Composition,Assembly, and Trafficking of a Wheat Xylan Synthase Complex

Xylans play an important role in plant cell wall integrity and have many industrial applications. Characterization of xylan synthase (XS) complexes responsible for the synthesis of these polymers is currently lacking. We recently purified XS activity from etiolated wheat (Triticum aestivum) seedlings. To further characterize this purified activity, we analyzed its protein composition and assembly. Proteomic analysis identified six main proteins: two glycosyltransferases (GTs) TaGT43-4 and TaGT47-13; two putative mutases (TaGT75-3 and TaGT75-4) and two non-GTs; a germin-like protein (TaGLP); and a vernalization related protein (TaVER2). Coexpression of TaGT43-4, TaGT47-13, TaGT75-3, and TaGT75-4 in Pichia pastoris confirmed that these proteins form a complex. Confocal microscopy showed that all these proteins interact in the endoplasmic reticulum (ER) but the complexes accumulate in Golgi, and TaGT43-4 acts as a scaffold protein that holds the other proteins. Furthermore, ER export of the complexes is dependent of the interaction between TaGT43-4 and TaGT47-13. Immunogold electron microscopy data support the conclusion that complex assembly occurs at specific areas of the ER before export to the Golgi. A di-Arg motif and a long sequence motif within the transmembrane domains were found conserved at the NH₂-terminal ends of TaGT43-4 and homologous proteins from diverse taxa. These conserved motifs may control the forward trafficking of the complexes and their accumulation in the Golgi. Our findings indicate that xylan synthesis in grasses may involve a new regulatory mechanism linking complex assembly with forward trafficking and provide new insights that advance our understanding of xylan biosynthesis and regulation in plants.It is believed that Golgi-localized, multiprotein complexes synthesize plant hemicellulosic polysaccharides, including xylans. Such complexes are not well characterized in plants (Zeng et al., 2010; Atmodjo et al., 2011; Chou et al., 2012), which is in sharp contrast with mammalian and yeast cells (Jungmann and Munro, 1998; McCormick et al., 2000; Giraudo et al., 2001). Xylans are the most abundant plant hemicellulosic polysaccharides on Earth and play an important role in the integrity of cell walls, which is a key factor in plant growth. Any mutations affecting xylan backbone biosynthesis seem to result in abnormal growth of plants due mostly to thinning and weakening of secondary xylem walls, described as the irregular xylem (irx) phenotype. Thus, characterizing the xylan synthase complex (XSC) would have an impact on plant improvement, as well as many industrial applications related to food, feed, and biofuel production (Yang and Wyman, 2004; Faik, 2010). Although the Arabidopsis (Arabidopsis thaliana) irx mutants have revealed the involvement of several glycosyltransferase (GT) gene families in xylan biosynthesis (Brown et al., 2007, 2009; Lee et al., 2007, 2010; Wu et al., 2009, 2010), no XSCs have been purified/isolated from Arabidopsis tissues, and we still do not know whether some of the identified Arabidopsis GTs can assemble into functional XSCs. Furthermore, if GTs do assemble into XSCs, we don’t know the mechanisms by which plant cells control their assembly and cellular trafficking. In contrast to dicots, xylan synthase activity was recently immunopurified from etiolated wheat (Triticum aestivum) microsomes (Zeng et al., 2010). This purified wheat XS activity was shown to catalyze three activities, xylan-glucuronosyltransferase (XGlcAT), xylan-xylosyltransferase (XXylT), and xylan-arabinofuranosyltranferase (XAT), which work synergistically to synthesize xylan-type polymers in vitro (Zeng et al., 2008, 2010). This work focuses on describing protein composition, assembly, and trafficking of this purified wheat XS activity.In all eukaryotes, proteins of the secretory pathway (including GTs) are synthesized in the endoplasmic reticulum (ER) and modified as they go through the Golgi cisternae. Most proteins exit the ER from ER export sites (ERESs; Hanton et al., 2009) and use a signal-based sorting mechanism that allows them to be selectively recruited into vesicles coated by coat protein II complexes (Barlowe, 2003; Beck et al., 2008). For many Golgi-resident type II membrane proteins, di-Arg motifs, such as RR, RXR, and RRR located in their cytosolic NH₂-terminal ends, have been shown to be required for their ER export (Giraudo et al., 2003; Czlapinski and Bertozzi, 2006; Schoberer et al., 2009; Tu and Banfield, 2010). Interestingly, di-Arg motifs located ∼40 amino acids from the membrane on the cytosolic side can also be used to retrieve some type II ER-resident proteins from cis-Golgi (Schutze et al., 1994; Hardt et al., 2003; Boulaflous et al., 2009). In contrast to the signal-based sorting mechanism involved in trafficking between the ER and Golgi, the steady-state localization/retention of proteins (including GTs) in the Golgi is thought to occur through vesicular cycling. Cycling is influenced by various mechanisms, including the length and composition of the transmembrane domain (TMD) of type II GTs (Bretscher and Munro, 1993; Colley, 1997; van Vliet et al., 2003; Sousa et al., 2003; Sharpe et al., 2010), and the oligomerization/aggregation of GTs (kin hypothesis), which suggests that formation of homo- or heterooligomers of GTs in the Golgi may prevent their recruitment into clathrin-coated vesicles (Machamer, 1991; Nilsson et al., 1993; Weisz et al., 1993; Cole et al., 1996). Some Golgi-resident GTs are predicted to have a cleavable NH₂-terminal secretion signal peptide (SP) and would therefore exist as soluble proteins in the Golgi lumen. To maintain their proper Golgi localization, these processed GTs are likely part of multiprotein complexes anchored to integral membrane proteins. The fact that homologs of many of the trafficking proteins from mammalian and yeast cells are found in plants indicates that trafficking machineries of the plant secretory pathway are likely conserved (d’Enfert et al., 1992; Bar-Peled and Raikhel, 1997; Batoko et al., 2000; Pimpl et al., 2000; Phillipson et al., 2001; Hawes et al., 2008).It is becoming increasingly evident that understanding the mechanisms controlling protein-protein interaction, sorting, and trafficking of polysaccharide synthases (including XSCs) will help elucidate how plants regulate cell wall synthesis and deposition during their development. To this end, we believe that the purified wheat XS activity (Zeng et al., 2010) is an excellent model for this type of study. In this work, proteomics was used to determine the protein composition of the purified XS activity. Confocal microscopy and immunogold transmission electron microscopy (TEM) were used to investigate the assembly and trafficking of the complex. Our proteomics data showed that the purified activity contains two GTs, TaGT43-4 and TaGT47-13, two putative mutases, TaGT75-3 and TaGT75-4, and two non-GT proteins: a germin-like protein (TaGLP) belonging to cupin superfamily and a protein specific to monocots annotated as wheat vernalization-related protein 2 (TaVER2). Microscopy analyses revealed that all these proteins interact in the ER, but the assembled complexes accumulate in the Golgi. Export of these complexes from the ER is controlled by the interaction between TaGT43-4 and TaGT47-13. Characterization of the wheat XSC and its trafficking furthers our understanding of xylan biosynthesis in grasses and helps elucidate how polysaccharide synthase complexes are assembled, sorted, and maintained in different compartments of the secretory pathway.     

