatively, jasmonate could promote anther stomata opening as the JA-Ile mimic coronatine does in leaves during Pseudomonas syringae infections (Melotto et al. 2006, Gimenez-Ibanez et al. 2017). Interaction of Jasmonate Signaling with Auxin and Gibberellin Several works have concluded that the hormones auxin and gibberellin control jasmonate synthesis through the regulation of DAD1 expression at the onset of stamen maturation (Nagpal et al. 2005, Cheng et al. 2009, Tabata et al. 2010, Reeves et al. 2012, Cecchetti et al. 2013). However, no direct effect of auxin or gibberellin signaling on the promoter of DAD1 has been yet reported. Moreover, as detailed below, jasmonate treatments are insufficient to rescue the stamen maturation defects of mutants impaired in auxin or gibberellin signaling (Nagpal et al. 2005, Cheng et al. 2009). Therefore, we favor the more open interpretation that these hormones work first and foremost to complete the development of different stamen cell types, which thereby become ‘competent’ to activate DAD1 expression and jasmonate synthesis. In this model, interrupting auxin or gibberellin signaling indirectly blocks jasmonate accumulation and, therefore, responses such as the jasmonate transcriptional signature. The model is summarized in Fig. 4 and detailed below. Fig. 4 Open in new tabDownload slide Interaction model of jasmonate with auxin and gibberellin during stamen maturation. We propose that gibberellin (GA) and auxin allow normal filament development, which is required to activate jasmonate (JA) synthesis via DAD1. Moreover, gibberellin potentially promotes AGAMOUS function, which may also induce DAD1 expression. The low specific levels of auxin required to activate ARF6/8 function might be reached through catabolism with a DAO enzyme or through downregulation of the auxin synthesis genes YUCCA 2/6 mediated by a putative FTIP-like factor. ARF6/8 also contributes independently to anther opening by activating MYB26 expression. See main text for further details. Fig. 4 Open in new tabDownload slide Interaction model of jasmonate with auxin and gibberellin during stamen maturation. We propose that gibberellin (GA) and auxin allow normal filament development, which is required to activate jasmonate (JA) synthesis via DAD1. Moreover, gibberellin potentially promotes AGAMOUS function, which may also induce DAD1 expression. The low specific levels of auxin required to activate ARF6/8 function might be reached through catabolism with a DAO enzyme or through downregulation of the auxin synthesis genes YUCCA 2/6 mediated by a putative FTIP-like factor. ARF6/8 also contributes independently to anther opening by activating MYB26 expression. See main text for further details. To investigate the relationship between gibberellin and jasmonate signaling during stamen development, Cheng et al. (2009) used the gibberellin synthesis mutant ga1, which is practically devoid of gibberellins and displays pleiotropic developmental defects. In addition to having shorter stamen filament cells, ga1 fails to produce viable pollen because microspores do not separate properly after meiosis nor undergo mitosis, and eventually degenerate. The anther wall tissue also develops abnormally and collapses (Cheng et al. 2004). In contrast, the pollen in jasmonate mutants almost reaches maturity, arriving at the tricellular stage but losing viability afterward, and anther wall tissues are normal until late in development (Sanders et al. 2000, Cecchetti et al. 2013). This suggests that lack of gibberellin arrests several aspects of stamen development long before jasmonate signaling is activated. Consequently, jasmonate levels and part of the jasmonate transcriptional signature are lower in ga1. This correlates with an 80% reduction in the transcripts of DAD1, indicating that ga1 arrested stamens are unable to initiate jasmonate synthesis (Cheng et al. 2009). It should be noted that Cheng et al. (2009) also considered the reduced expression of LOX1 in ga1 as correlative with the low jasmonate production. However, LOX1 is a 9-lipoxygenase that does not participate in jasmonate synthesis, a job performed exclusively by 13-lipoxygenases and specifically by LOX3 and LOX4 in stamens. Interestingly, treating ga1 flowers with jasmonate does activate the expression of MYB21 and MYB24 (Cheng et al. 2009), suggesting that reproductive tissues are competent to respond to jasmonate before stamen maturation, but the limiting step for signaling activation is jasmonate production. On the other hand, even if exogenous jasmonate activates signaling in the ga1 background, this is insufficient to rescue the stamen development arrest of the mutant. This emphasizes that jasmonate synthesis is just one of several aspects that are blocked in ga1 and that have to take place before the stamen maturation program is initiated (Cheng et al. 2009). These authors also suggest the intriguing possibility that gibberellin signaling may activate the expression of AGAMOUS, which in turn would directly control DAD1 expression. All flower organs of a double mutant defective in the auxin response factors ARF6 and ARF8 arrest at stage 12, shortly before the onset of stamen maturation. Consequently, both filament elongation and anther dehiscence fail to occur (Nagpal et al. 2005). These defects are only partial and variable in single arf6 or arf8 mutants indicating that these factors act in part redundantly. The arrest of arf6 arf8 stamens seems to happen later than that of ga1, although no histological description has been reported. Thus, it is not clear if and when arf6 arf8 pollen is defective, or if arf6 arf8 anthers are affected in other processes necessary for dehiscence but independent of jasmonate signaling, such as endothecium lignification. However, class 1 KNOX genes, which are known as repressors of cell differentiation, are ectopically expressed in arf6 arf8 flowers and this seems to partly account for their arrest (Tabata et al. 2010). Moreover, reducing ARF6/8 activity appears to block filament vasculature at the procambium stage, because a procambial marker shows an expanded expression in the double mutant (Rubio-Somoza and Weigel 2013). Similar to ga1, jasmonate levels in arf6 arf8 are reduced and correlated with a lack of DAD1 expression (Nagpal et al. 2005, Tabata et al. 2010). In contrast to ga1, jasmonate application does rescue anther opening in arf6 arf8 but not filament elongation, so it has been concluded that the low jasmonate accumulation in this mutant is only responsible for one defect but not the other (Nagpal et al. 2005). We propose that ARF6 and ARF8 are required for the correct development of filament cells, which are likely the source, via DAD1 activation, of the jasmonate that triggers both filament elongation and anther opening. In this model, arf6 arf8 filament cells remain immature, not competent to initiate the synthesis of jasmonate or to respond to it. Instead, anther tissues do mature and are obviously responsive to jasmonate but unable to synthesize it independently. In this sense, the work on ARF6 and ARF8 would support that anthers are normally not capable of triggering jasmonate synthesis, for which they are fully dependent on filaments. The potential requirement of ARF6 and ARF8 for the completion of vasculature development in the filament is remarkable, because jasmonate synthesis genes and enzymes can be expressed in vascular tissues (Hause et al. 2003a; Gasperini et al. 2015), which may be a suitable location for jasmonate production if transport to the anther is needed. Recent work has shown that the function of ARF8 in filament elongation is most likely effected through specific splice variants, mainly ARF8.4 with perhaps a minor contribution of ARF8.2 (Ghelli et al. 2018). Inducing higher levels of ARF8.4 in wild-type plants increases final stamen length slightly but significantly, and it restores the mild 15% reduction in filament length of the arf8-7 single mutant. Moreover, induction of ARF8.4 in arf8-7 recovers the expression of Aux/IAA19, an auxin-responsive gene that Ghelli et al. consider a master regulator of filament elongation. However, this interpretation is inconsistent with the function of Aux/IAA19 as a repressor of auxin signaling. Analogously to JAZs in jasmonate signaling, Aux/IAA proteins repress the function of ARFs until auxin accumulation and sensing target them for degradation (Chapman and Estelle 2009). Accordingly, the dominant MASSUGU2 mutations disrupt the degron motif of Aux/IAA19, which renders it stable and insensitive to auxin degradation (Tatematsu et al. 2004). Thus, filament growth is delayed in MASSUGU2 (Tashiro et al. 2009), similar to arf6 or arf8 single mutants, likely because ARF6/8 remain partially repressed. Therefore, Aux/IAA19 is a repressor of the filament cell development program enabled by ARF6 and ARF8 and has to be eliminated through auxin perception to activate ARF function. In addition, ARF6/8 probably induce the expression of not only Aux/IAA19 but also at least five other Aux/IAAs [c.f. microarray data in Reeves et al. (2012)]. Again, in analogy to jasmonate signaling, Aux/IAA induction probably serves a negative feedback role to limit (not stimulate) auxin-ARF output. It is also a stereotypical transcriptional response that is a practical readout of auxin-ARF signaling activation (Chapman and Estelle 2009). Ectopically inducing higher levels of both ARF8.2 and ARF8.4 causes precocious anther dehiscence in wild-type and arf8-7 anthers (Ghelli et al. 2018). This phenomenon was correlated with an earlier endothecium lignification mediated by ARF8.4 and with an increased DAD1 expression brought about through ARF8.2. As with any other overexpression or gain-of-function phenotype, this result shows some potential functions of ARF8.2 and ARF8.4, but this does not necessarily mean that they are actually performing these functions when expressed under their endogenous promoter. Therefore, it will be important to test if ARF8.2 is sufficient to rescue DAD1 expression and, therefore, jasmonate biosynthesis, in the full loss-of-function mutant arf6 arf8. Moreover, the potential role of ARF6/8 in endothecium lignification suggests that this process is likely impaired in arf6 arf8. However, as mentioned earlier, this and other histological features have not been yet reported for this mutant. If such a defect is found, it will also be necessary to test if ARF8.4 suffices to drive endothecium lignification in an arf6 arf8 double mutant. As mentioned earlier, the canonical auxin perception mechanism is expected to activate ARF6 and ARF8. This involves degradation of Aux/IAA repressors after auxin-mediated interaction with TIR1/ABF F-box proteins. Accordingly, a tir1 afb1 afb2 afb3 quadruple mutant shows approximately 25% reduced filament elongation with respect to the wild type (Cecchetti et al. 2008), reminiscent of the arf8-7 mutant. However, in stark contrast to the indehiscence of arf6 arf8, the tir1 afb1 afb2 afb3 mutant shows approximately 90% precocious anther dehiscence at stage 12, when is rarely observed in the wild type (Cecchetti et al. 2008). The triple mutant tir1 afb2 afb3 and the single mutant afb1 also display this phenotype at early stage 12, although at lower frequencies, 35 and 10%, respectively (Cecchetti et al. 2013). Furthermore, pollen grains also mature prematurely in all three mutants. In agreement with these phenotypes, artificially increasing the output of auxin signaling with exogenous auxin application or ectopic expression of the Agrobacterium rolB oncogene can delay anther dehiscence in Arabidopsis or tobacco, respectively (Cecchetti et al. 2004, Cecchetti et al. 2008, Cecchetti et al. 2013). In rice, excess accumulation of the bioactive auxin IAA blocks anther dehiscence of the dao mutant, impaired in an enzyme that catabolizes IAA, and of the Osftip7 mutant, affected in a protein essential for the downregulation of the auxin synthesis gene OsYUCCA4 (Zhao et al. 2013, Song et al. 2018). Collectively, this evidence supports that auxin signaling negatively regulates anther dehiscence and pollen viability. However, such conclusion seems to contradict the positive role of the auxin-dependent factors ARF6 and ARF8 in anther opening. Similar to the ectopic expression of ARF8.2 and ARF8.4, the precocious anther dehiscence in the tir1/afb mutants is due to an earlier expression of MYB26, which causes accelerated endothecium lignification, and to elevated jasmonate levels associated with higher DAD1 expression (Cecchetti et al. 2013). This deepens the contradiction because two seemingly opposite events (blocking auxin signaling and ectopically activating ARF8, an auxin-dependent factor) cause the same molecular and phenotypic effects. The root of this inconsistency may be the way Cecchetti et al. interpret the kinetics of auxin and auxin-induced DR5 reporters that they have observed in anthers (Cecchetti et al. 2008, Cecchetti et al. 2013). Both auxin and DR5 reporters peak after meiosis (stage 10) and then decline when endothecium lignification is complete (stage 11). Auxin concentration further decreases, and DR5 reporters are not detectable at the onset of maturation (stage 12). Moreover, DR5 reporters are inactive in tir1/afb multiple mutants. Thus, Cecchetti et al. (2008) imply that auxin signaling in anthers is normally switched off before maturation starts to allow anther opening. We propose instead that the reduced but substantial auxin content at stage 12 (Cecchetti et al. 2013) does drive a lower signaling output that is specifically required to activate ARF6 and ARF8 at appropriate levels in anthers (Fig. 4). It is also possible that the different affinities of each F-box receptor for different natural auxins or auxin-related molecules may determine the specific level of signaling required for ARF6/8 function during stamen maturation. However, this degree of auxin signaling is likely insufficient to activate DR5, a promoter that does not report all auxin responses (Liao et al. 2015, Chandler 2016). Instead, Aux/IAA19, another auxin response gene likely activated by ARF6 and ARF8, shows a strong, later expression (Tashiro et al. 2009) that may reflect auxin signaling during stamen maturation more accurately. In our interpretation, auxin perception is not abolished but only reduced in the single, triple or quadruple tir1/afb mutants to a level that allows earlier ARF6/8 activation and anther opening. The partial reduction (∼25%) of filament growth in the tir1 afb1 afb2 afb3 quadruple mutant supports this idea because it suggests that some auxin signaling is still occurring through AFB4 and AFB5. These are the remaining functional AFB genes that probably step in to activate ARF6/8 and allow significant filament elongation. Our interpretation predicts that the precocious anther dehiscence of tir1 afb1 afb2 afb3 requires ARF6/8 function. This can be unequivocally tested by introducing the tir1/afb mutations in an arf8 or arf6 arf8 background. Furthermore, we predict that completely removing auxin perception should not cause precocious anther dehiscence but rather prevent it entirely as in arf6 arf8. Recent work has generated even higher-order tir1/afb mutants that are viable and may allow to test this hypothesis (Prigge et al. 2019). Finally, the inhibitory effect of high auxin (signaling) levels in anther dehiscence may be due in part to excess ARF6/8 activity, which has also been shown to negatively impact the progression of this process (Wu et al. 2006, Zheng et al. 2019). Concluding Remarks The maturing Arabidopsis stamen has been a great model to uncover some of the important components of jasmonate signaling in general. Thus, there is clear genetic support for many of the jasmonate-related factors required for stamen maturation, including most of the specific enzymes of jasmonate synthesis, the hormone perception coreceptor and the transcription factors translating jasmonate signaling into gene expression changes. However, many interesting questions remain regarding (i) the cell-specific sites of jasmonate production and perception in stamens; (ii) the significance of the feedback expression of jasmonate-related genes; (iii) the actual cellular functions activated by gene expression to ultimately drive pollen viability, filament elongation and anther opening; and (iv) the interaction of jasmonate with other hormones during stamen maturation. We have outlined those questions and proposed ways to answer them. Continued research on the late stages of stamen development will not only refine our knowledge of jasmonate signaling but also uncover general principles for the workings of plant hormones, particularly how signaling results in specific developmental responses. Funding This work was supported by core funding from the Max Planck Society to the Max Planck Institute for Plant Breeding Research. References Acosta I.F. , Farmer E.E. ( 2010 ) Jasmonates . Arabidopsis Book 8 : e0129 . 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This is an Open Access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted reuse, distribution, and reproduction in any medium, provided the original work is properly cited. © The Author(s) 2019. Published by Oxford University Press on behalf of Japanese Society of Plant Physiologists.

