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.     

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.     

Hydrogen Sulfide Regulates Inward-Rectifying K+ Channels in Conjunction with Stomatal Closure

Hydrogen sulfide (H₂S) is the third biological gasotransmitter, and in animals, it affects many physiological processes by modulating ion channels. H₂S has been reported to protect plants from oxidative stress in diverse physiological responses. H₂S closes stomata, but the underlying mechanism remains elusive. Here, we report the selective inactivation of current carried by inward-rectifying K⁺ channels of tobacco (Nicotiana tabacum) guard cells and show its close parallel with stomatal closure evoked by submicromolar concentrations of H₂S. Experiments to scavenge H₂S suggested an effect that is separable from that of abscisic acid, which is associated with water stress. Thus, H₂S seems to define a unique and unresolved signaling pathway that selectively targets inward-rectifying K⁺ channels.Hydrogen sulfide (H₂S) is a small bioactive gas that has been known for centuries as an environmental pollutant (Reiffenstein et al., 1992). H₂S is soluble in both polar and, especially, nonpolar solvents (Wang, 2002), and has recently come to be recognized as the third member of a group of so-called biological gasotransmitters. Most importantly, H₂S shows both physical and functional similarities to the other gasotransmitters nitric oxide (NO) and carbon monoxide (Wang, 2002), and it has been shown to participate in diverse physiological processes in animals, including cardioprotection, neuromodulation, inflammation, apoptosis, and gastrointestinal functions among others (Kabil et al., 2014). Less is known about H₂S molecular targets and its modes of action. H₂S can directly modify specific targets through protein sulfhydration (the addition of an -SH group to thiol moiety of proteins; Mustafa et al., 2009) or reaction with metal centers (Li and Lancaster, 2013). It can also act indirectly, reacting with NO to form nitrosothiols (Whiteman et al., 2006; Li and Lancaster, 2013). Among its molecular targets, H₂S has been reported to regulate ATP-dependent K⁺ channels (Yang et al., 2005), Ca²⁺-activated K⁺ channels, T- and L-type Ca²⁺ channels, and transient receptor potential channels (Tang et al., 2010; Peers et al., 2012), suggesting H₂S as a key regulator of membrane ion transport.In plants, H₂S is produced enzymatically by the desulfhydration of l-Cys to form H₂S, pyruvate, and ammonia in a reaction catalyzed by the enzyme l-Cys desulfhydrase (Riemenschneider et al., 2005a, 2005b), DES1, that has been characterized in Arabidopsis (Arabidopsis thaliana; Alvarez et al., 2010). Alternatively, H₂S can be produced from d-Cys by d-Cys desulfhydrase (Riemenschneider et al., 2005a, 2005b) and in cyanide metabolism by β-cyano-Ala synthase (García et al., 2010). H₂S action was originally related to pathogenesis resistance (Bloem et al., 2004), but in the last decade it has been proven to have an active role in signaling, participating in key physiological processes, such as germination and root organogenesis (Zhang et al., 2008, 2009a), heat stress (Li et al., 2013a, 2013b), osmotic stress (Zhang et al., 2009b), and stomatal movement (García-Mata and Lamattina, 2010; Lisjak et al., 2010, 2011; Jin et al., 2013). Moreover, H₂S was reported to participate in the signaling of plant hormones, including abscisic acid (ABA; García-Mata and Lamattina, 2010; Lisjak et al., 2010; Jin et al., 2013; Scuffi et al., 2014), ethylene (Hou et al., 2013), and auxin (Zhang et al., 2009a).ABA is an important player in plant physiology. Notably, upon water stress, ABA triggers a complex signaling network to restrict the loss of water through the transpiration stream, balancing these needs with those of CO₂ for carbon assimilation. In the guard cells that surround the stomatal pore, ABA induces an increase of cytosolic-free Ca²⁺ concentration ([Ca2+]_cyt), elevates cytosolic pH (pH_i), and activates the efflux of anions, mainly chloride, through S- and R-type anion channels. The increase in [Ca2+]_cyt inactivates inward-rectifying K⁺ channels (I_KIN); anion efflux depolarizes the plasma membrane, and together with the rise in pH_i, it activates K⁺ efflux through outward-rectifying K⁺ channels (I_KOUT; Blatt, 2000; Schroeder et al., 2001). These changes in ion flux, in turn, generate an osmotically driven reduction in turgor and volume and closure of the stomatal pore. All three gasotransmitters have been implicated in regulating the activity of guard cell ion channels, but direct evidence is available only for NO (Garcia-Mata et al., 2003; Sokolovski et al., 2005). Here, we have used two-electrode voltage clamp measurements to study the role of H₂S in the regulation of the guard cell K⁺ channels of tobacco (Nicotiana tabacum). Our results show that H₂S selectively inactivates I_KIN and that this action parallels that of stomatal closure. These results confirm H₂S as a unique factor regulating guard cell ion transport and indicate that H₂S acts in a manner separable from that of ABA.     

