Long-Term Evolution of Nucleotide-Binding Site-Leucine-Rich Repeat Genes: Understanding Gained from and beyond the Legume Family

Proper utilization of plant disease resistance genes requires a good understanding of their short- and long-term evolution. Here we present a comprehensive study of the long-term evolutionary history of nucleotide-binding site (NBS)-leucine-rich repeat (LRR) genes within and beyond the legume family. The small group of NBS-LRR genes with an amino-terminal RESISTANCE TO POWDERY MILDEW8 (RPW8)-like domain (referred to as RNL) was first revealed as a basal clade sister to both coiled-coil-NBS-LRR (CNL) and Toll/Interleukin1 receptor-NBS-LRR (TNL) clades. Using Arabidopsis (Arabidopsis thaliana) as an outgroup, this study explicitly recovered 31 ancestral NBS lineages (two RNL, 21 CNL, and eight TNL) that had existed in the rosid common ancestor and 119 ancestral lineages (nine RNL, 55 CNL, and 55 TNL) that had diverged in the legume common ancestor. It was shown that, during their evolution in the past 54 million years, approximately 94% (112 of 119) of the ancestral legume NBS lineages experienced deletions or significant expansions, while seven original lineages were maintained in a conservative manner. The NBS gene duplication pattern was further examined. The local tandem duplications dominated NBS gene gains in the total number of genes (more than 75%), which was not surprising. However, it was interesting from our study that ectopic duplications had created many novel NBS gene loci in individual legume genomes, which occurred at a significant frequency of 8% to 20% in different legume lineages. Finally, by surveying the legume microRNAs that can potentially regulate NBS genes, we found that the microRNA-NBS gene interaction also exhibited a gain-and-loss pattern during the legume evolution.To combat the constant challenges by pathogens, plants have evolved a sophisticated two-layered defense system, in which proteins encoded by disease RESISTANCE (R) genes serve to sense pathogen invasion signals and to elicit defense responses (Jones and Dangl, 2006; McDowell and Simon, 2006; Bent and Mackey, 2007). Over 140 R genes have been characterized from different flowering plants, which confer resistance to a large array of pathogens, including bacteria, fungi, oomycetes, viruses, and nematodes (Liu et al., 2007; Yang et al., 2013). Among these, about 80% belong to the NBS-LRR class, which encodes a central nucleotide-binding site (NBS) domain and a C-terminal leucine-rich repeat (LRR) domain. Based on whether their N termini are homologous to the Toll/Interleukin1 receptor (TIR), the angiosperm NBS-LRR genes are further divided into the TIR-NBS-LRR (TNL) subclass and the non-TIR-NBS-LRR (nTNL) subclass (Meyers et al., 1999; Bai et al., 2002; Cannon et al., 2002). The latter has been also called CC-NBS-LRR (CNL) subclass, since a coiled-coil (CC) structure is often detected at the N terminus (Meyers et al., 2003). Interestingly, a small group of nTNL genes have an N-terminal RPW8-like domain with a transmembrane region before the CC structure (Xiao et al., 2001). This group of RPW8-NBS-LRR (RNL) genes has been usually viewed as a specific lineage of CNLs (Bonardi et al., 2011; Collier et al., 2011); however, its real phylogenetic relationship with CNLs and TNLs requires further investigation.NBS-LRR genes have been surveyed in many sequenced genomes of flowering plants, including four monocots: rice (Oryza sativa), maize (Zea mays), sorghum (Sorghum bicolor), and Brachypodium distachyon; one basal eudicot: Nelumbo nucifera; two asterid species: potato (Solanum tuberosum) and tomato (Solanum lycopersicum); and 14 rosids: Vitis vinifera, Populus trichocarpa, Ricinus communis, Medicago truncatula, soybean (Glycine max), Lotus japonicus, Cucumis sativus, Cucumis melo, Citrullus lanatus, Gossypium raimondii, Carica papaya, Arabidopsis (Arabidopsis thaliana), Arabidopsis lyrata, and Brassica rapa (Bai et al., 2002; Meyers et al., 2003; Monosi et al., 2004; Zhou et al., 2004; Yang et al., 2006, 2008b; Ameline-Torregrosa et al., 2008; Mun et al., 2009; Porter et al., 2009; Chen et al., 2010; Li et al., 2010a, 2010b; Guo et al., 2011; Zhang et al., 2011; Jupe et al., 2012; Lozano et al., 2012; Luo et al., 2012; Tan and Wu, 2012; Andolfo et al., 2013; Jia et al., 2013; Lin et al., 2013; Wan et al., 2013; Wei et al., 2013; Wu et al., 2014). Variable numbers (from dozens to hundreds) of NBS-LRR genes were reported from these genomes, making one wonder: how did these genes evolve so variably during flowering plant speciation?Comparative genomic studies conducted in the available genome sequences of closely related species or subspecies revealed that a significant proportion of NBS-LRR genes are not shared. For example, 70 NBS-LRR genes between Arabidopsis and A. lyrata show the presence/absence of polymorphisms (Chen et al., 2010; Guo et al., 2011). Moreover, synteny analysis revealed that, among 363 NBS-LRR gene loci in indica (cv 93-11) and japonica (cv Nipponbare) rice, 124 loci exist in only one genome (Luo et al., 2012). Unequal crossover, homologous repair, and nonhomologous repair are the three ways that NBS-LRR gene deletions are caused in rice genomes (Luo et al., 2012).In many surveyed genomes, the majority of NBS-LRR genes are found in a clustered organization (physically close to each other), with the rest exhibited as singletons. Many clusters are homogenous, with only duplicated members occupying the same phylogenetic lineage, whereas heterogenous clusters comprise members from distantly related clades (Meyers et al., 2003). Leister (2004) defined three types of NBS gene duplications: local tandem, ectopic, and segmental duplications. Although a general agreement on the widespread occurrence of local tandem duplications can be reached by various genome survey studies, the relative importance of ectopic and segmental duplications has been seldom investigated since the earliest surveys of the Arabidopsis genome (Richly et al., 2002; Baumgarten et al., 2003; Meyers et al., 2003; McDowell and Simon, 2006).With more genomic data available in certain angiosperm families, NBS-LRR genes should be further investigated among phylogenetically distant species to fill the gaps in the understanding of their long-term evolutionary patterns. The legume family contains many economically important crop species, such as clover (Trifolium spp.), soybean, peanut (Arachis hypogaea), and common bean (Phaseolus vulgaris). Although these legumes are constantly threatened by various pathogens, only a few functional legume R genes have been characterized, and all of them belong to the NBS-LRR class (Ashfield et al., 2004; Hayes et al., 2004; Gao et al., 2005; Seo et al., 2006; Yang et al., 2008a; Meyer et al., 2009). Therefore, it would be interesting to investigate the NBS-LRR gene repertoire among different legume species. Here, we carried out a comprehensive analysis of NBS-LRR genes in four divergent legume genomes, M. truncatula, pigeon pea (Cajanus cajan), common bean, and soybean, which shared a common ancestor approximately 54 million years ago (MYA; Fig. 1; Lavin et al., 2005). Approximately 1,000 nTNL and 667 TNL subclass NBS-encoding genes were identified in our study. Their genomic distribution, organization modes, phylogenetic relationships, and syntenic patterns were examined to obtain insight into the long-term evolutionary patterns of NBS-LRR genes.Open in a separate windowFigure 1.The phylogenetic tree of four investigated legume species (M. truncatula, pigeon pea, common bean, and soybean). Two WGD events are indicated with triangles: one occurred approximately 59 MYA in the common ancestor of the four investigated legumes, and the other occurred approximately 13 MYA in the Glycine spp. lineage alone (Schmutz et al., 2010). The numbers at the branch nodes indicate divergence times (Lavin et al., 2005; Stefanovic et al., 2009). [See online article for color version of this figure.]     

The Presence of Fucogalactoxyloglucan and Its Synthesis in Rice Indicates Conserved Functional Importance in Plants

