Rhamnolipids Elicit Defense Responses and Induce Disease Resistance against Biotrophic, Hemibiotrophic, and Necrotrophic Pathogens That Require Different Signaling Pathways in Arabidopsis and Highlight a Central Role for Salicylic Acid

Plant resistance to phytopathogenic microorganisms mainly relies on the activation of an innate immune response usually launched after recognition by the plant cells of microbe-associated molecular patterns. The plant hormones, salicylic acid (SA), jasmonic acid, and ethylene have emerged as key players in the signaling networks involved in plant immunity. Rhamnolipids (RLs) are glycolipids produced by bacteria and are involved in surface motility and biofilm development. Here we report that RLs trigger an immune response in Arabidopsis (Arabidopsis thaliana) characterized by signaling molecules accumulation and defense gene activation. This immune response participates to resistance against the hemibiotrophic bacterium Pseudomonas syringae pv tomato, the biotrophic oomycete Hyaloperonospora arabidopsidis, and the necrotrophic fungus Botrytis cinerea. We show that RL-mediated resistance involves different signaling pathways that depend on the type of pathogen. Ethylene is involved in RL-induced resistance to H. arabidopsidis and to P. syringae pv tomato whereas jasmonic acid is essential for the resistance to B. cinerea. SA participates to the restriction of all pathogens. We also show evidence that SA-dependent plant defenses are potentiated by RLs following challenge by B. cinerea or P. syringae pv tomato. These results highlight a central role for SA in RL-mediated resistance. In addition to the activation of plant defense responses, antimicrobial properties of RLs are thought to participate in the protection against the fungus and the oomycete. Our data highlight the intricate mechanisms involved in plant protection triggered by a new type of molecule that can be perceived by plant cells and that can also act directly onto pathogens.In their environment, plants are challenged by potentially pathogenic microorganisms. In response, they express a set of defense mechanisms including preformed structural and chemical barriers, as well as an innate immune response quickly activated after microorganism perception (Boller and Felix, 2009). Plant innate immunity is triggered after recognition by pattern recognition receptors of conserved pathogen- or microbe-associated molecular patterns (PAMPs or MAMPs, respectively) or by plant endogenous molecules released by pathogen invasion and called danger-associated molecular patterns (Boller and Felix, 2009; Dodds and Rathjen, 2010). This first step of recognition leads to the activation of MAMP-triggered immunity (MTI). Successful pathogens can secrete effectors that interfere or suppress MTI, resulting in effector-triggered susceptibility. A second level of perception involves the direct or indirect recognition by specific receptors of pathogen effectors leading to effector-triggered immunity (ETI; Boller and Felix, 2009; Dodds and Rathjen, 2010). Whereas MTI and ETI are thought to involve common signaling network, ETI is usually quantitatively stronger than MTI and associated with more sustained and robust immune responses (Katagiri and Tsuda, 2010; Tsuda and Katagiri, 2010).The plant hormones, salicylic acid (SA), jasmonic acid (JA), and ethylene (ET) have emerged as key players in the signaling networks involved in MTI and ETI (Robert-Seilaniantz et al., 2007; Tsuda et al., 2009; Katagiri and Tsuda, 2010; Mersmann et al., 2010; Tsuda and Katagiri, 2010; Robert-Seilaniantz et al., 2011). Interactions between these signal molecules allow the plant to activate and/or modulate an appropriate spectrum of responses, depending on the pathogen lifestyle, necrotroph or biotroph (Glazebrook, 2005; Koornneef and Pieterse, 2008). It is assumed that JA and ET signaling pathways are important for resistance to necrotrophic fungi including Botrytis cinerea and Alternaria brassicicola (Thomma et al., 2001; Ferrari et al., 2003; Glazebrook, 2005). Infection of Arabidopsis (Arabidopsis thaliana) with B. cinerea causes the induction of the JA/ET responsive gene PLANT DEFENSIN1.2 (PDF1.2; Penninckx et al., 1996; Zimmerli et al., 2001). Induction of PDF1.2 by B. cinerea is blocked in ethylene-insensitive2 (ein2) and coronatine-insensitive1 (coi1) mutants that are respectively defective in ET and JA signal transduction pathways. Moreover, ein2 and coi1 plants are highly susceptible to B. cinerea infection (Thomma et al., 1998; Thomma et al., 1999). JA/ET-dependent responses do not seem to be usually induced during resistance to biotrophs, but they can be effective if they are stimulated prior to pathogen challenge (Glazebrook, 2005). Plants impaired in SA signaling are highly susceptible to biotrophic and hemibiotrophic pathogens. Following pathogen infection, SA hydroxylase (NahG), enhanced disease susceptibility5 (eds5), or SA induction-deficient2 (sid2) plants are unable to accumulate high SA levels and they display heightened susceptibility to Pseudomonas syringae pv tomato (Pst), Hyaloperonospora arabidopsidis, or Erysiphe orontii (Delaney et al., 1994; Lawton et al., 1995; Wildermuth et al., 2001; Nawrath et al., 2002; Vlot et al., 2009). Mutants that are insensitive to SA, such as nonexpressor of PATHOGENESIS-RELATED (PR) genes1 (npr1), have enhanced susceptibility to these pathogens (Cao et al., 1994; Glazebrook et al., 1996; Shah et al., 1997; Dong, 2004). According to some reports, plant defense against necrotrophs also involves SA. Arabidopsis plants expressing the nahG gene and infected with B. cinerea show larger lesions compared with wild-type plants (Govrin and Levine, 2002). In tobacco (Nicotiana tabacum), acidic isoforms of PR3 and PR5 gene that are specifically induced by SA (Ménard et al., 2004) are up-regulated after challenge by B. cinerea (El Oirdi et al., 2010). Resistance to some necrotrophs like Fusarium graminearum involves both SA and JA signaling pathways (Makandar et al., 2010). It is assumed that SA and JA signaling can be antagonistic (Bostock, 2005; Koornneef and Pieterse, 2008; Pieterse et al., 2009; Thaler et al., 2012). In Arabidopsis, SA inhibits JA-dependent resistance against A. brassicicola or B. cinerea (Spoel et al., 2007; Koornneef et al., 2008). Recent studies demonstrated that ET modulates the NPR1-mediated antagonism between SA and JA (Leon-Reyes et al., 2009; Leon-Reyes et al., 2010a) and suppression by SA of JA-responsive gene expression is targeted at a position downstream of the JA biosynthesis pathway (Leon-Reyes et al., 2010b). Synergistic effects of SA- and JA-dependent signaling are also well documented (Schenk et al., 2000; van Wees et al., 2000; Mur et al., 2006) and induction of some defense responses after pathogen challenge requires intact JA, ET, and SA signaling pathways (Campbell et al., 2003).Isolated MAMPs trigger defense responses that also require the activation of SA, JA, and ET signaling pathways (Tsuda et al., 2009; Katagiri and Tsuda, 2010). For instance, treatment with the flagellin peptide flg22 induces many SA-related genes including SID2, EDS5, NPR1, and PR1 (Ferrari et al., 2007; Denoux et al., 2008), causes SA accumulation (Tsuda et al., 2008; Wang et al., 2009), and activates ET signaling (Bethke et al., 2009; Mersmann et al., 2010). Local application of lipopolysaccharides elevates the level of SA (Mishina and Zeier, 2007). The oomycete Pep13 peptide induces defense responses in potato (Solanum tuberosum) that require both SA and JA (Halim et al., 2009). Although signaling networks induced by isolated MAMPs are well documented, the contribution of SA, JA, and ET in MAMP- or PAMP-induced resistance to biotrophs and necrotrophs is poorly understood.Rhamnolipids (RLs) are glycolipids produced by various bacteria species including some Pseudomonas and Burkholderia species. They are essential for bacterial surface motility and biofilm development (Vatsa et al., 2010; Chrzanowski et al., 2012). RLs are potent stimulators of animal immunity (Vatsa et al., 2010). They have recently been shown to elicit plant defense responses and to induce resistance against B. cinerea in grapevine (Vitis vinifera; Varnier et al., 2009). They also participate to biocontrol activity of the plant beneficial bacteria Pseudomonas aeruginosa PNA1 against oomycetes (Perneel et al., 2008). However, the signaling pathways used by RLs to stimulate plant innate immunity are not known. To gain more insights into RL-induced MTI, we investigated RL-triggered defense responses and resistance to the necrotrophic fungus B. cinerea, the biotroph oomycete H. arabidopsidis, and the hemibiotroph bacterium Pst in Arabidopsis. Our results show that RLs trigger an innate immune response in Arabidopsis that protects the plant against these different lifestyle pathogens. We demonstrate that RL-mediated resistance involves separated signaling sectors that depend on the type of pathogen. In plants challenged by RLs, SA has a central role and participates to the restriction of the three pathogens. ET is fully involved in RL-induced resistance to the biotrophic oomycete and to the hemibiotrophic bacterium whereas JA is essential for the resistance to the necrotrophic fungus.     

