Phene Synergism between Root Hair Length and Basal Root Growth Angle for Phosphorus Acquisition

Shallow basal root growth angle (BRGA) increases phosphorus acquisition efficiency by enhancing topsoil foraging because in most soils, phosphorus is concentrated in the topsoil. Root hair length and density (RHL/D) increase phosphorus acquisition by expanding the soil volume subject to phosphorus depletion through diffusion. We hypothesized that shallow BRGA and large RHL/D are synergetic for phosphorus acquisition, meaning that their combined effect is greater than the sum of their individual effects. To evaluate this hypothesis, phosphorus acquisition in the field in Mozambique was compared among recombinant inbred lines of common bean (Phaseolus vulgaris) having four distinct root phenotypes: long root hairs and shallow basal roots, long root hairs and deep basal roots, short root hairs and shallow basal roots, and short root hairs and deep basal roots. The results revealed substantial synergism between BRGA and RHL/D. Compared with short-haired, deep-rooted phenotypes, long root hairs increased shoot biomass under phosphorus stress by 89%, while shallow roots increased shoot biomass by 58%. Genotypes with both long root hairs and shallow roots had 298% greater biomass accumulation than short-haired, deep-rooted phenotypes. Therefore, the utility of shallow basal roots and long root hairs for phosphorus acquisition in combination is twice as large as their additive effects. We conclude that the anatomical phene of long, dense root hairs and the architectural phene of shallower basal root growth are synergetic for phosphorus acquisition. Phene synergism may be common in plant biology and can have substantial importance for plant fitness, as shown here.Suboptimal phosphorus availability is a primary limitation to plant growth in terrestrial ecosystems (Vance et al., 2003). Large areas of tropical and subtropical soils in Africa, Latin America, and Asia have phosphorus availability limited by low total phosphorus content as well as high phosphorus fixation (Sanchez and Uehara, 1980). The use of phosphorus fertilizer to correct phosphorus deficiency is only a partial solution, since phosphorus fertilizers are costly, nonrenewable, potentially harmful to the environment, and often marginally effective in tropical soils because of immobilization by the soil (Cathcart, 1980). Therefore, the development of crop cultivars with enhanced ability to acquire phosphorus is an important strategy to increase agricultural productivity in low-input agroecosystems and to reduce input requirements in intensive agriculture (Vance et al., 2003; Gahoonia and Nielsen, 2004; Lambers et al., 2006; Lynch, 2007, 2011).Several root phenes (i.e. basic units of the phenotype; Serebrovsky, 1925; Lynch, 2011; for discussion, see York et al., 2013) enhance phosphorus acquisition, including root architectural phenes for topsoil foraging (Lynch and Brown, 2001), such as shallow root growth angles (Liao et al., 2004; Ho et al., 2005), increased basal root whorl number (Lynch and Brown, 2012; Miguel et al., 2013), and adventitious rooting (Miller et al., 2003); phenes to enhance soil exploitation, including root hair length and density (RHL/D; Bates and Lynch, 2000a, 2000b, 2001; Ma et al., 2001a; Gahoonia and Nielsen, 2004; Yan et al., 2004) and phosphorus-solubilizing root exudates (Ryan et al., 2001); mycorrhizal symbioses (Smith and Read, 2008); and phenes that reduce the metabolic cost of soil exploration (Lynch and Ho, 2005), such as root etiolation and root cortical aerenchyma (Fan et al., 2003; Postma and Lynch, 2010, 2011). It is probable that interactions among these phenes are important in determining the phosphorus acquisition of integrated phenotypes. Results from the structural-functional model SimRoot indicate that RHL/D, the distance from the root tip to the first appearance of root hairs, and the pattern of root hair-bearing epidermal cells (trichoblasts) among non-hair-bearing cells (atrichoblasts) are synergetic for phosphorus acquisition in Arabidopsis (Arabidopsis thaliana; Ma et al., 2001b). Another SimRoot study showed that on low-phosphorus soils, the utility of root cortical aerenchyma in maize (Zea mays) may be 2.9 times greater in plants with increased lateral branching density than in plants with normal branching (Postma and Lynch, 2011). Morphological, anatomical, symbiotic, and biochemical phenes expressed by root axes should have significant synergies with architectural phenes, since architectural phenes determine the position of root axes in time and space and, therefore, the soil domain in which spatially localized phenes are expressed (Lynch, 2011).Phosphorus availability is greater in the topsoil, with a steep decline with depth. Therefore, root architectural phenes that increase topsoil foraging can improve phosphorus acquisition (Lynch and Brown, 2001). Root shallowness regulated by basal root growth angle (BRGA) has been demonstrated to be of particular importance for topsoil foraging (Bonser et al., 1996; Liao et al., 2001; Rubio et al., 2001; Ho et al., 2005). These studies show that common bean (Phaseolus vulgaris) genotypes with smaller BRGA (i.e. shallower roots) have better performance in low-phosphorus soils. Shallow root distribution is also important for phosphorus acquisition in maize (Zhu et al., 2005).RHL/D are also important for phosphorus acquisition (Bates and Lynch, 2000a, 2000b, 2001; Gahoonia and Nielsen, 2004). Since phosphorus mobility in soil is governed by diffusion