Endomembrane Trafficking Protein SEC24A Regulates Cell Size Patterning in Arabidopsis

Size is a critical property of a cell, but how it is determined is still not well understood. The sepal epidermis of Arabidopsis (Arabidopsis thaliana) contains cells with a diversity of sizes ranging from giant cells to small cells. Giant cells have undergone endoreduplication, a specialized cell cycle in which cells replicate their DNA but fail to divide, becoming polyploid and enlarged. Through forward genetics, we have identified a new mutant with ectopic giant cells covering the sepal epidermis. Surprisingly, the mutated gene, SEC24A, encodes a coat protein complex II vesicle coat subunit involved in endoplasmic reticulum-to-Golgi trafficking in the early secretory pathway. We show that the ectopic giant cells of sec24a-2 are highly endoreduplicated and that their formation requires the activity of giant cell pathway genes LOSS OF GIANT CELLS FROM ORGANS, DEFECTIVE KERNEL1, and Arabidopsis CRINKLY4. In contrast to other trafficking mutants, cytokinesis appears to occur normally in sec24a-2. Our study reveals an unexpected yet specific role of SEC24A in endoreduplication and cell size patterning in the Arabidopsis sepal.Size is a fundamental characteristic of a cell, but how cell size is determined is still not well understood in most living organisms (Marshall et al., 2012). Cells of different types typically have characteristic sizes, indicating that size is carefully regulated to fit cell functions during differentiation. At the simplest level, cell size is determined by growth and division. Although many factors regulating these two processes have been studied, how they are comprehensively regulated to achieve specific size outcomes remains unclear.The sepal of Arabidopsis (Arabidopsis thaliana) is an excellent model to study the regulation of cell size because it exhibits a characteristic pattern of giant cells interspersed in between small cells. The giant cells are large cells that span about one-fifth the length of the sepal (approximately 360 μm), while the smallest cells only reach to about 10 μm (Roeder et al., 2010). Previously, we have shown that variability in cell division times is sufficient to produce the cell size pattern (Roeder et al., 2010). The giant cells stop dividing and enter endoreduplication, a specialized cell cycle in which the cell replicates its DNA but skips mitosis to continue growing (Edgar and Orr-Weaver, 2001; Sugimoto-Shirasu and Roberts, 2003; Inzé and De Veylder, 2006; Breuer et al., 2010). Alongside the giant cells, the smaller cells continue dividing mitotically. Giant cells and small cells are different cell types, as they can be distinguished by the expression pattern of two independent enhancers. Furthermore, mutant screens have shown that genes involved in epidermal specification and cell cycle regulation are crucial for sepal cell size patterning. DEFECTIVE KERNEL1 (DEK1), Arabidopsis thaliana MERISTEM LAYER1 (ATML1), Arabidopsis CRINKLY4 (ACR4), and HOMEODOMAIN GLABROUS11 first establish the identity of giant cells, and then the cyclin dependent kinase inhibitor LOSS OF GIANT CELLS FROM ORGANS (LGO) influences the probability with which cells enter endoreduplication. Endoreduplication can further suppress the identity of small cells through an unknown mechanism (Roeder et al., 2010, 2012). The number of giant cells influences the curvature of the sepal, which is important for protecting the flower (Roeder et al., 2012). Therefore, cell size patterning ensures the protective role of sepals at the physiological level.The secretory pathway in eukaryotes is crucial for cells to maintain membrane homeostasis and protein localization. Proteins destined for the cell surface are first translated on the rough endoplasmic reticulum (ER) and then incorporated into coat protein complex II (COPII) vesicles that bud from ER membranes on the way to the Golgi apparatus. COPII machinery is highly conserved in eukaryotes, and each COPII component acts sequentially on the surface of the ER (Bickford et al., 2004; Marti et al., 2010; Zanetti et al., 2012). Vesicle coat assembly is initiated by SEC12, an ER membrane-anchored guanine nucleotide exchange factor (Barlowe and Schekman, 1993). SEC12 exchanges GDP with GTP on the small GTPase Secretion-associated RAS-related protein1 (SAR1), which increases the membrane affinity of SAR1. The ER membrane-bound SAR1 subsequently brings the SEC23/SEC24 subunits to form the prebudding complex, and eventually SEC13/SEC31 are recruited to increase rigidity of the COPII vesicle coat (Nakano and Muramatsu, 1989; Barlowe et al., 1994; Shaywitz et al., 1997; Aridor et al., 1998; Kuehn et al., 1998; Stagg et al., 2006; Copic et al., 2012). For COPII vesicles to fuse with the target membrane, superfamily N-ethylmaleimide-sensitive factor adaptor protein receptors (SNAREs) must be incorporated by SEC24 (Mossessova et al., 2003; Lipka et al., 2007; Mancias and Goldberg, 2008). In addition to its role in SNARE packaging, SEC24 also binds and loads secretory cargo proteins (Miller et al., 2003). Both the cargo and SNARE specificities are determined by the correspondence between the SEC24 isoform and the various ER export signals of cargoes and SNAREs (Barlowe., 2003; Miller et al., 2003; Mossessova et al., 2003; Mancias and Goldberg, 2008). The Arabidopsis genome encodes four SEC24 isoforms, SEC24A to SEC24D; how they differentially regulate trafficking is unknown (Bassham et al., 2008). Likewise, SEC24-cargo/SNARE interactions remain elusive in plants.Secretion defects in plants often lead to cell division defects due to the unique mechanisms of plants cytokinesis (Sylvester, 2000; Jürgens, 2005). In many eukaryotes other than plants, cytokinesis is accomplished by contraction of the cleavage furrow at the division plane. By contrast, cytokinesis in plants requires de novo secretion of vesicles to the division plane, after formation of the phragmoplast as the scaffold for delivery. Homotypic vesicle fusion sets up the early cell plate, which then expands laterally by fusing with other arriving vesicles (Balasubramanian et al., 2004; Jürgens, 2005; Reichardt et al., 2007). Hence, disruption