Decreased Mitochondrial Activities of Malate Dehydrogenase and Fumarase in Tomato Lead to Altered Root Growth and Architecture via Diverse Mechanisms

Transgenic tomato (Solanum lycopersicum) plants in which either mitochondrial malate dehydrogenase or fumarase was antisense inhibited have previously been characterized to exhibit altered photosynthetic metabolism. Here, we demonstrate that these manipulations also resulted in differences in root growth, with both transgenics being characterized by a dramatic reduction of root dry matter deposition and respiratory activity but opposite changes with respect to root area. A range of physiological, molecular, and biochemical experiments were carried out in order to determine whether changes in root morphology were due to altered metabolism within the root itself, alterations in the nature of the transformants'' root exudation, consequences of alteration in the efficiency of photoassimilate delivery to the root, or a combination of these factors. Grafting experiments in which the transformants were reciprocally grafted to wild-type controls suggested that root length and area were determined by the aerial part of the plant but that biomass was not. Despite the transgenic roots displaying alteration in the expression of phytohormone-associated genes, evaluation of the levels of the hormones themselves revealed that, with the exception of gibberellins, they were largely unaltered. When taken together, these combined experiments suggest that root biomass and growth are retarded by root-specific alterations in metabolism and gibberellin contents. These data are discussed in the context of current models of root growth and biomass partitioning.The structure of the plant tricarboxylic acid (TCA) cycle has been established for decades (Beevers, 1961), and in vitro studies have established regulatory properties of many of its component enzymes (Budde and Randall, 1990; Millar and Leaver, 2000; Studart-Guimarães et al., 2005). That said, relatively little is known, as yet, regarding how this important pathway is regulated in vivo (Fernie et al., 2004a; Sweetlove et al., 2007). Indeed, even fundamental questions concerning the degree to which this pathway operates in illuminated leaves (Tcherkez et al., 2005; Nunes-Nesi et al., 2007a) and the influence it has on organic acid levels in fruits (Burger et al., 2003) remain contentious. Furthermore, in contrast to many other pathways of primary metabolism, the TCA cycle has been subjected to relatively few molecular physiological studies. To date, the functions of pyruvate dehydrogenase, citrate synthase, aconitase, isocitrate dehydrogenase, succinyl-CoA ligase, fumarase, and malate dehydrogenase have been studied via this approach (Landschütze et al., 1995; Carrari et al., 2003; Yui et al., 2003; Nunes-Nesi et al., 2005, 2007a; Lemaitre et al., 2007; Studart-Guimarães et al., 2007); however, several of these studies were relatively cursory. Despite this fact, they generally corroborate one another, with at least two studies providing clear evidence for an important role of the TCA cycle in flower development (Landschütze et al., 1995; Yui et al., 2003) or in the coordination of photosynthetic and respiratory metabolisms of the illuminated leaf (Carrari et al., 2003; Nunes-Nesi et al., 2005, 2007a).In our own studies on tomato (Solanum lycopersicum), we have observed that modulation of fumarase and mitochondrial malate dehydrogenase activities leads to contrasting shoot phenotypes, with the former displaying stunted growth while the later exhibited an enhanced photosynthetic performance (Nunes-Nesi et al., 2005, 2007a). We were able to demonstrate that the stunted-growth phenotype observed in aerial parts of the fumarase plants was a consequence of altered stomatal function (Nunes-Nesi et al., 2007a), whereas the increased photosynthetic performance of the mitochondrial malate dehydrogenase seems likely to be mediated by the alterations in ascorbate metabolism exhibited by these plants (Nunes-Nesi et al., 2005; Urbanczyk-Wochniak et al., 2006). In keeping with the altered rates of photosynthesis in these antisense plants, the fruit yield of fumarase and mitochondrial malate dehydrogenase plants was decreased and increased, respectively. However, the root biomass of both transgenics was significantly reduced (Nunes-Nesi et al., 2005, 2007a). These observations were somewhat surprising given that it is estimated that 30% to 60% of net photosynthate is transported to root organs (Merckx et al., 1986; Nguyen et al., 1999; Singer et al., 2003). When taken together, these results suggest that the root phenotype must result from either an impairment of translocation or a root-specific effect. Neither of these explanations is without precedence, with inhibition of the expression of Suc transporters (Riesmeier et al., 1993; Gottwald et al., 2000) resulting in dramatically impaired root growth while organic acid exudation itself has been implicated in a wide range of root organ functions, including nutrient acquisition (de la Fuente et al., 1997; Imas et al., 1997; Neumann and Römheld, 1999; López-Bucio et al., 2000; Anoop et al., 2003; Delhaize et al., 2004), metal sequestration (Gillooly et al., 1983; de la Fuente et al., 1997; Cramer and Titus, 2001), and microbial proliferation in the rhizosphere (Lugtenberg et al., 1999; Weisskopf et al., 2005). In addition to the putative mechanisms listed above, the TCA cycle could be anticipated to play a vital role in meeting the high energy demands of nitrogen fixation and polymer biosynthesis associated with rapidly growing heterotrophic organs (Pradet and Raymond, 1983; Dieuaide-Noubhani et al., 1997; Stasolla et al., 2003; Deuschle et al., 2006). In keeping with this theory, alteration of the energy status of roots and other heterotrophic tissue has been documented to positively correlate with elevated biomass production (Anekonda, 2001; Regierer et al., 2002; Carrari et al., 2003; Lovas et al., 2003; Geigenberger et al., 2005). Here, we performed a detailed physiological, molecular, and biochemical evaluation of whole plant and root metabolism of the mitochondrial malate dehydrogenase and fumarate antisense tomato lines. In this manner, we broadly assessed biochemical changes in the root, including the levels of several major phytohormones, as well as dissected which characteristics were influenced by aerial parts of the plant. The results obtained are discussed both with respect to the regulation of the TCA cycle per se and within the context of the determination of root morphology and growth.     