Multiscale Metabolic Modeling: Dynamic Flux Balance Analysis on a Whole-Plant Scale

Plant metabolism is characterized by a unique complexity on the cellular, tissue, and organ levels. On a whole-plant scale, changing source and sink relations accompanying plant development add another level of complexity to metabolism. With the aim of achieving a spatiotemporal resolution of source-sink interactions in crop plant metabolism, a multiscale metabolic modeling (MMM) approach was applied that integrates static organ-specific models with a whole-plant dynamic model. Allowing for a dynamic flux balance analysis on a whole-plant scale, the MMM approach was used to decipher the metabolic behavior of source and sink organs during the generative phase of the barley (Hordeum vulgare) plant. It reveals a sink-to-source shift of the barley stem caused by the senescence-related decrease in leaf source capacity, which is not sufficient to meet the nutrient requirements of sink organs such as the growing seed. The MMM platform represents a novel approach for the in silico analysis of metabolism on a whole-plant level, allowing for a systemic, spatiotemporally resolved understanding of metabolic processes involved in carbon partitioning, thus providing a novel tool for studying yield stability and crop improvement.Plants are of vital significance as a source of food (Grusak and DellaPenna, 1999; Rogalski and Carrer, 2011), feed (Lu et al., 2011), energy (Tilman et al., 2006; Parmar et al., 2011), and feedstocks for the chemical industry (Metzger and Bornscheuer, 2006; Kinghorn et al., 2011). Given the close connection between plant metabolism and the usability of plant products, there is a growing interest in understanding and predicting the behavior and regulation of plant metabolic processes. In order to increase crop quality and yield, there is a need for methods guiding the rational redesign of the plant metabolic network (Schwender, 2009).Mathematical modeling of plant metabolism offers new approaches to understand, predict, and modify complex plant metabolic processes. In plant research, the issue of metabolic modeling is constantly gaining attention, and different modeling approaches applied to plant metabolism exist, ranging from highly detailed quantitative to less complex qualitative approaches (for review, see Giersch, 2000; Morgan and Rhodes, 2002; Poolman et al., 2004; Rios-Estepa and Lange, 2007).A widely used modeling approach is flux balance analysis (FBA), which allows the prediction of metabolic capabilities and steady-state fluxes under different environmental and genetic backgrounds using (non)linear optimization (Orth et al., 2010). Assuming steady-state conditions, FBA has the advantage of not requiring the knowledge of kinetic parameters and, therefore, can be applied to model detailed, large-scale systems. In recent years, the FBA approach has been applied to several different plant species, such as maize (Zea mays; Dal’Molin et al., 2010; Saha et al., 2011), barley (Hordeum vulgare; Grafahrend-Belau et al., 2009b; Melkus et al., 2011; Rolletschek et al., 2011), rice (Oryza sativa; Lakshmanan et al., 2013), Arabidopsis (Arabidopsis thaliana; Poolman et al., 2009; de Oliveira Dal’Molin et al., 2010; Radrich et al., 2010; Williams et al., 2010; Mintz-Oron et al., 2012; Cheung et al., 2013), and rapeseed (Brassica napus; Hay and Schwender, 2011a, 2011b; Pilalis et al., 2011), as well as algae (Boyle and Morgan, 2009; Cogne et al., 2011; Dal’Molin et al., 2011) and photoautotrophic bacteria (Knoop et al., 2010; Montagud et al., 2010; Boyle and Morgan, 2011). These models have been used to study different aspects of metabolism, including the prediction of optimal metabolic yields and energy efficiencies (Dal’Molin et al., 2010; Boyle and Morgan, 2011), changes in flux under different environmental and genetic backgrounds (Grafahrend-Belau et al., 2009b; Dal’Molin et al., 2010; Melkus et al., 2011), and nonintuitive metabolic pathways that merit subsequent experimental investigations (Poolman et al., 2009; Knoop et al., 2010; Rolletschek et al., 2011). Although FBA of plant metabolic models was shown to be capable of reproducing experimentally determined flux distributions (Williams et al., 2010; Hay and Schwender, 2011b) and generating new insights into metabolic behavior, capacities, and efficiencies (Sweetlove and Ratcliffe, 2011), challenges remain to advance the utility and predictive power of the models.Given that many plant metabolic functions are based on interactions between different subcellular compartments, cell types, tissues, and organs, the reconstruction of organ-specific models and the integration of these models into interacting multiorgan and/or whole-plant models is a prerequisite to get insight into complex plant metabolic processes organized on a whole-plant scale (e.g. source-sink interactions). Almost all FBA models of plant metabolism are restricted to one cell type (Boyle and Morgan, 2009; Knoop et al., 2010; Montagud et al., 2010; Cogne et al., 2011; Dal’Molin et al., 2011), one tissue or one organ (Grafahrend-Belau et al., 2009b; Hay and Schwender, 2011a, 2011b; Pilalis et al., 2011; Mintz-Oron et al., 2012), and only one model exists taking into account the interaction between two cell types by specifying the interaction between mesophyll and bundle sheath cells in C4 photosynthesis (Dal’Molin et al., 2010). So far, no model representing metabolism at the whole-plant scale exists.Considering whole-plant metabolism raises the problem of taking into account temporal and environmental changes in metabolism during plant development and growth. Although classical static FBA is unable to predict the dynamics of metabolic processes, as the network analysis is based on steady-state solutions, time-dependent processes can be taken into account by extending the classical static FBA to a dynamic flux balance analysis (dFBA), as proposed by Mahadevan et al. (2002). The static (SOA) and dynamic optimization approaches introduced in this work provide a framework for analyzing the transience of metabolism by integrating kinetic expressions to dynamically constrain exchange fluxes. Due to the requirement of knowing or estimating a large number of kinetic parameters, so far dFBA has only been applied to a plant metabolic model once, to study the photosynthetic metabolism in the chloroplasts of C3 plants by a simplified model of five biochemical reactions (Luo et al., 2009). Integrating a dynamic model into a static FBA model is an alternative approach to perform dFBA.In this study, a multiscale metabolic modeling (MMM) approach was applied with the aim of achieving a spatiotemporal resolution of cereal crop plant metabolism. To provide a framework for the in silico analysis of the metabolic dynamics of barley on a whole-plant scale, the MMM approach integrates a static multiorgan FBA model and a dynamic whole-plant multiscale functional plant model (FPM) to perform dFBA. The performance of the novel whole-plant MMM approach was tested by studying source-sink interactions during the seed developmental phase of barley plants.     