Harada,, Kyohei;Arizono,, Takatoshi;Sato,, Ryoichi;Trinh, Mai Duy, Luu;Hashimoto,, Akira;Kono,, Masaru;Tsujii,, Masaru;Uozumi,, Nobuyuki;Takaichi,, Shinichi;Masuda,, Shinji

doi: 10.1093/pcp/pcz203

Lim,, HyunSeok;Tanaka,, Ayumi;Tanaka,, Ryouichi;Ito,, Hisashi

doi: 10.1093/pcp/pcz157pmid: 31392311

Abstract In plants, chlorophyll (Chl) a and b are interconvertible by the action of three enzymes—chlorophyllide a oxygenase, Chl b reductase (CBR) and 7-hydroxymethyl chlorophyll a reductase (HCAR). These reactions are collectively referred to as the Chl cycle. In plants, this cyclic pathway ubiquitously exists and plays essential roles in acclimation to different light conditions at various developmental stages. By contrast, only a limited number of cyanobacteria species produce Chl b, and these include Prochlorococcus, Prochloron, Prochlorothrix and Acaryochloris. In this study, we investigated a possible existence of the Chl cycle in Chl b synthesizing cyanobacteria by testing in vitro enzymatic activities of CBR and HCAR homologs from Prochlorothrix hollandica and Acaryochloris RCC1774. All of these proteins show respective CBR and HCAR activity in vitro, indicating that both cyanobacteria possess the potential to complete the Chl cycle. It is also found that CBR and HCAR orthologs are distributed only in the Chl b-containing cyanobacteria that habitat shallow seas or freshwater, where light conditions change dynamically, whereas they are not found in Prochlorococcus species that usually habitat environments with fixed lighting. Taken together, our results implicate a possibility that the Chl cycle functions for light acclimation in Chl b-containing cyanobacteria. Introduction Chlorophyll (Chl) plays crucial roles in photosynthesis by harvesting light energy and driving electron transfer (Renger and Schlodder 2010). Photosynthetic organisms have to change their light-harvesting capacity and number of photosystems to perform efficient and safe photosynthesis under changing light environments (Webb and Melis 1995, Dietzel et al. 2008). To achieve these photosystem restructurings, synthesis and degradation of Chl must be finely regulated in response to light environments (Tanaka et al. 2001, Masuda et al. 2003). Green plants contain Chl a and Chl b, which have different absorbance spectra and contribute to the use of a wide range of light spectra (Chen 2014). Chl a is responsible for charge separation in photosynthetic reaction centers and is a major light-harvesting pigment both in the core (CP43/47 of photosystem II and PsaA/B of photosystem I) and the peripheral antenna complexes. By contrast, Chl b is responsible only for light harvesting. The localization of Chl b in light-harvesting systems is different among green plants (Kunugi et al. 2016). Photosynthetic organisms adapted to high light levels, such as Chlamydomonas and land plants, contain Chl b only in the peripheral antenna complexes, whereas Chl b exists in both the core and the peripheral antenna complexes in green algae living in deep seas. The green plants alter their antenna size by changing the Chl a/b ratio (Bailey et al. 2001). When the plants grow under low light conditions, the plants have a low Chl a/b ratio and large antenna size. Therefore, the Chl a/b ratio must be regulated in response to changes in the light environment. Green plants employ various strategies to regulate the Chl a/b ratio. Chlorophyllide (Chlide) a oxygenase (CAO) is responsible for Chl b synthesis. CAO protein levels are finely regulated at the transcriptional level and protein degradation rates (Tanaka and Tanaka 2005, Yamasato et al. 2005, Nakagawara et al. 2007). Another important reaction for regulating the Chl a/b ratio is the conversion of Chl b to Chl a (Tanaka and Tanaka 2007). In this pathway, Chl b is converted to 7-hydroxymethyl chlorophyll a (HMChl a) by Chl b reductase (CBR; Kusaba et al. 2007), followed by the reduction to Chl a by HMChl a reductase [7-hydroxymethyl chlorophyll a reductase (HCAR); Meguro et al. 2011; Fig. 1]. This pathway is responsible not only for regulation of the Chl a/b ratio but also for the degradation of Chl b during senescence because Chl b must be converted to Chl a before degradation (Hörtensteiner et al. 1995). Therefore, interconversion of Chl a and Chl b (the Chl cycle) plays a crucial role in various developmental stages in green plants. Fig. 1 Open in new tabDownload slide Chl metabolic pathway. Outline arrow, black arrow and dashed arrow represent the Chl synthesis pathway, Chl cycle and Chl degradation pathway, respectively. Dashed circle indicates the reaction site. Chl Syn, Chl synthase. Fig. 1 Open in new tabDownload slide Chl metabolic pathway. Outline arrow, black arrow and dashed arrow represent the Chl synthesis pathway, Chl cycle and Chl degradation pathway, respectively. Dashed circle indicates the reaction site. Chl Syn, Chl synthase. Chl b is not present as free pigment but as light-harvesting Chl a/b-protein complexes (LHC) in the chloroplast. Mutant analysis with Arabidopsis (Horie et al. 2009) and rice (Kusaba et al. 2007) demonstrated that LHC is not degraded in the CBR mutant during senescence. In vitro experiments indicated that Chl b in LHC can be a substrate of recombinant CBR (Horie et al. 2009). These experiments indicate that Chl b in LHC is the primary substrate of CBR in chloroplasts. CBR might have evolved to be able to catalyze Chl b in LHC. Chl b is found not only in eukaryotic green plants but also in some cyanobacterial lineages, such as Prochlorococcus, Prochloron and Prochlorothrix (Palenik and Haselkorn 1992). Recently, a novel species of Acaryochloris was found that has Chl b instead of Chl d (Partensky et al. 2018). These organisms containing Chl b do not form a single cluster in the phylogenetic tree but appear independently in the cyanobacterial lineage (Partensky et al. 2018). Cyanobacteria do not have LHC. Instead, Chl b is incorporated into the prochlorophyte Chl b-binding protein (PCB; Bibby et al. 2003, Bumba et al. 2005), which is not phylogenetically related to LHC (La Roche et al. 1996). The localization of Chl b in Acaryochloris has not been reported, but it might be present in PCB, as in Prochlorococcus and Prochlorothrix. Chl metabolic pathways, including biosynthesis, the Chl cycle and degradation, have been determined (Tanaka and Tanaka 2005, Hörtensteiner 2006, Nagata et al. 2007), and major enzymes for these pathways have been identified in land plants (Nagata et al. 2005, Hauenstein et al. 2016, Shimoda et al. 2016). Pathway and enzymes of Chl biosynthesis are also determined in cyanobacteria. Although Chl degradation is an important process for cyanobacteria, the enzymes and pathway of Chl degradation have not been determined. As for the Chl cycle, Chl b is synthesized by CAO in cyanobacteria, as in green plants (Satoh and Tanaka 2006). However, it is not evident whether these cyanobacteria have a Chl b-to-a conversion pathway because CBR and HCAR of cyanobacteria could not be proposed only by the sequence similarity and phylogenetic tree. In this study, we constructed phylogenetic trees of CBR, HCAR and their homologous genes and determined the candidates of these genes in Prochlorothrix hollandica and Acaryochloris RCC1774. Then, we tried to detect the enzymatic activities using the recombinant proteins of these candidate genes and found the CBR and HCAR activities, implicating the presence of the Chl cycle in cyanobacteria containing Chl b. We discuss the functions and evolution of the Chl cycle in cyanobacteria. Results Pigment composition of P. hollandica and Acaryochloris RCC1774 Photosynthetic pigments of P. hollandica and Acaryochloris RCC1774 were examined by HPLC (Fig. 2A). Pigment compositions were similar to those in previous reports (Takaichi et al. 2012, Partensky et al. 2018). The elution profiles of HPLC were similar between the two organisms with some differences. Acaryochloris RCC1774 and P. hollandica have α- and β-carotene, respectively. Both of them have ε,ε-carotene (Fig. 2B). Chl a/b ratios of Acaryochloris RCC1774 and P. hollandica were 6.8 and 13.5, respectively. Acaryochloris RCC1774 accumulates divinyl protochlorophyllide a, which is generally a precursor of Chl biosynthesis, whereas P. hollandica does not accumulate this pigment. The absorbance spectrum of this pigment is similar to that of Chl c; therefore, it may function for light harvesting in Acaryochloris RCC1774 like it does in Prochloron (Larkum et al. 1994). Fig. 2 Open in new tabDownload slide Pigment compositions of Acaryochloris RCC1774 and P. hollandica. (A) HPLC profiles of the pigment monitored at 440 nm. The peaks were identified by their retention time and spectrum. (B) The absorption spectrum of ε,ε-carotene detected in Acaryochloris RCC1774. Fig. 2 Open in new tabDownload slide Pigment compositions of Acaryochloris RCC1774 and P. hollandica. (A) HPLC profiles of the pigment monitored at 440 nm. The peaks were identified by their retention time and spectrum. (B) The absorption spectrum of ε,ε-carotene detected in Acaryochloris RCC1774. Phylogenetic analysis of the Chl cycle enzymes CAO catalyzes the oxidation of a methyl group on Chl a to a formyl group (Oster et al. 2000), and it is responsible for the formation of Chl b in all organisms containing Chl b (Tomitani et al. 1999). The phylogenetic tree of CAO (Supplementary Fig. S1) demonstrates the phylogenetic relationship of these organisms. The CAO gene of green plants and cyanobacteria shares the common ancestor suggesting that the CAO gene has been transferred from an ancestral cyanobacterium to plants via the endosymbiosis event. CBR catalyzes the reduction of a formyl group on Chl b to a hydroxymethyl group, which is the first step of Chl b-to-a conversion. CBR belongs to a short-chain dehydrogenase family that has an enormous number of members and is greatly diversified (Kallberg et al. 2002). CBR homologs are widely distributed not only in green plants but also in other organisms, including red algae, diatoms, cyanobacteria and photosynthetic bacteria (Fig. 3A). Green plants have two CBRs, Non-Yellow Coloring 1 (NYC1) and NYC1-Like (NOL), whereas a group of green algae, Pracinophytes (Ostreococcus and Micromonas), has only one CBR gene. The phylogenetic analysis did not clearly show to which group Ostreococcus and Micromonas CBRs belong. These CBRs have no membrane-spanning helix, which is a characteristic feature of NOL, suggesting that they belong to the NOL members. Fig. 3 Open in new tabDownload slide Phylogenetic trees of CBR, HCAR and BciB. The phylogenetic trees were constructed by maximum likelihood. The numbers at each node represent the bootstrap value and the number of amino acid substitutions per site is illustrated by the scale bar. (A) Phylogenetic tree of CBR. Arabidopsis NYC1 (AT4G13250); Arabidopsis NOL (AT5G04900); Oryza sativa Japonica Group NOL (XP_015628274.1); O. sativa Japonica Group NYC1 (XP_015621887.1); Klebsormidium nitens NYC1 (GAQ77737.1); K. nitens NOL (GAQ87774.1); Coccomyxa subellipsoidea C-169 NYC1 (XP_005652224.1); C. subellipsoidea C-169 NOL (XP_005646276.1); Pelodictyon phaeoclathratiforme (WP_012508106.1); Ostreococcus lucimarinus CCE9901(XP_001415854.1 ); Micromonas commoda (XP_002508859.1); Chlamydomonas reinhardtii NYC1 (XP_001697080.1); C. reinhardtii NOL (XP_001701347.1); Vitrella brassicaformis CCMP3155 (CEM24690.1); Candidatus Heimdallarchaeota archaeon AB_125 (OLS31532.1); Chondrus