rather than mass flow, phosphorus uptake by roots is limited by localized phosphorus depletion in the rhizosphere (Barber, 1995). Long root hairs extend the phosphorus depletion zone surrounding the root, thereby increasing the total amount of phosphorus accessible by the roots and phosphorus acquisition. In many plant species, the length and density of root hairs increase in response to low phosphorus availability (Bates and Lynch, 1996; Ma et al., 2001a). Increased RHL/D increases phosphorus accumulation in Arabidopsis growing in low-phosphorus conditions (Bates and Lynch, 2000a, 2000b), and mutants lacking root hairs have reduced phosphorus acquisition (Bates and Lynch, 2000b; Gahoonia et al., 2001). Species that develop more and/or longer root hairs (e.g. Lolium perenne) are more efficient in accessing inorganic phosphorus from soils and thus show greater growth response to phosphorus fertilization than species that lack these traits (e.g. Podocarpus totara). Genotypic variation for root hairs is associated with increased phosphorus acquisition in several species, including barley (Hordeum vulgare; Gahoonia and Nielsen, 2004), common bean (Miguel, 2004; Yan et al., 2004), and maize (Zhu et al., 2010).We hypothesize that the utilities of BRGA and RHL/D for phosphorus acquisition are synergetic. Root hairs will be more valuable for phosphorus acquisition if located in surface soil horizons by arising from roots with a shallow growth angle; shallow roots will have greater benefit for phosphorus acquisition if they have long and dense hairs. Therefore, genotypes possessing long, dense root hairs on shallow roots should have greater phosphorus acquisition than genotypes with either long root hairs on deep roots or short root hairs on shallow roots. We expect the combined benefit of long root hairs and shallow root growth angles to exceed the sum of their individual effects, since they permit greater exploitation of soil strata with the greatest phosphorus availability.In this study, we evaluated the potential synergism between the architectural phene of BRGA and the morphological phene of RHL/D for phosphorus acquisition by comparison of contrasting phenotypes of common bean growing in a weathered tropical soil.     

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.     

Focus on Ethylene: Ethylene Biosynthesis Is Promoted by Very-Long-Chain Fatty Acids during Lysigenous Aerenchyma Formation in Rice Roots

In rice (Oryza sativa) roots, lysigenous aerenchyma, which is created by programmed cell death and lysis of cortical cells, is constitutively formed under aerobic conditions, and its formation is further induced under oxygen-deficient conditions. Ethylene is involved in the induction of aerenchyma formation. reduced culm number1 (rcn1) is a rice mutant in which the gene encoding the ATP-binding cassette transporter RCN1/OsABCG5 is defective. Here, we report that the induction of aerenchyma formation was reduced in roots of rcn1 grown in stagnant deoxygenated nutrient solution (i.e. under stagnant conditions, which mimic oxygen-deficient conditions in waterlogged soils). 1-Aminocyclopropane-1-carboxylic acid synthase (ACS) is a key enzyme in ethylene biosynthesis. Stagnant conditions hardly induced the expression of ACS1 in rcn1 roots, resulting in low ethylene production in the roots. Accumulation of saturated very-long-chain fatty acids (VLCFAs) of 24, 26, and 28 carbons was reduced in rcn1 roots. Exogenously supplied VLCFA (26 carbons) increased the expression level of ACS1 and induced aerenchyma formation in rcn1 roots. Moreover, in rice lines in which the gene encoding a fatty acid elongase, CUT1-LIKE (CUT1L; a homolog of the gene encoding Arabidopsis CUT1, which is required for cuticular wax production), was silenced, both ACS1 expression and aerenchyma formation were reduced. Interestingly, the expression of ACS1, CUT1L, and RCN1/OsABCG5 was induced predominantly in the outer part of roots under stagnant conditions. These results suggest that, in rice under oxygen-deficient conditions, VLCFAs increase ethylene production by promoting 1-aminocyclopropane-1-carboxylic acid biosynthesis in the outer part of roots, which, in turn, induces aerenchyma formation in the root cortex.Aerenchyma formation is a morphological adaptation of plants to complete submergence and waterlogging of the soil, and facilitates internal gas diffusion (Armstrong, 1979; Jackson and Armstrong, 1999; Colmer, 2003; Voesenek et al., 2006; Bailey-Serres and Voesenek, 2008; Licausi and Perata, 2009; Sauter, 2013; Voesenek and Bailey-Serres, 2015). To adapt to waterlogging in soil, rice (Oryza sativa) develops lysigenous aerenchyma in shoots (Matsukura et al., 2000; Colmer and Pedersen, 2008; Steffens et al., 2011) and roots (Jackson et al., 1985b; Justin and Armstrong, 1991; Kawai et al., 1998), which is formed by programmed cell death and subsequent lysis of some cortical cells (Jackson and Armstrong, 1999; Evans, 2004; Yamauchi et al., 2013). In rice roots, lysigenous aerenchyma is constitutively formed under aerobic conditions (Jackson et al., 1985b), and its formation is further induced under oxygen-deficient conditions (Colmer et al., 2006; Shiono et al., 2011). The former and latter are designated constitutive and inducible lysigenous aerenchyma formation, respectively (Colmer and Voesenek, 2009). The gaseous plant hormone ethylene regulates adaptive growth responses of plants to submergence (Voesenek and Blom, 1989; Voesenek et al., 1993; Visser et al., 1996a,b; Lorbiecke and Sauter, 1999; Hattori