The predominant structure of the hemicellulose xyloglucan (XyG) found in the cell walls of dicots is a fucogalactoXyG with an XXXG core motif, whereas in the Poaceae (grasses and cereals), the structure of XyG is less xylosylated (XXGG_n core motif) and lacks fucosyl residues. However, specialized tissues of rice (Oryza sativa) also contain fucogalactoXyG. Orthologous genes of the fucogalactoXyG biosynthetic machinery of Arabidopsis (Arabidopsis thaliana) are present in the rice genome. Expression of these rice genes, including fucosyl-, galactosyl-, and acetyltransferases, in the corresponding Arabidopsis mutants confirmed their activity and substrate specificity, indicating that plants in the Poaceae family have the ability to synthesize fucogalactoXyG in vivo. The data presented here provide support for a functional conservation of XyG structure in higher plants.The plant cell wall protects and structurally supports plant cells. The wall consists of a variety of polymers, including polysaccharides, the polyphenol lignin, and glycoproteins. One of the major polysaccharides present in the primary walls (i.e. walls of growing cells) in dicots is xyloglucan (XyG), which consists of a β-1,4-glucan backbone with xylosyl substituents. XyG binds noncovalently to cellulose microfibrils and thereby, is thought to act as a spacer molecule, hindering cellulose microfibrils to aggregate (Carpita and Gibeaut, 1993; Pauly et al., 1999a; Bootten et al., 2004; Cosgrove, 2005; Hayashi and Kaida, 2011; Park and Cosgrove, 2012).The side-chain substitutions on XyG can be structurally diverse depending on plant species, tissue type, and developmental stage of the tissue (Pauly et al., 2001; Hoffman et al., 2005; Peña et al., 2008; Hsieh and Harris, 2009, 2012; Lampugnani et al., 2013; Schultink et al., 2014). A one-letter code nomenclature has been established to specify the XyG side-chain substitutions (Fry et al., 1993; Tuomivaara et al., 20145). According to this nomenclature, an unsubstituted glucosyl residue is indicated by a G, whereas a glucosyl residue substituted with a xylosyl moiety is shown as an X. In most dicots, the xylosyl residue can be further substituted with a galactosyl residue (L), which in turn, can be further decorated with a fucosyl residue (F) and/or an acetyl group (F/L). In some species, the xylosyl residue can be substituted with an arabinosyl moiety (S), and the backbone glucosyl residue can be O-acetylated (G; Jia et al., 2003; Hoffman et al., 2005).Numerous genes have been identified in Arabidopsis (Arabidopsis thaliana) that are involved in fucogalactoXyG biosynthesis (Fig. 1; Pauly et al., 2013; Schultink et al., 2014). The glucan backbone is thought to be synthesized by cellulose synthase-like C (CSLC) family proteins, such as AtCSLC4, as shown by in vitro activity data (Cocuron et al., 2007). Several xylosyltransferases (XXTs) from glycosyl transferase family 34 (GT34) are thought to be responsible for XyG xylosylation. Five of these XXTs in Arabidopsis seem to have XXT activity on XyG in vitro (Faik et al., 2002; Zabotina et al., 2008; Vuttipongchaikij et al., 2012; Mansoori et al., 2015). MURUS3 (MUR3) represents a galactosyltransferase that transfers galactosyl moieties specifically to xylosyl residues adjacent to an unsubstituted glucosyl residue on an XXXG unit, converting it to XXLG, whereas Xyloglucan L-side chain galactosyl Transferase2 (XLT2) was identified as another galactosyltransferase transferring a galactosyl moiety specifically to the second xylosyl residue, resulting in XLXG (Madson et al., 2003; Jensen et al., 2012). Both MUR3 and XLT2 belong to GT47 (Li et al., 2004). MUR2/FUCOSYLTRANSFERASE1 (FUT1) from GT37 was found to harbor fucosyltransferase activity, transferring Fuc from GDP-Fuc to a galactosyl residue adjacent to the unsubstituted glucosyl residue (i.e. onto XXLG but not onto XLXG; Perrin et al., 1999; Vanzin et al., 2002). O-acetylation of the galactosyl residue is mediated by Altered Xyloglucan4 (AXY4) and AXY4L, both of which belong to the Trichome Birefringence-Like (TBL) protein family (Bischoff et al., 2010; Gille et al., 2011; Gille and Pauly, 2012).Open in a separate windowFigure 1.Schematic structures of two types of XyGs and known biosynthetic proteins in Arabidopsis (Hsieh and Harris, 2009; Pauly et al., 2013). The corresponding one-letter code for XyG is shown below the pictograms (Fry et al., 1993; Tuomivaara et al., 2015).XyG found throughout land plants exhibits structural diversity with respect to side-chain substitution patterns (Schultink et al., 2014). Most dicots, such as Arabidopsis, and the noncommelinoid monocots possess a fucogalactoXyG of the XXXG-type XyG structure as shown in Figure 1. However, plant species in the Solanaceae and Poaceae as well as the moss Physcomitrella patens contain a different XyG structure with a reduced level of xylosylation, resulting in an XXGG_n core motif (York et al., 1996; Kato et al., 2004; Gibeaut et al., 2005; Jia et al., 2005; Peña et al., 2008; Hsieh and Harris, 2009). In addition, the glucan backbone can be O-acetylated in plants of Solanaceae and Poaceae families (Gibeaut et al., 2005; Jia et al., 2005). XyG from Solanaceae with an XXGG core motif can be further arabinosylated and/or galactosylated (Jia et al., 2005). No XyGs with an XXGG_n motif backbone have been reported to be fucosylated.The function of structural diversity of XyG substitutions, such as fucosylation and/or altered xylosylation pattern, remains enigmatic. Removing the terminal fucosyl or acetyl moieties in the corresponding Arabidopsis mutants does not lead to any change in plant growth and development (Vanzin et al., 2002; Gille et al., 2011). However, removing galactosyl residues as well as fucosyl and acetyl moieties in the Arabidopsis xlt2 mur3.1 double mutant results in a dwarfed plant (Jensen et al., 2012; Kong et al., 2015). Replacing the galactosyl moiety with an arabinofuranosyl residue by, for example, expressing a tomato (Solanum lycopersicum) arabinosyltransferase in the Arabidopsis xlt2 mur3.1 mutant rescues the growth phenotype and restores wall biomechanics, indicating that galactosylation and arabinosylation in XyG have an equivalent function (Schultink et al., 2013). Recently, fucosylated XyG structures were found in the pollen tubes of tobacco (Nicotiana alata) and tomato, indicating that fucogalactoXyG is likely also present in other Solanaceae plants, albeit restricted to specific tissues (Lampugnani et al., 2013; Dardelle et al., 2015). Although there is circumstantial evidence that fucogalactoXyG is present in cell suspension cultures of rice (Oryza sativa) and cell suspension cultures of fescue (Festuca arundinaceae; McDougall and Fry, 1994; Peña et al., 2008), fucogalactoXyG has not been found in any physiologically relevant plant tissues of members of the Poaceae (Kato et al., 1982; Watanabe et al., 1984; Gibeaut et al., 2005; Hsieh and Harris, 2009; Brennan and Harris, 2011). Here, we provide chemical and genetic evidence that fucogalactoXyG is, indeed, present in plant tissues of a grass (rice) and prove that the rice genome harbors the genes that could be part of the synthetic machinery necessary to produce fucogalactoXyG.     

Spore Density Determines Infection Strategy by the Plant Pathogenic Fungus Plectosphaerella cucumerina