crispus (XP_005716045.1); Chlorobaculum tepidum (WP_010932815.1); Chlorobaculum limnaeum (WP_069809375.1); Fistulifera solaris (GAX23003.1); P. hollandica (WP_081599361.1); Acaryochloris RCC1774 (1) (WP_110987898.1); Acaryochloris RCC1774 (2) (WP_110986784.1); Synechocystis PCC6803 (WP_041428273.1). (B) Phylogenetic tree of BciB and HCAR. C. reinhardtii (PNW76723.1); M. commoda (XP_002503439.1); O. lucimarinus CCE9901 (XP_001416225.1); C. subellipsoidea C-169 (XP_005648937.1); K. nitens (GAQ88093.1); Physcomitrella patens (XP_024368500.1); Selaginella moellendorffii (XP_024538910.1); O. sativa Japonica Group (XP_015636785.1); Arabidopsis HCAR (AT1G04620.1); Acaryochloris RCC1774 HCAR (WP_110987361.1); P. hollandica HCAR (WP_017711629.1); Acaryochloris RCC1774 BciB (PZD72398.1); P. hollandica BciB (WP_044076442.1); Synechocystis PCC6803 (WP_010873198.1); Cyanidioschyzon merolae strain 10D (XP_005534820.1); Galdieria sulphuraria (XP_005706147.1); Chloroherpeton thalassium (WP_012499756.1); Chlorobium limicola (WP_059139293.1); P. phaeoclathratiforme BU-1 (ACF44893.1). Fig. 3 Open in new tabDownload slide Phylogenetic trees of CBR, HCAR and BciB. The phylogenetic trees were constructed by maximum likelihood. The numbers at each node represent the bootstrap value and the number of amino acid substitutions per site is illustrated by the scale bar. (A) Phylogenetic tree of CBR. Arabidopsis NYC1 (AT4G13250); Arabidopsis NOL (AT5G04900); Oryza sativa Japonica Group NOL (XP_015628274.1); O. sativa Japonica Group NYC1 (XP_015621887.1); Klebsormidium nitens NYC1 (GAQ77737.1); K. nitens NOL (GAQ87774.1); Coccomyxa subellipsoidea C-169 NYC1 (XP_005652224.1); C. subellipsoidea C-169 NOL (XP_005646276.1); Pelodictyon phaeoclathratiforme (WP_012508106.1); Ostreococcus lucimarinus CCE9901(XP_001415854.1 ); Micromonas commoda (XP_002508859.1); Chlamydomonas reinhardtii NYC1 (XP_001697080.1); C. reinhardtii NOL (XP_001701347.1); Vitrella brassicaformis CCMP3155 (CEM24690.1); Candidatus Heimdallarchaeota archaeon AB_125 (OLS31532.1); Chondrus crispus (XP_005716045.1); Chlorobaculum tepidum (WP_010932815.1); Chlorobaculum limnaeum (WP_069809375.1); Fistulifera solaris (GAX23003.1); P. hollandica (WP_081599361.1); Acaryochloris RCC1774 (1) (WP_110987898.1); Acaryochloris RCC1774 (2) (WP_110986784.1); Synechocystis PCC6803 (WP_041428273.1). (B) Phylogenetic tree of BciB and HCAR. C. reinhardtii (PNW76723.1); M. commoda (XP_002503439.1); O. lucimarinus CCE9901 (XP_001416225.1); C. subellipsoidea C-169 (XP_005648937.1); K. nitens (GAQ88093.1); Physcomitrella patens (XP_024368500.1); Selaginella moellendorffii (XP_024538910.1); O. sativa Japonica Group (XP_015636785.1); Arabidopsis HCAR (AT1G04620.1); Acaryochloris RCC1774 HCAR (WP_110987361.1); P. hollandica HCAR (WP_017711629.1); Acaryochloris RCC1774 BciB (PZD72398.1); P. hollandica BciB (WP_044076442.1); Synechocystis PCC6803 (WP_010873198.1); Cyanidioschyzon merolae strain 10D (XP_005534820.1); Galdieria sulphuraria (XP_005706147.1); Chloroherpeton thalassium (WP_012499756.1); Chlorobium limicola (WP_059139293.1); P. phaeoclathratiforme BU-1 (ACF44893.1). CBR homologs of red algae and diatoms are most closely related to green plant CBRs phylogenetically, although they do not produce Chl b. Interestingly, green sulfur bacteria also have CBR homologs. Genes highly homologous to CBR were found only in photosynthetic organisms, suggesting that homologous genes are related to photosynthesis. CBR homologs of P. hollandica and Acaryochloris RCC1774 were most distantly related to green plant CBRs phylogenetically in this tree and formed a cluster with other cyanobacterial CBR homologs (Fig. 3A), although these two cyanobacteria have Chl b. This distribution profile of cyanobacterial CBR homologs does not indicate whether they are genuine orthologs that encode functional CBR or not. In the Chl cycle, HMChl a is converted to Chl a by HCAR. We retrieved its homologous genes using Arabidopsis HCAR (AT1G04620) as a query. Many homologous proteins were found in photosynthetic eukaryotes and cyanobacteria. The phylogenetic tree of these proteins was separated into two clusters, one is HCAR and the other is 8-vinyl reductase (BciB, also known as F-DVR; Ito et al. 2008, Liu and Bryant 2011). Green plants have HCAR but not BciB because they employ BciA (also known as N-DVR) instead of BciB. Cyanobacteria use BciB instead of BciA and most of them have neither Chl b nor HCAR. However, cyanobacteria containing Chl b, P. hollandica and Acaryochloris RCC1774, have two homologous genes, one of which belongs to the HCAR cluster and the other to the BciB cluster (Fig. 3B). This phylogenetic analysis implicates that P. hollandica and Acaryochloris RCC1774 have HCAR. Enzymatic analysis of the candidate genes of P. hollandica and Acaryochloris RCC1774 To clarify whether CBR homologs of P. hollandica and Acaryochloris RCC1774 have CBR activity or not, the recombinant proteins encoded by these genes were expressed in Escherichia coli. Immunoblotting analysis showed that these proteins were successfully expressed in E. coli and found in a soluble fraction (Fig. 4). Fig. 4 Open in new tabDownload slide Immunoblotting analysis of recombinant proteins using a specific antibody against the histidine tag. Candidate genes of CBR, HCAR or BciB were expressed in E. coli and soluble fraction of the cell lysate was analyzed by immunoblotting. The markers for molecular size are shown in the left side. The cell lysate of E. coli having an empty vector (pET30a) was employed for the negative standard. PhCBR, P. hollandica CBR; AcaCBR, Acaryochloris RCC1774 CBR; PhHCAR, P. hollandica HCAR; AcaHCAR, Acaryochloris RCC1774 HCAR; PhBciB, P. hollandica BciB; AcaBciB, Acaryochloris RCC1774 BciB; SynBciB, Synechocystis BciB; AtNOL, Arabidopsis NOL; AtHCAR, Arabidopsis HCAR. Fig. 4 Open in new tabDownload slide Immunoblotting analysis of recombinant proteins using a specific antibody against the histidine tag. Candidate genes of CBR, HCAR or BciB were expressed in E. coli and soluble fraction of the cell lysate was analyzed by immunoblotting. The markers for molecular size are shown in the left side. The cell lysate of E. coli having an empty vector (pET30a) was employed for the negative standard. PhCBR, P. hollandica CBR; AcaCBR, Acaryochloris RCC1774 CBR; PhHCAR, P. hollandica HCAR; AcaHCAR, Acaryochloris RCC1774 HCAR; PhBciB, P. hollandica BciB; AcaBciB, Acaryochloris RCC1774 BciB; SynBciB, Synechocystis BciB; AtNOL, Arabidopsis NOL; AtHCAR, Arabidopsis HCAR. In P. hollandica, we found one candidate gene, WP_081599361, for a potential CBR gene by phylogenetic analysis. When the recombinant protein of WP_081599361 was incubated with Chl b, we detected two new peaks in the HPLC chromatograms for pigment analysis (Fig. 5A, line 5), neither of which were found in the negative control experiment (Fig. 5A, line 4). The retention times and absorption spectra of Peak 1 and Peak 2 matched those of HMChl a and its C13-2 epimer form, i.e. HMChl a′ (Fig. 5B). The activity was diminished by the depletion of NADPH (Fig. 5A, line 6) indicating that the protein uses NADPH as a reductant like Arabidopsis CBR. Fig. 5 Open in new tabDownload slide Enzymatic analysis of CBR of P. hollandica and Acaryochloris RCC1774. (A) HPLC profiles of the pigments after incubation of Chl with recombinant proteins. Chl b was incubated with the lysate of E. coli expressing PhCBR and AcaCBR with or without NADPH. Detected peaks of each pigment at 440 nm were identified by their retention time and absorption spectrum. Arabidopsis NOL was used as a positive control. 1, Chl b; 2, HMChl a; 3, lysate of E. coli expressing Arabidopsis NOL and Chl b, incubated with NADPH; 4, lysate of E. coli having pET30a and Chl b, incubated with NADPH; 5, lysate of E. coli expressing PhCBR and Chl b, incubated with NADPH; 6, lysate of E. coli expressing PhCBR and Chl b, incubated without NADPH; 7, lysate of E. coli expressing AcaCBR and Chl b, incubated with NADPH; 8, lysate of E. coli expressing AcaCBR and Chl b, incubated without NADPH. AtNOL, Arabidopsis NOL; PhCBR, P. hollandica CBR; AcaCBR, Acaryochloris RCC1774 CBR. (B) The absorption spectrum of HMChl a, where Peak 1 and Peak 2 were found in the reaction mixture of the lysate of E. coli expressing PhCBR and NADPH, as illustrated in (A). Fig. 5 Open in new tabDownload slide Enzymatic analysis of CBR of P. hollandica and Acaryochloris RCC1774. (A) HPLC profiles of the pigments after incubation of Chl with recombinant proteins. Chl b was incubated with the lysate of E. coli expressing PhCBR and AcaCBR with or without NADPH. Detected peaks of each pigment at 440 nm were identified by their retention time and absorption spectrum. Arabidopsis NOL was used as a positive control. 1, Chl b; 2, HMChl a; 3, lysate of E. coli expressing Arabidopsis NOL and Chl b, incubated with NADPH; 4, lysate of E. coli having pET30a and Chl b, incubated with NADPH; 5, lysate of E. coli expressing PhCBR and Chl b, incubated with NADPH; 6, lysate of E. coli expressing PhCBR and Chl b, incubated without NADPH; 7, lysate of E. coli expressing AcaCBR and Chl b, incubated with NADPH; 8, lysate of E. coli expressing AcaCBR and Chl b, incubated without NADPH. AtNOL, Arabidopsis NOL; PhCBR, P. hollandica CBR; AcaCBR, Acaryochloris RCC1774 CBR. (B) The absorption spectrum of HMChl a, where Peak 1 and Peak 2 were found in the reaction mixture of the lysate of E. coli expressing PhCBR and NADPH, as illustrated in (A). Acaryochloris RCC1774 has two candidates, WP_110987898 and WP_110986784, for a potential CBR gene. It should be noted that the PZD74811 locus encodes a peptide sequence similar to the second half of the CBR sequences but it appears to lack the first half. Therefore, we did not include this gene for further analysis. We examined the recombinant protein of WP_110986784 activity and found its CBR activity (Fig 5A, line 7). When NADPH was omitted from the assay mixture, these two peaks did not appear (Fig. 5A, line 8), which indicate that the appearance of these peaks is dependent on enzymatic activity. Taken together, we concluded that the gene product of Acaryochloris RCC1773 WP_110986784 has CBR activity. By contrast, we could not detect CBR activity with the gene product of WP_110987898. These results indicate that P. hollandica and Acaryochloris RCC1774 have CBR orthologs. A significant amount of the epimer type of HMChl a was formed after incubation of Chl a with P. hollandica and Acaryochloris RCC1774 CBRs (Fig. 5A). By contrast, HPLC analysis showed that Chl a′ (an epimer of Chl a) was not accumulated in a large amount in cells (Fig. 2). We speculate that the production of this epimer could be derived from Chl b′ produced by incubation with E. coli lysate, and then it was converted to HMChl a′ by CBR. Alternatively, the discrepancy could be explained by assuming some mechanism that suppresses the CBR-mediated formation of HMChl a′ in the cell. Phylogenetic analysis suggests that P. hollandica WP_017711629 and Acaryochloris RCC1774 WP_110987361 are HCARs. Recombinant proteins of these genes were expressed in E. coli, and the cell extract was incubated with HMChl a. HPLC analysis showed that HMChl a was converted to Chl a by these proteins (Fig. 6A, line 6 and line 8). The production of Chl a was also confirmed by the detection of the pigment by fluorescence at 680 nm with excitation at 430 nm (Supplementary Fig. S2). These results indicate that the gene products of P. hollandica WP_017711629 and Acaryochloris RCC1774 WP_110987361 have HCAR activity. Taken together, it is shown that P. hollandica and Acaryochloris RCC1774 have the potential to complete the Chl cycle. Fig. 6 Open in new tabDownload slide Enzymatic analysis of HCAR of P. hollandica and Acaryochloris RCC1774. (A) HPLC profiles of the pigments after incubation of Chl with recombinant proteins. HMChl a was incubated with the lysate of E. coli expressing PhHCAR and AcaHCAR with and without NADPH, FNR and Fd. Detected peaks of each pigment at 440 nm were identified by their retention time and absorption spectrum. 