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

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

The Type II NADPH Dehydrogenase Facilitates Cyclic Electron Flow,Energy-Dependent Quenching,and Chlororespiratory Metabolism during Acclimation of Chlamydomonas reinhardtii to Nitrogen Deprivation

When photosynthetic organisms are deprived of nitrogen (N), the capacity to grow and assimilate carbon becomes limited, causing a decrease in the productive use of absorbed light energy and likely a rise in the cellular reduction state. Although there is a scarcity of N in many terrestrial and aquatic environments, a mechanistic understanding of how photosynthesis adjusts to low-N conditions and the enzymes/activities integral to these adjustments have not been described. In this work, we use biochemical and biophysical analyses of photoautotrophically grown wild-type and mutant strains of Chlamydomonas reinhardtii to determine the integration of electron transport pathways critical for maintaining active photosynthetic complexes even after exposure of cells to N deprivation for 3 d. Key to acclimation is the type II NADPH dehydrogenase, NDA2, which drives cyclic electron flow (CEF), chlororespiration, and the generation of an H⁺ gradient across the thylakoid membranes. N deprivation elicited a doubling of the rate of NDA2-dependent CEF, with little contribution from PGR5/PGRL1-dependent CEF. The H⁺ gradient generated by CEF is essential to sustain nonphotochemical quenching, while an increase in the level of reduced plastoquinone would promote a state transition; both are necessary to down-regulate photosystem II activity. Moreover, stimulation of NDA2-dependent chlororespiration affords additional relief from the elevated reduction state associated with N deprivation through plastid terminal oxidase-dependent water synthesis. Overall, rerouting electrons through the NDA2 catalytic hub in response to photoautotrophic N deprivation sustains cell viability while promoting the dissipation of excess excitation energy through quenching and chlororespiratory processes.Oxygenic photosynthesis involves the conversion of light energy into chemical bond energy by plants, green algae, and cyanobacteria and the use of that energy to fix CO₂. The photosynthetic electron transport system, located in thylakoid membranes, involves several major protein complexes: PSII (water-plastoquinone oxidoreductase), cytochrome b₆f (cyt b6f; plastoquinone-plastocyanin oxidoreductase), PSI (plastocyanin-ferredoxin oxidoreductase), and the ATP synthase (CF_oCF₁). Light energy absorbed by the photosynthetic apparatus is used to establish both linear electron flow (LEF) and cyclic electron flow (CEF), which drive the production of ATP and NADPH, the chemical products of the light reactions needed for CO₂ fixation in the Calvin-Benson-Bassham (CBB) cycle.With the absorption of light energy by pigment-protein complexes associated with PSII, energy is funneled into unique chlorophyll (Chl) molecules located in the PSII reaction center (RC), where it can elicit a charge separation that generates a large enough oxidizing potential to extract electrons from water. In LEF, electrons from PSII RCs are transferred sequentially along a set of electron carriers, initially reducing the plastoquinone (PQ) pool, then the cyt b₆f complex, and subsequently the lumenal electron carrier plastocyanin (PC). Light energy absorbed by PSI excites a special pair of Chl molecules (P700), causing a charge separation that generates the most negative redox potential in nature (Nelson and Yocum, 2006). The energized electron, which is replaced by electrons from PC, is sequentially transferred to ferredoxin and ferredoxin NADP⁺ reductase, generating reductant in the form of NADPH.Electron transport from water to NADPH in LEF is accompanied by the transport of H⁺ into the thylakoid lumen. For each water molecule oxidized, two H⁺ are released in the thylakoid lumen. In addition, H⁺ are moved into the lumen by the transfer of electrons through cyt b₆f (Q cycle). H⁺ accumulation in the thylakoid lumen dramatically alters the lumenal pH, and the transmembrane H⁺ gradient (ΔpH) together with the transmembrane ion gradient constitute the proton motive force (pmf), which drives ATP formation by ATP synthase (Mitchell, 1961, 1966, 2011). This pmf also promotes other cellular processes, including the dissipation of excess absorbed excitation energy as heat in a photoprotective process (see below; Li et al., 2009; Erickson et al., 2015). The NADPH and ATP molecules generated by LEF and CEF fuel the synthesis of reduced carbon backbones (in the CBB cycle) used in the production of many cellular metabolites and fixed carbon storage polymers.A basic role for CEF is to increase the ATP-NADPH ratio, which can satisfy the energy requirements of the cell and augment the synthesis of ATP by LEF, which is required to sustain CO₂ fixation by the CBB cycle (Allen, 2003; Kramer et al., 2004; Iwai et al., 2010; Alric, 2014). There are two distinct CEF pathways identified in plants and algae. In both pathways, electrons flow from the PQ pool through cyt b₆f to reduce the oxidized form of P700 (P700⁺). In one CEF pathway, electrons are transferred back to the PQ pool prior to the formation of NADPH. This route involves the proteins PGR5 and PGRL1 (DalCorso et al., 2008; Tolleter et al., 2011; Hertle et al., 2013) and is termed PGR5/L1-dependent CEF. A second route for CEF includes an NADPH dehydrogenase that oxidizes NADPH (product of LEF) to NADP⁺, simultaneously reducing the PQ (Allen, 2003; Kramer et al., 2004; Rumeau et al., 2007). The reduced PQ pool is then oxidized by cyt b₆f, causing H⁺ translocation into the thylakoid lumen, followed by the transfer of electrons to P700⁺ via PC. In the green alga Chlamydomonas reinhardtii, this second route for CEF involves a type II NADPH dehydrogenase (NDA2; Jans et al., 2008; Desplats et al., 2009).Oxygenic photosynthetic organisms have inhabited the planet for approximately 3 billion years and have developed numerous strategies to acclimate to environmental fluctuations. These acclimation processes confer flexibility to the photosynthetic machinery, allowing it to adjust to changes in conditions that impact the