of secretion in plants can often result in cytokinesis defects. For instance, a mutation in the SNARE KNOLLE leads to enlarged embryo cells with multiple nuclei (Lukowitz et al., 1996).Another common phenotype observed in secretion-deficient plants is abnormal auxin responses. The phytohormone auxin acts as a prominent signal in Arabidopsis development, and auxin influx/efflux carriers are essential in directing auxin transport and creating local maxima in an auxin gradient (Reinhardt et al., 2003; Heisler et al., 2005; Jönsson et al., 2006; Smith et al., 2006; Vanneste and Friml, 2009). To maintain appropriate auxin gradients, the subcellular localization of auxin carriers must be delicately regulated. Thus, auxin responses are highly sensitive to trafficking perturbations in plants (Geldner et al., 2003; Grunewald and Friml, 2010).Here, we have identified a new mutant with ectopic giant cells. Through positional cloning, we determined that the mutation occurs in the SEC24A gene, which encodes the cargo-binding subunit of the COPII vesicle complex. In addition to altered cell size, this unique sec24a-2 allele shows pleiotropic defects, including dwarfism, which have not been reported previously for other SEC24A alleles (Faso et al., 2009; Nakano et al., 2009; Conger et al., 2011). Although the mutant is developmentally aberrant, both cytokinesis and auxin response appear normal in sec24a-2, unlike other transport mutants. Instead, we find SEC24A regulates cell size specifically via the giant cell development pathway. Thus, our data reveal an unexpected role of SEC24A in endoreduplication and cell size patterning in the Arabidopsis sepal.     

Spore Density Determines Infection Strategy by the Plant Pathogenic Fungus Plectosphaerella cucumerina

Necrotrophic and biotrophic pathogens are resisted by different plant defenses. While necrotrophic pathogens are sensitive to jasmonic acid (JA)-dependent resistance, biotrophic pathogens are resisted by salicylic acid (SA)- and reactive oxygen species (ROS)-dependent resistance. Although many pathogens switch from biotrophy to necrotrophy during infection, little is known about the signals triggering this transition. This study is based on the observation that the early colonization pattern and symptom development by the ascomycete pathogen Plectosphaerella cucumerina (P. cucumerina) vary between inoculation methods. Using the Arabidopsis (Arabidopsis thaliana) defense response as a proxy for infection strategy, we examined whether P. cucumerina alternates between hemibiotrophic and necrotrophic lifestyles, depending on initial spore density and distribution on the leaf surface. Untargeted metabolome analysis revealed profound differences in metabolic defense signatures upon different inoculation methods. Quantification of JA and SA, marker gene expression, and cell death confirmed that infection from high spore densities activates JA-dependent defenses with excessive cell death, while infection from low spore densities induces SA-dependent defenses with lower levels of cell death. Phenotyping of Arabidopsis mutants in JA, SA, and ROS signaling confirmed that P. cucumerina is differentially resisted by JA- and SA/ROS-dependent defenses, depending on initial spore density and distribution on the leaf. Furthermore, in situ staining for early callose deposition at the infection sites revealed that necrotrophy by P. cucumerina is associated with elevated host defense. We conclude that P. cucumerina adapts to early-acting plant defenses by switching from a hemibiotrophic to a necrotrophic infection program, thereby gaining an advantage of immunity-related cell death in the host.Plant pathogens are often classified as necrotrophic or biotrophic, depending on their infection strategy (Glazebrook, 2005; Nishimura and Dangl, 2010). Necrotrophic pathogens kill living host cells and use the decayed plant tissue as a substrate to colonize the plant, whereas biotrophic pathogens parasitize living plant cells by employing effector molecules that suppress the host immune system (Pel and Pieterse, 2013). Despite this binary classification, the majority of pathogenic microbes employ a hemibiotrophic infection strategy, which is characterized by an initial biotrophic phase followed by a necrotrophic infection strategy at later stages of infection (Perfect and Green, 2001). The pathogenic fungi Magnaporthe grisea, Sclerotinia sclerotiorum, and Mycosphaerella graminicola, the oomycete Phytophthora infestans, and the bacterial pathogen Pseudomonas syringae are examples of hemibiotrophic plant pathogens (Perfect and Green, 2001; Koeck et al., 2011; van Kan et al., 2014; Kabbage et al., 2015).Despite considerable progress in our understanding of plant resistance to necrotrophic and biotrophic pathogens (Glazebrook, 2005; Mengiste, 2012; Lai and Mengiste, 2013), recent debate highlights the dynamic and complex interplay between plant-pathogenic microbes and their hosts, which is raising concerns about the use of infection strategies as a static tool to classify plant pathogens. For instance, the fungal genus Botrytis is often labeled as an archetypal necrotroph, even though there is evidence that it can behave as an endophytic fungus with a biotrophic lifestyle (van Kan et al., 2014). The rice blast fungus Magnaporthe oryzae, which is often classified as a hemibiotrophic leaf pathogen (Perfect and Green, 2001; Koeck et al., 2011), can adopt a purely biotrophic lifestyle when infecting root tissues (Marcel et al., 2010). It remains unclear which signals are responsible for the switch from biotrophy to necrotrophy and whether these signals rely solely on the physiological state of the pathogen, or whether host-derived signals play a role as well (Kabbage et al., 2015).The plant hormones salicylic acid (SA) and jasmonic acid (JA) play a central role in the activation of plant defenses (Glazebrook, 2005; Pieterse et al., 2009, 2012). The first evidence that biotrophic and necrotrophic pathogens are resisted by different immune responses came from Thomma et al. (1998), who demonstrated that Arabidopsis (Arabidopsis thaliana) genotypes impaired in SA signaling show enhanced susceptibility to the biotrophic