Evolution of Conifer Diterpene Synthases: Diterpene Resin Acid Biosynthesis in Lodgepole Pine and Jack Pine Involves Monofunctional and Bifunctional Diterpene Synthases

Abscisic acid (ABA) induces stomatal closure and inhibits light-induced stomatal opening. The mechanisms in these two processes are not necessarily the same. It has been postulated that the ABA receptors involved in opening inhibition are different from those involved in closure induction. Here, we provide evidence that four recently identified ABA receptors (PYRABACTIN RESISTANCE1 [PYR1], PYRABACTIN RESISTANCE-LIKE1 [PYL1], PYL2, and PYL4) are not sufficient for opening inhibition in Arabidopsis (Arabidopsis thaliana). ABA-induced stomatal closure was impaired in the pyr1/pyl1/pyl2/pyl4 quadruple ABA receptor mutant. ABA inhibition of the opening of the mutant’s stomata remained intact. ABA did not induce either the production of reactive oxygen species and nitric oxide or the alkalization of the cytosol in the quadruple mutant, in accordance with the closure phenotype. Whole cell patch-clamp analysis of inward-rectifying K⁺ current in guard cells showed a partial inhibition by ABA, indicating that the ABA sensitivity of the mutant was not fully impaired. ABA substantially inhibited blue light-induced phosphorylation of H⁺-ATPase in guard cells in both the mutant and the wild type. On the other hand, in a knockout mutant of the SNF1-related protein kinase, srk2e, stomatal opening and closure, reactive oxygen species and nitric oxide production, cytosolic alkalization, inward-rectifying K⁺ current inactivation, and H⁺-ATPase phosphorylation were not sensitive to ABA.The phytohormone abscisic acid (ABA), which is synthesized in response to abiotic stresses, plays a key role in the drought hardiness of plants. Reducing transpirational water loss through stomatal pores is a major ABA response (Schroeder et al., 2001). ABA promotes the closure of open stomata and inhibits the opening of closed stomata. These effects are not simply the reverse of one another (Allen et al., 1999; Wang et al., 2001; Mishra et al., 2006).A class of receptors of ABA was identified (Ma et al., 2009; Park et al., 2009; Santiago et al., 2009; Nishimura et al., 2010). The sensitivity of stomata to ABA was strongly decreased in quadruple and sextuple mutants of the ABA receptor genes PYRABACTIN RESISTANCE/PYRABACTIN RESISTANCE-LIKE/REGULATORY COMPONENT OF ABSCISIC ACID RECEPTOR (PYR/PYL/RCAR; Nishimura et al., 2010; Gonzalez-Guzman et al., 2012). The PYR/PYL/RCAR receptors are involved in the early ABA signaling events, in which a sequence of interactions of the receptors with PROTEIN PHOSPHATASE 2Cs (PP2Cs) and subfamily 2 SNF1-RELATED PROTEIN KINASES (SnRK2s) leads to the activation of downstream ABA signaling targets in guard cells (Cutler et al., 2010; Kim et al., 2010; Weiner et al., 2010). Studies of Commelina communis and Vicia faba suggested that the ABA receptors involved in stomatal opening are not the same as the ABA receptors involved in stomatal closure (Allan et al., 1994; Anderson et al., 1994; Assmann, 1994; Schwartz et al., 1994). The roles of PYR/PYL/RCAR in either stomatal opening or closure remained to be elucidated.Blue light induces stomatal opening through the activation of plasma membrane H⁺-ATPase in guard cells that generates an inside-negative electrochemical gradient across the plasma membrane and drives K⁺ uptake through voltage-dependent inward-rectifying K⁺ channels (Assmann et al., 1985; Shimazaki et al., 1986; Blatt, 1987; Schroeder et al., 1987; Thiel et al., 1992). Phosphorylation of the penultimate Thr of the plasma membrane H⁺-ATPase is a prerequisite for blue light-induced activation of the H⁺-ATPase (Kinoshita and Shimazaki, 1999, 2002). ABA inhibits H⁺-ATPase activity through dephosphorylation of the penultimate Thr in the C terminus of the H⁺-ATPase in guard cells, resulting in prevention of the opening (Goh et al., 1996; Zhang et al., 2004; Hayashi et al., 2011). Inward-rectifying K⁺ currents (I_Kin) of guard cells are negatively regulated by ABA in addition to through the decline of the H⁺ pump-driven membrane potential difference (Schroeder and Hagiwara, 1989; Blatt, 1990; McAinsh et al., 1990; Schwartz et al., 1994; Grabov and Blatt, 1999; Saito et al., 2008). This down-regulation of ion transporters by ABA is essential for the inhibition of stomatal opening.A series of second messengers has been shown to mediate ABA-induced stomatal closure. Reactive oxygen species (ROS) produced by NADPH oxidases play a crucial role in ABA signaling in guard cells (Pei et al., 2000; Zhang et al., 2001; Kwak et al., 2003; Sirichandra et al., 2009; Jannat et al., 2011). Nitric oxide (NO) is an essential signaling component in ABA-induced stomatal closure (Desikan et al., 2002; Guo et al., 2003; Garcia-Mata and Lamattina, 2007; Neill et al., 2008). Alkalization of cytosolic pH in guard cells is postulated to mediate ABA-induced stomatal closure in Arabidopsis (Arabidopsis thaliana) and Pisum sativum and Paphiopedilum species (Irving et al., 1992; Gehring et al., 1997; Grabov and Blatt, 1997; Suhita et al., 2004; Gonugunta et al., 2008). These second messengers transduce environmental signals to ion channels and ion transporters that create the driving force for stomatal movements (Ward et al., 1995; MacRobbie, 1998; Garcia-Mata et al., 2003).In this study, we examined the mobilization of second messengers, the inactivation of I_Kin, and the suppression of H⁺-ATPase phosphorylation evoked by ABA in Arabidopsis mutants to clarify the downstream signaling events of ABA signaling in guard cells. The mutants included a quadruple mutant of PYR/PYL/RCARs, pyr1/pyl1/pyl2/pyl4, and a mutant of a SnRK2 kinase, srk2e.     

Heterodimeric Capping Protein from Arabidopsis Is a Membrane-Associated,Actin-Binding Protein

