Kinesins Have a Dual Function in Organizing Microtubules during Both Tip Growth and Cytokinesis in Physcomitrella patens

Microtubules (MTs) play a crucial role in the anisotropic deposition of cell wall material, thereby affecting the direction of growth. A wide range of tip-growing cells display highly polarized cell growth, and MTs have been implicated in regulating directionality and expansion. However, the molecular machinery underlying MT dynamics in tip-growing plant cells remains unclear. Here, we show that highly dynamic MT bundles form cyclically in the polarized expansion zone of the moss Physcomitrella patens caulonemal cells through the coalescence of growing MT plus ends. Furthermore, the plant-specific kinesins (KINID1) that are is essential for the proper MT organization at cytokinesis also regulate the turnover of the tip MT bundles as well as the directionality and rate of cell growth. The plus ends of MTs grow toward the expansion zone, and KINID1 is necessary for the stability of a single coherent focus of MTs in the center of the zone, whose formation coincides with the accumulation of KINID1. We propose that KINID-dependent MT bundling is essential for the correct directionality of growth as well as for promoting growth per se. Our findings indicate that two localized cell wall deposition processes, tip growth and cytokinesis, previously believed to be functionally and evolutionarily distinct, share common and plant-specific MT regulatory components.     

MICROTUBULE-ASSOCIATED PROTEIN65 Is Essential for Maintenance of Phragmoplast Bipolarity and Formation of the Cell Plate in Physcomitrella patens

The phragmoplast, a plant-specific apparatus that mediates cytokinesis, mainly consists of microtubules (MTs) arranged in a bipolar fashion, such that their plus ends interdigitate at the equator. Membrane vesicles are thought to move along the MTs toward the equator and fuse to form the cell plate. Although several genes required for phragmoplast MT organization have been identified, the mechanisms that maintain the bipolarity of phragmoplasts remain poorly understood. Here, we show that engaging phragmoplast MTs in a bipolar fashion in protonemal cells of the moss Physcomitrella patens requires the conserved MT cross-linking protein MICROTUBULE-ASSOCIATED PROTEIN65 (MAP65). Simultaneous knockdown of the three MAP65s expressed in those cells severely compromised MT interdigitation at the phragmoplast equator after anaphase onset, resulting in the collapse of the phragmoplast in telophase. Cytokinetic vesicles initially localized to the anaphase midzone as normal but failed to further accumulate in the next several minutes, although the bipolarity of the MT array was preserved. Our data indicate that the presence of bipolar MT arrays is insufficient for vesicle accumulation at the equator and further suggest that MAP65-mediated MT interdigitation is a prerequisite for maintenance of bipolarity of the phragmoplast and accumulation and/or fusion of cell plate–destined vesicles at the equatorial plane.     

Affinity Purification and Characterization of Functional Tubulin from Cell Suspension Cultures of Arabidopsis and Tobacco

Microtubules assemble into several distinct arrays that play important roles in cell division and cell morphogenesis. To decipher the mechanisms that regulate the dynamics and organization of this versatile cytoskeletal component, it is essential to establish in vitro assays that use functional tubulin. Although plant tubulin has been purified previously from protoplasts by reversible taxol-induced polymerization, a simple and efficient purification method has yet to be developed. Here, we used a Tumor Overexpressed Gene (TOG) column, in which the tubulin-binding domains of a yeast (Saccharomyces cerevisiae) TOG homolog are immobilized on resin, to isolate functional plant tubulin. We found that several hundred micrograms of pure tubulin can readily be purified from cell suspension cultures of tobacco (Nicotiana tabacum) and Arabidopsis (Arabidopsis thaliana). The tubulin purified by the TOG column showed high assembly competence, partly because of low levels of polymerization-inhibitory phosphorylation of α-tubulin. Compared with porcine brain tubulin, Arabidopsis tubulin is highly dynamic in vitro at both the plus and minus ends, exhibiting faster shrinkage rates and more frequent catastrophe events, and exhibits frequent spontaneous nucleation. Furthermore, our study shows that an internal histidine tag in α-tubulin can be used to prepare particular isotypes and specifically engineered versions of α-tubulin. In contrast to previous studies of plant tubulin, our mass spectrometry and immunoblot analyses failed to detect posttranslational modification of the isolated Arabidopsis tubulin or detected only low levels of posttranslational modification. This novel technology can be used to prepare assembly-competent, highly dynamic pure tubulin from plant cell cultures.Microtubules (MTs) are important cytoskeletal polymers that are conserved in eukaryotic cells and are assembled from α- and β-tubulin heterodimers (Desai and Mitchison, 1997). In plants, MTs have important functions in essential cellular processes, such as cell division, and in cell morphogenesis. MTs in plant cells adopt several distinct higher order arrays and are remodeled in response to the cell cycle, developmental programs, and environmental cues (Hashimoto, 2015). Genetic, molecular, and cell biological approaches have been used to identify cellular factors that regulate the organization and dynamics of plant MTs. Considerable effort has been devoted to simulating the organization of cortical MT arrays by computational modeling.Cell-free in vitro studies are essential for the biochemical characterization of various MT regulators and for elucidating the mechanistic principles underlying the versatility of this dynamic polymer in cellular functions. The purification of sufficient amounts of assembly-competent tubulin is a prerequisite for these in vitro studies. Tubulin is traditionally purified from mammalian brains, since these tissues contain sufficiently high concentrations of tubulin to allow MT assembly in crude cell extracts. Polymerized MTs and their associated MT-binding proteins are separated from other cellular proteins by sedimentation. Pelleted MTs are then depolymerized upon drug washout under MT-destabilizing conditions, such as high concentrations of salt and calcium and low temperature. A few rounds of assembly-disassembly cycles highly enrich for tubulin and copurify MT-associated proteins, which can subsequently be removed by column chromatography (Borisy et al., 1975). Tubulin has also been purified from several plant sources (Morejohn and Fosket, 1982; Mizuno, 1985; Jiang et al., 1992; Bokros et al., 1993; Moore et al., 1997). However, since tubulin concentrations are low in plant cells, taxol, which stabilizes MTs, is generally included in the polymerization buffer, and cytoplasm-rich miniprotoplasts, which lack vacuoles, are sometimes used as starting material (Hamada et al., 2013). Since it is technically challenging to isolate assembly-competent pure tubulin from nonneural sources (Sackett et al., 2010), general plant science laboratories may hesitate to prepare plant tubulin themselves.Although the primary amino acid sequences of eukaryotic tubulins are fairly well conserved and the biophysical mechanisms of MT assembly and disassembly are thought to be similar for all MTs, the kinetics of MT dynamic instability differ for MTs assembled from animal and plant tubulin (Moore et al., 1997). Interactions with MT-interacting proteins may differ for tubulins isolated from different biological sources, as reported for the MT-dependent activation of kinesin (Alonso et al., 2007). Posttranslational modifications of tubulin, which generate distinct tubulin signatures and may modulate the functions of MT-interacting proteins, such as kinesin (Sirajuddin et al., 2014), are extensive in brain tubulin (Janke, 2014) but may be quantitatively and qualitatively different in plant tubulin. Furthermore, MT nucleation by the γ-tubulin ring complex shows a strong preference for tubulin from the same species (Kollman et al., 2015). Thus, it is important to use plant tubulin, and not brain tubulin, for in vitro studies of plant MTs.Tubulin is folded by a series of molecular chaperones to form an αβ-tubulin heterodimer in which one structural GTP is embedded in the interdimer interface (Lundin et al., 2010). The requirement of these eukaryote-specific chaperones precludes the use of prokaryotic expression systems for synthesizing properly folded and functional tubulin. Bacterially synthesized tubulin can be folded in rabbit reticulocyte lysate to produce functional tubulin, but with moderate yields (Shah et al., 2001). A yeast (Saccharomyces cerevisiae) expression system has been developed to produce modified yeast tubulin (Uchimura et al., 2006; Johnson et al., 2011), but this system is not suitable for the synthesis of animal (Sirajuddin et al., 2014) and plant (our unpublished data) tubulin. A baculovirus-insect cell expression system was recently reported to yield functional human tubulin (Minoura et al., 2013).Tubulin-binding proteins have been used to develop affinity-purification columns. The TOG domains (named after the human MT regulator, colonic and hepatic Tumor Overexpressed Gene [ch-TOG]) are among the best-characterized tubulin-binding domains. ch-TOG and orthologs from other eukaryotes bind to the growing plus ends of MTs and accelerate MT growth (Al-Bassam and Chang, 2011). TOG domains from the yeast ortholog Stu2 were recently used to affinity purify assembly-competent tubulin from fungal and animal sources (Widlund et al., 2012). In this study, we demonstrate that a TOG-based affinity column can be used to purify functional tubulin from tobacco (Nicotiana tabacum) and Arabidopsis (Arabidopsis thaliana). We examined the posttranslational modifications of the isolated tubulins by mass spectrometry and immunoblot analysis and showed that a His-tagged Arabidopsis tubulin isotype could be purified using this column. These results show that wild-type and recombinant functional tubulin from plant sources can be isolated efficiently.     

Xyloglucan Deficiency Disrupts Microtubule Stability and Cellulose Biosynthesis in Arabidopsis,Altering Cell Growth and Morphogenesis