pathogen Hyaloperonospora arabidopsidis (formerly known as Peronospora parastitica), while JA-insensitive genotypes were more susceptible to the necrotrophic fungus Alternaria brassicicola. In subsequent years, the differential effectiveness of SA- and JA-dependent defense mechanisms has been confirmed in different plant-pathogen interactions, while additional plant hormones, such as ethylene, abscisic acid (ABA), auxins, and cytokinins, have emerged as regulators of SA- and JA-dependent defenses (Bari and Jones, 2009; Cao et al., 2011; Pieterse et al., 2012). Moreover, SA- and JA-dependent defense pathways have been shown to act antagonistically on each other, which allows plants to prioritize an appropriate defense response to attack by biotrophic pathogens, necrotrophic pathogens, or herbivores (Koornneef and Pieterse, 2008; Pieterse et al., 2009; Verhage et al., 2010).In addition to plant hormones, reactive oxygen species (ROS) play an important regulatory role in plant defenses (Torres et al., 2006; Lehmann et al., 2015). Within minutes after the perception of pathogen-associated molecular patterns, NADPH oxidases and apoplastic peroxidases generate early ROS bursts (Torres et al., 2002; Daudi et al., 2012; O’Brien et al., 2012), which activate downstream defense signaling cascades (Apel and Hirt, 2004; Torres et al., 2006; Miller et al., 2009; Mittler et al., 2011; Lehmann et al., 2015). ROS play an important regulatory role in the deposition of callose (Luna et al., 2011; Pastor et al., 2013) and can also stimulate SA-dependent defenses (Chaouch et al., 2010; Yun and Chen, 2011; Wang et al., 2014; Mammarella et al., 2015). However, the spread of SA-induced apoptosis during hyperstimulation of the plant immune system is contained by the ROS-generating NADPH oxidase RBOHD (Torres et al., 2005), presumably to allow for the sufficient generation of SA-dependent defense signals from living cells that are adjacent to apoptotic cells. Nitric oxide (NO) plays an additional role in the regulation of SA/ROS-dependent defense (Trapet et al., 2015). This gaseous molecule can stimulate ROS production and cell death in the absence of SA while preventing excessive ROS production at high cellular SA levels via S-nitrosylation of RBOHD (Yun et al., 2011). Recently, it was shown that pathogen-induced accumulation of NO and ROS promotes the production of azelaic acid, a lipid derivative that primes distal plants for SA-dependent defenses (Wang et al., 2014). Hence, NO, ROS, and SA are intertwined in a complex regulatory network to mount local and systemic resistance against biotrophic pathogens. Interestingly, pathogens with a necrotrophic lifestyle can benefit from ROS/SA-dependent defenses and associated cell death (Govrin and Levine, 2000). For instance, Kabbage et al. (2013) demonstrated that S. sclerotiorum utilizes oxalic acid to repress oxidative defense signaling during initial biotrophic colonization, but it stimulates apoptosis at later stages to advance necrotrophic colonization. Moreover, SA-induced repression of JA-dependent resistance not only benefits necrotrophic pathogens but also hemibiotrophic pathogens after having switched from biotrophy to necrotrophy (Glazebrook, 2005; Pieterse et al., 2009, 2012).Plectosphaerella cucumerina ((P. cucumerina, anamorph Plectosporum tabacinum) anamorph Plectosporum tabacinum) is a filamentous ascomycete fungus that can survive saprophytically in soil by decomposing plant material (Palm et al., 1995). The fungus can cause sudden death and blight disease in a variety of crops (Chen et al., 1999; Harrington et al., 2000). Because P. cucumerina can infect Arabidopsis leaves, the P. cucumerina-Arabidopsis interaction has emerged as a popular model system in which to study plant defense reactions to necrotrophic fungi (Berrocal-Lobo et al., 2002; Ton and Mauch-Mani, 2004; Carlucci et al., 2012; Ramos et al., 2013). Various studies have shown that Arabidopsis deploys a wide range of inducible defense strategies against P. cucumerina, including JA-, SA-, ABA-, and auxin-dependent defenses, glucosinolates (Tierens et al., 2001; Sánchez-Vallet et al., 2010; Gamir et al., 2014; Pastor et al., 2014), callose deposition (García-Andrade et al., 2011; Gamir et al., 2012, 2014; Sánchez-Vallet et al., 2012), and ROS (Tierens et al., 2002; Sánchez-Vallet et al., 2010; Barna et al., 2012; Gamir et al., 2012, 2014; Pastor et al., 2014). Recent metabolomics studies have revealed large-scale metabolic changes in P. cucumerina-infected Arabidopsis, presumably to mobilize chemical defenses (Sánchez-Vallet et al., 2010; Gamir et al., 2014; Pastor et al., 2014). Furthermore, various chemical agents have been reported to induce resistance against P. cucumerina. These chemicals include β-amino-butyric acid, which primes callose deposition and SA-dependent defenses, benzothiadiazole (BTH or Bion; Görlach et al., 1996; Ton and Mauch-Mani, 2004), which activates SA-related defenses (Lawton et al., 1996; Ton and Mauch-Mani, 2004; Gamir et al., 2014; Luna et al., 2014), JA (Ton and Mauch-Mani, 2004), and ABA, which primes ROS and callose deposition (Ton and Mauch-Mani, 2004; Pastor et al., 2013). However, among all these studies, there is increasing controversy about the exact signaling pathways and defense responses contributing to plant resistance against P. cucumerina. While it is clear that JA and ethylene contribute to basal resistance against the fungus, the exact roles of SA, ABA, and ROS in P. cucumerina resistance vary between studies (Thomma et al., 1998; Ton and Mauch-Mani, 2004; Sánchez-Vallet et al., 2012; Gamir et al., 2014).This study is based on the observation that the disease phenotype during P. cucumerina infection differs according to the inoculation method used. We provide evidence that the fungus follows a hemibiotrophic infection strategy when infecting from relatively low spore densities on the leaf surface. By contrast, when challenged by localized host defense to relatively high spore densities, the fungus switches to a necrotrophic infection program. Our study has uncovered a novel strategy by which plant-pathogenic fungi can take advantage of the early immune response in the host plant.     