The actin cytoskeleton is a major regulator of cell morphogenesis and responses to biotic and abiotic stimuli. The organization and activities of the cytoskeleton are choreographed by hundreds of accessory proteins. Many actin-binding proteins are thought to be stimulus-response regulators that bind to signaling phospholipids and change their activity upon lipid binding. Whether these proteins associate with and/or are regulated by signaling lipids in plant cells remains poorly understood. Heterodimeric capping protein (CP) is a conserved and ubiquitous regulator of actin dynamics. It binds to the barbed end of filaments with high affinity and modulates filament assembly and disassembly reactions in vitro. Direct interaction of CP with phospholipids, including phosphatidic acid, results in uncapping of filament ends in vitro. Live-cell imaging and reverse-genetic analyses of cp mutants in Arabidopsis (Arabidopsis thaliana) recently provided compelling support for a model in which CP activity is negatively regulated by phosphatidic acid in vivo. Here, we used complementary biochemical, subcellular fractionation, and immunofluorescence microscopy approaches to elucidate CP-membrane association. We found that CP is moderately abundant in Arabidopsis tissues and present in a microsomal membrane fraction. Sucrose density gradient separation and immunoblotting with known compartment markers were used to demonstrate that CP is enriched on membrane-bound organelles such as the endoplasmic reticulum and Golgi. This association could facilitate cross talk between the actin cytoskeleton and a wide spectrum of essential cellular functions such as organelle motility and signal transduction.The cellular levels of membrane-associated lipids undergo dynamic changes in response to developmental and environmental stimuli. Different species of phospholipids target specific proteins and this often affects the activity and/or subcellular localization of these lipid-binding proteins. One such membrane lipid, phosphatidic acid (PA), serves as a second messenger and regulates multiple developmental processes in plants, including seedling development, root hair growth and pattern formation, pollen tube growth, leaf senescence, and fruit ripening. PA levels also change during various stress responses, including high salinity and dehydration, pathogen attack, and cold tolerance (Testerink and Munnik, 2005, 2011; Wang, 2005; Li et al., 2009). In mammalian cells, PA is critical for vesicle trafficking events, such as vesicle budding from the Golgi apparatus, vesicle transport, exocytosis, endocytosis, and vesicle fusion (Liscovitch et al., 2000; Freyberg et al., 2003; Jenkins and Frohman, 2005).The actin cytoskeleton and a plethora of actin-binding proteins (ABPs) are well-known targets and transducers of lipid signaling (Drøbak et al., 2004; Saarikangas et al., 2010; Pleskot et al., 2013). For example, several ABPs have the ability to bind phosphoinositide lipids, such as phosphatidylinositol 4,5-bisphosphate [PtdIns(4,5)P₂]. The severing or actin filament depolymerizing proteins such as villin, cofilin, and profilin are inhibited when bound to PtdIns(4,5)P₂. One ABP appears to be strongly regulated by another phospholipid; human gelsolin binds to lysophosphatidic acid and its filament severing and barbed-end capping activities are inhibited by this biologically active lipid (Meerschaert et al., 1998). Gelsolin is not, however, regulated by PA (Meerschaert et al., 1998), nor are profilin (Lassing and Lindberg, 1985), α-actinin (Fraley et al., 2003), or chicken CapZ (Schafer et al., 1996).The heterodimeric capping protein (CP) from Arabidopsis (Arabidopsis thaliana) also binds to and its activity is inhibited by phospholipids, including both PtdIns(4,5)P₂ and PA (Huang et al., 2003, 2006). PA and phospholipase D activity have been implicated in the actin-dependent tip growth of root hairs and pollen tubes (Ohashi et al., 2003; Potocký et al., 2003; Samaj et al., 2004; Monteiro et al., 2005a; Pleskot et al., 2010). Exogenous application of PA causes an elevation of actin filament levels in suspension cells, pollen, and Arabidopsis epidermal cells (Lee et al., 2003; Potocký et al., 2003; Huang et al., 2006; Li et al., 2012; Pleskot et al., 2013). Capping protein (CP) binds to the barbed end of actin filaments with high (nanomolar) affinity, dissociates quite slowly, and prevents the addition of actin subunits at this end (Huang et al., 2003, 2006; Kim et al., 2007). In the presence of phospholipids, AtCP is not able to bind to the barbed end of actin filaments (Huang et al., 2003, 2006). Furthermore, capped filament ends are uncapped by the addition of PA, allowing actin assembly from a pool of profilin-actin (Huang et al., 2006). Collectively, these data lead to a simple model whereby CP, working in concert with profilin-actin, serves to maintain tight regulation of actin assembly at filament barbed ends (Huang et al., 2006; Blanchoin et al., 2010; Henty-Ridilla et al., 2013; Pleskot et al., 2013). Furthermore, the availability of CP for filament ends can be modulated by fluxes in signaling lipids. Genetic evidence for this model was recently obtained by analyzing the dynamic behavior of actin filament ends in living Arabidopsis epidermal cells after treatment with exogenous PA (Li et al., 2012). Specifically, changes in the architecture of cortical actin arrays and dynamics of individual actin filaments that are induced by PA treatment were found to be attenuated in cp mutant cells (Li et al., 2012; Pleskot et al., 2013).Structural characterization of chicken CapZ demonstrates that the α- and β-subunits of the heterodimer form a compact structure resembling a mushroom with pseudo-two-fold rotational symmetry (Yamashita et al., 2003). Actin- and phospholipid-binding sites are conserved on the C-terminal regions, sometimes referred to as tentacles, which comprise amphipathic α-helices (Cooper and Sept, 2008; Pleskot et al., 2012). Coarse-grained molecular dynamics (CG-MD) simulations recently revealed the mechanism of chicken and AtCP association with membranes (Pleskot et al., 2012). AtCP interacts specifically with lipid bilayers through interactions between PA and the amphipathic helix of the α-subunit tentacle. Extensive polar contacts between lipid headgroups and basic residues on CP (including K278, which is unique to plant CP), as well as partial embedding of nonpolar groups into the lipid bilayer, are observed (Pleskot et al., 2012). Moreover, a glutathione S-transferase fusion protein containing the C-terminal 38 amino acids from capping protein α subunit (CPA) is sufficient to bind PA-containing liposomes in vitro (Pleskot et al., 2012). Collectively, these findings lead us to predict that AtCP will behave like a membrane-associated protein in plant cells.Additional evidence from animal and microbial cells supports the association of CP with biological membranes. In Acanthamoeba castellanii, CP is localized primarily to the hyaline ectoplasm in a region of the cytoplasm just under the plasma membrane that contains a high concentration of actin filaments (Cooper et al., 1984). Localization of CP with regions rich in actin filaments and with membranes was supported by subcellular fractionation experiments, in which CP was associated with a crude membrane fraction that included plasma membrane (Cooper et al., 1984). Further evidence demonstrates that CP localizes to cortical actin patches at sites of new cell wall growth in budding yeast (Saccharomyces cerevisiae), including the site of bud emergence. By contrast, CP did not colocalize with actin cables in S. cerevisiae (Amatruda and Cooper, 1992). CP may localize to these sites by direct interactions with membrane lipids, through binding the ends of actin filaments, or by association with another protein different from actin. In support of this hypothesis, GFP-CP fusion proteins demonstrate that sites of actin assembling in living cells contain both CP and the actin-related protein2/3 (Arp2/3) complex, and CP is located in two types of structures: (1) motile regions of the cell periphery, which reflect movement of the edge of the lamella during extension and ruffling; and (2) dynamic spots within the lamella (Schafer et al., 1998). CP has been colocalized to the F-actin patches in fission yeast (Schizosaccharomyces pombe; Kovar et al., 2005), which promotes Arp2/3-dependent nucleation and branching and limits the extent of filament elongation (Akin and Mullins, 2008). These findings lend additional support for a model whereby CP cooperates with the Arp2/3 complex to regulate actin dynamics (Nakano and Mabuchi, 2006). Activities and localization of other plant ABPs are linked to membranes. Membrane association has been linked to the assembly status of the ARP2/3 complex, an actin filament nucleator, in Arabidopsis (Kotchoni et al., 2009). SPIKE1 (SPK1), a Rho of plants (Rop)-guanine nucleotide exchange factor (GEF) and peripheral membrane protein, maintains the homeostasis of the early secretory pathway and signal integration during morphogenesis through specialized domains in the endoplasmic reticulum (ER; Zhang et al., 2010). Furthermore, Nck-associated protein1 (NAP1), a component of the suppressor of cAMP receptor/WASP-family verprolin homology protein (SCAR/WAVE) complex, strongly associates with membranes and is particularly enriched in ER membranes (Zhang et al., 2013a). Finally, a superfamily of plant ABPs, called NETWORKED proteins, was recently discovered; these link the actin cytoskeleton to various cellular membranes (Deeks et al., 2012; Hawkins et al., 2014; Wang et al., 2014).In this work, we demonstrate that CP is a membrane-associated protein in Arabidopsis. To our knowledge, this is the first direct evidence for CP-membrane association in plants. This interaction likely targets CP to cellular compartments such as the ER and Golgi. This unique location may allow CP to remodel the actin cytoskeleton in the vicinity of endomembrane compartments and/or to respond rapidly to fluxes in signaling lipids.     