Xyloglucan constitutes most of the hemicellulose in eudicot primary cell walls and functions in cell wall structure and mechanics. Although Arabidopsis (Arabidopsis thaliana) xxt1 xxt2 mutants lacking detectable xyloglucan are viable, they display growth defects that are suggestive of alterations in wall integrity. To probe the mechanisms underlying these defects, we analyzed cellulose arrangement, microtubule patterning and dynamics, microtubule- and wall-integrity-related gene expression, and cellulose biosynthesis in xxt1 xxt2 plants. We found that cellulose is highly aligned in xxt1 xxt2 cell walls, that its three-dimensional distribution is altered, and that microtubule patterning and stability are aberrant in etiolated xxt1 xxt2 hypocotyls. We also found that the expression levels of microtubule-associated genes, such as MAP70-5 and CLASP, and receptor genes, such as HERK1 and WAK1, were changed in xxt1 xxt2 plants and that cellulose synthase motility is reduced in xxt1 xxt2 cells, corresponding with a reduction in cellulose content. Our results indicate that loss of xyloglucan affects both the stability of the microtubule cytoskeleton and the production and patterning of cellulose in primary cell walls. These findings establish, to our knowledge, new links between wall integrity, cytoskeletal dynamics, and wall synthesis in the regulation of plant morphogenesis.The primary walls of growing plant cells are largely constructed of cellulose and noncellulosic matrix polysaccharides that include hemicelluloses and pectins (Carpita and Gibeaut, 1993; Somerville et al., 2004; Cosgrove, 2005). Xyloglucan (XyG) is the most abundant hemicellulose in the primary walls of eudicots and is composed of a β-1,4-glucan backbone with side chains containing Xyl, Gal, and Fuc (Park and Cosgrove, 2015). XyG is synthesized in the Golgi apparatus before being secreted to the apoplast, and its biosynthesis requires several glycosyltransferases, including β-1,4-glucosyltransferase, α-1,6-xylosyltransferase, β-1,2-galactosyltransferase, and α-1,2-fucosyltransferase activities (Zabotina, 2012). Arabidopsis (Arabidopsis thaliana) XYLOGLUCAN XYLOSYLTRANSFERASE1 (XXT1) and XXT2 display xylosyltransferase activity in vitro (Faik et al., 2002; Cavalier and Keegstra, 2006), and strikingly, no XyG is detectable in the walls of xxt1 xxt2 double mutants (Cavalier et al., 2008; Park and Cosgrove, 2012a), suggesting that the activity of XXT1 and XXT2 are required for XyG synthesis, delivery, and/or stability.Much attention has been paid to the interactions between cellulose and XyG over the past 40 years. Currently, there are several hypotheses concerning the nature of these interactions (Park and Cosgrove, 2015). One possibility is that XyGs bind directly to cellulose microfibrils (CMFs). Recent data indicating that crystalline cellulose cores are surrounded with hemicelluloses support this hypothesis (Dick-Pérez et al., 2011). It is also possible that XyG acts as a spacer-molecule to prevent CMFs from aggregating in cell walls (Anderson et al., 2010) or as an adapter to link cellulose with other cell wall components, such as pectin (Cosgrove, 2005; Cavalier et al., 2008). XyG can be covalently linked to pectin (Thompson and Fry, 2000; Popper and Fry, 2005, 2008), and NMR data demonstrate that pectins and cellulose might interact to a greater extent than XyG and cellulose in native walls (Dick-Pérez et al., 2011). Alternative models exist for how XyG-cellulose interactions influence primary wall architecture and mechanics. One such model posits that XyG chains act as load-bearing tethers that bind to CMFs in primary cell walls to form a cellulose-XyG network (Carpita and Gibeaut, 1993; Pauly et al., 1999; Somerville et al., 2004; Cosgrove, 2005). However, results have been accumulating against this tethered network model, leading to an alternative model in which CMFs make direct contact, in some cases mediated by a monolayer of xyloglucan, at limited cell wall sites dubbed “biomechanical hotspots,” which are envisioned as the key sites of cell wall loosening during cell growth (Park and Cosgrove, 2012a; Wang et al., 2013; Park and Cosgrove, 2015). Further molecular, biochemical, and microscopy experiments are required to help distinguish which aspects of the load-bearing, spacer/plasticizer, and/or hotspot models most accurately describe the functions of XyG in primary walls.Cortical microtubules (MTs) direct CMF deposition by guiding cellulose synthase complexes in the plasma membrane (Baskin et al., 2004; Paredez et al., 2006; Emons et al., 2007; Sánchez-Rodriguez et al., 2012), and the patterned deposition of cellulose in the wall in turn can help determine plant cell anisotropic growth and morphogenesis (Baskin, 2005). Disruption of cortical MTs by oryzalin, a MT-depolymerizing drug, alters the alignment of CMFs, suggesting that MTs contribute to CMF organization (Baskin et al., 2004). CELLULOSE SYNTHASE (CESA) genes, including CESA1, CESA3, and CESA6, are required for normal CMF synthesis in primary cell walls (Kohorn et al., 2006; Desprez et al., 2007), and accessory proteins such as COBRA function in cellulose production (Lally et al., 2001). Live-cell imaging from double-labeled YFP-CESA6; CFP-ALPHA-1 TUBULIN (TUA1) Arabidopsis seedlings provides direct evidence that cortical MTs determine the trajectories of cellulose synthesis complexes (CSCs) and patterns of cellulose deposition (Paredez et al., 2006). Additionally, MT organization affects the rotation of cellulose synthase trajectories in the epidermal cells of Arabidopsis hypocotyls (Chan et al., 2010). Recently, additional evidence for direct guidance of CSCs by MTs has been provided by the identification of CSI1/POM2, which binds to both MTs and CESAs (Bringmann et al., 2012; Li et al., 2012). MICROTUBULE ORGANIZATION1 (MOR1) is essential for cortical MT organization (Whittington et al., 2001), but disruption of cortical MTs in the mor1 mutant does not greatly affect CMF organization (Sugimoto et al., 2003), and oryzalin treatment does not abolish CSC motility (Paredez et al., 2006).Conversely, the organization of cortical MTs can be affected by cellulose synthesis. Treatment with isoxaben, a cellulose synthesis inhibitor, results in disorganized cortical MTs in tobacco cells, suggesting that inhibition of cellulose synthesis affects MT organization (Fisher and Cyr, 1998), and treatment with 2,6-dichlorobenzonitrile, another cellulose synthesis inhibitor, alters MT organization in mor1 plants (Himmelspach et al., 2003). Cortical MT orientation in Arabidopsis roots is also altered in two cellulose synthesis-deficient mutants, CESA6^52-isx and kor1-3, suggesting that CSC activity can affect MT arrays (Paredez et al., 2008). Together, these results point to a bidirectional relationship between cellulose synthesis/patterning and MT organization.MTs influence plant organ morphology, but the detailed mechanisms by which they do so are incompletely understood. The dynamics and stability of cortical MTs are also affected by MT-associated proteins (MAPs). MAP18 is a MT destabilizing protein that depolymerizes MTs (Wang et al., 2007), MAP65-1 functions as a MT crosslinker, and MAP70-1 functions in MT assembly (Korolev et al., 2005; Lucas et al., 2011). MAP70-5 stabilizes existing MTs to maintain their length, and its overexpression induces right-handed helical growth (Korolev et al., 2007); likewise, MAP20 overexpression results in helical cell twisting (Rajangam et al., 2008). CLASP promotes microtubule stability, and its mutant is hypersensitive to microtubule-destabilizing drug oryzalin (Ambrose et al., 2007). KATANIN1 (KTN1) is a MT-severing protein that can sever MTs into short fragments and promote the formation of thick MT bundles that ultimately depolymerize (Stoppin-Mellet et al., 2006), and loss of KTN1 function results in reduced responses to mechanical stress (Uyttewaal et al., 2012). In general, cortical MT orientation responds to mechanical signals and can be altered by applying force directly to the shoot apical meristem (Hamant et al., 2008). The application of external mechanical pressure to Arabidopsis leaves also triggers MT bundling (Jacques et al., 2013). Kinesins, including KINESIN-13A (KIN-13A) and FRAGILE FIBER1 (FRA1), have been implicated in cell wall synthesis (Cheung and Wu, 2011; Fujikura et al., 2014). The identification of cell wall receptors and sensors is beginning to reveal how plant cell walls sense and respond to external signals (Humphrey et al., 2007; Ringli, 2010); some of them, such as FEI1, FEI2, THESEUS1 (THE1), FERONIA (FER), HERCULES RECEPTOR KINASE1 (HERK1), WALL ASSOCIATED KINASE1 (WAK1), WAK2, and WAK4, have been characterized (Lally et al., 2001; Decreux and Messiaen, 2005; Kohorn et al., 2006; Xu et al., 2008; Guo et al., 2009; Cheung and Wu, 2011). However, the relationships between wall integrity, cytoskeletal dynamics, and wall synthesis have not yet been fully elucidated.In this study, we analyzed CMF patterning, MT patterning and dynamics, and cellulose biosynthesis in the Arabidopsis xxt1 xxt2 double mutant that lacks detectable XyG and displays altered growth (Cavalier et al., 2008; Park and Cosgrove, 2012a). To investigate whether and how XyG deficiency affects the organization of CMFs and cortical MTs, we observed CMF patterning in xxt1 xxt2 mutants and Col (wild-type) controls using atomic force microscopy (AFM), field emission scanning electron microscopy (FESEM), transmission electron microscopy (TEM), and confocal microscopy (Hodick and Kutschera, 1992; Derbyshire et al., 2007; Anderson et al., 2010; Zhang et al., 2014). We also generated transgenic Col and xxt1 xxt2 lines expressing GFP-MAP4 (Marc et al., 1998) and GFP-CESA3 (Desprez et al., 2007), and analyzed MT arrays and cellulose synthesis using live-cell imaging. Our results show that the organization of CMFs is altered, that MTs in xxt1 xxt2 mutants are aberrantly organized and are more sensitive to external mechanical pressure and the MT-depolymerizing drug oryzalin, and that cellulose synthase motility and cellulose content are decreased in xxt1 xxt2 mutants. Furthermore, real-time quantitative RT-PCR measurements indicate that the enhanced sensitivity of cortical MTs to mechanical stress and oryzalin in xxt1 xxt2 plants might be due to altered expression of MT-stabilizing and wall receptor genes. Together, these data provide insights into the connections between the functions of XyG in wall assembly, the mechanical integrity of the cell wall, cytoskeleton-mediated cellular responses to deficiencies in wall biosynthesis, and cell and tissue morphogenesis.     