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.     

Engineering Triterpene and Methylated Triterpene Production in Plants Provides Biochemical and Physiological Insights into Terpene Metabolism

Linear, branch-chained triterpenes, including squalene (C30), botryococcene (C30), and their methylated derivatives (C31–C37), generated by the green alga Botryococcus braunii race B have received significant attention because of their utility as chemical and biofuel feedstocks. However, the slow growth habit of B. braunii makes it impractical as a production system. In this study, we evaluated the potential of generating high levels of botryococcene in tobacco (Nicotiana tabacum) plants by diverting carbon flux from the cytosolic mevalonate pathway or the plastidic methylerythritol phosphate pathway by the targeted overexpression of an avian farnesyl diphosphate synthase along with two versions of botryococcene synthases. Up to 544 µg g⁻¹ fresh weight of botryococcene was achieved when this metabolism was directed to the chloroplasts, which is approximately 90 times greater than that accumulating in plants engineered for cytosolic production. To test if methylated triterpenes could be produced in tobacco, we also engineered triterpene methyltransferases (TMTs) from B. braunii into wild-type plants and transgenic lines selected for high-level triterpene accumulation. Up to 91% of the total triterpene contents could be converted to methylated forms (C31 and C32) by cotargeting the TMTs and triterpene biosynthesis to the chloroplasts, whereas only 4% to 14% of total triterpenes were methylated when this metabolism was directed to the cytoplasm. When the TMTs were overexpressed in the cytoplasm of wild-type plants, up to 72% of the total squalene was methylated, and total triterpene (C30+C31+C32) content was elevated 7-fold. Altogether, these results point to innate mechanisms controlling metabolite fluxes, including a homeostatic role for squalene.Terpenes and terpenoids represent a distinct class of natural products (Buckingham, 2003) that are derived from two universal five-carbon precursors: isopentenyl diphosphate (IPP) and dimethylallyl diphosphate (DMAPP). In eukaryotic fungi and animals, IPP and DMAPP are synthesized via the mevalonate (MVA) pathway, whereas in prokaryotes, they are synthesized via the methylerythritol phosphate (MEP) pathway. In higher plants, the pathways are present in separate compartments and are believed to operate independently. The MVA pathway in the cytoplasm is predominantly responsible for sesquiterpene (C15), triterpene (C30), and polyprenol (greater than C45) biosynthesis and associated with the endoplasmic reticulum (ER) system. The MEP pathway resides in plastids and is dedicated to monoterpenes (C10), diterpenes (C20), carotenoids (C40), and long-chain phytol biosynthesis. All these compounds are usually produced by plants for a variety of physiological (i.e. hormones, aliphatic membrane anchors, and maintaining membrane structure) and ecological (i.e. defense compounds and insect/animal attractants) roles (Kempinski et al., 2015). Terpenes are also important for various industrial applications, ranging from flavors and fragrances (Schwab et al., 2008) to medicines (Dewick, 2009; Niehaus et al., 2011; Shelar, 2011).The utility of terpenes as chemical and biofuel feedstocks has also received considerable attention recently. Isoprenoid-derived biofuels include farnesane (Renninger and McPhee, 2008; Rude and Schirmer, 2009), bisabolene (Peralta-Yahya et al., 2011), pinene dimers (Harvey et al., 2010), isopentenal (Withers et al., 2007), and botryococcene (Moldowan and Seifert, 1980; Hillen et al., 1982; Glikson et al., 1989; Mastalerz and Hower, 1996). The richness of branches within these hydrocarbon scaffolds correlate with their high-energy content, which enables them to serve as suitable alternatives to crude petroleum (Peralta-Yahya and Keasling, 2010). Indeed, some of them are already major contributors to current-day petroleum-based fuels. One of the best examples of this is the triterpene oil accumulating in the green alga Botryococcus braunii race B, which is considered a major progenitor to oil and coal shale deposits (Moldowan and Seifert, 1980). This alga has been well studied, and the major constituents of its prodigious hydrocarbon oil are a group of triterpenes including squalene (C30), organism-specific botryococcene (C30), methylated squalene (C31–C34), and methylated botryococcene (C31–C37; Metzger et al., 1988; Huang and Poulter, 1989; Okada et al., 1995), which can be readily converted into all classes of combustible fuels under hydrocracking conditions (Hillen et al., 1982).The unique biosynthetic mechanism for the triterpenes in B. braunii was recently described by Niehaus et al. (2011), and a series of novel squalene synthase-like genes were identified (Fig. 1). In short, squalene synthase-like enzyme, SSL-1, performs a head-to-head condensation of two farnesyl diphosphate (FPP) molecules into presqualene diphosphate, followed by a reductive rearrangement to yield squalene (C30) by the enzyme SSL-2, or is converted by SSL-3 to form botryococcene through a different reductive rearrangement (Niehaus et al., 2011). Methylated derivatives are the dominant triterpene species generated by B. braunii race B (Metzger, 1985; Metzger et al., 1988), and these derivatives are known to yield higher quality fuels due to their high energy content and the hydrocracking products derived by virtue of having more hydrocarbon branches. Triterpene methyltransferases (TMTs) that can methylate squalene and botryococcene have been successfully characterized by Niehaus et al. (2012). TRITERPENE METHYLTRANSFERASE1 (TMT-1) and TMT-2 prefer squalene C30 as their substrate for the production of monomethylated (C31) or dimethylated (C32) squalene, while TMT-3 prefers botryococcene