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.     

Phototropins Function in High-Intensity Blue Light-Induced Hypocotyl Phototropism in Arabidopsis by Altering Cytosolic Calcium

Phototropins (phot1 and phot2), the blue light receptors in plants, regulate hypocotyl phototropism in a fluence-dependent manner. Especially under high fluence rates of blue light (HBL), the redundant function mediated by both phot1 and phot2 drastically restricts the understanding of the roles of phot2. Here, systematic analysis of phototropin-related mutants and overexpression transgenic lines revealed that HBL specifically induced a transient increase in cytosolic Ca²⁺ concentration ([Ca2+]_cyt) in Arabidopsis (Arabidopsis thaliana) hypocotyls and that the increase in [Ca2+]_cyt was primarily attributed to phot2. Pharmacological and genetic experiments illustrated that HBL-induced Ca²⁺ increases were modulated differently by phot1 and phot2. Phot2 mediated the HBL-induced increase in [Ca2+]_cyt mainly by an inner store-dependent Ca²⁺-release pathway, not by activating plasma membrane Ca²⁺ channels. Further analysis showed that the increase in [Ca2+]_cyt was possibly responsible for HBL-induced hypocotyl phototropism. An inhibitor of auxin efflux carrier exhibited significant inhibitions of both phototropism and increases in [Ca2+]_cyt, which indicates that polar auxin transport is possibly involved in HBL-induced responses. Moreover, PHYTOCHROME KINASE SUBSTRATE1 (PKS1), the phototropin-related signaling element identified, interacted physically with phototropins, auxin efflux carrier PIN-FORMED1 and calcium-binding protein CALMODULIN4, in vitro and in vivo, respectively, and HBL-induced phototropism was impaired in pks multiple mutants, indicating the role of the PKS family in HBL-induced phototropism. Together, these results provide new insights into the functions of phototropins and highlight a potential integration point through which Ca²⁺ signaling-related HBL modulates hypocotyl phototropic responses.Blue light (BL) is a key factor controlling plant growth and morphogenesis. Recent genetics investigations using Arabidopsis (Arabidopsis thaliana) have revealed that the BL receptors phototropin1 (phot1) and phot2 mediate BL-induced plant movements such as phototropism, chloroplast relocation, stomatal opening, leaf flattening, and leaf positioning responses (Inoue et al., 2010). Most of these responses are mediated redundantly by both phot1 and phot2 (Kinoshita et al., 2001; Sakamoto and Briggs, 2002), but some responses are mediated by either phot1 or phot2 (Sakai et al., 2001; Suetsugu et al., 2005). In addition, several lines of evidence have indicated that phot2 might negatively regulate the phot1-mediated response (de Carbonnel et al., 2010) and vice versa (Harada et al., 2003, 2013).One of the numerous physiological processes controlled by BL is phototropism. Phototropism enables plants to bend toward incident light by perceiving the direction, wavelength, and intensity of incident light so that they are able to obtain optimum light. Genetic evidence has shown that both phot1 and phot2 redundantly function to regulate hypocotyl phototropism in a fluence-dependent manner (Sakai et al., 2001). Phot1 functions at both low (0.01–1 μmol m⁻² s⁻¹) and high (greater than 1 μmol m⁻² s⁻¹) fluence rates to mediate phototropic responses, but phot2 functions only at high fluence rates (Inada et al., 2004). The functional specification of phot1 and phot2 could be attributed to the differences in signal intermediates between phot1 and phot2 signaling pathways.Genetic analysis has illustrated that phot1 mediates hypocotyl phototropism via its downstream signal transducers NONPHOTOTROPIC HYPOCOTYL3 (NPH3; Motchoulski and Liscum, 1999), ROOT PHOTOTROPISM2 (RPT2; Sakai et al., 2000), and NONPHOTOTROPIC HYPOCOTYL4/AUXIN RESPONSE FACTOR7 (NPH4/ARF7; Harper et al., 2000), resulting in the asymmetric distribution of auxin and the induction of a phototropic response in higher plants. Recently, studies have demonstrated that PHYTOCHROME KINASE SUBSTRATE (PKS) proteins are required for hypocotyl phototropism and that PKS1 binds PHOT1 and NPH3 in vivo (Lariguet et al., 2006). In addition, ATP-BINDING CASSETTE B19 (ABCB19), a newly identified auxin transporter, has been reported to interact with phot1 to regulate the BL-dependent phototropism (Christie et al., 2011). However, little is known about phot2-mediated phototropism for functional specialization, especially under high fluence rates of blue light (HBL), although several lines of evidence have shown that phot2- and phot1-mediated signaling pathways share some intermediates in BL responses (Kimura and Kagawa, 2006; Christie, 2007). Previous researches have suggested that phot1 acts not only positively in the presence of RPT2 but also negatively in its absence during the phototropic response of hypocotyls at high fluence rates, suggesting that RPT2 modulates the function of phot1. However, RPT2 does not act in the phot2-mediated pathway (Inada et al., 2004). More recently, RCN1-1, the A1 subunit of Ser/Thr PROTEIN PHOSPHATASE2A (PP2A), has been identified to interact with phot2. While reduced PP2A activity enhances the activity of phot2, it does not enhance either phot1 dephosphorylation or the activity of phot1 in mediating phototropism (Tseng and Briggs, 2010).Besides these signal intermediates noted above, phototropins may also confer their effects through the change of ion homeostasis. Ca²⁺ is a case in point. Recent reports have demonstrated that phototropins mediate the mobilization of Ca²⁺ in response to BL and that phot1 and phot2 mediate Ca²⁺ increases with distinctive mechanism in leaf cells according to the changes of ambient light intensity (Harada and Shimazaki, 2007). Under low fluence rates of BL, phot1 solely mediated Ca²⁺ influx through the channels in the plasma membrane. Under HBL, the increase in cytosolic Ca²⁺ concentration ([Ca2+]_cyt) is primarily attributed to phot2-dependent Ca²⁺ release from the internal calcium stores as well as the plasma membrane Ca²⁺ channels. Interestingly, the inhibitory effects of phospholipase C (PLC) inhibitors on the BL-induced responses in the wild type are larger than those in the phot1 single mutant, which indicates that there are some functional interactions between phot1 and phot2 to induce the elevation of cytosolic Ca²⁺ (Harada et al., 2003).However, until now, the function of Ca²⁺ in the phototropin-mediated phototropism signaling process has remained largely unknown. Pharmacological experiments indicate that changes in [Ca2+]_cyt are required for the phot1-mediated inhibition of hypocotyl growth but not for phot1-mediated phototropism (Folta et al., 2003). Otherwise, electrophysiological studies indicate that phototropic bending involves changes in ion fluxes, including calcium (Babourina et al., 2004). Such divergent responses show that the link between phototropins and calcium has not been firmly established in the case of hypocotyl phototropism. In phototropism, the phot1-dependent relocalization of the auxin efflux carrier PIN-FORMED1 (PIN1) is required for auxin redistribution (Blakeslee et al., 2004), and the PINOID kinase influences the relocalization of PIN1 (Friml et al., 2004). Given that both the calmodulin-related protein TCH3 and the calcium-binding protein AtPBP1 can bind to the PINOID kinase (Benjamins et al., 2003), it would appear that the cross talk among phototropins, auxin, and calcium is an important event for phototropism.Here, we show that HBL induces increases in [Ca2+]_cyt, which are mostly attributed to the function of phot2, and that the increases in [Ca2+]_cyt are required for HBL-induced phototropism in Arabidopsis hypocotyls. We also demonstrate that PKS1 may integrate phototropins with auxin transport in phot2-dependent Ca²⁺ signaling, and we discuss the possible molecular link between phototropins and other potential signal elements in HBL-induced phototropism.     