On the Inside

Cellulose synthase complexes (CSCs) at the plasma membrane (PM) are aligned with cortical microtubules (MTs) and direct the biosynthesis of cellulose. The mechanism of the interaction between CSCs and MTs, and the cellular determinants that control the delivery of CSCs at the PM, are not yet well understood. We identified a unique small molecule, CESA TRAFFICKING INHIBITOR (CESTRIN), which reduces cellulose content and alters the anisotropic growth of Arabidopsis (Arabidopsis thaliana) hypocotyls. We monitored the distribution and mobility of fluorescently labeled cellulose synthases (CESAs) in live Arabidopsis cells under chemical exposure to characterize their subcellular effects. CESTRIN reduces the velocity of PM CSCs and causes their accumulation in the cell cortex. The CSC-associated proteins KORRIGAN1 (KOR1) and POM2/CELLULOSE SYNTHASE INTERACTIVE PROTEIN1 (CSI1) were differentially affected by CESTRIN treatment, indicating different forms of association with the PM CSCs. KOR1 accumulated in bodies similar to CESA; however, POM2/CSI1 dissociated into the cytoplasm. In addition, MT stability was altered without direct inhibition of MT polymerization, suggesting a feedback mechanism caused by cellulose interference. The selectivity of CESTRIN was assessed using a variety of subcellular markers for which no morphological effect was observed. The association of CESAs with vesicles decorated by the trans-Golgi network-localized protein SYNTAXIN OF PLANTS61 (SYP61) was increased under CESTRIN treatment, implicating SYP61 compartments in CESA trafficking. The properties of CESTRIN compared with known CESA inhibitors afford unique avenues to study and understand the mechanism under which PM-associated CSCs are maintained and interact with MTs and to dissect their trafficking routes in etiolated hypocotyls.Plant cell expansion and anisotropic cell growth are driven by vacuolar turgor pressure and cell wall extensibility, which in a dynamic and restrictive manner direct cell morphogenesis (Baskin, 2005). Cellulose is the major load-bearing component of the cell wall and is thus a major determinant for anisotropic growth (Baskin, 2001). Cellulose is made up of β-1,4-linked glucan chains that may aggregate to form microfibrils holding 18 to 36 chains (Somerville, 2006; Fernandes et al., 2011; Jarvis, 2013; Newman et al., 2013; Thomas et al., 2013). In contrast to cell wall structural polysaccharides, including pectin and hemicellulose, which are synthesized by Golgi-localized enzymes, cellulose is synthesized at the plasma membrane (PM) by cellulose synthase complexes (CSCs; Somerville, 2006; Scheller and Ulvskov, 2010; Atmodjo et al., 2013). The cellulose synthases (CESAs) are the principal catalytic units of cellulose biosynthesis and in higher plants are organized into globular rosettes (Haigler and Brown, 1986). For their biosynthetic function, each primary cell wall CSC requires a minimum of three catalytic CESA proteins (Desprez et al., 2007; Persson et al., 2007).On the basis of observations that cellulose microfibrils align with cortical microtubules (MTs) and that MT disruption leads to a loss of cell expansion, it was hypothesized that cortical MTs guide the deposition and, therefore, the orientation of cellulose (Green, 1962; Ledbetter and Porter, 1963; Baskin, 2001; Bichet et al., 2001; Sugimoto et al., 2003; Baskin et al., 2004; Wasteneys and Fujita, 2006). Confocal microscopy of CESA fluorescent fusions has advanced our understanding of CESA trafficking and dynamics. CSCs are visualized as small particles moving within the plane of the PM, with an average velocity of approximately 200 to 400 nm min⁻¹. Their movement in linear tracks along cortical MTs (Paredez et al., 2006) supports the MT-cellulose alignment hypothesis.Our current understanding of cellulose synthesis suggests that CESAs are assembled into CSCs in either the endoplasmic reticulum (ER) or the Golgi apparatus and trafficked by vesicles to the PM (Bashline et al., 2014; McFarlane et al., 2014). The presence of CESAs in isolated Golgi and vesicles from the trans-Golgi network (TGN) has been established by proteomic studies (Dunkley et al., 2006; Drakakaki et al., 2012; Nikolovski et al., 2012; Parsons et al., 2012; Groen et al., 2014). Their localization at the TGN has been corroborated by electron microscopy and colocalization with TGN markers, such as vacuolar H⁺-ATP synthase subunit a1 (VHA-a1), and the Soluble NSF Attachment Protein Receptor (SNARE) protein SYNTAXIN OF PLANTS41 (SYP41), SYP42, and SYP61 (Crowell et al., 2009; Gutierrez et al., 2009; Drakakaki et al., 2012). A population of post-Golgi compartments carrying CSCs, referred to as microtubule-associated cellulose synthase compartments (MASCs) or small cellulose synthase compartments (SmaCCs), may be associated with MTs or actin filaments and are thought to be directly involved in either CSC delivery to, or internalization from, the PM (Crowell et al., 2009; Gutierrez et al., 2009).In addition to the CESAs, auxiliary proteins have been identified that play a vital role in the cellulose-synthesizing machinery. These include COBRA (Roudier et al., 2005), the endoglucanase KORRIGAN1 (KOR1; Lane et al., 2001; Lei et al., 2014b; Vain et al., 2014), and the recently identified POM-POM2/CELLULOSE SYNTHASE INTERACTIVE PROTEIN1 (POM2/CSI1; Gu et al., 2010; Bringmann et al., 2012). The latter protein functions as a linker between the cortical MTs and CSCs, as genetic lesions in POM2/CSI1 result in a lower incidence of coalignment between CSCs and cortical MTs (Bringmann et al., 2012). Given the highly regulated process of cellulose biosynthesis and deposition, it can be expected that many more accessory proteins participate in the delivery of CSCs and their interaction with MTs. Identification of these unique CSC-associated proteins can ultimately provide clues for the mechanisms behind cell growth and cell shape formation.Arabidopsis (Arabidopsis thaliana) mutants with defects in the cellulose biosynthetic machinery exhibit a loss of anisotropic growth, which results in organ swelling. This phenotype may be used as a diagnostic tool in genetic screens to identify cellulose biosynthetic and CSC auxiliary proteins (Mutwil et al., 2008). Chemical inhibitors complement genetic lesions to perturb, study, and control the cellular and physiological function of proteins (Drakakaki et al., 2009). A plethora of bioactive small molecules have been identified, and their analytical use contributes to our understanding of cellulose biosynthesis and CESA subcellular behavior (for review, see Brabham and Debolt, 2012). Small molecule treatment can induce distinct characteristic subcellular CESA patterns that can be broadly grouped into three categories (Brabham and Debolt, 2012). The first is characterized by the depletion of CESAs from the PM and their accumulation in cytosolic compartments, as observed for the herbicide isoxaben {N-[3-(1-ethyl-1-methylpropyl)-5-isoxazolyl]-2,6-dimethyoxybenzamide}, CGA 325615 [1-cyclohexyl-5-(2,3,4,5,6-pentafluorophe-noxyl)-1λ4,2,4,6-thiatriazin-3-amine], thaxtomin A (4-nitroindol-3-yl containing 2,5-dioxopiperazine), AE F150944 [N2-(1-ethyl-3-phenylpropyl)-6-(1-fluoro-1-methylethyl)-1,3,5-triazine-2,4-di-amine], and quinoxyphen [4-(2-bromo-4,5-dimethoxyphenyl)-3,4-dihydro-¹H-benzo-quinolin-2-one]; (Paredez et al., 2006; Bischoff et al., 2009; Crowell et al., 2009; Gutierrez et al., 2009; Harris et al., 2012). The second displays hyperaccumulation of CESAs at the PM, as seen for the herbicides dichlobenil (2,6-dichlorobenzonitrile) and indaziflam {N-[(1R,2S)-2,3-dihydro-2,6-dimethyl-¹H-inden-1-yl)-6-(1-fluoroethyl]-1,3,5-triazine-2,4-diamine} (Herth, 1987; DeBolt et al., 2007b; Brabham et al., 2014). The third exhibits disturbance of both CESAs and MTs and alters CESA trajectories at the PM, as exemplified by morlin (7-ethoxy-4-methylchromen-2-one; DeBolt et al., 2007a). Unique compounds inducing a phenotype combining CESA accumulation in intermediate compartments and disruption of CSC-MT interactions can contribute to both the identification of the accessory proteins linking CSCs with MTs and the vesicular delivery mechanisms of CESAs.In this study, we identified and characterized a unique cellulose deposition inhibitor, the small molecule CESA TRAFFICKING INHIBITOR (CESTRIN), which affects the localization pattern of CSCs and their interacting proteins in a unique way. The induction of cytoplasmic CESTRIN bodies might provide further clues for trafficking routes that carry CESAs to the PM.     

Distribution of Transglutaminase in Pear Pollen Tubes in Relation to Cytoskeleton and Membrane Dynamics