Necrotrophic and biotrophic pathogens are resisted by different plant defenses. While necrotrophic pathogens are sensitive to jasmonic acid (JA)-dependent resistance, biotrophic pathogens are resisted by salicylic acid (SA)- and reactive oxygen species (ROS)-dependent resistance. Although many pathogens switch from biotrophy to necrotrophy during infection, little is known about the signals triggering this transition. This study is based on the observation that the early colonization pattern and symptom development by the ascomycete pathogen Plectosphaerella cucumerina (P. cucumerina) vary between inoculation methods. Using the Arabidopsis (Arabidopsis thaliana) defense response as a proxy for infection strategy, we examined whether P. cucumerina alternates between hemibiotrophic and necrotrophic lifestyles, depending on initial spore density and distribution on the leaf surface. Untargeted metabolome analysis revealed profound differences in metabolic defense signatures upon different inoculation methods. Quantification of JA and SA, marker gene expression, and cell death confirmed that infection from high spore densities activates JA-dependent defenses with excessive cell death, while infection from low spore densities induces SA-dependent defenses with lower levels of cell death. Phenotyping of Arabidopsis mutants in JA, SA, and ROS signaling confirmed that P. cucumerina is differentially resisted by JA- and SA/ROS-dependent defenses, depending on initial spore density and distribution on the leaf. Furthermore, in situ staining for early callose deposition at the infection sites revealed that necrotrophy by P. cucumerina is associated with elevated host defense. We conclude that P. cucumerina adapts to early-acting plant defenses by switching from a hemibiotrophic to a necrotrophic infection program, thereby gaining an advantage of immunity-related cell death in the host.Plant pathogens are often classified as necrotrophic or biotrophic, depending on their infection strategy (Glazebrook, 2005; Nishimura and Dangl, 2010). Necrotrophic pathogens kill living host cells and use the decayed plant tissue as a substrate to colonize the plant, whereas biotrophic pathogens parasitize living plant cells by employing effector molecules that suppress the host immune system (Pel and Pieterse, 2013). Despite this binary classification, the majority of pathogenic microbes employ a hemibiotrophic infection strategy, which is characterized by an initial biotrophic phase followed by a necrotrophic infection strategy at later stages of infection (Perfect and Green, 2001). The pathogenic fungi Magnaporthe grisea, Sclerotinia sclerotiorum, and Mycosphaerella graminicola, the oomycete Phytophthora infestans, and the bacterial pathogen Pseudomonas syringae are examples of hemibiotrophic plant pathogens (Perfect and Green, 2001; Koeck et al., 2011; van Kan et al., 2014; Kabbage et al., 2015).Despite considerable progress in our understanding of plant resistance to necrotrophic and biotrophic pathogens (Glazebrook, 2005; Mengiste, 2012; Lai and Mengiste, 2013), recent debate highlights the dynamic and complex interplay between plant-pathogenic microbes and their hosts, which is raising concerns about the use of infection strategies as a static tool to classify plant pathogens. For instance, the fungal genus Botrytis is often labeled as an archetypal necrotroph, even though there is evidence that it can behave as an endophytic fungus with a biotrophic lifestyle (van Kan et al., 2014). The rice blast fungus Magnaporthe oryzae, which is often classified as a hemibiotrophic leaf pathogen (Perfect and Green, 2001; Koeck et al., 2011), can adopt a purely biotrophic lifestyle when infecting root tissues (Marcel et al., 2010). It remains unclear which signals are responsible for the switch from biotrophy to necrotrophy and whether these signals rely solely on the physiological state of the pathogen, or whether host-derived signals play a role as well (Kabbage et al., 2015).The plant hormones salicylic acid (SA) and jasmonic acid (JA) play a central role in the activation of plant defenses (Glazebrook, 2005; Pieterse et al., 2009, 2012). The first evidence that biotrophic and necrotrophic pathogens are resisted by different immune responses came from Thomma et al. (1998), who demonstrated that Arabidopsis (Arabidopsis thaliana) genotypes impaired in SA signaling show enhanced susceptibility to the biotrophic pathogen Hyaloperonospora arabidopsidis (formerly known as Peronospora parastitica), while JA-insensitive genotypes were more susceptible to the necrotrophic fungus Alternaria brassicicola. In subsequent years, the differential effectiveness of SA- and JA-dependent defense mechanisms has been confirmed in different plant-pathogen interactions, while additional plant hormones, such as ethylene, abscisic acid (ABA), auxins, and cytokinins, have emerged as regulators of SA- and JA-dependent defenses (Bari and Jones, 2009; Cao et al., 2011; Pieterse et al., 2012). Moreover, SA- and JA-dependent defense pathways have been shown to act antagonistically on each other, which allows plants to prioritize an appropriate defense response to attack by biotrophic pathogens, necrotrophic pathogens, or herbivores (Koornneef and Pieterse, 2008; Pieterse et al., 2009; Verhage et al., 2010).In addition to plant hormones, reactive oxygen species (ROS) play an important regulatory role in plant defenses (Torres et al., 2006; Lehmann et al., 2015). Within minutes after the perception of pathogen-associated molecular patterns, NADPH oxidases and apoplastic peroxidases generate early ROS bursts (Torres et al., 2002; Daudi et al., 2012; O’Brien et al., 2012), which activate downstream defense signaling cascades (Apel and Hirt, 2004; Torres et al., 2006; Miller et al., 2009; Mittler et al., 2011; Lehmann et al., 2015). ROS play an important regulatory role in the deposition of callose (Luna et al., 2011; Pastor et al., 2013) and can also stimulate SA-dependent defenses (Chaouch et al., 2010; Yun and Chen, 2011; Wang et al., 2014; Mammarella et al., 2015). However, the spread of SA-induced apoptosis during hyperstimulation of the plant immune system is contained by the ROS-generating NADPH oxidase RBOHD (Torres et al., 2005), presumably to allow for the sufficient generation of SA-dependent defense signals from living cells that are adjacent to apoptotic cells. Nitric oxide (NO) plays an additional role in the regulation of SA/ROS-dependent defense (Trapet et al., 2015). This gaseous molecule can stimulate ROS production and cell death in the absence of SA while preventing excessive ROS production at high cellular SA levels via S-nitrosylation of RBOHD (Yun et al., 2011). Recently, it was shown that pathogen-induced accumulation of NO and ROS promotes the production of azelaic acid, a lipid derivative that primes distal plants for SA-dependent defenses (Wang et al., 2014). Hence, NO, ROS, and SA are intertwined in a complex regulatory network to mount local and systemic resistance against biotrophic pathogens. Interestingly, pathogens with a necrotrophic lifestyle can benefit from ROS/SA-dependent defenses and associated cell death (Govrin and Levine, 2000). For instance, Kabbage et al. (2013) demonstrated that S. sclerotiorum utilizes oxalic acid to repress oxidative defense signaling during initial biotrophic colonization, but it stimulates apoptosis at later stages to advance necrotrophic colonization. Moreover, SA-induced repression of JA-dependent resistance not only benefits necrotrophic pathogens but also hemibiotrophic pathogens after having switched from biotrophy to necrotrophy (Glazebrook, 2005; Pieterse et al., 2009, 2012).Plectosphaerella cucumerina ((P. cucumerina, anamorph Plectosporum tabacinum) anamorph Plectosporum tabacinum) is a filamentous ascomycete fungus that can survive saprophytically in soil by decomposing plant material (Palm et al., 1995). The fungus can cause sudden death and blight disease in a variety of crops (Chen et al., 1999; Harrington et al., 2000). Because P. cucumerina can infect Arabidopsis leaves, the P. cucumerina-Arabidopsis interaction has emerged as a popular model system in which to study plant defense reactions to necrotrophic fungi (Berrocal-Lobo et al., 2002; Ton and Mauch-Mani, 2004; Carlucci et al., 2012; Ramos et al., 2013). Various studies have shown that Arabidopsis deploys a wide range of inducible defense strategies against P. cucumerina, including JA-, SA-, ABA-, and auxin-dependent defenses, glucosinolates (Tierens et al., 2001; Sánchez-Vallet et al., 2010; Gamir et al., 2014; Pastor et al., 2014), callose deposition (García-Andrade et al., 2011; Gamir et al., 2012, 2014; Sánchez-Vallet et al., 2012), and ROS (Tierens et al., 2002; Sánchez-Vallet et al., 2010; Barna et al., 2012; Gamir et al., 2012, 2014; Pastor et al., 2014). Recent metabolomics studies have revealed large-scale metabolic changes in P. cucumerina-infected Arabidopsis, presumably to mobilize chemical defenses (Sánchez-Vallet et al., 2010; Gamir et al., 2014; Pastor et al., 2014). Furthermore, various chemical agents have been reported to induce resistance against P. cucumerina. These chemicals include β-amino-butyric acid, which primes callose deposition and SA-dependent defenses, benzothiadiazole (BTH or Bion; Görlach et al., 1996; Ton and Mauch-Mani, 2004), which activates SA-related defenses (Lawton et al., 1996; Ton and Mauch-Mani, 2004; Gamir et al., 2014; Luna et al., 2014), JA (Ton and Mauch-Mani, 2004), and ABA, which primes ROS and callose deposition (Ton and Mauch-Mani, 2004; Pastor et al., 2013). However, among all these studies, there is increasing controversy about the exact signaling pathways and defense responses contributing to plant resistance against P. cucumerina. While it is clear that JA and ethylene contribute to basal resistance against the fungus, the exact roles of SA, ABA, and ROS in P. cucumerina resistance vary between studies (Thomma et al., 1998; Ton and Mauch-Mani, 2004; Sánchez-Vallet et al., 2012; Gamir et al., 2014).This study is based on the observation that the disease phenotype during P. cucumerina infection differs according to the inoculation method used. We provide evidence that the fungus follows a hemibiotrophic infection strategy when infecting from relatively low spore densities on the leaf surface. By contrast, when challenged by localized host defense to relatively high spore densities, the fungus switches to a necrotrophic infection program. Our study has uncovered a novel strategy by which plant-pathogenic fungi can take advantage of the early immune response in the host plant.     

The Arabidopsis Chloroplast Stromal N-Terminome: Complexities of Amino-Terminal Protein Maturation and Stability

Protein amino (N) termini are prone to modifications and are major determinants of protein stability in bacteria, eukaryotes, and perhaps also in chloroplasts. Most chloroplast proteins undergo N-terminal maturation, but this is poorly understood due to insufficient experimental information. Consequently, N termini of mature chloroplast proteins cannot be accurately predicted. This motivated an extensive characterization of chloroplast protein N termini in Arabidopsis (Arabidopsis thaliana) using terminal amine isotopic labeling of substrates and mass spectrometry, generating nearly 14,000 tandem mass spectrometry spectra matching to protein N termini. Many nucleus-encoded plastid proteins accumulated with two or three different N termini; we evaluated the significance of these different proteoforms. Alanine, valine, threonine (often in N-α-acetylated form), and serine were by far the most observed N-terminal residues, even after normalization for their frequency in the plastid proteome, while other residues were absent or highly underrepresented. Plastid-encoded proteins showed a comparable distribution of N-terminal residues, but with a higher frequency of methionine. Infrequent residues (e.g. isoleucine, arginine, cysteine, proline, aspartate, and glutamate) were observed for several abundant proteins (e.g. heat shock proteins 70 and 90, Rubisco large subunit, and ferredoxin-glutamate synthase), likely reflecting functional regulation through their N termini. In contrast, the thylakoid lumenal proteome showed a wide diversity of N-terminal residues, including those typically associated with instability (aspartate, glutamate, leucine, and phenylalanine). We propose that, after cleavage of the chloroplast transit peptide by stromal processing peptidase, additional processing by unidentified peptidases occurs to avoid unstable or otherwise unfavorable N-terminal residues. The possibility of a chloroplast N-end rule is discussed.Following synthesis, most proteins undergo various N-terminal (Nt) protein modifications, including removal of the Nt Met and signal peptide, N-terminal α-acetylation (NAA), ubiquitination, and acylations. These Nt modifications play an important role in the regulation of cellular functions. The N terminus of proteins has also been shown to be a major determinant of protein stability in bacteria (Varshavsky, 2011), eukaryotes (Graciet et al., 2009), mitochondria, and perhaps in plastids/chloroplasts (Apel et al., 2010; Nishimura et al., 2013; van Wijk, 2015). The role of the N terminus in protein stability is conceptualized in the N-end rule, which states that certain amino acids, when exposed at the N terminus of a protein, act as triggers for degradation (Bachmair et al., 1986; Dougan et al., 2012; Tasaki et al., 2012; Gibbs et al., 2014).Most of the approximately 3,000 plastid proteins are nucleus encoded (n-encoded) and are targeted to the plastid through an Nt chloroplast transit peptide (cTP). After import, the cTP is cleaved by the stromal processing peptidase (SPP; Richter and Lamppa, 1998; Trösch and Jarvis, 2011). The consensus site of cTP cleavage by SPP is only loosely defined, and the rules, mechanisms, and enzymes for possible subsequent processing, stabilization, and other posttranslational modifications (PTMs) are not well characterized (for discussion, see van Wijk, 2015). The exact N terminus is unknown for many chloroplast proteins and cannot be accurately predicted, because SPP specificity is not sufficiently understood (Emanuelsson et al., 2000; Zybailov et al., 2008) and probably also because additional Nt processing occurs for many chloroplast proteins (Fig. 1A). The approximately 85 plastid-encoded (p-encoded) proteins typically first undergo cotranslational Nt deformylation, followed by N-terminal Met excision (NME; Giglione et al., 2009; Fig. 1B); both these PTMs are required for normal plastid/chloroplast development and protein stability (Dirk et al., 2001, 2002; Giglione et al., 2003; Meinnel et al., 2006). Both n-encoded and p-encoded proteins can undergo NAA inside the plastid (Zybailov et al., 2008; Fig. 1). Postulated functions of NAA in eukaryotes include the mediation of protein location, assembly, and stability (Jones and O’Connor, 2011; Starheim et al., 2012; Hoshiyasu et al., 2013; Xu et al., 2015), thereby affecting a variety of processes, including drought tolerance in Arabidopsis (Arabidopsis thaliana; Linster et al., 2015).Open in a separate windowFigure 1.Conceptual illustration of Nt maturation of n-encoded and p-encoded proteins. Ac, Acetylated; MAP, Met amino peptidase; NAT, N-acetyltransferase; N-term, N-terminal; PDF, peptide deformylase. A, Nt maturation of n-encoded plastid proteins including removal of cTP by SPP and potential subsequent Nt modifications. B, Nt maturation of p-encoded proteins. *, The removal depends on the penultimate residue, generally following the N-terminal Met Excision (NME) rule; **, N-terminal acetylation typically occurs only for selected residues; “Results”).Typical proteomics work flows generally yield only partial coverage of protein sequences, and it is often difficult to know which peptides represent the true N termini (Nti) or C termini. Systematic identification of Nti or C termini requires specific labeling and enrichment strategies, such as combined fractional diagonal chromatography, developed by Gevaert and colleagues (Staes et al., 2011), and terminal amine isotopic labeling of substrates (TAILS), developed by the group of Overall (Kleifeld et al., 2011; Lange and Overall, 2013). These strategies allow the identification of different Nt proteoforms and were recently also applied to plants (Tsiatsiani et al., 2013; Carrie et al., 2015; Kohler et al., 2015; Zhang et al., 2015) and diatoms (Huesgen et al., 2013). We previously reported on Nti of chloroplast proteins based on tandem mass spectrometry (MS/MS) analysis, but because no Nt enrichment/labeling technique was used, only those that underwent NAA could be considered bona fide Nti (Zybailov et al., 2008). Nt Edman degradation sequencing was systematically carried out for thylakoid lumen proteins (Peltier et al., 2000, 2002) but not for stromal proteins or chloroplast membrane proteins with their Nti exposed to the stroma. The Nti of thylakoid lumen proteins are mostly generated by lumenal peptidases (Hsu et al., 2011; Midorikawa et al., 2014), and the thylakoid lumen contains a different set of peptidases than the stroma; hence, rules for Nt maturation and stability are likely different than those for stroma-exposed proteins.The objective of this study was to systematically determine the Nti of stroma-exposed chloroplast proteins of Arabidopsis (the N-terminome) and to provide a baseline for understanding Nt protein maturation and protein stability in the chloroplast stroma. To that end, we applied the TAILS technique and determined the Nti of approximately 250 chloroplast proteins by mass spectrometry (MS). We observed that many n-encoded plastid proteins accumulated with two or even three different Nt residues, in many cases both with and without NAA. The extent of accumulation of different Nt proteoforms is surprising and will be discussed. The p-encoded proteins generally showed very similar Nt residues as compared with the n-encoded proteins, with the exception of Met. Our data show that small, apolar, or hydroxylated residues (Ala, Val, Ser, and Thr) are the most frequent Nt residues of stromal proteins, whereas other residues are strictly avoided or are only present for very specific proteins likely to aid in their function. Chloroplast protein degradation products were also detected, with enrichment for peptides generated by cleavage between Arg and Thr residues. We present testable hypotheses for understanding Nt processing and maturation, stability, and a possible N-end rule in chloroplast stroma.     