metabolic/energetic state of the organism and, most importantly, the formation of reactive oxygen species that may damage the photosynthetic apparatus and other cellular components (Li et al., 2009). Several ways in which the photosynthetic apparatus adjusts to environmental fluctuations have been established. A well-studied acclimation process, nonphotochemical quenching (NPQ), reduces the excitation pressure on PSII when oxidized downstream electron acceptors are not available (Eberhard et al., 2008; Li et al., 2009; Erickson et al., 2015). Several processes constitute NPQ, as follows. (1) qT, which involves the physical movement of light-harvesting complexes (LHCs) from one photosystem to another (this is also designated state transitions; Rochaix, 2014). (2) qE, which involves thermal dissipation of the excitation energy. This energy-dependent process requires an elevated ΔpH and involves an LHC-like protein, LHCSR3 (in C. reinhardtii) or PSBS (in plants), as well as the accumulation of specific xanthophylls (mainly lutein in C. reinhardtii and zeaxanthin in plants; Niyogi et al., 1997b; Li et al., 2000, 2004; Peers et al., 2009). (3) qZ, which is energy independent and involves the accumulation of zeaxanthin (Dall’Osto et al., 2005; Nilkens et al., 2010). (4) qI, which promotes quenching following physical damage to PSII core subunits (Aro et al., 1993). Additional mechanisms that can impact LEF and CEF are the synthesis and degradation of pigment molecules, changes in levels of RC and antenna complexes, and the control of electron distribution between LEF and CEF as the energetic demands of the cell change (Allen, 2003; Kramer et al., 2004). In addition, electrons can be consumed by mitochondrial and chlororespiratory activities (Bennoun, 1982; Peltier and Cournac, 2002; Johnson et al., 2014; Bailleul et al., 2015). The latter mainly involves the plastid terminal oxidase PTOX2, which catalyzes the oxidation of the PQ pool and the reduction of oxygen and H⁺ to form water molecules (Houille-Vernes et al., 2011; Nawrocki et al., 2015).Photosynthetic processes also must be modulated as organisms experience changes in the levels of available nutrients (Grossman and Takahashi, 2001). The macronutrient nitrogen (N), which represents 3% to 5% of the dry weight of photosynthetic organisms, is required to synthesize many biological molecules (e.g. amino acids, nucleic acids, and various metabolites) and also participates in posttranslational modifications of proteins (e.g. S-nitrosylation; Romero-Puertas et al., 2013). Importantly, N is highly abundant in chloroplasts in the form of DNA, ribosomes, Chl, and polypeptides (e.g. Rubisco and LHCs; Evans, 1989; Raven, 2013). Furthermore, there is a strong integration between N and carbon assimilation. During N limitation under photoautotrophic conditions, the inability of the organism to synthesize amino acids and other N-containing molecules necessary for cell growth and division can feed back to inhibit both carbon fixation by the CBB cycle and electron transport processes and also can negatively impact the expression of genes encoding key CBB cycle enzymes (Terashima and Evans, 1988; Huppe and Turpin, 1994; Nunes-Nesi et al., 2010).C. reinhardtii is a well-established model organism in which to study photosynthesis and acclimation processes, including acclimation to nutrient limitation (Wykoff et al., 1998; Grossman and Takahashi, 2001; Moseley et al., 2006; Grossman et al., 2009; Terauchi et al., 2010; Aksoy et al., 2013). This unicellular alga grows rapidly as a photoheterotroph (on fixed carbon in the light) or as a heterotroph (on fixed carbon in the dark), has completely sequenced nuclear, chloroplast, and mitochondrial genomes, can be used for classical genetic analyses, and is haploid, which makes some aspects of molecular manipulation (e.g. the generation of knockout mutants) easier (Merchant et al., 2007; Blaby et al., 2014). In the past few years, there have been many studies on the ways in which C. reinhardtii responds to N deprivation (Bulté and Wollman, 1992; Blaby et al., 2013; Goodenough et al., 2014; Schmollinger et al., 2014; Wei et al., 2015; Juergens et al., 2015). Cells deprived of N under photoheterotrophic conditions (i.e. acetate as an external carbon source) minimize the use of N (referred to as N sparing) and induce mechanisms associated with scavenging N from both external and internal pools, all of which eventually lead to proteome modifications and an elevated carbon-N ratio (Schmollinger et al., 2014). Acclimation under photoheterotrophic conditions also causes dramatic modifications of cellular metabolism and energetics: photosynthesis is down-regulated at multiple levels, with a portion of its N content recycled (mainly Chl and polypeptides of the photosynthetic apparatus), while there is enhanced accumulation of mitochondrial complexes leading to increased respiratory activity (Schmollinger et al., 2014; Juergens et al., 2015). Additionally, while fixed carbon cannot be used for growth in the absence of N, it may be stored as starch and triacylglycerol (Work et al., 2010; Siaut et al., 2011; Davey et al., 2014; Goodenough et al., 2014).In contrast to the acclimation of photoheterotrophically grown C. reinhardtii to N deprivation, little is known about how the photosynthetic machinery in this alga adjusts in response to N deprivation under photoautotrophic conditions, when the cells absolutely require photosynthetic energy generation for maintenance. Specifically, we sought to understand how photosynthesis adjusts to metabolic restrictions that slow down the CBB cycle, which in turn could cause the accumulation of photoreductant, particularly NADPH, as the demand for electrons declines (Peltier and Schmidt, 1991; Rumeau et al., 2007). Based on analyses of mutants and the use of spectroscopic and fluorescence measurements, we established a critical role for NDA2 in the acclimation of C. reinhardtii to N deprivation under photoautotrophic conditions, including (1) an augmented capacity for alternative routes of electron utilization (which decrease the NADPH-NADP⁺ ratio) based on increased NDA2-dependent CEF and chlororespiration, and (2) elevated qE, which relies on the H⁺ gradient generated by NDA2-dependent CEF.     