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

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

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

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

Surveying Rubisco Diversity and Temperature Response to Improve Crop Photosynthetic Efficiency

The threat to global food security of stagnating yields and population growth makes increasing crop productivity a critical goal over the coming decades. One key target for improving crop productivity and yields is increasing the efficiency of photosynthesis. Central to photosynthesis is Rubisco, which is a critical but often rate-limiting component. Here, we present full Rubisco catalytic properties measured at three temperatures for 75 plants species representing both crops and undomesticated plants from diverse climates. Some newly characterized Rubiscos were naturally “better” compared to crop enzymes and have the potential to improve crop photosynthetic efficiency. The temperature response of the various catalytic parameters was largely consistent across the diverse range of species, though absolute values showed significant variation in Rubisco catalysis, even between closely related species. An analysis of residue differences among the species characterized identified a number of candidate amino acid substitutions that will aid in advancing engineering of improved Rubisco in crop systems. This study provides new insights on the range of Rubisco catalysis and temperature response present in nature, and provides new information to include in models from leaf to canopy and ecosystem scale.In a changing climate and under pressure from a population set to hit nine billion by 2050, global food security will require massive changes to the way food is produced, distributed, and consumed (Ort et al., 2015). To match rising demand, agricultural production must increase by 50 to 70% in the next 35 years, and yet the gains in crop yields initiated by the green revolution are slowing, and in some cases, stagnating (Long and Ort, 2010; Ray et al., 2012). Among a number of areas being pursued to increase crop productivity and food production, improving photosynthetic efficiency is a clear target, offering great promise (Parry et al., 2007; von Caemmerer et al., 2012; Price et al., 2013; Ort et al., 2015). As the gatekeeper of carbon entry into the biosphere and often acting as the rate-limiting step of photosynthesis, Rubisco, the most abundant enzyme on the planet (Ellis, 1979), is an obvious and important target for improving crop photosynthetic efficiency.Rubisco is considered to exhibit comparatively poor catalysis, in terms of catalytic rate, specificity, and CO₂ affinity (Tcherkez et al., 2006; Andersson, 2008), leading to the suggestion that even small increases in catalytic efficiency may result in substantial improvements to carbon assimilation across a growing season (Zhu et al., 2004; Parry et al., 2013; Galmés et al., 2014a; Carmo-Silva et al., 2015). If combined with complimentary changes such as optimizing other components of the Calvin Benson or photorespiratory cycles (Raines, 2011; Peterhansel et al., 2013; Simkin et al., 2015), optimized canopy architecture (Drewry et al., 2014), or introducing elements of a carbon concentrating mechanism (Furbank et al., 2009; Lin et al., 2014a; Hanson et al., 2016; Long et al., 2016), Rubisco improvement presents an opportunity to dramatically increase the photosynthetic efficiency of crop plants (McGrath and Long, 2014; Long et al., 2015; Betti et al., 2016). A combination of the available strategies is essential for devising tailored solutions to meet the varied requirements of different crops and the diverse conditions under which they are typically grown around the world.Efforts to engineer an improved Rubisco have not yet produced a “super Rubisco” (Parry et al., 2007; Ort et al., 2015). However, advances in engineering precise changes in model systems continue to provide important developments that are increasing our understanding of Rubisco catalysis (Spreitzer et al., 2005; Whitney et al., 2011a, 2011b; Morita et al., 2014; Wilson et al., 2016), regulation (Andralojc et al., 2012; Carmo-Silva and Salvucci, 2013; Bracher et al., 2015), and biogenesis (Saschenbrecker et al., 2007; Whitney and Sharwood, 2008; Lin et al., 2014b; Hauser et al., 2015; Whitney et al., 2015).A complementary approach is to understand and exploit Rubisco natural diversity. Previous characterization of Rubisco from a limited number of species has not only demonstrated significant differences in the underlying catalytic parameters, but also suggests that further undiscovered diversity exists in nature and that the properties of some of these enzymes could be beneficial if present in crop plants (Carmo-Silva et al., 2015). Recent studies clearly illustrate the variation possible among even closely related species (Galmés et al., 2005, 2014b, 2014c; Kubien et al., 2008; Andralojc et al., 2014; Prins et al., 2016).Until recently, there have been relatively few attempts to characterize the consistency, or lack thereof, of temperature effects on in vitro Rubisco catalysis (Sharwood and Whitney, 2014), and often studies only consider a subset of Rubisco catalytic properties. This type of characterization is particularly important for future engineering efforts, enabling specific temperature effects to be factored into any attempts to modify crops for a future climate. In addition, the ability to coanalyze catalytic properties and DNA or amino acid sequence provides the opportunity to correlate sequence and biochemistry to inform engineering studies (Christin et al., 2008; Kapralov et al., 2011; Rosnow et al., 2015). While the amount of gene sequence information available grows rapidly with improving technology, knowledge of the corresponding biochemical variation resulting has yet to be determined (Cousins et al., 2010; Carmo-Silva et al., 2015; Sharwood and Whitney, 2014; Nunes-Nesi et al., 2016).This study aimed to characterize the catalytic properties of Rubisco from diverse species, comprising a broad range of monocots and dicots from diverse environments. The temperature dependence of Rubisco catalysis was evaluated to tailor Rubisco engineering for crop improvement in specific environments. Catalytic diversity was analyzed alongside the sequence of the Rubisco large subunit gene, rbcL, to identify potential catalytic switches for improving photosynthesis and productivity. In vitro results were compared to the average temperature of the warmest quarter in the regions where each species grows to investigate the role of temperature in modulating Rubisco catalysis.     