Association of the P6 Protein of Cauliflower mosaic virus with Plasmodesmata and Plasmodesmal Proteins

The P6 protein of Cauliflower mosaic virus (CaMV) is responsible for the formation of inclusion bodies (IBs), which are the sites for viral gene expression, replication, and virion assembly. Moreover, recent evidence indicates that ectopically expressed P6 inclusion-like bodies (I-LBs) move in association with actin microfilaments. Because CaMV virions accumulate preferentially in P6 IBs, we hypothesized that P6 IBs have a role in delivering CaMV virions to the plasmodesmata. We have determined that the P6 protein interacts with a C2 calcium-dependent membrane-targeting protein (designated Arabidopsis [Arabidopsis thaliana] Soybean Response to Cold [AtSRC2.2]) in a yeast (Saccharomyces cerevisiae) two-hybrid screen and have confirmed this interaction through coimmunoprecipitation and colocalization assays in the CaMV host Nicotiana benthamiana. An AtSRC2.2 protein fused to red fluorescent protein (RFP) was localized to the plasma membrane and specifically associated with plasmodesmata. The AtSRC2.2-RFP fusion also colocalized with two proteins previously shown to associate with plasmodesmata: the host protein Plasmodesmata-Localized Protein1 (PDLP1) and the CaMV movement protein (MP). Because P6 I-LBs colocalized with AtSRC2.2 and the P6 protein had previously been shown to interact with CaMV MP, we investigated whether P6 I-LBs might also be associated with plasmodesmata. We examined the colocalization of P6-RFP I-LBs with PDLP1-green fluorescent protein (GFP) and aniline blue (a stain for callose normally observed at plasmodesmata) and found that P6-RFP I-LBs were associated with each of these markers. Furthermore, P6-RFP coimmunoprecipitated with PDLP1-GFP. Our evidence that a portion of P6-GFP I-LBs associate with AtSRC2.2 and PDLP1 at plasmodesmata supports a model in which P6 IBs function to transfer CaMV virions directly to MP at the plasmodesmata.Through the years, numerous studies have focused on the characterization of viral replication sites within the cell, as well as how plant virus movement proteins (MPs) modify the plasmodesmata to facilitate cell-to-cell movement (for review, see Benitez-Alfonso et al., 2010; Laliberté and Sanfaçon, 2010; Niehl and Heinlein, 2011; Ueki and Citovsky, 2011; Verchot, 2012). It is accepted that plant virus replication is associated with host membranes, and at some point, the viral genomic nucleic acid must be transferred from the site of replication in the cell to the plasmodesmata. This step could involve transport from a distant site within the cell, or alternatively, it may be that replication is coupled with transport at the entrance of the plasmodesmata (Tilsner et al., 2013). However, even with the latter model, there is ample evidence that the viral proteins necessary for replication or cell-to-cell movement utilize intracellular trafficking pathways within the cell to become positioned at the plasmodesma. These pathways may involve microfilaments, microtubules, or specific endomembranes that participate in macromolecular transport pathways, or combinations of these elements (Harries et al., 2010; Schoelz et al., 2011; Patarroyo et al., 2012; Peña and Heinlein, 2012; Tilsner and Oparka 2012; Liu and Nelson, 2013).The P6 protein of Cauliflower mosaic virus (CaMV) is one viral protein that had not been considered to play a role in viral movement until recently. P6 is the most abundant protein component of the amorphous, electron-dense inclusion bodies (IBs) present during virus infection (Odell and Howell, 1980; Shockey et al., 1980). Ectopic expression of P6 in Nicotiana benthamiana leaves resulted in the formation of inclusion-like bodies (I-LBs) that were capable of intracellular movement along actin microfilaments. Furthermore, treatment of Nicotiana edwardsonii leaves with latrunculin B abolished the formation of CaMV local lesions, suggesting that intact microfilaments are required for CaMV infection (Harries et al., 2009a). A subsequent paper showed that P6 physically interacts with Chloroplast Unusual Positioning1 (CHUP1), a plant protein localized to the chloroplast outer membrane that contributes to movement of chloroplasts on microfilaments in response to changes in light intensity (Oikawa et al., 2003, 2008; Angel et al., 2013). The implication was that P6 might hijack CHUP1 to facilitate movement of the P6 IBs on microfilaments. Silencing of CHUP1 in N. edwardsonii, a host for CaMV, slowed the rate of local lesion formation, suggesting that CHUP1 contributes to intracellular movement of CaMV (Angel et al., 2013).In addition to its role in intracellular trafficking, the P6 protein has been shown to have at least four other distinct functions in the viral infection cycle. P6-containing IBs induced during virus infection are likely virion factories, as they are the primary site for CaMV protein synthesis, genome replication, and assembly of virions (Hohn and Fütterer, 1997). Second, P6 interacts with host ribosomes to facilitate reinitiation of translation of genes on the polycistronic 35S viral RNA, a process called translational transactivation (Bonneville et al., 1989; Park et al., 2001; Ryabova et al., 2002). The translational transactivator region of P6 (Fig. 1) defines the essential sequences required for translational transactivation (DeTapia et al., 1993). Third, P6 is an important pathogenicity determinant. P6 functions as an avirulence determinant in some solanaceous and cruciferous species (Daubert et al., 1984; Schoelz et al., 1986; Hapiak et al., 2008) and is a chlorosis symptom determinant in susceptible hosts (Daubert et al., 1984; Baughman et al., 1988; Goldberg et al., 1991; Cecchini et al., 1997). Finally, P6 has the capacity to compromise host defenses, as it is a suppressor of RNA silencing and cell death (Love et al., 2007; Haas et al., 2008), and it modulates signaling by salicylic acid, jasmonic acid, ethylene, and auxin (Geri et al., 2004; Love et al., 2012; Laird et al., 2013). Domain D1 of P6 has been shown to be necessary but not sufficient for suppression of silencing and salicylic acid-mediated defenses (Laird et al., 2013).Open in a separate windowFigure 1.CaMV and host constructs used for confocal microscopy or coimmunoprecipitation (co-IP). A, Structure of CaMV P6 and Arabidopsis (Arabidopsis thaliana) Soybean Response to Cold (AtSRC2.2) proteins. The functions of P6 domains D1 to D4 tested for interaction with AtSRC2.2 are indicated by the shaded boxes. The Mini TAV is the minimal region for the translational transactivation function. The NLSa sequence corresponds to the nuclear localization signal of influenza virus. The NLS sequence corresponds to the nuclear localization signal of human ribosomal protein L22. B, Structure of P6 (Angel et al., 2013), AtSRC2.2, PDLP (Thomas et al., 2008), and CaMV MP fusions developed for confocal microscopy and/or co-IP. aa, Amino acid.Because P6-containing IBs are the site for virion accumulation and they are capable of movement, they may be responsible for delivering virions to the CaMV MP located at the plasmodesmata (for review, see Schoelz et al., 2011). The vast majority of CaMV virions accumulate in association with P6-containing IBs. Furthermore, P6 physically interacts with the CaMV capsid and MP, as well as the two proteins necessary for aphid transmission, P2 and P3 (Himmelbach et al., 1996; Ryabova et al., 2002; Hapiak et al., 2008; Lutz et al., 2012). Recent studies have indicated that P6 IBs serve as a reservoir for virions, in which the virions may be rapidly transferred to P2 electron-lucent IBs for acquisition by aphids (Bak et al., 2013). It stands to reason that P6 IBs may also serve as a reservoir for CaMV virions to be transferred to the CaMV MP in the plasmodesmata.CaMV virions move from cell to cell through plasmodesmata modified into tubules through the function of its MP (Perbal et al., 1993; Kasteel et al., 1996). However, studies have suggested that CaMV virions do not appear to directly interact with the MP. Instead, the MP interacts with the CaMV P3 protein (also known as the virion-associated protein [VAP]), which forms a trimeric structure that is anchored into the virions (Leclerc et al., 1998; Leclerc et al., 2001). Electron microscopy studies have indicated that MP and VAP colocalize with virions only at the entrance to or within the plasmodesmata, and it has been suggested that the VAP/virion complex travels to the plasmodesmata independently from the MP (Stavolone et al., 2005). Consequently, there is a need for a second CaMV protein such as P6 to fulfill the role of delivery of virions to the plasmodesmata (Schoelz et al., 2011).Additional studies have shown that the CaMV MP is incorporated into vesicles and is trafficked on the endomembrane system to reach the plasmodesma (Carluccio et al., 2014). These authors suggest that the CaMV MP is recycled in a vesicular transport pathway between plasmodesmata and early endosome compartments. The CaMV MP interacts with µA-Adaptin (Carluccio et al., 2014) and Movement Protein-Interacting7 (Huang et al., 2001), two proteins shown to have a role in vesicular trafficking. Once the MP arrives at plasmodesmata, it interacts with the Plasmodesmata-Localized Protein (PDLP) proteins, which comprise a family of eight proteins associated with plasmodesmata (Amari et al., 2010). In addition to its interaction with CaMV MP, PDLP1 interacts with the 2B protein of Grapevine fan leaf virus (GFLV) at the base of tubules formed by the 2B protein. Furthermore, an Arabidopsis transfer DNA (T-DNA) mutant line in which three PDLP genes had been knocked out (pdlp1-pdlp2-pdlp3) responded to GFLV and CaMV inoculation with a delayed infection (Amari et al., 2010). This has led to the suggestion that the PDLPs might act as receptors for the MPs of the tubule-forming viruses such as GFLV and CaMV (Amari et al., 2010, 2011).To better understand the function of the P6 protein during CaMV intracellular movement, we have utilized a yeast (Saccharomyces cerevisiae) two-hybrid assay to identify host proteins that interact with CaMV P6. We show that P6 physically interacts with a C2-calcium-dependent protein (designated AtSRC2.2). AtSRC2.2 is a membrane-bound protein that is capable of forming punctate spots associated with plasmodesmata. The localization of AtSRC2.2 with plasmodesmata led to an analysis of interactions between P6 I-LBs, AtSRC2.2, PDLP1, and the CaMV MP and also revealed that a portion of P6 I-LBs are found adjacent to plasmodesmata. These results provide further evidence for a model in which P6 IBs are capable of delivery of virions to plasmodesmata for their transit to other host cells.     