Phosphatidylinositol 4-Phosphate Negatively Regulates Chloroplast Division in Arabidopsis

Chloroplast division is performed by the constriction of envelope membranes at the division site. Although constriction of a ring-like protein complex has been shown to be involved in chloroplast division, it remains unknown how membrane lipids participate in the process. Here, we show that phosphoinositides with unknown function in envelope membranes are involved in the regulation of chloroplast division in Arabidopsis thaliana. PLASTID DIVISION1 (PDV1) and PDV2 proteins interacted specifically with phosphatidylinositol 4-phosphate (PI4P). Inhibition of phosphatidylinositol 4-kinase (PI4K) decreased the level of PI4P in chloroplasts and accelerated chloroplast division. Knockout of PI4Kβ2 expression or downregulation of PI4Kα1 expression resulted in decreased levels of PI4P in chloroplasts and increased chloroplast numbers. PI4Kα1 is the main contributor to PI4P synthesis in chloroplasts, and the effect of PI4K inhibition was largely abolished in the pdv1 mutant. Overexpression of DYNAMIN-RELATED PROTEIN5B (DRP5B), another component of the chloroplast division machinery, which is recruited to chloroplasts by PDV1 and PDV2, enhanced the effect of PI4K inhibition, whereas overexpression of PDV1 and PDV2 had additive effects. The amount of DRP5B that associated with chloroplasts increased upon PI4K inhibition. These findings suggest that PI4P is a regulator of chloroplast division in a PDV1- and DRP5B-dependent manner.     

Chloroplast Activity and 3′phosphadenosine 5′phosphate Signaling Regulate Programmed Cell Death in Arabidopsis

Programmed cell death (PCD) is a crucial process both for plant development and responses to biotic and abiotic stress. There is accumulating evidence that chloroplasts may play a central role during plant PCD as for mitochondria in animal cells, but it is still unclear whether they participate in PCD onset, execution, or both. To tackle this question, we have analyzed the contribution of chloroplast function to the cell death phenotype of the myoinositol phosphate synthase1 (mips1) mutant that forms spontaneous lesions in a light-dependent manner. We show that photosynthetically active chloroplasts are required for PCD to occur in mips1, but this process is independent of the redox state of the chloroplast. Systematic genetic analyses with retrograde signaling mutants reveal that 3′-phosphoadenosine 5′-phosphate, a chloroplast retrograde signal that modulates nuclear gene expression in response to stress, can inhibit cell death and compromises plant innate immunity via inhibition of the RNA-processing 5′-3′ exoribonucleases. Our results provide evidence for the role of chloroplast-derived signal and RNA metabolism in the control of cell death and biotic stress response.Programmed cell death (PCD) is a universal process in multicellular organisms, contributing to the controlled and active degradation of the cell. In plants, PCD is required for processes as diverse as development, self-incompatibility, and stress response. One well-documented example is the induction of PCD upon pathogen attack, allowing the confinement of the infection, and resistance of the plant. The signaling events leading to the onset of PCD have been extensively studied: pathogen recognition triggers activation of mitogen-activated protein kinase cascades, as well as production of reactive oxygen species (ROS) and salicylic acid (SA), which lead to a hypersensitive response (Coll et al., 2011).From a cellular point of view, several classes of plant PCD have been described and compared with the ones found in animal cells (van Doorn, 2011). PCD is thought to have evolved independently in plants and animals, and genes underlying these mechanisms are therefore poorly conserved between the two kingdoms. However, most cellular features are conserved between plant and animal PCD that are both characterized by cell shrinkage, chromatin condensation, DNA laddering, mitochondria permeabilization, and depolarization (Dickman and Fluhr, 2013). In animal cells, mitochondria play a central role in the regulation of apoptosis (Czabotar et al., 2014; Mariño et al., 2014), and this role is likely shared between the two kingdoms (Lord and Gunawardena, 2012). That said, additional mitochondria-independent PCD pathways have clearly evolved in plants.Genetic approaches have greatly contributed to our understanding of cellular pathways governing PCD in plants. For example, the isolation of lesion mimic mutants (LMMs), in which cell death occurs spontaneously, has allowed the identification of several negative regulators of cell death (for review, see Bruggeman et al., 2015b). Interestingly, lesion formation is light dependent in several of these mutants, which include one of the best characterized LMMs—lesions simulating disease1 (lsd1; Dietrich et al., 1994). The LSD1 protein is required for plant acclimation to excess excitation energy (Mateo et al., 2004): when plants are exposed to excessive amounts of light, the redox status of the plastoquinone pool in the chloroplastic electron transfer chain is thought to influence LSD1-dependent signaling to modulate cell death (Mühlenbock et al., 2008). Additionally, we have previously identified the myoinositol phosphate synthase1 (mips1) mutant as a LMM, in which lesion formation is also light dependent (Meng et al., 2009). This mutant is deficient in the myoinositol (MI) phosphate synthase that catalyzes the first committed step of MI biosynthesis and displays pleiotropic defects such as reduced root growth, abnormal vein development, and spontaneous cell death on leaves, together with severe growth reduction after lesions begin to develop (Meng et al., 2009; Donahue et al., 2010). The light-dependent PCD in the mips1 mutant, as observed for lsd1, suggests that chloroplasts may play a role in the MI-dependent cell death regulation. Accumulating evidence suggests that chloroplasts may play a central role in PCD regulation like mitochondria in animal cells (Wang and Bayles, 2013). First, as described in the case of lsd1, excess light energy received by the chloroplast can function as a trigger for PCD. Furthermore, singlet oxygen (¹O₂), a ROS, can activate the EXECUTER1 (EX1) and EX2 proteins in the chloroplasts to initiate PCD (Lee et al., 2007). Likewise, ROS generated by chloroplasts play a major role for PCD onset during nonhost interaction between tobacco (Nicotiana tabacum) and Xanthomonas campestris (Zurbriggen et al., 2009). Finally, functional chloroplasts have also been shown to be required for PCD in cell suspensions (Gutierrez et al., 2014) and in a number of LMMs (Mateo et al., 2004; Meng et al., 2009; Bruggeman et al., 2015b). Thus, chloroplasts are now recognized as important components of plant defense response against pathogens (Stael et al., 2015) and are proposed to function with mitochondria in the execution of PCD (Van Aken and Van Breusegem, 2015). However, the exact signaling and metabolic contribution of chloroplasts to PCD remain to be elucidated. Furthermore, cross talk between chloroplasts and mitochondria does occur, such as during photorespiration (Sunil et al., 2013), but whether such communication functions sequentially or in parallel in the control of PCD remains to be determined (Van Aken and Van Breusegem, 2015).To further investigate how chloroplasts contribute to the regulation of cell death, we performed both forward and reverse genetics on the mips1 mutant. An extragenic secondary mutation in divinyl protochlorophyllide 8-vinyl reductase involved in chlorophyll biosynthesis leads to chlorophyll deficiency that abolishes the mips1 cell death phenotype, as do changes in CO₂ availability. These findings provide evidence for a link between photosynthetic activity and PCD induction in mips1. Additionally, we investigated the contribution of several retrograde signaling pathways (Chan et al., 2015) to the control of PCD in mips1. This process was independent of GENOMES UNCOUPLED (GUN) and EX signaling pathways, but we found that the SAL1-PAP_XRN retrograde signaling pathway inhibits cell death as well as basal defense reactions in Arabidopsis (Arabidopsis thaliana).     