Capturing Arabidopsis Root Architecture Dynamics with root-fit Reveals Diversity in Responses to Salinity

The plant root is the first organ to encounter salinity stress, but the effect of salinity on root system architecture (RSA) remains elusive. Both the reduction in main root (MR) elongation and the redistribution of the root mass between MRs and lateral roots (LRs) are likely to play crucial roles in water extraction efficiency and ion exclusion. To establish which RSA parameters are responsive to salt stress, we performed a detailed time course experiment in which Arabidopsis (Arabidopsis thaliana) seedlings were grown on agar plates under different salt stress conditions. We captured RSA dynamics with quadratic growth functions (root-fit) and summarized the salt-induced differences in RSA dynamics in three growth parameters: MR elongation, average LR elongation, and increase in number of LRs. In the ecotype Columbia-0 accession of Arabidopsis, salt stress affected MR elongation more severely than LR elongation and an increase in LRs, leading to a significantly altered RSA. By quantifying RSA dynamics of 31 different Arabidopsis accessions in control and mild salt stress conditions, different strategies for regulation of MR and LR meristems and root branching were revealed. Different RSA strategies partially correlated with natural variation in abscisic acid sensitivity and different Na⁺/K⁺ ratios in shoots of seedlings grown under mild salt stress. Applying root-fit to describe the dynamics of RSA allowed us to uncover the natural diversity in root morphology and cluster it into four response types that otherwise would have been overlooked.Salt stress is known to affect plant growth and productivity as a result of its osmotic and ionic stress components. Osmotic stress imposed by salinity is thought to act in the early stages of the response, by reducing cell expansion in growing tissues and causing stomatal closure to minimize water loss. The build-up of ions in photosynthetic tissues leads to toxicity in the later stages of salinity stress and can be reduced by limiting sodium transport into the shoot tissue and compartmentalization of sodium ions into the root stele and vacuoles (Munns and Tester, 2008). The effect of salt stress on plant development was studied in terms of ion accumulation, plant survival, and signaling (Munns et al., 2012; Hasegawa, 2013; Pierik and Testerink, 2014). Most studies focus on traits in the aboveground tissues, because minimizing salt accumulation in leaf tissue is crucial for plant survival and its productivity. This approach has led to the discovery of many genes underlying salinity tolerance (Munns and Tester, 2008; Munns et al., 2012; Hasegawa, 2013; Maathuis, 2014). Another way to estimate salinity stress tolerance is by studying the rate of main root (MR) elongation of seedlings transferred to medium supplemented with high salt concentration. This is how Salt Overly Sensitive mutants were identified, being a classical example of genes involved in salt stress signaling and tolerance (Hasegawa, 2013; Maathuis, 2014). The success of this approach is to be explained by the important role that the root plays in salinity tolerance. Roots not only provide anchorage and ensure water and nutrient uptake, but also act as a sensory system, integrating changes in nutrient availability, water content, and salinity to adjust root morphology to exploit available resources to the maximum capacity (Galvan-Ampudia et al., 2013; Gruber et al., 2013). Understanding the significance of environmental modifications of root system architecture (RSA) for plant productivity is one of the major challenges of modern agriculture (de Dorlodot et al., 2007; Den Herder et al., 2010; Pierik and Testerink, 2014).The RSA of dicotyledonous plants consists of an embryonically derived MR and lateral roots (LRs) that originate from xylem pole pericycle cells of the MR, or from LRs in the case of higher-order LRs. Root growth and branching is mainly guided through the antagonistic action of two plant hormones: auxin and cytokinins (Petricka et al., 2012). Under environmental stress conditions, the synthesis of abscisic acid (ABA), ethylene, and brassinosteroids is known to be induced and to modulate the growth of MRs and LRs (Achard et al., 2006; Osmont et al., 2007; Achard and Genschik, 2009; Duan et al., 2013; Geng et al., 2013). In general, lower concentrations of salt were observed to slightly induce MR and LR elongation, whereas higher concentrations resulted in decreased growth of both MRs and LRs (Wang et al., 2009; Zolla et al., 2010). The reduction of growth is a