Calcineurin B-Like Protein-Interacting Protein Kinase CIPK21 Regulates Osmotic and Salt Stress Responses in Arabidopsis

The role of calcium-mediated signaling has been extensively studied in plant responses to abiotic stress signals. Calcineurin B-like proteins (CBLs) and CBL-interacting protein kinases (CIPKs) constitute a complex signaling network acting in diverse plant stress responses. Osmotic stress imposed by soil salinity and drought is a major abiotic stress that impedes plant growth and development and involves calcium-signaling processes. In this study, we report the functional analysis of CIPK21, an Arabidopsis (Arabidopsis thaliana) CBL-interacting protein kinase, ubiquitously expressed in plant tissues and up-regulated under multiple abiotic stress conditions. The growth of a loss-of-function mutant of CIPK21, cipk21, was hypersensitive to high salt and osmotic stress conditions. The calcium sensors CBL2 and CBL3 were found to physically interact with CIPK21 and target this kinase to the tonoplast. Moreover, preferential localization of CIPK21 to the tonoplast was detected under salt stress condition when coexpressed with CBL2 or CBL3. These findings suggest that CIPK21 mediates responses to salt stress condition in Arabidopsis, at least in part, by regulating ion and water homeostasis across the vacuolar membranes.Drought and salinity cause osmotic stress in plants and severely affect crop productivity throughout the world. Plants respond to osmotic stress by changing a number of cellular processes (Xiong et al., 1999; Xiong and Zhu, 2002; Bartels and Sunkar, 2005; Boudsocq and Lauriére, 2005). Some of these changes include activation of stress-responsive genes, regulation of membrane transport at both plasma membrane (PM) and vacuolar membrane (tonoplast) to maintain water and ionic homeostasis, and metabolic changes to produce compatible osmolytes such as Pro (Stewart and Lee, 1974; Krasensky and Jonak, 2012). It has been well established that a specific calcium (Ca²⁺) signature is generated in response to a particular environmental stimulus (Trewavas and Malhó, 1998; Scrase-Field and Knight, 2003; Luan, 2009; Kudla et al., 2010). The Ca²⁺ changes are primarily perceived by several Ca²⁺ sensors such as calmodulin (Reddy, 2001; Luan et al., 2002), Ca²⁺-dependent protein kinases (Harper and Harmon, 2005), calcineurin B-like proteins (CBLs; Luan et al., 2002; Batistič and Kudla, 2004; Pandey, 2008; Luan, 2009; Sanyal et al., 2015), and other Ca²⁺-binding proteins (Reddy, 2001; Shao et al., 2008) to initiate various cellular responses.Plant CBL-type Ca²⁺ sensors interact with and activate CBL-interacting protein kinases (CIPKs) that phosphorylate downstream components to transduce Ca²⁺ signals (Liu et al., 2000; Luan et al., 2002; Batistič and Kudla, 2004; Luan, 2009). In several plant species, multiple members have been identified in the CBL and CIPK family (Luan et al., 2002; Kolukisaoglu et al., 2004; Pandey, 2008; Batistič and Kudla, 2009; Weinl and Kudla, 2009; Pandey et al., 2014). Involvement of specific CBL-CIPK pair to decode a particular type of signal entails the alternative and selective complex formation leading to stimulus-response coupling (D’Angelo et al., 2006; Batistič et al., 2010).Several CBL and CIPK family members have been implicated in plant responses to drought, salinity, and osmotic stress based on genetic analysis of Arabidopsis (Arabidopsis thaliana) mutants (Zhu, 2002; Cheong et al., 2003, 2007; Kim et al., 2003; Pandey et al., 2004, 2008; D’Angelo et al., 2006; Qin et al., 2008; Tripathi et al., 2009; Held et al., 2011; Tang et al., 2012; Drerup et al., 2013; Eckert et al., 2014). A few CIPKs have also been functionally characterized by gain-of-function approach in crop plants such as rice (Oryza sativa), pea (Pisum sativum), and maize (Zea mays) and were found to be involved in osmotic stress responses (Mahajan et al., 2006; Xiang et al., 2007; Yang et al., 2008; Tripathi et al., 2009; Zhao et al., 2009; Cuéllar et al., 2010).In this report, we examined the role of the Arabidopsis CIPK21 gene in osmotic stress response by reverse genetic analysis. The loss-of-function mutant plants became hypersensitive to salt and mannitol stress conditions, suggesting that CIPK21 is involved in the regulation of osmotic stress response in Arabidopsis. These findings are further supported by an enhanced tonoplast targeting of the cytoplasmic CIPK21 through interaction with the vacuolar Ca²⁺ sensors CBL2 and CBL3 under salt stress condition.     