Cell Growth Defect Factor1/CHAPERONE-LIKE PROTEIN OF POR1 Plays a Role in Stabilization of Light-Dependent Protochlorophyllide Oxidoreductase in Nicotiana benthamiana and Arabidopsis

Angiosperms require light for chlorophyll biosynthesis because one reaction in the pathway, the reduction of protochlorophyllide (Pchlide) to chlorophyllide, is catalyzed by the light-dependent protochlorophyllide oxidoreductase (POR). Here, we report that Cell growth defect factor1 (Cdf1), renamed here as CHAPERONE-LIKE PROTEIN OF POR1 (CPP1), an essential protein for chloroplast development, plays a role in the regulation of POR stability and function. Cdf1/CPP1 contains a J-like domain and three transmembrane domains, is localized in the thylakoid and envelope membranes, and interacts with POR isoforms in chloroplasts. CPP1 can stabilize POR proteins with its holdase chaperone activity. CPP1 deficiency results in diminished POR protein accumulation and defective chlorophyll synthesis, leading to photobleaching and growth inhibition of plants under light conditions. CPP1 depletion also causes reduced POR accumulation in etioplasts of dark-grown plants and as a result impairs the formation of prolamellar bodies, which subsequently affects chloroplast biogenesis upon illumination. Furthermore, in cyanobacteria, the CPP1 homolog critically regulates POR accumulation and chlorophyll synthesis under high-light conditions, in which the dark-operative Pchlide oxidoreductase is repressed by its oxygen sensitivity. These findings and the ubiquitous presence of CPP1 in oxygenic photosynthetic organisms suggest the conserved nature of CPP1 function in the regulation of POR.     

A transit peptide-like sorting signal at the C terminus directs the Bienertia sinuspersici preprotein receptor Toc159 to the chloroplast outer membrane

Although Toc159 is known to be one of the key GTPase receptors for selective recognition of chloroplast preproteins, the mechanism for its targeting to the chloroplast surface remains unclear. To compare the targeting of these GTPase receptors, we identified two Toc159 isoforms and a Toc34 from Bienertia sinuspersici, a single-cell C₄ species with dimorphic chloroplasts in individual chlorenchyma cells. Fluorescent protein tagging and immunogold studies revealed that the localization patterns of Toc159 were distinctive from those of Toc34, suggesting different targeting pathways. Bioinformatics analyses indicated that the C-terminal tails (CTs) of Toc159 possess physicochemical and structural properties of chloroplast transit peptides (cTPs). These results were further confirmed by fluorescent protein tagging, which showed the targeting of CT fusion proteins to the chloroplast surface. The CT of Bs Toc159 in reverse orientation functioned as a cleavable cTP that guided the fluorescent protein to the stroma. Moreover, a Bs Toc34 mutant protein was retargeted to the chloroplast envelope using the CTs of Toc159 or reverse sequences of other cTPs, suggesting their conserved functions. Together, our data show that the C terminus and the central GTPase domain represent a novel dual domain–mediated sorting mechanism that might account for the partitioning of Toc159 between the cytosol and the chloroplast envelope for preprotein recognition.     

A Dominant Mutation in the Light-Oxygen and Voltage2 Domain Vicinity Impairs Phototropin1 Signaling in Tomato