Transglutaminases (TGases) are ubiquitous enzymes that take part in a variety of cellular functions. In the pollen tube, cytoplasmic TGases are likely to be involved in the incorporation of primary amines at selected peptide-bound glutamine residues of cytosolic proteins (including actin and tubulin), while cell wall-associated TGases are believed to regulate pollen tube growth. Using immunological probes, we identified TGases associated with different subcellular compartments (cytosol, membranes, and cell walls). Binding of cytosolic TGase to actin filaments was shown to be Ca²⁺ dependent. The membrane TGase is likely associated with both Golgi-derived structures and the plasma membrane, suggesting a Golgi-based exocytotic delivery of TGase. Association of TGase with the plasma membrane was also confirmed by immunogold transmission electron microscopy. Immunolocalization of TGase indicated that the enzyme was present in the growing region of pollen tubes and that the enzyme colocalizes with cell wall markers. Bidimensional electrophoresis indicated that different TGase isoforms were present in distinct subcellular compartments, suggesting either different roles or different regulatory mechanisms of enzyme activity. The application of specific inhibitors showed that the distribution of TGase in different subcellular compartments was regulated by both membrane dynamics and cytoskeleton integrity, suggesting that delivery of TGase to the cell wall requires the transport of membranes along cytoskeleton filaments. Taken together, these data indicate that a cytoplasmic TGase interacts with the cytoskeleton, while a different TGase isoform, probably delivered via a membrane/cytoskeleton-based transport system, is secreted in the cell wall of pear (Pyrus communis) pollen tubes, where it might play a role in the regulation of apical growth.Transglutaminases (TGases [EC 2.3.2.13]; protein-Gln γ-glutamyltransferase) are a family of ubiquitous Ca²⁺-activated enzymes that are involved in animal cell morphogenesis and differentiation, apoptosis, cell death, inflammation, cell migration, and wound healing (Griffin et al., 2002; Mehta et al., 2006; Beninati et al., 2009). TGases are associated with different subcellular compartments, such as cytosol, plasma membrane, nucleus, mitochondria, and extracellular matrix. The specific localization of TGases is likely to determine both the biochemical activity and the type of proteins and/or substrates with which TGases react (Park et al., 2010). The distribution profile of TGase is affected by Ca²⁺, since the enzyme is preferentially associated with the lysosome compartment of liver cells in the absence of Ca²⁺ (Juprelle-Soret et al., 1984).TGase was initially detected in association with the cytosol, with the particulate (probably the microsomal) fraction (Birckbichler et al., 1976), and with the nucleus of animal cells (Remington and Russell, 1982). The association of TGase with the plasma membrane was related to its activity in promoting cell adhesion and to the interaction of cells with the extracellular matrix, while the presence of TGase in the nucleus is likely related to cell apoptosis (Griffin et al., 2002). How TGase is delivered to its final destination in animal cells remains to be clarified. Since the cytoskeleton is essential for the correct positioning of proteins in the cells, this interplay has often been studied in terms of potential substrates of TGase activity (Griffin et al., 2002). For example, the TGase-mediated incorporation of polyamines (PAs) stimulates actin polymerization (Takashi, 1988; Griffin et al., 2002). TGase was also found to associate with myosin in stress fibers of vascular smooth cells (Chowdhury et al., 1997). The association between TGase and microtubules (MTs) was initially studied in view of the importance of MTs in Alzheimer’s disease (Griffin et al., 2002), whereas the dynamics of MTs is also likely to be controlled by TGase (Al-Jallad et al., 2011). Interestingly, MTs are also a substrate of TGase activity in cells committed to apoptosis (Piredda et al., 1999). TGase was also shown to posttranslationally modify MT-associated proteins such as tau (Griffin et al., 2002).Information about the localization and function of TGases in plant cells is limited. Following the early evidence of an enzyme-based incorporation of PAs in plants (Serafini-Fracassini et al., 1988), a number of reports described the presence and role of TGase in nonphotosynthetic/photosynthetic tissues and in isolated chloroplasts (Serafini-Fracassini and Del Duca, 2008, and refs. therein). Attempts have also been made to examine the differences and similarities between plant and animal TGases. For example, a tobacco (Nicotiana tabacum) TGase was proposed to be involved in the programmed cell death (PCD) of the flower corolla (Della Mea et al., 2007); in such a case, TGase is likely to be released into the cell wall by a Golgi vesicle-based transport. Plant TGases might also be involved in protection against viruses (Del Duca et al., 2007) and in the self-incompatibility (SI) response involving pollen and stigma during sexual reproduction (Del Duca et al., 2010). Recently, different TGase isoforms were detected in meristematic apices of Jerusalem artichoke (Helianthus tuberosus) tuber sprouts (Beninati et al., 2013).The pollen tube is a widely investigated tip-growing plant cell (Lee and Yang, 2008). Studies are generally aimed at clarifying the many aspects related either to its growth or to rejection by the stigma/style. Early evidence for a role of PAs during pollen tube emergence (Bagni et al., 1981) was confirmed through the detection of PA binding via a Ca²⁺-activated TGase activity (Del Duca et al., 1997) and later by the identification of actin and tubulin as substrates of purified pollen TGase (Del Duca et al., 2009). In pollen, the enzyme affected the polymerization state and activity of actin filaments (AFs) and MTs (Del Duca et al., 2009) and existed as both soluble and cell wall associated (Di Sandro et al., 2010). Visualization of fluorescently labeled TGase products indicated that the cross-linking activity of TGase occurred at the apex of pollen tubes, in a basal region close to the pollen grain and within the pollen grain itself (Iorio et al., 2008). The enzyme was found as a soluble cytoplasmic form likely involved in the regulation of unspecified physiological processes (possibly associated with the cytoskeleton; Del Duca et al., 2009).Although the association of pollen TGases with organelles/vesicles has not been reported, an extracellular form of a Ca²⁺-dependent TGase was shown to be involved in pollen tube growth (likely as a modulator of cell wall building and strengthening). Moreover, pollen TGase was secreted in the incubation medium during germination, where it might catalyze the cross linking of PAs with secreted proteins (Di Sandro et al., 2010). This suggests that pollen TGase may be secreted through a vesicle-based mechanism. Finally, a TGase activity was also observed in planta, consistent with a possible role of TGase during tube migration through the style (Di Sandro et al., 2010) or in the SI response of pollen tubes (Del Duca et al., 2010).The pollen tube is an excellent model to study how a given plant protein is either secreted or delivered to its final destination. Although we know that actin and tubulin are substrates of TGase activity, and that the active enzyme is located in the cell wall and released outside, how TGase is distributed in the cells and how this process is dependent on cytoskeleton and membrane dynamics remain unknown. Here, we wanted to study in detail the localization and distribution of TGase in growing pollen tubes of pear (Pyrus communis) in relation to both cytoskeleton and membrane dynamics. The aim was to shed light on the mechanism by which TGase is transported and secreted, a process that is still not well understood even in animal cells. Specific antibodies that cross react with the TGase of pollen tubes were used to localize the enzyme in different membrane compartments and in the cell wall. The use of specific inhibitors indicated that the delivery of extracellular TGase is dependent on both AFs and membrane dynamics. Analysis by bidimensional electrophoresis (2-DE) showed that distinct TGase isoforms are associated with different cell compartments, suggesting that TGase might be differently regulated according to its position in the cell. Together, these data may contribute to our understanding of the mechanisms underlying pollen tube growth, an essential aspect of fertilization processes.     

Noninvasive Evaluation of Heavy Metal Uptake and Storage in Micoralgae Using a Fluorescence Resonance Energy Transfer-Based Heavy Metal Biosensor

We have developed a fluorescence resonance energy transfer (FRET)-based heavy metal biosensor for the quantification of bioavailable free heavy metals in the cytoplasm of the microalga Chlamydomonas reinhardtii. The biosensor is composed of an end-to-end fusion of cyan fluorescent protein (CFP), chicken metallothionein II (MT-II), and yellow fluorescent protein (YFP). In vitro measurements of YFP/CFP fluorescence emission ratios indicated that the addition of metals to the purified biosensor enhanced FRET between CFP and YFP, consistent with heavy metal-induced folding of MT-II. A maximum YFP/CFP FRET ratio of 2.8 was observed in the presence of saturating concentrations of heavy metals. The sensitivity of the biosensor was greatest for Hg²⁺ followed by Cd²⁺ ≈ Pb²⁺ > Zn²⁺ > Cu²⁺. The heavy metal biosensor was unresponsive to metals that do not bind to MT-II (Na⁺ and Mg²⁺). When expressed in C. reinhardtii, we observed a differential metal-dependent response to saturating external concentrations (1.6 mm) of heavy metals (Pb²⁺ > Cd²⁺) that was unlike that observed for the isolated biosensor (in vitro). Significantly, analysis of metal uptake kinetics indicated that equilibration of the cytoplasm with externally applied heavy metals occurred within seconds. Our results also indicated that algae have substantial buffering capacity for free heavy metals in their cytosol, even at high external metal concentrations.Many proteins utilize metals to stabilize their structures or as cofactors to catalyze redox and other chemical reactions. Metals such as zinc, copper, iron, magnesium, cobalt, and manganese are required by most living organisms for their normal cellular functions. Essential metals are often present at low concentrations in the environment, however, and must be imported into cells, often at the expense of energy (Hanikenne et al., 2005; Merchant et al., 2006). In contrast to essential metals, toxic metals such as cadmium, lead, and mercury can disrupt cellular functions by competing with essential metals for their metal-binding sites and/or by altering the redox state of cells. Exposure of organisms to high concentrations of toxic metals can impair their cellular functions, growth, and reproduction. To prevent metal-induced cellular anomalies, organisms have evolved a variety of strategies to reduce the toxicity of heavy metals. One such strategy involves the selective binding of toxic metals in the cytoplasm by metal-binding proteins and other small molecules. As discussed below, both enzymatically and ribosomally synthesized Cys-rich peptides, including phytochelatins and metallothioneins (MTs), are utilized by a variety of organisms to sequester toxic heavy metals, including cadmium, mercury, lead, silver, and gold. The peptides may also serve as storage reserves for essential metals such as copper and zinc (Cobbett and Goldsbrough, 2002).Phytochelatins are enzymatically synthesized polypeptides containing repeating units of (γ-Glu-Cys)_n-Gly, where n = 2 to 11 (Rauser, 1990), whereas MTs are genetically encoded, ribosomally synthesized polypeptides (Cobbett and Goldsbrough, 2002). MTs have molecular mass values ranging from 6 to 7 kD and contain approximately 20 conserved Cys residues (Cobbett and Goldsbrough, 2002; Romero-Isart and Vasák, 2002). Metals are characteristically bound to MT via the thiolate sulfur ligands of Cys residues (Kägi, 1991). It is estimated that the metal-saturated MT contains about 10% thiolate sulfur and bound metals by mass (Romero-Isart and Vasák, 2002). Structural analyses of metal-free and metal-complexed MTs demonstrated that MTs undergo a structural transition from a metal-free random-coil structure to a metal-bound compact dumbbell-shaped structure having metal saturated α- and β-domains (Pearce et al., 2000; Romero-Isart and Vasak, 2002; Hong and Maret, 2003). The N-terminal β-domain binds three metal ion equivalents, and the C-terminal α-domain binds four metal ion equivalents (Romero-Isart and Vasák, 2002; Vasák, 2005). Furthermore, several decades of work on MTs have provided a great deal of information regarding their metal-binding affinity, specificity, and domain selectivity for select metals (Cobbett and Goldsbrough, 2002; Romero-Isart and Vasák, 2002; Vasák, 2005).Fluorescence resonance energy transfer (FRET) involves the nonradioactive transfer of energy between the excited state of a luminescent or fluorescent donor molecule and a nearby acceptor molecule that has overlapping excited state transitions. Proteins that are modified to have efficient energy donor and acceptor domains and that undergo structural changes upon binding a specific ligand are good candidates for FRET-based sensors. For ligand-specific FRET-based biosensors, the distance and/or the orientation between the energy donor and acceptor molecules is changed upon ligand binding in a concentration-dependent manner (Selvin, 1995; Weiss, 2000; Hong and Maret, 2003; Looger et al., 2005). Relevant to this discussion, a FRET-based biosensor with GFP variants fused to MT was previously shown to be an effective means to monitor metal release during nitric oxide-induced signaling in endothelial cells (Pearce et al., 2000).Unicellular algae such as Chlamydomonas species are often found in areas that might be contaminated by toxic heavy metals (Merchant et al., 2006). Chlamydomonas species have also been shown to sequester toxic metals (e.g. cadmium and mercury) and have potential use for bioremediation of these metals (Cai et al., 1999; Adhiya et al., 2002; Siripornadulsil et al., 2002; He et al., 2011; Priyadarshani et al., 2011). To determine the kinetics and selectivity of exogenous heavy metal uptake as well as free heavy metal concentration in the cytoplasm of Chlamydomonas species, we developed an MT, FRET-based metal-binding sensor and expressed this in the cytoplasm of the unicellular green alga Chlamydomonas reinhardtii. We demonstrate that heavy metal uptake is rapid in C. reinhardtii and that cytoplasmic free heavy metal concentrations are substantially lower than exogenous free heavy metal concentrations, implying that heavy metals are rapidly sequestered by various biological molecules in the cell.     