Characterization of the Possible Roles for B Class MADS Box Genes in Regulation of Perianth Formation in Orchid

To investigate sepal/petal/lip formation in Oncidium Gower Ramsey, three paleoAPETALA3 genes, O. Gower Ramsey MADS box gene5 (OMADS5; clade 1), OMADS3 (clade 2), and OMADS9 (clade 3), and one PISTILLATA gene, OMADS8, were characterized. The OMADS8 and OMADS3 mRNAs were expressed in all four floral organs as well as in vegetative leaves. The OMADS9 mRNA was only strongly detected in petals and lips. The mRNA for OMADS5 was only strongly detected in sepals and petals and was significantly down-regulated in lip-like petals and lip-like sepals of peloric mutant flowers. This result revealed a possible negative role for OMADS5 in regulating lip formation. Yeast two-hybrid analysis indicated that OMADS5 formed homodimers and heterodimers with OMADS3 and OMADS9. OMADS8 only formed heterodimers with OMADS3, whereas OMADS3 and OMADS9 formed homodimers and heterodimers with each other. We proposed that sepal/petal/lip formation needs the presence of OMADS3/8 and/or OMADS9. The determination of the final organ identity for the sepal/petal/lip likely depended on the presence or absence of OMADS5. The presence of OMADS5 caused short sepal/petal formation. When OMADS5 was absent, cells could proliferate, resulting in the possible formation of large lips and the conversion of the sepal/petal into lips in peloric mutants. Further analysis indicated that only ectopic expression of OMADS8 but not OMADS5/9 caused the conversion of the sepal into an expanded petal-like structure in transgenic Arabidopsis (Arabidopsis thaliana) plants.The ABCDE model predicts the formation of any flower organ by the interaction of five classes of homeotic genes in plants (Yanofsky et al., 1990; Jack et al., 1992; Mandel et al., 1992; Goto and Meyerowitz, 1994; Jofuku et al., 1994; Pelaz et al., 2000, 2001; Theißen and Saedler, 2001; Pinyopich et al., 2003; Ditta et al., 2004; Jack, 2004). The A class genes control sepal formation. The A, B, and E class genes work together to regulate petal formation. The B, C, and E class genes control stamen formation. The C and E class genes work to regulate carpel formation, whereas the D class gene is involved in ovule development. MADS box genes seem to have a central role in flower development, because most ABCDE genes encode MADS box proteins (Coen and Meyerowitz, 1991; Weigel and Meyerowitz, 1994; Purugganan et al., 1995; Rounsley et al., 1995; Theißen and Saedler, 1995; Theißen et al., 2000; Theißen, 2001).The function of B group genes, such as APETALA3 (AP3) and PISTILLATA (PI), has been thought to have a major role in specifying petal and stamen development (Jack et al., 1992; Goto and Meyerowitz, 1994; Krizek and Meyerowitz, 1996; Kramer et al., 1998; Hernandez-Hernandez et al., 2007; Kanno et al., 2007; Whipple et al., 2007; Irish, 2009). In Arabidopsis (Arabidopsis thaliana), mutation in AP3 or PI caused identical phenotypes of second whorl petal conversion into a sepal structure and third flower whorl stamen into a carpel structure (Bowman et al., 1989; Jack et al., 1992; Goto and Meyerowitz, 1994). Similar homeotic conversions for petal and stamen were observed in the mutants of the AP3 and PI orthologs from a number of core eudicots such as Antirrhinum majus, Petunia hybrida, Gerbera hybrida, Solanum lycopersicum, and Nicotiana benthamiana (Sommer et al., 1990; Tröbner et al., 1992; Angenent et al., 1993; van der Krol et al., 1993; Yu et al., 1999; Liu et al., 2004; Vandenbussche et al., 2004; de Martino et al., 2006), from basal eudicot species such as Papaver somniferum and Aquilegia vulgaris (Drea et al., 2007; Kramer et al., 2007), as well as from monocot species such as Zea mays and Oryza sativa (Ambrose et al., 2000; Nagasawa et al., 2003; Prasad and Vijayraghavan, 2003; Yadav et al., 2007; Yao et al., 2008). This indicated that the function of the B class genes AP3 and PI is highly conserved during evolution.It has been thought that B group genes may have arisen from an ancestral gene through multiple gene duplication events (Doyle, 1994; Theißen et al., 1996, 2000; Purugganan, 1997; Kramer et al., 1998; Kramer and Irish, 1999; Lamb and Irish, 2003; Kim et al., 2004; Stellari et al., 2004; Zahn et al., 2005; Hernandez-Hernandez et al., 2007). In the gymnosperms, there was a single putative B class lineage that duplicated to generate the paleoAP3 and PI lineages in angiosperms (Kramer et al., 1998; Theißen et al., 2000; Irish, 2009). The paleoAP3 lineage is composed of AP3 orthologs identified in lower eudicots, magnolid dicots, and monocots (Kramer et al., 1998). Genes in this lineage contain the conserved paleoAP3- and PI-derived motifs in the C-terminal end of the proteins, which have been thought to be characteristics of the B class ancestral gene (Kramer et al., 1998; Tzeng and Yang, 2001; Hsu and Yang, 2002). The PI lineage is composed of PI orthologs that contain a highly conserved PI motif identified in most plant species (Kramer et al., 1998). Subsequently, there was a second duplication at the base of the core eudicots that produced the euAP3 and TM6 lineages, which have been subject to substantial sequence changes in eudicots during evolution (Kramer et al., 1998; Kramer and Irish, 1999). The paleoAP3 motif in the C-terminal end of the proteins was retained in the TM6 lineage and replaced by a conserved euAP3 motif in the euAP3 lineage of most eudicot species (Kramer et al., 1998). In addition, many lineage-specific duplications for paleoAP3 lineage have occurred in plants such as orchids (Hsu and Yang, 2002; Tsai et al., 2004; Kim et al., 2007; Mondragón-Palomino and Theißen, 2008, 2009; Mondragón-Palomino et al., 2009), Ranunculaceae, and Ranunculales (Kramer et al., 2003; Di Stilio et al., 2005; Shan et al., 2006; Kramer, 2009).Unlike the A or C class MADS box proteins, which form homodimers that regulate flower development, the ability of B class proteins to form homodimers has only been reported in gymnosperms and in the paleoAP3 and PI lineages of some monocots. For example, LMADS1 of the lily Lilium longiflorum (Tzeng and Yang, 2001), OMADS3 of the orchid Oncidium Gower Ramsey (Hsu and Yang, 2002), and