In higher plants, blue light (BL) phototropism is primarily controlled by the phototropins, which are also involved in stomatal movement and chloroplast relocation. These photoresponses are mediated by two phototropins, phot1 and phot2. Phot1 mediates responses with higher sensitivity than phot2, and phot2 specifically mediates chloroplast avoidance and dark positioning responses. Here, we report the isolation and characterization of a Nonphototropic seedling1 (Nps1) mutant of tomato (Solanum lycopersicum). The mutant is impaired in low-fluence BL responses, including chloroplast accumulation and stomatal opening. Genetic analyses show that the mutant locus is dominant negative in nature. In dark-grown seedlings of the Nps1 mutant, phot1 protein accumulates at a highly reduced level relative to the wild type and lacks BL-induced autophosphorylation. The mutant harbors a single glycine-1484-to-alanine transition in the Hinge1 region of a phot1 homolog, resulting in an arginine-to-histidine substitution (R495H) in a highly conserved A′α helix proximal to the light-oxygen and voltage2 domain of the translated gene product. Significantly, the R495H substitution occurring in the Hinge1 region of PHOT1 abolishes its regulatory activity in Nps1 seedlings, thereby highlighting the functional significance of the A′α helix region in phototropic signaling of tomato.Being sessile in nature, plants have developed diverse sets of sensory mechanisms, integrating external cues such as light, water, and temperature to adapt their growth and development to the ambient environment. Plants have evolved a cohort of photoreceptors such as red/far-red light-sensing phytochromes (Chen and Chory, 2011), UV-A/blue light (BL)-sensing phototropins (Christie, 2007; Holland et al., 2009; Suetsugu and Wada, 2013), cryptochromes (Yu et al., 2010; Liu et al., 2011), Zeitlupe (ZTL)/Flavin-binding, Kelch repeat, F-box protein1/light-oxygen and voltage (LOV)-kelch protein2 members of the ZTL/ADAGIO putative family of photoreceptors (Suetsugu and Wada, 2013), and UV-B light-sensing UV-B resistance8 (Heijde and Ulm, 2012), enabling them to sense nearly the full range of the solar spectrum. One of the most visually obvious photoresponses of flowering plants involves the growth and orientation of organs toward or away from light, particularly during the early stages of growth and the establishment of seedlings (Iino, 1990) and during gap-filling situations in dense canopy conditions (Ballaré, 1999) for optimizing photosynthesis and interspecies/intraspecies competition. Several studies involving the relative effectiveness of different wavelengths of the solar spectrum as well as monitoring of lateral differences in light intensity revealed that the directional growth of plants is specifically mediated by the UV-A/blue region of the visible spectrum. Molecular genetic analysis of Arabidopsis (Arabidopsis thaliana) mutants inhibited in hypocotyl curvature toward BL revealed that, among the UV-A light-/BL-specific photoreceptors, the phototropins perceive ambient light as a cue for directional growth (Liscum and Briggs, 1995; Kagawa et al., 2001).Phototropins have been identified in several plant species, ranging from the green alga Chlamydomonas reinhardtii to higher plants (Briggs et al., 2001). To date, two members of the phototropins have been reported from higher plants, phot1 and phot2, which share sequence homology (Sakai et al., 2001). Physiological analyses with Arabidopsis mutants lacking phot1 and phot2 have revealed that, in addition to regulating the hypocotyl curvature of seedlings toward BL (Huala et al., 1997; Christie et al., 1998), phototropins also regulate a diverse range of responses in flowering plants (Christie and Murphy, 2013; Hohm et al., 2013). These responses include chloroplast movements (Sakai et al., 2001), nuclear positioning (Iwabuchi et al., 2007), stomatal opening (Kinoshita et al., 2001), sun tracking (Inoue et al., 2008b), leaf expansion (Ohgishi et al., 2004), leaf movements (Inoue et al., 2005), leaf photomorphogenesis (Kozuka et al., 2011), leaf flattening (Sakamoto and Briggs, 2002), and the rapid inhibition of the growth of etiolated hypocotyls (Folta and Spalding, 2001).While both phot1 and phot2 overlap in function in regulating phototropism, chloroplast accumulation, leaf expansion, and stomatal opening, they also exhibit differential photosensitivity to BL, where phot1 is more sensitive to low-fluence BL than phot2. Both phot1 and phot2 redundantly regulate the chloroplast accumulation toward low-fluence BL, and phot2 exclusively regulates the chloroplast avoidance from high-fluence BL (Jarillo et al., 2001; Kagawa et al., 2001), while phot1 solely mediates the rapid inhibition of the elongation of etiolated hypocotyls (Folta and Spalding, 2001). Analysis of mutants downstream of blue light perception by phototropins revealed that the phototropin signaling branches out at an early step, and phot1 and phot2 trigger distinct photoresponses recruiting multiple signaling partners (Christie and Murphy, 2013; Hohm et al., 2013).Molecular characterizations have shown that phototropins are plasma membrane-associated Ser/Thr kinases containing a photosensory domain (Briggs and Christie, 2002) in the N-terminal region composed of two LOV domains (LOV1 and LOV2) and the kinase domain at the C-terminal end. The LOV1 and LOV2 domains bind the FMN as chromophore and are responsible for BL sensing by phototropin. Although phototropins characteristically possess two LOV domains, the photoregulation of phototropin activity is predominantly mediated by LOV2 (Christie, 2007). The exposure to BL also causes adduct formation between the FMN and the Cys residue in LOV domains and leads to the phosphorylation of phototropin, which is believed to be the primary step in the transmission of phototropic signals (Christie et al., 1998; Sakai et al., 2000). To decipher the functions of different domains of phototropins, many different substitution mutants of phototropins have been generated, which have enabled the elucidation of the functional significance of the different domains (Matsuoka and Tokutomi, 2005; Jones et al., 2007; Kong et al., 2007; Inoue et al., 2008a). Inoue et al. (2008a) showed that the BL-induced autophosphorylation of Ser-851 in the C-terminal kinase domain of phototropin is the primary step for initiating stomatal opening, phototropism, chloroplast accumulation, and leaf flattening. Mutational studies also revealed that the photosensory N-terminal domain of phototropin acts as a kinase inhibitor, where the LOV2 domain inhibits the activity of kinase domain by binding to it, and BL exposure is required for the dissociation of the LOV2 domain, enabling phosphorylation of the kinase domain (Matsuoka and Tokutomi, 2005; Jones et al., 2007).While our current understanding of phototropism has been greatly facilitated by the isolation of phototropins and their signaling mutants, the phot mutants identified to date are loss-of-function alleles. The lack of dominant-negative alleles or alleles with increased sensitivity to phototropic stimulus has hindered exploration into the roles of different domains of phot proteins in regulating phototropic signaling. In addition, the dearth of mutants defective in phototropin or phototropin-mediated responses has been a major bottleneck in furthering our understanding of the function of phototropins in crop species. Although phototropin homologs have been identified from a variety of crop species, including oat (Avena sativa; Zacherl et al., 1998), rice (Oryza sativa; Kanegae et al., 2000), and tomato (Solanum lycopersicum; Sharma et al., 2007; Sharma and Sharma, 2007), only the coleoptile phototropism1 mutant of rice has been isolated, which is defective in BL phototropism (Haga et al., 2005).Here, we report that in a mutant screen for nonphototropic seedlings under continuous BL, we recovered a strong dominant-negative mutation of phot1. The dominant-negative mutations are useful to elucidate redundant functions, as mutant protein in addition to suppressing its own functions can also suppress the function of its partners. The characterization of this new phot1 mutant revealed that the dominant activity is caused by the substitution of an Arg residue located in the A′α helix in the Hinge1 region between the LOV1 and LOV2 domains. Our study shows the functional importance of the A′α helix (Halavaty and Moffat, 2007) in regulating phot1-mediated signaling in tomato.     

HYPERSENSITIVE TO HIGH LIGHT1 Interacts with LOW QUANTUM YIELD OF PHOTOSYSTEM II1 and Functions in Protection of Photosystem II from Photodamage in Arabidopsis

Under high-irradiance conditions, plants must efficiently protect photosystem II (PSII) from damage. In this study, we demonstrate that the chloroplast protein HYPERSENSITIVE TO HIGH LIGHT1 (HHL1) is expressed in response to high light and functions in protecting PSII against photodamage. Arabidopsis thaliana hhl1 mutants show hypersensitivity to high light, drastically decreased PSII photosynthetic activity, higher nonphotochemical quenching activity, a faster xanthophyll cycle, and increased accumulation of reactive oxygen species following high-light exposure. Moreover, HHL1 deficiency accelerated the degradation of PSII core subunits under high light, decreasing the accumulation of PSII core subunits and PSII–light-harvesting complex II supercomplex. HHL1 primarily localizes in the stroma-exposed thylakoid membranes and associates with the PSII core monomer complex through direct interaction with PSII core proteins CP43 and CP47. Interestingly, HHL1 also directly interacts, in vivo and in vitro, with LOW QUANTUM YIELD OF PHOTOSYSTEM II1 (LQY1), which functions in the repair and reassembly of PSII. Furthermore, the hhl1 lqy1 double mutants show increased photosensitivity compared with single mutants. Taken together, these results suggest that HHL1 forms a complex with LQY1 and participates in photodamage repair of PSII under high light.     