β-Galactosyl Yariv Reagent Binds to the β-1,3-Galactan of Arabinogalactan Proteins

Yariv phenylglycosides [1,3,5-tri(p-glycosyloxyphenylazo)-2,4,6-trihydroxybenzene] are a group of chemical compounds that selectively bind to arabinogalactan proteins (AGPs), a type of plant proteoglycan. Yariv phenylglycosides are widely used as cytochemical reagents to perturb the molecular functions of AGPs as well as for the detection, quantification, purification, and staining of AGPs. However, the target structure in AGPs to which Yariv phenylglycosides bind has not been determined. Here, we identify the structural element of AGPs required for the interaction with Yariv phenylglycosides by stepwise trimming of the arabinogalactan moieties using combinations of specific glycoside hydrolases. Whereas the precipitation with Yariv phenylglycosides (Yariv reactivity) of radish (Raphanus sativus) root AGP was not reduced after enzyme treatment to remove α-l-arabinofuranosyl and β-glucuronosyl residues and β-1,6-galactan side chains, it was completely lost after degradation of the β-1,3-galactan main chains. In addition, Yariv reactivity of gum arabic, a commercial product of acacia (Acacia senegal) AGPs, increased rather than decreased during the repeated degradation of β-1,6-galactan side chains by Smith degradation. Among various oligosaccharides corresponding to partial structures of AGPs, β-1,3-galactooligosaccharides longer than β-1,3-galactoheptaose exhibited significant precipitation with Yariv in a radial diffusion assay on agar. A pull-down assay using oligosaccharides cross linked to hydrazine beads detected an interaction of β-1,3-galactooligosaccharides longer than β-1,3-galactopentaose with Yariv phenylglycoside. To the contrary, no interaction with Yariv was detected for β-1,6-galactooligosaccharides of any length. Therefore, we conclude that Yariv phenylglycosides should be considered specific binding reagents for β-1,3-galactan chains longer than five residues, and seven residues are sufficient for cross linking, leading to precipitation of the Yariv phenylglycosides.Arabinogalactan proteins (AGPs) are a type of plant proteoglycans consisting of a Hyp-rich core protein and large arabinogalactan (AG) moieties (Fincher et al., 1983; Nothnagel, 1997). Although there are many molecular species of AGP differentiated by their core proteins, the AG moieties commonly comprise β-1,3-galactan main chains and β-1,6-galactan side chains, to which l-Ara and other auxiliary sugars, such as GlcA, 4-O-methyl-GlcA, l-Fuc, l-Rha, and Xyl, are attached (Fincher et al., 1983; Nothnagel, 1997; Seifert and Roberts, 2007). A commercial product of AGPs prepared from the acacia (Acacia senegal) tree is known as gum arabic and utilized as a food stabilizer. In the Japanese herbal remedy Juzen-Taiho-To, AGs from Astragalus membranaceus are the active ingredient (Majewska-Sawka and Nothnagel, 2000; Kiyohara et al., 2002). In intact plants, AGPs are implicated in various physiological events and serve as extracellular constituents and signaling molecules. For instance, an AGP from stylar transmitting tissue attracts pollen tubes and stimulates their elongation in tobacco (Nicotiana tabacum; Cheung et al., 1995).Yariv phenylglycosides [1,3,5-tri(p-glycosyloxyphenylazo)-2,4,6-trihydroxybenzene] are a group of chemical compounds that were initially developed as carbohydrate antigens for the purification of anti-glycoside antibody and sugar-binding protein (Yariv et al., 1962, 1967a). It then turned out that Yariv phenylglycosides specifically precipitate AGPs (Yariv et al., 1967b; Jermyn and Yeow, 1975). The specific interaction of AGPs with Yariv phenylglycosides forming brown-red precipitate is called Yariv reactivity and has been recognized as an important criterion in the definition of AGPs, even though a number of AGPs do not exhibit Yariv reactivity. Nevertheless, the structure involved in the interaction with Yariv phenylglycoside is presumed to be conserved in many AGPs. The interaction of Yariv phenylglycosides with AGP depends on the glycosyl residues attached to the phenylazotrihydroxybenzene core. In particular, β-glucosyl Yariv phenylglycoside (β-Glc-Yariv) and β-galactosyl Yariv phenylglycoside (β-Gal-Yariv) bind to AGPs, whereas α-glucosyl Yariv and α-galactosyl Yariv (α-Gal-Yariv) do not bind to AGPs (Jermyn and Yeow, 1975; Larkin, 1977, 1978; Nothnagel and Lyon, 1986). Because of the specific interaction with the β-glycosyl Yariv phenylglycosides (β-Yarivs), AGPs were formerly called “β-lectins” (Jermyn and Yeow, 1975; Gleeson and Jermyn, 1979; Nothnagel and Lyon, 1986).The β-Yarivs are useful tools for staining, detection, and quantification of AGPs. Using β-Glc-Yariv, β-lectins were shown to exist in angiosperm, gymnosperm, fern, moss, and liverwort, illustrating the wide distribution of AGPs in the plant kingdom (Jermyn and Yeow, 1975; Clarke et al., 1978). In addition, β-Yarivs are also used as chemical reagents in the purification of AGPs. A nonclassical AGP, xylogen, which is a signaling molecule inducing the differentiation to tracheary elements, has been purified from the culture medium of zinnia (Zinnia elegans) cells by precipitation with β-Glc-Yariv (Motose et al., 2004). As the treatment with β-Yarivs causes the perturbation of various physiological processes in plants, β-Yarivs are reliable cytochemical reagents to explore AGP functions. Application of β-Yarivs to cultured cells of Arabidopsis (Arabidopsis thaliana) induced programmed cell death, demonstrating the involvement of AGPs in the determination of cell fate (Gao and Showalter, 1999). In tobacco cultured cells, the treatment with β-Yarivs has indicated a possible role of AGPs in the orientation of cortical microtubules and the polymerization of F-actin (Sardar et al., 2006).Although Yariv phenylglycosides have been extensively utilized in studies of AGPs over 40 years, the identification of the target structures on AGPs required for β-Yariv reactivity remains elusive (Nothnagel, 1997; Seifert and Roberts, 2007). It has been proposed that β-Yarivs bind to the Hyp-rich core protein, based on the observation that deglycosylation treatment with hydrogen fluoride did not abolish the Yariv reactivity of gum arabic and a tobacco AGP (Akiyama et al., 1984). To the contrary, other reports have asserted the importance of the carbohydrate moieties for Yariv reactivity (Komalavilas et al., 1991). However, with regard to the specific carbohydrate structure required for interaction with β-Yarivs, the results were not always consistent: neither α-l-arabinofuranosyl residues nor β-1,6-galactan side chains were found to be involved in Yariv reactivity of AGPs from Gladiolus spp., radish (Raphanus sativus), and grape (Vitis vinifera; Gleeson and Clarke, 1979; Tsumuraya et al., 1987; Saulnier et al., 1992); partial acid hydrolysis to remove α-l-arabinofuranosyl residues diminished Yariv reactivity of a rose (Rosa spp.) AGP (Komalavilas et al., 1991); and mugwort (Artemisia vulgaris) pollen O-glycans consisting of a β-1,6-galactan core and branched α-l-arabinofuranosyl side chains precipitated with β-Glc-Yariv (Léonard et al., 2005). Accordingly, it has also been suggested that Yariv reactivity depends on the overall physical and chemical properties rather than a specific structural feature of AGPs.In this study, we demonstrate that the peptide component of AGPs is not required for Yariv reactivity. By sequentially trimming the AG moieties of AGPs with sets of specific glycoside hydrolases, we show that β-Gal-Yariv binds to the β-1,3-galactan main chains of radish root AGP. We confirm that β-1,6-galactan side chains are not necessary for Yariv reactivity, we identify β-1,3-galactopentaose (β-1,3-Gal₅) as the smallest carbohydrate structure to interact with β-Gal-Yariv, and we show that β-1,3-galactoheptaose (β-1,3-Gal₇) or longer β-1,3-galactosyl chains are required for the formation of insoluble precipitate with Yariv phenylglycoside. Based on computational modeling, a possible interaction mechanism between β-Gal-Yariv and β-1,3-galactan is suggested.     

Patterning and Lifetime of Plasma Membrane-Localized Cellulose Synthase Is Dependent on Actin Organization in Arabidopsis Interphase Cells

The actin and microtubule cytoskeletons regulate cell shape across phyla, from bacteria to metazoans. In organisms with cell walls, the wall acts as a primary constraint of shape, and generation of specific cell shape depends on cytoskeletal organization for wall deposition and/or cell expansion. In higher plants, cortical microtubules help to organize cell wall construction by positioning the delivery of cellulose synthase (CesA) complexes and guiding their trajectories to orient newly synthesized cellulose microfibrils. The actin cytoskeleton is required for normal distribution of CesAs to the plasma membrane, but more specific roles for actin in cell wall assembly and organization remain largely elusive. We show that the actin cytoskeleton functions to regulate the CesA delivery rate to, and lifetime of CesAs at, the plasma membrane, which affects cellulose production. Furthermore, quantitative image analyses revealed that actin organization affects CesA tracking behavior at the plasma membrane and that small CesA compartments were associated with the actin cytoskeleton. By contrast, localized insertion of CesAs adjacent to cortical microtubules was not affected by the actin organization. Hence, both actin and microtubule cytoskeletons play important roles in regulating CesA trafficking, cellulose deposition, and organization of cell wall biogenesis.Plant cells are surrounded by a flexible yet durable extracellular matrix that makes up the cell wall. This structure offers mechanical strength that counters osmotically driven turgor pressure, is an important factor for water movement in plants, acts as a physical barrier against pathogens (Somerville et al., 2004), and is a determining factor for plant cell morphogenesis. Hence, the cell wall plays a central role in plant biology.Two main types of cell walls can typically be distinguished: the primary and the secondary cell wall. The major load-bearing component in both of these cell walls is the β-1,4-linked glucan polymer cellulose (Somerville et al., 2004). Cellulose polymers are synthesized by plasma membrane (PM)-localized cellulose synthase (CesA) complexes (Mueller and Brown, 1980), which contain several CesA subunits with similar amino acid sequences (Mutwil et al., 2008a). The primary wall CesA complexes are believed to be assembled in the Golgi and are subsequently delivered to the PM via vesicular trafficking (Gutierrez et al., 2009), sometimes associated with Golgi pausing (Crowell et al., 2009). Furthermore, the primary wall CesA complexes are preferentially inserted into the PM at sites that coincide with cortical microtubules (MTs), which subsequently guide cellulose microfibril deposition (Gutierrez et al., 2009). Hence, the cortical MT array is a determinant for multiple aspects of primary wall cellulose production.The actin cytoskeleton plays a crucial role in organized deposition of cell wall polymers in many cell types, including cellulose-related polymers and pectins in tip-growing cells, such as pollen tubes and root hairs (Hu et al., 2003; Chen et al., 2007). Thus, actin-depolymerizing drugs and genetic manipulation of ACTIN genes impair directed expansion of tip-growing cells and long-distance transport of Golgi bodies with vesicles to growing regions (Ketelaar et al., 2003; Szymanski, 2005). In diffusely growing cells in roots and hypocotyls, loss of anisotropic growth has also been observed in response to mutations to vegetative ACTIN genes and to actin-depolymerizing and -stabilizing drugs (Baluska et al., 2001; Kandasamy et al., 2009). While actin is clearly important for cell wall assembly, it is less clear what precise roles it plays.One well-known function of actin in higher plants is to support intracellular movement of cytoplasmic organelles via actomyosin-based motility (Geisler et al., 2008; Szymanski, 2009). During primary wall synthesis in interphase cells, treatment with the actin assembly inhibitor latrunculin B (LatB) led to inhibition of Golgi motility and pronounced inhomogenities in CesA density at the PM (Crowell et al., 2009; Gutierrez et al., 2009) that coincided with the density of underlying and immobile Golgi bodies (Gutierrez et al., 2009). These results suggested that Golgi motility is important for CesA distribution (Gutierrez et al., 2009). The actin cytoskeleton also appears to be important for secondary wall cellulose microfibril deposition. For example, longitudinal actin filaments (AFs) define the movement of secondary wall CesA-containing Golgi bodies in developing xylem vessels (Wightman and Turner, 2008). In addition, it has been proposed that the AFs also can regulate the delivery of the secondary wall CesA complex to the PM via pausing of the Golgi (Wightman and Turner, 2008). It is therefore clear that actin organization is important for CesA distribution and for the pattern of cellulose microfibril deposition.Despite the above findings, very few reports have undertaken detailed studies to elucidate the role of the actin cytoskeleton in the distribution and trafficking of specific proteins in plant cells. Here, we have investigated the intracellular trafficking of CesA-containing vesicles and delivery of CesAs to the PM, in the context of the actin cytoskeleton. We quantitatively demonstrate that the organization of the actin cytoskeleton regulates CesA-containing Golgi distribution and the exocytic and endocytic rate of the CesAs. However, actin organization has no effect on the localized insertion of CesAs at sites of MTs at the PM.     