as its substrate for the biosynthesis of monomethylated (C31) or dimethylated (C32) botryococcene (Fig. 1). These TMTs are believed to be insoluble enzymes; they exhibit large hydrophobic areas, and their activities were only observed in vitro using yeast microsomal preparations (no activity was observed when expressed in bacteria; Niehaus et al., 2012).Open in a separate windowFigure 1.Depiction of the catalytic roles of novel SSL and TMT enzymes in B. braunii race B and their putative contributions to the triterpene constituents (Niehaus et al., 2011; Niehaus et al., 2012). SSL-1 catalyzes the condensation of two farnesyl diphosphate (FPP) molecules to presqualene diphosphate (PSPP), which is converted to either squalene or botryococcene by SSL-2 or SSL-3, respectively. Squalene can also be synthesized directly from the condensation of two FPP molecules catalyzed by squalene synthase (SQS). TMT-1 and TMT-2 transfer the methyl donor group from S-adenosylmethionine (SAM) to squalene to form monomethylated and dimethylated squalene, whereas TMT-3 acts on botryococcene to form monomethylated and dimethylated botryococcene (Niehaus et al., 2012).Like the majority of identified methyltransferases, these TMTs utilize the methyl donor S-adenosyl methionine (SAM), which is ubiquitous in prokaryotes and eukaryotes (Scheer et al., 2011; Liscombe et al., 2012). In plants, SAM is one of the most abundant cofactors (Fontecave et al., 2004; Sauter et al., 2013) and is synthesized exclusively in the cytosol (Wallsgrove et al., 1983; Ravanel et al., 1998, 2004; Bouvier et al., 2006). While it is used predominantly as a methyl donor in the methylation reaction (Ravanel et al., 2004), it also serves as the primary precursor for the biosynthesis of ethylene (Wang et al., 2002b), polyamines (Kusano et al., 2008), and nicotianamine (Takahashi et al., 2003), which play a variety of important roles for plant growth and development (Huang et al., 2012; Sauter et al., 2013). The SAM present in organelles, like the chloroplast, appears to be imported from the cytosol by specific SAM/S-adenosylhomocysteine exchange transporters that reside on the envelope membranes of plastids (Ravanel et al., 2004; Bouvier et al., 2006). The imported SAM is involved in the biogenesis of Asp-derived amino acids (Curien et al., 1998; Jander and Joshi, 2009; Sauter et al., 2013) and serves as the methyl donor for the methylation of macromolecules, such as plastid DNA (Nishiyama et al., 2002; Ahlert et al., 2009) and proteins (Houtz et al., 1989; Niemi et al., 1990; Ying et al., 1999; Trievel et al., 2003; Alban et al., 2014), and small molecule metabolites, such as prenylipids (e.g. plastoquinone, tocopherol, chlorophylls, and phylloquinone; Bouvier et al., 2005, 2006; DellaPenna, 2005).Although plants and microbes are the natural sources for useful terpenes, most of them are produced in very small amounts and often as complex mixtures. In contrast, B. braunii produces large quantities of triterpenes, but its slow growth makes it undesirable as a viable production platform (Niehaus et al., 2011). Nevertheless, metabolic engineering and synthetic biology offer many strategies to manipulate terpene metabolism in various biological systems to achieve high-value terpene production with high yield and high fidelity for particular practical applications (Nielsen and Keasling, 2011). Many successes have been achieved in engineering valuable terpenes in heterotrophic microbes, such as Escherichia coli (Nishiyama et al., 2002; Martin et al., 2003; Ajikumar et al., 2010) and Saccharomyces cerevisiae (Ro et al., 2006; Takahashi et al., 2007; Westfall et al., 2012; Zhuang and Chappell, 2015). The strategies developed in these efforts usually take advantage of specific microbe strains whose innate biosynthetic machinery is genetically modified to accumulate certain prenyldiphosphate precursors (e.g. IPP or FPP), which can be utilized by other introduced terpene synthase(s) for the production of the desired terpene(s). For example, greater than 900 mg L⁻¹ bisabolene was produced when bisabolene synthase genes from plants were introduced into FPP-overproducing E. coli or S. cerevisiae strains (Peralta-Yahya et al., 2011). High levels of farnesane production for diesel fuels were also achieved by reductive hydrogenation of its precursor farnesene, which was generated from a genetically engineered yeast (e.g. Saccharomyces cerevisiae) strain using plant farnesene synthases (Renninger and McPhee, 2008; Ubersax and Platt, 2010). However, terpene production using microbial platforms is still dependent on exogenous feedstocks (i.e. sugars) and elaborate production facilities, both of which add significantly to their production costs.Compared with microbial systems, engineering terpene production in plant systems seems like an attractive target as well. This is because plants can take advantage of photosynthesis by using atmospheric CO₂ as their carbon resource instead of relying on exogenous carbon feedstocks. Moreover, crop plants such as tobacco (Nicotiana tabacum) can generate a large amount of green tissues efficiently when grown for biomass production (Schillberg et al., 2003; Andrianov et al., 2010), making them a robust, sustainable, and scalable platform for large-scale terpene production. Nonetheless, compared with microbial platforms, there are only a few examples of elevating terpene production in bioengineered plants. This is due partly to higher plants being complex multicellular organisms, in which terpene metabolism generally utilizes more complex innate machinery that can be compartmentalized intracellularly and to cell/tissue specificities (Lange and Ahkami, 2013; Kempinski et al., 2015). Significant efforts have been made to overcome these obstacles to improve the production of valuable terpenes in plants, including monoterpenes (Lücker et al., 