1, HMChl a; 2, Chl a; 3, divinyl Chl a; 4, lysate of E. coli having Arabidopsis HCAR and HMChl a, incubated with NADPH, FNR and Fd; 5, lysate of E. coli having pET30a and HMChl a, incubated with NADPH, FNR and Fd; 6, lysate of E. coli expressing PhHCAR and HMChl a, incubated with NADPH, FNR and Fd; 7, lysate of E. coli expressing PhHCAR and HMChl a, incubated without reductant; 8, lysate of E. coli expressing AcaHCAR and HMChl a, incubated with NADPH, FNR, and Fd; 9, lysate of E. coli expressing AcaHCAR and HMChl a, incubated without reductant. DV-Chl a, divinyl Chl a. AtHCAR, Arabidopsis HCAR; PhHCAR, P. hollandica HCAR; AcaHCAR, Acaryochloris RCC1774 HCAR. (B) HPLC profiles of the pigments after incubation of Chl with recombinant proteins. Divinyl Chl a was incubated with the lysate of E. coli expressing PhHCAR, AcaHCAR and SynBciB with NADPH, FNR and Fd. Detected peaks of each pigment at 440 nm were identified by their retention time and absorption spectrum. 1, lysate of E. coli expressing PhHCAR and divinyl Chl a, incubated with NADPH, FNR, and Fd; 2, lysate of E. coli expressing AcaHCAR and divinyl Chl a, incubated with NADPH, FNR, and Fd; 3, lysate of E. coli expressing SynBciB and divinyl Chl a, incubated with NADPH, FNR, and Fd. SynBciB, Synechocystis BciB. (C) The absorption spectrum of Chl a, where Peak 1 and Peak 2 were found in the reaction mixture of the lysate of E. coli expressing PhHCAR and AcaHCAR with reductant, as illustrated in (A), and Peak3, Peak 4 and Peak 5 were found in the reaction mixture of the lysate of E. coli expressing PhHCAR, AcaHCAR and SynBciB with reductant, as illustrated in (B). Fig. 6 Open in new tabDownload slide Enzymatic analysis of HCAR of P. hollandica and Acaryochloris RCC1774. (A) HPLC profiles of the pigments after incubation of Chl with recombinant proteins. HMChl a was incubated with the lysate of E. coli expressing PhHCAR and AcaHCAR with and without NADPH, FNR and Fd. Detected peaks of each pigment at 440 nm were identified by their retention time and absorption spectrum. 1, HMChl a; 2, Chl a; 3, divinyl Chl a; 4, lysate of E. coli having Arabidopsis HCAR and HMChl a, incubated with NADPH, FNR and Fd; 5, lysate of E. coli having pET30a and HMChl a, incubated with NADPH, FNR and Fd; 6, lysate of E. coli expressing PhHCAR and HMChl a, incubated with NADPH, FNR and Fd; 7, lysate of E. coli expressing PhHCAR and HMChl a, incubated without reductant; 8, lysate of E. coli expressing AcaHCAR and HMChl a, incubated with NADPH, FNR, and Fd; 9, lysate of E. coli expressing AcaHCAR and HMChl a, incubated without reductant. DV-Chl a, divinyl Chl a. AtHCAR, Arabidopsis HCAR; PhHCAR, P. hollandica HCAR; AcaHCAR, Acaryochloris RCC1774 HCAR. (B) HPLC profiles of the pigments after incubation of Chl with recombinant proteins. Divinyl Chl a was incubated with the lysate of E. coli expressing PhHCAR, AcaHCAR and SynBciB with NADPH, FNR and Fd. Detected peaks of each pigment at 440 nm were identified by their retention time and absorption spectrum. 1, lysate of E. coli expressing PhHCAR and divinyl Chl a, incubated with NADPH, FNR, and Fd; 2, lysate of E. coli expressing AcaHCAR and divinyl Chl a, incubated with NADPH, FNR, and Fd; 3, lysate of E. coli expressing SynBciB and divinyl Chl a, incubated with NADPH, FNR, and Fd. SynBciB, Synechocystis BciB. (C) The absorption spectrum of Chl a, where Peak 1 and Peak 2 were found in the reaction mixture of the lysate of E. coli expressing PhHCAR and AcaHCAR with reductant, as illustrated in (A), and Peak3, Peak 4 and Peak 5 were found in the reaction mixture of the lysate of E. coli expressing PhHCAR, AcaHCAR and SynBciB with reductant, as illustrated in (B). Catalytic specificity of HCAR and BciB Previous studies showed that HCAR and BciB show sequence similarity (Ito and Tanaka 2014). A phylogenetic analysis indicates that HCAR arose within the cluster of BciB during evolution. Interestingly, it was shown that cyanobacterial BciB has promiscuous HCAR activity (Ito and Tanaka 2014), which might have enabled the enzyme evolution from BciB to HCAR. By contrast, green plant HCAR has no 8-vinyl reductase activity, although these two enzymes have high sequence similarity, which is hypothesized to be a result of evolutional fitting to the new substrate (HMChl a; Ito and Tanaka 2014). We hypothesized that promiscuous activity (such as HCAR activity in Synechocystis BciB) is only subjected to evolutional selection when this activity competes with the genuine activity, such as that in green algae and plants. To test this hypothesis, we examined the substrate specificity of the four gene products, Acaryochloris RCC1774 BciB (PZD72398), P. hollandica BciB (WP_044076442), Acaryochloris RCC1774 HCAR (WP_110987361) and P. hollandica HCAR (WP_017711629). As a control experiment, Synechocystis PCC6803 (hereafter Synechocystis) BciB (Slr1923) was used in the HCAR and BciB assays. In this study, the same volumes of the soluble fraction of E. coli lysate were used for the enzymatic assay. Exceptionally, the extract containing recombinant Synechocystis BciB was diluted by four times to adjust the protein levels (Fig. 4), because the protein level of Synechocystis BciB was much higher than those of other proteins. We tested promiscuous HCAR activity of BciB from P. hollandica and Acaryochloris RCC1774. Instead of HMChl a, we used 7-hydroxymethyl chlorophyllide a (HMChlide a) for the HCAR assay of BciB because our previous experiment (Ito and Tanaka 2014) showed that Synechocystis BciB prefers this pigment to HMChl a in the in vitro HCAR assay. We confirmed that the recombinant BciB proteins of P. hollandica and Acaryochloris RCC1774 are able to use Chlide as a substrate for 8-vinyl group reduction (BciB) reaction (Supplementary Fig. S3). After incubation of BciB proteins of P. hollandica and Acaryochloris RCC1774 with HMChlide a, Chlide a was not detected (Fig. 7B, line 3 and line 4), whereas BciB of Synechocystis produced Chlide a after incubation (Fig. 7B, line 5). Fig. 7 Open in new tabDownload slide Enzymatic analysis of BciB of P. hollandica and Acaryochloris RCC1774. (A) HPLC profiles of the pigments after incubation of Chl with recombinant proteins. Divinyl Chl a was incubated with the lysate of E. coli expressing PhBciB and AcaBciB with or without NADPH, FNR and Fd. Detected peaks of each pigment at 440 nm were identified by their retention time and absorption spectrum. 1, Divinyl Chl a; 2, Chl a; 3, lysate of E. coli having pET30a and divinyl Chl a, incubated with NADPH, FNR and Fd; 4, lysate of E. coli expressing PhBciB and divinyl Chl a, incubated with NADPH, FNR and Fd; 5, lysate of E. coli expressing PhBciB and divinyl Chl a, incubated without reductant; 6. lysate of E. coli expressing AcaBciB and divinyl Chl a, incubated with NADPH, FNR and Fd; 7, lysate of E. coli expressing AcaBciB and divinyl Chl a, incubated without reductant. (B) HPLC profiles of the pigments after incubation of Chlide with recombinant proteins. HMChlide a was incubated with the lysate of E. coli expressing PhBciB, AcaBciB and SynBciB with NADPH, FNR and Fd. Detected peaks of each pigment at 680 nm fluorescence were identified by their retention time. The arrow indicates the produced Chlide a. 1, HMChlide a; 2, Chlide a; 3, lysate of E. coli expressing PhBciB and HMChlide a, incubated with NADPH, FNR and Fd; 4, lysate of E. coli expressing AcaBciB and HMChlide a, incubated with NADPH, FNR and Fd; 5, lysate of E. coli expressing SynBciB and HMChlide a, incubated with NADPH, FNR, and Fd. DV-Chl, divinyl Chl a; PhBciB, P. hollandica BciB; AcaBciB, Acaryochloris RCC1774 BciB; SynBciB, Synechocystis BciB. (C) The absorption spectrum of Chl a, where Peak 1, Peak 2 and Peak 3 were found in the reaction mixture of the lysate of E. coli expressing PhBciB and AcaBciB with or without reductant, as illustrated in (A). Fig. 7 Open in new tabDownload slide Enzymatic analysis of BciB of P. hollandica and Acaryochloris RCC1774. (A) HPLC profiles of the pigments after incubation of Chl with recombinant proteins. Divinyl Chl a was incubated with the lysate of E. coli expressing PhBciB and AcaBciB with or without NADPH, FNR and Fd. Detected peaks of each pigment at 440 nm were identified by their retention time and absorption spectrum. 1, Divinyl Chl a; 2, Chl a; 3, lysate of E. coli having pET30a and divinyl Chl a, incubated with NADPH, FNR and Fd; 4, lysate of E. coli expressing PhBciB and divinyl Chl a, incubated with NADPH, FNR and Fd; 5, lysate of E. coli expressing PhBciB and divinyl Chl a, incubated without reductant; 6. lysate of E. coli expressing AcaBciB and divinyl Chl a, incubated with NADPH, FNR and Fd; 7, lysate of E. coli expressing AcaBciB and divinyl Chl a, incubated without reductant. (B) HPLC profiles of the pigments after incubation of Chlide with recombinant proteins. HMChlide a was incubated with the lysate of E. coli expressing PhBciB, AcaBciB and SynBciB with NADPH, FNR and Fd. Detected peaks of each pigment at 680 nm fluorescence were identified by their retention time. The arrow indicates the produced Chlide a. 1, HMChlide a; 2, Chlide a; 3, lysate of E. coli expressing PhBciB and HMChlide a, incubated with NADPH, FNR and Fd; 4, lysate of E. coli expressing AcaBciB and HMChlide a, incubated with NADPH, FNR and Fd; 5, lysate of E. coli expressing SynBciB and HMChlide a, incubated with NADPH, FNR, and Fd. DV-Chl, divinyl Chl a; PhBciB, P. hollandica BciB; AcaBciB, Acaryochloris RCC1774 BciB; SynBciB, Synechocystis BciB. (C) The absorption spectrum of Chl a, where Peak 1, Peak 2 and Peak 3 were found in the reaction mixture of the lysate of E. coli expressing PhBciB and AcaBciB with or without reductant, as illustrated in (A). Subsequently, we tested the 8-vinyl reductase (BciB) activity of HCAR recombinant proteins. After incubation of HCARs of P. hollandica and Acaryochloris RCC1774 with divinyl Chl a, the monovinyl form of Chl a was not detected (Fig. 6B). We noted that the absorbance spectra of minor peaks (Peak 3 and Peak 4) were similar to those of divinyl Chl a (Fig. 6C), indicating that they are most likely divinyl Chl a′, which might have been formed during the assay. The lack of promiscuous BciB activity in HCAR recombinant proteins was also examined using divinyl Chlide a as a substrate. We confirmed that these proteins are able to reduce HMChlide a (Supplementary Fig. S4, line 5 and line 7) in our assay conditions, suggesting that they accept Chlide as a substrate. The monovinyl form of Chlide a was not detected after the 8-vinyl reductase (BciB) assay (Supplementary Fig. S4, line 12 and line 13). The results show that P. hollandica and Acaryochloris RCC1774 HCARs do not have BciB activity in these assay conditions. Collectively, our results indicate that HCAR from P. hollandica and Acaryochloris RCC1774 did not show detectable promiscuous