Sequence Motifs in Transit Peptides Act as Independent Functional Units and Can Be Transferred to New Sequence Contexts

A large number of nuclear-encoded proteins are imported into chloroplasts after they are translated in the cytosol. Import is mediated by transit peptides (TPs) at the N termini of these proteins. TPs contain many small motifs, each of which is critical for a specific step in the process of chloroplast protein import; however, it remains unknown how these motifs are organized to give rise to TPs with diverse sequences. In this study, we generated various hybrid TPs by swapping domains between Rubisco small subunit (RbcS) and chlorophyll a/b-binding protein, which have highly divergent sequences, and examined the abilities of the resultant TPs to deliver proteins into chloroplasts. Subsequently, we compared the functionality of sequence motifs in the hybrid TPs with those of wild-type TPs. The sequence motifs in the hybrid TPs exhibited three different modes of functionality, depending on their domain composition, as follows: active in both wild-type and hybrid TPs, active in wild-type TPs but inactive in hybrid TPs, and inactive in wild-type TPs but active in hybrid TPs. Moreover, synthetic TPs, in which only three critical motifs from RbcS or chlorophyll a/b-binding protein TPs were incorporated into an unrelated sequence, were able to deliver clients to chloroplasts with a comparable efficiency to RbcS TP. Based on these results, we propose that diverse sequence motifs in TPs are independent functional units that interact with specific translocon components at various steps during protein import and can be transferred to new sequence contexts.The chloroplasts of plant cells have more than 3,000 different types of proteins involved in their functions (Leister, 2003; Li and Chiu, 2010), and more than 90% of these proteins are encoded in the nucleus and translated by cytosolic ribosomes (Li and Chiu, 2010; Lee et al., 2013a). Consequently, one of the most critical processes in chloroplast proteome biogenesis is the specific, posttranslational delivery of these nuclear-encoded proteins to chloroplasts (Jarvis, 2008; Li and Chiu, 2010; Lee et al., 2013a, 2014). Delivery to chloroplasts requires a specific targeting signal whose form depends on the type of protein and its location in the chloroplast. Most proteins imported into the chloroplast contain an N-terminal transit peptide (TP) as a targeting signal (Lee et al., 2006, 2008, 2013a; Chotewutmontri et al., 2012; Li and Teng, 2013). The TP is cleaved off after import into the chloroplast; thus, the proteins that still contain the TP are called preproteins. Despite progress made in previous studies (Lee et al., 2008; Chotewutmontri et al., 2012; Li and Teng, 2013), the types of information encoded by the long TPs, as well as how this information determines translocation through the import channel, remain to be elucidated.One long-lasting question regarding the mechanism of TP-mediated protein import is how TPs can specifically deliver proteins into chloroplasts. In striking contrast to endoplasmic reticulum (ER)-targeting signals, TPs are highly diverse at the primary sequence level and do not converge toward a consensus sequence. The leader sequence, which contains the N-terminal ER-targeting signal, is composed of a stretch of hydrophobic amino acids ranging from 15 to 20 residues. Although the exact sequence is highly variable, the residues tend to be hydrophobic, making a high degree of hydrophobicity a common characteristic feature for both luminal and membrane proteins. Despite their diversity in primary sequence, TPs also share certain characteristics that serve as the basis for the software prediction of chloroplast proteins; these features include an amino acid composition with a high concentration of hydroxylated residues and a lack of acidic residues (Bruce, 2000; Bhushan et al., 2006), an unfolded and extended structure, an α-helix-containing secondary structure that may be induced by binding to the lipids of chloroplasts (Wienk et al., 1999; Bruce, 2000), and an abundance of Pro residues that may contribute to the unstructured nature of TPs (Pilon et al., 1995; Bruce, 2000; Zybailov et al., 2008).These features provide insight into the sequence information carried by TPs. However, we are still far from fully understanding how TPs function in the mechanism of protein import into chloroplasts. Recent studies have identified sequence motifs by analyzing various deletion and substitution mutants (Pilon et al., 1995; Lee et al., 2006, 2008, 2013a; Chotewutmontri et al., 2012). These motifs, or domains, are thought to be involved in the interaction with components of the translocon (Chotewutmontri et al., 2012; Li and Teng, 2013). Moreover, multiple sequence motifs function individually, or in a combinatorial manner, during specific steps of the import process (Lee et al., 2006, 2008, 2009a). In addition, certain motifs share functional redundancy, or are additive or synergistic. However, despite the progress in identifying sequence motifs from different TPs, it remains unknown how the large number of diverse TPs, as a whole, can deliver proteins to chloroplasts. In ER targeting, the targeting machinery recognizes hydrophobicity, a common feature of the leader sequences, but not the primary sequence (Hessa et al., 2005). Therefore, leader sequences with different primary sequences can be recognized by the same molecular machinery. However, in contrast to the leader sequences, the TPs of chloroplast preproteins contain different sets of sequence motifs (Lee et al., 2006, 2008). These observations raise several questions, including (1) how the large number of TPs with different sets of sequence motifs can be recognized by only a few import receptors (Li and Chiu, 2010; Lee et al., 2013a; Li and Teng, 2013), and (2) how TPs can have such diverse sequences while still retaining their function.In this study, we investigated the design principles of TPs with diverse primary sequences. Using TPs of the Rubisco small subunit (RbcS) and chlorophyll a/b-binding protein (Cab) proteins, which have completely different primary sequences and functional motifs (Lee et al., 2006, 2008), we generated hybrid TPs and examined their activities in chloroplast protein import within protoplasts. We provide evidence that sequence motifs are independent functional units that interact with various components of the translocon during import into chloroplasts and can be transferred to new sequence contexts. However, the functionalities as well as the activities of these motifs are greatly dependent on the overall sequence context of, and their positions in, TPs. In addition, we demonstrated that functional synthetic transit peptides (SynTPs) can be generated by incorporating only a few sequence motifs from RbcS and Cab TPs into an unrelated sequence.     