Unidirectional Movement of Cellulose Synthase Complexes in Arabidopsis Seed Coat Epidermal Cells Deposit Cellulose Involved in Mucilage Extrusion,Adherence, and Ray Formation

CELLULOSE SYNTHASE5 (CESA5) synthesizes cellulose necessary for seed mucilage adherence to seed coat epidermal cells of Arabidopsis (Arabidopsis thaliana). The involvement of additional CESA proteins in this process and details concerning the manner in which cellulose is deposited in the mucilage pocket are unknown. Here, we show that both CESA3 and CESA10 are highly expressed in this cell type at the time of mucilage synthesis and localize to the plasma membrane adjacent to the mucilage pocket. The isoxaben resistant1-1 and isoxaben resistant1-2 mutants affecting CESA3 show defects consistent with altered mucilage cellulose biosynthesis. CESA3 can interact with CESA5 in vitro, and green fluorescent protein-tagged CESA5, CESA3, and CESA10 proteins move in a linear, unidirectional fashion around the cytoplasmic column of the cell, parallel with the surface of the seed, in a pattern similar to that of cortical microtubules. Consistent with this movement, cytological evidence suggests that the mucilage is coiled around the columella and unwinds during mucilage extrusion to form a linear ray. Mutations in CESA5 and CESA3 affect the speed of mucilage extrusion and mucilage adherence. These findings imply that cellulose fibrils are synthesized in an ordered helical array around the columella, providing a distinct structure to the mucilage that is important for both mucilage extrusion and adherence.The epidermal cells of Arabidopsis (Arabidopsis thaliana) seed coats produce two distinct secondary cell walls: pectin-rich mucilage and cellulose-rich columellae (Western et al., 2000). When seeds are hydrated, mucilage expands rapidly, rupturing the outer tangential cell wall and forming a mucilage capsule that surrounds the seed. Seed coat mucilage is composed primarily of rhamnogalacturonan I (RG I) and also contains homogalacturonan (HG), hemicelluloses (such as xylans and glucomannans), and cellulose (for review, see Haughn and Western, 2012). Extruded mucilage consists of an outer, nonadherent fraction and an inner, adherent fraction (Western et al., 2000, 2001; Macquet et al., 2007a). The adherent and nonadherent mucilage layers differ in the amount of methylesterified HG (Rautengarten et al., 2008; Saez-Aguayo et al., 2013; Voiniciuc et al., 2013), galactans (Dean et al., 2007; Macquet et al., 2007b), arabinans (Arsovski et al., 2009), mannans (Yu et al., 2014), and cellulose (Harpaz-Saad et al., 2011; Mendu et al., 2011; Sullivan et al., 2011), all of which influence the physical properties of the layers.Adherent mucilage has a distinct structure, which can be examined using cell wall dyes and antibodies. When treated with cellulose-specific dyes, densely stained rays extend from the top of each columella to the outer edge of the adherent layer, many cell lengths above the seed surface (Mendu et al., 2011; Sullivan et al., 2011). Cytological evidence indicates that cellulose, pectins, and mannans are components of the ray (Haughn and Western, 2012; Griffiths et al., 2014; North et al., 2014; Yu et al., 2014), although the exact manner in which they are assembled is unknown.Cellulose is abundant in mucilage rays and mediates adherence. Loss-of-function mutations in CELLULOSE SYNTHASE5 (CESA5) result in reduced cellulose levels and increased detachment of mucilage from the seed (Harpaz-Saad et al., 2011; Mendu et al., 2011; Sullivan et al., 2011; Griffiths et al., 2014). How a reduction in cellulose results in a loss of adherence is still unknown, but it likely involves interaction with other mucilage components such as pectin and arabinogalactan proteins (Griffiths et al., 2014). Since cesa5 mutants still have some cellulose in the rays of the adherent mucilage halo (Mendu et al., 2011; Sullivan et al., 2011), additional cellulose synthases must be involved in mucilage cellulose biosynthesis.The Arabidopsis genome encodes 10 different CESAs (Delmer, 1999; Richmond and Somerville, 2000). Multiple lines of evidence suggest that three different CESAs are required to form one active cellulose synthase complex (CSC; for review, see Somerville, 2006). CSCs are membrane-bound protein complexes that synthesize cellulose microfibrils in the apoplast (for review, see Somerville, 2006; Endler and Persson, 2011; Lei et al., 2012). CESA1, CESA3, and CESA6 are considered the core components of the primary wall CSC (Desprez et al., 2007; Persson et al., 2007). CESA2, CESA5, and CESA9 are partially redundant to CESA6 in primary wall biosynthesis, and genetic evidence suggests that each of these CESA polypeptides can form a functional CSC with CESA3 and CESA1 (Desprez et al., 2007; Persson et al., 2007). CESA10 is expressed in young plants, stems, floral tissue, and the base of rosette leaves (Beeckman et al., 2002; Doblin et al., 2002), but its function in cellulose biosynthesis is unclear. Other cesa mutant lines have been examined for altered mucilage phenotypes (cesa1, radially swollen1 [Burn et al., 2002; Sullivan et al., 2011], cesa2, cesa6, and cesa9 [Mendu et al., 2011]; CESA3, je5 [Sullivan et al., 2011] and cesa10-1 [Sullivan et al., 2011]); to date, only CESA5 has been shown to be required for cellulose biosynthesis during mucilage deposition.Two mutant alleles of CESA3, isoxaben resistant1-1 (ixr1-1) and ixr1-2, were isolated in a screen for resistance to the herbicide isoxaben (Scheible et al., 2001). Isoxaben inhibits the incorporation of Glc into the emerging cellulose polymer and is considered a potent and specific inhibitor of cellulose biosynthesis (Heim et al., 1990). Homozygous ixr1-1 and ixr1-2 lines show increased resistance to the herbicide, and the mutations causing this resistance were mapped to the genomic locus of CESA3 (Heim et al., 1990; Scheible et al., 2001). The ixr1-1 and ixr1-2 mutations cause amino acid substitutions near the C terminus of the CESA3 protein. ixr1-1 causes a Gly-to-Asn substitution (G998A) located in a transmembrane domain, while ixr1-2 contains a Thr-to-Ile substitution (T942I) in an apoplastic region of the protein between two transmembrane domains (Scheible et al., 2001). Recently, the ixr1-2 allele was shown to affect the velocity of CSCs in the plasma membrane, which consequently modifies cellulose crystallinity in the cell wall (Harris et al., 2012). It is not exactly clear how the ixr1-1 mutation affects cellulose biosynthesis. The effects of either of these mutations on seed coat mucilage have not been investigated.Since mucilage is composed primarily of pectins with smaller amounts of cellulose, seed coat epidermal cells represent an excellent system to study cellulose biosynthesis and interactions between cellulose and other wall components in muro. In this study, we investigated how cellulose is synthesized and deposited in seed coat epidermal cells. We show that at least three different CESA proteins are highly expressed in the seed coat epidermis during mucilage biosynthesis. These CESAs are oriented and move in a linear fashion around the cytoplasmic column of each cell in an identical pattern to cortical microtubules. In addition, we provide evidence that the adherent mucilage has a helical structure that expands and unwinds during extrusion to form the mucilage ray. We propose that during seed coat epidermal cell development, the biosynthesis of cellulose predetermines the structure of rays in the adherent mucilage layer.     

