Origin of β-Carotene-Rich Plastoglobuli in Dunaliella bardawil

The halotolerant microalgae Dunaliella bardawil accumulates under nitrogen deprivation two types of lipid droplets: plastoglobuli rich in β-carotene (βC-plastoglobuli) and cytoplasmatic lipid droplets (CLDs). We describe the isolation, composition, and origin of these lipid droplets. Plastoglobuli contain β-carotene, phytoene, and galactolipids missing in CLDs. The two preparations contain different lipid-associated proteins: major lipid droplet protein in CLD and the Prorich carotene globule protein in βC-plastoglobuli. The compositions of triglyceride (TAG) molecular species, total fatty acids, and sn-1+3 and sn-2 positions in the two lipid pools are similar, except for a small increase in palmitic acid in plastoglobuli, suggesting a common origin. The formation of CLD TAG precedes that of βC-plastoglobuli, reaching a maximum after 48 h of nitrogen deprivation and then decreasing. Palmitic acid incorporation kinetics indicated that, at early stages of nitrogen deprivation, CLD TAG is synthesized mostly from newly formed fatty acids, whereas in βC-plastoglobuli, a large part of TAG is produced from fatty acids of preformed membrane lipids. Electron microscopic analyses revealed that CLDs adhere to chloroplast envelope membranes concomitant with appearance of small βC-plastoglobuli within the chloroplast. Based on these results, we propose that CLDs in D. bardawil are produced in the endoplasmatic reticulum, whereas βC-plastoglobuli are made, in part, from hydrolysis of chloroplast membrane lipids and in part, by a continual transfer of TAG or fatty acids derived from CLD.Eukaryotic cells accumulate neutral lipids in different tissues mainly in the form of lipid droplets (Murphy, 2012). Most lipid droplets consist of a core of triglycerides (TAGs) and/or sterol esters coated by a phospholipids monolayer and embedded with proteins (Zweytick et al., 2000). Plants accumulate TAGs in different tissues, primarily in seeds but also in fruit, such as palm oil, flowers, and leaves. The best characterized system for TAG metabolism is oil seeds, in which TAG serves as the major carbon and energy reservoir to be used during germination (Huang, 1992, 1996). Recent studies show that lipid droplets are not just static pools of lipids but have diverse metabolic functions (Farese and Walther, 2009). In addition, plants also contain plastoglobuli, small chloroplastic lipid droplets consisting primarily of storage lipids and pigments. Proteome analyses of plastoglobuli suggest that they are involved in synthesis and degradation of lipids, pigments, and coenzymes (Ytterberg et al., 2006; Lundquist et al., 2012). It has been shown that plant plastoglobuli are associated with thylakoid membranes (Austin et al., 2006; Ytterberg et al., 2006).It is not entirely clear where the TAGs are synthesized in the plant cell. Until recently, it has been assumed that most TAGs are made in the endoplasmatic reticulum (ER) from fatty acids, which are mostly synthesized in the chloroplast and imported to the cytoplasm (Joyard et al., 2010). However, the recent identification of the enzyme diacylglycerol acyl transferase in plant plastoglobuli (Lundquist et al., 2012) suggests that TAG may be synthesized directly in chloroplasts, although direct evidence is missing. TAG may be synthesized also from galactolipid fatty acids during stress or senescence by phytyl ester synthases, which catalyze acyl transesterification from galactolipids to TAGs (Lippold et al., 2012). Phosphatidyl choline (PC) plays a major role in acyl transfer of newly synthesized fatty acids from the chloroplast into TAGs at the ER in plants (Bates et al., 2009). An indication for the origin of glycerolipids in plants is the identity of the fatty acids at the sn-2 position: if it originates in the chloroplast, it is mostly C16:0, whereas if it was made in the ER, it is mostly C:18 (Heinz and Roughan, 1983).Many species of unicellular microalgae can accumulate large amounts of TAGs under growth-limiting conditions, such as nitrogen deprivation (Shifrin and Chisholm, 1981; Roessler, 1990; Avron and Ben-Amotz, 1992; Thompson, 1996). In green microalgae (Chlorophyceae), TAGs are usually synthesized and accumulated in cytoplasmatic lipid droplets (CLDs; Murphy, 2012), although in some cases, such as in Chlamydomonas reinhardtii starchless mutants, they also accumulate in chloroplasts (Fan et al., 2011; Goodson et al., 2011). Recent studies indicate that the CLDs are closely associated with ER membranes and possibly, chloroplast envelope membranes as well (Goodson et al., 2011; Peled et al., 2012).Green microalgae also contain two distinct types of chloroplastic lipid droplets. The first type is plastoglobuli, similar in morphology to higher plants plastoglobuli (Bréhélin et al., 2007; Kessler and Vidi, 2007). The second type is the eyespot (stigma), part of the visual system in microalgae. The eyespot is composed of a cluster of β-carotene-containing lipid droplets organized in several layers between grana membranes in the chloroplast (Häder and Lebert, 2009; Kreimer, 2009). Recent proteomic analysis of algal eyespot proteins revealed that they contain diverse structural proteins, lipid and carotenoid metabolizing enzymes, transporters, and signal transduction components (Schmidt et al., 2006).The origin of TAG in microalgae is still not clear. In C. reinhardtii, it was found that the major fatty acids in the sn-2 position are 16:0, which according to the plant dogma, is made in the chloroplast (Fan et al., 2011). In C. reinhardtii, which lacks PC, monogalactosyldiacylglycerol (MGDG) was proposed to replace PC in the mobilization of fatty acids from plastidal galactoglycerolipids into TAG based on mutation of a galactoglycerolipid lipase (Li et al., 2012). Based on these results and others, it has been proposed that, in C. reinhardtii, triglycerides are primarily produced in the chloroplast or combined with ER (Li et al., 2012; Liu and Benning, 2013).Plants and algae lipid droplets contain structural major proteins localized at the lipid droplet periphery, and their major function seems to be stabilization and prevention of fusion (Huang, 1992, 1996; Katz et al., 1995; Frandsen et al., 2001; Liu et al., 2009). In plant seed oils, the major classes of lipid droplet proteins are oleosins and caleosins, which have a characteristic hydrophobic loop with a conserved three Pro domain (Hsieh and Huang, 2004; Capuano et al., 2007; Purkrtova et al., 2008; Tzen, 2012). Oleosin and caleosin analogs were also recently identified in some green microalgal species (Lin et al., 2012; Vieler et al., 2012; Huang et al., 2013). However, the most abundant lipid droplets proteins in green algae (Chloropyceae) are a new family of major lipid droplet proteins (MLDPs) structurally distinct from plant oleosins and caleosins (Moellering and Benning, 2010; Peled et al., 2011; Davidi et al., 2012). Plastoglobules have different major lipid-associated proteins termed plastoglobules-associated protein-fibrillins, which form a distinct protein family with no sequence or structural similarities to oleosins (Kim and Huang, 2003). We have previously identified in the plastoglobuli rich in β-carotene (βC-plastoglobuli) a lipid-associated protein termed carotene globule protein (CGP), whose degradation destabilized the lipid droplets (Katz et al., 1995). The proteome of C. reinhardtii lipid droplet indicates that algal CLDs also contain several enzymes, suggesting that they are involved in lipid metabolism (Nguyen et al., 2011).The halotolerant green algae Dunaliella bardawil and Dunaliella salina ‘Teodoresco’ are unique in that they accumulate under high light stress or nitrogen deprivation large amounts of plastidic lipid droplets (βC-plastoglobuli), which consist of TAG and two isomers of β-carotene, all trans and 9-cis (Ben-Amotz et al., 1982, 1988). D. bardawil also accumulates CLD under the same stress conditions, similar to other green algae (Davidi et al., 2012). It has been shown that the function of βC-plastoglobuli is to protect the photosynthetic system against photoinhibition (Ben-Amotz et al., 1989). The enzymatic pathway for β-carotene synthesis in D. bardawil and D. salina has been partly identified, but the subcellular localization of β-carotene biosynthesis is not known (Jin and Polle, 2009). The synthesis of β-carotene depends on TAG biosynthesis (Rabbani et al., 1998); however, the origin of βC-plastoglobuli is not known. Are they formed within the chloroplast, or are they made in the cytoplasm? Is the TAG in βC-plastoglobuli and CLD identical or different, and where is it formed?D. bardawil is an excellent model organism for isolation of lipid droplet for several reasons. First, D. bardawil contains large amounts of both CLD and βC-plastoglobuli (Ben-Amotz et al., 1982; Fried et al., 1982), making it possible to obtain sufficient amounts of proteins and lipids from the two types of lipid pools for detailed analyses. Second, Dunaliella do not have a rigid cell wall and can be lysed by a gentle osmotic shock, which does not rupture the chloroplast. Therefore, it is possible to sequentially release pure CLD and βC-plastoglobuli by a two-step lysis (Katz et al., 1995). Third, D. bardawil seems to lack the eyespot structure, which can be clearly observed in other Dunaliella spp. even in a light microscope or by electron microscopy, but has never been observed in D. bardawil by us. It avoids the risk of cross contamination of βC-plastoglobuli with eyespot proteins. Fourth, the availability of protein markers for the major lipid droplet-associated proteins, CGPs and MLDPs, enabled both good immunolocalization and careful monitoring of the purity of the preparations by western analysis.In this work, we describe the purification, lipid compositions, and protein profiles of two lipid pools from D. bardawil: CLD and plastidic βC-plastoglobuli. A detailed proteomic analysis of these lipid droplets will be described in another work. Combined with detailed electron microscopy studies, these results led to surprising conclusions regarding the origin of the plastidic βC-plastoglobuli.     