Cytoplasmic Nucleation and Atypical Branching Nucleation Generate Endoplasmic Microtubules in Physcomitrella patens

The mechanism underlying microtubule (MT) generation in plants has been primarily studied using the cortical MT array, in which fixed-angled branching nucleation and katanin-dependent MT severing predominate. However, little is known about MT generation in the endoplasm. Here, we explored the mechanism of endoplasmic MT generation in protonemal cells of Physcomitrella patens. We developed an assay that utilizes flow cell and oblique illumination fluorescence microscopy, which allowed visualization and quantification of individual MT dynamics. MT severing was infrequently observed, and disruption of katanin did not severely affect MT generation. Branching nucleation was observed, but it showed markedly variable branch angles and was occasionally accompanied by the transport of nucleated MTs. Cytoplasmic nucleation at seemingly random locations was most frequently observed and predominated when depolymerized MTs were regrown. The MT nucleator γ-tubulin was detected at the majority of the nucleation sites, at which a single MT was generated in random directions. When γ-tubulin was knocked down, MT generation was significantly delayed in the regrowth assay. However, nucleation occurred at a normal frequency in steady state, suggesting the presence of a γ-tubulin-independent backup mechanism. Thus, endoplasmic MTs in this cell type are generated in a less ordered manner, showing a broader spectrum of nucleation mechanisms in plants.     

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

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

Probing Arabidopsis Chloroplast Diacylglycerol Pools by Selectively Targeting Bacterial Diacylglycerol Kinase to Suborganellar Membranes

Diacylglycerol (DAG) is an intermediate in metabolism of both triacylglycerols and membrane lipids. Probing the steady-state pools of DAG and understanding how they contribute to the synthesis of different lipids is important when designing plants with altered lipid metabolism. However, traditional methods of assaying DAG pools are difficult, because its abundance is low and because fractionation of subcellular membranes affects DAG pools. To manipulate and probe DAG pools in an in vivo context, we generated multiple stable transgenic lines of Arabidopsis (Arabidopsis thaliana) that target an Escherichia coli DAG kinase (DAGK) to each leaflet of each chloroplast envelope membrane. E. coli DAGK is small, self inserts into membranes, and has catalytic activity on only one side of each membrane. By comparing whole-tissue lipid profiles between our lines, we show that each line has an individual pattern of DAG, phosphatidic acid, phosphatidylcholine, and triacylglycerol steady-state levels, which supports an individual function of DAG in each membrane leaflet. Furthermore, conversion of DAG in the leaflets facing the chloroplast intermembrane space by DAGK impairs plant growth. As a result of DAGK presence in the outer leaflet of the outer envelope membrane, phosphatidic acid accumulation is not observed, likely because it is either converted into other lipids or removed to other membranes. Finally, we use the outer envelope-targeted DAGK line as a tool to probe the accessibility of DAG generated in response to osmotic stress.Diacylglycerol (DAG) is a central metabolite in plant lipid metabolism. Its glycerol backbone is modified with two acyl chains. If a third acyl chain is added, triacylglycerol (TAG) is formed, whereas if a head group is added, it is converted into polar lipids such as a galactolipid. In green tissues, the majority of DAG is used as an intermediate in galactolipid synthesis, because the extensive thylakoid membrane system consists of approximately 85% galactolipids (Block et al., 1983). Although under normal conditions the galactolipids are exclusively chloroplastic, in Arabidopsis (Arabidopsis thaliana), the DAG used to make galactolipids is derived from assembly pathways in both the chloroplast and the endoplasmic reticulum (ER; Benning, 2009). In both pathways, the bulk of the fatty acids are synthesized in the chloroplast stroma (Browse et al., 1986) in the following order of abundance: 18:1, 16:0, and 18:0 (Wallis and Browse, 2002).In the chloroplast pathway, these fatty acids are directly attached to a glycerol-3-P, generating first lyso-phosphatidic acid (l-PtdOH) and then phosphatidic acid (PtdOH) in the inner leaflet of the chloroplast inner envelope (Fig. 1; Frentzen et al., 1983). The acyltransferases involved are specific to the extent that the sn-2 position of the glycerol backbone predominantly receives a 16:0 fatty acid. PtdOH is then used directly for phosphatidylglycerol (PtdGro) synthesis (Babiychuk et al., 2003) or converted to DAG by a PtdOH phosphatase (Joyard and Douce, 1977). The PtdOH phosphatase activity is known to be associated with the inner envelope, though which leaflet is obscured by the fact that DAG can efficiently flip across membranes (Hamilton et al., 1991) and the actual enzyme has not been unambiguously identified and located (Nakamura et al., 2007). However, the leaflet associations of two other enzymes that use DAG in the inner envelope have been established. MGD1, which uses DAG to synthesize the most abundant galactolipid, monogalactosyldiacylglycerol (MGDG), is on the outer leaflet of the inner envelope membrane (Xu et al., 2005), while SQD2, which uses DAG to generate the less abundant sulfolipid, sulfoquinovosyldiacylglycerol (SQDG), is located on the inner leaflet of the inner envelope membrane (Tietje and Heinz, 1998). Also associated with the inner envelope membrane are a number of fatty acid desaturases, including FAD4, FAD5, FAD6, FAD7, and FAD8 (Joyard et al., 2010). Two of these are specific, generating lipids with signature desaturations: FAD4 desaturates only the 16:0 fatty acid of PtdGro, giving plastidic PtdGro a distinct 16:1 Δ3 trans moiety (Browse et al., 1985; Gao et al., 2009), and FAD5 desaturates primarily the 16:0 fatty acid of MGDG, producing 16:1 Δ7 cis (Kunst et al., 1989). The remaining desaturases are less specific, with little preference for head group or acyl tail. They further desaturate 16:1 or 18:1 in the cis conformation to 16:2 or 18:2 (FAD6; Browse et al., 1989) and on to 16:3 or 18:3 (FAD7 and FAD8; Wallis and Browse, 