Hitching a Ride on Vesicles: Cauliflower Mosaic Virus Movement Protein Trafficking in the Endomembrane System

The transport of a viral genome from cell to cell is enabled by movement proteins (MPs) targeting the cell periphery to mediate the gating of plasmodesmata. Given their essential role in the development of viral infection, understanding the regulation of MPs is of great importance. Here, we show that cauliflower mosaic virus (CaMV) MP contains three tyrosine-based sorting signals that interact with an Arabidopsis (Arabidopsis thaliana) μA-adaptin subunit. Fluorophore-tagged MP is incorporated into vesicles labeled with the endocytic tracer N-(3-triethylammoniumpropyl)-4-(6-(4-(diethylamino)phenyl)hexatrienyl)pyridinium dibromide. The presence of at least one of the three endocytosis motifs is essential for internalization of the protein from the plasma membrane to early endosomes, for tubule formation, and for CaMV infection. In addition, we show that MP colocalizes in vesicles with the Rab GTPase AtRAB-F2b, which is resident in prevacuolar late endosomal compartments that deliver proteins to the vacuole for degradation. Altogether, these results demonstrate that CaMV MP traffics in the endocytic pathway and that virus viability depends on functional host endomembranes.Membrane trafficking is essential in eukaryotic cells. Cellular membranes serve as a delivery system for newly synthesized proteins such as transporters and receptors exiting the endoplasmic reticulum after proper folding. They then transit through the Golgi complex, reaching the plasma membrane (PM) or the tonoplast via intermediate endomembrane compartments. Receptors and transporters returning from the PM are either recycled or targeted to the vacuole for degradation. Delivery and recycling sorting pathways overlap in the trans-Golgi network (TGN)/early endosome (EE), an intermediate compartment for both exocytosis and endocytosis (Reyes et al., 2011). In plant systems, the endoplasmic reticulum and PM provide membrane continuity between cells through the connections made by plasmodesmata (PD), cytoplasmic channels that regulate traffic in the symplasm (Maule et al., 2011).The selective transport of macromolecules between different compartments of the endomembrane system is mediated by coat proteins promoting the generation of small cargo-trafficking coated vesicles (Spang, 2008). The recognition and recruitment of cargo proteins are mediated by so-called adaptor complexes (AP complexes [AP-1–AP-4]; Robinson, 2004) one of which, AP-1, is localized on the TGN/EE and endosomes, whereas AP-2 is in the PM. The μ-subunit of AP complexes is devoted to cargo protein selection via a specific and well-characterized interaction with a Tyr-sorting signal, YXXΦ, where Φ is a bulky hydrophobic residue and X is any amino acid (Bonifacino and Dell’Angelica, 1999). YXXΦ motifs are present in the cytoplasmic tail of many proteins integral to the PM and TGN/EE and have been found in the movement proteins (MPs) of some viruses (Laporte et al., 2003; Haupt et al., 2005). Plant viruses are obligate parasites that exploit host components to move within the cell and from cell to cell into the vascular system for systemic invasion of the host. Virus movement, which requires the passage of macromolecules through PD connections, is mediated by one or more virus-encoded MPs with the help of the host cytoskeleton and/or endomembranes (Harries et al., 2010). While most MPs act to increase the size exclusion limit of PD to facilitate the passage of the viral nucleoprotein complex, other MPs are assembled in tubules that pass inside highly modified PD and transport encapsidated particles through their lumen.Here, we focus on this second group of tubule-forming MPs and examine the intracellular trafficking of cauliflower mosaic virus (CaMV) MP. The MP encoded by CaMV forms tubules guiding encapsidated virus particle cell-to-cell transport via an indirect MP-virion interaction (Stavolone et al., 2005; Sánchez-Navarro et al., 2010). However, how CaMV MP (and the other tubule-forming MPs) targets the PM and forms tubules remains to be elucidated. Tubule-forming MPs do not require an intact cytoskeleton for PM targeting (Huang et al., 2000; Pouwels et al., 2002) and/or tubule formation (Laporte et al., 2003). However, suppression of tubule formation upon treatment with brefeldin A (BFA), a specific inhibitor of secretion or endocytosis, suggests the involvement of the endomembrane system in correct functioning of some tubule-forming MPs (Huang et al., 2000; Laporte et al., 2003). In this study, we examined the three Tyr-sorting motifs in CaMV MP and show that each of the three domains interacts directly with subunit μ of an Arabidopsis (Arabidopsis thaliana) AP complex. Mutations in these domains revert in the viral context to maintain CaMV viability. MP is found in endosomal compartments labeled by AtRAB-F2b (ARA7) and N-(3-triethylammoniumpropyl)-4-(6-(4-(diethylamino)phenyl)hexatrienyl)pyridinium dibromide (FM4-64). The presence of at least one functional YXXΦ domain is essential for the localization of MP to endosomes and for tubule assembly but is not required for MP targeting to the PM. We provide several lines of evidence to show CaMV MP trafficking in the endocytic pathway. Our findings are discussed in the light of the recent demonstration that the TGN/EE functions as a major hub controlling secretory and endocytic pathways in plants.     

Rapid Severing and Motility of Chloroplast-Actin Filaments Are Required for the Chloroplast Avoidance Response in Arabidopsis

Phototropins (phot1 and phot2 in Arabidopsis thaliana) relay blue light intensity information to the chloroplasts, which move toward weak light (the accumulation response) and away from strong light (the avoidance response). Chloroplast-actin (cp-actin) filaments are vital for mediating these chloroplast photorelocation movements. In this report, we examine in detail the cp-actin filament dynamics by which the chloroplast avoidance response is regulated. Although stochastic dynamics of cortical actin fragments are observed on the chloroplasts, the basic mechanisms underlying the disappearance (including severing and turnover) of the cp-actin filaments are regulated differently from those of cortical actin filaments. phot2 plays a pivotal role in the strong blue light–induced severing and random motility of cp-actin filaments, processes that are therefore essential for asymmetric cp-actin formation for the avoidance response. In addition, phot2 functions in the bundling of cp-actin filaments that is induced by dark incubation. By contrast, the function of phot1 is dispensable for these responses. Our findings suggest that phot2 is the primary photoreceptor involved in the rapid reorganization of cp-actin filaments that allows chloroplasts to change direction rapidly and control the velocity of the avoidance movement according to the light’s intensity and position.     

Salicylic Acid Regulates Arabidopsis Microbial Pattern Receptor Kinase Levels and Signaling

In Arabidopsis thaliana, responses to pathogen-associated molecular patterns (PAMPs) are mediated by cell surface pattern recognition receptors (PRRs) and include the accumulation of reactive oxygen species, callose deposition in the cell wall, and the generation of the signal molecule salicylic acid (SA). SA acts in a positive feedback loop with ACCELERATED CELL DEATH6 (ACD6), a membrane protein that contributes to immunity. This work shows that PRRs associate with and are part of the ACD6/SA feedback loop. ACD6 positively regulates the abundance of several PRRs and affects the responsiveness of plants to two PAMPs. SA accumulation also causes increased levels of PRRs and potentiates the responsiveness of plants to PAMPs. Finally, SA induces PRR- and ACD6-dependent signaling to induce callose deposition independent of the presence of PAMPs. This PAMP-independent effect of SA causes a transient reduction of PRRs and ACD6-dependent reduced responsiveness to PAMPs. Thus, SA has a dynamic effect on the regulation and function of PRRs. Within a few hours, SA signaling promotes defenses and downregulates PRRs, whereas later (within 24 to 48 h) SA signaling upregulates PRRs, and plants are rendered more responsive to PAMPs. These results implicate multiple modes of signaling for PRRs in response to PAMPs and SA.     