Proteome Analysis of Cytoplasmatic and Plastidic β-Carotene Lipid Droplets in Dunaliella bardawil

The halotolerant green alga Dunaliella bardawil is unique in that it accumulates under stress two types of lipid droplets: cytoplasmatic lipid droplets (CLD) and β-carotene-rich (βC) plastoglobuli. Recently, we isolated and analyzed the lipid and pigment compositions of these lipid droplets. Here, we describe their proteome analysis. A contamination filter and an enrichment filter were utilized to define core proteins. A proteome database of Dunaliella salina/D. bardawil was constructed to aid the identification of lipid droplet proteins. A total of 124 and 42 core proteins were identified in βC-plastoglobuli and CLD, respectively, with only eight common proteins. Dunaliella spp. CLD resemble cytoplasmic droplets from Chlamydomonas reinhardtii and contain major lipid droplet-associated protein and enzymes involved in lipid and sterol metabolism. The βC-plastoglobuli proteome resembles the C. reinhardtii eyespot and Arabidopsis (Arabidopsis thaliana) plastoglobule proteomes and contains carotene-globule-associated protein, plastid-lipid-associated protein-fibrillins, SOUL heme-binding proteins, phytyl ester synthases, β-carotene biosynthesis enzymes, and proteins involved in membrane remodeling/lipid droplet biogenesis: VESICLE-INDUCING PLASTID PROTEIN1, synaptotagmin, and the eyespot assembly proteins EYE3 and SOUL3. Based on these and previous results, we propose models for the biogenesis of βC-plastoglobuli and the biosynthesis of β-carotene within βC-plastoglobuli and hypothesize that βC-plastoglobuli evolved from eyespot lipid droplets.Lipid droplets are the least characterized organelles in both mammalian and plant cells, and they were considered until a few years ago as passive storage compartments for triglycerides (TAG), sterol esters, and some pigments. However, recent studies have shown that they have diverse metabolic functions (Goodman, 2008; Farese and Walther, 2009; Murphy, 2012). Proteomic analyses in plants and some microalgae have shown that lipid droplets in the cytoplasm and in the chloroplast contain a large diversity of proteins including both structural proteins and many enzymes, indicating that they take an active metabolic role in the synthesis, degradation, and mobilization of glycerolipids, sterols, and pigments as well as in regulatory functions that have not yet been clarified (Schmidt et al., 2006; Ytterberg et al., 2006; Nguyen et al., 2011; Lundquist et al., 2012b; Eugeni Piller et al., 2014). A major limitation for determining the proteomes of lipid droplets, particularly in microalgae, is the purity and the homogeneity of the preparation. Green microalgae, for example, may contain three distinct pools of lipid droplets in one cell: the cytoplasmatic lipid droplets (CLD), the major neutral lipid pool, which are induced under stress conditions such as nitrogen limitation or at the stationary growth phase (Wang et al., 2009); plastoglobules, which are smaller lipid droplets within the chloroplast that have been shown to change in size and number under stress conditions and seem to be involved in stress resistance, metabolite transport, and the regulation of photosynthetic electron transport (Bréhélin et al., 2007; Besagni and Kessler, 2013); and the eyespot structure, part of the visual system in green algae, composed of one or several layers of lipid droplets, characterized by their orange color resulting from a high content of β-carotene (Kreimer, 2009). Disruption of microalgal cells, which is required for the isolation of the lipid droplets, usually involves harsh treatments such as sonication, mixing with glass beads, or use of a French press that breaks not only the cell membrane but also the chloroplast. Therefore, it is almost impossible to separate the different lipid droplet classes by the subsequent density gradient centrifugation, making it difficult to assign the origin of identified proteins. The other major difficulty is contamination by proteins released during cell lysis and fractionation, which associate and copurify with lipid droplets. These include cytoplasmic, chloroplastic, and mitochondrial proteins (Moellering and Benning, 2010; James et al., 2011; Nguyen et al., 2011; Nojima et al., 2013). Purification of isolated lipid droplets from loosely associated proteins is possible by treatments with detergents, high salt, and chaotropic agents (Jolivet et al., 2004; Nguyen et al., 2011); however, the danger in such treatments is that they also remove native loosely associated proteins from the lipid droplets.In this work, we tried to circumvent these problems by choosing a special algal species that is suitable for controlled cell lysis and fractionation and by utilizing two different contamination filters.The alga we selected, Dunaliella bardawil, is unique in that it accumulates large amounts of two different types of lipid droplets, CLD and β-carotene-rich (βC) plastoglobuli, under stress conditions (Davidi et al., 2014). The lack of a rigid cell wall in this alga allows lysis of the plasma membrane by a gentle osmotic shock, releasing CLD but leaving the chloroplast intact (Katz et al., 1995). This enables the recovery of large quantities of the two types of highly purified lipid droplets by differential lysis. In a recent study, we described the isolation and lipid compositions of these two lipid pools and showed that they have similar TAG compositions but different lipid-associated major proteins (Davidi et al., 2014).The high nutritional and pharmacological value of β-carotene for humans has promoted intensive research aimed to clarify its biosynthesis and regulation in plants and also led to attempts to increase β-carotene levels by genetic manipulations in crop plants such as tomato (Solanum lycopersicum; Rosati et al., 2000; Giorio et al., 2007) or by the creation of Golden rice (Oryza sativa; Ye et al., 2000). However, the capacity of plants to store β-carotene is limited, and in this respect, D. bardawil is an exceptional example of an organism that can accumulate large amounts of this pigment, up to 10% of its dry weight. This is enabled by the compartmentation and storage of this lipophilic pigment in specialized plastoglobules. Also, the unusual isomeric composition, consisting of around 50% 9-cis- and 50% all-trans-isomers (Ben-Amotz et al., 1982, 1988), is probably of major importance in this respect, due to the better solubility of the cis-isomer in lipids, which enables the storage of high concentrations exceeding 50% of the lipid droplets. The localization of carotenoid biosynthesis in plants appears to be tissue specific: in green tissues, it takes place in chloroplast membranes, probably within the inner chloroplast envelope membrane (Joyard et al., 2009), whereas in carotenoid-accumulating fruits, such as tomato or bell pepper (Capsicum annuum), it takes place in specialized organelles derived from chromoplasts (Siddique et al., 2006; Barsan et al., 2010). In green microalgae, there are at least two types of carotenoid-accumulating organelles: CLD and eyespot. Algae such as Haematococcus pluvialis and Chlorella zofigiensis accumulate carotenoids within CLD. In H. pluvialis, the major pigment, astaxanthin, is synthesized initially in the chloroplast as β-carotene and then transferred to CLD, where it is oxidized and hydroxylated to astaxanthin (Grünewald et al., 2001). The eyespot, which is composed of one or several layers of small β-carotene-containing lipid droplets, has been shown by proteomic analysis to include part of the β-carotene biosynthesis enzymes, indicating that β-carotene is probably synthesized within these lipid droplets (Schmidt et al., 2006). Similarly, plant chromoplasts also contain carotenoid biosynthesis enzymes (Schmidt et al., 2006; Ytterberg et al., 2006; Schapire et al., 2009). D. bardawil and Dunaliella salina are unique in that they accumulate large amounts of β-carotene within βC-plastoglobuli. A special focus in this work was the identification of the β-carotene biosynthesis machinery in D. bardawil. It is not known if the synthesis takes place inside the lipid βC-plastoglobuli or in chloroplast envelope membranes. Since D. bardawil also contains β-carotene and xanthophylls at the photosynthetic system, it is interesting to know whether the β-carotene that accumulates under stress in βC-plastoglobuli is produced by the constitutive carotenoid biosynthetic pathway or by a different stress-induced enzymatic system.     

The Arabidopsis Endosomal Sorting Complex Required for Transport III Regulates Internal Vesicle Formation of the Prevacuolar Compartment and Is Required for Plant Development