Arabidopsis 3-Ketoacyl-Coenzyme A Synthase9 Is Involved in the Synthesis of Tetracosanoic Acids as Precursors of Cuticular Waxes,Suberins, Sphingolipids,and Phospholipids

Very-long-chain fatty acids (VLCFAs) with chain lengths from 20 to 34 carbons are involved in diverse biological functions such as membrane constituents, a surface barrier, and seed storage compounds. The first step in VLCFA biosynthesis is the condensation of two carbons to an acyl-coenzyme A, which is catalyzed by 3-ketoacyl-coenzyme A synthase (KCS). In this study, amino acid sequence homology and the messenger RNA expression patterns of 21 Arabidopsis (Arabidopsis thaliana) KCSs were compared. The in planta role of the KCS9 gene, showing higher expression in stem epidermal peels than in stems, was further investigated. The KCS9 gene was ubiquitously expressed in various organs and tissues, including roots, leaves, and stems, including epidermis, silique walls, sepals, the upper portion of the styles, and seed coats, but not in developing embryos. The fluorescent signals of the KCS9::enhanced yellow fluorescent protein construct were merged with those of BrFAD2::monomeric red fluorescent protein, which is an endoplasmic reticulum marker in tobacco (Nicotiana benthamiana) epidermal cells. The kcs9 knockout mutants exhibited a significant reduction in C24 VLCFAs but an accumulation of C20 and C22 VLCFAs in the analysis of membrane and surface lipids. The mutant phenotypes were rescued by the expression of KCS9 under the control of the cauliflower mosaic virus 35S promoter. Taken together, these data demonstrate that KCS9 is involved in the elongation of C22 to C24 fatty acids, which are essential precursors for the biosynthesis of cuticular waxes, aliphatic suberins, and membrane lipids, including sphingolipids and phospholipids. Finally, possible roles of unidentified KCSs are discussed by combining genetic study results and gene expression data from multiple Arabidopsis KCSs.Very-long-chain fatty acids (VLCFAs) are fatty acids of 20 or more carbons in length and are essential precursors of functionally diverse lipids, cuticular waxes, aliphatic suberins, phospholipids, sphingolipids, and seed oils in the Brassicaceae. These lipids are involved in various functions, such as acting as protective barriers between plants and the environment, impermeable barriers to water and ions, energy-storage compounds in seeds, structural components of membranes, and lipid signaling, which is involved in the hypersensitive response (Pollard et al., 2008; Kunst and Samuels, 2009; Franke et al., 2012). VLCFAs are synthesized by the microsomal fatty acid elongase complex, which catalyzes the cyclic addition of a C2 moiety obtained from malonyl-CoA to C16 or C18 acyl-CoA. The fatty acid elongation process has been shown to proceed through a series of four reactions: condensation of the C2 carbon moiety to acyl-CoA by 3-ketoacyl coenzyme A synthase (KCS), reduction of KCS by 3-ketoacyl coenzyme A reductase (KCR), dehydration of 3-hydroxyacyl-CoA by 3-hydroxyacyl-CoA dehydratase (PAS2), and reduction of trans-2,3-enoyl-CoA by trans-2-enoyl-CoA reductase (ECR). Except for KCS isoforms with redundancy, disruption of KCR1, ECR/ECERIFERUM10 (CER10), or PAS2 exhibited severe morphological abnormalities and embryo lethality, suggesting that VLCFA homeostasis is essential for plant developmental processes (Zheng et al., 2005; Bach et al., 2008; Beaudoin et al., 2009).Cuticular waxes that cover plant aerial surfaces are known to be involved in limiting nonstomatal water loss and gaseous exchanges (Boyer et al., 1997; Riederer and Schreiber, 2001), repelling lipophilic pathogenic spores and dust (Barthlott and Neinhuis, 1997), and protecting plants from UV light (Reicosky and Hanover, 1978). VLCFAs that are synthesized in the epidermal cells are either directly used or further modified into aldehydes, alkanes, secondary alcohols, ketones, primary alcohols, and wax esters for the synthesis of cuticular waxes. Reverse genetic analysis and Arabidopsis (Arabidopsis thaliana) epidermal peel microarray analysis (Suh et al., 2005) has enabled the research community to identify the functions of many genes involved in cuticular wax biosynthesis (Kunst and Samuels, 2009): CER1 (Bourdenx et al., 2011; Bernard et al., 2012), WAX2/CER3 (Chen et al., 2003; Rowland et al., 2007; Bernard et al., 2012), and MAH1(Greer et al., 2007; Wen and Jetter, 2009) have been shown to be involved in the decarbonylation pathway to form aldehydes, alkanes, secondary alcohols, and ketones, and acyl-coenzyme A reductase (FAR; Aarts et al., 1997; Rowland et al., 2006) and WSD1 (Li et al., 2008) have been shown to be involved in the decarboxylation pathway for the synthesis of primary alcohols and wax esters. The export of wax precursors to the extracellular space is mediated by a heterodimer of the ATP-binding cassette transporters in the plasma membrane (Pighin et al., 2004; Bird et al., 2007; McFarlane et al., 2010). In addition, glycosylphosphatidylinositol-anchored LTP (LTPG1) and LTPG2 contribute either directly or indirectly to the export of cuticular wax (DeBono et al., 2009; Lee et al., 2009; Kim et al., 2012).VLCFAs that are synthesized in the endodermis of primary roots, seed coats, and the chalaza-micropyle region of seeds are used as precursors for the synthesis of aliphatic suberins. The suberin layer is known to function as a barrier against uncontrolled water, gas, and ion loss and provides protection from environmental stresses and pathogens (Pollard et al., 2008; Franke et al., 2012). For aliphatic suberin biosynthesis, the ω-carbon of the VLCFAs is oxidized by the fatty acyl ω-hydroxylase (Xiao et al., 2004; Li et al., 2007; Höfer et al., 2008; Molina et al., 2008, 2009; Compagnon et al., 2009; Li-Beisson et al., 2009), and the ω-hydroxy VLCFAs are further oxidized into α,ω-dicarboxylic acids by the HOTHEAD-like oxidoreductase (Kurdyukov et al., 2006). α,ω-Dicarboxylic acids are acylated to glycerol-3-P via acyl-CoA:glycerol-3-P acyltransferase (Beisson et al., 2007; Li et al., 2007; Li-Beisson et al., 2009; Yang et al., 2010) or to ferulic acid. In addition, C18, C20, and C22 fatty acids are also reduced by FAR enzymes to primary fatty alcohols, which are a common component in root suberin (Vioque and Kolattukudy, 1997). Finally, the aliphatic suberin precursors are likely to be extensively polymerized and cross linked with the polysaccharides or lignins in the cell wall.In addition, VLCFAs are found in sphingolipids, including glycosyl inositolphosphoceramides, glycosylceramides, and ceramides and phospholipids, such as phosphatidylethanolamine (PE) and phosphatidyl-Ser (PS), which are present in the extraplastidial membrane (Pata et al., 2010; Yamaoka et al., 2011). For sphingolipid biosynthesis, VLCFA-CoAs and Ser are condensed to form 3-keto-sphinganine, which is subsequently reduced to produce sphinganine, a long chain base (LCB). LCBs are known to be further modified by 4-hydroxylation, 4-desaturation, and 8-desaturation (Lynch and Dunn, 2004; Chen et al., 2006, 2012; Pata et al., 2010). The additional VLCFAs are linked with 4-hydroxy LCBs via an amino group to form ceramides (Chen et al., 2008). The presence of VLCFA in sphingolipids may contribute to an increase of their hydrophobicity, membrane leaflet interdigitation, and the transition from a fluid to a gel phase, which is required for microdomain formation. In plants, PS is synthesized from CDP-diacylglycerol and Ser by PS synthase or through an exchange reaction between a phospholipid head group and Ser by a calcium-dependent base-exchange-type PS synthase (Vincent et al., 1999; Yamaoka et al., 2011). PE biosynthesis proceeds through decarboxylation via PS decarboxylase (Nerlich et al., 2007), the phosphoethanolamine transfer from CDP-ethanolamine to diacylglycerol (Kennedy pathway), and the exchange of the head group of PE with Ser via a base-exchange enzyme (Marshall and Kates, 1973). In particular, PS containing a relatively large amount of VLCFAs is enriched in endoplasmic reticulum (ER)-derived vesicles that may function in stabilizing small (70- to 80-nm-diameter) vesicles (Vincent et al., 2001).During the fatty acid elongation process, the first committed step is the condensation of C₂ units to acyl-CoA by KCS. Arabidopsis harbors a large family containing 21 KCS members (Joubès et al., 2008). Characterization of Arabidopsis KCS mutants with defects in VLCFA synthesis revealed in planta roles and substrate specificities (based on differences in carbon chain length and degree of unsaturation) of KCSs. For example, FAE1, a seed-specific condensing enzyme, was shown to catalyze C20 and C22 VLCFA biosynthesis for seed storage lipids (James et al., 1995). KCS6/CER6/CUT1 and KCS5/CER60 are involved in the elongation of fatty acyl-CoAs longer than C28 VLCFA for cuticular waxes in epidermis and pollen coat lipids (Millar et al., 1999; Fiebig et al., 2000; Hooker et al., 2002). KCS20 and KCS2/DAISY are functionally redundant in the two-carbon elongation to C22 VLCFA, which is required for cuticular wax and root suberin biosynthesis (Franke et al., 2009; Lee et al., 2009). When KCS1 and KCS9 were expressed in yeast (Saccharomyces cerevisiae), KCS1 showed broad substrate specificity for saturated and monounsaturated C16 to C24 acyl-CoAs and KCS9 utilized the C16 to C22 acyl-CoAs (Trenkamp et al., 2004; Blacklock and Jaworski, 2006; Paul et al., 2006). Recently, CER2 encoding putative BAHD acyltransferase was reported to be a fatty acid elongase that was involved in the elongation of C28 fatty acids for the synthesis of wax precursors (Haslam et al., 2012).In this study, the expression patterns and subcellular localization of KCS9 were examined, and an Arabidopsis kcs9 mutant was isolated to investigate the roles of KCS9 in planta. Diverse classes of lipids, including cuticular waxes, aliphatic suberins, and sphingolipids, as well as fatty acids in various organs were analyzed from the wild type, the kcs9 mutant, and complementation lines. The combined results of this study revealed that KCS9 is involved in the elongation of C22 to C24 fatty acids, which are essential precursors for the biosynthesis of cuticular waxes, aliphatic suberins, and membrane lipids, including sphingolipids. To the best of our knowledge, this is the first study where a KCS9 isoform involved in sphingolipid biosynthesis was identified.     