result of the inhibition of cell cycle progression and a reduction in root apical meristem size (West et al., 2004). However, conflicting results were presented for the effect of salinity on lateral root density (LRD; Wang et al., 2009; Zolla et al., 2010; Galvan-Ampudia and Testerink, 2011). Some studies suggest that mild salinity enhances LR initiation or emergence events, thereby affecting patterning, whereas other studies imply that salinity arrests LR development. The origin of those contradictory observations could be attributable to studying LR initiation and density at single time points, rather than observing the dynamics of LR development, because LR formation changes as a function of root growth rate (De Smet et al., 2012). The dynamics of LR growth and development were characterized previously for the MR region formed before the salt stress exposure, identifying the importance of ABA in early growth arrest of postemerged LRs in response to salt stress (Duan et al., 2013). The effect of salt on LR emergence and initiation was found to differ for MR regions formed prior and subsequent to salinity exposure (Duan et al., 2013), consistent with LR patterning being determined at the root tip (Moreno-Risueno et al., 2010). Yet the effect of salt stress on the reprogramming of the entire RSA on a longer timescale remains elusive.Natural variation in Arabidopsis (Arabidopsis thaliana) is a great source for dissecting the genetic components underlying phenotypic diversity (Trontin et al., 2011; Weigel, 2012). Genes underlying phenotypic plasticity of RSA to environmental stimuli were also found to have high allelic variation leading to differences in root development between different Arabidopsis accessions (Rosas et al., 2013). Supposedly, genes responsible for phenotypic plasticity of the root morphology to different environmental conditions are under strong selection for adaptation to local environments. Various populations of Arabidopsis accessions were used to study natural variation in ion accumulation and salinity tolerance (Rus et al., 2006; Jha et al., 2010; Katori et al., 2010; Roy et al., 2013). In addition, a number of studies focusing on the natural variation in RSA have been published, identifying quantitative trait loci and allelic variation for genes involved in RSA development under control conditions (Mouchel et al., 2004; Meijón et al., 2014) and nutrient-deficient conditions (Chevalier et al., 2003; Gujas et al., 2012; Gifford et al., 2013; Kellermeier et al., 2013; Rosas et al., 2013). Exploring natural variation not only expands the knowledge of genes and molecular mechanisms underlying biological processes, but also provides insight on how plants adapt to challenging environmental conditions (Weigel, 2012) and whether the mechanisms are evolutionarily conserved. The early growth arrest of newly emerged LRs upon exposure to salt stress was observed to be conserved among the most commonly used Arabidopsis accessions Columbia-0 (Col-0), Landsberg erecta, and Wassilewskija (Ws; Duan et al., 2013). By studying salt stress responses of the entire RSA and a wider natural variation in root responses to stress, one could identify new morphological traits that are under environmental selection and possibly contribute to stress tolerance.In this work, we not only identify the RSA components that are responsive to salt stress, but we also describe the natural variation in dynamics of salt-induced changes leading to redistribution of root mass and different root morphology. The growth dynamics of MRs and LRs under different salt stress conditions were described by fitting a set of quadratic growth functions (root-fit) to individual RSA components. Studying salt-induced changes in RSA dynamics of 31 Arabidopsis accessions revealed four major strategies conserved among the accessions. Those four strategies were due to differences in salt stress sensitivity of individual RSA components (i.e. growth rates of MRs and LRs, and increases in the number of emerged LRs). This diversity in root morphology responses caused by salt stress was observed to be partially associated with differences in ABA, but not ethylene sensitivity. In addition, we observed that a number of accessions exhibiting a relatively strong inhibition of LR elongation showed a smaller increase in the Na⁺/K⁺ ratio in shoot tissue after exposure to salt stress. Our results imply that different RSA strategies identified in this study reflect diverse adaptations to different soil conditions and thus might contribute to efficient water extraction and ion compartmentalization in their native environments.     