et al., 2009; Steffens and Sauter, 2009; van Veen et al., 2013). Ethylene also induces lysigenous aerenchyma formation in roots of some gramineous plants (Drew et al., 2000; Shiono et al., 2008). The treatment of roots with ethylene or its precursor (1-aminocyclopropane-1-carboxylic acid [ACC]) stimulates aerenchyma formation in rice (Justin and Armstrong, 1991; Colmer et al., 2006; Yukiyoshi and Karahara, 2014), maize (Zea mays; Drew et al., 1981; Jackson et al., 1985a; Takahashi et al., 2015), and wheat (Triticum aestivum; Yamauchi et al., 2014a,b). Moreover, treatment of roots with inhibitors of ethylene action or ethylene biosynthesis effectively blocks aerenchyma formation under hypoxic conditions in maize (Drew et al., 1981; Konings, 1982; Jackson et al., 1985a; Rajhi et al., 2011).Ethylene biosynthesis is accomplished by two main successive enzymatic reactions: conversion of S-adenosyl-Met to ACC by 1-aminocyclopropane-1-carboxylic acid synthase (ACS), and conversion of ACC to ethylene by 1-aminocyclopropane-1-carboxylic acid oxidase (ACO; Yang and Hoffman, 1984). The activities of both enzymes are enhanced during aerenchyma formation under hypoxic conditions in maize root (He et al., 1996). Since the ACC content in roots of maize is increased by oxygen deficiency and is strongly correlated with ethylene production (Atwell et al., 1988), ACC biosynthesis is essential for ethylene production during aerenchyma formation in roots. In fact, exogenously supplied ACC induced ethylene production in roots of maize (Drew et al., 1979; Konings, 1982; Atwell et al., 1988) and wheat (Yamauchi et al., 2014b), even under aerobic conditions. Ethylene production in plants is inversely related to oxygen concentration (Yang and Hoffman, 1984). Under anoxic conditions, the oxidation of ACC to ethylene by ACO, which requires oxygen, is almost completely repressed (Yip et al., 1988; Tonutti and Ramina, 1991). Indeed, anoxic conditions stimulate neither ethylene production nor aerenchyma formation in maize adventitious roots (Drew et al., 1979). Therefore, it is unlikely that the root tissues forming inducible aerenchyma are anoxic, and that the ACO-mediated step is repressed. Moreover, aerenchyma is constitutively formed in rice roots even under aerobic conditions (Jackson et al., 1985b), and thus, after the onset of waterlogging, oxygen can be immediately supplied to the apical regions of roots through the constitutively formed aerenchyma.Very-long-chain fatty acids (VLCFAs; ≥20 carbons) are major constituents of sphingolipids, cuticular waxes, and suberin in plants (Franke and Schreiber, 2007; Kunst and Samuels, 2009). In addition to their structural functions, VLCFAs directly or indirectly participate in several physiological processes (Zheng et al., 2005; Reina-Pinto et al., 2009; Roudier et al., 2010; Ito et al., 2011; Nobusawa et al., 2013; Tsuda et al., 2013), including the regulation of ethylene biosynthesis (Qin et al., 2007). During fiber cell elongation in cotton ovules, ethylene biosynthesis is enhanced by treatment with saturated VLCFAs, especially 24-carbon fatty acids, and is suppressed by an inhibitor of VLCFA biosynthesis (Qin et al., 2007). The first rate-limiting step in VLCFA biosynthesis is condensation of acyl-CoA with malonyl-CoA by β-ketoacyl-CoA synthase (KCS; Joubès et al., 2008). KCS enzymes are thought to determine the substrate and tissue specificities of fatty acid elongation (Joubès et al., 2008). The Arabidopsis (Arabidopsis thaliana) genome has 21 KCS genes (Joubès et al., 2008). In the Arabidopsis cut1 mutant, which has a defect in the gene encoding CUT1 that is required for cuticular wax production (i.e. one of the KCS genes), the expression of AtACO genes and growth of root cells were reduced when compared with the wild type (Qin et al., 2007). Furthermore, expression of the AtACO genes was rescued by exogenously supplied saturated VLCFAs (Qin et al., 2007). These observations imply that VLCFAs or their derivatives work as regulatory factors for gene expression during some physiological processes in plants.reduced culm number1 (rcn1) was first identified as a rice mutant with a low tillering rate in a paddy field (Takamure and Kinoshita, 1985; Yasuno et al., 2007). The rcn1 (rcn1-2) mutant has a single nucleotide substitution in the gene encoding a member of the ATP-binding cassette (ABC) transporter subfamily G, RCN1/OsABCG5, causing an Ala-684Pro substitution (Yasuno et al., 2009). The mutation results in several mutant phenotypes, although the substrates of RCN1/OsABCG5 have not been determined (Ureshi et al., 2012; Funabiki et al., 2013; Matsuda et al., 2014). We previously found that the rcn1 mutant has abnormal root morphology, such as shorter root length and brownish appearance of roots, under stagnant (deoxygenated) conditions (which mimics oxygen-deficient conditions in waterlogged soils). We also found that the rcn1 mutant accumulates less of the major suberin monomers originating from VLCFAs in the outer part of adventitious roots, and this results in a reduction of a functional apoplastic barrier in the root hypodermis (Shiono et al., 2014a).The objective of this study was to elucidate the molecular basis of inducible aerenchyma formation. To this end, we examined lysigenous aerenchyma formation and ACC, ethylene, and VLCFA accumulation and their biosyntheses in rcn1 roots. Based on the results of these studies, we propose that VLCFAs are involved in inducible aerenchyma formation through the enhancement of ethylene biosynthesis in rice roots.     