Patterns of Metabolite Changes Identified from Large-Scale Gene Perturbations in Arabidopsis Using a Genome-Scale Metabolic Network

Metabolomics enables quantitative evaluation of metabolic changes caused by genetic or environmental perturbations. However, little is known about how perturbing a single gene changes the metabolic system as a whole and which network and functional properties are involved in this response. To answer this question, we investigated the metabolite profiles from 136 mutants with single gene perturbations of functionally diverse Arabidopsis (Arabidopsis thaliana) genes. Fewer than 10 metabolites were changed significantly relative to the wild type in most of the mutants, indicating that the metabolic network was robust to perturbations of single metabolic genes. These changed metabolites were closer to each other in a genome-scale metabolic network than expected by chance, supporting the notion that the genetic perturbations changed the network more locally than globally. Surprisingly, the changed metabolites were close to the perturbed reactions in only 30% of the mutants of the well-characterized genes. To determine the factors that contributed to the distance between the observed metabolic changes and the perturbation site in the network, we examined nine network and functional properties of the perturbed genes. Only the isozyme number affected the distance between the perturbed reactions and changed metabolites. This study revealed patterns of metabolic changes from large-scale gene perturbations and relationships between characteristics of the perturbed genes and metabolic changes.Rational and quantitative assessment of metabolic changes in response to genetic modification (GM) is an open question and in need of innovative solutions. Nontargeted metabolite profiling can detect thousands of compounds, but it is not easy to understand the significance of the changed metabolites in the biochemical and biological context of the organism. To better assess the changes in metabolites from nontargeted metabolomics studies, it is important to examine the changed metabolites in the context of the genome-scale metabolic network of the organism.Metabolomics is a technique that aims to quantify all the metabolites in a biological system (Nikolau and Wurtele, 2007; Nicholson and Lindon, 2008; Roessner and Bowne, 2009). It has been used widely in studies ranging from disease diagnosis (Holmes et al., 2008; DeBerardinis and Thompson, 2012) and drug discovery (Cascante et al., 2002; Kell, 2006) to metabolic reconstruction (Feist et al., 2009; Kim et al., 2012) and metabolic engineering (Keasling, 2010; Lee et al., 2011). Metabolomic studies have demonstrated the possibility of identifying gene functions from changes in the relative concentrations of metabolites (metabotypes or metabolic signatures; Ebbels et al., 2004) in various species including yeast (Saccharomyces cerevisiae; Raamsdonk et al., 2001; Allen et al., 2003), Arabidopsis (Arabidopsis thaliana; Brotman et al., 2011), tomato (Solanum lycopersicum; Schauer et al., 2006), and maize (Zea mays; Riedelsheimer et al., 2012). Metabolomics has also been used to better understand how plants interact with their environments (Field and Lake, 2011), including their responses to biotic and abiotic stresses (Dixon et al., 2006; Arbona et al., 2013), and to predict important agronomic traits (Riedelsheimer et al., 2012). Metabolite profiling has been performed on many plant species, including angiosperms such as Arabidopsis, poplar (Populus trichocarpa), and Catharanthus roseus (Sumner et al., 2003; Rischer et al., 2006), basal land plants such as Selaginella moellendorffii and Physcomitrella patens (Erxleben et al., 2012; Yobi et al., 2012), and Chlamydomonas reinhardtii (Fernie et al., 2012; Davis et al., 2013). With the availability of whole genome sequences of various species, metabolomics has the potential to become a useful tool for elucidating the functions of genes using large-scale systematic analyses (Fiehn et al., 2000; Saito and Matsuda, 2010; Hur et al., 2013).Although metabolomics data have the potential for identifying the roles of genes that are associated with metabolic phenotypes, the biochemical mechanisms that link functions of genes with metabolic phenotypes are still poorly characterized. For example, we do not yet know the principles behind how perturbing the expression of a single gene changes the metabolic system as a whole. Large-scale metabolomics data have provided useful resources for linking phenotypes to genotypes (Fiehn et al., 2000; Roessner et al., 2001; Tikunov et al., 2005; Schauer et al., 2006; Lu et al., 2011; Fukushima et al., 2014). For example, Lu et al. (2011) compared morphological and metabolic phenotypes from more than 5,000 Arabidopsis chloroplast mutants using gas chromatography (GC)- and liquid chromatography (LC)-mass spectrometry (MS). Fukushima et al. (2014) generated metabolite profiles from various characterized and uncharacterized mutant plants and clustered the mutants with similar metabolic phenotypes by conducting multidimensional scaling with quantified metabolic phenotypes. Nonetheless, representation and analysis of such a large amount of data remains a challenge for scientific discovery (Lu et al., 2011). In addition, these studies do not examine the topological and functional characteristics of metabolic changes in the context of a genome-scale metabolic network. To understand the relationship between genotype and metabolic phenotype, we need to investigate the metabolic changes caused by perturbing the expression of a gene in a genome-scale metabolic network perspective, because metabolic pathways are not independent biochemical factories but are components of a complex network (Berg et al., 2002; Merico et al., 2009).Much progress has been made in the last 2 decades to represent metabolism at a genome scale (Terzer et al., 2009). The advances in genome sequencing and emerging fields such as biocuration and bioinformatics enabled the representation of genome-scale metabolic network reconstructions for model organisms (Bassel et al., 