The Pseudomonas syringae Effector HopQ1 Promotes Bacterial Virulence and Interacts with Tomato 14-3-3 Proteins in a Phosphorylation-Dependent Manner

A key virulence strategy of bacterial pathogens is the delivery of multiple pathogen effector proteins into host cells during infection. The Hrp outer protein Q (HopQ1) effector from Pseudomonas syringae pv tomato (Pto) strain DC3000 is conserved across multiple bacterial plant pathogens. Here, we investigated the virulence function and host targets of HopQ1 in tomato (Solanum lycopersicum). Transgenic tomato lines expressing dexamethasone-inducible HopQ1 exhibited enhanced disease susceptibility to virulent Pto DC3000, the Pto ΔhrcC mutant, and decreased expression of a pathogen-associated molecular pattern-triggered marker gene after bacterial inoculation. HopQ1-interacting proteins were coimmunoprecipitated and identified by mass spectrometry. HopQ1 can associate with multiple tomato 14-3-3 proteins, including TFT1 and TFT5. HopQ1 is phosphorylated in tomato, and four phosphorylated peptides were identified by mass spectrometry. HopQ1 possesses a conserved mode I 14-3-3 binding motif whose serine-51 residue is phosphorylated in tomato and regulates its association with TFT1 and TFT5. Confocal microscopy and fractionation reveal that HopQ1 exhibits nucleocytoplasmic localization, while HopQ1 dephosphorylation mimics exhibit more pronounced nuclear localization. HopQ1 delivered from Pto DC3000 was found to promote bacterial virulence in the tomato genotype Rio Grande 76R. However, the HopQ1(S51A) mutant delivered from Pto DC3000 was unable to promote pathogen virulence. Taken together, our data demonstrate that HopQ1 enhances bacterial virulence and associates with tomato 14-3-3 proteins in a phosphorylation-dependent manner that influences HopQ1’s subcellular localization and virulence-promoting activities in planta.The ability to detect and mount a defense response against pathogenic microbes is vital for plant survival. Plants rely on both passive and active defenses to ward off microbial pathogens. Physical barriers, such as the cell wall and cuticle, as well as chemical barriers provide a first line of defense against microbial colonization. Unlike animals, plants do not possess a circulating immune system and rely on innate immunity for active defenses against microbial pathogens (Spoel and Dong, 2012). Plants use surface-localized receptors to recognize conserved pathogen-associated molecular patterns (PAMPs), such as bacterial flagellin, resulting in pattern-triggered immunity (PTI; Zipfel et al., 2006). Plants also use primarily intracellular nucleotide-binding domain, Leu-rich repeat containing (NLR) immune receptors to recognize pathogen effectors delivered into host cells during infection (Spoel and Dong, 2012). NLR activation results in effector-triggered immunity (ETI). A signature of ETI is the hypersensitive response (HR), a form of programmed cell death occurring at the site of infection.In order to cause disease and suppress host defense responses, gram-negative bacterial pathogens deliver effector proteins into host cells via the type III secretion system (TTSS). Plant pathogenic bacteria deliver a large number (20–40) of effectors into host cells during infection (Cui et al., 2009). Collectively, effectors are required for bacterial virulence (Lindgren et al., 1986). However, knockouts affecting individual effectors frequently have phenotypes that are subtle, likely due to functional redundancy (Cunnac et al., 2011). Alternatively, individual effectors may play an important role in bacterial survival under conditions that are not typically analyzed in the laboratory or act cooperatively with one another. Progress in understanding individual effectors’ contributions to virulence has been made by generating transgenic plants that express effectors. Multiple effectors have been shown to suppress plant innate immunity and promote bacterial growth when either transiently or stably expressed in plants (Jamir et al., 2004; Guo et al., 2009). Effector