We have established an efficient transient expression system with several vacuolar reporters to study the roles of endosomal sorting complex required for transport (ESCRT)-III subunits in regulating the formation of intraluminal vesicles of prevacuolar compartments (PVCs)/multivesicular bodies (MVBs) in plant cells. By measuring the distributions of reporters on/within the membrane of PVC/MVB or tonoplast, we have identified dominant negative mutants of ESCRT-III subunits that affect membrane protein degradation from both secretory and endocytic pathways. In addition, induced expression of these mutants resulted in reduction in luminal vesicles of PVC/MVB, along with increased detection of membrane-attaching vesicles inside the PVC/MVB. Transgenic Arabidopsis (Arabidopsis thaliana) plants with induced expression of ESCRT-III dominant negative mutants also displayed severe cotyledon developmental defects with reduced cell size, loss of the central vacuole, and abnormal chloroplast development in mesophyll cells, pointing out an essential role of the ESCRT-III complex in postembryonic development in plants. Finally, membrane dissociation of ESCRT-III components is important for their biological functions and is regulated by direct interaction among Vacuolar Protein Sorting-Associated Protein20-1 (VPS20.1), Sucrose Nonfermenting7-1, VPS2.1, and the adenosine triphosphatase VPS4/SUPPRESSOR OF K⁺ TRANSPORT GROWTH DEFECT1.Endomembrane trafficking in plant cells is complicated such that secretory, endocytic, and recycling pathways are usually integrated with each other at the post-Golgi compartments, among which, the trans-Golgi network (TGN) and prevacuolar compartment (PVC)/multivesicular body (MVB) are best studied (Tse et al., 2004; Lam et al., 2007a, 2007b; Müller et al., 2007; Foresti and Denecke, 2008; Hwang, 2008; Otegui and Spitzer, 2008; Robinson et al., 2008; Richter et al., 2009; Ding et al., 2012; Gao et al., 2014). Following the endocytic trafficking of a lipophilic dye, FM4-64, the TGN and PVC/MVB are sequentially labeled and thus are defined as the early and late endosome, respectively, in plant cells (Lam et al., 2007a; Chow et al., 2008). While the TGN is a tubular vesicular-like structure that may include several different microdomains and fit its biological function as a sorting station (Chow et al., 2008; Kang et al., 2011), the PVC/MVB is 200 to 500 nm in size with multiple luminal vesicles of approximately 40 nm (Tse et al., 2004). Membrane cargoes destined for degradation are sequestered into these tiny luminal vesicles and delivered to the lumen of the lytic vacuole (LV) via direct fusion between the PVC/MVB and the LV (Spitzer et al., 2009; Viotti et al., 2010; Cai et al., 2012). Therefore, the PVC/MVB functions between the TGN and LV as an intermediate organelle and decides the fate of membrane cargoes in the LV.In yeast (Saccharomyces cerevisiae), carboxypeptidase S (CPS) is synthesized as a type II integral membrane protein and sorted from the Golgi to the lumen of the vacuole (Spormann et al., 1992). Genetic analyses on the trafficking of CPS have led to the identification of approximately 17 class E genes (Piper et al., 1995; Babst et al., 1997, 2002a, 2002b; Odorizzi et al., 1998; Katzmann et al., 2001) that constitute the core endosomal sorting complex required for transport (ESCRT) machinery. The evolutionarily conserved ESCRT complex consists of several functionally different subcomplexes, ESCRT-0, ESCRT-I, ESCRT-II, and ESCRT-III and the ESCRT-III-associated/Vacuolar Protein Sorting4 (VPS4) complex. Together, they form a complex protein-protein interaction network that coordinates sorting of cargoes and inward budding of the membrane on the MVB (Hurley and Hanson, 2010; Henne et al., 2011). Cargo proteins carrying ubiquitin signals are thought to be passed from one ESCRT subcomplex to the next, starting with their recognition by ESCRT-0 (Bilodeau et al., 2002, 2003; Hislop and von Zastrow, 2011; Le Bras et al., 2011; Shields and Piper, 2011; Urbé, 2011). ESCRT-0 recruits the ESCRT-I complex, a heterotetramer of VPS23, VPS28, VPS37, and MVB12, from the cytosol to the endosomal membrane (Katzmann et al., 2001, 2003). The C terminus of VPS28 interacts with the N terminus of VPS36, a member of the ESCRT-II complex (Kostelansky et al., 2006; Teo et al., 2006). Then, cargoes passed from ESCRT-I and ESCRT-II are concentrated in certain membrane domains of the endosome by ESCRT-III, which includes four coiled-coil proteins and is sufficient to induce the membrane invagination (Babst et al., 2002b; Saksena et al., 2009; Wollert et al., 2009). Finally, the ESCRT components are disassociated from the membrane by the adenosine triphosphatase (ATPase) associated with diverse cellular activities (AAA) VPS4/SUPPRESSOR OF K+ TRANSPORT GROWTH DEFECT1 (SKD1) before releasing the internal vesicles (Babst et al., 1997, 1998).Putative homologs of ESCRT-I–ESCRT-III and ESCRT-III-associated components have been identified in plants, except for ESCRT-0, which is only present in Opisthokonta (Winter and Hauser, 2006; Leung et al., 2008; Schellmann and Pimpl, 2009). To date, only a few plant ESCRT components have been studied in detail. The Arabidopsis (Arabidopsis thaliana) AAA ATPase SKD1 localized to the PVC/MVB and showed ATPase activity that was regulated by Lysosomal Trafficking Regulator-Interacting Protein5, a plant homolog of Vps Twenty Associated1 Protein (Haas et al., 2007). Expression of the dominant negative form of SKD1 caused an increase in the size of the MVB and a reduction in the number of internal vesicles (Haas et al., 2007). This protein also contributes to the maintenance of the central vacuole and might be associated with cell cycle regulation, as leaf trichomes expressing its dominant negative mutant form lost the central vacuole and frequently contained multiple nuclei (Shahriari et al., 2010). Double null mutants of CHARGED MULTIVESICULAR BODY PROTEIN, chmp1achmp1b, displayed severe growth defects and were seedling lethal. This may be due to the mislocalization of plasma membrane (PM) proteins, including those involved in auxin transport such as PINFORMED1, PINFORMED2, and AUXIN-RESISTANT1, from the vacuolar degradation pathway to the tonoplast of the LV (Spitzer et al., 2009).Plant ESCRT components usually contain several homologs, with the possibility of functional redundancy. Single mutants of individual ESCRT components may not result in an obvious phenotype, whereas knockout of all homologs of an ESCRT component by generating double or triple mutants may be lethal to the plant. As a first step to carry out systematic analysis on each ESCRT complex in plant cells, here, we established an efficient analysis system to monitor the localization changes of four vacuolar reporters that accumulate either in the lumen (LRR84A-GFP, EMP12-GFP, and aleurain-GFP) or on the tonoplast (GFP-VIT1) of the LV and identified several ESCRT-III dominant negative mutants. We reported that ESCRT-III subunits were involved in the release of PVC/MVB’s internal vesicles from the limiting membrane and were required for membrane protein degradation from secretory and endocytic pathways. In addition, transgenic Arabidopsis plants with induced expression of ESCRT-III dominant negative mutants showed severe cotyledon developmental defects. We also showed that membrane dissociation of ESCRT-III subunits was regulated by direct interaction with SKD1.     

The Role of a Potassium Transporter OsHAK5 in Potassium Acquisition and Transport from Roots to Shoots in Rice at Low Potassium Supply Levels

In plants, K transporter (KT)/high affinity K transporter (HAK)/K uptake permease (KUP) is the largest potassium (K) transporter family; however, few of the members have had their physiological functions characterized in planta. Here, we studied OsHAK5 of the KT/HAK/KUP family in rice (Oryza sativa). We determined its cellular and tissue localization and analyzed its functions in rice using both OsHAK5 knockout mutants and overexpression lines in three genetic backgrounds. A β-glucuronidase reporter driven by the OsHAK5 native promoter indicated OsHAK5 expression in various tissue organs from root to seed, abundantly in root epidermis and stele, the vascular tissues, and mesophyll cells. Net K influx rate in roots and K transport from roots to aerial parts were severely impaired by OsHAK5 knockout but increased by OsHAK5 overexpression in 0.1 and 0.3 mm K external solution. The contribution of OsHAK5 to K mobilization within the rice plant was confirmed further by the change of K concentration in the xylem sap and K distribution in the transgenic lines when K was removed completely from the external solution. Overexpression of OsHAK5 increased the K-sodium concentration ratio in the shoots and salt stress tolerance (shoot growth), while knockout of OsHAK5 decreased the K-sodium concentration ratio in the shoots, resulting in sensitivity to salt stress. Taken together, these results demonstrate that OsHAK5 plays a major role in K acquisition by roots faced with low external K and in K upward transport from roots to shoots in K-deficient rice plants.Potassium (K) is one of the three most important macronutrients and the most abundant cation in plants. As a major osmoticum in the vacuole, K drives the generation of turgor pressure, enabling cell expansion. In the vascular tissue, K is an important participant in the generation of root pressure (for review, see Wegner, 2014 [including his new hypothesis]). In the phloem, K is critical for the transport of photoassimilates from source to sink (Marschner, 1996; Deeken et al., 2002; Gajdanowicz et al., 2011). In addition, enhancing K absorption and decreasing sodium (Na) accumulation is a major strategy of glycophytes in salt stress tolerance (Maathuis and Amtmann, 1999; Munns and Tester, 2008; Shabala and Cuin, 2008).Plants acquire K through K-permeable proteins at the root surface. Since available K concentration in the soil may vary by 100-fold, plants have developed multiple K uptake systems for adapting to this variability (Epstein et al., 1963; Grabov, 2007; Maathuis, 2009). In a classic K uptake experiment in barley (Hordeum vulgare), root K absorption has been described as a high-affinity and low-affinity biphasic transport process (Epstein et al., 1963). It is generally assumed that the low-affinity transport system (LATS) in the roots mediates K uptake in the millimolar range and that the activity of this system is insensitive to external K concentration (Maathuis and Sanders, 1997; Chérel et al., 2014). In contrast, the high-affinity transport system (HATS) was rapidly up-regulated when the supply of exogenous K was halted (Glass, 1976; Glass and Dunlop, 1978).The membrane transporters for K flux identified in plants are generally classified into three channels and three transporter families based on phylogenetic analysis (Mäser et al., 2001; Véry and Sentenac, 2003; Lebaudy et al., 2007; Alemán et al., 2011). For K uptake, it was predicted that, under most circumstances, K transporters function as HATS, while K-permeable channels mediate LATS (Maathuis and Sanders, 1997). However, a root-expressed K channel in Arabidopsis (Arabidopsis thaliana), Arabidopsis K Transporter1 (AKT1), mediates K absorption over a wide range of external K concentrations (Sentenac et al., 1992; Lagarde et al., 1996; Hirsch et al., 1998; Spalding et al., 1999), while evidence is accumulating that many K transporters, including members of the K transporter (KT)/high affinity K transporter (HAK)/K uptake permease (KUP) family, are low-affinity K transporters (Quintero and Blatt, 1997; Senn et al., 2001), implying that functions of plant K channels and transporters overlap at different K concentration ranges.Out of the three families of K transporters, cation proton antiporter (CPA), high affinity K/Na transporter (HKT), and KT/HAK/KUP, CPA was characterized as a K⁺(Na⁺)/H⁺ antiporter, HKT may cotransport Na and K or transport Na only (Rubio et al., 1995; Uozumi et al., 2000), while KT/HAK/KUP were predicted to be H⁺-coupled K⁺ symporters (Mäser et al., 2001; Lebaudy et al., 2007). KT/HAK/KUP were named by different researchers who first identified and cloned them (Quintero and Blatt, 1997; Santa-María et al., 1997). In plants, the KT/HAK/KUP family is the largest K transporter family, including 13 members in Arabidopsis and 27 members in the rice (Oryza sativa) genome (Rubio et al., 2000; Mäser et al., 2001; Bañuelos et al., 2002; Gupta et al., 2008). Sequence alignments show that genes of this family share relatively low homology to each other. The KT/HAK/KUP family was divided into four major clusters (Rubio et al., 2000; Gupta et al., 2008), and in cluster I and II, they were further separated into A and B groups. Genes of cluster I or II likely exist in all plants, cluster III is composed of genes from both Arabidopsis and rice, while cluster IV includes only four rice genes (Grabov, 2007; Gupta et al., 2008).The functions of KT/HAK/KUP were studied mostly in heterologous expression systems. Transporters of cluster I, such as AtHAK5, HvHAK1, OsHAK1, and OsHAK5, are localized in the plasma membrane (Kim et al., 1998; Bañuelos et al., 2002; Gierth et al., 2005) and exhibit high-affinity K uptake in the yeast Saccharomyces cerevisiae (Santa-María et al., 1997; Fu and Luan, 1998; Rubio et al., 2000) and in Escherichia coli (Horie et al., 2011). Transporters of cluster II, like AtKUP4 (TINY ROOT HAIRS1, TRH1), HvHAK2, OsHAK2, OsHAK7, and OsHAK10, could not complement the K uptake-deficient yeast (Saccharomyces cerevisiae) but were able to mediate K fluxes in a bacterial mutant; they might be tonoplast transporters (Senn et al., 2001; Bañuelos et al., 2002; Rodríguez-Navarro and Rubio, 2006). The function of transporters in clusters III and IV is even less known (Grabov, 2007).Existing data suggest that some KT/HAK/KUP transporters also may respond to salinity stress (Maathuis, 2009). The cluster I transporters of HvHAK1 mediate Na influx (Santa-María et al., 1997), while AtHAK5 expression is inhibited by Na (Rubio et al., 2000; Nieves-Cordones et al., 2010). Expression of OsHAK5 in tobacco (Nicotiana tabacum) BY2 cells enhanced the salt tolerance of these cells by accumulating more K without affecting their Na content (Horie et al., 2011).There are only scarce reports on the physiological function of KT/HAK/KUP in planta. In Arabidopsis, mutation of AtKUP2 (SHORT HYPOCOTYL3) resulted in a short hypocotyl, small leaves, and a short flowering stem (Elumalai et al., 2002), while a loss-of-function mutation of AtKUP4 (TRH1) resulted in short root hairs and a loss of gravity response in the root (Rigas et al., 2001; Desbrosses et al., 2003; Ahn et al., 2004). AtHAK5 is the only system currently known to mediate K uptake at concentrations below 0.01 mm (Rubio et al., 2010) and provides a cesium uptake pathway (Qi et al., 2008). AtHAK5 and AtAKT1 are the two major physiologically relevant molecular entities mediating K uptake into roots in the range between 0.01 and 0.05 mm (Pyo et al., 2010; Rubio et al., 2010). AtAKT1 may contribute to K uptake within the K concentrations that belong to the high-affinity system described by Epstein et al. (1963).Among all 27 members of the KT/HAK/KUP family in rice, OsHAK1, OsHAK5, OsHAK19, and OsHAK20 were grouped in cluster IB (Gupta et al., 2008). These four rice HAK members share 50.9% to 53.4% amino acid identity with AtHAK5. OsHAK1 was expressed in the whole plant, with maximum expression in roots, and was up-regulated by K deficiency; it mediated high-affinity K uptake in yeast (Bañuelos et al., 2002). In this study, we examined the tissue-specific localization and the physiological functions of OsHAK5 in response to variation in K supply and to salt stress in rice. By comparing K uptake and translocation in OsHAK5 knockout (KO) mutants and in OsHAK5-overexpressing lines with those in their respective wild-type lines supplied with different K concentrations, we found that OsHAK5 not only mediates high-affinity K acquisition but also participates in root-to-shoot K transport as well as in K-regulated salt tolerance.     