2012). Genome-scale metabolic models have been built and applied broadly from microbes to plants. The first step toward modeling a genome-scale metabolism in a plant species started with developing a genome-scale metabolic pathway database for Arabidopsis (AraCyc; Mueller et al., 2003) from reference pathway databases (Kanehisa and Goto, 2000; Karp et al., 2002; Zhang et al., 2010). Genome-scale metabolic pathway databases have been built for several plant species (Mueller et al., 2005; Zhang et al., 2005, 2010; Urbanczyk-Wochniak and Sumner, 2007; May et al., 2009; Dharmawardhana et al., 2013; Monaco et al., 2013, 2014; Van Moerkercke et al., 2013; Chae et al., 2014; Jung et al., 2014). Efforts have been made to develop predictive genome-scale metabolic models using enzyme kinetics and stoichiometric flux-balance approaches (Sweetlove et al., 2008). de Oliveira Dal’Molin et al. (2010) developed a genome-scale metabolic model for Arabidopsis and successfully validated the model by predicting the classical photorespiratory cycle as well as known key differences between redox metabolism in photosynthetic and nonphotosynthetic plant cells. Other genome-scale models have been developed for Arabidopsis (Poolman et al., 2009; Radrich et al., 2010; Mintz-Oron et al., 2012), C. reinhardtii (Chang et al., 2011; Dal’Molin et al., 2011), maize (Dal’Molin et al., 2010; Saha et al., 2011), sorghum (Sorghum bicolor; Dal’Molin et al., 2010), and sugarcane (Saccharum officinarum; Dal’Molin et al., 2010). These predictive models have the potential to be applied broadly in fields such as metabolic engineering, drug target discovery, identification of gene function, study of evolutionary processes, risk assessment of genetically modified crops, and interpretations of mutant phenotypes (Feist and Palsson, 2008; Ricroch et al., 2011).Here, we interrogate the metabotypes caused by 136 single gene perturbations of Arabidopsis by analyzing the relative concentration changes of 1,348 chemically identified metabolites using a reconstructed genome-scale metabolic network. We examine the characteristics of the changed metabolites (the metabolites whose relative concentrations were significantly different in mutants relative to the wild type) in the metabolic network to uncover biological and topological consequences of the perturbed genes.     

Focus on Ethylene: Ethylene and the Regulation of Physiological and Morphological Responses to Nutrient Deficiencies

To cope with nutrient deficiencies, plants develop both morphological and physiological responses. The regulation of these responses is not totally understood, but some hormones and signaling substances have been implicated. It was suggested several years ago that ethylene participates in the regulation of responses to iron and phosphorous deficiency. More recently, its role has been extended to other deficiencies, such as potassium, sulfur, and others. The role of ethylene in so many deficiencies suggests that, to confer specificity to the different responses, it should act through different transduction pathways and/or in conjunction with other signals. In this update, the data supporting a role for ethylene in the regulation of responses to different nutrient deficiencies will be reviewed. In addition, the results suggesting the action of ethylene through different transduction pathways and its interaction with other hormones and signaling substances will be discussed.When plants suffer from a mineral nutrient deficiency, they develop morphological and physiological responses (mainly in their roots) aimed to facilitate the uptake and mobilization of the limiting nutrient. After the nutrient has been acquired in enough quantity, these responses need to be switched off to avoid toxicity and conserve energy. In recent years, different plant hormones (e.g. ethylene, auxin, cytokinins, jasmonic acid, abscisic acid, brassinosteroids, GAs, and strigolactones) have been implicated in the regulation of these responses (Romera et al., 2007, 2011, 2015; Liu et al., 2009; Rubio et al., 2009; Kapulnik et al., 2011; Kiba et al., 2011; Iqbal et al., 2013; Zhang et al., 2014).Before the 1990s, there were several publications relating ethylene and nutrient deficiencies (cited in Lynch and Brown [1997] and Romera et al. [1999]) without establishing a direct implication of ethylene in the regulation of nutrient deficiency responses. In 1994, Romera and Alcántara (1994) published an article in Plant Physiology suggesting a role for ethylene in the regulation of Fe deficiency responses. In 1999, Borch et al. (1999) showed the participation of ethylene in the regulation of P deficiency responses. Since then, evidence has been accumulating in support of a role for ethylene in the regulation of both Fe (Romera et al., 1999, 2015; Waters and Blevins, 2000; Lucena et al., 2006; Waters et al., 2007; García et al., 2010, 2011, 2013, 2014; Yang et al., 2014) and P deficiency responses (Kim et al., 2008; Lei et al., 2011; Li et al., 2011; Nagarajan and Smith, 2012; Wang et al., 2012, 2014c). Both Fe and P may be poorly available in most soils, and plants develop similar responses under their deficiencies (Romera and Alcántara, 2004; Zhang et al., 2014). More recently, a role for ethylene has been extended to other deficiencies, such as K (Shin and Schachtman, 2004; Jung et al., 2009; Kim et al., 2012), S (Maruyama-Nakashita et al., 2006; Wawrzyńska et al., 2010; Moniuszko et al., 2013), and B (Martín-Rejano et al., 2011). Ethylene has also been implicated in both N deficiency and excess (Tian et al., 2009; Mohd-Radzman et al., 2013; Zheng et al., 2013), and its participation in Mg deficiency has been suggested (Hermans et al., 2010).In this update, we will review the information supporting a role for ethylene in the regulation of different nutrient deficiency responses. For information relating ethylene to other aspects of plant mineral nutrition, such as N₂ fixation and responses to excess of nitrate or essential heavy metals, the reader is referred to other reviews (for review, see Maksymiec, 2007; Mohd-Radzman et al., 2013; Steffens, 2014).     