Transposon-Mediated Alteration of TaMATE1B Expression in Wheat Confers Constitutive Citrate Efflux from Root Apices

The TaMATE1B gene (for multidrug and toxic compound extrusion) from wheat (Triticum aestivum) was isolated and shown to encode a citrate transporter that is located on the plasma membrane. TaMATE1B expression in roots was induced by iron deficiency but not by phosphorus deficiency or aluminum treatment. The coding region of TaMATE1B was identical in a genotype showing citrate efflux from root apices (cv Carazinho) to one that lacked citrate efflux (cv Egret). However, sequence upstream of the coding region differed between these two genotypes in two ways. The first difference was a single-nucleotide polymorphism located approximately 2 kb upstream from the start codon in cv Egret. The second difference was an 11.1-kb transposon-like element located 25 bp upstream of the start codon in cv Carazinho that was absent from cv Egret. The influence of these polymorphisms on TaMATE1B expression was investigated using fusions to green fluorescent protein expressed in transgenic lines of rice (Oryza sativa). Fluorescence measurements in roots of rice indicated that 1.5- and 2.3-kb regions upstream of TaMATE1B in cv Carazinho (which incorporated 3′ regions of the transposon-like element) generated 20-fold greater expression in the apical 1 mm of root compared with the native promoter in cv Egret. By contrast, fluorescence in more mature tissues was similar in both cultivars. The presence of the single-nucleotide polymorphism alone consistently generated 2-fold greater fluorescence than the cv Egret promoter. We conclude that the transposon-like element in cv Carazinho extends TaMATE1B expression to the root apex, where it confers citrate efflux and enhanced aluminum tolerance.Intraspecific variation in aluminum (Al3+) tolerance is evident in many crop species and has been shown to be under either simple or complex genetic control (Ma et al., 2004; Magalhaes et al., 2007; Krill et al., 2010; Famoso et al., 2011). In wheat (Triticum aestivum), Al3+ tolerance is primarily associated with efflux of organic anions from root tips, with at least two independent mechanisms being involved. The first and most widely distributed among genotypes is the Al3+-activated efflux of malate from root apices (Delhaize et al., 1993, Ryan et al., 1995). More recently, a second and less prevalent mechanism has been identified that relies on the constitutive efflux of citrate from root apices (Ryan et al., 2009).Organic anions protect roots by chelating and detoxifying Al3+ in the apoplast and rhizosphere around sensitive root apices (Delhaize et al., 1993, 2012; Ryan et al., 2001). For example, Al3+-tolerant genotypes of wheat release significantly more malate than sensitive genotypes (Ryan et al., 1995). The efflux of malate from root apices is mediated by the TaALMT1 (for Al3+-activated malate transporter) gene, which resides on chromosome 4DL (Raman et al., 2005, 2008; Zhou et al., 2007). TaALMT1 encodes an Al3+-activated anion channel permeable to malate located on the plasma membrane of root cells (Sasaki et al., 2004; Yamaguchi et al., 2005; Piñeros et al., 2008; Zhang et al., 2008). Similarly, ALMT1 genes contribute to Al3+ tolerance in Arabidopsis (Arabidopsis thaliana), oilseed rape (Brassica napus), and rye (Secale cereale; Ryan et al., 2011). While all members of the ALMT family characterized to date encode transport proteins, only a minority are involved in Al3+ tolerance. The others contribute to physiological processes related to ionic relations and osmotic adjustment (Kovermann et al., 2007; Gruber et al., 2010; Meyer et al., 2010, 2011; Sasaki et al., 2010).In many plant species, citrate efflux from root apices also confers Al3+ tolerance. The genes controlling citrate efflux were first identified in sorghum (Sorghum bicolor; SbMATE [for multidrug and toxic compound extrusion]; Magalhaes et al., 2007) and barley (Hordeum vulgare; HvAACT1; Furukawa et al., 2007), with additional genes subsequently identified in Arabidopsis (Liu et al., 2009), maize (Zea mays; Maron et al., 2010), rice (Yokosho et al., 2009), and rice bean (Vigna umbellate; Yang et al., 2011). These genes belong to a large and ubiquitous family of MATE genes that were first identified in prokaryotes (Hvorup et al., 2003). SbMATE and HvAACT1 are part of a subset of plant MATE genes that facilitate citrate efflux (Liu et al., 2009; Magalhaes, 2010) with roles in Al3+ tolerance and iron (Fe) nutrition. For example, FRD3 from Arabidopsis and OsFRDL1 from rice both encode MATE proteins located in vascular tissue, where they release citrate into the xylem to enable Fe transport to shoots (Durrett et al., 2007). Recent evidence suggests that the Al3+ tolerance gene in barley, HvAACT1, was coopted from an original function in Fe nutrition by a random mutation. Al3+-tolerant genotypes of barley have a 1-kb insertion in the 5′ untranslated region (UTR) of the HvAACT1 coding region that alters its expression pattern. The insertion extends HvAACT1 expression to the root apices, which, in the presence