PeMADS4 of the orchid Phalaenopsis equestris (Tsai et al., 2004) in the paleoAP3 lineage, LRGLOA and LRGLOB of the lily Lilium regale (Winter et al., 2002), TGGLO of the tulip Tulipa gesneriana (Kanno et al., 2003), and PeMADS6 of the orchid P. equestris (Tsai et al., 2005) in the PI lineage, and GGM2 of the gymnosperm Gnetum gnemon (Winter et al., 1999) were able to form homodimers that regulate flower development. Proteins in the euAP3 lineage and in most paleoAP3 lineages were not able to form homodimers and had to interact with PI to form heterodimers in order to regulate petal and stamen development in various plant species (Schwarz-Sommer et al., 1992; Tröbner et al., 1992; Riechmann et al., 1996; Moon et al., 1999; Winter et al., 2002; Kanno et al., 2003; Vandenbussche et al., 2004; Yao et al., 2008). In addition to forming dimers, AP3 and PI were able to interact with other MADS box proteins, such as SEPALLATA1 (SEP1), SEP2, and SEP3, to regulate petal and stamen development (Pelaz et al., 2000; Honma and Goto, 2001; Theißen and Saedler, 2001; Castillejo et al., 2005).Orchids are among the most important plants in the flower market around the world, and research on MADS box genes has been reported for several species of orchids during the past few years (Lu et al., 1993, 2007; Yu and Goh, 2000; Hsu and Yang, 2002; Yu et al., 2002; Hsu et al., 2003; Tsai et al., 2004, 2008; Xu et al., 2006; Guo et al., 2007; Kim et al., 2007; Chang et al., 2009). Unlike the flowers in eudicots, the nearly identical shape of the sepals and petals as well as the production of a unique lip in orchid flowers make them a very special plant species for the study of flower development. Four clades (1–4) of genes in the paleoAP3 lineage have been identified in several orchids (Hsu and Yang, 2002; Tsai et al., 2004; Kim et al., 2007; Mondragón-Palomino and Theißen, 2008, 2009; Mondragón-Palomino et al., 2009). Several works have described the possible interactions among these four clades of paleoAP3 genes and one PI gene that are involved in regulating the differentiation and formation of the sepal/petal/lip of orchids (Tsai et al., 2004; Kim et al., 2007; Mondragón-Palomino and Theißen, 2008, 2009). However, the exact mechanism that involves the orchid B class genes remains unclear and needs to be clarified by more experimental investigations.O. Gower Ramsey is a popular orchid with important economic value in cut flower markets. Only a few studies have been reported on the role of MADS box genes in regulating flower formation in this plant species (Hsu and Yang, 2002; Hsu et al., 2003; Chang et al., 2009). An AP3-like MADS gene that regulates both floral formation and initiation in transgenic Arabidopsis has been reported (Hsu and Yang, 2002). In addition, four AP1/AGAMOUS-LIKE9 (AGL9)-like MADS box genes have been characterized that show novel expression patterns and cause different effects on floral transition and formation in Arabidopsis (Hsu et al., 2003; Chang et al., 2009). Compared with other orchids, the production of a large and well-expanded lip and five small identical sepals/petals makes O. Gower Ramsey a special case for the study of the diverse functions of B class MADS box genes during evolution. Therefore, the isolation of more B class MADS box genes and further study of their roles in the regulation of perianth (sepal/petal/lip) formation during O. Gower Ramsey flower development are necessary. In addition to the clade 2 paleoAP3 gene OMADS3, which was previously characterized in our laboratory (Hsu and Yang, 2002), three more B class MADS box genes, OMADS5, OMADS8, and OMADS9, were characterized from O. Gower Ramsey in this study. Based on the different expression patterns and the protein interactions among these four orchid B class genes, we propose that the presence of OMADS3/8 and/or OMADS9 is required for sepal/petal/lip formation. Further sepal and petal formation at least requires the additional presence of OMADS5, whereas large lip formation was seen when OMADS5 expression was absent. Our results provide a new finding and information pertaining to the roles for orchid B class MADS box genes in the regulation of sepal/petal/lip formation.     

PLASTID MOVEMENT IMPAIRED1 and PLASTID MOVEMENT IMPAIRED1-RELATED1 Mediate Photorelocation Movements of Both Chloroplasts and Nuclei

Organelle movement and positioning play important roles in fundamental cellular activities and adaptive responses to environmental stress in plants. To optimize photosynthetic light utilization, chloroplasts move toward weak blue light (the accumulation response) and escape from strong blue light (the avoidance response). Nuclei also move in response to strong blue light by utilizing the light-induced movement of attached plastids in leaf cells. Blue light receptor phototropins and several factors for chloroplast photorelocation movement have been identified through molecular genetic analysis of Arabidopsis (Arabidopsis thaliana). PLASTID MOVEMENT IMPAIRED1 (PMI1) is a plant-specific C2-domain protein that is required for efficient chloroplast photorelocation movement. There are two PLASTID MOVEMENT IMPAIRED1-RELATED (PMIR) genes, PMIR1 and PMIR2, in the Arabidopsis genome. However, the mechanism in which PMI1 regulates chloroplast and nuclear photorelocation movements and the involvement of PMIR1 and PMIR2 in these organelle movements remained unknown. Here, we analyzed chloroplast and nuclear photorelocation movements in mutant lines of PMI1, PMIR1, and PMIR2. In mesophyll cells, the pmi1 single mutant showed severe defects in both chloroplast and nuclear photorelocation movements resulting from the impaired regulation of chloroplast-actin filaments. In pavement cells, pmi1 mutant plants were partially defective in both plastid and nuclear photorelocation movements, but pmi1pmir1 and pmi1pmir1pmir2 mutant lines lacked the blue light-induced movement responses of plastids and nuclei completely. These results indicated that PMI1 is essential for chloroplast and nuclear photorelocation movements in mesophyll cells and that both PMI1 and PMIR1 are indispensable for photorelocation movements of plastids and thus, nuclei in pavement cells.In plants, organelles move within