Structure of Cellulose Microfibrils in Primary Cell Walls from Collenchyma

In the primary walls of growing plant cells, the glucose polymer cellulose is assembled into long microfibrils a few nanometers in diameter. The rigidity and orientation of these microfibrils control cell expansion; therefore, cellulose synthesis is a key factor in the growth and morphogenesis of plants. Celery (Apium graveolens) collenchyma is a useful model system for the study of primary wall microfibril structure because its microfibrils are oriented with unusual uniformity, facilitating spectroscopic and diffraction experiments. Using a combination of x-ray and neutron scattering methods with vibrational and nuclear magnetic resonance spectroscopy, we show that celery collenchyma microfibrils were 2.9 to 3.0 nm in mean diameter, with a most probable structure containing 24 chains in cross section, arranged in eight hydrogen-bonded sheets of three chains, with extensive disorder in lateral packing, conformation, and hydrogen bonding. A similar 18-chain structure, and 24-chain structures of different shape, fitted the data less well. Conformational disorder was largely restricted to the surface chains, but disorder in chain packing was not. That is, in position and orientation, the surface chains conformed to the disordered lattice constituting the core of each microfibril. There was evidence that adjacent microfibrils were noncovalently aggregated together over part of their length, suggesting that the need to disrupt these aggregates might be a constraining factor in growth and in the hydrolysis of cellulose for biofuel production.Growth and form in plants are controlled by the precisely oriented expansion of the walls of individual cells. The driving force for cell expansion is osmotic, but the rate and direction of expansion are controlled by the mechanical properties of the cell wall (Szymanski and Cosgrove, 2009). Expanding, primary cell walls are nanocomposite materials in which long microfibrils of cellulose, a few nanometers in diameter, run through a hydrated matrix of xyloglucans, pectins, and other polymers (Knox, 2008; Mohnen, 2008; Szymanski and Cosgrove, 2009; Scheller and Ulvskov, 2010). Native cellulose microfibrils are partially crystalline (Nishiyama, 2009; Fernandes et al., 2011). Formerly, primary wall cellulose was thought to have a unique crystal structure called cellulose IV₁ (Dinand et al., 1996), but NMR evidence suggests the presence of forms similar to the better characterized cellulose Iα and Iβ crystalline forms together with large quantities of less ordered cellulose (Wickholm et al., 1998; Sturcová et al., 2004; Wada et al., 2004). Nevertheless, cellulose is much more ordered than any other component of the primary cell wall (Bootten et al., 2004), in keeping with its key role of providing strength and controlling growth.The stiffness of the cell wall is greatest in the direction of the cellulose microfibrils, where growth is directional and the predominant microfibril orientation is usually transverse to the growth direction (Green, 1999; MacKinnon et al., 2006; Szymanski and Cosgrove, 2009). Expansion of the cell wall then requires either widening of the spacing between microfibrils (Marga et al., 2005) or slippage between them (Cosgrove, 2005), or both, and the microfibrils reorient toward the direction of growth (Anderson et al., 2010). Polymer cross bridges between microfibrils (McCann et al., 1990) are thought to resist these deformations of the cell wall nanostructure and, thus, to control the rate of growth. Until recently, most attention was focused on bridging xyloglucans, hydrogen bonded to microfibril surfaces (Scheller and Ulvskov, 2010). However, there is evidence that not all xyloglucans are appropriately positioned (Fujino et al., 2000; Park and Cosgrove, 2012a) and that other bridging polymers may be involved (Zykwinska et al., 2007). It has also been suggested that bundles of aggregated microfibrils, not single microfibrils, might be the key structural units in primary cell walls (Anderson et al., 2010), as in wood (Fahlén and Salmén, 2005; Fernandes et al., 2011). If so, single microfibrils could bridge between microfibril bundles. In summary, the growth of plant cells is not well understood, and we need more information on how cellulose orientation is controlled and on the nature of the bridging polymers, the cellulose surfaces to which these polymers bind, and the cohesion between microfibril surfaces that might mediate aggregation.Cellulose microfibrils are synthesized at the cell surface by large enzyme complexes having hexagonal symmetry, sometimes called “rosettes” (Somerville, 2006). Each complex contains multiple cellulose synthases that differ between primary cell walls and wood, although the appearance of the complexes is similar (Somerville, 2006; Atanassov