of Al3+, causes citrate efflux from the apices and enhanced Al3+ tolerance (Fujii et al., 2012).Evidence that citrate efflux confers Al3+ tolerance in some genotypes of wheat was provided by Ryan et al. (2009). The trait was mapped to the long arm of chromosome 4B and cosegregated with an EST showing 94% sequence identity to HvAACT1. This EST was expressed only in the root apices of wheat lines that showed citrate efflux (e.g. cv Carazinho; Ryan et al., 2009), suggesting that a MATE gene encodes citrate efflux from root apices. Here, we describe the isolation and characterization of a MATE gene from wheat named TaMATE1B that encodes a citrate transporter located on the plasma membrane. A large transposable element-like sequence was found to be inserted near the start of the TaMATE1B coding region specifically in genotypes that release citrate. We demonstrate that the fragment functions as a promoter that extends TaMATE1B expression to root apices and discuss how this mutation recruited a gene to confer a new phenotype associated with Al3+ tolerance.     

ROOT HAIR DEFECTIVE3 Family of Dynamin-Like GTPases Mediates Homotypic Endoplasmic Reticulum Fusion and Is Essential for Arabidopsis Development

In all eukaryotic cells, the endoplasmic reticulum (ER) forms a tubular network whose generation requires the fusion of ER membranes. In Arabidopsis (Arabidopsis thaliana), the membrane-bound GTPase ROOT HAIR DEFECTIVE3 (RHD3) is a potential candidate to mediate ER fusion. In addition, Arabidopsis has two tissue-specific isoforms of RHD3, namely RHD3-like (RL) proteins, and their function is not clear. Here, we show that a null allele of RHD3, rhd3-8, causes growth defects and shortened root hairs. A point mutant, rhd3-1, exhibits a more severe growth phenotype than the null mutant, likely because it exerts a dominant-negative effect on the RL proteins. Genetic analysis reveals that the double deletion of RHD3 and RL1 is lethal and that the rhd3 rl2 plants produce no viable pollen, suggesting that the RL proteins are redundant to RHD3. RHD3 family proteins can replace Sey1p, the homolog of RHD3 in yeast (Saccharomyces cerevisiae), in the maintenance of ER morphology, and they are able to fuse membranes both in vivo and in vitro. Our results suggest that RHD3 proteins mediate ER fusion and are essential for plant development and that the formation of the tubular ER network is of general physiological significance.In all eukaryotic cells, the endoplasmic reticulum (ER) comprises a continuous membrane system of sheets and tubules (Baumann and Walz, 2001; Shibata et al., 2006). ER tubules frequently connect through homotypic membrane fusion to form a reticular network (Lee and Chen, 1988; Prinz et al., 2000; Du et al., 2004). ER fusion in metazoans is mediated by the atlastins (ATLs), a class of dynamin-like, membrane-bound GTPases (Hu et al., 2009; Orso et al., 2009). ATL possesses a cytoplasmic N-terminal GTPase domain, followed by a helical domain, two closely spaced transmembrane domains, and a C-terminal cytosolic tail. ATL proteins localize mostly to ER tubules and they interact with the tubule-shaping proteins, reticulons and DP1 (Hu et al., 2009). A role for the ATLs in ER fusion is suggested by the fact that depletion of ATLs leads to long, nonbranched ER tubules in cultured cells (Hu et al., 2009) and to ER fragmentation in Drosophila melanogaster (Orso et al., 2009), possibly due to insufficient fusion between the tubules. Nonbranched ER tubules are also observed upon the expression of dominant-negative ATL mutants (Hu et al., 2009). In addition, antibodies to ATL inhibit ER network formation in Xenopus laevis egg extracts (Hu et al., 2009). Moreover, proteoliposomes containing purified D. melanogaster ATL undergo GTP-dependent fusion in vitro (Orso et al., 2009; Bian et al., 2011). The physiological significance of ER fusion is supported by the observation that mutations in human ATL1, the dominant isoform in the brain, cause hereditary spastic paraplegia (Zhao et al., 2001), a neurodegenerative disease characterized by axon shortening in corticospinal motor neurons and progressive spasticity and weakness of the lower limbs (Salinas et al., 2008).Many organisms lack ATL homologs. In yeast (Saccharomyces cerevisiae), another dynamin-like GTPase, Sey1p, has been found to share the same signature motifs and membrane topology as ATL (Hu et al., 2009). Recent work suggests that Sey1p mediates ER membrane fusion both in vivo and in vitro (Anwar et al., 2012). Cells lacking Sey1p grow normally (Hu et al., 2009), but additional mutation of an ER SNARE Ufe1p, which probably represents an alternative ER fusion mechanism in yeast, causes severe growth defects (Anwar et al., 2012). In Arabidopsis (Arabidopsis thaliana), the potential functional ortholog of ATL appears to be ROOT HAIR DEFECTIVE3 (RHD3; Hu et al., 2009), which was initially discovered by a genetic screen of root hair-defective mutants (Schiefelbein and Somerville, 1990). It is sequence related to Sey1p over the entire length (Wang et al., 1997; Brands and Ho, 2002). Mutations of RHD3 cause short