Focus on Water: Leaf Shrinkage with Dehydration: Coordination with Hydraulic Vulnerability and Drought Tolerance

Leaf shrinkage with dehydration has attracted attention for over 100 years, especially as it becomes visibly extreme during drought. However, little has been known of its correlation with physiology. Computer simulations of the leaf hydraulic system showed that a reduction of hydraulic conductance of the mesophyll pathways outside the xylem would cause a strong decline of leaf hydraulic conductance (K_leaf). For 14 diverse species, we tested the hypothesis that shrinkage during dehydration (i.e. in whole leaf, cell and airspace thickness, and leaf area) is associated with reduction in K_leaf at declining leaf water potential (Ψ_leaf). We tested hypotheses for the linkage of leaf shrinkage with structural and physiological water relations parameters, including modulus of elasticity, osmotic pressure at full turgor, turgor loss point (TLP), and cuticular conductance. Species originating from moist habitats showed substantial shrinkage during dehydration before reaching TLP, in contrast with species originating from dry habitats. Across species, the decline of K_leaf with mild dehydration (i.e. the initial slope of the K_leaf versus Ψ_leaf curve) correlated with the decline of leaf thickness (the slope of the leaf thickness versus Ψ_leaf curve), as expected based on predictions from computer simulations. Leaf thickness shrinkage before TLP correlated across species with lower modulus of elasticity and with less negative osmotic pressure at full turgor, as did leaf area shrinkage between full turgor and oven desiccation. These findings point to a role for leaf shrinkage in hydraulic decline during mild dehydration, with potential impacts on drought adaptation for cells and leaves, influencing plant ecological distributions.As leaves open their stomata to capture CO₂ for photosynthesis, water is lost to transpiration, which needs to be replaced by flow through the hydraulic system. The leaf hydraulic system has two components, which act essentially in series: the pathways for water movement through the xylem from the petiole to leaf minor veins, and those through the living bundle sheath and mesophyll cells to the sites of evaporation (Tyree and Zimmermann, 2002; Sack et al., 2004; Sack and Holbrook, 2006). The decline in leaf hydraulic conductance (K_leaf) with dehydration may thus depend on both components. The importance of the xylem component is well established. Vein xylem embolism and cell collapse have been observed in dehydrating leaves (Salleo et al., 2001; Cochard et al., 2004a; Johnson et al., 2009), and computer modeling and experimental work showed that species with high major vein length per leaf area (VLA; i.e. for the first three vein-branching orders) were more resistant to hydraulic decline, providing more pathways around embolisms (Scoffoni et al., 2011). However, the physical impacts of dehydration on the extraxylem pathways have not been studied, even though in turgid leaves these pathways account for 26% to 88% of leaf hydraulic resistance (i.e. of 1/K_leaf), depending on species (Sack et al., 2003a; Cochard et al., 2004b). The aim of this study was to determine whether leaf shrinkage during dehydration relates to the decline of K_leaf as well as the structural determinants of leaf shrinkage.The shrinkage of leaves with dehydration has drawn attention for over 100 years. Leaves shrink in their area (Bogue, 1892; Gardner and Ehlig, 1965; Jones, 1973; Tang and Boyer, 2007; Blonder et al., 2012) and, considered in relative terms, even more strongly in their thickness (Fig. 1; Meidner, 1952; Gardner and Ehlig, 1965; Downey and Miller, 1971; Syvertsen and Levy, 1982; Saini and Rathore, 1983; Burquez, 1987; McBurney, 1992; Sancho-Knapik et al., 2010, 2011). Leaves fluctuate in thickness daily and seasonally according to transpiration (Kadoya et al., 1975; Tyree and Cameron, 1977; Fensom and Donald, 1982; Rozema et al., 1987; Ogaya and Peñuelas, 2006; Seelig et al., 2012). Indeed, the relation of leaf thickness to water status is so tight that using leaf thickness to guide irrigation has led to water savings of up to 45% (Seelig et al., 2012).Open in a separate windowFigure 1.Sketches of a fully turgid leaf (A) versus a strongly dehydrated leaf (B; drawings based on leaf cross sections of sunflower in Fellows and Boyer, 1978). Note the strong reduction in leaf thickness, cell thickness, and intercellular airspaces in the dehydrated leaf. Epidermal cells are shrunk in the dehydrated leaf, inducing whole-leaf area shrinkage. Note that this sketch represents shrinkage for a typical drought-sensitive species. Many species such as oaks (Quercus spp.) will experience less thickness shrinkage and an increase in intercellular airspace (see “Discussion”). [See online article for color version of this figure.]Previous studies of leaf shrinkage with progressive dehydration have tended to focus on single or few species. These studies showed that thickness declines with water status in two phases. Before the bulk leaf turgor loss point (TLP; leaf water potential [Ψ_leaf] at TLP) is reached, the slope of leaf thickness versus Ψ_leaf or relative water content (RWC) is shallower than past TLP for most species (Meidner, 1955, Kennedy and Booth, 1958, Burquez, 1987, McBurney, 1992, Sancho-Knapik et al., 2010, 2011). This is because before TLP, declining Ψ_leaf is strongly driven by declines in turgor pressure, which have a relatively low impact on cell and airspace volume, whereas past the TLP, declining Ψ_leaf depends only on solute concentration, which increases in inverse proportion as cell water volume declines while airspaces may shrink or expand (Tyree and Hammel, 1972, Sancho-Knapik et al., 2011). However, the steepness of the slope of leaf thickness versus Ψ_leaf before TLP seems to vary strongly across species (Meidner, 1955; Kennedy and Booth, 1958; Fellows and Boyer, 1978; Burquez, 1987; Colpitts and Coleman, 1997; Sancho-Knapik et al., 2010).A high leaf cell volume and turgor is crucial to physiological processes (Boyer, 1968; Lawlor and Cornic, 2002). Shrinkage may affect cell connectivity and water transport (Sancho-Knapik et al., 2011). However, no studies have tested for a possible relationship of leaf shrinkage with the decline of K_leaf during dehydration. Such an association would arise if, across species, shrinkage occurred simultaneously with vein xylem embolism or if tissue shrinkage led to declines in the extraxylem hydraulic conductance.To refine our hypotheses, we modified a computer model of the leaf hydraulic system (Cochard et al., 2004b; McKown et al., 2010; Scoffoni et al., 2011) to predict the impact of losses of xylem and extraxylem conductance on the response of K_leaf to dehydration. We characterized the degree of leaf shrinkage in thickness, in the thickness of cells and airspaces within the leaf, and in leaf area for 14 species diverse in phylogeny, leaf traits, and drought tolerance. We hypothesized that loss of extraxylem hydraulic conductance should have a greater impact on K_leaf at less negative water potentials when xylem tensions are too weak to trigger embolism and induce dramatic declines in K_leaf. We hypothesized that species with greater degrees of shrinkage before TLP would experience greater loss of K_leaf. Furthermore, we hypothesized that species from moist habitats would have greater degrees of shrinkage.For insight into the mechanisms and consequences of leaf shrinkage, we also investigated the relationships of 18 indices of leaf shrinkage with a wide range of aspects of leaf structure and composition, including gross morphology, leaf venation architecture, parameters of pressure-volume curves, and leaf water storage. We hypothesized that, across species, shrinkage in whole leaf, cell, and intercellular airspace thickness would be lower for species with greater allocation to structural rigidity and osmotic concentration, and thus shrinkage would be positively correlated with a lower modulus of elasticity (ε), less negative osmotic pressure at full turgor (π_o), lower leaf mass per area (LMA), and lower leaf density. Additionally, we tested the longstanding hypothesis that species with higher major VLA and/or minor VLA (i.e. the fourth and higher vein-branching orders) would shrink less in area and/or thickness with dehydration (Gardner and Ehlig, 1965). Finally, we tested the ability of dehydrated leaves to recover in size with rehydration. We hypothesized that recovery would be greater for mildly than for strongly dehydrated leaves and that species with greater leaf shrinkage would be better able to recover from shrinkage.     