Bisphosphonate Inhibitors Reveal a Large Elasticity of Plastidic Isoprenoid Synthesis Pathway in Isoprene-Emitting Hybrid Aspen

Recently, a feedback inhibition of the chloroplastic 1-deoxy-d-xylulose 5-phosphate (DXP)/2-C-methyl-d-erythritol 4-phosphate (MEP) pathway of isoprenoid synthesis by end products dimethylallyl diphosphate (DMADP) and isopentenyl diphosphate (IDP) was postulated, but the extent to which DMADP and IDP can build up is not known. We used bisphosphonate inhibitors, alendronate and zoledronate, that inhibit the consumption of DMADP and IDP by prenyltransferases to gain insight into the extent of end product accumulation and possible feedback inhibition in isoprene-emitting hybrid aspen (Populus tremula × Populus tremuloides). A kinetic method based on dark release of isoprene emission at the expense of substrate pools accumulated in light was used to estimate the in vivo pool sizes of DMADP and upstream metabolites. Feeding with fosmidomycin, an inhibitor of DXP reductoisomerase, alone or in combination with bisphosphonates was used to inhibit carbon input into DXP/MEP pathway or both input and output. We observed a major increase in pathway intermediates, 3- to 4-fold, upstream of DMADP in bisphosphonate-inhibited leaves, but the DMADP pool was enhanced much less, 1.3- to 1.5-fold. In combined fosmidomycin/bisphosphonate treatment, pathway intermediates accumulated, reflecting cytosolic flux of intermediates that can be important under strong metabolic pull in physiological conditions. The data suggested that metabolites accumulated upstream of DMADP consist of phosphorylated intermediates and IDP. Slow conversion of the huge pools of intermediates to DMADP was limited by reductive energy supply. These data indicate that the DXP/MEP pathway is extremely elastic, and the presence of a significant pool of phosphorylated intermediates provides an important valve for fine tuning the pathway flux.Isoprenoids constitute a versatile class of compounds fulfilling major physiological functions. They are formed by two pathways in plants, the mevalonate (MVA) pathway in the cytosol (Gershenzon and Croteau, 1993) and the 1-deoxy-d-xylulose 5-phosphate (DXP)/2-C-methyl-d-erythritol 4-phosphate (MEP) pathway in plastids (Gershenzon and Croteau, 1993; Jomaa et al., 1999; Li and Sharkey, 2013b). The MVA pathway is primarily responsible for the synthesis of sesquiterpenes (C15), triterpenes (C30) including brassinosteroids, and even larger molecules such as dolichols (Bick and Lange, 2003; Li and Sharkey, 2013b; Rajabi Memari et al., 2013; Rosenkranz and Schnitzler, 2013). The DXP/MEP pathway is responsible for the synthesis of the simplest isoprenoids, isoprene and 2-methyl-3-buten-2-ol (C5), monoterpenes (C10), diterpenes (C20) including gibberellins and phytol residue of chlorophylls, and tetraterpenes (C40) including carotenoids (Rodríguez-Concepción and Boronat, 2002; Roberts, 2007).Given that in plants both pathways produce ultimately the same substrates, dimethylallyl diphosphate (DMADP) and isopentenyl diphosphate (IDP), the pertinent question is to what extent the two pathways can exchange metabolites (Rodríguez-Concepción and Boronat, 2002). There is evidence of a certain exchange of IDP between cytosolic and plastidic compartments, although the contribution of IDP from one compartment to the pathway flux in the other seems to be relatively minor (Schwender et al., 2001; Rodríguez-Concepción and Boronat, 2002; Bick and Lange, 2003). Some studies have further demonstrated that the exchange of IDP is fully bidirectional (De-Eknamkul and Potduang, 2003; Rodríguez-Concepción, 2006), whereas other studies suggest that IDP export from plastids to cytosol operates with a greater efficiency than the opposite transport (Hemmerlin et al., 2003; Laule et al., 2003). However, although the overall intercompartmental exchange of isoprenoid substrates to pathway flux in the given compartment might seem minor under nonstressed conditions, the importance of cross talk among the pathways might increase under stress conditions that specifically inhibit isoprenoid synthesis in one pathway. In fact, the DXP/MEP pathway is strongly linked to photosynthetic metabolism, and therefore, inhibition of photosynthesis under stressful conditions such as heat stress or drought or photoinhibition could inhibit the synthesis of isoprenoids when they are most needed to fulfill their protective function (Loreto and Schnitzler, 2010; Niinemets, 2010; Possell and Loreto, 2013). There is some evidence demonstrating a certain cooperativity among the two isoprenoid synthesis pathways under conditions leading to a reduction of the activity of one of them (Piel et al., 1998; Jux et al., 2001; Page et al., 2004; Rodríguez-Concepción, 2006), but the capacity for such a replacement of function and regulation is poorly understood.Recent studies using genetically modified plants accumulating end products of the DXP/MEP pathway or using natural variation in product accumulation have demonstrated the existence of a potentially important feedback regulation of the DXP/MEP pathway flux by primary end products of the pathway (Banerjee et al., 2013; Ghirardo et al., 2014; Wright et al., 2014). In particular, binding of DMADP and perhaps IDP to DXP synthase, the first enzyme in the DXP/MEP pathway, leads to downregulation of the pathway flux when the end products cannot be used, such as under stress conditions. However, the strength of such a feedback regulation can be importantly modified by accumulation of phosphorylated intermediates of the pathway, such that DMADP and IDP do not accumulate. Previous studies have demonstrated that there is a certain pool of phosphorylated intermediates in vivo, and that this pool can strongly increase under certain conditions, including experimental and genetic modification of DXP/MEP pathway input and output (Li et al., 2011; Rasulov et al., 2011; Li and Sharkey, 2013a; Ghirardo et al., 2014; Wright et al., 2014).It has further been shown that 2-C-methyl-d-erythritol 2,4-cyclodiphosphate (ME-cDP) is the metabolite accumulating in the plastids, and this accumulation can buffer DMADP and IDP changes in the case of varying DXP/MEP pathway input and consumption (Li and Sharkey, 2013a; Wright et al., 2014). A significant part of ME-cDP might even escape to cytosol, implying the existence of another interesting link between cytosolic and chloroplastic processes in isoprenoid synthesis (Wright et al., 2014). Furthermore, as ME-cDP is an important signaling molecule eliciting a number of gene expression responses (Xiao et al., 2012), accumulation of ME-cDP in plastids and flux to cytosol and further to the nucleus is particularly interesting from the perspective of long-term regulation of isoprenoid synthesis, and suggests a coordination of cellular stress responses by the plastidial isoprenoid synthesis pathway.Isoprene-emitting species constitute an exciting model system where a very large DXP/MEP pathway flux goes to isoprene synthesis under physiological conditions (Li and Sharkey, 2013b; Sharkey et al., 2013). In isoprene-emitting species, there is a concomitant use of the primary substrate DMADP between the plastidic synthesis of isoprene and isoprenoids with a larger molecular size, such as phytol residue of chlorophyll (C20) and carotenoids (C40; Ghirardo et al., 2014; Rasulov et al., 2014), and different from nonemitting species, isoprene emitters seem to support a much larger pool of DMADP without the onset of feedback inhibition (Ghirardo et al., 2014; Wright et al., 2014). However, it is poorly understood how inhibition of one branch of the pathway (isoprene versus larger isoprenoids) affects the other, to what extent it can lead to accumulation of phosphorylated intermediates, how it affects the overall pathway flux through the feedback regulation, and what is the possible role of cytosolic import and export of intermediates. These are all relevant questions to gain insight into the control of the partitioning of pathway flux between isoprene and larger isoprenoids and to understand the biological role of isoprene emission.Studies using metabolic inhibitors to deconvolute the factors involved in pathway regulation and understand the biological role of isoprene have so far used inhibitors that block the early steps of the corresponding pathways. In particular, fosmidomycin, a specific inhibitor of DXP reductoisomerase, the enzyme responsible for the synthesis of MEP from DXP, has been used to inhibit the DXP/MEP pathway (Loreto and Velikova, 2001; Sharkey et al., 2001; Loreto et al., 2004). In addition, lovastatin (mevinolin), the inhibitor of 3-hydroxy-3-methylglutaryl-CoA reductase that controls the MVA pathway flux, has been used to study the cooperativity of the two pathways (e.g. Laule et al., 2003; Mansouri and Salari, 2014). However, these inhibitors are not suitable to understand how end product accumulation can alter the pathway flux.Bisphosphonates constitute a promising class of inhibitors that could be particularly apt for studies on the effects of the inhibition of the end points of the pathway. They have been demonstrated to inhibit cytosolic farnesyl diphosphate (FDP) synthase activity (Oberhauser et al., 1998; Cromartie et al., 1999; van Beek et al., 1999; Bergstrom et al., 2000; Burke et al., 2004), аs well as geranyl diphosphate (GDP) and geranylgeranyl diphosphate (GGDP) synthase activities (Oberhauser et al., 1998; Cromartie et al., 1999; Kloer et al., 2006; No et al., 2012; Lindert et al., 2013). To our knowledge, bisphosphonates have not been used to study the effects of end product accumulation on the pathway flux in isoprene-emitting species, with the exception of one study that investigated the development of isoprene emission capacity through leaf ontogeny (Rasulov et al., 2014).A limitation with any inhibitor study could be a certain nonspecificity, inhibition of additional nondesired reactions, but so far there are no data on such nonspecificity of bisphosphonates. However, there is evidence that diphosphate and its analogs are inhibitors of any ferredoxin (Fd)-dependent reaction (Forti and Meyer, 1969; Bojko and Więckowski, 1999). This could be potentially relevant given that DXP/MEP pathway-reducing steps, at the level of 4-hydroxy-3-methyl-2-(E)-butenyl diphosphate (HMBDP) synthase (HDS) and reductase (HDR), directly accept electrons from Fd in light (Eisenreich et al., 2001; Seemann et al., 2006; Li and Sharkey, 2013a). In addition to the DXP/MEP pathway, inhibition at the level of Fd could also affect photosynthetic reactions and thereby alter energy supply for the DXP/MEP pathway.In this study, we have investigated the effects of inhibition of the initial and final steps of the DXP/MEP pathway by fosmidomycin and bisphosphonate inhibitors alendronate and zoledronate in a strong isoprene emitter hybrid aspen (Populus tremula × Populus tremuloides). Alendronate is a highly specific inhibitor of GDP (Lange et al., 2001; Burke et al., 2004) and FDP synthases (Bergstrom et al., 2000; Burke et al., 2004), and a less specific inhibitor of GGDP synthase (Szabo et al., 2002). Zoledronate operates similarly to alendronate, but is a much stronger inhibitor, being operationally active in concentrations several orders of magnitude less than alendronate (Lange et al., 2001; Henneman et al., 2011; Wasko, 2011). A unique in vivo method was used to study dynamic changes in DMADP and phosphorylated intermediate pool sizes (Rasulov et al., 2009a, 2011; Li et al., 2011), and different inhibitors were applied alone or in sequence to study the regulation of the pathway flux in conditions when the flux out of the pathway or into the pathway is curbed and when both the input and the output are curbed. Dynamic model calculations were used to quantitatively evaluate the significance of the cytosolic intermediate input into chloroplastic isoprenoid synthesis under different conditions of the DXP/MEP pathway entrance and exit, and to evaluate the possible nonspecific inhibition of other steps controlling DXP/MEP pathway flux. The study demonstrates the important regulation of DXP/MEP pathway input and output under conditions of end product accumulation and partial cooperativity among chloroplastic and cytosolic isoprenoid synthesis pathways.     