activity. These results support our evolutional fitting hypothesis mentioned above. Discussion The Chl cycle in cyanobacteria containing Chl b We showed in this report that cyanobacteria containing Chl b, P. hollandica and Acaryochloris RCC1774, have CAO, CBR and HCAR orthologs, and the latter two of them showed CBR and HCAR enzymatic activities, respectively (Figs. 5, 6). These results indicate that cyanobacteria containing Chl b have the potential to complete the Chl cycle. Acaryochloris RCC1774 and P. hollandica grow in shallow seas and freshwater, respectively, where light conditions change dynamically. It is likely that these two organisms need to control the antenna size in response to light conditions, as in green algae. In P. hollandica, Chl b is localized in Chl a/b-binding proteins (encoded by PCB genes) which are not related to LHC (Bumba et al. 2005, Herbstová et al. 2010). The antenna size and Chl a/b ratio are dynamically changed in response to light intensity in P. hollandica (Burger-Wiersma and Post 1989). However, the mechanisms of antenna size regulation are different between P. hollandica and green plants, because the Chl a/b ratio becomes low under high light conditions in P. hollandica (Burger-Wiersma and Post 1989) but high in green plants. There are no reports concerning the changes in Chl a/b ratio in Acaryochloris RCC1774, whereas other Acaryochloris species containing Chl d change Chl d/a ratios in response to light conditions (Duxbury et al. 2009). If a change in the Chl a/b ratio is linked to the antenna size regulation in those cyanobacteria, the Chl cycle might play a role in light acclimation. In plants, the Chl b-to-a conversion is an important step for Chl breakdown during leaf senescence (Kusaba et al. 2007), because the Chl-degrading enzymes of plants cannot catabolize Chl b-type pigments that contain a formyl group at the C7 position. It is tempting to speculate that the Chl cycle could also play an important role in Chl b degradation in cyanobacteria containing Chl b. Physiological and genetic evidence will be necessary to determine if the Chl cycle functions for light acclimation and Chl degradation in these cyanobacteria. We could not find any HCAR or BciB homologs in Prochlorococcus. This cyanobacterium uses divinyl Chl instead of monovinyl Chl (Barrera-Rojas et al. 2018); therefore, a lack of BciB homologs would be reasonable. Our analysis presented in this study indicates that Prochlorococcus does not have the Chl cycle either. It is reported that Prochlorococcus has evolved into high-light-adapted and low-light-adapted strains (Rocap et al. 2003), which have genetically adapted to different light niches. The low-light-adapted Prochlorococcus strain is found in deep seas. It has a large number of PCB genes and a low Chl a/b ratio. By contrast, the high-light-adapted Prochlorococcus strain has a high Chl a/b ratio. Therefore, there is no strong demand for an ability to change their antenna size dynamically. It is also reported that Prochlorococcus evolved to reduce their genome size to decrease the demand of nitrogen in oligotrophic environments (García-Fernández et al. 2004), which might also account for the absence of the Chl cycle in Prochlorococcus. One potential drawback in losing the Chl cycle might be the loss of the ability to degrade Chl b. This is because one of the enzymes of Chl degradation pathway (pheophorbide a oxygenase) does not catalyze the oxidation of the substrate if the pheophorbide has a formyl group on C7 position (Hörtensteiner et al. 1995). At present, it is not clear whether Prochlorococcus has a novel Chl b-to-a conversion pathway or a distinct Chl breakdown pathway, which is able to catabolize Chl b without converting it to Chl a. It is also possible to speculate that cyanobacterial pheophorbide a oxygenase accepts Chl b-derived pigments, as Auxenochlorella protothecoides (Chlorella protothecoides) produces red Chl catabolites with the C7 formyl group (Iturraspe et al. 1994). Otherwise, it is also possible to assume that the bacterium simply excretes Chl b-type molecules. Further studies are needed to elucidate Chl degradation in Prochlorococcus. Acaryochloris containing Chl b has the genes having the enzymatic activities to complete the Chl cycle by which Chl a and Chl b are interconverted. Other Acaryochloris produces Chl d instead of Chl b (Partensky et al. 2018). These Acaryochloris are expected to have the interconversion pathway of Chl a and Chl d, which would be beneficial for altering the Chl a/d ratio corresponding to the intensity of far-red light (Duxbury et al. 2009). It should be noted that Acaryochloris marina has both BciA and BciB and these two genes are considered to be involved in Chl biosynthesis (Chen et al. 2016). Identification of the enzymes responsible for the Chl a and Chl d interconversion is indispensable for understanding the acclimation of Acaryochloris containing Chl d. Evolution of the enzymes of the Chl cycle It is suggested that the promiscuous activity of the enzymes is kept at a low level if the activity is harmful to the cells (Khersonsky and Tawfik 2010). The BciB of Synechocystis has high sequence similarity to green plant HCAR and the BciB has promiscuous HCAR activity (Ito and Tanaka 2014). This promiscuous activity is not harmful to cyanobacteria because they do not have the substrate (HMChl a) for this promiscuous activity in the cell. This broad substrate specificity and catalytic activity would contribute to the enzyme to have high catalytic activity of the primary reaction because high specificity accompanies the low catalytic activity (Khersonsky and Tawfik 2010). In this study, Synechocystis BciB showed promiscuous HCAR activities, whereas neither of P. hollandica and Acaryochloris RCC1774 BciBs showed this activity in our assay conditions. Nevertheless, the possibility cannot be excluded that these BciB homologs show HCAR under different assay conditions. Khersonsky and Tawfik (2010) assume that if a promiscuous activity is harmful, the enzyme evolves to lose this activity. By contrast, if the promiscuous activity is not harmful, the activity will be retained (Khersonsky and Tawfik 2010). We hypothesize that the BciB of P. hollandica and Acaryochloris RCC1774 have evolved to gain higher substrate specificity and have lost their promiscuous HCAR activity, in contrast to Synechocystis BciB. We speculate that having both genuine and promiscuous HCAR activities in the same cell might interfere with the regulation of the Chl cycle in Chl b-containing cyanobacteria, while promiscuous HCAR activity should not have any harm in Chl b-less cyanobacteria. The same phenomenon is observed with HCAR of the green plants and cyanobacteria containing Chl b. These HCARs have no BciB activity because these organisms have another 8-vinyl reductase. Our phylogenetic analysis (Fig. 3B) shows that BciB and HCAR homologs form different clades. One scenario to explain this result is that BciB was duplicated in a cyanobacterium and evolved to possess more specific substrate recognition ability of HMChl a in Chl b-containing cyanobacteria. It is possible to assume that a common ancestor of Chl b-containing cyanobacteria had a duplicated BciB ortholog, or that a BciB ortholog, which had HCAR activity was horizontally transferred to one of the ancestors of Chl b-containing cyanobacteria. HCARs of cyanobacteria and green plants form a single cluster, suggesting that green plant HCAR evolved from cyanobacterial HCAR. Considering the similar tree topologies of CAO and HCAR sequences, it is tempting to hypothesize that HCAR might be transferred with CAO during an endosymbiotic event. The phylogenetic tree of CBR is more complicated: CBRs do not form a single cluster (Fig. 3A). It is not evident from the phylogenetic tree whether green plant CBR is derived from cyanobacterial CBR or appeared independently. In either case, green plant CBR is assumed to be derived from cyanobacterial genes. In eukaryotes, red algae and brown algae have homologs of green plant CBR. The tree topology of CBR indicates that CBR homologs of red and brown algae are evolved from CBR. It is not clear at this stage whether an ancestor of those algae that do not produce Chl b but contain CBR homologs had synthesized Chl b or not. It should be noted that we also found homologs in green filamentous photosynthetic bacteria. A plausible explanation for the distribution of CBR homologs in the organisms that do not produce Chl b would be that those organisms obtain CBR homologs by either vertical or horizontal transfer at a certain stage of evolution, but they evolved to show other enzymatic activities than the reduction of Chl b. It is known that the short-chain dehydrogenase family to which CBR belongs shows remarkable diversification to catalyze various reactions (Kavanagh et al. 2008). The distribution of CBR may illustrate one example of such diversification of the short-chain dehydrogenase family. Materials and Methods Phylogenetic tree analysis The protein sequences were obtained from the databases Phytozome (https://phytozome.jgi.doe.gov/pz/portal.html) and NCBI (https://www.ncbi.nlm.nih.gov/). We used a broad variety of organisms from prokaryotes to green plants to construct a reasonable phylogenetic tree. CBR, HCAR and its homolog sequences were digested by M-Coffee and evaluated (Moretti et al. 2007). In the M-Coffee analysis, we only employed the amino acid residues which assessed the ‘good’ (displayed by red color) and the leftovers, which were tagged either ‘BAD’ or ‘AVG’, were clipped off from the alignment in each protein (Supplementary Tables S1, S2). The phylogenetic trees were constructed by using the maximum likelihood model and IQ-TREE version 1.6.9 (Trifinopoulos et al. 2016). Phylogenetic analysis was operated based on the 1,000 bootstrap replicants in ultrafast mode. The best-fitting amino acid substitution model was searched and employed automatically; LG+G4 was applied for both proteins. Strains and culture conditions Acaryochloris RCC1774 was obtained from the Roscoff Culture Collection. Acaryochloris RCC1774 and P. hollandica were grown in IMK medium (Nihon Pharmaceutical, Tokyo, Japan) and BG11 medium at 23°C under a 16 h photoperiod at a light condition of 2.5 µmol photons·m−2·s−1 without shaking. Extraction and analysis of pigments Cells were harvested by centrifuging at 20,000 × g for 1 min. The pellet was suspended in methanol and centrifuged at 20,000 × g for 10 min. The supernatant was immediately subjected to HPLC equipped with a diode array detector (SPD-M10A, Shimadzu, Kyoto, Japan) or a fluorescence detector (RF-20A, Shimadzu). The pigments were separated through a Symmetry C8 column (⁠ 4.6 × 150 mm ⁠, Waters, Milford, MA) with a gradient from eluent A [methanol:acetonitrile:aqueous pyridine solution (0.25 M pyridine; 50:25:25 v:v:v)] to eluent B [methanol:acetonitrile:acetone (20:60:20 v:v:v)] at a flow rate of 1 ml·min−1 at 40°C (Shimoda et al. 2016). Detected peaks of each pigment at 440 nm were identified by their retention time and absorption spectrum. The absorption spectrum of each peak was corrected from the background absorbance. Cloning of CBR, HCAR and BciB from Acaryochloris RCC1774 and P. hollandica Each coding region of CBR, HCAR and BciB derived from Acaryochloris RCC1774 (WP_110986784.1, PZD72038.1 and PZD72398.1) and P. hollandica (WP_081599361.1, WP_017711629.1 and WP_044076442.1) was amplified by polymerase