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

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

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

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

A New LxxxA Motif in the Transmembrane Helix3 of Maize Aquaporins Belonging to the Plasma Membrane Intrinsic Protein PIP2 Group Is Required for Their Trafficking to the Plasma Membrane

Aquaporins play important roles in maintaining plant water status under challenging environments. The regulation of aquaporin density in cell membranes is essential to control transcellular water flows. This work focuses on the maize (Zea mays) plasma membrane intrinsic protein (ZmPIP) aquaporin subfamily, which is divided into two sequence-related groups (ZmPIP1s and ZmPIP2s). When expressed alone in mesophyll protoplasts, ZmPIP2s are efficiently targeted to the plasma membrane, whereas ZmPIP1s are retained in the endoplasmic reticulum (ER). A protein domain-swapping approach was utilized to demonstrate that the transmembrane domain3 (TM3), together with the previously identified N-terminal ER export diacidic motif, account for the differential localization of these proteins. In addition to protoplasts, leaf epidermal cells transiently transformed by biolistic particle delivery were used to confirm and refine these results. By generating artificial proteins consisting of a single transmembrane domain, we demonstrated that the TM3 of ZmPIP1;2 or ZmPIP2;5 discriminates between ER and plasma membrane localization, respectively. More specifically, a new LxxxA motif in the TM3 of ZmPIP2;5, which is highly conserved in plant PIP2s, was shown to regulate its anterograde routing along the secretory pathway, particularly its export from the ER.Aquaporins are of major importance to plant physiology, being essential for the regulation of transcellular water movement during growth and development (Maurel et al., 2008; Gomes et al., 2009; Heinen et al., 2009; Prado and Maurel, 2013; Chaumont and Tyerman, 2014). Aquaporins are small membrane proteins consisting of six transmembrane (TM) domains connected by five loops (A–E), and N and C termini facing the cytosol (Fig. 1A). They assemble as homotetramers and/or heterotetramers in the membrane, with each monomer acting as an independent water channel (Murata et al., 2000; Fetter et al., 2004; Gomes et al., 2009). Aquaporins form a highly divergent protein family in plants (Chaumont et al., 2001; Johanson et al., 2001), and this work focuses on the maize (Zea mays) plasma membrane intrinsic protein (ZmPIP) family (Chaumont et al., 2001). The regulation of the subcellular localization of these proteins is a key process controlling their density in the plasma membrane (PM) and, hence, their physiological roles (Hachez et al., 2013).Open in a separate windowFigure 1.Swapping TM3 of ZmPIP2;5 with that of ZmPIP1;2 retains the protein in intracellular structures. A, Cartoons representing the chimeric proteins composed of ZmPIP2;5, in which each TM has been replaced by the corresponding TM from ZmPIP1;2. All proteins are drawn with the cytosolic domains facing down. ZmPIP2;5 and ZmPIP1;2 portions are shown in black and white, respectively. All chimeras were fused to the C terminus of mYFP, which is not displayed for clarity purposes. B, Confocal microscopy images of maize mesophyll protoplasts transiently coexpressing mYFP-tagged ZmPIP2;5-PIP1;2 TM chimeric proteins (green) and the ER marker mCFP:HDEL (cyan). FM4-64 was added as a PM marker (red). Arrowheads in image 13 indicate accumulation of the protein in punctate structures that are not labeled by mCFP:HDEL. The localization patterns of the proteins of interest are representative of a total of at least 22 cells from three independent experiments. C, Confocal microscopy images of a maize mesophyll protoplast transiently expressing mYFP:ZmPIP2;5-TM3_PIP1;2 (green) and ST:mCFP (magenta). Arrowheads indicate colocalization in Golgi stacks. The images are representative of a total of 17 cells from two independent experiments. Bar = 5 µm.PIP aquaporins cluster in two groups (PIP1s and PIP2s), which are highly conserved across species (Kammerloher et al., 1994; Chaumont et al., 2000, 2001; Johanson et al., 2001; Anderberg et al., 2012). We previously showed that the maize PIP1 and PIP2 isoforms exhibit different water channel activities when expressed in Xenopus laevis oocytes, with only PIP2s increasing the membrane water permeability coefficient (P_f; Chaumont et al., 2000). However, when ZmPIP1 and ZmPIP2 are coexpressed, the isoforms physically interact to modify their stability and trafficking to the oocyte membrane, and synergistically increase the oocyte P_f (Fetter et al., 2004). Similar synergistic interactions between PIP1s and PIP2s have been reported in numerous plant species (Temmei et al., 2005; Mahdieh et al., 2008; Vandeleur et al., 2009; Bellati et al., 2010; Ayadi et al., 2011; Horie et al., 2011; Yaneff et al., 2014).PIPs were originally thought to be exclusively localized in the PM and were named accordingly (Kammerloher et al., 1994). However, recent experiments have shown that not all PIPs are located to the PM under all conditions, and that regulation of PIP subcellular localization is a highly dynamic process involving protein interactions (Boursiac et al., 2005, 2008; Zelazny et al., 2007, 2009; Uehlein et al., 2008; Besserer et al., 2012; Luu et al., 2012). When expressed singly in maize leaf mesophyll protoplasts, fluorescently tagged ZmPIP1s and ZmPIP2s differ in their subcellular localization. ZmPIP1s are retained in the endoplasmic reticulum (ER), whereas ZmPIP2s are targeted to the PM (Zelazny et al., 2007). However, upon coexpression, ZmPIP1s are relocalized from the ER to the PM, where they perfectly colocalize with ZmPIP2s. This relocalization results from their physical interaction as demonstrated by Förster resonance energy transfer/fluorescence lifetime imaging microscopy and immunoprecipitation experiments (Zelazny et al., 2007). These results indicate that ZmPIP2s, but not ZmPIP1s, possess signals that allow them to be delivered to the PM, and that hetero-oligomerization is required for ZmPIP1 trafficking to the PM. Interestingly, a diacidic motif (DxE, Asp-any amino acid-Glu) located in the N terminus of ZmPIP2;4, ZmPIP2;5, and Arabidopsis (Arabidopsis thaliana) AtPIP2;1 was shown to be required to exit the ER (Zelazny et al., 2009; Sorieul et al., 2011). Diacidic motifs interact with Secretory protein24, which is thought to be the main cargo-selection protein of the Coat proteinII complex that mediates vesicle formation at ER export sites (Miller et al., 2003). However, not all PM-localized PIP2s contain a diacidic ER export signal (Zelazny et al., 2009). In addition, swapping the N-terminal region of ER-retained ZmPIP1;2 with that of PM-localized ZmPIP2;5, which contains the functional diacidic motif, is not sufficient to trigger ER export of the protein (Zelazny et al., 2009). This result suggests that other export signals might be present in PIP2s and/or ER retention signals might be present in PIP1s elsewhere than in the N terminus.To identify new signals regulating ZmPIP1 and ZmPIP2 protein trafficking along the secretory pathway, we used a protein domain swapping-based approach and identified the TM3 as an important region that discriminates between ER-retained ZmPIP1;2 and PM-localized ZmPIP2;5. Specific mutations in the TM3 region of ZmPIP2;5 allowed the identification of a new ZmPIP2-conserved LxxxA motif, which regulates its export from the ER.     