Arabidopsis Small Rubber Particle Protein Homolog SRPs Play Dual Roles as Positive Factors for Tissue Growth and Development and in Drought Stress Responses

Lipid droplets (LDs) act as repositories for fatty acids and sterols, which are used for various cellular processes such as energy production and membrane and hormone synthesis. LD-associated proteins play important roles in seed development and germination, but their functions in postgermination growth are not well understood. Arabidopsis (Arabidopsis thaliana) contains three SRP homologs (SRP1, SRP2, and SRP3) that share sequence identities with small rubber particle proteins of the rubber tree (Hevea brasiliensis). In this report, the possible cellular roles of SRPs in postgermination growth and the drought tolerance response were investigated. Arabidopsis SRPs appeared to be LD-associated proteins and displayed polymerization properties in vivo and in vitro. SRP-overexpressing transgenic Arabidopsis plants (35S:SRP1, 35S:SRP2, and 35S:SRP3) exhibited higher vegetative and reproductive growth and markedly better tolerance to drought stress than wild-type Arabidopsis. In addition, constitutive over-expression of SRPs resulted in increased numbers of large LDs in postgermination seedlings. In contrast, single (srp1, 35S:SRP2-RNAi, and srp3) and triple (35S:SRP2-RNAi/srp1srp3) loss-of-function mutant lines exhibited the opposite phenotypes. Our results suggest that Arabidopsis SRPs play dual roles as positive factors in postgermination growth and the drought stress tolerance response. The possible relationships between LD-associated proteins and the drought stress response are discussed.Environmental stresses, including drought, high salinity, oxidative stress, and unfavorable temperatures, profoundly affect the growth and development of higher plants. Because of their sessile life cycle, plants have developed self-protective mechanisms to increase their tolerance to short- and long-term stresses by triggering diverse sets of signal transduction pathways and activating stress-responsive genes. The genetic and cellular mechanisms in response to abiotic stress have been widely documented in higher plants (Shinozaki and Yamaguchi-Shinozaki, 1996; Bray, 1997; Ishitani et al., 1997; Zhu, 2002; Bohnert et al., 2006; Shinozaki and Yamaguchi-Shinozaki, 2007; Vij and Tyagi, 2007).Lipid droplets (LDs) are dynamic subcellular organelles enclosed by a monolayer of phospholipid. LDs act as repositories for fatty acids and sterols, which are used for energy production and membrane and hormone synthesis. LDs are also involved in various cellular processes, including intracellular protein storage, stress responses, and lipid signaling (Bartz et al., 2007; Zehmer et al., 2009; Carman, 2012; Herker and Ott, 2012; Murphy, 2012; Sun et al., 2013; Kory et al., 2015). LDs bud from the endoplasmic reticulum (ER), where they become enriched with triacylglycerols and subsequently enlarged, until they pinch off to form an LD (Chapman et al., 2012; Chapman and Ohlrogge, 2012; Jacquier et al., 2013). Several reports suggest that LD-associated proteins, such as fat-specific protein 27 (FSP27), SEIPIN, and PERILIPIN1 (Plin1), are key regulators of LD formation in mammals, Drosophila, and yeasts (Farese and Walther, 2009; Xu et al., 2012; Yang et al., 2012). After budding from the ER, LDs fuse with each other and expand. In adipocytes, Plin1 functions as an enhancer of FSP27-mediated lipid transfer and LD growth, indicating that Plin1 and FSP27 participate in LD formation and fusion (Sun et al., 2013). Enlarged LDs provide surfaces to allow the attachment of numerous LD-associated proteins, which are later displaced during shrinkage of LDs by lipolysis (Kory et al., 2015).Because LDs are mainly present in seeds, studies on LD-associated proteins in higher plants have focused on seed development and germination (Chapman et al., 2013; Gidda et al., 2013; Horn et al., 2013; Szymanski et al., 2014). For example, oleosins regulate LD size in Arabidopsis (Arabidopsis thaliana) seed development (Siloto et al., 2006). Arabidopsis SEIPINs modulate LD proliferation and neutral lipid accumulation in developing seeds (Cai et al., 2015). On the other hand, the cellular roles of LD-associated proteins in postgermination growth remain largely unraveled.CaSRP1 (Capsicum annuum stress-related protein 1) was previously identified as a hot pepper small rubber particle protein (SRPPs) homolog (Hong and Kim, 2005). CaSRP1 was induced in response to water stress in hot pepper plants. Constitutive over-expression of CaSRP1 in transgenic Arabidopsis plants resulted in elevated growth and increased drought tolerance relative to wild-type Arabidopsis (Kim et al., 2010). CaSRP1 is evolutionarily related to SRPPs in rubber-producing plants (Wititsuwannakul et al., 2008). Rubber particles are single-membrane organelles that store rubber (cis-1,4-polyisoprene). Although rubber particles and LDs have different lipid compositions, their basic architectures are similar (Cornish et al., 1999). Thus, SRPP homologs may have common properties in the formation and biogenesis of rubber particles and/or LDs in rubber-producing and non-rubber-producing plants.In this report, we identified and characterized three SRPP homologs, SRP1, SRP2, and SRP3, in Arabidopsis. The SRP genes were differentially expressed in various tissues and induced by abscisic acid (ABA) and a broad spectrum of abiotic stress, including drought, high salinity, and low temperature. SRP-overexpressing transgenic Arabidopsis plants (35S:SRP1, 35S:SRP2, and 35S:SRP3) exhibited higher vegetative and reproductive growth and markedly better tolerance to drought stress than wild-type Arabidopsis plants. In addition, ectopic expression of SRPs resulted in increased numbers of large LDs in postgermination seedlings. In contrast, single (srp1, 35S:SRP2-RNAi, and srp3) and triple (35S:SRP2-RNAi/srp1srp3) loss-of-function mutant lines showed the opposite phenotypes. Arabidopsis SRPs appeared to be LD-associated proteins and displayed polymerization properties in vivo and in vitro. These results are discussed in light of the suggestion that Arabidopsis SRPs play dual roles as positive factors in postgermination growth and drought stress response. The possible relationships between LD-associated proteins and stress tolerance response are also discussed.     