chain reaction from the genomic DNA of each species. The primers used for amplification are shown in Supplementary Table S3. Amplified genes are cloned into pET-30a (+) vectors (Novagen, Madison, WI) containing a His6 tag at the C terminus using the NdeI and XhoI sites through an in-fusion cloning system (Clontech, Mountain View, CA). Synechocystis BciB (Slr1923) was prepared as reported previously (Ito et al. 2008). Expression and detection of recombinant proteins The constructed plasmids for protein expression were introduced into E. coli (BL21). Escherichia coli was grown and recombinant protein was expressed in an auto-induction medium (Studier 2005) at 18°C with 130 rpm shaking. When the cell was fully saturated, 500 μl of the cell was harvested by centrifuge at 20,000 × g for 2 min. The pellet was resuspended with 500 μl of BugBuster Protein extraction reagents (Novagen) with 0.1% benzonase (Novagen). Immunoblotting analysis was employed to determine the expression of the recombinant proteins because expression levels were too low to be detected by Coomassie Blue Brilliant staining. After centrifugation at 20,000 × g for 10 min, the supernatant of the E. coli lysate was mixed with the same amount of sample buffer for SDS-PAGE [125 mM Tris-HCl, pH 6.8, 4% (w/v) SDS, 10% (w/v) sucrose, 5% (v/v) 2-mercaptoehanol, and a little bit of bromophenol blue], and subjected to SDS-PAGE. For immunoblotting analysis, proteins were transferred to a polyvinylidene difluoride film. Using the antibodies for Histidine tag (Anti-His-tag mAb-HRP-DirecT, MBL, Nagoya, Japan) and Western Lighting Plus-ECL (PerkinElmer Life Science, Waltham, MA), proteins were detected by fluorescence imaging. Preparation of the Chl derivatives Divinyl Chl was prepared from a Synechocystis mutant that lacks BciB (Slr1923; Ito et al. 2008). HMChl a was prepared through the reduction of Chl b using NaBH4 according to a previous report (Holt 1959). Chlide derivatives were prepared by removing the phytyl chain through hydrolysis by chlorophyllase (Ito and Tanaka 2014). Enzyme assay Escherichia coli lysate (50 µl) prepared as described above was used for the enzymatic assay. For CBR analysis, we added 1 µl of 50 mM NADPH. For HCAR and BciB analysis, we provided 1 µl of spinach ferredoxin-NADP+ reductase (FNR; 0.1 mg·ml−1, Sigma-Aldrich, St. Louis, MO), 1 µl of spinach ferredoxin (Fd; 1 mg·ml−1, Sigma-Aldrich) and 1 µl of 50 mM NADPH to E. coli lysate. The pigments were solubilized with DMSO and used 0.5 µl of the solution, which contains 500 pmol of pigments. The mixture was incubated at 37°C for 1 h, and the reaction was stopped by adding 200 µl of acetone. After centrifugation at 20,000 × g for 10 min, the supernatant was analyzed by HPLC. Funding JSPS, KAKENHI [15H04381 to A.T.; 16H06554 and 17K07431 to R.T.; and 17K07430 to H.I.]. Acknowledgments We thank Dr. Atsushi Takabayashi (Hokkaido University) for assistance with phylogenetic analysis. Disclosures The authors have no conflicts of interest to declare. References Bailey S. , Walters R. , Jansson S. , Horton P. ( 2001 ) Acclimation of Arabidopsis thaliana to the light environment: the existence of separate low light and high light responses . 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For permissions, please email: [email protected] This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Ohashi-Ito,, Kyoko;Iwamoto,, Kuninori;Nagashima,, Yoshinobu;Kojima,, Mikiko;Sakakibara,, Hitoshi;Fukuda,, Hiroo

doi: 10.1093/pcp/pcz156pmid: 31392340

Abstract The phytohormone auxin governs various developmental processes in plants including vascular formation. Auxin transport and biosynthesis are important factors in determining auxin distribution in tissues. Although the role of auxin transport in vein pattern formation is widely recognized, that of auxin biosynthesis in vascular development is poorly understood. Heterodimer complexes comprising two basic helix–loop–helix protein families, LONESOME HIGHWAY (LHW) and TARGET OF MONOPTEROS5 (TMO5)/TMO5-LIKE1 (T5L1), are master transcriptional regulators of the initial process of vascular development. The LHW–TMO5/T5L1 dimers regulate vascular initial cell production, vascular cell proliferation and xylem fate determination in the embryo and root apical meristem (RAM). In this study, we investigated the function of local auxin biosynthesis in initial vascular development in RAM. Results showed that LHW–T5L1 upregulated the expression of YUCCA4 (YUC4), a key auxin biosynthesis gene. The expression of YUC4 was essential for promoting xylem differentiation and vascular cell proliferation in RAM. Conversely, auxin biosynthesis was required for maintaining the expression levels of LHW, TMO5/T5L1 and their targets. Our results suggest that local auxin biosynthesis forms a positive feedback loop for fine-tuning the level of LHW–TMO5/T5L1, which is necessary for initiating vascular development. Introduction The formation of vascular initial cells first occurs in the lower middle part of the embryo at the globular stage. Recent studies demonstrate that heterodimer complexes comprising LONESOME HIGHWAY (LHW) and TARGET OF MONOPTEROS5 (TMO5)/TMO5-LIKE1 (T5L1) regulate cell divisions for the generation of vascular initial cells in an embryo, and upregulate cytokinin biosynthesis, which contributes to the proliferation of vascular stem cells in embryos and root apical meristem (RAM) (Ohashi-Ito and Bergmann 2007, De Rybel et al. 2013, Ohashi-Ito et al. 2013, De Rybel et al. 2014, Ohashi-Ito et al. 2014). The LHW–TMO5/T5L1 dimers also establish xylem cell identity (Katayama et al. 2015). Thus, LHW–TMO5/T5L1 are considered as key complexes regulating the initial process of vascular development in an embryo and RAM. In addition to LHW–TMO5/T5L1, auxin is another key regulator of initial vascular development (Cano-Delgado et al. 2010). Therefore, it is reasonable to speculate that LHW–TMO5/T5L1 and auxin signaling pathways are associated with each other in the initial process of vascular development. Indeed, TMO5 is a target of MONOPTEROS/AUXIN RESPONSE FACTOR5 (MP/ARF5; Schlereth et al. 2010) and is auxin-inducible. However, the functional relationship between LHW–TMO5/T5L1 and auxin and the role of transcription factor-dependent auxin biosynthesis in the initiation of vascular development are poorly understood. We found that YUCCA4 (YUC4) was listed as a candidate downstream gene of LHW–T5L1 in a transcriptome analysis of LHW–T5L1 overexpressing cells (Ohashi-Ito et al. 2014). Because YUC4 is well known as a key auxin biosynthetic enzyme (Cheng et al. 2006, Cheng et al. 2007), we here investigated the functional relationship between LHW–TMO5/T5L1 and auxin in initial vascular development, placing YUC4 at the heart of the analysis. Results To elucidate the role of YUC4 in vascular development, we first determined whether the expression of YUC4 was regulated by LHW and T5L1 in Arabidopsis culture cells simultaneously overexpressing LHW and T5L1-GFP under the control of an estrogen-inducible promoter. The expression of YUC4 was conspicuously upregulated >70-fold at 24 h after the addition of estrogen, with high expression levels at 48 and 72 h (Fig. 1A). Next, we performed chromatin immunoprecipitation (ChIP)-PCR assay using culture cells, which harboring the estrogen-inducible LHW/T5L1-GFP or the estrogen-inducible YFP. The estrogen-inducible YFP line was used as a control, and immunoprecipitates of this sample were the background level. Results of ChIP-PCR analysis showed that YUC4 promoter fragments, but not coding sequence fragments, were enriched in the immunoprecipitates of LHW and T5L1-GFP overexpressing cells (Fig. 1B, C), suggesting that T5L1 directly binds to the YUC4 promoter in the presence of LHW. Next, we generated plants harboring both pYUC4::GUS and estrogen-inducible LHW and T5L1-GFP constructs. Although GUS activity driven by the YUC4 promoter was restricted to the vascular region of the root in the absence of estrogen, ectopic GUS activity was observed in various root cells when the expression of LHW and T5L1 was induced after the addition of estrogen (Fig. 1D, E), indicating that the combination of LHW and T5L1 can induce YUC4 expression. Next, we examined the expression patterns of pYUC4::nuclear-localized YFP (nYFP) in wild-type and lhw roots. The nYFP signal driven by the YUC4 promoter was observed in metaxylem precursor cells in wild-type RAM (Fig. 1F, H). Because LHW is broadly expressed in RAM and TMO5 and T5L1 are expressed in xylem precursor cells, the LHW–T5L1 heterodimer is considered to function in xylem precursor cells in RAM (Ohashi-Ito and Bergmann 2007, De Rybel et al. 2013). In RAM, therefore, the YUC4 expressing cells overlapped with LHW–T5L1 functioning cells. In lhw RAM, the number of YUC4 expressing cells was dramatically decreased (Fig. 1G, I). Together, these results imply that LHW–T5L1 directly regulates YUC4 expression in the RAM. Fig. 1 Open in new tabDownload slide LHW–T5L1 regulates YUC4 expression. (A) Quantitative reverse transcription-PCR (RT-PCR) analysis of YUC4 expression levels in cultured cells harboring estrogen-inducible LHW and T5L1-GFP with (black) or without (light blue) estrogen. Data represent the mean ± standard deviation (SD). The mean is the average of three independent biological replicates. Significant differences were determined using Student’s t-test and are indicated with asterisks (***P < 0.001). (B) Schematic representation of the YUC4 gene. Black boxes indicate exons, and black bars indicate positions of primers used in the ChIP-PCR experiment. (C) Quantitative ChIP-PCR analysis using cells including estrogen-inducible LHW and T5L1-GFP (light blue, black), and cells including estrogen-inducible YFP (yellow, gray) with estrogen (black, gray) and without estrogen (light blue, yellow). Data represent mean ± SD. Significant differences between LHW–T5L1 Est− vs. YFP Est− and LHW–T5L1 Est+ vs. YFP Est+ were determined using Student’s t-test and are indicated with asterisks (**P < 0.01). The mean is the average of three independent biological replicates. (D, E) Expression patterns of pYUC4::GUS in the differentiation zone of roots of 8-day-old plants harboring estrogen-inducible LHW–T5L1 treated with (E) or without (D) estrogen for 24 h. Scale bar = 100 µm. (F–I) Expression patterns of pYUC4::nYFP in 5-day-old wild-type (F) and lhw (G) root tips. Cross-sections of (F) and (G) are shown in (H) and (I), respectively. Asterisks indicate protoxylem cells. Scale bar = 100 µm. Fig. 1 Open in new tabDownload slide LHW–T5L1 regulates YUC4 expression. (A) Quantitative reverse transcription-PCR (RT-PCR) analysis of YUC4 expression levels in cultured cells harboring estrogen-inducible LHW and T5L1-GFP with (black) or without (light blue) estrogen. Data represent the mean ± standard deviation (SD). The mean is the average of three independent biological replicates. Significant differences were determined using Student’s t-test and are indicated with asterisks (***P < 0.001). (B) Schematic representation of the YUC4 gene. Black boxes indicate exons, and black bars indicate positions of primers used in the ChIP-PCR experiment. (C) Quantitative ChIP-PCR analysis using cells including estrogen-inducible LHW and T5L1-GFP (light blue, black), and cells including estrogen-inducible YFP (yellow, gray) with estrogen (black, gray) and without estrogen (light blue, yellow). Data represent mean ± SD. Significant differences between