2004; Ohara et al., 2010; Lange et al., 2011), sesquiterpenes (Aharoni et al., 2003; Kappers et al., 2005; Wu et al., 2006; Davidovich-Rikanati et al., 2008), diterpenes (Besumbes et al., 2004; Anterola et al., 2009), and triterpenes (Inagaki et al., 2011; Wu et al., 2012). Among these, engineering terpene metabolism into a subcellular organelle, where the engineered enzymes/pathways can utilize unlimited/unregulated precursors as substrates, appears most successful. For example, Wu et al. (2006, 2012) expressed an avian farnesyl diphosphate synthase (FPS) with foreign sesquiterpene/triterpene synthases targeted to the plastid to divert the IPP/DMAPP pool from the plastidic MEP pathway to synthesize high levels of the novel sesquiterpenes patchoulol and amorpha-4,11-diene up to 30 µg g⁻¹ fresh weight and the triterpene squalene up to 1,000 µg g⁻¹ fresh weight. This strategy appears to be particularly robust because it avoids possible endogenous regulation of sesquiterpene and triterpene biosynthesis, which occurs normally in the cytoplasm, and relies upon more plastic precursor pools of IPP/DMAPP inherent in the plastid, which are primarily derived from the local CO₂ fixation (Wright et al., 2014).The goal of this study was to evaluate the prospects for engineering advanced features of triterpene metabolism from B. braunii into tobacco and, thus, to probe the innate intricacies of isoprenoid metabolism in plants. In order to achieve this, we first introduced the key steps of botryococcene biosynthesis into specific subcellular compartments of tobacco cells under the direction of constitutive or trichome-specific promoters. The transgenic lines expressing the enzymes in the chloroplast were found to accumulate the highest levels of botryococcene. Triterpene methyltransferases were next introduced into the same intracellular compartments of selected high-triterpene-accumulating lines. A high yield of methylated triterpenes was also achieved in transgenic lines when the TMTs were targeted to the chloroplast. Through careful comparison of the levels of triterpenes and the methylated triterpene products in the various transgenic lines, we have also gained a deeper insight into the subcellular distribution of the triterpene products in these transgenic lines as well as a better understanding of methylation metabolism for specialized metabolites in particular compartments. These findings all contribute to our understanding of the regulatory elements that control carbon flux through the innate terpene biosynthetic pathways operating in plants.     

The Endoplasmic Reticulum Is a Reservoir for WAVE/SCAR Regulatory Complex Signaling in the Arabidopsis Leaf

During plant cell morphogenesis, signal transduction and cytoskeletal dynamics interact to locally organize the cytoplasm and define the geometry of cell expansion. The WAVE/SCAR (for WASP family verprolin homologous/suppressor of cyclic AMP receptor) regulatory complex (W/SRC) is an evolutionarily conserved heteromeric protein complex. Within the plant kingdom W/SRC is a broadly used effector that converts Rho-of-Plants (ROP)/Rac small GTPase signals into Actin-Related Protein2/3 and actin-dependent growth responses. Although the components and biochemistry of the W/SRC pathway are well understood, a basic understanding of how cells partition W/SRC into active and inactive pools is lacking. In this paper, we report that the endoplasmic reticulum (ER) is an important organelle for W/SRC regulation. We determined that a large intracellular pool of the core W/SRC subunit NAP1, like the known positive regulator of W/SRC, the DOCK family guanine nucleotide-exchange factor SPIKE1 (SPK1), localizes to the surface of the ER. The ER-associated NAP1 is inactive because it displays little colocalization with the actin network, and ER localization requires neither activating signals from SPK1 nor a physical association with its W/SRC-binding partner, SRA1. Our results indicate that in Arabidopsis (Arabidopsis thaliana) leaf pavement cells and trichomes, the ER is a reservoir for W/SRC signaling and may have a key role in the early steps of W/SRC assembly and/or activation.The W/SRC (for WASP family verprolin homologous/suppressor of cAMP receptor regulatory complex) and Actin-Related Protein (ARP)2/3 complex are part of an evolutionarily conserved Rho-of-Plants (ROP)/Rac small GTPase signal transduction cascade that controls actin-dependent morphogenesis in a wide variety of tissues and developmental contexts (Smith and Oppenheimer, 2005; Szymanski, 2005; Yalovsky et al., 2008). Many of the components and regulatory relationships among the complexes were discovered based on the stage-specific cell-swelling and -twisting phenotypes of the distorted class of Arabidopsis (Arabidopsis thaliana) trichome mutants (Szymanski et al., 1999; Zhang et al., 2005, 2008; Djakovic et al., 2006; Le et al., 2006; Uhrig et al., 2007). However, in both maize (Zea mays) and Arabidopsis, W/SRC and/or ARP2/3 are required for normal pavement cell morphogenesis (Frank and Smith, 2002; Mathur et al., 2003b; Brembu et al., 2004). Compared with other Arabidopsis pavement cell mutants, the shape defects of the distorted group are relatively mild. However, the distorted mutants and spike1 (spk1) differ from most other morphology mutants in that they display gaps in the shoot epidermis, most frequently at the interface of pavement cells and stomata (Qiu et al., 2002; Le et al., 2003; Li et al., 2003; Mathur et al., 2003b; Zhang et al., 2005; Djakovic et al., 2006). The cell gaps may reflect either uncoordinated growth between neighboring cells or defective cortical actin-dependent secretion of polysaccharides and/or proteins that promote cell-cell adhesion (Smith and Oppenheimer, 2005; Hussey et al., 2006; Leucci et al., 2007).In tip-growing cells, there is a strict requirement for actin to organize the