the cell and become appropriately positioned to accomplish their functions and adapt to the environment (for review, see Wada and Suetsugu, 2004). Light-induced chloroplast movement (chloroplast photorelocation movement) is one of the best characterized organelle movements in plants (Suetsugu and Wada, 2012). Under weak light conditions, chloroplasts move toward light to capture light efficiently (the accumulation response; Zurzycki, 1955). Under strong light conditions, chloroplasts escape from light to avoid photodamage (the avoidance response; Kasahara et al., 2002; Sztatelman et al., 2010; Davis and Hangarter, 2012; Cazzaniga et al., 2013). In most green plant species, these responses are induced primarily by the blue light receptor phototropin (phot) in response to a range of wavelengths from UVA to blue light (approximately 320–500 nm; for review, see Suetsugu and Wada, 2012; Wada and Suetsugu, 2013; Kong and Wada, 2014). Phot-mediated chloroplast movement has been shown in land plants, such as Arabidopsis (Arabidopsis thaliana; Jarillo et al., 2001; Kagawa et al., 2001; Sakai et al., 2001), the fern Adiantum capillus-veneris (Kagawa et al., 2004), the moss Physcomitrella patens (Kasahara et al., 2004), and the liverwort Marchantia polymorpha (Komatsu et al., 2014). Two phots in Arabidopsis, phot1 and phot2, redundantly mediate the accumulation response (Sakai et al., 2001), whereas phot2 primarily regulates the avoidance response (Jarillo et al., 2001; Kagawa et al., 2001; Luesse et al., 2010). M. polymorpha has only one phot that mediates both the accumulation and avoidance responses (Komatsu et al., 2014), although two or more phots mediate chloroplast photorelocation movement in A. capillus-veneris (Kagawa et al., 2004) and P. patens (Kasahara et al., 2004). Thus, duplication and functional diversification of PHOT genes have occurred during land plant evolution, and plants have gained a sophisticated light sensing system for chloroplast photorelocation movement.In general, movements of plant organelles, including chloroplasts, are dependent on actin filaments (for review, see Wada and Suetsugu, 2004). Most organelles common in eukaryotes, such as mitochondria, peroxisomes, and Golgi bodies, use the myosin motor for their movements, but there is no clear evidence that chloroplast movement is myosin dependent (for review, see Suetsugu et al., 2010a). Land plants have innovated a novel actin-based motility system that is specialized for chloroplast movement as well as a photoreceptor system (for review, see Suetsugu et al., 2010a; Wada and Suetsugu, 2013; Kong and Wada, 2014). Chloroplast-actin (cp-actin) filaments, which were first found in Arabidopsis, are short actin filaments specifically localized around the chloroplast periphery at the interface between the chloroplast and the plasma membrane (Kadota et al., 2009). Strong blue light induces the rapid disappearance of cp-actin filaments and then, their subsequent reappearance preferentially at the front region of the moving chloroplasts. This asymmetric distribution of cp-actin filaments is essential for directional chloroplast movement (Kadota et al., 2009; Kong et al., 2013a). The greater the difference in the amount of cp-actin filaments between the front and rear regions of chloroplasts becomes, the faster the chloroplasts move, in which the magnitude of the difference is determined by fluence rate (Kagawa and Wada, 2004; Kadota et al., 2009; Kong et al., 2013a). Strong blue light-induced disappearance of cp-actin filaments is regulated in a phot2-dependent manner before the intensive polymerization of cp-actin filaments at the front region occurs (Kadota et al., 2009; Ichikawa et al., 2011; Kong et al., 2013a). This phot2-dependent response contributes to the greater difference in the amount of cp-actin filaments between the front and rear regions of chloroplasts. Similar behavior of cp-actin filaments has also been observed in A. capillus-veneris (Tsuboi and Wada, 2012) and P. patens (Yamashita et al., 2011).Like chloroplasts, nuclei also show light-mediated movement and positioning (nuclear photorelocation movement) in land plants (for review, see Higa et al., 2014b). In gametophytic cells of A. capillus-veneris, weak light induced the accumulation responses of both chloroplasts and nuclei, whereas strong light induced avoidance responses (Kagawa and Wada, 1993, 1995; Tsuboi et al., 2007). However, in mesophyll cells of Arabidopsis, strong blue light induced both chloroplast and nuclear avoidance responses, but weak blue light induced only the chloroplast accumulation response (Iwabuchi et al., 2007, 2010; Higa et al., 2014a). In Arabidopsis pavement cells, small numbers of tiny plastids were found and showed autofluorescence under the confocal laser-scanning microscopy (Iwabuchi et al., 2010; Higa et al., 2014a). Hereafter, the plastid in the pavement cells is called the pavement cell plastid. Strong blue light-induced avoidance responses of pavement cell plastids and nuclei were induced in a phot2-dependent manner, but the accumulation response was not detected for either organelle (Iwabuchi et al., 2007, 2010; Higa et al., 2014a). In both Arabidopsis and A. capillus-veneris, phots mediate nuclear photorelocation movement, and phot2 mediates the nuclear avoidance response (Iwabuchi et al., 2007, 2010; Tsuboi et al., 2007). The nuclear avoidance response is dependent on actin filaments in both mesophyll and pavement cells of Arabidopsis (Iwabuchi et al., 2010). Recently, it was shown that the nuclear avoidance response relies on cp-actin-dependent movement of pavement cell plastids, where nuclei are associated with pavement cell plastids of Arabidopsis (Higa et al., 2014a). In mesophyll cells, nuclear avoidance response is likely dependent on cp-actin filament-mediated chloroplast movement, because the mutants deficient in chloroplast movement were also defective in nuclear avoidance response (Higa et al., 2014a). Thus, phots mediate both chloroplast (and pavement cell plastid) and nuclear photorelocation movement by regulating cp-actin filaments.Molecular genetic analyses of Arabidopsis mutants deficient in chloroplast photorelocation movement have identified many molecular factors