LHW–T5L1 Est− vs. YFP Est− and LHW–T5L1 Est+ vs. YFP Est+ were determined using Student’s t-test and are indicated with asterisks (**P < 0.01). The mean is the average of three independent biological replicates. (D, E) Expression patterns of pYUC4::GUS in the differentiation zone of roots of 8-day-old plants harboring estrogen-inducible LHW–T5L1 treated with (E) or without (D) estrogen for 24 h. Scale bar = 100 µm. (F–I) Expression patterns of pYUC4::nYFP in 5-day-old wild-type (F) and lhw (G) root tips. Cross-sections of (F) and (G) are shown in (H) and (I), respectively. Asterisks indicate protoxylem cells. Scale bar = 100 µm. Next, we characterized the phenotype of a loss-of-function yuc4 mutant, which had a T-DNA insertion in the 5′-UTR region of YUC4 and the transcript level of YUC4 in yuc4 was greatly reduced compared with that of the wild type (Supplementary Fig. S1). In yuc4 plants, the root vasculature showed a diarch pattern, which consists of two protoxylem and protophloem poles, with normally differentiated protoxylem and metaxylem vessels (Fig. 2A, B, F, G), similar to that observed in wild-type Arabidopsis roots. However, yuc4 roots contained fewer vascular stem cells (procambial cells) than wild-type roots, although the number of xylem and phloem cells was similar to that in wild-type roots (Fig. 2J, K). To further examine the role of YUC4 in vascular development, we generated lhw yuc4 double mutant plants, because the lhw mutant exhibited major defects in vascular development and thus it was suitable for the analysis of vascular development as a sensitive background. The phenotype of about one-third of the lhw yuc4 roots was similar to that of lhw roots, such as a monarch pattern vasculature; however, in the remaining two-thirds of the lhw yuc4 roots, xylem vessel differentiation was partially arrested because of the delay of differentiation, which was a novel phenotype (Fig. 2C-E, H, I). This result implies that LHW is not the only regulator of YUC4 and other factors including LHW-family proteins may also regulate YUC4 expression. The arrest of xylem vessel differentiation in lhw yuc4 mutant roots was complemented by the expression of pYUC4::YUC4 (Fig. 2L). To determine whether YUC4 expression in RAM is required for proper vessel differentiation, we overexpressed YUC4 under the control of a RAM-specific promoter RGFR1 (Shinohara et al. 2016) in the lhw yuc4 double mutant background. The expression of pRGFR1::YUC4 complemented the arrested differentiation phenotype of xylem vessels (Fig. 2M). To further examine the importance of auxin biosynthesis in vascular development, we observed plants that are treated with kynurenine, an inhibitor of auxin synthesis. The width of a stele in RAM was significantly decreased in kynurenine-treated plants (Supplementary Fig. S2A–C). In addition, delay and partial arrest of xylem differentiation frequently occurred in kynurenine-treated lhw roots (Supplementary Fig. S2D, E). These results suggest that auxin synthesis is required for promoting vascular cell proliferation and proper xylem differentiation. Fig. 2 Open in new tabDownload slide Phenotypes observed in the vascular region of yuc4 roots. (A–E) Images of vascular tissue in 7-day-old wild-type (A), yuc4 (B), lhw (C) and lhw yuc4 (D, E) differentiation zone of roots. Scale bar = 50 µm. The fraction of samples showing similar patterns is presented. (F–I) Cross-sections of the vascular region in 7-day-old wild-type (F), yuc4 (G), lhw (H) and lhw yuc4 (I) differentiation zone of roots. Procambial cells, xylem precursor cells and phloem cells are indicated in yellow, green and blue, respectively. Cell types were judged by their morphological features. Scale bar = 20 µm. (J) Number of vascular cells in wild-type, yuc4, lhw and lhw yuc4 roots. Data represent mean ± SD; n = 14 (wild-type), 13 (yuc4), 12 (lhw) and 8 (lhw yuc4). The means, indicated by different letter superscripts in each column, were significantly different according to one-way ANOVA with the Tukey–Kramer post hoc test (P < 0.01). (K) Number of xylem, phloem and vascular stem cells in wild-type (black) and yuc4 (gray) roots. Data represent mean ± SD; n = 13. **P < 0.01 (Student’s t-test). (L, M) Images of vascular tissue in 7-day-old lhw yuc4 differentiation zone of roots harboring pYUC4::YUC4 (L) or pRGFR1::YUC4 (M). Scale bar = 50 µm. The fraction of samples showing similar patterns is presented. Fig. 2 Open in new tabDownload slide Phenotypes observed in the vascular region of yuc4 roots. (A–E) Images of vascular tissue in 7-day-old wild-type (A), yuc4 (B), lhw (C) and lhw yuc4 (D, E) differentiation zone of roots. Scale bar = 50 µm. The fraction of samples showing similar patterns is presented. (F–I) Cross-sections of the vascular region in 7-day-old wild-type (F), yuc4 (G), lhw (H) and lhw yuc4 (I) differentiation zone of roots. Procambial cells, xylem precursor cells and phloem cells are indicated in yellow, green and blue, respectively. Cell types were judged by their morphological features. Scale bar = 20 µm. (J) Number of vascular cells in wild-type, yuc4, lhw and lhw yuc4 roots. Data represent mean ± SD; n = 14 (wild-type), 13 (yuc4), 12 (lhw) and 8 (lhw yuc4). The means, indicated by different letter superscripts in each column, were significantly different according to one-way ANOVA with the Tukey–Kramer post hoc test (P < 0.01). (K) Number of xylem, phloem and vascular stem cells in wild-type (black) and yuc4 (gray) roots. Data represent mean ± SD; n = 13. **P < 0.01 (Student’s t-test). (L, M) Images of vascular tissue in 7-day-old lhw yuc4 differentiation zone of roots harboring pYUC4::YUC4 (L) or pRGFR1::YUC4 (M). Scale bar = 50 µm. The fraction of samples showing similar patterns is presented. Indole-3-acetic acid (IAA), a major auxin, is synthesized from tryptophan in two steps, and YUC4 is one of the enzymes, which catalyze the second step in this pathway (Mashiguchi et al. 2011). To determine whether the LHW–T5L1 dimer promotes auxin biosynthesis, we monitored changes in IAA concentration after the induction of LHW and T5L1-GFP. The amount of IAA started to increase at 72 h after the induction of LHW–T5L1, and was accumulated to high levels at 96 h, whereas the amount of IAA did not change when LHW–T5L1 overexpression was not induced (Fig. 3A). This result confirms that LHW–T5L1 can activate auxin biosynthesis. Fig. 3 Open in new tabDownload slide Auxin biosynthesis maintains the expression level of LHW–TMO5-related genes. (A) Concentration of IAA in cells harboring estrogen-inducible LHW and T5L1-GFP with estrogen (black) and without estrogen (gray). Data represent mean ± SD (n = 5). (B, C) Quantitative RT-PCR analysis of expression levels of LHW, TMO5 and T5L1 (B), and LOG3, LOG4 and ACL5 (C), in 7-day-old root tips treated with 100 µM kynurenine (Kyn) or water (control) for 0, 24 and 48 h. Data represent mean ± SD (*P < 0.05, **P < 0.01; Student’s t-test). The mean is the average of three independent biological replicates. (D, E) Quantitative RT-PCR analysis of expression levels of LHW, TMO5 and T5L1 (D) and LOG3, LOG4 and ACL5 (E) in 7-day-old wild-type and yuc4 root tips. Data represent mean ± SD (*P < 0.05, **P < 0.01; Student’s t-test). The mean is the average of three independent biological replicates. Fig. 3 Open in new tabDownload slide Auxin biosynthesis maintains the expression level of LHW–TMO5-related genes. (A) Concentration of IAA in cells harboring estrogen-inducible LHW and T5L1-GFP with estrogen (black) and without estrogen (gray). Data represent mean ± SD (n = 5). (B, C) Quantitative RT-PCR analysis of expression levels of LHW, TMO5 and T5L1 (B), and LOG3, LOG4 and ACL5 (C), in 7-day-old root tips treated with 100 µM kynurenine (Kyn) or water (control) for 0, 24 and 48 h. Data represent mean ± SD (*P < 0.05, **P < 0.01; Student’s t-test). The mean is the average of three independent biological replicates. (D, E) Quantitative RT-PCR analysis of expression levels of LHW, TMO5 and T5L1 (D) and LOG3, LOG4 and ACL5 (E) in 7-day-old wild-type and yuc4 root tips. Data represent mean ± SD (*P < 0.05, **P < 0.01; Student’s t-test). The mean is the average of three independent biological replicates. The expression of TMO5 and T5L1 is under the regulation of auxin because these genes are targets of MP/ARF5 (Schlereth et al. 2010). We hypothesized that auxin biosynthesis promoted by LHW–T5L1 is needed for maintaining the expression level of LHW and T5L1. To test this hypothesis, we first examined the expression level of LHW, TMO5 and T5L1 and their target genes such as LOG3, LOG4 and ACL5 in plants treated with kynurenine for 48 h. After starting the kynurenine treatment, the fluorescence of DR5, an auxin response marker (Heisler et al. 2005), was decreased at 3 h, very weak at 6 h and almost undetectable at 24 h (Fig. 4G–L). The kynurenine treatment caused significant reduction in the expression level of LHW, TMO5 and T5L1 and their target genes, LOG3, LOG4 and ACL5 (Fig. 3B, C). Especially, the T5L1 expression level was greatly reduced. We also monitored the temporal expression pattern of pT5L1-nYFP after the inhibition of auxin biosynthesis with kynurenine. The nYFP signal greatly decreased at 3 h after kynurenine treatment and was completely abolished by 24 h (Fig. 4A–F). These results strongly support our hypothesis of a positive feedback loop in which auxin biosynthesis promotes the expression of LHW, TMO5 and T5L1, which in turn promotes the expression of their target genes. Gene expression analysis indicated that LOG4 and ACL5 were significantly downregulated in yuc4 root tips compared with wild-type root tips (Fig. 3E), although no significant difference was detected in the expression levels of LHW, TMO5 and T5L1 between wild-type and yuc4 root tips (Fig. 3D). Although these data support our hypothesis at least partially, they also indicate that YUC4 may not be the only player of auxin biosynthesis in LHW–TMO5/T5L1-dependent vascular events. Previous studies showed abnormal vein patterns in leaves of quadruple mutants of yuc genes, including yuc4 (Cheng et al. 2006, Cheng et al. 2007), suggesting that other YUC genes are also involved in initial vascular development. Fig. 4 Open in new tabDownload slide The inhibition of auxin biosynthesis changes gene expression patterns in vascular cells. (A–R) Expression patterns of T5L1 (A–F), DR5 (G–L) and TCSn (M–R) in the RAM of 5-day-old plants treated with or without 100 µM kynurenine (Kyn) for 3, 6 and 24 h. Roots were stained with propidium iodide (purple). Scale bar = 100 µm. Fig. 4 Open in new tabDownload slide The inhibition of auxin biosynthesis changes gene expression patterns in vascular cells. (A–R) Expression patterns of T5L1 (A–F), DR5 (G–L) and TCSn (M–R) in the RAM of 5-day-old plants treated with or without 100 µM kynurenine (Kyn) for 3, 6 and 24 h. Roots were stained with propidium iodide (purple). Scale bar = 100 µm. In both auxin-related experiments, auxin reduction decreased LOG gene expression. Therefore, we examined the expression of a cytokinin response marker, TCSn (Zurcher et al. 2013), as an output of LHW–T5L1 activity. The TCSn signal decreased gradually from 6 to 24 h after the start of the kynurenine treatment, as expected (Fig. 4M–R). This result also suggests that the inhibition of auxin biosynthesis causes the decrease of the LHW–TMO5/T5L1 activity because we previously showed that the TCSn signal is absent at the vascular region in lhw and log3 