trafficking and secretion activities of the cell to restrict growth to the apex. In Arabidopsis, the W/SRC-ARP2/3 pathway is not an essential tip growth component, because null alleles of both W/SRC and ARP2/3 subunits do not cause noticeable pollen tube or root hair phenotypes (Le et al., 2003; Djakovic et al., 2006). However, reverse genetic analysis of the W/SRC subunit BRK1 and ARP2/3 in the tip-growing protonemal cells of Physcomitrella patens revealed the obvious importance of this pathway (Harries et al., 2005; Perroud and Quatrano, 2008). Along similar lines, in two different legume species, W/SRC subunits are required for a normal root nodulation response to symbiotic bacteria (Yokota et al., 2009; Miyahara et al., 2010), indicating a conditional importance for this pathway in root hair growth. These genetic studies centered on the W/SRC and ARP2/3 pathways, in addition to those that involve a broader collection of actin-based morphology mutants (Smith and Oppenheimer, 2005; Blanchoin et al., 2010), are defining important cytoskeletal proteins and new interactions with the endomembrane system during morphogenesis. However, it is not completely clear how unstable actin filaments and actin bundle networks dictate the growth patterns of cells (Staiger et al., 2009).The difficulty of understanding the functions of specific actin arrays can be explained, in part, by the fact that plant cells that employ a diffuse growth mechanism have highly unstable cortical actin filaments and large actin bundles that do not have a geometry that obviously relates to the direction of growth or a specific subcellular activity (Blanchoin et al., 2010). This is in contrast to the cortical endocytic actin patches in yeast (Saccharomyces cerevisiae; Evangelista et al., 2002; Kaksonen et al., 2003) and cortical meshworks in the lamellipodia of crawling cells (Pollard and Borisy, 2003) that reveal subcellular locations where actin works to locally control membrane dynamics. In thick-walled plant cells, the magnitude of the forces that accompany turgor-driven cell expansion exceed those that could be generated by actin polymerization by orders of magnitude (Szymanski and Cosgrove, 2009). Localized cell wall loosening or the assembly of an anisotropic cell wall generates asymmetric yielding responses to turgor-induced stress (Baskin, 2005; Cosgrove, 2005). Therefore, the actin-based control of cell boundary dynamics is indirect, and the actin cytoskeleton influences cell shape change, in part, by actin and/or myosin-dependent trafficking of hormone transporters (Geldner et al., 2001) and organelles (Prokhnevsky et al., 2008), including those that control the localized delivery of protein complexes and polysaccharides that pattern the cell wall (Leucci et al., 2007; Gutierrez et al., 2009). In this scheme for actin-based growth control, the actin network dynamically rearranges at spatial scales that span from approximately 1- to 10-µm subcellular domains that may locally position organelles (Cleary, 1995; Gibbon et al., 1999; Szymanski et al., 1999) to the more than 100-µm actin bundle networks that operate at the spatial scales of entire cells (Gutierrez et al., 2009; Dyachok et al., 2011). It is clear from the work of several laboratories that the W/SRC and ARP2/3 protein complexes are required to organize cortical actin and actin bundle networks in trichomes (Szymanski et al., 1999; Le et al., 2003; Deeks et al., 2004; Zhang et al., 2005) and cylindrical epidermal cells (Mathur et al., 2003b; Dyachok et al., 2008, 2011). A key challenge now is to understand how plant cells deploy these approximately 10- to 20-nm heteromeric protein complexes to influence the patterns of growth at cellular scales.The genetic and biochemical control of ARP2/3 is complicated, but this is a tractable problem in plants, because the pathway is relatively simple compared with most other species in which it has been characterized. For example, in organisms ranging from yeast to humans, there are multiple types of ARP2/3 activators, protein complexes, and pathways that activate ARP2/3 (Welch and Mullins, 2002; Derivery and Gautreau, 2010). However, the maize and Arabidopsis genomes encode only WAVE/SCAR homologous proteins that can potently activate ARP2/3 (Frank et al., 2004; Basu et al., 2005). Detailed genetic and biochemical analyses of the WAVE/SCAR gene family in Arabidopsis demonstrated that the plant activators function interchangeably within the context of the W/SRC and define the lone pathway for ARP2/3 activation (Zhang et al., 2008). Bioinformatic analyses are consistent with a prominent role for W/SRC in the angiosperms, because in general, WASH complex subunits, which are structurally similar to WAVE/SCAR proteins, are largely absent from the higher plant genomes, while WAVE/SCAR genes are highly conserved (Kollmar et al., 2012).The components and regulatory schemes of the W/SRC-ARP2/3 pathway in Arabidopsis and P. patens are conserved compared with vertebrate species that employ these same protein complexes (Szymanski, 2005). For example, mutant complementation tests indicate that human W/SRC and ARP2/3 complex subunits can substitute for the Arabidopsis proteins (Mathur et al., 2003b). Furthermore, biochemical assays of Arabidopsis W/SRC (Basu et al., 2004; El-Assal et al., 2004; Frank et al., 2004; Le et al., 2006; Uhrig et al., 2007) and ARP2/3 assembly (Kotchoni et al., 2009) have shown that the binary interactions among W/SRC subunits and ARP2/3 complex assembly mechanisms are indistinguishable from those that have been observed for human W/SRC (Gautreau et al., 2004) and yeast ARP2/3 (Winter et al., 1999). After an initial period of controversy concerning the biochemical control of W/SRC, it is now apparent that vertebrate W/SRC (Derivery et al., 2009; Ismail et al., 2009), like the ARP2/3 complex (Machesky et al., 1999), is intrinsically