Composition,Assembly, and Trafficking of a Wheat Xylan Synthase Complex

Xylans play an important role in plant cell wall integrity and have many industrial applications. Characterization of xylan synthase (XS) complexes responsible for the synthesis of these polymers is currently lacking. We recently purified XS activity from etiolated wheat (Triticum aestivum) seedlings. To further characterize this purified activity, we analyzed its protein composition and assembly. Proteomic analysis identified six main proteins: two glycosyltransferases (GTs) TaGT43-4 and TaGT47-13; two putative mutases (TaGT75-3 and TaGT75-4) and two non-GTs; a germin-like protein (TaGLP); and a vernalization related protein (TaVER2). Coexpression of TaGT43-4, TaGT47-13, TaGT75-3, and TaGT75-4 in Pichia pastoris confirmed that these proteins form a complex. Confocal microscopy showed that all these proteins interact in the endoplasmic reticulum (ER) but the complexes accumulate in Golgi, and TaGT43-4 acts as a scaffold protein that holds the other proteins. Furthermore, ER export of the complexes is dependent of the interaction between TaGT43-4 and TaGT47-13. Immunogold electron microscopy data support the conclusion that complex assembly occurs at specific areas of the ER before export to the Golgi. A di-Arg motif and a long sequence motif within the transmembrane domains were found conserved at the NH₂-terminal ends of TaGT43-4 and homologous proteins from diverse taxa. These conserved motifs may control the forward trafficking of the complexes and their accumulation in the Golgi. Our findings indicate that xylan synthesis in grasses may involve a new regulatory mechanism linking complex assembly with forward trafficking and provide new insights that advance our understanding of xylan biosynthesis and regulation in plants.It is believed that Golgi-localized, multiprotein complexes synthesize plant hemicellulosic polysaccharides, including xylans. Such complexes are not well characterized in plants (Zeng et al., 2010; Atmodjo et al., 2011; Chou et al., 2012), which is in sharp contrast with mammalian and yeast cells (Jungmann and Munro, 1998; McCormick et al., 2000; Giraudo et al., 2001). Xylans are the most abundant plant hemicellulosic polysaccharides on Earth and play an important role in the integrity of cell walls, which is a key factor in plant growth. Any mutations affecting xylan backbone biosynthesis seem to result in abnormal growth of plants due mostly to thinning and weakening of secondary xylem walls, described as the irregular xylem (irx) phenotype. Thus, characterizing the xylan synthase complex (XSC) would have an impact on plant improvement, as well as many industrial applications related to food, feed, and biofuel production (Yang and Wyman, 2004; Faik, 2010). Although the Arabidopsis (Arabidopsis thaliana) irx mutants have revealed the involvement of several glycosyltransferase (GT) gene families in xylan biosynthesis (Brown et al., 2007, 2009; Lee et al., 2007, 2010; Wu et al., 2009, 2010), no XSCs have been purified/isolated from Arabidopsis tissues, and we still do not know whether some of the identified Arabidopsis GTs can assemble into functional XSCs. Furthermore, if GTs do assemble into XSCs, we don’t know the mechanisms by which plant cells control their assembly and cellular trafficking. In contrast to dicots, xylan synthase activity was recently immunopurified from etiolated wheat (Triticum aestivum) microsomes (Zeng et al., 2010). This purified wheat XS activity was shown to catalyze three activities, xylan-glucuronosyltransferase (XGlcAT), xylan-xylosyltransferase (XXylT), and xylan-arabinofuranosyltranferase (XAT), which work synergistically to synthesize xylan-type polymers in vitro (Zeng et al., 2008, 2010). This work focuses on describing protein composition, assembly, and trafficking of this purified wheat XS activity.In all eukaryotes, proteins of the secretory pathway (including GTs) are synthesized in the endoplasmic reticulum (ER) and modified as they go through the Golgi cisternae. Most proteins exit the ER from ER export sites (ERESs; Hanton et al., 2009) and use a signal-based sorting mechanism that allows them to be selectively recruited into vesicles coated by coat protein II complexes (Barlowe, 2003; Beck et al., 2008). For many Golgi-resident type II membrane proteins, di-Arg motifs, such as RR, RXR, and RRR located in their cytosolic NH₂-terminal ends, have been shown to be required for their ER export (Giraudo et al., 2003; Czlapinski and Bertozzi, 2006; Schoberer et al., 2009; Tu and Banfield, 2010). Interestingly, di-Arg motifs located ∼40 amino acids from the membrane on the cytosolic side can also be used to retrieve some type II ER-resident proteins from cis-Golgi (Schutze et al., 1994; Hardt et al., 2003; Boulaflous et al., 2009). In contrast to the signal-based sorting mechanism involved in trafficking between the ER and Golgi, the steady-state localization/retention of proteins (including GTs) in the Golgi is thought to occur through vesicular cycling. Cycling is influenced by various mechanisms, including the length and composition of the transmembrane domain (TMD) of type II GTs (Bretscher and Munro, 1993; Colley, 1997; van Vliet et al., 2003; Sousa et al., 2003; Sharpe et al., 2010), and the oligomerization/aggregation of GTs (kin hypothesis), which suggests that formation of homo- or heterooligomers of GTs in the Golgi may prevent their recruitment into clathrin-coated vesicles (Machamer, 1991; Nilsson et al., 1993; Weisz et al., 1993; Cole et al., 1996). Some Golgi-resident GTs are predicted to have a cleavable NH₂-terminal secretion signal peptide (SP) and would therefore exist as soluble proteins in the Golgi lumen. To maintain their proper Golgi localization, these processed GTs are likely part of multiprotein complexes anchored to integral membrane proteins. The fact that homologs of many of the trafficking proteins from mammalian and yeast cells are found in plants indicates that trafficking machineries of the plant secretory pathway are likely conserved (d’Enfert et al., 1992; Bar-Peled and Raikhel, 1997; Batoko et al., 2000; Pimpl et al., 2000; Phillipson et al., 2001; Hawes et al., 2008).It is becoming increasingly evident that understanding the mechanisms controlling protein-protein interaction, sorting, and trafficking of polysaccharide synthases (including XSCs) will help elucidate how plants regulate cell wall synthesis and deposition during their development. To this end, we believe that the purified wheat XS activity (Zeng et al., 2010) is an excellent model for this type of study. In this work, proteomics was used to determine the protein composition of the purified XS activity. Confocal microscopy and immunogold transmission electron microscopy (TEM) were used to investigate the assembly and trafficking of the complex. Our proteomics data showed that the purified activity contains two GTs, TaGT43-4 and TaGT47-13, two putative mutases, TaGT75-3 and TaGT75-4, and two non-GT proteins: a germin-like protein (TaGLP) belonging to cupin superfamily and a protein specific to monocots annotated as wheat vernalization-related protein 2 (TaVER2). Microscopy analyses revealed that all these proteins interact in the ER, but the assembled complexes accumulate in the Golgi. Export of these complexes from the ER is controlled by the interaction between TaGT43-4 and TaGT47-13. Characterization of the wheat XSC and its trafficking furthers our understanding of xylan biosynthesis in grasses and helps elucidate how polysaccharide synthase complexes are assembled, sorted, and maintained in different compartments of the secretory pathway.     