log4 mutant root tips (Ohashi-Ito et al. 2014). Discussion In this study, we showed that LHW–T5L1, which is a key regulator of vascular development in RAM, promotes auxin biosynthesis. It is shown that auxin biosynthesis in RAM is essential for metaxylem differentiation (Ursache et al. 2014). Our results also showed that auxin biosynthesis in RAM promoted xylem differentiation. Thus, the control of xylem differentiation driven by auxin biosynthesis may occur quite early in vascular development. It is reported that LHW–TMO5/T5L1 positively regulate the biosynthesis of two phytohormones or phytohormone-like molecules, in addition to auxin. One of these molecules is cytokinin, which spreads into neighboring cells and induces the division of vascular cells, whereas the other is thermospermine, which functions in the negative feedback loop affecting LHW–TMO5/T5L1 (De Rybel et al. 2014, Ohashi-Ito et al. 2014, Katayama et al. 2015, Vera-Sirera et al. 2015). Thus, LHW–TMO5/T5L1 regulate the synthesis of at least three phytohormones or phytohormone-like molecules. Among these, auxin and thermospermine act in a cell-autonomous manner, whereas cytokinin acts in a noncell-autonomous manner. A combination of these two types of signal molecules may be important for generating a new tissue to spatiotemporally coordinate cell division and differentiation. Our results suggest that LHW–T5L1 and auxin biosynthesis are tightly related in the initial process of vascular development, and these two factors mutually regulate each other. The LHW–T5L1 dimer upregulates YUC4 and activates auxin biosynthesis. Conversely, auxin promotes the expression of LHW, TMO5 and T5L1. Therefore, it is likely that LHW–T5L1/TMO5 and YUC4-dependent auxin biosynthesis form a positive feedback loop. Although it is well known that auxin flow is regulated through a positive feedback loop (Linh et al. 2018), the concept that auxin biosynthesis is a component of a positive feedback loop is novel in vascular development. The LHW–T5L1/TMO5 dimers also form a negative feedback loop through the upregulation of ACL5 and SACL family genes, which encode a thermospermine synthase and bHLH proteins, respectively (Katayama et al. 2015, Vera-Sirera et al. 2015). The synthesized thermospermine promotes the production of SACL proteins, which interact with LHW and block the formation of the active LHW–T5L1/TMO5 heterodimers. This negative feedback regulation suppresses the function of LHW–T5L1/TMO5 at the protein level, whereas the positive feedback regulation, comprising auxin biosynthesis, promotes the function of LHW–T5L1/TMO5 at the transcript level. Although mechanisms underlying the positive and negative feedback regulations of LHW–T5L1/TMO5 are different, both these regulations likely control the output of LHW–T5L1/TMO5 as a whole. In other words, the balance between positive and negative feedback regulations may be responsible for fine-tuning the output of LHW–T5L1/TMO5 (Supplementary Fig. S3). We note, however, that other YUC4-dependent transcription factor than LHW–TMO5/T5L1 may be also involved in the initial process of vascular development, because the number of vascular cells decreased in yuc4, despite the fact that expression levels of LHW, TMO5 and T5L1 were not significantly changed between wild type and yuc4. In this case, this unknown factor might be reduced in yuc4 mutants, resulting in a decrease in the expression level of LOG4, which promotes cell division by producing cytokinin. ATHB8 is a possible candidate of this factor because ATHB8 is auxin-inducible (Ursache et al. 2014) and regulates ACL5 (Baima et al. 2014), which expression was lower in yuc4. Previous studies showed that local auxin biosynthesis plays crucial roles in many aspects of plant development such as the formation of floral organs, the apical–basal axis formation in embryos and the maintenance of root stem cell niche (Cheng et al. 2006, Zhao 2008, Robert et al. 2013, Brumos et al. 2018). In some of these processes, local auxin biosynthesis contributes to generating auxin maximum. Our previous study showed that lateral root primordia in lhw mutants failed to confine auxin maximum in vascular cells and the quiescent center (Ohashi-Ito et al. 2013). Therefore, LHW–TMO5/T5L1-mediated local auxin biosynthesis and positive feedback regulations may contribute to the confined auxin maximum formation during vascular development. In summary, we propose a model in which auxin biosynthesis is activated in the initial process of vascular development, and forms a positive feedback loop to control the level of LHW–T5L1/TMO5. The level of LHW–T5L1/TMO5 is also regulated by a negative feedback loop consisting of LHW–T5L1/TMO5, ACL5 and SACLs. Because initial vascular development, including the proliferation of vascular stem cells and determination of xylem cell identity, is dominated by LHW–T5L1/TMO5, it appears that adjusting the output of LHW–T5L1/TMO5 appropriately through the double feedback regulations is critical for the coordinated development of vascular tissues. Materials and Methods Plant materials and growth conditions Arabidopsis thaliana accession Columbia, yuc4 (SALK_047083C), lhw (SALK_079402), DR5 rev::3x VENUS-N7 (Heisler et al. 2005) and TCSn::GFP (Zurcher et al. 2013) were used. Seeds were sown on half-strength Murashige and Skoog (1/2 MS) agar plates containing 1% sucrose, 0.8% agar and appropriate antibiotics (MS plates), incubated at 4°C for 2–3 d, and then moved to an incubator for growth under continuous light at 22°C. For kynurenine treatment, plants were grown on MS plates for 5 d and then transferred onto MS plates including 100 µM kynurenine. Arabidopsis culture cells harboring estrogen-inducible LHW and T5L1-GFP established in the previous study were used (Ohashi-Ito et al. 2014). Estrogen-inducible YFP line was established according to previously described methods (Ohashi-Ito et al. 2010). To induce LHW and T5L1-GFP, or YFP expression, 1 ml of MS medium containing 0.5 µM estradiol was supplemented to a 0.5 ml aliquot of 10-day-old transformant cell culture and then cultured on a rotary shaker at 124 rpm at 22°C in the dark. DNA manipulation The promoter sequences of YUC4 (2.2 kb), RGFR1 (2.5 kb) and T5L1 (3.0 kb) were amplified by PCR and cloned into pENTR/D-TOPO cloning vector (Thermo Fisher Scientific, Waltham, MA, USA) to generate entry vectors. The promoter sequences were integrated into pBGYN vector (Kubo et al. 2005) or pGWB434 vector, which included intron-GUS for promoter analysis, and was gift from T. Nakagawa, using LR Clonase II Enzyme Mix (Life Technologies). The coding sequence of YUC4 was amplified by PCR with YUC4cds-AscI-F and YUC4cds-AscI-R primers and digested with AscI restriction enzyme (New England Biolabs, Beverly, MA, USA). This DNA fragment was ligated with AscI-digested entry vectors to generate pYUC4::YUC4 or pRGFR1::YUC4, which were transferred into pGWB1 vector (Nakagawa et al. 2007) using LR Clonase II Enzyme Mix. Primers used in this study are listed in Supplementary Table S1. RNA isolation and quantitative RT-PCR Total RNA isolation and cDNA synthesis were performed according to previously described methods (Ohashi-Ito et al. 2010). Quantitative RT-PCR was performed using LightCycler480 Probes Master (Roche Diagnostics, Mannheim, Germany) with sets of Universal ProbeLibrary probes (Roche) and primers listed in Supplementary Table S1 on a LightCycler480 instrument II (Roche) according to the manufacturer’s protocol. Standard curves for each primer set were created by the second derivative maximum method using 10-fold serial dilutions of cDNA obtained from one of the samples. Expression levels of each gene calculated from the standard curves were converted to expression ratio relative to those of the UBQ10 reference gene. Chromatin immunoprecipitation-PCR Cell fixation was performed according to the method described by Morohashi and Grotewold (2009) and Morohashi et al. (2012), and chromatin shearing was performed according to the method described by Ohashi-Ito et al. (2014). Cultured cells were fixed for 20 min in fix buffer [0.4 M sucrose, 10 mM Tris-HCl, pH 8.0, 1 mM EDTA, 1% protease inhibitor cocktail for plant cell and extracts (Sigma–Aldrich, St. Louis, MO, USA), and 1% formaldehyde]. After addition of 0.1 M glycine and incubation for 5 min with gentle shaking, cells were collected by centrifugation, washed twice with ice-cold water, frozen in liquid nitrogen and stored at −80°C. Frozen cells collected from 2-ml cell cultures were ground using Shake Master neo (Biomedical Science, Tokyo, Japan) at 1,500 rpm for 2 min, and suspended in 1 ml of lysis buffer [50 mM HEPES, pH 7.5, 150 mM NaCl, 1 mM EDTA, 1% Triton X-100, 0.1% deoxycholate, 0.1% SDS, 1 mM phenylmethylsulfonyl fluoride, 10 mM sodium butyrate, 1% protease inhibitor cocktail for plant cell and extracts (Sigma-Aldrich)]. Subsequently, the suspension was sonicated for 10 min with Covaris S220 and centrifuged at 15,000 × g for 10 min at 4°C. Immunoprecipitation was performed with 240 µl aliquots of the supernatants and 8 µl of anti-GFP antibody (ab290, Abcam, Cambridge, UK) using OneDay ChIP kit (Diagenode, Liège, Belgium) according to the manufacturer’s protocol. Input samples were prepared from 24 µl aliquots of the supernatants. Real-time quantitative PCR was run using LightCycler480 Probes Master (Roche) with sets of Universal ProbeLibrary probes (Roche) and primers listed in Supplementary Table S1 on a LightCycler480 instrument II (Roche) according to the manufacturer’s protocol. Quantification of endogenous IAA levels The culture cells harboring the estrogen-inducible LHW and T5L1-GFP transgene were used for the quantification of endogenous IAA levels. Induced-cells were treated with 5 µM estrogen for 24 h, washed with culture medium three times, and then continued to culture. Quantification was performed according to Kojima et al. (2009). Histological analysis and fluorescence imaging GUS staining of 8-day-old seedlings was performed as described previously (Ohashi-Ito et al. 2010). DIC images were taken using a BM5500 microscope (Leica Microsystems, Nussloch, Germany). Technovit sections were prepared using 7-day-old roots according to previously described methods (Ohashi-Ito et al. 2013) and stained with Toluidine Blue O. Sections were prepared from roots at 3 mm above the root tips. Fluorescence images were taken using a FV1200 confocal microscope (Olympus, Tokyo, Japan). For staining the plasma membrane, roots were incubated in 0.01 mg/ml propidium iodide. Funding The Ministry of Education, Culture, Sports, Science and Technology of Japan [25113004 to K.O.-I. and 15H05958 to H.F.] and the Japan Society for the Promotion of Science [16H06377 to H.F.]. Acknowledgments We thank Minobu Shimizu, Yukiko Sugisawa and Sumika Tsuji-Tsukinoki for technical support. We also thank Nam-Hai Chua, Bruno Müller and ABRC for materials. Disclosures The authors have no conflicts of interest to declare. References Baima S. , Forte V. , Possenti M. , Penalosa A. , Leoni G. , Salvi S. , et al. 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Cuesta-Seijo, Jose Antonio; De Porcellinis, Alice Jara; Valente, Angela Hørdum; Striebeck, Alexander; Voss, Cynthia; Marri, Lucia; Hansson, Andreas; Jansson, Anita M; Dinesen, Malene Hessellund; Fangel, Jonatan Ulrik; Harholt, Jesper; Popovic, Milan; Thieme, Mercedes; Hochmuth, Anton; Zeeman, Samuel C; Mikkelsen, Teis Nørgaard; Jørgensen, Rikke Bagger;