2002). The combined actions of these FADs result in the highly desaturated fatty acid profiles seen for most chloroplast lipids.Open in a separate windowFigure 1.Overview of DAG pools in the chloroplast envelope membranes. Processes that are known to have activity feeding into or withdrawing from DAG pools in the chloroplast envelope membranes are shown. Enzymes are indicated, and their substrates and products are connected with black arrows. However, for space reasons, not all reactants are shown. Membrane leaflets are indicated, and enzymes with known membrane topology are displayed correctly, while those without known topology are displayed in the center of the appropriate membrane. The acyl group preferred by each l-PtdOH acyltransferase is given in parentheses. Proposed processes transporting lipids from the ER to the chloroplast are shown with dashed arrows. Enzymes are as follows: 1, ATS1; 2, ATS2; 3, lipid phosphate phosphatase γ; 4, MGD1; 5, SQD2; 6, cytosolic phospholipases; 7, MGD2 or MGD3; 8, SFR2; 9, acyl-CoA:glycerol-3-P acyltransferase; 10, l-PtdOH acyltransferase; 11, PtdOH phosphatase; 12, cytidine diphosphate-choline:DAG cholinephosphotransferase; 13, TGD4; and 14, TGD1, TGD 2, TGD3 lipid transport complex. OE, Chloroplast outer envelope membrane; IE, chloroplast inner envelope membrane; ACP, acyl carrier protein. [See online article for color version of this figure.]In unstressed plants, DAG seems to be used primarily in the inner chloroplast envelope. However, several conditions are known to cause extensive DAG use in the chloroplast outer envelope. During phosphate deprivation, MGD2 and MDG3 synthesize MGDG from DAG on the outer leaflet of the outer envelope membrane (Kobayashi et al., 2009). The DAG backbones are probably supplied from the phosphatidylcholine (PtdCho) pool by phospholipase activity, which was shown to be simultaneously up-regulated (Andersson et al., 2004; Nakamura et al., 2005). DAG is also generated during freezing stress by a galactolipid:galactolipid galactosyltransferase named Sensitive to FReezing2 (SFR2). This enzyme transfers the galactosyl head group of MGDG onto another MGDG, giving rise to digalactosyldiacylglycerol (DGDG) and DAG (Moellering et al., 2010). The DAG is subsequently sequestered into a lipid droplet by formation of TAG by an as yet unidentified enzyme.In the ER pathway, fatty acids synthesized in the chloroplast stroma are exported through a still poorly defined mechanism to the ER and activated to acyl-CoAs. Acyltransferases sequentially catalyze formation of l-PtdOH and PtdOH from glycerol-3-P and acyl-CoAs. Again, the acyltransferase working on the sn-2 position of the glycerol backbone is specific, but unlike the chloroplast isoform, it prefers an 18:1 carbon fatty acid (Frentzen et al., 1983). Newly generated PtdOH can be converted to PtdGro or phosphatidyl inositol (PtdIns) (Collin et al., 1999) or hydrolyzed to DAG (Shimojima et al., 2009). DAG can then be further metabolized to TAG and PtdCho. PtdCho acyl groups (18:1/18:1 and 18:1/16:0) are desaturated sequentially by desaturases FAD2 (Okuley et al., 1994) and FAD3 (Browse et al., 1993). These desaturases prefer PtdCho as substrate. The acyl chains modified on PtdCho are transferred to other ER lipids, including DAG, as a result of continual acyl editing of the PtdCho pool (Bates et al., 2012). Furthermore, PtdOH and many of the other extraplastidic phospholipids can be converted to DAG by action of phospholipases (Shimojima et al., 2009). These have as yet partially defined roles in response to stress or recycling of membrane lipids (Testerink and Munnik, 2005).Glycerolipid precursors generated by de novo synthesis, acyl editing, and possibly stress conditions in the ER are transported to the chloroplast by a mechanism that is likely to involve at least two putative lipid transporters: trigalactosyldiacylglycerol4 (TGD4) in the chloroplast outer envelope membrane and the TGD1, TGD2, and TGD3 complex in the inner envelope membrane (Wang and Benning, 2012). The actual lipid species transported remains unclear, but PtdCho, lyso-phosphatidylcholine, PtdOH, and DAG have been discussed in the literature (Andersson and Dörmann, 2009). The DAG moieties are then fully incorporated into all plastidic lipids except PtdGro, presumably using the same pathways that metabolize plastidic DAG, described above. Because of the preference of chloroplast and ER sn-2 acyltransferases for 16 or 18 carbon fatty acids, respectively, the origin of the DAG moieties can be distinguished by positional analysis of the acyl groups on the glycerol backbone (Roughan and Slack, 1982). In Arabidopsis, the chloroplast and ER lipid synthesis pathways contribute nearly equally to mature chloroplast lipids (Browse et al., 1986; Mongrand et al., 1998). Thus, the DAG pools described so far in the chloroplast inner and outer envelope membranes are each of dual origin.A challenge for the analysis of the different DAG pools is that this compound is not a bilayer-forming lipid and thus does not accumulate stably to high levels. Furthermore, during any lengthy fractionation procedure, its levels can be expected to alter, as DAG-modifying enzymes exist in multiple membranes. Moreover, because DAG is quickly metabolized and may have efficient transport systems (Dong et al., 2012), it is difficult to confirm whether metabolizing enzymes are accessing the same or separate DAG pools.To probe different DAG pools of chloroplast membranes in vivo, we have generated a series of stable transgenic Arabidopsis lines in which we target an Escherichia coli DAG kinase (DAGK) to each leaflet of the chloroplast envelope membranes. The basic utility of this approach was previously shown by targeting a DAGK to the chloroplast in tobacco (Nicotiana tabacum) using a single construct fusing the bacterial protein to the Rubisco small subunit N-terminal peptide (Fritz et al., 2007). Here, we present a full phenotypic analysis of these lines, determining which chloroplast membranes show steady-state alterations of DAG and PtdOH levels predicted by ectopic DAGK activity. We further determine the accessibility of DAG pools generated on the outer leaflet of the chloroplast outer envelope membrane during osmotic stress. Having this system established in Arabidopsis will allow characterization of DAG pools in multiple lipid mutant lines.     