et al., 2009). The simultaneous synthesis, from the same end, of all the chains in a native cellulose microfibril is why they are parallel (Nishiyama et al., 2002, 2003), in contrast to the entropically favored antiparallel structure found in man-made celluloses like rayon (Langan et al., 2001). The number of chains in a microfibril and the number of cellulose synthases in the synthetic complex are evidently related. It is commonly assumed that the number of chains is divisible by six, matching the hexagonal rosette symmetry, and 36-chain models (Himmel et al., 2007) bounded by the hydrophilic [110] and [1-10] crystal faces, as in algal celluloses (Bergenstråhle et al., 2008), have been widely adopted. The assembly and orientation of cellulose are connected, as several cellulose synthase mutants have phenotypes defective in cellulose orientation and plant form as well as depleted in cellulose content (Paredez et al., 2008). In certain other mutant lines, the crystallinity of the microfibrils appears to be affected (Fujita et al., 2011; Harris et al., 2012; Sánchez-Rodríguez et al., 2012).Therefore, a detailed understanding of the structure of primary wall cellulose microfibrils would help us to understand cellulose synthesis as well as the growth and structural mechanics of living plants (Burgert and Fratzl, 2009). Primary cell walls and their cellulose skeletons also affect food quality characteristics like the crispness of salad vegetables and apples (Malus domestica; Jarvis, 2011). When biofuels are produced from lignocellulosic biomass, lignification leads to recalcitrance (Himmel et al., 2007), but some of the cell types in Miscanthus spp., switchgrass (Panicum virgatum), and arable crop residues have only primary walls with no lignin, and recalcitrance then depends on the nature of the cellulose microfibrils (Beckham et al., 2011).A relatively detailed structure has recently been proposed for the microfibrils of spruce (Picea spp.) wood (Fernandes et al., 2011), which are 3.0 nm in diameter, allowing space for only about 24 cellulose chains. Evidence from x-ray diffraction supported a “rectangular” shape (Matthews et al., 2006) bounded by the [010] and [200] faces. There was considerable disorder increasing toward the surface, and the microfibrils were aggregated into bundles about 15 to 20 nm across, with some, but not all, of the lateral interfaces being resistant to water (Fernandes et al., 2011). Disordered domains are a feature of other strong biological materials such as spider silk (van Beek et al., 2002).Therefore, it is of interest whether any of these features of wood cellulose might also be found in the cellulose microfibrils of primary (growing) cell walls. It would be particularly useful to characterize the disorder known to be present in primary wall microfibrils, that is, to define how cellulose that is not measured as “crystalline” differs from crystalline cellulose. Many of the experiments leading toward a structure for wood cellulose were dependent on exceptionally uniform orientation of the cellulose microfibrils (Sturcová et al., 2004; Fernandes et al., 2011). However, in growing cell walls, the microfibrils are not uniformly oriented. When microfibrils are first laid down at the inner face of the primary cell wall, their orientation is normally transverse to the direction of growth, but as the cell wall expands, the microfibrils reorient so that the orientation distribution, integrated across the thickness of the expanded cell wall, becomes progressively closer to random (Cosgrove, 2005; MacKinnon et al., 2006).This technical problem does not apply to the cell walls of celery (Apium graveolens) collenchyma, which are similar in composition to other primary cell walls but have their microfibrils oriented relatively uniformly along the cell axis (Sturcová et al., 2004; Kennedy et al., 2007a, 2007b). Some structural information on celery collenchyma cellulose has already been derived from spectroscopic and scattering experiments (Sturcová et al., 2004; Kennedy et al., 2007a, 2007b), confirming the disorder expected in a primary wall cellulose. Some of these experiments were analogous to what has been done on spruce cellulose (Fernandes et al., 2011), but insufficient data are available to specify the number of chains in each primary wall microfibril, the nature and location of the disorder, and the presence or absence of direct contact between microfibrils. Here, we report x-ray and neutron scattering and spectroscopic experiments addressing these questions and leading to a proposed structure for primary wall cellulose microfibrils. Characterizing a structure containing so much disorder presented unusual challenges, but disorder appears to be central to the enigmatic capacity of primary wall cellulose to provide high strength and yet to permit and control growth.     