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.     

Specialization of Oleosins in Oil Body Dynamics during Seed Development in Arabidopsis Seeds

Oil bodies (OBs) are seed-specific lipid storage organelles that allow the accumulation of neutral lipids that sustain plantlet development after the onset of germination. OBs are covered with specific proteins embedded in a single layer of phospholipids. Using fluorescent dyes and confocal microscopy, we monitored the dynamics of OBs in living Arabidopsis (Arabidopsis thaliana) embryos at different stages of development. Analyses were carried out with different genotypes: the wild type and three mutants affected in the accumulation of various oleosins (OLE1, OLE2, and OLE4), three major OB proteins. Image acquisition was followed by a detailed statistical analysis of OB size and distribution during seed development in the four dimensions (x, y, z, and t). Our results indicate that OB size increases sharply during seed maturation, in part by OB fusion, and then decreases until the end of the maturation process. In single, double, and triple mutant backgrounds, the size and spatial distribution of OBs are modified, affecting in turn the total lipid content, which suggests that the oleosins studied have specific functions in the dynamics of lipid accumulation.The seed is a complex, specific structure that allows a quiescent plant embryo to cope with unfavorable germinating conditions and also permits dissemination of the species. To achieve these functions, seeds accumulate reserve compounds that will ensure the survival of the embryo and fuel the growth of the plantlet upon germination. Accumulation of lipids occurs in many eukaryotic cells and is a rather common means of storing carbon and energy. Lipid droplets (LDs) can be found in all eukaryotes, such as yeast (Saccharomyces cerevisiae; Leber et al., 1994), mammals (Murphy, 2001; Hodges and Wu, 2010), Caenorhabditis elegans (Zhang et al., 2010; Mak, 2012), Drosophila melanogaster (Beller et al., 2006, 2010), and plants (Hsieh and Huang, 2004), but also in prokaryotes (Wältermann et al., 2005). The basic structure of an LD is a core of neutral lipids covered by a phospholipid monolayer. LDs differ between species by the set of proteins covering their surface, the nature of the lipids stored, and their turnover. Nevertheless, they apparently always ensure the same function in the cell (i.e. energy storage; Murphy, 2012). In Brassicacea species such as Arabidopsis (Arabidopsis thaliana), seed reserves are mainly composed of carbohydrates, proteins, and lipids (Baud et al., 2002). The lipids are primarily stored as triacylglycerols (TAGs) in LDs, more commonly called oil bodies (OBs; Hsieh and Huang, 2004; Chapman et al., 2012; Chapman and Ohlrogge, 2012) of diameter 0.5 to 2 µm (Tzen et al., 1993).The protein composition of seed OBs has been determined for several plant species, including Brassica napus (Katavic et al., 2006; Jolivet et al., 2009) and Arabidopsis (Jolivet et al., 2004; D’Andréa et al., 2007; Vermachova et al., 2011). In Arabidopsis, 10 proteins have been identified, and seed-specific oleosins represent up to 79% of the OB proteins (Jolivet et al., 2004; D’Andréa et al., 2007; Vermachova et al., 2011). Oleosins are rather small proteins of 18.5 to 21.2 kD with a specific and highly conserved central hydrophobic domain of 72 amino acid residues flanked by hydrophilic domains of variable size and amino acid composition (Qu and Huang, 1990; Tzen et al., 1990, 1992; Huang, 1996; Hsieh and Huang, 2004). It is generally agreed that oleosins cover the OB surface, with their central hydrophobic domain inserted in the TAG through the phospholipid layer (Tzen and Huang, 1992). Besides their structural function in OBs, oleosins may serve as docking stations for other proteins at its surface (Wilfling et al., 2013) and may participate in the biosynthesis and mobilization of plant oils (Parthibane et al., 2012a, 2012b). Oleosins are probably involved in OB stability (Leprince et al., 1998; Shimada et al., 2008) and in the regulation of OB repulsion (Heneen et al., 2008), preventing the coalescence of OBs into a single organelle (Schmidt and Herman, 2008). Nevertheless, the precise functions of oleosins in OB biogenesis and dynamics have not yet been established.Global analysis of seed lipids can be performed using gas chromatography (Li et al., 2006), which allows the precise determination of both lipid content and fatty acid composition. Recently, direct organelle mass spectrometry has been used to visualize the lipid composition of cotton (Gossypium hirsutum) seed OBs (Horn et al., 2011). Nevertheless, in both cases, the methods are destructive. To observe lipid accumulation at the subcellular level, well-known nondestructive techniques for lipid visualization have been adapted to seeds. Third harmonic generation microscopy (Débarre et al., 2006) and label-free coherent anti-Stokes Raman scattering microscopy (Paar et al., 2012) allow dyeless observation of LDs but require very specific equipment. Magnetic resonance imaging enables topographic analysis of lipid distribution in cereal grains (Neuberger et al., 2008) and in submillimeter-sized seeds like those of tobacco (Nicotiana tabacum; Fuchs et al., 2013). Nevertheless, the use of fluorescent dyes such as Nile Red (Greenspan and Fowler, 1985), BODIPY (Pagano et al., 1991), or LipidTOX (Invitrogen) associated with confocal microscopy is also a powerful way to monitor LDs in living organisms.Despite knowledge accumulated on this topic (Brasaemle and Wolins, 2012; Chapman et al., 2012), little is known about OB dynamics during seed maturation. In this article, we investigate this question by monitoring the evolution of OBs in living Arabidopsis embryos over time. This analysis showed a marked change in OB size at 9 to 10 d after flowering (DAF). We then examined single, double, and triple mutants of the major oleosins found in developing seeds (OLE1 [At4g25140], OLE2 [At5g40420], and OLE4 [At3g01570]; Jolivet et al., 2004). We analyzed the OB dynamics in these mutant backgrounds as if they would contain only these three proteins. We show that the lack of specific oleosins influences the dynamics and distribution of OBs during seed maturation, which in turn affects lipid accumulation. These results pave the way for analyzing specific functions of oleosins in the synthesis, growth, and evolution of OBs.     

Remodeling of the Infection Chamber before Infection Thread Formation Reveals a Two-Step Mechanism for Rhizobial Entry into the Host Legume Root Hair