New Arabidopsis Advanced Intercross Recombinant Inbred Lines Reveal Female Control of Nonrandom Mating

Female control of nonrandom mating has never been genetically established, despite being linked to inbreeding depression and sexual selection. In order to map the loci that control female-mediated nonrandom mating, we constructed a new advanced intercross recombinant inbred line (RIL) population derived from a cross between Arabidopsis (Arabidopsis thaliana) accessions Vancouver (Van-0) and Columbia (Col-0) and mapped quantitative trait loci (QTLs) responsible for nonrandom mating and seed yield traits. We genotyped a population of 490 RILs. A subset of these lines was used to construct an expanded map of 1,061.4 centimorgans with an average interval of 6.7 ± 5.3 centimorgans between markers. QTLs were then mapped for female- and male-mediated nonrandom mating and seed yield traits. To map the genetic loci responsible for female-mediated nonrandom mating and seed yield, we performed mixed pollinations with genetically marked Col-0 pollen and Van-0 pollen on RIL pistils. To map the loci responsible for male-mediated nonrandom mating and seed yield, we performed mixed pollinations with genetically marked Col-0 and RIL pollen on Van-0 pistils. Composite interval mapping of these data identified four QTLs that control female-mediated nonrandom mating and five QTLs that control female-mediated seed yield. We also identified four QTLs that control male-mediated nonrandom mating and three QTLs that control male-mediated seed yield. Epistasis analysis indicates that several of these loci interact. To our knowledge, the results of these experiments represent the first time female-mediated nonrandom mating has been genetically defined.The process of pollination offers plants the opportunity to selectively breed. For example, in pollinations that include more than one pollen population, pollen often show differential siring ability. This process is called nonrandom mating. Although pollen may fail in pollinations because they are self pollen in an obligate outcrossing plant or pollen from a different species, we focus our studies on differential siring ability of compatible, conspecific mates (Hogenboom, 1973, 1975; Williams et al., 1999; de Nettancourt, 2001; Husband et al., 2002; Wheeler et al., 2009; Meng et al., 2011; Nasrallah, 2011). Nonrandom mating at this level has received intense interest for its potential to avoid inbreeding depression and its potential to be the result of sexual selection (Charnov, 1979; Mulcahy, 1979; Willson, 1979; Queller, 1983; Stephenson and Bertin, 1983; Willson and Burley, 1983; Marshall and Ellstrand, 1986; Charlesworth and Charlesworth, 1987; Mulcahy and Mulcahy, 1987; Cruzan, 1990; Quesada et al., 1993; Snow, 1994; Paschke et al., 2002; Skogsmyr and Lankinen, 2002; Stephenson et al., 2003; Armbruster and Rogers, 2004; Bernasconi et al., 2004; Lankinen and Armbruster, 2007). Despite a long history of theoretical and experimental attention, very little is known about the underlying genetics that govern the process (Carlson et al., 2011).One challenge in understanding the genetics of nonrandom mating lies in its complexity, potentially involving multiple distinct pathways specific to either female or male tissues. Physiologically, postpollination nonrandom mating may be a result of intrinsic differences in pollen competitive abilities (male-mediated nonrandom mating). A number of experimental strategies have been employed to demonstrate male-mediated control of nonrandom mating. For example, experiments in radish (Raphanus sativus) found that some pollen sire more seeds than others in mixed pollinations across a range of maternal plants, demonstrating consistency of male function (Marshall and Ellstrand, 1986, 1988; Mitchell and Marshall, 1998). More direct measures of male function, such as in vitro and in vivo pollen tube growth rates, verify variation in male function and demonstrable impact on nonrandom mating (Snow and Spira, 1991a, 1991b; Pasonen et al., 1999; Skogsmyr and Lankinen, 1999; Stephenson et al., 2001; Lankinen and Skogsmyr, 2002; Lankinen et al., 2009). Finally, recent work in our laboratory has directly mapped the genetic loci responsible for the control of male-mediated nonrandom mating in Arabidopsis (Arabidopsis thaliana; Carlson et al., 2011).Alternatively, or concurrently, nonrandom mating can be the result of differential interaction between the female tissue and competing pollen populations or seeds (female-mediated nonrandom mating). Establishing the female role in nonrandom mating has been more challenging, as most study designs involve the deposition of pollen from multiple donors and thus include the confounding variable of pollen competition. Despite this challenge, a number of experimental strategies have been devised to explore the role of the female in nonrandom mating. For example, a number of studies demonstrate that maternal identity influences nonrandom mating patterns (Marshall and Ellstrand, 1986, 1988; Snow and Mazer, 1988; Johnston, 1993; Marshall et al., 2000; Carlson et al., 2009, 2013). Studies have also established that manipulation of watering or nutrient regimes of maternal plants changes the patterns and magnitude of nonrandom mating (Marshall and Diggle, 2001; Shaner and Marshall, 2003; Haileselassie et al., 2005; Marshall et al., 2007). These studies and others implicate the identity and condition of the female in the process of nonrandom mating. Despite a long history of research, genetic control of female-mediated nonrandom mating has never been demonstrated, and the identity of the genes involved remains unexplored.In previous work, we developed a system in Arabidopsis to assay nonrandom mating and showed its utility for genetically mapping the loci responsible (Carlson et al., 2009, 2011). Pursuing the genetics of nonrandom mating in a largely selfing plant such as Arabidopsis provides both theoretical and practical advantages. First, outcrossing plants carry higher levels of heterozygosity that produce pollen populations that display different phenotypes because of segregating alleles. This complicates genetic analysis. Also, in outcrossing plants that carry genetic load, reproductive success is context dependent. Pollinations with self pollen or pollen from genetically similar plants often lead to poor reproductive outcomes. For example, in mixed pollinations in generally outcrossing self-compatible plants that include self pollen, self pollen often sire a disproportionally low number of seeds (Bateman, 1956; Weller and Ornduff, 1977; Bowman, 1987; Eckert and Barrett, 1994; Jones, 1994; Hauser and Siegismund, 2000; Teixeira et al., 2009), but other findings have been reported (Sork and Schemske, 1992; Johnston, 1993). Thus, in outcrossing plants, gene variants that influence reproductive success, parental relatedness, and segregating heterozygosity all influence reproductive outcomes. Two of these factors are essentially eliminated by studying plant populations that have historically selfed. As outcrossing populations become increasingly self-fertilizing, they both lose heterozygosity, and their genetic load is purged (Lande and Schemske, 1985; Schemske and Lande, 1985; Charlesworth and Charlesworth, 1987; Lande et al., 1994; Byers and Waller, 1999; Crnokrak and Barrett, 2002). This is the case for Arabidopsis, whose tested populations show relatively low levels of heterozygosity and little evidence for the early-acting inbreeding depression that is indicative of genetic load (Bakker et al., 2006; Bomblies et al., 2010; Platt et al., 2010; Carlson et al., 2013). Thus, this system provides an excellent opportunity to identify and explore the genetic variation in differential reproduction that develops or persists in plant populations unrelated to inbreeding depression.Using this system, we previously identified potential female control of nonrandom mating in mixed pollinations between Vancouver (Van-0) and Columbia (Col-0) accessions of Arabidopsis (Carlson et al., 2009). When Van-0 and genetically marked Col-0 (Col-NPTII) pollen compete on Col-0 pistils, Col-NPTII pollen sire 43% of the progeny, while Van-0 pollen sire 57%. When these pollen compete on Van-0 pistils, Col-NPTII pollen sire 67.5% of the progeny, while Van-0 pollen sire 32.5%. This system offers us, to our knowledge for the first time, the opportunity to genetically define female-mediated nonrandom mating and map the loci responsible.In order to genetically map female control of nonrandom mating, we constructed a new advanced intercross recombinant inbred line (RIL) mapping population derived from a cross between Van-0 and Col-0 accessions of Arabidopsis. RILs are powerful tools that allow high-resolution genetic mapping of loci that direct complex traits. Each RIL contains chromosomes that are defined homozygous patchworks of parental DNA, in this case Van-0 and Col-0. By phenotyping these lines, we can statistically associate nonrandom mating and seed yield phenotypes with chromosomal regions. We chose these two accessions because (1) our previous experiments predict clear female control of nonrandom mating and (2) we have previously mapped male-mediated nonrandom mating controls using a Col-4/Landsberg mapping population (a population that does not display female control of nonrandom mating; Carlson et al., 2011). Thus, this new population provides us the opportunity to map loci that control female nonrandom mating and investigate the degree of conservation of loci that affect male-mediated nonrandom mating. We use this new mapping population to perform quantitative trait locus (QTL) mapping and identify multiple loci that direct both female- and male-mediated control of nonrandom mating and seed yield traits.     

Freeze-Thaw Stress: Effects of Temperature on Hydraulic Conductivity and Ultrasonic Activity in Ten Woody Angiosperms