Nontransgenic Genome Modification in Plant Cells

Zinc finger nucleases (ZFNs) are a powerful tool for genome editing in eukaryotic cells. ZFNs have been used for targeted mutagenesis in model and crop species. In animal and human cells, transient ZFN expression is often achieved by direct gene transfer into the target cells. Stable transformation, however, is the preferred method for gene expression in plant species, and ZFN-expressing transgenic plants have been used for recovery of mutants that are likely to be classified as transgenic due to the use of direct gene-transfer methods into the target cells. Here we present an alternative, nontransgenic approach for ZFN delivery and production of mutant plants using a novel Tobacco rattle virus (TRV)-based expression system for indirect transient delivery of ZFNs into a variety of tissues and cells of intact plants. TRV systemically infected its hosts and virus ZFN-mediated targeted mutagenesis could be clearly observed in newly developed infected tissues as measured by activation of a mutated reporter transgene in tobacco (Nicotiana tabacum) and petunia (Petunia hybrida) plants. The ability of TRV to move to developing buds and regenerating tissues enabled recovery of mutated tobacco and petunia plants. Sequence analysis and transmission of the mutations to the next generation confirmed the stability of the ZFN-induced genetic changes. Because TRV is an RNA virus that can infect a wide range of plant species, it provides a viable alternative to the production of ZFN-mediated mutants while avoiding the use of direct plant-transformation methods.Methods for genome editing in plant cells have fallen behind the remarkable progress made in whole-genome sequencing projects. The availability of reliable and efficient methods for genome editing would foster gene discovery and functional gene analyses in model plants and the introduction of novel traits in agriculturally important species (Puchta, 2002; Hanin and Paszkowski, 2003; Reiss, 2003; Porteus, 2009). Genome editing in various species is typically achieved by integrating foreign DNA molecules into the target genome by homologous recombination (HR). Genome editing by HR is routine in yeast (Saccharomyces cerevisiae) cells (Scherer and Davis, 1979) and has been adapted for other species, including Drosophila, human cell lines, various fungal species, and mouse embryonic stem cells (Baribault and Kemler, 1989; Venken and Bellen, 2005; Porteus, 2007; Hall et al., 2009; Laible and Alonso-González, 2009; Tenzen et al., 2009). In plants, however, foreign DNA molecules, which are typically delivered by direct gene-transfer methods (e.g. Agrobacterium and microbombardment of plasmid DNA), often integrate into the target cell genome via nonhomologous end joining (NHEJ) and not HR (Ray and Langer, 2002; Britt and May, 2003).Various methods have been developed to indentify and select for rare site-specific foreign DNA integration events or to enhance the rate of HR-mediated DNA integration in plant cells. Novel T-DNA molecules designed to support strong positive- and negative-selection schemes (e.g. Thykjaer et al., 1997; Terada et al., 2002), altering the plant DNA-repair machinery by expressing yeast chromatin remodeling protein (Shaked et al., 2005), and PCR screening of large numbers of transgenic plants (Kempin et al., 1997; Hanin et al., 2001) are just a few of the experimental approaches used to achieve HR-mediated gene targeting in plant species. While successful, these approaches, and others, have resulted in only a limited number of reports describing the successful implementation of HR-mediated gene targeting of native and transgenic sequences in plant cells (for review, see Puchta, 2002; Hanin and Paszkowski, 2003; Reiss, 2003; Porteus, 2009; Weinthal et al., 2010).HR-mediated gene targeting can potentially be enhanced by the induction of genomic double-strand breaks (DSBs). In their pioneering studies, Puchta et al. (1993, 1996) showed that DSB induction by the naturally occurring rare-cutting restriction enzyme I-SceI leads to enhanced HR-mediated DNA repair in plants. Expression of I-SceI and another rare-cutting restriction enzyme (I-CeuI) also led to efficient NHEJ-mediated site-specific mutagenesis and integration of foreign DNA molecules in plants (Salomon and Puchta, 1998; Chilton and Que, 2003; Tzfira et al., 2003). Naturally occurring rare-cutting restriction enzymes thus hold great promise as a tool for genome editing in plant cells (Carroll, 2004; Pâques and Duchateau, 2007). However, their wide application is hindered by the tedious and next to impossible reengineering of such enzymes for novel DNA-target specificities (Pâques and Duchateau, 2007).A viable alternative to the use of rare-cutting restriction enzymes is the zinc finger nucleases (ZFNs), which have been used for genome editing in a wide range of eukaryotic species, including plants (e.g. Bibikova et al., 2001; Porteus and Baltimore, 2003; Lloyd et al., 2005; Urnov et al., 2005; Wright et al., 2005; Beumer et al., 2006; Moehle et al., 2007; Santiago et al., 2008; Shukla et al., 2009; Tovkach et al., 2009; Townsend et al., 2009; Osakabe et al., 2010; Petolino et al., 2010; Zhang et al., 2010). Here too, ZFNs have been used to enhance DNA integration via HR (e.g. Shukla et al., 2009; Townsend et al., 2009) and as an efficient tool for the induction of site-specific mutagenesis (e.g. Lloyd et al., 2005; Zhang et al., 2010) in plant species. The latter is more efficient and simpler to implement in plants as it does not require codelivery of both ZFN-expressing and donor DNA molecules and it relies on NHEJ—the dominant DNA-repair machinery in most plant species (Ray and Langer, 2002; Britt and May, 2003).ZFNs are artificial restriction enzymes composed of a fusion between an artificial Cys₂His₂ zinc-finger protein DNA-binding domain and the cleavage domain of the FokI endonuclease. The DNA-binding domain of ZFNs can be engineered to recognize a variety of DNA sequences (for review, see Durai et al., 2005; Porteus and Carroll, 2005; Carroll et al., 2006). The FokI endonuclease domain functions as a dimer, and digestion of the target DNA requires proper alignment of two ZFN monomers at the target site (Durai et al., 2005; Porteus and Carroll, 2005; Carroll et al., 2006). Efficient and coordinated expression of both monomers is thus required for the production of DSBs in living cells. Transient ZFN expression, by direct gene delivery, is the method of choice for targeted mutagenesis in human and animal cells (e.g. Urnov et al., 2005; Beumer et al., 2006; Meng et al., 2008). Among the different methods used for high and efficient transient ZFN delivery in animal and human cell lines are plasmid injection (Morton et al., 2006; Foley et al., 2009), direct plasmid transfer (Urnov et al., 2005), the use of integrase-defective lentiviral vectors (Lombardo et al., 2007), and mRNA injection (Takasu et al., 2010).In plant species, however, efficient and strong gene expression is often achieved by stable gene transformation. Both transient and stable ZFN expression have been used in gene-targeting experiments in plants (Lloyd et al., 2005; Wright et al., 2005; Maeder et al., 2008; Cai et al., 2009; de Pater et al., 2009; Shukla et al., 2009; Tovkach et al., 2009; Townsend et al., 2009; Osakabe et al., 2010; Petolino et al., 2010; Zhang et al., 2010). In all cases, direct gene-transformation methods, using polyethylene glycol, silicon carbide whiskers, or Agrobacterium, were deployed. Thus, while mutant plants and tissues could be recovered, potentially without any detectable traces of foreign DNA, such plants were generated using a transgenic approach and are therefore still likely to be classified as transgenic. Furthermore, the recovery of mutants in many cases is also dependent on the ability to regenerate plants from protoplasts, a procedure that has only been successfully applied in a limited number of plant species. Therefore, while ZFN technology is a powerful tool for site-specific mutagenesis, its wider implementation for plant improvement may be somewhat limited, both by its restriction to certain plant species and by legislative restrictions imposed on transgenic plants.Here we describe an alternative to direct gene transfer for ZFN delivery and for the production of mutated plants. Our approach is based on the use of a novel Tobacco rattle virus (TRV)-based expression system, which is capable of systemically infecting its host and spreading into a variety of tissues and cells of intact plants, including developing buds and regenerating tissues. We traced the indirect ZFN delivery in infected plants by activation of a mutated reporter gene and we demonstrate that this approach can be used to recover mutated plants.