and wavy root hairs (Schiefelbein and Somerville, 1990; Wang et al., 1997; Stefano et al., 2012) and defects in cell expansion (Wang et al., 2002).Despite the sequence homology between Sey1p and RHD3, it was reported that Sey1p could not replace RHD3 in plants and vice versa (Chen et al., 2011). Therefore, it is not clear whether RHD3 can mediate ER fusion. Another complication in plants is that the Arabidopsis RHD3 family also contains two RHD3-like (RL) proteins (Hu et al., 2003): RL1 is expressed only in pollen, whereas RL2 is expressed ubiquitously, but both are present at very low levels. Deletion of either RL protein causes no detectable defects in root hair development or overall growth (Chen et al., 2011). Whether RL proteins support the role of RHD3 in a tissue-specific manner remains to be investigated.Here, we have analyzed the function of RHD3 and RL proteins in Arabidopsis. We show that RHD3 and the two RL proteins play redundant roles but function during different stages of Arabidopsis development. In addition, we show that RHD3 proteins can functionally replace Sey1p in yeast and mediate ER membrane fusion.     

Remodeling of the Infection Chamber before Infection Thread Formation Reveals a Two-Step Mechanism for Rhizobial Entry into the Host Legume Root Hair

In many legumes, root entry of symbiotic nitrogen-fixing rhizobia occurs via host-constructed tubular tip-growing structures known as infection threads (ITs). Here, we have used a confocal microscopy live-tissue imaging approach to investigate early stages of IT formation in Medicago truncatula root hairs (RHs) expressing fluorescent protein fusion reporters. This has revealed that ITs only initiate 10 to 20 h after the completion of RH curling, by which time major modifications have occurred within the so-called infection chamber, the site of bacterial entrapment. These include the accumulation of exocytosis (M. truncatula Vesicle-Associated Membrane Protein721e)- and cell wall (M. truncatula EARLY NODULIN11)-associated markers, concomitant with radial expansion of the chamber. Significantly, the infection-defective M. truncatula nodule inception-1 mutant is unable to create a functional infection chamber. This underlines the importance of the NIN-dependent phase of host cell wall remodeling that accompanies bacterial proliferation and precedes IT formation, and leads us to propose a two-step model for rhizobial infection initiation in legume RHs.Legumes possess the remarkable capacity to improve their nutrition by establishing a nitrogen-fixing root nodule symbiosis (RNS) with soil bacteria collectively called rhizobia. In many legumes such as Medicago truncatula, rhizobia penetrate across the root epidermis and outer cortex to reach the differentiating nodule tissues via sequentially constructed transcellular compartments known as infection threads (ITs; Gage, 2004). It is now well established that this mode of entry through specialized infection compartments, often referred to as accommodation, is shared with the more ancient arbuscular mycorrhizal (AM) symbiosis, from which the legume-Rhizobium RNS is thought to have evolved (Parniske, 2008; Markmann and Parniske, 2009). Furthermore, strong evidence indicates that the signaling and cellular mechanisms underlying IT formation in legumes are closely related to those used for infection compartment formation during AM infection of epidermal and outer cortical tissues (Bapaume and Reinhardt, 2012; Oldroyd, 2013).Rhizobial infection is set in motion after an initial molecular dialogue between symbiotic partners, in which rhizobial lipo-chitooligosaccharide (LCO) Nod factors (NFs) are key signaling molecules (for review, see Oldroyd, 2013). Host responses to NF signaling include rapid and sustained nuclear-associated Ca²⁺ oscillations (Ca²⁺ spiking; Ehrhardt et al., 1996; Oldroyd and Downie, 2006; Sieberer et al., 2009; Capoen et al., 2011) and the rapid expression of early epidermal marker genes such as M. truncatula EARLY NODULIN11 (Charron et al., 2004). The activation of nuclear Ca²⁺ spiking is one of the most characteristic features of the so-called common symbiotic signaling pathway, common to both RNS and AM (Kistner and Parniske, 2002; Singh and Parniske, 2012). Whereas these preinfection responses to NFs are observed in the majority of elongating root hairs (RHs) early after rhizobial inoculation (Journet et al., 2001; Wais et al., 2002), ITs are only formed in a small subset of RHs, and MtENOD11 expression is strongly activated at these rhizobial infection sites (Journet et al., 2001; Boisson-Dernier et al., 2005).ITs are tubular plant-derived structures delimited by a membrane that is contiguous with the RH plasmalemma and a layer of cell wall-like material, thus isolating the rhizobia from the host cell cytoplasm (Gage, 2004). These apoplastic infection compartments are progressively constructed along the length of the RH with their growing tip connected via a cytoplasmic bridge to the migrating RH nucleus. This broad cytoplasmic column provides the cell machinery for tip growth, which involves targeted exocytosis of