The Rubisco Small Subunit Is Involved in Tobamovirus Movement and Tm-22-Mediated Extreme Resistance

The multifunctional movement protein (MP) of Tomato mosaic tobamovirus (ToMV) is involved in viral cell-to-cell movement, symptom development, and resistance gene recognition. However, it remains to be elucidated how ToMV MP plays such diverse roles in plants. Here, we show that ToMV MP interacts with the Rubisco small subunit (RbCS) of Nicotiana benthamiana in vitro and in vivo. In susceptible N. benthamiana plants, silencing of NbRbCS enabled ToMV to induce necrosis in inoculated leaves, thus enhancing virus local infectivity. However, the development of systemic viral symptoms was delayed. In transgenic N. benthamiana plants harboring Tobacco mosaic virus resistance-2² (Tm-2²), which mediates extreme resistance to ToMV, silencing of NbRbCS compromised Tm-2²-dependent resistance. ToMV was able to establish efficient local infection but was not able to move systemically. These findings suggest that NbRbCS plays a vital role in tobamovirus movement and plant antiviral defenses.Plant viruses use at least one movement protein (MP) to facilitate viral spread between plant cells via plasmodesmata (PD; Lucas and Gilbertson, 1994; Ghoshroy et al., 1997). Among viral MPs, the MP of tobamoviruses, such as Tobacco mosaic virus (TMV) and its close relative Tomato mosaic virus (ToMV), is the best characterized. TMV MP specifically accumulates in PD and modifies the plasmodesmatal size exclusion limit in mature source leaves or tissues (Wolf et al., 1989; Deom et al., 1990; Ding et al., 1992). TMV MP and viral genomic RNA form a mobile ribonucleoprotein complex that is essential for cell-to-cell movement of viral infection (Watanabe et al., 1984; Deom et al., 1987; Citovsky et al., 1990, 1992; Kiselyova et al., 2001; Kawakami et al., 2004; Waigmann et al., 2007). TMV MP also enhances intercellular RNA silencing (Vogler et al., 2008) and affects viral symptom development, host range, and host susceptibility to virus (Dardick et al., 2000; Bazzini et al., 2007). Furthermore, ToMV MP is identified as an avirulence factor that is recognized by tomato (Solanum lycopersicum) resistance proteins Tobacco mosaic virus resistance-2 (Tm-2) and Tm-2² (Meshi et al., 1989; Lanfermeijer et al., 2004). Indeed, tomato Tm-2² confers extreme resistance against TMV and ToMV in tomato plants and even in heterologous tobacco (Nicotiana tabacum) plants (Lanfermeijer et al., 2003, 2004).To date, several host factors that interact with TMV MP have been identified. These TMV MP-binding host factors include cell wall-associated proteins such as pectin methylesterase (Chen et al., 2000), calreticulin (Meshi et al., 1989), ANK1 (Ueki et al., 2010), and the cellular DnaJ-like protein MPIP1 (Shimizu et al., 2009). Many cytoskeletal components such as actin filaments (McLean et al., 1995), microtubules (Heinlein et al., 1995), and the microtubule-associated proteins MPB2C (Kragler et al., 2003) and EB1a (Brandner et al., 2008) also interact with TMV MP. Most of these factors are involved in TMV cell-to-cell movement.Rubisco catalyzes the first step of CO₂ assimilation in photosynthesis and photorespiration. The Rubisco holoenzyme is a heteropolymer consisting of eight large subunits (RbCLs) and eight small subunits (RbCSs). RbCL was reported to interact with the coat protein of Potato virus Y (Feki et al., 2005). Both RbCS and RbCL were reported to interact with the P3 proteins encoded by several potyviruses, including Shallot yellow stripe virus, Onion yellow dwarf virus, Soybean mosaic virus, and Turnip mosaic virus (Lin et al., 2011). Proteomic analysis of the plant-virus interactome revealed that RbCS participates in the formation of virus complexes of Rice yellow mottle virus (Brizard et al., 2006). However, the biological function of Rubisco in viral infection remains unknown.In this study, we show that RbCS plays an essential role in virus movement, host susceptibility, and Tm-2²-mediated extreme resistance in the ToMV-host plant interaction.     