Structural Characterization of Arabidopsis Leaf Arabinogalactan Polysaccharides

Proteins decorated with arabinogalactan (AG) have important roles in cell wall structure and plant development, yet the structure and biosynthesis of this polysaccharide are poorly understood. To facilitate the analysis of biosynthetic mutants, water-extractable arabinogalactan proteins (AGPs) were isolated from the leaves of Arabidopsis (Arabidopsis thaliana) plants and the structure of the AG carbohydrate component was studied. Enzymes able to hydrolyze specifically AG were utilized to release AG oligosaccharides. The released oligosaccharides were characterized by high-energy matrix-assisted laser desorption ionization-collision-induced dissociation mass spectrometry and polysaccharide analysis by carbohydrate gel electrophoresis. The Arabidopsis AG is composed of a β-(1→3)-galactan backbone with β-(1→6)-d-galactan side chains. The β-(1→6)-galactan side chains vary in length from one to over 20 galactosyl residues, and they are partly substituted with single α-(1→3)-l-arabinofuranosyl residues. Additionally, a substantial proportion of the β-(1→6)-galactan side chain oligosaccharides are substituted at the nonreducing termini with single 4-O-methyl-glucuronosyl residues via β-(1→6)-linkages. The β-(1→6)-galactan side chains are occasionally substituted with α-l-fucosyl. In the fucose-deficient murus1 mutant, AGPs lack these fucose modifications. This work demonstrates that Arabidopsis mutants in AGP structure can be identified and characterized. The detailed structural elucidation of the AG polysaccharides from the leaves of Arabidopsis is essential for insights into the structure-function relationships of these molecules and will assist studies on their biosynthesis.Arabinogalactans (AGs) are structurally complex large-branched polysaccharides attached to Hyp residues of many plant cell wall polypeptides. Most proteins glycosylated with AGs (AGPs) have both AG glycosylated domains (glycomodules) and structural or enzymatic domains. However, typical AGPs commonly contain less than 10% protein, suggesting that the AG is the functional part of the molecule (Clarke et al., 1979; Fincher et al., 1983; Kieliszewski and Lamport, 1994; Borner et al., 2003; Xu et al., 2008). Hyp is the most characteristic amino acid present at the glycosylated domain of the AGP, but other amino acids such as Ser, Ala, and Thr are also very common. Type II AG polysaccharides share common structural features based on a β-(1→3)-galactan backbone with β-(1→6)-linked galactan side chains and can be found both on AGPs and rhamnogalacturonan-I (RG-I) pectin (Renard et al., 1991). The galactopyranosyl (Galp) residues can be further substituted with l-arabinofuranosyl (l-Araf) and occasionally also l-rhamnosyl (l-Rha), l-fucosyl (l-Fuc), and glucuronosyl (GlcA; with or without 4-O-methylation) residues (Tsumuraya et al., 1988; Tan et al., 2004; Tryfona et al., 2010). (Sugars mentioned in this work belong to the D-series unless otherwise stated.)The structure of AGs is poorly characterized, and this is mainly due to the great heterogeneity of glycan structures, not only between different AGPs but also even on the same peptide sequence in the same tissue (Estévez et al., 2006). The glycan structure can also be different depending on the developmental stage and tissue type (Tsumuraya et al., 1988), adding to the great heterogeneity of these molecules and therefore limiting their detailed characterization. Molecular and biochemical evidence has indicated that AGPs have specific functions during root formation, promotion of somatic embryogenesis (van Hengel et al., 2002), and attraction of pollen tubes to the style (Cheung et al., 1995). In addition, enhanced secretion efficiency or stability in the cell wall are properties that the AG may confer on the glycosylated protein (Borner et al., 2003). However, it has been difficult to differentiate one species of AGP from another in plant tissues and to assign specific roles to individual AGPs.l-Fuc is present in AGPs in Arabidopsis (Arabidopsis thaliana; van Hengel et al., 2002), radish (Raphanus sativus; Nakamura et al., 1984; Tsumuraya et al., 1984a, 1984b, 1988), and several other dicot plants such as thyme (Thymus vulgaris; Chun et al., 2001) and celery (Apium graveolens; Lin et al., 2011). Reduction in l-Fuc by 40% in roots of murus1 (mur1) plants resulted in a decrease of 50% in root cell elongation, and eel lectin binding assays suggested that the phenotype was the result of alterations in the composition of root AGPs (van Hengel and Roberts, 2002). An α-(1→2)-fucosyltransferase (FUT) activity for radish primary root AGPs has been described, where an α-l-Araf-(1→3)-β-Galp-(1→6)-Galp trisaccharide was used as exogenous substrate acceptor to mimic an AG polysaccharide in the enzymatic assay (Misawa et al., 1996). Linkage analysis, reactivity with eel lectin, and digestion with α-(1→2)-fucosidase indicated that the l-Fuc residues added are terminal and attached via an α-linkage to the C-2 position of an adjacent l-Araf residue (Nakamura et al., 1984; Tsumuraya et al., 1984a, 1984b, 1988). Recently, Wu et al. (2010) identified AtFUT4 and AtFUT6 genes encoding FUT proteins specific to AGPs, but the structures of the fucosylated AG generated have not been fully characterized.To gain insights into the synthesis and function of plant AGPs, it would be useful to have mutants altered in their carbohydrate moieties. However, no AG-specific biosynthetic mutants have been characterized, and this, among other reasons, is due to the very limited knowledge of the structure of Arabidopsis AGs (Qu et al., 2008). Moreover, characterization of AG in candidate mutants remains challenging. Even though the structures of some AGs have been proposed using NMR and sugar linkage analyses, the complete structural elucidation of a native AG still remains a formidable task, because NMR spectroscopy and methylation analysis have been largely used to provide information regarding the amount and type of linkages between adjacent glycosyl residues, and AG heterogeneity can confound attempts to build complete structural models. Recently, a modular structure was proposed for AGs on heterologously expressed proteins in tobacco (Nicotiana tabacum; Tan et al., 2010). Tan et al. (2010) proposed that approximately 15-residue repeating blocks of decorated β-(1→3)-trigalactosyl subunits connected by β-(1→6)-linkages were the building blocks of type II AG polysaccharides and concluded that these molecules are far less complex than commonly supposed. Most characterized β-(1→6)-galactan side chains in AGs are reported to be short, of one or two residues (Neukom and Markwalder, 1975; Gane et al., 1995; Gaspar et al., 2001). On the contrary, there are reports of long β-(1→6)-galactan side chains in radish root AGPs (Haque et al., 2005). Similarly, we recently found evidence that wheat (Triticum aestivum) flour endosperm AGP extracts contained long β-(1→6)-galactan side chains heavily substituted with l-Araf at C-3 (Tryfona et al., 2010). This partial structure of the carbohydrate component of wheat flour AGP isolated from water extracts of wheat endosperm was elucidated utilizing a combination of analytical approaches, such as the use of enzymes able to release oligosaccharides specifically from AGs, high-energy matrix-assisted laser desorption ionization (MALDI)-collision-induced dissociation (CID) mass spectrometry (MS), and polysaccharide analysis by carbohydrate gel electrophoresis (PACE; Tryfona et al., 2010). In this work, we applied these techniques to study the carbohydrate component of Arabidopsis leaf AGPs. AG-specific enzyme digestion products were analyzed by PACE and MS, allowing a partial structure to be proposed. We show that endogenous Arabidopsis leaf AG is composed of a β-(1→3)-galactan backbone with β-(1→6)-galactan side chains. These side chains are substituted with l-Araf residues via α-(1→3)-linkages and can vary in length from one up to at least 20 Galp residues. We also found that the β-(1→6)-galactan side chains are substituted mainly with 4-O-methyl-glucuronosyl (4-O-Me-GlcA) at their nonreducing termini, while occasional l-Fuc substitutions were also present via α-(1→2)-linkages on l-Araf residues. In addition, AG oligosaccharides from leaves of the mur1 mutant were identified, and their structures were compared with those isolated from wild-type plants.     

The Regulatory Properties of Rubisco Activase Differ among Species and Affect Photosynthetic Induction during Light Transitions

Rubisco’s catalytic chaperone, Rubisco activase (Rca), uses the energy from ATP hydrolysis to restore catalytic competence to Rubisco. In Arabidopsis (Arabidopsis thaliana), inhibition of Rca activity by ADP is fine tuned by redox regulation of the α-isoform. To elucidate the mechanism for Rca regulation in species containing only the redox-insensitive β-isoform, the response of activity to ADP was characterized for different Rca forms. When assayed in leaf extracts, Rubisco activation was significantly inhibited by physiological ratios of ADP to ATP in species containing both α-Rca and β-Rca (Arabidopsis and camelina [Camelina sativa]) or just the β-Rca (tobacco [Nicotiana tabacum]). However, Rca activity was insensitive to ADP inhibition in an Arabidopsis transformant, rwt43, which expresses only Arabidopsis β-Rca, although not in a transformant of Arabidopsis that expresses a tobacco-like β-Rca. ATP hydrolysis by recombinant Arabidopsis β-Rca was much less sensitive to inhibition by ADP than recombinant tobacco β-Rca. Mutation of 17 amino acids in the tobacco β-Rca to the corresponding Arabidopsis residues reduced ADP sensitivity. In planta, Rubisco deactivated at low irradiance except in the Arabidopsis rwt43 transformant containing an ADP-insensitive Rca. Induction of CO₂ assimilation after transition from low to high irradiance was much more rapid in the rwt43 transformant compared with plants containing ADP-sensitive Rca forms. The faster rate of photosynthetic induction and a greater enhancement of growth under a fluctuating light regime by the rwt43 transformant compared with wild-type Arabidopsis suggests that manipulation of Rca regulation might provide a strategy for enhancing photosynthetic performance in certain variable light environments.The activity of Rubisco, the enzyme that catalyzes CO₂ assimilation in photosynthesis, is regulated by Rubisco activase (Rca), a specific catalytic chaperone (Spreitzer and Salvucci, 2002; Portis, 2003). Like other AAA+ ATPases (Snider et al., 2008), Rca uses the energy from ATP hydrolysis to remodel the conformation of its target protein, Rubisco. The conformational changes induced by Rca restore catalytic competence to Rubisco active sites that have been inactivated by the unproductive binding of sugar phosphates, including the substrate ribulose 1,5-bisphosphate (RuBP; Wang and Portis, 1992). Because of the requirement for ATP hydrolysis and the inhibition of activity by ADP (Robinson and Portis, 1988, 1989), Rca adjusts the rate of CO₂ fixation to the rates of electron transport activity via changes in the activation state of Rubisco (Salvucci et al., 1985). As a result of this coordinate regulation, the light response of Rubisco activation closely resembles the light response of CO₂ assimilation, and the levels of RuBP under steady-state conditions are relatively constant over a wide range of irradiance levels (Perchorowicz et al., 1981; Dietz and Heber, 1984).Many plant species express two isoforms of Rca, α and β, that are both active in ATP hydrolysis and Rubisco activation (Shen et al., 1991; Salvucci et al., 2003). In some plant species, these isoforms are the products of an alternative splicing event that generates two polypeptides, which are identical except for a 20- to 30-amino acid extension at the C terminus of the longer α-isoform (Werneke et al., 1989). In cotton (Gossypium hirsutum), soybean (Glycine max), and presumably other plant species, separate genes encode the two isoforms of Rca (Salvucci et al., 2003; Yin et al., 2010). In these species, the amino acid sequences of the overlapping regions of the α- and β-polypeptides are very similar, and the C-terminal extension of the longer α-isoform is similar to the extension produced by alternative splicing (Supplemental Fig. S1).Our current understanding of the role of the two Rca isoforms is based primarily on investigations with Arabidopsis (Arabidopsis thaliana; Zhang and Portis, 1999; Zhang et al., 2001; Wang and Portis, 2006). The C-terminal extension of the α-Rca contains two redox-regulated Cys residues that are modulated by thioredoxin f (Zhang and Portis, 1999). When these residues are oxidized to a disulfide, the affinity for ATP decreases and enzyme activity is more sensitive to inhibition by ADP. Physiological ratios of ADP to ATP significantly inhibit the activity of the Arabidopsis α-Rca when in the oxidized state, but inhibition is much less when this isoform has been reduced by thioredoxin. In contrast, the shorter Arabidopsis β-Rca is not redox regulated and is less sensitive to inhibition by ADP (Zhang and Portis, 1999). Mixing experiments with recombinant Rca have shown that the properties of α-Rca are conferred to the heterooligomer, providing a mechanism for redox regulating the Rca holoenzyme (Zhang et al., 2001). In this way, Rca is similar to the chloroplastic glyceraldehyde 3-P dehydrogenase (GAPDH), which also has both redox-regulated (GAP-B) and non-redox-regulated (GAP-A) forms that differ by a C-terminal extension (Baalmann et al., 1996). Like Rca (Zhang and Portis, 1999; Zhang et al., 2001), redox regulation of two Cys residues in the extension exerts master control over the mixed GAPDH oligomer.Some plant species, including members of the Solanaceae family, as well as maize (Zea mays) and green algae, express only the shorter β-Rca (Salvucci et al., 1987). The β-Rca in these species is not responsive to redox regulation, even though the activation state of Rubisco in these plants is modulated by irradiance (Salvucci and Anderson, 1987) and seems to be associated with the redox status of the chloroplast (Ruuska et al., 2000). With GAPDH, all higher plants appear to have both chloroplastic isoforms, but the non-redox-sensitive form, Gap-A, can be regulated indirectly by thioredoxin through the binding of the small chloroplast protein CP12 (Trost et al., 2006). By analogy, a similar association with CP12 or a CP12-like protein could provide a means of conferring redox sensitivity to β-Rca in species that have only this Rca isoform. However, no association of Rca was observed when the native CP12 complex and other high-molecular-mass species were isolated from tobacco (Nicotiana tabacum) chloroplasts (Carmo-Silva et al., 2011b).In this study, the regulation of β-Rca activity was examined both in vivo and in vitro in plant species that contain only one (i.e. β-) or both (α- and β-) Rca isoforms. The response of enzyme activity to physiological ratios of ADP to ATP was measured for the native Rca in leaf extracts, as well as for recombinant Arabidopsis and tobacco enzymes, to determine the sensitivity of β-Rca to ADP in different species. In addition, experiments were conducted with transgenic Arabidopsis plants containing variants of β-Rca to determine the link between Rca regulation and photosynthetic induction. The results suggest a new strategy for enhancing photosynthetic performance under variable light environments based on altering the regulatory properties of Rca to increase the rate of photosynthetic induction.     