membrane and extracellular materials to the growing apex of the IT (Oldroyd et al., 2011; Bapaume and Reinhardt, 2012). It is presumed that this cytoplasmic bridge shares an equivalent role to the prepenetration apparatus (PPA) formed at the onset of AM fungal infection (Genre et al., 2005, 2008). We now know that the IT tip region is formed in advance of rhizobial colonization and is progressively populated by dividing rhizobia that also physically move down the thread (Gage, 2004; Fournier et al., 2008). It has been proposed that the matrix of the growing IT tip is initially in a fluid or gel-like state compatible with bacterial growth and movement (Brewin, 2004; Fournier et al., 2008). This relative plasticity could result in part from the presence of atypical extracellular (glyco) proteins such as the repetitive Pro-rich proteins MtENOD11/MtENOD12 because their low Tyr content is presumed to limit cross linking to other wall components (Scheres et al., 1990; Pichon et al., 1992; Journet et al., 2001).Nevertheless, the mechanism by which rhizobial IT formation is initiated in RHs is not clear. Whereas AM fungal hyphae form contact structures called hyphopodia on the exposed surface of nonhair epidermal cells prior to PPA formation and perifungal infection compartment formation (Genre et al., 2005), rhizobial entry requires that the bacteria first become entrapped between RH walls. Attachment of rhizobia close to a growing RH tip induces a continuous reorientation of tip growth, most likely the result of localized NF production (Esseling et al., 2003), eventually leading to RH curling and subsequent bacterial entrapment within a closed chamber in the center of the curl (Catoira et al., 2001; Geurts et al., 2005). Rhizobial entrapment can also occur between the cell walls of two touching RHs (Dart, 1974; Gage, 2004).The closed chamber in curled RHs has often been termed the infection pocket (e.g. Murray, 2011; Guan et al., 2013). However, because this term is also used to designate a quite different and larger structure formed in root subepidermal tissues of legumes during intercellular infection after crack entry and involving localized cell death (Goormachtig et al., 2004), we propose to use the term infection chamber to describe the unique enclosure formed during rhizobial RH infection.After entrapment, it has been proposed that rhizobia multiply to form a so-called microcolony (Gage et al., 1996; Limpens et al., 2003), and that IT polar growth initiates in front of this microcolony by local invagination of the RH plasmalemma combined with exocytosis of extracellular materials (Gage, 2004). Furthermore, it has been suggested that localized degradation of the chamber wall would allow the rhizobia to access the newly formed IT (Callaham and Torrey, 1981; Turgeon and Bauer, 1985). However, a detailed investigation of this particular stage of rhizobial infection is lacking, particularly concerning when and where the rhizobia/cell wall interface becomes modified. Such studies have been limited until now, notably because ITs develop only in a low proportion of curled RHs (Dart, 1974).To attempt to answer this question, we have used a live-tissue imaging approach developed for in vivo confocal microscopy in M. truncatula (Fournier et al., 2008; Cerri et al., 2012; Sieberer et al., 2012) and particularly well adapted to time-lapse studies of the initial stages of rhizobial infection, including RH curling and IT formation. To investigate modifications occurring at the RH interface with the enclosed rhizobia during these early stages, we prepared M. truncatula plants expressing fluorescent protein fusions aimed at detecting both exocytosis activity and cell wall remodeling during the initial construction of the IT apoplastic compartment. To this end, we made use of the M. truncatula Vesicle-Associated Membrane Protein721e (MtVAMP721e; Ivanov et al., 2012), recently shown to label exocytosis sites both in growing RHs and during AM colonization (Genre et al., 2012), as well as the infection- and cell wall-associated MtENOD11 Pro-rich glycoprotein (Journet et al., 2001). Our experiments have revealed that IT development in curled RHs only initiates after a lengthy interval of 10 to 20 h, during which sustained exocytosis and MtENOD11 secretion to the infection chamber are associated with radial expansion as well as remodeling of the surrounding walls. Importantly, it was found that the infection-defective M. truncatula nodule inception-1 (Mtnin-1) mutant (Marsh et al., 2007) is impaired in chamber remodeling. Our findings led us to propose a new model for IT formation in which the infection chamber first differentiates into a globular apoplastic compartment displaying similarities to the future IT, and in which the enclosed rhizobia multiply. This is then followed by a switch from radial to tubular growth corresponding to tip-driven IT growth and associated movement of rhizobia into the extending thread. Importantly, this two-step model no longer requires that the host cell wall is degraded to allow access of the colonizing rhizobia to the newly initiated IT.     