expression can also result in avirulent phenotypes when a plant NLR receptor recognizes a cognate effector and mounts an HR. Such an HR phenotype can be used to dissect important effector domains required for plant recognition and enzymatic activity.Elucidating effector targets and enzymatic activity is necessary in order to understand how they act to subvert plant immune responses and can provide elegant insight into biological processes. Significant progress has been made in elucidating the enzymatic activity of a subset of effectors. Some of the most well-characterized effectors come from Pseudomonas syringae pv tomato (Pto), the causal agent of bacterial speck on tomato (Solanum lycopersicum) and Arabidopsis (Arabidopsis thaliana). Multiple effectors can suppress immune responses by directly targeting PAMP receptors (AvrPto and AvrPtoB) or by interfering with downstream signaling processes (AvrB, AvrPphB, and HopAI1; Cui et al., 2009, 2010). The HopU1 effector interferes with RNA metabolism (Fu et al., 2007), and the HopI1 effector targets heat-shock proteins in the plant chloroplast (Jelenska et al., 2010).14-3-3s are conserved eukaryotic proteins that bind a diverse set of phosphorylated client proteins, typically at one of three distinct 14-3-3 binding motifs (Bridges and Moorhead, 2005). There are common recognition motifs for 14-3-3 proteins that contain phosphorylated Ser or Thr residues, but binding to nonphosphorylated ligands and to proteins lacking consensus motifs has been reported (Henriksson et al., 2002; Smith et al., 2011). The 14-3-3 mode I consensus motif is RXXpS/pTX and that of mode II is RXXXpS/pTXP, where X can be any amino acid and p indicates the site of phosphorylation (Smith et al., 2011). 14-3-3 proteins can also bind to the extreme C termini of proteins at the RXXpS/pTX-COOH mode III consensus motif (Smith et al., 2011). Interaction with 14-3-3s can regulate protein activity by influencing client subcellular localization, structure, and protein-protein interactions (Bridges and Moorhead, 2005). Recently, the Xanthomonas campestris XopN effector was shown to target tomato 14-3-3 isoforms, which facilitates its interaction with the tomato atypical receptor kinase1 and suppresses PTI (Kim et al., 2009; Taylor et al., 2012). Other 14-3-3s have also been shown to play a role during plant defense responses. The tomato TFT7 14-3-3 interacts with multiple mitogen-activated protein kinases to positively regulate HR induced by ETI (Oh and Martin, 2011). The Arabidopsis 14-3-3 isoform λ interacts with the RPW8.2 powdery mildew receptor and is required for complete RPW8.2-mediated resistance (Yang et al., 2009).In this study, we investigated the function of the Pto HopQ1 (for Hrp outer protein Q [also known as HopQ1-1]) effector in tomato. HopQ1 is an active effector that is transcribed and translocated via the TTSS (Schechter et al., 2004). HopQ1 induces cell death when expressed in Nicotiana benthamiana and therefore contributes to differences in host range in P. syringae pathovars on Nicotiana spp. (Wei et al., 2007; Ferrante et al., 2009). HopQ1 was also reported to slightly enhance disease symptoms (approximately 0.2 log) and bacterial virulence on bean (Phaseolus vulgaris) when expressed from P. syringae pv tabaci (Ferrante et al., 2009). Here, we generated transgenic tomato plants expressing HopQ1 that exhibited enhanced susceptibility to virulent Pto as well as the Pto ΔhrcC mutant. HopQ1-interacting proteins were identified from tomato using coimmunoprecipitations coupled with mass spectrometry. Multiple 14-3-3 proteins were identified. HopQ1 possesses a 14-3-3 binding motif whose Ser residue is phosphorylated in planta and affects its association with the tomato 14-3-3s TFT1 and TFT5. Mutation of HopQ1’s 14-3-3 binding motif affected its ability to promote bacterial virulence. Taken together, these results indicate that phosphorylation and subsequent interaction with tomato 14-3-3 proteins affect HopQ1’s virulence-promoting activities and subcellular localization.     