In many legumes, root entry of symbiotic nitrogen-fixing rhizobia occurs via host-constructed tubular tip-growing structures known as infection threads (ITs). Here, we have used a confocal microscopy live-tissue imaging approach to investigate early stages of IT formation in Medicago truncatula root hairs (RHs) expressing fluorescent protein fusion reporters. This has revealed that ITs only initiate 10 to 20 h after the completion of RH curling, by which time major modifications have occurred within the so-called infection chamber, the site of bacterial entrapment. These include the accumulation of exocytosis (M. truncatula Vesicle-Associated Membrane Protein721e)- and cell wall (M. truncatula EARLY NODULIN11)-associated markers, concomitant with radial expansion of the chamber. Significantly, the infection-defective M. truncatula nodule inception-1 mutant is unable to create a functional infection chamber. This underlines the importance of the NIN-dependent phase of host cell wall remodeling that accompanies bacterial proliferation and precedes IT formation, and leads us to propose a two-step model for rhizobial infection initiation in legume RHs.Legumes possess the remarkable capacity to improve their nutrition by establishing a nitrogen-fixing root nodule symbiosis (RNS) with soil bacteria collectively called rhizobia. In many legumes such as Medicago truncatula, rhizobia penetrate across the root epidermis and outer cortex to reach the differentiating nodule tissues via sequentially constructed transcellular compartments known as infection threads (ITs; Gage, 2004). It is now well established that this mode of entry through specialized infection compartments, often referred to as accommodation, is shared with the more ancient arbuscular mycorrhizal (AM) symbiosis, from which the legume-Rhizobium RNS is thought to have evolved (Parniske, 2008; Markmann and Parniske, 2009). Furthermore, strong evidence indicates that the signaling and cellular mechanisms underlying IT formation in legumes are closely related to those used for infection compartment formation during AM infection of epidermal and outer cortical tissues (Bapaume and Reinhardt, 2012; Oldroyd, 2013).Rhizobial infection is set in motion after an initial molecular dialogue between symbiotic partners, in which rhizobial lipo-chitooligosaccharide (LCO) Nod factors (NFs) are key signaling molecules (for review, see Oldroyd, 2013). Host responses to NF signaling include rapid and sustained nuclear-associated Ca²⁺ oscillations (Ca²⁺ spiking; Ehrhardt et al., 1996; Oldroyd and Downie, 2006; Sieberer et al., 2009; Capoen et al., 2011) and the rapid expression of early epidermal marker genes such as M. truncatula EARLY NODULIN11 (Charron et al., 2004). The activation of nuclear Ca²⁺ spiking is one of the most characteristic features of the so-called common symbiotic signaling pathway, common to both RNS and AM (Kistner and Parniske, 2002; Singh and Parniske, 2012). Whereas these preinfection responses to NFs are observed in the majority of elongating root hairs (RHs) early after rhizobial inoculation (Journet et al., 2001; Wais et al., 2002), ITs are only formed in a small subset of RHs, and MtENOD11 expression is strongly activated at these rhizobial infection sites (Journet et al., 2001; Boisson-Dernier et al., 2005).ITs are tubular plant-derived structures delimited by a membrane that is contiguous with the RH plasmalemma and a layer of cell wall-like material, thus isolating the rhizobia from the host cell cytoplasm (Gage, 2004). These apoplastic infection compartments are progressively constructed along the length of the RH with their growing tip connected via a cytoplasmic bridge to the migrating RH nucleus. This broad cytoplasmic column provides the cell machinery for tip growth, which involves targeted exocytosis of membrane and extracellular materials to the growing apex of the IT (Oldroyd et al., 2011; Bapaume and Reinhardt, 2012). It is presumed that this cytoplasmic bridge shares an equivalent role to the prepenetration apparatus (PPA) formed at the onset of AM fungal infection (Genre et al., 2005, 2008). We now know that the IT tip region is formed in advance of rhizobial colonization and is progressively populated by dividing rhizobia that also physically move down the thread (Gage, 2004; Fournier et al., 2008). It has been proposed that the matrix of the growing IT tip is initially in a fluid or gel-like state compatible with bacterial growth and movement (Brewin, 2004; Fournier et al., 2008). This relative plasticity could result in part from the presence of atypical extracellular (glyco) proteins such as the repetitive Pro-rich proteins MtENOD11/MtENOD12 because their low Tyr content is presumed to limit cross linking to other wall components (Scheres et al., 1990; Pichon et al., 1992; Journet et al., 2001).Nevertheless, the mechanism by which rhizobial IT formation is initiated in RHs is not clear. Whereas AM fungal hyphae form contact structures called hyphopodia on the exposed surface of nonhair epidermal cells prior to PPA formation and perifungal infection compartment formation (Genre et al., 2005), rhizobial entry requires that the bacteria first become entrapped between RH walls. Attachment of rhizobia close to a growing RH tip induces a continuous reorientation of tip growth, most likely the result of localized NF production (Esseling et al., 2003), eventually leading to RH curling and subsequent bacterial entrapment within a closed chamber in the center of the curl (Catoira et al., 2001; Geurts et al., 2005). Rhizobial entrapment can also occur between the cell walls of two touching RHs (Dart, 1974; Gage, 2004).The closed chamber in curled RHs has often been termed the infection pocket (e.g. Murray, 2011; Guan et al., 2013). However, because this term is also used to designate a quite different and larger structure formed in root subepidermal tissues of legumes during intercellular infection after crack entry and involving localized cell death (Goormachtig et al., 2004), we propose to use the term infection chamber to describe the unique enclosure formed during rhizobial RH infection.After entrapment, it has been proposed that rhizobia multiply to form a so-called microcolony (Gage et al., 1996; Limpens et al., 2003), and that IT polar growth initiates in front of this microcolony by local invagination of the RH plasmalemma combined with exocytosis of extracellular materials (Gage, 2004). Furthermore, it has been suggested that localized degradation of the chamber wall would allow the rhizobia to access the newly formed IT (Callaham and Torrey, 1981; Turgeon and Bauer, 1985). However, a detailed investigation of this particular stage of rhizobial infection is lacking, particularly concerning when and where the rhizobia/cell wall interface becomes modified. Such studies have been limited until now, notably because ITs develop only in a low proportion of curled RHs (Dart, 1974).To attempt to answer this question, we have used a live-tissue imaging approach developed for in vivo confocal microscopy in M. truncatula (Fournier et al., 2008; Cerri et al., 2012; Sieberer et al., 2012) and particularly well adapted to time-lapse studies of the initial stages of rhizobial infection, including RH curling and IT formation. To investigate modifications occurring at the RH interface with the enclosed rhizobia during these early stages, we prepared M. truncatula plants expressing fluorescent protein fusions aimed at detecting both exocytosis activity and cell wall remodeling during the initial construction of the IT apoplastic compartment. To this end, we made use of the M. truncatula Vesicle-Associated Membrane Protein721e (MtVAMP721e; Ivanov et al., 2012), recently shown to label exocytosis sites both in growing RHs and during AM colonization (Genre et al., 2012), as well as the infection- and cell wall-associated MtENOD11 Pro-rich glycoprotein (Journet et al., 2001). Our experiments have revealed that IT development in curled RHs only initiates after a lengthy interval of 10 to 20 h, during which sustained exocytosis and MtENOD11 secretion to the infection chamber are associated with radial expansion as well as remodeling of the surrounding walls. Importantly, it was found that the infection-defective M. truncatula nodule inception-1 (Mtnin-1) mutant (Marsh et al., 2007) is impaired in chamber remodeling. Our findings led us to propose a new model for IT formation in which the infection chamber first differentiates into a globular apoplastic compartment displaying similarities to the future IT, and in which the enclosed rhizobia multiply. This is then followed by a switch from radial to tubular growth corresponding to tip-driven IT growth and associated movement of rhizobia into the extending thread. Importantly, this two-step model no longer requires that the host cell wall is degraded to allow access of the colonizing rhizobia to the newly initiated IT.     

Distinct Detoxification Mechanisms Confer Resistance to Mesotrione and Atrazine in a Population of Waterhemp

Previous research reported the first case of resistance to mesotrione and other 4-hydroxyphenylpyruvate dioxygenase (HPPD) herbicides in a waterhemp (Amaranthus tuberculatus) population designated MCR (for McLean County mesotrione- and atrazine-resistant). Herein, experiments were conducted to determine if target site or nontarget site mechanisms confer mesotrione resistance in MCR. Additionally, the basis for atrazine resistance was investigated in MCR and an atrazine-resistant but mesotrione-sensitive population (ACR for Adams County mesotrione-sensitive but atrazine-resistant). A standard sensitive population (WCS for Wayne County herbicide-sensitive) was also used for comparison. Mesotrione resistance was not due to an alteration in HPPD sequence, HPPD expression, or reduced herbicide absorption. Metabolism studies using whole plants and excised leaves revealed that the time for 50% of absorbed mesotrione to degrade in MCR was significantly shorter than in ACR and WCS, which correlated with previous phenotypic responses to mesotrione and the quantity of the metabolite 4-hydroxy-mesotrione in excised leaves. The cytochrome P450 monooxygenase inhibitors malathion and tetcyclacis significantly reduced mesotrione metabolism in MCR and corn (Zea mays) excised leaves but not in ACR. Furthermore, malathion increased mesotrione activity in MCR seedlings in greenhouse studies. These results indicate that enhanced oxidative metabolism contributes significantly to mesotrione resistance in MCR. Sequence analysis of atrazine-resistant (MCR and ACR) and atrazine-sensitive (WCS) waterhemp populations detected no differences in the psbA gene. The times for 50% of absorbed atrazine to degrade in corn, MCR, and ACR leaves were shorter than in WCS, and a polar metabolite of atrazine was detected in corn, MCR, and ACR that cochromatographed with a synthetic atrazine-glutathione conjugate. Thus, elevated rates of metabolism via distinct detoxification mechanisms contribute to mesotrione and atrazine resistance within the MCR population.Waterhemp (Amaranthus tuberculatus) is a troublesome annual weed species in midwestern U.S. corn (Zea mays) and soybean (Glycine max) production. The change to production systems with limited tillage has favored waterhemp germination and growth (Hager et al., 2002). Waterhemp seeds are small, and one female plant can produce up to one million seeds (Steckel et al., 2003), which endow waterhemp with an effective short-distance dispersal mechanism. In addition, multiple herbicide resistance mechanisms in waterhemp are facilitated by its dioecious biology and wind-pollinated flowers (Steckel, 2007). The long-distance flow of pollen may be one of the main reasons that multiple herbicide resistance in waterhemp has become widespread in the United States (Liu et al., 2012).Mesotrione (2-[4-(methylsulfonyl)-2-nitrobenzoyl]-1,3-cyclohexanedione) belongs to the triketone class of 4-hydroxyphenylpyruvate dioxygenase (HPPD)-inhibiting herbicides (Beaudegnies et al., 2009). Molecular information regarding plant HPPD gene sequences and expression patterns is limited (for review, see Pallett, 2000; Kim and Petersen, 2002; Riechers and Stanford, 2002; Matringe et al., 2005), and only a single expressed HPPD gene was detected in waterhemp (Riggins et al., 2010). Herbicidal activity of mesotrione in sensitive plants is due to competitive inhibition of the HPPD enzyme, which is a key enzyme in the biosynthesis of tocopherols and plastoquinone. Plastoquinone is an electron acceptor for the phytoene desaturase reaction in the pathway of carotenoid biosynthesis and also serves as an electron acceptor in PSII (Hess, 2000). Tocopherols and carotenoids are responsible for the detoxification of reactive oxygen species and scavenging of free radicals in plant tissues (Maeda and DellaPenna, 2007; Triantaphylidès and Havaux, 2009; Mène-Saffrané and DellaPenna, 2010), and carotenoids also protect chlorophyll from photooxidation (Cazzonelli and Pogson, 2010). Following mesotrione treatment, carotenoid biosynthesis is inhibited in sensitive plants, resulting in bleaching and necrosis. In particular, new leaves and meristems are primarily affected due to the need for protective carotenoids and tocopherols in photosynthetic tissues (Triantaphylidès and Havaux, 2009) and the systemic nature of mesotrione, which is translocated in the phloem (Mitchell et al., 2001; Beaudegnies et al., 2009).There are two main mechanisms of herbicide resistance in plants: (1) target site alterations, such as mutations that affect herbicide-binding kinetics or amplification of the target site gene (Powles and Yu, 2010), and (2) nontarget site mechanisms, including metabolism, translocation, and sequestration (Yuan et al., 2007; Powles and Yu, 2010). Metabolic detoxification is a common nontarget-based mechanism for herbicide resistance, which typically may result from elevated levels of cytochrome P450 monooxygenase (P450) or glutathione S-transferase (GST) activity (Powles and Yu, 2010; Délye et al., 2011). In addition to conferring resistance in weeds, these enzymes also confer natural tolerance in crops (Kreuz et al., 1996; Riechers et al., 2010). Similar to tolerant sorghum (Sorghum bicolor) lines (Abit and Al-Khatib, 2009), corn is tolerant to mesotrione via rapid metabolism (i.e. ring hydroxylation catalyzed by P450 activity) in combination with slower uptake relative to sensitive weeds and a less sensitive form of the HPPD enzyme in grasses relative to dicots (Hawkes et al., 2001; Mitchell et al., 2001).Atrazine (2-chloro-4-(ethylamino)-6-(isopropylamino)-S-triazine) is a symmetrical triazine herbicide commonly used in corn to selectively control annual dicot weeds. Atrazine disrupts electron transport by competing with plastoquinone for the secondary electron-accepting plastoquinone-binding site on the D1 protein of PSII in chloroplasts (Hess, 2000). Atrazine resistance in weeds can be due to a mutation in the psbA gene that causes a Ser-Gly substitution at amino acid position 264 of the D1 protein (Hirschberg and McIntosh, 1983; Devine and Preston, 2000). Corn and grain sorghum are naturally tolerant to atrazine via the rapid metabolism of atrazine through conjugation with reduced glutathione (GSH; Frear and Swanson, 1970; Lamoureux et al., 1973), which is catalyzed by GST activities (Shimabukuro et al., 1971). Enhanced metabolism of atrazine and simazine in weedy species has been reported in Abutilon theophrasti, Lolium rigidum, and Alopecurus myosuroides due to either GST- or P450-mediated detoxification mechanisms (Burnet et al., 1993; Gray et al., 1996; Cummins et al., 1999; Délye et al., 2011).A population of waterhemp (designated MCR for McLean County mesotrione- and atrazine-resistant) from Illinois is resistant to HPPD inhibitors (Hausman et al., 2011) and atrazine as well as to acetolactate synthase (ALS)-inhibiting herbicides. A different population of waterhemp (designated ACR for Adams County mesotrione-sensitive but atrazine-resistant; Patzoldt et al., 2005) that is atrazine resistant but sensitive to mesotrione (Hausman et al., 2011) and a waterhemp population (designated WCS for Wayne County herbicide-sensitive; Patzoldt et al., 2005) that is sensitive to both mesotrione and atrazine (Hausman et al., 2011) were used in comparison with MCR in this research. MCR displayed 10- and 35-fold resistance to mesotrione in comparison with ACR and WCS, respectively, in greenhouse studies (Hausman et al., 2011). In addition, waterhemp populations with similar patterns of multiple resistance have recently been identified (Hausman et al., 2011; McMullan and Green, 2011; Heap, 2012). However, the mechanisms of resistance to mesotrione and atrazine in these waterhemp populations are currently unknown. Therefore, the objective of this study was to determine if the multiple-herbicide-resistant phenotype of MCR (in regard to mesotrione and atrazine resistance) is due to either target site or nontarget site mechanisms.     