The Rubisco Small Subunit Is Involved in Tobamovirus Movement and Tm-22-Mediated Extreme Resistance

The multifunctional movement protein (MP) of Tomato mosaic tobamovirus (ToMV) is involved in viral cell-to-cell movement, symptom development, and resistance gene recognition. However, it remains to be elucidated how ToMV MP plays such diverse roles in plants. Here, we show that ToMV MP interacts with the Rubisco small subunit (RbCS) of Nicotiana benthamiana in vitro and in vivo. In susceptible N. benthamiana plants, silencing of NbRbCS enabled ToMV to induce necrosis in inoculated leaves, thus enhancing virus local infectivity. However, the development of systemic viral symptoms was delayed. In transgenic N. benthamiana plants harboring Tobacco mosaic virus resistance-2² (Tm-2²), which mediates extreme resistance to ToMV, silencing of NbRbCS compromised Tm-2²-dependent resistance. ToMV was able to establish efficient local infection but was not able to move systemically. These findings suggest that NbRbCS plays a vital role in tobamovirus movement and plant antiviral defenses.Plant viruses use at least one movement protein (MP) to facilitate viral spread between plant cells via plasmodesmata (PD; Lucas and Gilbertson, 1994; Ghoshroy et al., 1997). Among viral MPs, the MP of tobamoviruses, such as Tobacco mosaic virus (TMV) and its close relative Tomato mosaic virus (ToMV), is the best characterized. TMV MP specifically accumulates in PD and modifies the plasmodesmatal size exclusion limit in mature source leaves or tissues (Wolf et al., 1989; Deom et al., 1990; Ding et al., 1992). TMV MP and viral genomic RNA form a mobile ribonucleoprotein complex that is essential for cell-to-cell movement of viral infection (Watanabe et al., 1984; Deom et al., 1987; Citovsky et al., 1990, 1992; Kiselyova et al., 2001; Kawakami et al., 2004; Waigmann et al., 2007). TMV MP also enhances intercellular RNA silencing (Vogler et al., 2008) and affects viral symptom development, host range, and host susceptibility to virus (Dardick et al., 2000; Bazzini et al., 2007). Furthermore, ToMV MP is identified as an avirulence factor that is recognized by tomato (Solanum lycopersicum) resistance proteins Tobacco mosaic virus resistance-2 (Tm-2) and Tm-2² (Meshi et al., 1989; Lanfermeijer et al., 2004). Indeed, tomato Tm-2² confers extreme resistance against TMV and ToMV in tomato plants and even in heterologous tobacco (Nicotiana tabacum) plants (Lanfermeijer et al., 2003, 2004).To date, several host factors that interact with TMV MP have been identified. These TMV MP-binding host factors include cell wall-associated proteins such as pectin methylesterase (Chen et al., 2000), calreticulin (Meshi et al., 1989), ANK1 (Ueki et al., 2010), and the cellular DnaJ-like protein MPIP1 (Shimizu et al., 2009). Many cytoskeletal components such as actin filaments (McLean et al., 1995), microtubules (Heinlein et al., 1995), and the microtubule-associated proteins MPB2C (Kragler et al., 2003) and EB1a (Brandner et al., 2008) also interact with TMV MP. Most of these factors are involved in TMV cell-to-cell movement.Rubisco catalyzes the first step of CO₂ assimilation in photosynthesis and photorespiration. The Rubisco holoenzyme is a heteropolymer consisting of eight large subunits (RbCLs) and eight small subunits (RbCSs). RbCL was reported to interact with the coat protein of Potato virus Y (Feki et al., 2005). Both RbCS and RbCL were reported to interact with the P3 proteins encoded by several potyviruses, including Shallot yellow stripe virus, Onion yellow dwarf virus, Soybean mosaic virus, and Turnip mosaic virus (Lin et al., 2011). Proteomic analysis of the plant-virus interactome revealed that RbCS participates in the formation of virus complexes of Rice yellow mottle virus (Brizard et al., 2006). However, the biological function of Rubisco in viral infection remains unknown.In this study, we show that RbCS plays an essential role in virus movement, host susceptibility, and Tm-2²-mediated extreme resistance in the ToMV-host plant interaction.     

Involvement of the Sieve Element Cytoskeleton in Electrical Responses to Cold Shocks