Evolution of Conifer Diterpene Synthases: Diterpene Resin Acid Biosynthesis in Lodgepole Pine and Jack Pine Involves Monofunctional and Bifunctional Diterpene Synthases

Abscisic acid (ABA) induces stomatal closure and inhibits light-induced stomatal opening. The mechanisms in these two processes are not necessarily the same. It has been postulated that the ABA receptors involved in opening inhibition are different from those involved in closure induction. Here, we provide evidence that four recently identified ABA receptors (PYRABACTIN RESISTANCE1 [PYR1], PYRABACTIN RESISTANCE-LIKE1 [PYL1], PYL2, and PYL4) are not sufficient for opening inhibition in Arabidopsis (Arabidopsis thaliana). ABA-induced stomatal closure was impaired in the pyr1/pyl1/pyl2/pyl4 quadruple ABA receptor mutant. ABA inhibition of the opening of the mutant’s stomata remained intact. ABA did not induce either the production of reactive oxygen species and nitric oxide or the alkalization of the cytosol in the quadruple mutant, in accordance with the closure phenotype. Whole cell patch-clamp analysis of inward-rectifying K⁺ current in guard cells showed a partial inhibition by ABA, indicating that the ABA sensitivity of the mutant was not fully impaired. ABA substantially inhibited blue light-induced phosphorylation of H⁺-ATPase in guard cells in both the mutant and the wild type. On the other hand, in a knockout mutant of the SNF1-related protein kinase, srk2e, stomatal opening and closure, reactive oxygen species and nitric oxide production, cytosolic alkalization, inward-rectifying K⁺ current inactivation, and H⁺-ATPase phosphorylation were not sensitive to ABA.The phytohormone abscisic acid (ABA), which is synthesized in response to abiotic stresses, plays a key role in the drought hardiness of plants. Reducing transpirational water loss through stomatal pores is a major ABA response (Schroeder et al., 2001). ABA promotes the closure of open stomata and inhibits the opening of closed stomata. These effects are not simply the reverse of one another (Allen et al., 1999; Wang et al., 2001; Mishra et al., 2006).A class of receptors of ABA was identified (Ma et al., 2009; Park et al., 2009; Santiago et al., 2009; Nishimura et al., 2010). The sensitivity of stomata to ABA was strongly decreased in quadruple and sextuple mutants of the ABA receptor genes PYRABACTIN RESISTANCE/PYRABACTIN RESISTANCE-LIKE/REGULATORY COMPONENT OF ABSCISIC ACID RECEPTOR (PYR/PYL/RCAR; Nishimura et al., 2010; Gonzalez-Guzman et al., 2012). The PYR/PYL/RCAR receptors are involved in the early ABA signaling events, in which a sequence of interactions of the receptors with PROTEIN PHOSPHATASE 2Cs (PP2Cs) and subfamily 2 SNF1-RELATED PROTEIN KINASES (SnRK2s) leads to the activation of downstream ABA signaling targets in guard cells (Cutler et al., 2010; Kim et al., 2010; Weiner et al., 2010). Studies of Commelina communis and Vicia faba suggested that the ABA receptors involved in stomatal opening are not the same as the ABA receptors involved in stomatal closure (Allan et al., 1994; Anderson et al., 1994; Assmann, 1994; Schwartz et al., 1994). The roles of PYR/PYL/RCAR in either stomatal opening or closure remained to be elucidated.Blue light induces stomatal opening through the activation of plasma membrane H⁺-ATPase in guard cells that generates an inside-negative electrochemical gradient across the plasma membrane and drives K⁺ uptake through voltage-dependent inward-rectifying K⁺ channels (Assmann et al., 1985; Shimazaki et al., 1986; Blatt, 1987; Schroeder et al., 1987; Thiel et al., 1992). Phosphorylation of the penultimate Thr of the plasma membrane H⁺-ATPase is a prerequisite for blue light-induced activation of the H⁺-ATPase (Kinoshita and Shimazaki, 1999, 2002). ABA inhibits H⁺-ATPase activity through dephosphorylation of the penultimate Thr in the C terminus of the H⁺-ATPase in guard cells, resulting in prevention of the opening (Goh et al., 1996; Zhang et al., 2004; Hayashi et al., 2011). Inward-rectifying K⁺ currents (I_Kin) of guard cells are negatively regulated by ABA in addition to through the decline of the H⁺ pump-driven membrane potential difference (Schroeder and Hagiwara, 1989; Blatt, 1990; McAinsh et al., 1990; Schwartz et al., 1994; Grabov and Blatt, 1999; Saito et al., 2008). This down-regulation of ion transporters by ABA is essential for the inhibition of stomatal opening.A series of second messengers has been shown to mediate ABA-induced stomatal closure. Reactive oxygen species (ROS) produced by NADPH oxidases play a crucial role in ABA signaling in guard cells (Pei et al., 2000; Zhang et al., 2001; Kwak et al., 2003; Sirichandra et al., 2009; Jannat et al., 2011). Nitric oxide (NO) is an essential signaling component in ABA-induced stomatal closure (Desikan et al., 2002; Guo et al., 2003; Garcia-Mata and Lamattina, 2007; Neill et al., 2008). Alkalization of cytosolic pH in guard cells is postulated to mediate ABA-induced stomatal closure in Arabidopsis (Arabidopsis thaliana) and Pisum sativum and Paphiopedilum species (Irving et al., 1992; Gehring et al., 1997; Grabov and Blatt, 1997; Suhita et al., 2004; Gonugunta et al., 2008). These second messengers transduce environmental signals to ion channels and ion transporters that create the driving force for stomatal movements (Ward et al., 1995; MacRobbie, 1998; Garcia-Mata et al., 2003).In this study, we examined the mobilization of second messengers, the inactivation of I_Kin, and the suppression of H⁺-ATPase phosphorylation evoked by ABA in Arabidopsis mutants to clarify the downstream signaling events of ABA signaling in guard cells. The mutants included a quadruple mutant of PYR/PYL/RCARs, pyr1/pyl1/pyl2/pyl4, and a mutant of a SnRK2 kinase, srk2e.     