Glycosylphosphatidylinositol (GPI) Modification Serves as a Primary Plasmodesmal Sorting Signal

Plasmodesmata (Pd) are membranous channels that serve as a major conduit for cell-to-cell communication in plants. The Pd-associated β-1,3-glucanase (BG_pap) and CALLOSE BINDING PROTEIN1 (PDCB1) were identified as key regulators of Pd conductivity. Both are predicted glycosylphosphatidylinositol-anchored proteins (GPI-APs) carrying a conserved GPI modification signal. However, the subcellular targeting mechanism of these proteins is unknown, particularly in the context of other GPI-APs not associated with Pd. Here, we conducted a comparative analysis of the subcellular targeting of the two Pd-resident and two unrelated non-Pd GPI-APs in Arabidopsis (Arabidopsis thaliana). We show that GPI modification is necessary and sufficient for delivering both BG_pap and PDCB1 to Pd. Moreover, the GPI modification signal from both Pd- and non-Pd GPI-APs is able to target a reporter protein to Pd, likely to plasma membrane microdomains enriched at Pd. As such, the GPI modification serves as a primary Pd sorting signal in plant cells. Interestingly, the ectodomain, a region that carries the functional domain in GPI-APs, in Pd-resident proteins further enhances Pd accumulation. However, in non-Pd GPI-APs, the ectodomain overrides the Pd targeting function of the GPI signal and determines a specific GPI-dependent non-Pd localization of these proteins at the plasma membrane and cell wall. Domain-swap analysis showed that the non-Pd localization is also dominant over the Pd-enhancing function mediated by a Pd ectodomain. In conclusion, our results indicate that segregation between Pd- and non-Pd GPI-APs occurs prior to Pd targeting, providing, to our knowledge, the first evidence of the mechanism of GPI-AP sorting in plants.Plant cells are interconnected with cross-wall membranous channels called plasmodesmata (Pd). Recent studies have shown that the region of the plasma membrane (PM) lining the Pd channel is a specialized membrane microdomain whose lipid and protein composition differs from the rest of the PM (Tilsner et al., 2011, 2016; Bayer et al., 2014; González-Solís et al., 2014; Grison et al., 2015). In a similar manner, the cell wall domain surrounding the Pd channel is specialized and, unlike the rest of the cell wall, is devoid of cellulose, rich in pectin, and contains callose (an insoluble β-1,3-glucan; Zavaliev et al., 2011; Knox and Benitez-Alfonso, 2014). In response to physiological signals, callose can be transiently deposited and degraded at Pd, which provides a mechanism for controlling the Pd aperture in diverse developmental and stress-related processes (Zavaliev et al., 2011). Control of Pd functioning is mediated by proteins that are specifically targeted to Pd. Plasmodesmal proteins localized to the PM domain of Pd can be integral transmembrane proteins, such as Pd-localized proteins (Thomas et al., 2008), the receptor kinase ARABIDOPSIS CRINKLY4 (Stahl et al., 2013), and callose synthases (Vatén et al., 2011). Alternatively, Pd proteins can associate with the membrane through a lipid modification like myristoylation (e.g. remorins; Raffaele et al., 2009) or be attached by a glycosylphosphatidylinositol (GPI) anchor (e.g. Pd-associated β-1,3-glucanases [BG_pap]; Levy et al., 2007; Rinne et al., 2011; Benitez-Alfonso et al., 2013), Pd-associated callose-binding proteins (PDCBs; Simpson et al., 2009), and LYSIN MOTIF DOMAIN-CONTAINING PROTEIN2 (LYM2; Faulkner et al., 2013).Among the known Pd proteins involved in Pd-specific callose degradation is BG_pap, a cell wall enzyme carrying a glycosyl hydrolase family 17 (GH17) module as its functional domain (Levy et al., 2007). Another group of proteins controlling callose dynamics at Pd are PDCBs that harbor a callose-binding domain termed carbohydrate-binding module 43 (CBM43), implicated in stabilizing callose at Pd (Simpson et al., 2009). Some β-1,3-glucanases may combine the two callose-modifying activities by harboring both GH17 and CBM43 functional domains, and several such proteins were shown to localize to Pd (Rinne et al., 2011; Benitez-Alfonso et al., 2013; Gaudioso-Pedraza and Benitez-Alfonso, 2014). A distinct feature of BG_pap and PDCBs is that both are predicted glycosylphosphatidylinositol-anchored proteins (GPI-APs). The GPI anchor is a form of posttranslational modification common to many cell surface proteins in all eukaryotes. GPI-APs are covalently attached to the outer leaflet of the PM through the GPI anchor. The basic structure of the anchor consists of ethanolamine phosphate, followed by a glycan chain of three Man residues and glucosamine, followed by phosphatidylinositol lipid moiety (EtNP-6Manα1-2Manα1-6Manα1-4GlcNα1-6myoinositol-1-P-lipid; Ferguson et al., 2009). All predicted GPI-APs carry an N-terminal secretion signal peptide (SP) similar to other secreted proteins. Distinctly, GPI-APs also carry a structurally conserved 25- to 30-residue C-terminal GPI attachment signal, which typically begins with a small amino acid (e.g. Ala, Asn, Asp, Cys, Gly, or Ser) termed omega, followed by a spacer region of five to 10 polar residues, and ending with a transmembrane segment of 15 to 20 hydrophobic residues (Ferguson et al., 2009). The entire region between the N-terminal and the C-terminal signals of a GPI-AP is termed the ectodomain and carries the protein’s functional domain(s). The GPI modification process takes place in the lumenal face of the endoplasmic reticulum (ER) in a cotranslational manner. Upon translocation into the ER, a GPI-AP is stabilized in the ER membrane by its C-terminal signal, which is concurrently cleaved after the omega amino acid, and a preassembled GPI anchor is covalently attached to the C terminus of the omega amino acid. After attachment to a protein, the GPI anchor undergoes a series of modifications (remodeling), both at the glucan chain and at the lipid moiety. Such remodeling is crucial for the sorting of GPI-APs in the secretory pathway and the subsequent lateral heterogeneity at the PM (Kinoshita, 2015). In particular, the addition of saturated fatty acid chains to the lipid moiety of the anchor leads to the enriched accumulation of GPI-APs in the PM microdomains, also termed lipid rafts (Muñiz and Zurzolo, 2014). In Arabidopsis (Arabidopsis thaliana), GPI modification has been predicted for 210 proteins of diverse functions at the PM or the cell wall or both (Borner et al., 2002). Despite extensive research on the GPI modification pathway and the function of GPI-APs in mammalian and yeast cells, such knowledge in plant systems is scarce. In particular, despite an emerging role of GPI-APs in the regulation of the cell wall domain of Pd, their subcellular targeting and compartmentalization mechanism have not been studied. In addition, it is not known how the targeting mechanism of Pd-resident GPI-APs is different from that of other classes of GPI-APs, which are not localized to Pd.In this study, we investigated the subcellular targeting mechanism of Pd-associated callose-modifying GPI-APs, BG_pap and PDCB1, and compared it with that of two unrelated non-Pd GPI-APs, ARABINOGALACTAN PROTEIN4 (AGP4) and LIPID TRANSFER PROTEIN1 (LTPG1). Using sequential fluorescent labeling of protein domains, we found that the C-terminal GPI modification signal present in both Pd- and non-Pd GPI-APs can function as a primary signal in targeting proteins to the Pd-enriched PM domain. Moreover, we show that while the GPI signal is sufficient for Pd targeting, the ectodomains in BG_pap and PDCB1 further enhance their accumulation at Pd. In contrast, the ectodomains in non-Pd GPI-APs mediate exclusion of the proteins from the Pd-enriched targeting pathway. The Pd exclusion effect was found to be dominant over the Pd-targeting function of the GPI signal and the Pd-enhancing function of the Pd ectodomain, and it possibly occurs prior to PM localization. Our findings thus uncover a novel Pd-targeting signal and provide, to our knowledge, the first evidence of the cellular mechanism that regulates the sorting of GPI-APs in plants.