This study dealt with the visualization of the sieve element (SE) cytoskeleton and its involvement in electrical responses to local cold shocks, exemplifying the role of the cytoskeleton in Ca²⁺-triggered signal cascades in SEs. High-affinity fluorescent phalloidin as well as immunocytochemistry using anti-actin antibodies demonstrated a fully developed parietal actin meshwork in SEs. The involvement of the cytoskeleton in electrical responses and forisome conformation changes as indicators of Ca²⁺ influx was investigated by the application of cold shocks in the presence of diverse actin disruptors (latrunculin A and cytochalasin D). Under control conditions, cold shocks elicited a graded initial voltage transient, ΔV₁, reduced by external La³⁺ in keeping with the involvement of Ca²⁺ channels, and a second voltage transient, ΔV₂. Cytochalasin D had no effect on ΔV₁, while ΔV₁ was significantly reduced with 500 nm latrunculin A. Forisome dispersion was triggered by cold shocks of 4°C or greater, which was indicative of an all-or-none behavior. Forisome dispersion was suppressed by incubation with latrunculin A. In conclusion, the cytoskeleton controls cold shock-induced Ca²⁺ influx into SEs, leading to forisome dispersion and sieve plate occlusion in fava bean (Vicia faba).It has been argued for a long time that sieve elements (SEs) are devoid of a cytoskeleton (Parthasarathy and Pesacreta, 1980; Thorsch and Esau, 1981; Evert, 1990), but more recent biochemical and cytological studies favor the opposite view. Actin as well as profilin were detected in phloem exudates of various monocot and dicot species (Schobert et al., 1998, 2000), while immunocytochemical tests showed the presence of actin and tubulin in phloem exudates of pumpkin (Cucurbita maxima; Kulikova and Puryaseva, 2002). Proteome analyses gave further credence to the occurrence of microfilaments in SEs in castor bean (Ricinus communis; profilin; Barnes et al., 2004), pumpkin (actin; Walz et al., 2004), canola (Brassica napus; actin, profilin1 and profilin2, actin-depolymerizing factor4; Giavalisco et al., 2006), and rice (Oryza sativa; actin1, actin-depolymerizing factor2, actin depolymerizing-factor3, and actin-depolymerizing factor6; Aki et al., 2008). Moreover, cytological evidence suggests residues of a cytoskeleton in SEs; fluorescent immunolabeling identified an actin/myosin system at the sieve plates (Chaffey and Barlow, 2002).Theoretical considerations also call for the presence of a cytoskeleton in SEs. Turnover and addressing of macromolecules (Fisher et al., 1992; Leineweber et al., 2000) requires a local distribution network in SEs. This function was attributed to an endoplasmic reticulum (ER) continuous to the ER strands running through pore plasmodesma units (Blackman et al., 1998) into the companion cells. Although such a mechanism is essentially conceivable, an interaction between the ER and cytoskeleton would provide a more conventional mode of intracellular distribution (Hepler et al., 1990; Boevink et al., 1998; Ueda et al., 2010; Yokota et al., 2011; Chen et al., 2012). Moreover, macromolecular trafficking through pore plasmodesma units (Lucas et al., 2001) was proposed to be executed by actin and myosin (Oparka, 2004), implying the presence of a cytoskeleton in SEs. Despite the massive circumstantial evidence, however, a complete cytoskeleton network and its spatial distribution in SEs have not been visually documented thus far.The existence of an SE cytoskeleton would raise questions regarding its task(s) in this highly specialized cell type. In other plant cells, the cytoskeleton was proposed to be engaged, among others, in ion channel operation and intracellular signaling (Trewavas and Malho, 1997; Mazars et al., 1997, and refs. therein; Thuleau et al., 1998; Örvar et al., 2000; Sangwan et al., 2001; Drøbak et al., 2004; Davies and Stankovic, 2006), as in animal cells (Janmey, 1998; Lange and Gartzke, 2006). For instance, K⁺ fluxes are regulated by actin dynamics (Hwang et al., 1997; Liu and Luan, 1998; Chérel, 2004), while Ca²⁺ influx into the cytoplasm appears to be mediated by voltage-dependent Ca²⁺-permeable channels associated with microtubules (Mazars et al., 1997; Thion et al., 1998) or by mechanosensitive channels possibly associated with microfilaments (Wang et al., 2004; Zhang et al., 2007).Both types of Ca²⁺-permeable channels probably reside in the SE plasma membrane (Knoblauch et al., 2001; Hafke et al., 2007, 2009; Furch et al., 2009), where they are likely involved in Ca²⁺-dependent systemic signaling (Furch et al., 2009; Hafke et al., 2009; van Bel et al., 2011; Hafke and van Bel, 2013). These channels are also putative initiators of Ca²⁺-induced signal transduction in SEs, leading to sieve-plate occlusion in response to local cold shocks (Thorpe et al., 2010). In fava bean (Vicia faba), Ca²⁺-dependent sieve tube occlusion by dispersion of special phloem-specific proteins (P-proteins) known as forisomes has been studied intensely (Knoblauch et al., 2001; Furch et al., 2007, 2009; Thorpe et al., 2010). Thus, apart from its distributive tasks, a cytoskeleton may be of major importance for intracellular signaling cascades in the highly specialized, sparsely equipped SEs.Our objective was to investigate the existence and spatial distribution of an SE cytoskeleton and its engagement in local signaling through Ca²⁺ influx brought about by cold shocks. This study dealt with the visualization of cytoskeletal components in intact sieve tubes using microinjection of fluorescent phalloidin and immunocytochemistry. Confocal laser-scanning micrography (CLSM) and transmission electron microscopy unequivocally showed a parietally located cylindrical actin meshwork. We demonstrated the engagement of the network in local cold shock-induced electrical responses and its association with Ca²⁺ influx, since we found effects of the Ca²⁺ channel blocker La³⁺ and of the cytoskeleton disruptor latrunculin A (LatA) on electrical signatures triggered by cold shocks and, by consequence, on forisome conformation changes.     