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.     

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.     

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.     

Salt Stress Reduces Root Meristem Size by Nitric Oxide-Mediated Modulation of Auxin Accumulation and Signaling in Arabidopsis

The development of the plant root system is highly plastic, which allows the plant to adapt to various environmental stresses. Salt stress inhibits root elongation by reducing the size of the root meristem. However, the mechanism underlying this process remains unclear. In this study, we explored whether and how auxin and nitric oxide (NO) are involved in salt-mediated inhibition of root meristem growth in Arabidopsis (Arabidopsis thaliana) using physiological, pharmacological, and genetic approaches. We found that salt stress significantly reduced root meristem size by down-regulating the expression of PINFORMED (PIN) genes, thereby reducing auxin levels. In addition, salt stress promoted AUXIN RESISTANT3 (AXR3)/INDOLE-3-ACETIC ACID17 (IAA17) stabilization, which repressed auxin signaling during this process. Furthermore, salt stress stimulated NO accumulation, whereas blocking NO production with the inhibitor N^ω-nitro-l-arginine-methylester compromised the salt-mediated reduction of root meristem size, PIN down-regulation, and stabilization of AXR3/IAA17, indicating that NO is involved in salt-mediated inhibition of root meristem growth. Taken together, these findings suggest that salt stress inhibits root meristem growth by repressing PIN expression (thereby reducing auxin levels) and stabilizing IAA17 (thereby repressing auxin signaling) via increasing NO levels.Due to agricultural practices and climate change, soil salinity has become a serious factor limiting the productivity and quality of agricultural crops (Zhu, 2007). Worldwide, high salinity in the soil damages approximately 20% of total irrigated lands and takes 1.5 million ha out of production each year (Munns and Tester, 2008). In general, high salinity affects plant growth and development by reducing plant water potential, altering nutrient uptake, and increasing the accumulation of toxic ions (Hasegawa et al., 2000; Munns, 2002; Zhang and Shi, 2013). Together, these effects severely reduce plant growth and survival.Because the root is the first organ to sense high salinity, salt stress plays a direct, important role in modulating root system architecture (Wang et al., 2009). For instance, salt stress negatively regulates root hair formation and gravitropism (Sun et al., 2008; Wang et al., 2008). The role of salt in lateral root formation depends on the NaCl concentration. While high NaCl levels inhibit lateral root formation, lower NaCl levels stimulate lateral root formation in an auxin-dependent manner (Zolla et al., 2010; Ji et al., 2013). The root meristem plays an essential role in sustaining root growth (Perilli et al., 2012). Salt stress inhibits primary root elongation by suppressing root meristem activity (West et al., 2004). However, how this inhibition occurs remains largely unclear.Plant hormones are important intermediary signaling compounds that function downstream of environmental stimuli. Among plant hormones, indole-3-acetic acid (IAA) is thought to play a fundamental role in root system architecture by regulating cell division, expansion, and differentiation. In Arabidopsis (Arabidopsis thaliana) root tips, a distal auxin maximum is formed and maintained by polar auxin transport (PAT), which determines the orientation and