Why have chloroplasts developed a unique motility system?

Organelle movement in plants is dependent on actin filaments with most of the organelles being transported along the actin cables by class XI myosins. Although chloroplast movement is also actin filament-dependent, a potential role of myosin motors in this process is poorly understood. Interestingly, chloroplasts can move in any direction and change the direction within short time periods, suggesting that chloroplasts use the newly formed actin filaments rather than preexisting actin cables. Furthermore, the data on myosin gene knockouts and knockdowns in Arabidopsis and tobacco do not support myosins'' XI role in chloroplast movement. Our recent studies revealed that chloroplast movement and positioning are mediated by the short actin filaments localized at chloroplast periphery (cp-actin filaments) rather than cytoplasmic actin cables. The accumulation of cp-actin filaments depends on kinesin-like proteins, KAC1 and KAC2, as well as on a chloroplast outer membrane protein CHUP1. We propose that plants evolved a myosin XI-independent mechanism of the actin-based chloroplast movement that is distinct from the mechanism used by other organelles.Key words: actin, Arabidopsis, blue light, kinesin, myosin, organelle movement, phototropinOrganelle movement and positioning are pivotal aspects of the intracellular dynamics in most eukaryotes. Although plants are sessile organisms, their organelles are quickly repositioned in response to fluctuating environmental conditions and certain endogenous signals. By and large, plant organelle movements and positioning are dependent on actin filaments, although microtubules play certain accessory roles in organelle dynamics.^1,2 Actin inhibitors effectively retard the movements of mitochondria,^3–6 peroxisomes,^5,7–11 Golgi stacks,^12,13 endoplasmic reticulum (ER),^14,15 and nuclei.^16–18 These organelles are co-aligned and associated with actin filaments.^{5,7,8,10–12,15,18} Recent progress in this field started to reveal the molecular motility system responsible for the organelle transport in plants.¹⁹Chloroplast movement is among the most fascinating models of organelle movement in plants because it is precisely controlled by ambient light conditions.^20,21 Weak light induces chloroplast accumulation response so that chloroplasts can capture photosynthetic light efficiently (Fig. 1A). Strong light induces chloroplast avoidance response to escape from photodamage (Fig. 1B).²² The blue light-induced chloroplast movement is mediated by the blue light receptor phototropin (phot). In some cryptogam plants, the red light-induced chloroplast movement is regulated by a chimeric phytochrome/phototropin photoreceptor neochrome.^23–25 In a model plant Arabidopsis, phot1 and phot2 function redundantly to regulate the accumulation response,²⁶ whereas phot2 alone is essential for the avoidance response.^27,28 Several additional factors regulating chloroplast movement were identified by analyses of Arabidopsis mutants deficient in chloroplast photorelocation.^29–32 In particular, identification of CHUP1 (chloroplast unusual positioning 1) revealed the connection between chloroplasts and actin filaments at the molecular level.²⁹ CHUP1 is a chloroplast outer membrane protein capable of interacting with F-actin, G-actin and profilin in vitro.^29,33,34 The chup1 mutant plants are defective in both the chloroplast movement and chloroplast anchorage to the plasma membrane,^22,29,33 suggesting that CHUP1 plays an important role in linking chloroplasts to the plasma membrane through the actin filaments. However, how chloroplasts move using the actin filaments and whether chloroplast movement utilizes the actin-based motility system similar to other organelle movements remained to be determined.Open in a separate windowFigure 1Schematic distribution patterns of chloroplasts in a palisade cell under different light conditions, weak (A) and strong (B) lights. Shown as a side view of mid-part of the cell and a top view with three different levels (i.e., top, middle and bottom of the cell). The cell was irradiated from the leaf surface shown as arrows. Weak light induces chloroplast accumulation response (A) and strong light induces the avoidance response (B).Here, we review the recent findings pointing to existence of a novel actin-based mechanisms for chloroplast movement and discuss the differences between the mechanism responsible for movement of chloroplasts and other organelles.     

MET1 Is a Thylakoid-Associated TPR Protein Involved in Photosystem II Supercomplex Formation and Repair in Arabidopsis

Photosystem II (PSII) requires constant disassembly and reassembly to accommodate replacement of the D1 protein. Here, we characterize Arabidopsis thaliana MET1, a PSII assembly factor with PDZ and TPR domains. The maize (Zea mays) MET1 homolog is enriched in mesophyll chloroplasts compared with bundle sheath chloroplasts, and MET1 mRNA and protein levels increase during leaf development concomitant with the thylakoid machinery. MET1 is conserved in C3 and C4 plants and green algae but is not found in prokaryotes. Arabidopsis MET1 is a peripheral thylakoid protein enriched in stroma lamellae and is also present in grana. Split-ubiquitin assays and coimmunoprecipitations showed interaction of MET1 with stromal loops of PSII core components CP43 and CP47. From native gels, we inferred that MET1 associates with PSII subcomplexes formed during the PSII repair cycle. When grown under fluctuating light intensities, the Arabidopsis MET1 null mutant (met1) showed conditional reduced growth, near complete blockage in PSII supercomplex formation, and concomitant increase of unassembled CP43. Growth of met1 in high light resulted in loss of PSII supercomplexes and accelerated D1 degradation. We propose that MET1 functions as a CP43/CP47 chaperone on the stromal side of the membrane during PSII assembly and repair. This function is consistent with the observed differential MET1 accumulation across dimorphic maize chloroplasts.     

Arabidopsis Phytochrome B Promotes SPA1 Nuclear Accumulation to Repress Photomorphogenesis under Far-Red Light

Phytochrome A (phyA) is the primary photoreceptor mediating deetiolation under far-red (FR) light, whereas phyB predominantly regulates light responses in red light. SUPPRESSOR OF PHYA-105 (SPA1) forms an E3 ubiquitin ligase complex with CONSTITUTIVE PHOTOMORPHOGENIC1 (COP1), which is responsible for the degradation of various photomorphogenesis-promoting factors, resulting in desensitization to light signaling. However, the role of phyB in FR light signaling and the regulatory pathway from light-activated phytochromes to the COP1-SPA1 complex are largely unknown. Here, we confirm that PHYB overexpression causes an etiolation response with reduced ELONGATED HYPOCOTYL5 (HY5) accumulation under FR light. Notably, phyB exerts its nuclear activities and promotes seedling etiolation in both the presence and absence of phyA in response to FR light. PhyB acts upstream of SPA1 and is functionally dependent on it in FR light signaling. PhyB interacts and forms a protein complex with SPA1, enhancing its nuclear accumulation under FR light. During the dark-to-FR transition, phyB is rapidly imported into the nucleus and facilitates nuclear SPA1 accumulation. These findings support the notion that phyB plays a role in repressing FR light signaling. Activity modulation of the COP1-SPA E3 complex by light-activated phytochromes is an effective and pivotal regulatory step in light signaling.     

The Cytosolic Nucleoprotein of the Plant-Infecting Bunyavirus Tomato Spotted Wilt Recruits Endoplasmic Reticulum–Resident Proteins to Endoplasmic Reticulum Export Sites

In contrast with animal-infecting viruses, few known plant viruses contain a lipid envelope, and the processes leading to their membrane envelopment remain largely unknown. Plant viruses with lipid envelopes include viruses of the Bunyaviridae, which obtain their envelope from the Golgi complex. The envelopment process is predominantly dictated by two viral glycoproteins (Gn and Gc) and the viral nucleoprotein (N). During maturation of the plant-infecting bunyavirus Tomato spotted wilt, Gc localizes at endoplasmic reticulum (ER) membranes and becomes ER export competent only upon coexpression with Gn. In the presence of cytosolic N, Gc remains arrested in the ER but changes its distribution from reticular into punctate spots. Here, we show that these areas correspond to ER export sites (ERESs), distinct ER domains where glycoprotein cargo concentrates prior to coat protein II vesicle–mediated transport to the Golgi. Gc concentration at ERES is mediated by an interaction between its cytoplasmic tail (CT) and N. Interestingly, an ER-resident calnexin provided with Gc-CT was similarly recruited to ERES when coexpressed with N. Furthermore, disruption of actin filaments caused the appearance of a larger amount of smaller ERES loaded with N-Gc complexes, suggesting that glycoprotein cargo concentration acts as a trigger for de novo synthesis of ERES.