Engineering Monolignol p-Coumarate Conjugates into Poplar and Arabidopsis Lignins

Lignin acylation, the decoration of hydroxyls on lignin structural units with acyl groups, is common in many plant species. Monocot lignins are decorated with p-coumarates by the polymerization of monolignol p-coumarate conjugates. The acyltransferase involved in the formation of these conjugates has been identified in a number of model monocot species, but the effect of monolignol p-coumarate conjugates on lignification and plant growth and development has not yet been examined in plants that do not inherently possess p-coumarates on their lignins. The rice (Oryza sativa) p-COUMAROYL-Coenzyme A MONOLIGNOL TRANSFERASE gene was introduced into two eudicots, Arabidopsis (Arabidopsis thaliana) and poplar (Populus alba × grandidentata), and a series of analytical methods was used to show the incorporation of the ensuing monolignol p-coumarate conjugates into the lignin of these plants. In poplar, specifically, the addition of these conjugates did not occur at the expense of the naturally incorporated monolignol p-hydroxybenzoates. Plants expressing the p-COUMAROYL-Coenzyme A MONOLIGNOL TRANSFERASE transgene can therefore produce monolignol p-coumarate conjugates essentially without competing with the formation of other acylated monolignols and without drastically impacting normal monolignol production.Lignification of plant cell walls prototypically involves the polymerization of the monolignols (MLs), p-coumaryl alcohol, coniferyl alcohol (CA), and sinapyl alcohol (SA), predominantly by stepwise radical coupling of each monomer to the phenolic end of the growing polymer (Sarkanen and Ludwig, 1971; Boerjan et al., 2003; Ralph et al., 2004). The contribution of various MLs to the lignins depends on plant species, cell type, plant tissue, and tissue age. Although the majority of the lignin polymer is derived from these three MLs, the lignification process has a high degree of metabolic plasticity (Boerjan et al., 2003; Ralph et al., 2004; Ralph, 2007; Vanholme et al., 2012). Of particular interest are ML conjugates in which the ester group can be acetate (Ac; Sarkanen et al., 1967; Ralph, 1996; Ralph and Lu, 1998; Del Río et al., 2007; del Río et al., 2008; Martínez et al., 2008), p-hydroxybenzoate (pBz; Venverloo, 1971; Monties and Lapierre, 1981; Landucci et al., 1992; Tomimura, 1992a, 1992b; Hibino et al., 1994; Sun et al., 1999; Kuroda et al., 2001; Lu et al., 2004, 2015; Morreel et al., 2004; Rencoret et al., 2013), p-coumarate (pCA; Monties and Lapierre, 1981; Ralph et al., 1994; Crestini and Argyropoulos, 1997; del Río et al., 2008, 2012a, 2012b; Withers et al., 2012; Rencoret et al., 2013; Petrik et al., 2014), or ferulate (FA; Grabber et al., 2008; Ralph, 2010; Wilkerson et al., 2014). In all cases, the MLs are acylated before polymerization as proven by the presence in the lignins of unique β-β coupling products that only arise when one or both of the MLs are acylated, preventing the formation of the typical resinols from internal trapping of the quinone methide intermediates by the γ-OH (Lu and Ralph, 2002, 2008; Del Río et al., 2007; Lu et al., 2015).The BAHD acyltransferase, FERULOYL-CoA MONOLIGNOL TRANSFERASE (FMT), was recently identified in Angelica sinensis and transformed into poplar (Populus alba × grandidentata), which naturally incorporates other acylated MLs, namely ML-pBz conjugates, into its lignin (Wilkerson et al., 2014). Plants that incorporate ML-FAs into their lignins have the potential to be particularly important economically, because their lignin backbones are permeated with readily cleavable ester bonds, facilitating lignin breakdown and removal under alkaline pretreatment conditions. Determining the extent to which ML-FAs are incorporated into the lignin polymer is, however, extremely difficult because of the diversity of products generated during the polymerization events, which is described in the supplemental information in Wilkerson et al., 2014.There is currently only one technique, derivatization followed by reductive cleavage (DFRC), that can release diagnostic chemical marker compounds from lignins containing ML-FAs (Lu and Ralph, 2014; Wilkerson et al., 2014). The DFRC method selectively cleaves β-ethers while leaving ester linkages intact. This technique was recently used to show that ML-FA conjugates are fully incorporated into the lignin of the FMT poplar (Wilkerson et al., 2014), but the extent of incorporation, the spatial distribution, the exact mechanism of delivery to the developing cell wall, and the efficiency of incorporation remain largely unknown.The biological role of pCA in lignin has been highly speculative. It is hypothesized that the pCA moieties may function as a radical sensitizer (Takahama and Oniki, 1996, 1997; Takahama et al., 1996; Ralph et al., 2004; Hatfield et al., 2008; Ralph, 2010). Peroxidases and/or laccases readily oxidize pCA to a radical but are poor oxidizers for SA. Free radicals of pCA readily undergo radical transfer to SA, which in turn, forms a homodimer or couples to the end of a growing polymer chain. Conjugating pCA to an ML, like SA, to form SA-pCA, the most prevalent ML-pCA conjugate in grasses, creates a compound with a built-in radical sensitizer that can participate in the polymerization event. The prevalence of these conjugates in potential biofuel crops and the impact that these ester-linked conjugates have on the lignin polymer during pretreatment and downstream fermentation processes have driven the search to find the genes and their enzymes responsible for acylating MLs in monocots (Withers et al., 2012; Marita et al., 2014; Petrik et al., 2014; Wilkerson et al., 2014).In rice (Oryza sativa), enzymes have been characterized that function specifically in the addition of pCA onto hemicelluloses (Bartley et al., 2013) or lignin (Withers et al., 2012; Petrik et al., 2014). The p-COUMAROYL-CoA MONOLIGNOL TRANSFERASE (PMT) was identified as one of many grass-specific BAHD acyltransferases produced by rice and found to coexpress with many ML biosynthetic enzymes (Withers et al., 2012). The enzyme preferentially forms a γ-ester through its specificity toward p-coumaroyl-CoA and an ML, and has kinetic efficiency with p-coumaryl alcohol > SA > CA. In most grasses, the PMT enzyme predominantly produces SA-pCA conjugates that are then incorporated into the lignin polymer (Petrik et al., 2014).To test the role of PMT during cell wall lignification, genetic manipulation of PMT genes has been performed in Brachypodium distachyon and maize (Zea mays), two model monocots. The suppression and overexpression of a BdPMT revealed the PMT to be involved only in the acylation of MLs before polymerization and not in the acylation of hemicelluloses (Petrik et al., 2014). RNA interference-mediated suppression of BdPMT resulted in decreased incorporation of ML-pCA conjugates into the cell wall without adversely affecting growth, height, or digestibility of the mature plants. Even deleterious mutations in the BdPMT gene, which resulted in a complete absence of pCA-acylating B. distachyon lignins, did not affect plant growth or development (Petrik et al., 2014). The arabinose-bound FA and pCA levels remained virtually unchanged in the PMT-misregulated plants, illustrating the specificity of the PMT enzyme for the p-coumaroyl-CoA substrate and its ML acylation. The PMT enzyme identified in maize (pCAT = ZmPMT) also displayed the highest catalytic efficiency with p-coumaroyl-CoA and SA as substrates (Marita et al., 2014). RNAi-mediated suppression of ZmPMT also resulted in decreased production of the ML conjugates. The effect on the lignin polymer when introducing PMT into plants that do not normally express a homologous enzyme is, however, unknown.pCAs, because they favor radical transfer over radical coupling, are overwhelmingly seen as free-phenolic pendant entities on the lignin polymer (Ralph et al., 1994; Ralph, 2010). As a result, the pCA itself can be completely quantified by simple saponification. The units to which the pCA is attached are, like their normal ML-derived counterparts, not fully releasable from lignin as identifiable monomers (during degradative reactions), but the pCA’s terminal location makes p-coumaroylated units more readily releasable and detectable than if they participated in lignification (as FAs do). Examining the effect of PMT and its resulting conjugates on lignification in plants that do not naturally produce such conjugates will contribute to our understanding of the role of PMT in lignification in general.In this study, we aimed to assess the ability of the model eudicot plants Arabidopsis (Arabidopsis thaliana) and poplar, neither of which naturally produces ML-pCA conjugates, to express a PMT gene and incorporate these novel conjugates into their cell wall lignins. We also investigated the effect that the introduction of PMT has on the native levels of ML-pBz conjugates in poplar lignin. Various analytical techniques were optimized and used to examine the cell walls of the transgenic plants for pCA conjugates and determine whether they were specifically incorporated into the lignin polymer in the cell wall.