HAPLESS13, the Arabidopsis μ1 Adaptin,Is Essential for Protein Sorting at the trans-Golgi Network/Early Endosome

In plant cells, secretory and endocytic routes intersect at the trans-Golgi network (TGN)/early endosome (EE), where cargos are further sorted correctly and in a timely manner. Cargo sorting is essential for plant survival and therefore necessitates complex molecular machinery. Adaptor proteins (APs) play key roles in this process by recruiting coat proteins and selecting cargos for different vesicle carriers. The µ1 subunit of AP-1 in Arabidopsis (Arabidopsis thaliana) was recently identified at the TGN/EE and shown to be essential for cytokinesis. However, little was known about other cellular activities affected by mutations in AP-1 or the developmental consequences of such mutations. We report here that HAPLESS13 (HAP13), the Arabidopsis µ1 adaptin, is essential for protein sorting at the TGN/EE. Functional loss of HAP13 displayed pleiotropic developmental defects, some of which were suggestive of disrupted auxin signaling. Consistent with this, the asymmetric localization of PIN-FORMED2 (PIN2), an auxin transporter, was compromised in the mutant. In addition, cell morphogenesis was disrupted. We further demonstrate that HAP13 is critical for brefeldin A-sensitive but wortmannin-insensitive post-Golgi trafficking. Our results show that HAP13 is a key link in the sophisticated trafficking network in plant cells.Plant cells contain sophisticated endomembrane compartments, including the endoplasmic reticulum, the Golgi, the trans-Golgi network (TGN)/early endosome (EE), the prevacuolar compartments/multivesicular bodies (PVC/MVB), various types of vesicles, and the plasma membrane (PM; Ebine and Ueda, 2009; Richter et al., 2009). Intracellular protein sorting between the various locations in the endomembrane system occurs in both secretory and endocytic routes (Richter et al., 2009; De Marcos Lousa et al., 2012). Vesicles in the secretory route start at the endoplasmic reticulum, passing through the Golgi before reaching the TGN/EE, while vesicles in the endocytic route start from the PM before reaching the TGN/EE (Dhonukshe et al., 2007; Viotti et al., 2010). The TGN/EE in Arabidopsis (Arabidopsis thaliana) is an independent and highly dynamic organelle transiently associated with the Golgi (Dettmer et al., 2006; Lam et al., 2007; Viotti et al., 2010), distinct from the animal TGN. Once reaching the TGN/EE, proteins delivered by their vesicle carriers are subject to further sorting, being incorporated either into vesicles that pass through the PVC/MVB before reaching the vacuole for degradation or into vesicles that enter the secretory pathway for delivery to the PM (Ebine and Ueda, 2009; Richter et al., 2009). Therefore, the TGN/EE is a critical sorting compartment that lies at the intersection of the secretory and endocytic routes.Fine-tuned control of intracellular protein sorting at the TGN/EE is essential for plant development (Geldner et al., 2003; Dhonukshe et al., 2007, 2008; Richter et al., 2007; Kitakura et al., 2011; Wang et al., 2013). An auxin gradient is crucial for pattern formation in plants, whose dynamic maintenance requires the polar localization of auxin efflux carrier PINs through endocytic recycling (Geldner et al., 2003; Blilou et al., 2005; Paciorek et al., 2005; Abas et al., 2006; Jaillais et al., 2006; Dhonukshe et al., 2007; Kleine-Vehn et al., 2008). Receptor-like kinases (RLKs) have also been recognized as major cargos undergoing endocytic trafficking, which are either recycled back to the PM or sent for vacuolar degradation (Geldner and Robatzek, 2008; Irani and Russinova, 2009). RLKs are involved in most if not all developmental processes of plants (De Smet et al., 2009).Intracellular protein sorting relies on sorting signals within cargo proteins and on the molecular machinery that recognizes sorting signals (Boehm and Bonifacino, 2001; Robinson, 2004; Dhonukshe et al., 2007). Adaptor proteins (AP) play a key role (Boehm and Bonifacino, 2001; Robinson, 2004) in the recognition of sorting signals. APs are heterotetrameric protein complexes composed of two large subunits (β and γ/α/δ/ε), a small subunit (σ), and a medium subunit (µ) that is crucial for cargo selection (Boehm and Bonifacino, 2001). APs associate with the cytoplasmic side of secretory and endocytic vesicles, recruiting coat proteins and recognizing sorting signals within cargo proteins for their incorporation into vesicle carriers (Boehm and Bonifacino, 2001). Five APs have been identified so far, classified by their components, subcellular localization, and function (Boehm and Bonifacino, 2001; Robinson, 2004; Hirst et al., 2011). Of the five APs, AP-1 associates with the TGN or recycling endosomes (RE) in yeast and mammals (Huang et al., 2001; Robinson, 2004), mediating the sorting of cargo proteins to compartments of the endosomal-lysosomal system or to the basolateral PM of polarized epithelial cells (Gonzalez and Rodriguez-Boulan, 2009). Knockouts of AP-1 components in multicellular organisms resulted in embryonic lethality (Boehm and Bonifacino, 2001; Robinson, 2004).We show here that the recently identified Arabidopsis µ1 adaptin AP1M2 (Park et al., 2013; Teh et al., 2013) is a key component in the cellular machinery mediating intracellular protein sorting at the TGN/EE. AP1M2 was previously named HAPLESS13 (HAP13), whose mutant allele hap13 showed male gametophytic lethality (Johnson et al., 2004). In recent quests for AP-1 in plants, HAP13/AP1M2 was confirmed as the Arabidopsis µ1 adaptin based on its interaction with other components of the AP-1 complex as well as its localization at the TGN (Park et al., 2013; Teh et al., 2013). A novel mutant allele of HAP13/AP1M2, ap1m2-1, was found to be defective in the intracellular distribution of KNOLLE, leading to defective cytokinesis (Park et al., 2013; Teh et al., 2013). However, it was not clear whether HAP13/AP1M2 mediated other cellular activities and their developmental consequences. Using the same mutant allele, we found that functional loss of HAP13 (hap13-1/ap1m2-1) resulted in a full spectrum of growth defects, suggestive of compromised auxin signaling and of defective RLK signaling. Cell morphogenesis was also disturbed in hap13-1. Importantly, hap13-1 was insensitive to brefeldin A (BFA) washout, indicative of defects in guanine nucleotide exchange factors for ADP-ribosylation factor (ArfGEF)-mediated post-Golgi trafficking. Furthermore, HAP13/AP1M2 showed evolutionarily conserved function during vacuolar fusion, providing additional support to its identity as a µ1 adaptin. These results demonstrate the importance of the Arabidopsis µ1 adaptin for intracellular protein sorting centered on the TGN/EE.