Natural Variation of Arabidopsis Root Architecture Reveals Complementing Adaptive Strategies to Potassium Starvation

Root architecture is a highly plastic and environmentally responsive trait that enables plants to counteract nutrient scarcities with different foraging strategies. In potassium (K) deficiency (low K), seedlings of the Arabidopsis (Arabidopsis thaliana) reference accession Columbia (Col-0) show a strong reduction of lateral root elongation. To date, it is not clear whether this is a direct consequence of the lack of K as an osmoticum or a triggered response to maintain the growth of other organs under limiting conditions. In this study, we made use of natural variation within Arabidopsis to look for novel root architectural responses to low K. A comprehensive set of 14 differentially responding root parameters were quantified in K-starved and K-replete plants. We identified a phenotypic gradient that links two extreme strategies of morphological adaptation to low K arising from a major tradeoff between main root (MR) and lateral root elongation. Accessions adopting strategy I (e.g. Col-0) maintained MR growth but compromised lateral root elongation, whereas strategy II genotypes (e.g. Catania-1) arrested MR elongation in favor of lateral branching. K resupply and histochemical staining resolved the temporal and spatial patterns of these responses. Quantitative trait locus analysis of K-dependent root architectures within a Col-0 × Catania-1 recombinant inbred line population identified several loci each of which determined a particular subset of root architectural parameters. Our results indicate the existence of genomic hubs in the coordinated control of root growth in stress conditions and provide resources to facilitate the identification of the underlying genes.The ability of plants to actively respond to nutrient scarcity with changes in root architecture is a fascinating phenomenon. Advances in root research and breeding efforts that focus on the enhancement of root traits have been recognized as principal goals to ensure those high yields necessary to feed an ever-growing human population (Hammer et al., 2009; Den Herder et al., 2010). Indeed, understanding the adaptations of root systems to environmental factors has been pointed out as a key issue in modern agriculture (Den Herder et al., 2010).Potassium (K) is the quantitatively most important cation for plant growth, as it serves as the major osmoticum for cell expansion (Leigh and Wyn Jones, 1984; Amtmann et al., 2006). Moreover, K is essential for many cellular and tissue processes, such as enzymatic activity, transport of minerals and metabolites, and regulation of stomatal aperture (Amtmann et al., 2006). Even in fertilized fields, rapid K uptake by plants can lead to K shortage in the root environment, especially early in the growth season. Root adaptations to K deficiency (low K) take place at the physiological (Armengaud et al., 2004; Shin and Schachtman, 2004; Alemán et al., 2011), metabolic (Armengaud et al., 2009a), and morphological levels. In a classic study, Drew (1975) showed an increase in overall lateral root (LR) growth of barley seedlings, even when K was supplied only to parts of the root system. Conversely, a typical response of Arabidopsis (Arabidopsis thaliana) Columbia (Col-0) seedlings to low K is the drastic reduction of LR elongation (Armengaud et al., 2004; Shin and Schachtman, 2004). Conflicting data have been published on the effect of low K on main root (MR) growth in the same species, ranging from no effect (Shin and Schachtman, 2004) to impaired MR elongation (Jung et al., 2009; Kim et al., 2010). Some components involved in K starvation responses have been identified, such as jasmonates (Armengaud et al., 2004, 2010), reactive oxygen species (Shin and Schachtman, 2004), and ethylene (Jung et al., 2009). However, the molecular identity of a root K sensor acting at the base of the signaling cascade is so far unknown.Genetic variation within species is a useful resource to dissect the genetic components determining phenotypes (Koornneef et al., 2004; Trontin et al., 2011; Weigel, 2012). Natural variation within Arabidopsis has been the basis for many studies on plant morphology, physiology, and development as well as stress response (Alonso-Blanco et al., 2009; Weigel, 2012). Natural variation of root traits such as primary root length (Mouchel et al., 2004; Loudet et al., 2005; Sergeeva et al., 2006), LR length (Loudet et al., 2005), and total root size (Fitz Gerald et al., 2006) have pinpointed genomic regions underlying the phenotypic variation via mapping of quantitative trait loci (QTLs) as a first step toward the identification of novel regulatory genes (Mouchel et al., 2004). This strategy has also been applied to environmental responses, such as growth responses to phosphate starvation (Reymond et al., 2006; Svistoonoff et al., 2007). However, despite their importance for plant growth and their strong effect on overall root architecture, root responses to K deficiency have not been genetically dissected.Here, we show that Arabidopsis accessions follow different strategies to adapt to K starvation. We present the quantification of a comprehensive set of root architectural parameters of Arabidopsis grown in K-sufficient and K-deficient media and the identification of genetic loci, each of which determines the response of a distinct subset of root architectural parameters to K starvation.