involved in signal transduction and/or motility systems as well as those involved in the photoreceptor system for chloroplast photorelocation movement (and thus, nuclear photorelocation movement; for review, see Suetsugu and Wada, 2012; Wada and Suetsugu, 2013; Kong and Wada, 2014). CHLOROPLAST UNUSUAL POSITIONING1 (CHUP1; Oikawa et al., 2003) and KINESIN-LIKE PROTEIN FOR ACTIN-BASED CHLOROPLAST MOVEMENT (KAC; Suetsugu et al., 2010b) are key factors for generating and/or maintaining cp-actin filaments. Both proteins are highly conserved in land plants and essential for the movement and attachment of chloroplasts to the plasma membrane in Arabidopsis (Oikawa et al., 2003, 2008; Suetsugu et al., 2010b), A. capillus-veneris (Suetsugu et al., 2012), and P. patens (Suetsugu et al., 2012; Usami et al., 2012). CHUP1 is localized on the chloroplast outer membrane and binds to globular and filamentous actins and profilin in vitro (Oikawa et al., 2003, 2008; Schmidt von Braun and Schleiff, 2008). Although KAC is a kinesin-like protein, it lacks microtubule-dependent motor activity but has filamentous actin binding activity (Suetsugu et al., 2010b). An actin-bundling protein THRUMIN1 (THRUM1) is required for efficient chloroplast photorelocation movement (Whippo et al., 2011) and interacts with cp-actin filaments (Kong et al., 2013a). chup1 and kac mutant plants were shown to lack detectable cp-actin filaments (Kadota et al., 2009; Suetsugu et al., 2010b; Ichikawa et al., 2011; Kong et al., 2013a). Similarly, cp-actin filaments were rarely detected in thrum1 mutant plants (Kong et al., 2013a), indicating that THRUM1 also plays an important role in maintaining cp-actin filaments.Other proteins J-DOMAIN PROTEIN REQUIRED FOR CHLOROPLAST ACCUMULATION RESPONSE1 (JAC1; Suetsugu et al., 2005), WEAK CHLOROPLAST MOVEMENT UNDER BLUE LIGHT1 (WEB1; Kodama et al., 2010), and PLASTID MOVEMENT IMPAIRED2 (PMI2; Luesse et al., 2006; Kodama et al., 2010) are involved in the light regulation of cp-actin filaments and chloroplast photorelocation movement. JAC1 is an auxilin-like J-domain protein that mediates the chloroplast accumulation response through its J-domain function (Suetsugu et al., 2005; Takano et al., 2010). WEB1 and PMI2 are coiled-coil proteins that interact with each other (Kodama et al., 2010). Although web1 and pmi2 were partially defective in the avoidance response, the jac1 mutation completely suppressed the phenotype of web1 and pmi2, suggesting that the WEB1/PMI2 complex suppresses JAC1 function (i.e. the accumulation response) under strong light conditions (Kodama et al., 2010). Both web1 and pmi2 showed impaired disappearance of cp-actin filaments in response to strong blue light (Kodama et al., 2010). However, the exact molecular functions of these proteins are unknown.In this study, we characterized mutant plants deficient in the PMI1 gene and two homologous genes PLASTID MOVEMENT IMPAIRED1-RELATED1 (PMIR1) and PMIR2. PMI1 was identified through molecular genetic analyses of pmi1 mutants that showed severe defects in chloroplast accumulation and avoidance responses (DeBlasio et al., 2005). PMI1 is a plant-specific C2-domain protein (DeBlasio et al., 2005; Zhang and Aravind, 2010), but its roles and those of PMIRs in cp-actin-mediated chloroplast and nuclear photorelocation movements remained unclear. Thus, we analyzed chloroplast and nuclear photorelocation movements in the single, double, and triple mutants of pmi1, pmir1, and pmir2.     

Plastidial NAD-Dependent Malate Dehydrogenase Is Critical for Embryo Development and Heterotrophic Metabolism in Arabidopsis

In illuminated chloroplasts, one mechanism involved in reduction-oxidation (redox) homeostasis is the malate-oxaloacetate (OAA) shuttle. Excess electrons from photosynthetic electron transport in the form of nicotinamide adenine dinucleotide phosphate, reduced are used by NADP-dependent malate dehydrogenase (MDH) to reduce OAA to malate, thus regenerating the electron acceptor NADP. NADP-MDH is a strictly redox-regulated, light-activated enzyme that is inactive in the dark. In the dark or in nonphotosynthetic tissues, the malate-OAA shuttle was proposed to be mediated by the constitutively active plastidial NAD-specific MDH isoform (pdNAD-MDH), but evidence is scarce. Here, we reveal the critical role of pdNAD-MDH in Arabidopsis (Arabidopsis thaliana) plants. A pdnad-mdh null mutation is embryo lethal. Plants with reduced pdNAD-MDH levels by means of artificial microRNA (miR-mdh-1) are viable, but dark metabolism is altered as reflected by increased nighttime malate, starch, and glutathione levels and a reduced respiration rate. In addition, miR-mdh-1 plants exhibit strong pleiotropic effects, including dwarfism, reductions in chlorophyll levels, photosynthetic rate, and daytime carbohydrate levels, and disordered chloroplast ultrastructure, particularly in developing leaves, compared with the wild type. pdNAD-MDH deficiency in miR-mdh-1 can be functionally complemented by expression of a microRNA-insensitive pdNAD-MDH but not NADP-MDH, confirming distinct roles for NAD- and NADP-linked redox homeostasis.Reduction-oxidation (redox) reactions play pivotal roles for most metabolic processes and occur in all cellular compartments. The origin of all reducing power in plants is the chloroplast thylakoid membrane system, where light-driven photosynthetic electron transport leads to the coupled formation of ATP and the reducing equivalent NADPH (Dietz and Pfannschmidt, 2011). Sudden changes in light intensity and withdrawal of ATP and NADPH for biosynthetic processes in varying amounts can potentially disturb the ATP:NADPH ratio. Maintaining this ratio within certain limits, however, is crucial for plant metabolism, because it avoids the accumulation of excess electrons and the production of cytotoxic reactive oxygen species and allows for the continued production of ATP (Apel and Hirt, 2004; Logan, 2006; Scheibe and Dietz, 2012). Accordingly, plants have several mechanisms to dissipate excess electrons, avoid damage to cellular components, and maintain redox homeostasis. These mechanisms include nonphotochemical energy quenching, chlororespiration, cyclic electron transport, and the Mehler reaction (Scheibe et al., 2005).Reducing equivalents in the form of dedicated electron carriers or reduced cofactors (e.g. ferredoxin and NADH) are not generally transported directly across membranes; however, they can be shuttled indirectly as malate in exchange for oxaloacetic acid (OAA). This redox-poising mechanism is known as the malate valve in illuminated plastids or more generally, the malate-OAA shuttle (Heber, 1974; Scheibe, 2004; Taniguchi and Miyake, 2012). The key enzyme of the malate-OAA shuttle is malate dehydrogenase (MDH), which catalyses the reversible interconversion of malate and OAA. Isoforms of MDH are present in various cell compartments (Gietl, 1992), and each isoform is specific to either cosubstrate NAD (NAD-MDH; EC 1.1.1.37) or NADP (NADP-MDH; EC 1.1.1.82). The Arabidopsis genome encodes eight putative NAD-MDH isoforms: two isoforms are peroxisomal MDH (PMDH; PMDH1 and PMDH2; Pracharoenwattana et al., 2007; Eubel et al., 2008), two isoforms are mitochondrial MDH (MMDH; MMDH1 and MMDH2; Millar et al., 2001; Lee et al., 2008; Tomaz et al., 2010), and one isoform is plastidial MDH (plastid-localized NAD-dependent MDH [pdNAD-MDH]; Berkemeyer et al., 1998). The remaining three isoforms have no detectable target sequence and are thought to be cytosolic MDH (CMDH; CMDH1, CMDH2, and CMDH3). The Arabidopsis genome also encodes an additional NADP-dependent isoform of MDH, which is localized to the plastid (Hebbelmann et al., 2012).The physiological role of the different isoforms depends on the subcellular localization and the different metabolic pathways occurring there. For instance, MMDH was reported to be involved in two processes that are at least partly mitochondrial: leaf respiration and photorespiration (Tomaz et al., 2010). An MMDH null mutant (mmdh1 mmdh2) was slow growing and showed elevated leaf respiration in the dark and the light, although photosynthetic capacity was not affected. Tomaz et al. (2010) proposed that MMDH uses NADH to reduce OAA to malate, which is then shuttled to the cytosol, rather than generate NADH to fuel mitochondrial respiration (Tomaz et al., 2010). PMDH might serve at least two different functions. First, during fatty acid β-oxidation, which generates NADH, PMDH is proposed to regenerate the electron acceptor NAD by reducing OAA to malate, which is then shuttled to the cytosol in exchange for OAA (Pracharoenwattana et al., 2007). Second, PMDH is thought to generate NADH during photorespiration by oxidation of malate imported from the cytosol (Reumann and Weber, 2006; Cousins et al., 2008). Arabidopsis mutants lacking PMDH (pmdh1 pmdh2) are severely impaired in β-oxidation, and seedling establishment is strongly impaired and dependent on the supply of exogenous sugar (Pracharoenwattana et al., 2007), a phenotype characteristic of β-oxidation mutants (Pinfield-Wells et al., 2005; Baker et al., 2006). However, after transfer of established pmdh1 pmdh2 seedlings to compost, they grew only slightly slower than wild-type plants (Pracharoenwattana et al., 2007).Until recently, genetic evidence for the roles of the plastidial MDH isoforms was scarce. In most C₄ plants, NADP-MDH is directly involved in CO₂ fixation, catalyzing the formation of the stable CO₂ carrier malate from the primary CO₂ fixation product OAA (Scheibe, 1987). However, in C₃ plants, NADP-MDH has long been proposed to have its major function in the malate valve, leading to shuttling of reducing power (as malate) from the chloroplast to the cytosol during the day and thereby regenerating the electron acceptor NADP inside the chloroplasts (Heber, 1974; Lance and Rustin, 1984; Scheibe, 1987). NADP-MDH is redox activated by thioredoxins in the light and essentially inactive in the dark (Scheibe, 1987; Buchanan and Balmer, 2005). The widely accepted belief that chloroplasts only possess this one strictly light-/redox-activated NADP-MDH temporarily led to the conclusion that the malate valve only works in illuminated chloroplasts (Berkemeyer et al., 1998; Scheibe, 2004). However, a recent study showed that Arabidopsis plants lacking NADP-MDH (nadp-mdh) were indistinguishable from wild-type plants, even under conditions that are supposed to provoke the accumulation of excess electrons and the production of cytotoxic reactive oxygen species (high light and short days; Hebbelmann et al., 2012). This finding indicates that NADP-MDH is not crucial for providing electron acceptors in chloroplasts, but it rather suggests that other mechanisms can counteract or prevent overreduction of the chloroplast.The existence of a second MDH isoform in plastids, which uses NAD as cofactor, has been questioned, because it could not be ruled out that NAD-MDH activity detected in isolated chloroplasts was caused by contamination from other organelles (Siebke et al., 1991; Backhausen et al., 1998). In 1998, Berkemeyer et al. (1998) reported the cloning, heterologous expression, and in vitro characterization of a pdNAD-MDH from Arabidopsis (At3g47520). In contrast to NADP-MDH, pdNAD-MDH is active under both light and dark conditions in isolated chloroplasts, and the activities of both enzymes are within the same range in the light (Backhausen et al., 1998; Berkemeyer et al., 1998). However, up to now, genetic evidence for the in vivo function of pdNAD-MDH is missing, and experimental data are scarce. Backhausen et al. (1998) showed that chloroplasts and heterotrophic chromoplasts isolated from different sources followed by incubation in the dark concomitantly produced 3-phosphoglycerate and malate on addition of dihydroxyacetone phosphate and OAA to the medium. It was proposed that 3-phosphoglycerate production was in a glycolytic step involving glyceraldehyde 3-phosphate dehydrogenase (GAPDH) and that pdNAD-MDH regenerates the electron acceptor NAD required by GAPDH through reduction of OAA to malate, thus operating the malate-OAA shuttle in the dark and in nongreen tissues (Scheibe, 2004; Taniguchi and Miyake, 2012).Here, we aimed to evaluate the function of this MDH isoform in plastid metabolism by analyzing Arabidopsis plants with a transposon insertion in the pdNAD-MDH gene and Arabidopsis plants with reduced pdNAD-MDH by means of artificial microRNA silencing.     