Freeze-thaw events can affect plant hydraulics by inducing embolism. This study analyzed the effect of temperature during the freezing process on hydraulic conductivity and ultrasonic emissions (UE). Stems of 10 angiosperms were dehydrated to a water potential at 12% percentage loss of hydraulic conductivity (PLC) and exposed to freeze-thaw cycles. The minimal temperature of the frost cycle correlated positively with induced PLC, whereby species with wider conduits (hydraulic diameter) showed higher freeze-thaw-induced PLC. Ultrasonic activity started with the onset of freezing and increased with decreasing subzero temperatures, whereas no UE were recorded during thawing. The temperature at which 50% of UE were reached varied between −9.1°C and −31.0°C across species. These findings indicate that temperatures during freezing are of relevance for bubble formation and air seeding. We suggest that species-specific cavitation thresholds are reached during freezing due to the temperature-dependent decrease of water potential in the ice, while bubble expansion and the resulting PLC occur during thawing. UE analysis can be used to monitor the cavitation process and estimate freeze-thaw-induced PLC.Xylem embolism is a limiting factor for plant survival and distribution (Choat et al., 2012; Charrier et al., 2013). Two major factors can induce embolism in the xylem of plants: drought and freeze stress. Freeze-thaw-induced embolism is caused by bubbles formed during freezing that then expand on thawing (Lemoine et al., 1999; Hacke and Sperry, 2001; Cruiziat et al., 2002; Tyree and Zimmermann, 2002). As wider conduits contain more gas and form larger bubbles, which expand at less negative tension, conduit diameter and xylem sap tension are critical for the formation of freeze-thaw-induced embolism (Davis et al., 1999; Pittermann and Sperry, 2003). Accordingly, Mayr and Sperry (2010) observed a loss of conductivity only when samples were under critical tension during thawing. Under drought stress, tension in the xylem sap increases the sensitivity to embolism generated by successive freeze-thaw cycles (Mayr et al., 2003, 2007).Ultrasonic emissions (UE) analysis can be used to detect cavitation events in wood. It is unclear how well related UE are to cavitation events, as they are extracted from continuous acoustic emissions and depend on set definitions. However, UE analysis has been proven effective for monitoring drought-induced embolism in the laboratory (Pena and Grace, 1986; Salleo and Lo Gullo, 1986; Borghetti et al., 1993; Salleo et al., 2000) as well as in field experiments (Ikeda and Ohtsu, 1992; Jackson et al., 1995; Jackson and Grace, 1996; Hölttä et al., 2005; Ogaya and Penuelas, 2007). In a cavitating conduit, signals are probably produced by the disruption of the water column and subsequent tension relaxation of cell walls.UE have also been detected during freezing events, but the origin of these signals was less clear. In some cases, UE were observed during thawing, which are thus probably related to embolism formation according to the classic thaw-expansion hypothesis (Mayr and Sperry, 2010); however, all species studied have produced UE on freezing, which cannot yet be explained (Raschi et al., 1989; Kikuta and Richter, 2003; Mayr et al., 2007; Mayr and Sperry, 2010; Mayr and Zublasing, 2010). The low solubility of gases in ice prompted the idea that air bubbles expulsed from the ice structure produce UE near the ice-liquid interface (Sevanto et al., 2012). As the water potential of ice is strongly temperature dependent, the minimum temperature during freezing might be a relevant factor. Numerous studies have analyzed UE patterns during freeze-thaw cycles in conifers (Mayr et al., 2007; Mayr and Sperry, 2010; Mayr and Zublasing, 2010) or angiosperms (Weiser and Wallner, 1988; Kikuta and Richter, 2003), but few of them measured embolism concomitantly. Percentage loss of hydraulic conductivity (PLC) was only measured in a few studies and only in conifers (Mayr et al., 2007; Mayr and Sperry, 2010).In this study, we analyzed the effect of freeze-thaw cycles on the hydraulic conductivity and ultrasonic activity in 10 angiosperm species. We hypothesized that (1) the extent of freeze-thaw-induced embolism depends on xylem anatomy (related to conduit diameter) and minimal temperature (related to the water potential of ice); (2) ultrasonic activity is also influenced by anatomy and temperature; and (3) PLC and UE are positively correlated. PLC was measured in 10 angiosperm species after freeze-thaw cycles at different minimal temperatures (−10 to −40°C). Furthermore, UE were recorded during a freeze-thaw cycle down to −40°C.     

The Genetics of Leaf Flecking in Maize and Its Relationship to Plant Defense and Disease Resistance

Physiological leaf spotting, or flecking, is a mild-lesion phenotype observed on the leaves of several commonly used maize (Zea mays) inbred lines and has been anecdotally linked to enhanced broad-spectrum disease resistance. Flecking was assessed in the maize nested association mapping (NAM) population, comprising 4,998 recombinant inbred lines from 25 biparental families, and in an association population, comprising 279 diverse maize inbreds. Joint family linkage analysis was conducted with 7,386 markers in the NAM population. Genome-wide association tests were performed with 26.5 million single-nucleotide polymorphisms (SNPs) in the NAM population and with 246,497 SNPs in the association population, resulting in the identification of 18 and three loci associated with variation in flecking, respectively. Many of the candidate genes colocalizing with associated SNPs are similar to genes that function in plant defense response via cell wall modification, salicylic acid- and jasmonic acid-dependent pathways, redox homeostasis, stress response, and vesicle trafficking/remodeling. Significant positive correlations were found between increased flecking, stronger defense response, increased disease resistance, and increased pest resistance. A nonlinear relationship with total kernel weight also was observed whereby lines with relatively high levels of flecking had, on average, lower total kernel weight. We present evidence suggesting that mild flecking could be used as a selection criterion for breeding programs trying to incorporate broad-spectrum disease resistance.The plant hypersensitive response (HR) is a form of programmed cell death (PCD) characterized by rapid, localized cell death at the point of attempted pathogen penetration, usually resulting in disease resistance (Coll et al., 2011). It is often associated with other responses, including ion fluxes, an oxidative burst, lipid peroxidation, and cell wall fortification (Hammond-Kosack and Jones, 1996). van Doorn et al. (2011) suggested that HR is a type of PCD sharing features with, but distinct from, both vacuolar cell death and necrosis.HR has been associated with resistance to almost every class of pathogen and pest, including bacteria, viruses, fungi, nematodes, insects, and parasitic plants (Wu and Baldwin, 2010), and generally is most effective against biotrophic pathogens, since biotrophs require a long-term feeding relationship with living host cells. It is generally mediated by dominant resistance (R) genes whose activation is triggered by the direct or indirect detection of specific pathogen-derived effector proteins (Bent and Mackey, 2007). R proteins are maintained in their inactive state if their corresponding effector is not present. Mutants in which HR is constitutively active have been identified in many plant species, including maize/corn (Zea mays; Walbot et al., 1983; Johal, 2007), Arabidopsis (Arabidopsis thaliana; Lorrain et al., 2003), barley (Hordeum vulgare; Wolter et al., 1993), and rice (Oryza sativa; Yin et al., 2000).One well-known class of plant mutants spontaneously form lesions (patches of dead or chlorotic cells) in the absence of any obvious injury, stress, or infection to the plant. Since these lesions in some cases resemble HR, they have been termed disease-lesion mimics (Neuffer and Calvert, 1975). These mutants, which we will here collectively term Les mutants, have been studied extensively, especially in maize (Walbot et al., 1983; Johal et al., 1995; Johal, 2007) and Arabidopsis (Coll et al., 2011). While some of these lesion phenotypes are indeed caused by perturbations in the plant defense response (Hu et al., 1996; Rustérucci et al., 2001), some of the genes underlying this mutant class affect various other pathways that cause cell death if their function is perturbed (Johal, 2007). For instance, the Arabidopsis gene acd2 and the maize gene lls1 are defective in chlorophyll degradation (Gray et al., 1997; Mach et al., 2001).We have defined leaf flecking as the mild, genetically determined spotting observed on many maize inbred cultivars (Vontimitta et al., 2015; Fig. 1). The trait is qualitatively and visually similar to, but quantitatively less severe than, Les mutant phenotypes. The distinction between what constitutes a flecking versus a mild Les trait is necessarily somewhat arbitrary, but for our purposes, we have defined any nonproliferating and distinct leaf-spotting phenotype as flecking.Open in a separate windowFigure 1.A, Examples of variation in the flecking phenotype among inbred lines, with severity increasing from left to right (flecking scores in parentheses, from 0 to 4, scored on a scale of 1–10). B, Leaves of the lines nearly isogenic to inbred Mo20W, into which specific indicated dominant Les mutant genes have been introgressed (Rp1-D21 mutation in an H95 inbred background). Photographs were taken in Clayton, North Carolina, 12 weeks after planting. This figure is adapted from Figure 1 of Vontimitta et al. (2015).Leaf flecking is familiar to most corn breeders, appearing in such well-known and widely used lines such as Mo17 (Zehr et al., 1994) and in several other species such as barley (Makepeace et al., 2007), wheat (Triticum aestivum; Nair and Tomar, 2001), and oat (Avena sativa; Ferdinandsen and Winge, 1930). Flecking tends to be more noticeable in inbreds compared with their derived hybrids (M. Goodman and W. Dolezal, personal communication). Anecdotally, it is often thought to be indicative of a constitutive low-level defense response and as a marker for increased disease resistance.In previous work, we and others have defined the genetic architectures associated with resistance to several maize diseases, including southern leaf blight (SLB; causal agent, Cochliobolus heterostrophus), northern leaf blight (NLB; causal agent, Exserohilum turcicum), and gray leaf spot (GLS; causal agent, Cercospora zeae-maydis; Kump et al., 2011; Poland et al., 2011; Wisser et al., 2011; Benson et al., 2015), and with the control of the maize HR (Chintamanani et al., 2010; Chaikam et al., 2011; Olukolu et al., 2013). For much of this work, we used two powerful mapping populations: the maize association population (Flint-Garcia et al., 2005), a collection of 302 diverse inbred lines with low linkage disequilibrium, and the 5,000-line nested association mapping (NAM) population (McMullen et al., 2009), which is made up of 25 200-line recombinant inbred line (RIL) subpopulations derived from crosses between the common parent B73 and 25 diverse inbreds. Using these populations, it is possible to both sample a diverse array of germplasm and map quantitative trait loci (QTLs) precisely, in some cases to the gene level (Tian et al., 2011; Cook et al., 2012; Hung et al., 2012; Larsson et al., 2013; Olukolu et al., 2013; Wang and Balint-Kurti, 2016).A recent study using 300 lines from the maize intermated B73 × Mo17 population advanced intercross line mapping population identified low but moderately significant positive correlations between increased flecking and increased disease resistance and defense response (Vontimitta et al., 2015). Loci associated with variation in flecking were mapped, although these loci did not colocalize with QTLs identified previously for disease resistance and defense response traits (Balint-Kurti et al., 2007, 2008, 2010; Olukolu et al., 2013). In this study, we have extended this work to examine the genetic basis of leaf flecking over a much more diverse set of maize germplasm using a substantially larger population. We mapped loci associated with variation in leaf flecking and identified candidate genes and pathways that may be involved in this phenotype. Additionally, we have examined the correlations between leaf flecking and disease resistance, the hypersensitive defense response, and total kernel weight.