extent of cell division in the root meristem as well as root pattern formation (Sabatini et al., 1999). PINFORMED (PIN) proteins, which are components of the auxin efflux machinery, regulate primary root elongation and root meristem size (Blilou et al., 2005; Dello Ioio et al., 2008; Yuan et al., 2013, 2014). The auxin signal transduction pathway is activated by direct binding of auxin to its receptor protein, TRANSPORT INHIBITOR RESPONSE1 (TIR1)/AUXIN SIGNALING F-BOX (AFB), promoting the degradation of Aux/IAA proteins, releasing auxin response factors (ARFs), and activating the expression of auxin-responsive genes (Gray et al., 2001; Dharmasiri et al., 2005a; Kepinski and Leyser, 2005). Aux/IAA proteins are short-lived, nuclear-localized proteins that play key roles in auxin signal activation and root growth modulation (Rouse et al., 1998). Other hormones and stresses often regulate auxin signaling by affecting Aux/IAA protein stability (Lim and Kunkel, 2004; Nemhauser et al., 2004; Wang et al., 2007; Kushwah and Laxmi, 2014).Nitric oxide (NO) is a signaling molecule with diverse biological functions in plants (He et al., 2004; Fernández-Marcos et al., 2011; Shi et al., 2012), including important roles in the regulation of root growth and development. NO functions downstream of auxin during the adventitious rooting process in cucumber (Cucumis sativus; Pagnussat et al., 2002). Exogenous auxin-induced NO biosynthesis is associated with nitrate reductase activity during lateral root formation, and NO is necessary for auxin-induced lateral root and root hair development (Pagnussat et al., 2002; Lombardo et al., 2006). Pharmacological and genetic analyses in Arabidopsis indicate that NO suppresses primary root growth and root meristem activity (Fernández-Marcos et al., 2011). Additionally, both exogenous application of the NO donor sodium nitroprusside (SNP) and overaccumulation of NO in the mutant chlorophyll a/b binding protein underexpressed1 (cue1)/nitric oxide overproducer1 (nox1) result in reduced PIN1 expression and auxin accumulation in root tips. The auxin receptors protein TIR1 is S-nitrosylated by NO, suggesting that this protein is a direct target of NO in the regulation of root development (Terrile et al., 2012).Because NO is a free radical, NO levels are dynamically regulated by endogenous and environmental cues. Many phytohormones, including abscisic acid, auxin, cytokinin, salicylic acid, jasmonic acid, and ethylene, induce NO biosynthesis (Zottini et al., 2007; Kolbert et al., 2008; Tun et al., 2008; García et al., 2011). In addition, many abiotic and biotic stresses or stimuli, such as cold, heat, salt, drought, heavy metals, and pathogens/elicitors, also stimulate NO biosynthesis (Zhao et al., 2009; Mandal et al., 2012). Salt stress stimulates NO and ONOO^– accumulation in roots (Corpas et al., 2009), but the contribution of NO to root meristem growth under salinity stress has yet to be examined in detail.In this study, we found that salt stress significantly down-regulated the expression of PIN genes and promoted AUXIN RESISTANT3 (AXR3)/IAA17 stabilization. Furthermore, salt stress stimulated NO accumulation, and pharmacological inhibition of NO biosynthesis compromised the salt-mediated reduction in root meristem size. Our results support a model in which salt stress reduces root meristem size by increasing NO accumulation, which represses PIN expression and stabilizes IAA17, thereby reducing auxin levels and repressing auxin signaling.     

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.