Large-Scale Phosphoprotein Analysis in Medicago truncatula Roots Provides Insight into in Vivo Kinase Activity in Legumes

Nitrogen fixation in legumes requires the development of root organs called nodules and their infection by symbiotic rhizobia. Over the last decade, Medicago truncatula has emerged as a major model plant for the analysis of plant-microbe symbioses and for addressing questions pertaining to legume biology. While the initiation of symbiosis and the development of nitrogen-fixing root nodules depend on the activation of a protein phosphorylation-mediated signal transduction cascade in response to symbiotic signals produced by the rhizobia, few sites of in vivo phosphorylation have previously been identified in M. truncatula. We have characterized sites of phosphorylation on proteins from M. truncatula roots, from both whole cell lysates and membrane-enriched fractions, using immobilized metal affinity chromatography and tandem mass spectrometry. Here, we report 3,457 unique phosphopeptides spanning 3,404 nonredundant sites of in vivo phosphorylation on 829 proteins in M. truncatula Jemalong A17 roots, identified using the complementary tandem mass spectrometry fragmentation methods electron transfer dissociation and collision-activated dissociation. With this being, to our knowledge, the first large-scale plant phosphoproteomic study to utilize electron transfer dissociation, analysis of the identified phosphorylation sites revealed phosphorylation motifs not previously observed in plants. Furthermore, several of the phosphorylation motifs, including LxKxxs and RxxSxxxs, have yet to be reported as kinase specificities for in vivo substrates in any species, to our knowledge. Multiple sites of phosphorylation were identified on several key proteins involved in initiating rhizobial symbiosis, including SICKLE, NUCLEOPORIN133, and INTERACTING PROTEIN OF DMI3. Finally, we used these data to create an open-access online database for M. truncatula phosphoproteomic data.Medicago truncatula has become a model for studying the biology of leguminous plants such as soybean (Glycine max), alfalfa (Medicago sativa), and clover (Trifolium spp.; Singh et al., 2007). Most members of this vast family have the ability to fix atmospheric nitrogen by virtue of an endosymbiotic association with rhizobial bacteria, through which legumes undergo nodulation, the process of forming root nodules (Jones et al., 2007). Legumes are central to modern agriculture and civilization because of their ability to grow in nitrogen-depleted soils and replenish nitrogen through crop rotation. Consequently, there is great interest in understanding the molecular events that allow legumes to recognize their symbionts, develop root nodules, and fix nitrogen. Nod factors are lipochitooligosaccharidic signals secreted by the rhizobia and are required, in most legumes, for intracellular infection and nodule development. In recent decades, an elegant combination of genetics, biochemistry, and cell biology has shown that Nod factors activate intricate signaling events within cells of legume roots, including protein phosphorylation cascades and intracellular ion fluxes (Oldroyd and Downie, 2008).Protein phosphorylation is a central mechanism of signal transfer in cells (Laugesen et al., 2006; Peck, 2006; Huber, 2007). Several characterized protein kinases are required for symbiosis signal transduction in M. truncatula roots (Lévy et al., 2004; Yoshida and Parniske, 2005; Smit et al., 2007). A recent antibody-based study of cultured M. truncatula cells observed protein phosphorylation changes at the proteomic level in response to fungal infection (Trapphoff et al., 2009); however, the target residues of the phosphorylation events were not determined. A variety of studies have determined in vitro phosphorylation sites on legume proteins and demonstrated the biological importance of the target residues by mutagenesis (Yoshida and Parniske, 2005; Arrighi et al., 2006; Lima et al., 2006; Miyahara et al., 2008; Yano et al., 2008). To our knowledge, only six sites of in vivo protein phosphorylation have been detected for M. truncatula (Laugesen et al., 2006; Lima et al., 2006; Wienkoop et al., 2008), demonstrating the need for the identification of endogenous protein phosphorylation sites in legume model organisms on a proteome-wide scale.While considerable advancements have been made in the global analysis of protein phosphorylation (Nita-Lazar et al., 2008; Macek et al., 2009; Piggee, 2009; Thingholm et al., 2009), phosphoproteomics in plants has lagged years behind that of the mammalian systems (Kersten et al., 2006, 2009; Peck, 2006), which have more fully sequenced genomes and better annotated protein predictions. Arabidopsis (Arabidopsis thaliana), the first plant genome sequenced (Arabidopsis Genome Initiative, 2000), is now predicted to have over 1,000 protein kinases (Finn et al., 2008), approximately twice as many as in human (Manning et al., 2002). Because many of the kinases in the commonly studied mammalian systems are not conserved in the plant kingdom, there is significant need for large-scale phosphoproteomic technologies to discern the intricacies of phosphorylation-mediated cell signaling in plants. With the high mass accuracy afforded by the linear ion trap-orbitrap hybrid mass spectrometer (Makarov et al., 2006; Yates et al., 2006), recent studies in Arabidopsis have reported 2,597 phosphopeptides from suspension cell culture (Sugiyama et al., 2008) and 3,029 phosphopeptides from seedlings (Reiland et al., 2009).All previous large-scale plant phosphoproteomic studies have relied solely on collision-activated dissociation (CAD) during tandem mass spectrometry (MS/MS) and have not taken advantage of the more recently developed methods (Kersten et al., 2009) electron capture dissociation (Kelleher et al., 1999) or electron transfer dissociation (ETD; Coon et al., 2004; Syka et al., 2004). Mapping sites of posttranslational modifications, such as phosphorylation, is often more straightforward using electron-based fragmentation methods, as they frequently produce a full spectrum of sequence-informative ions without causing neutral loss of the modifying functional groups (Meng et al., 2005; Chi et al., 2007; Khidekel et al., 2007; Molina et al., 2007; Wiesner et al., 2008; Chalkley et al., 2009; Swaney et al., 2009). With an ETD-enabled hybrid orbitrap mass spectrometer (McAlister et al., 2007, 2008), we previously compared the performance of CAD and ETD tandem MS for large-scale identification of phosphopeptides (Swaney et al., 2009). ETD identified a greater percentage of unique phosphopeptides and more frequently localized phosphorylation sites. Still, the low overlap of identified phosphopeptides indicates that the two methods are highly complementary. With this in mind, we recently developed a decision tree-driven tandem MS algorithm to select the optimal fragmentation method for each precursor (Swaney et al., 2008).Here, we utilize this technology to map sites of in vivo protein phosphorylation in roots of M. truncatula Jemalong A17 plants. Phosphoproteins, from both whole-cell lysate and membrane-enriched fractions, were analyzed after digestion with a variety of different enzymes individually. Utilizing the complementary fragmentation methods of ETD and CAD, we report 3,404 nonredundant phosphorylation sites at an estimated false discovery rate (FDR) of 1%. Analysis of these data revealed several phosphorylation motifs not previously observed in plants. The phosphorylation sites identified provide insight into the potential regulation of key proteins involved in rhizobial symbiosis, potential consensus sequences by which kinases recognize their substrates, and critical phosphorylation events that are conserved between plant species.