inactive and requires positive regulation by Rac and other factors to fully activate ARP2/3 (Ismail et al., 2009; Lebensohn and Kirschner, 2009; Chen et al., 2010). Although overexpression of dominant negative ROP mutants causes trichome swelling and a reduced trichome branch number (Fu et al., 2002), the involvement of ROPs in trichome morphogenesis has been difficult to prove with a loss-of-function ROP allele because so many ROPs are expressed in this cell type (Marks et al., 2009). Existing reports on ROP loss-of-function mutants demonstrate the importance of pavement cell morphogenesis but do not document a trichome phenotype (Fu et al., 2005; Xu et al., 2010). A recent report describes a clever strategy to generate ROP loss-of-function lines that used the ectopic expression of ROP-specific bacterial toxins. There was a strong association between inducible expression of the toxins and the appearance of trichomes with severe trichome swelling and reduced branch number phenotypes (Singh et al., 2012). Although the exact mechanism of ROP-dependent control of W/SRC remains to be determined, the results described above in combination with the detection of direct interactions between the ROPGEF SPK1, active forms of ROP, and W/SRC subunits (Basu et al., 2004, 2008; Uhrig et al., 2007) strongly suggest that W/SRC is a ROP effector complex.The major challenge in the field now is to better understand the cellular control of W/SRC and how the complex is partitioned into active and inactive pools. In mammalian cells that crawl on a solid substrate, current models propose that a cytosolic pool of inactive WAVE/SCAR proteins and W/SRC is locally recruited and activated at specific plasma membrane surfaces in response to signals from some unknown Rac guanine nucleotide-exchange factor (GEF), protein kinase, and/or lipid kinase (Oikawa et al., 2004; Lebensohn and Kirschner, 2009; Chen et al., 2010). However, in Drosophila melanogaster neurons (Bogdan and Klämbt, 2003) and cultured human melanoma cells (Steffen et al., 2004), there are large pools of W/SRC with a perinuclear or organelle-like punctate localization that has no obvious relationship to cell shape or motility, raising uncertainty about the cellular mechanisms of W/SRC activation and the importance of different subcellular pools of the complex.In plants, cell fractionation experiments indicate that SCAR1 and ARP2/3 have an increased association with membranes compared with their animal counterparts (Dyachok et al., 2008; Kotchoni et al., 2009). In tip-growing moss protonemal cells, both the W/SRC subunit BRK1 and ARP2/3 localize to a population of unidentified organelles within the apical zone (Perroud and Quatrano, 2008). Similar live-cell imaging experiments in Arabidopsis reported a plasma membrane localization for SCAR1 and BRK1 in a variety of shoot epidermal and root cortex, and their accumulation at young trichome branch tips and at three-way cell wall junctions may define subcellular domains for W/SRC-ARP2/3-dependent actin filament nucleation at the plasma membrane (Dyachok et al., 2008). However, to our knowledge, active W/SRC, defined here as the fraction of W/SRC that colocalizes with ARP2/3 or actin, has not been reported in plants, and the plasma membrane is not necessarily the only organelle involved in W/SRC regulation. For example, the reported accumulation of BRK1 and SCAR1 at three-way cell wall junctions has a punctate appearance at the cell cortex that may not simply correspond to the plasma membrane (Dyachok et al., 2008). Also, in young stage 4 trichomes, there was an uncharacterized pool of intracellular SCAR1, but not BRK1, that localized to relatively large punctate structures (Dyachok et al., 2008). The endoplasmic reticulum (ER) may also be involved in W/SRC regulation. The ER-localized DOCK family ROPGEF SPK1 (Zhang et al., 2010) physically associates with multiple W/SRC proteins (Uhrig et al., 2007; Basu et al., 2008) and, based on genetic criteria, is an upstream, positive regulator of the W/SRC-ARP2/3 pathway (Basu et al., 2008). In the leaf, one function of SPK1 is to promote normal trafficking between the ER and Golgi; however, arp2/3 mutants do not share ER-stress phenotypes with spk1 (Zhang et al., 2010), making it unclear if SPK1 and the ER are directly involved in W/SRC signaling.This paper focuses on the localization and control of the W/SRC subunit NAP1/GNARLED/NAPP/HEM1/2. Arabidopsis NAP1 directly interacts with the ROP/Rac effector subunit SRA1/PIROGI/KLUNKER/PIRP (Basu et al., 2004; El-Assal et al., 2004; Uhrig et al., 2007). Based on the equally severe syndrome of nap1 and arp2/3 null phenotypes, and double mutant analyses, the only known function of NAP1 is to positively regulate ARP2/3 (Brembu et al., 2004; Deeks et al., 2004; El-Din El-Assal et al., 2004; Li et al., 2004). The vertebrate SRA1-NAP1 dimer is important for W/SRC assembly (Gautreau et al., 2004) and forms an extended physical surface that trans-inhibits the C-terminal ARP2/3-activating domain of WAVE/SCAR (Chen et al., 2010). The plant NAP1 and SRA1 proteins share end-to-end amino acid conservation with their vertebrate homologs and may form a heterodimer with similar functions (Basu et al., 2004; El-Assal et al., 2004; Uhrig et al., 2007). We report here that Arabidopsis NAP1 is strongly associated with ER membranes. In a detailed series of localization experiments, we detect a complicated intracellular distribution of NAP1 among the ER, the nucleus, and unidentified submicrometer punctae. A large pool of ER-associated NAP1 is inactive, based on the low level of colocalization with actin.Its accumulation on the ER does not require activating signals from either SPK1 or SRA1. These data indicate that the ER is a reservoir for W/SRC signaling and suggest that early steps in the positive regulation of NAP1 and the W/SRC occur on the ER surface.