Alternative Splicing-Mediated Targeting of the Arabidopsis GLUTAMATE RECEPTOR3.5 to Mitochondria Affects Organelle Morphology

Since the discovery of 20 genes encoding for putative ionotropic glutamate receptors in the Arabidopsis (Arabidopsis thaliana) genome, there has been considerable interest in uncovering their physiological functions. For many of these receptors, neither their channel formation and/or physiological roles nor their localization within the plant cells is known. Here, we provide, to our knowledge, new information about in vivo protein localization and give insight into the biological roles of the so-far uncharacterized Arabidopsis GLUTAMATE RECEPTOR3.5 (AtGLR3.5), a member of subfamily 3 of plant glutamate receptors. Using the pGREAT vector designed for the expression of fusion proteins in plants, we show that a splicing variant of AtGLR3.5 targets the inner mitochondrial membrane, while the other variant localizes to chloroplasts. Mitochondria of knockout or silenced plants showed a strikingly altered ultrastructure, lack of cristae, and swelling. Furthermore, using a genetically encoded mitochondria-targeted calcium probe, we measured a slightly reduced mitochondrial calcium uptake capacity in the knockout mutant. These observations indicate a functional expression of AtGLR3.5 in this organelle. Furthermore, AtGLR3.5-less mutant plants undergo anticipated senescence. Our data thus represent, to our knowledge, the first evidence of splicing-regulated organellar targeting of a plant ion channel and identify the first cation channel in plant mitochondria from a molecular point of view.In vertebrates, ionotropic glutamate receptors (iGluRs in animals) are ligand-gated cation channels that mediate the majority of the excitatory neurotransmission in the central nervous system (Dingledine et al., 1999). In the model plant Arabidopsis (Arabidopsis thaliana), 20 genes encoding homologs of animal iGluRs have been identified (Lam et al., 1998). According to phylogenetic analyses, the Arabidopsis GLUTAMATE RECEPTOR (AtGLR) homologs can be subdivided into three separate subgroups (Chiu et al., 1999, 2002). Some evidence for the channel-forming ability by plant ionotropic glutamate receptors (iGLRs) has been obtained only recently, and only for AtGLR3.4 and AtGLR1.4 expressed in heterologous systems (Vincill et al., 2012; Tapken et al., 2013). Studies with transgenic plants suggested roles of members of the plant GLR family in Ca²⁺ fluxes (AtGLR2; Kim et al., 2001), coordination of mitotic activity in the root apical meristem (Li et al., 2006), regulation of abscisic acid biosynthesis and water balance (AtGLR1.1; Kang and Turano, 2003; Kang et al., 2004), carbon/nitrogen sensing (AtGLR1.1; Kang and Turano, 2003), resistance against fungal infection (Kang et al., 2006), leaf-to-leaf wound signaling (Mousavi et al., 2013), and lateral root initiation (Vincill et al., 2013). Application of antagonists and agonists of animal iGluRs revealed that plant GLRs might be involved in the regulation of root growth and branching (Walch-Liu et al., 2006), in light signal transduction (Lam et al., 1998), and in the response to aluminum (Sivaguru et al., 2003). In various plant cell types, the agonists Glu- and Gly-induced plasma membrane depolarization and a rise in intracellular Ca²⁺ concentration that were inhibited by blockers of nonselective cation channels (NSCCs) and by antagonists of animal iGluRs (Dennison and Spalding, 2000; Dubos et al., 2003; Meyerhoff et al., 2005; Krol et al., 2007; Kwaaitaal et al., 2011; Michard et al., 2011). Furthermore, Glu-activated cation currents in patch-clamped root protoplasts were inhibited by NSCC blockers such as La³⁺ and Gd³⁺ (Demidchik et al., 2004). Therefore, it was proposed that plant iGLRs can form Ca²⁺-permeable NSCCs, are inhibited by animal iGluR antagonists, and might contribute to the shaping of plant Ca²⁺ signaling (McAinsh and Pittman, 2009). Studies using AtGLR3.3 mutant plants showed that intracellular Ca²⁺ rise and membrane depolarization induced by Glu in Arabidopsis hypocotyls and root cells are correlated with the presence of AtGLR3.3 (Qi et al., 2006; Stephens et al., 2008).However, most plant iGLRs, when expressed in heterologous systems, do not give rise to any current (e.g. in Xenopus spp. oocytes) or are toxic to host cells (e.g. in mammalian cells; Davenport, 2002). Recently, to examine whether AtGLR homologs possess functional ion channel domains, Tapken and Hollmann (2008) transplanted the pore loop together with two adjacent intracellular loops of 17 AtGLR subunits into two rat iGluR subunits and tested the resulting chimeric receptors for ion channel activity in the heterologous expression system Xenopus spp. oocyte. They showed that AtGLR1.1 and AtGLR1.4 have functional ion pore domains. The AtGLR1.1 pores are permeable to Na⁺, K⁺, and Ca²⁺ and are blocked by the nonspecific cation channel blocker La³⁺ (Tapken and Hollmann, 2008). Recent work has demonstrated that the expression of full-length AtGLR1.4 in oocytes gives rise to an amino acid-activated, nonselective, calcium-permeable channel that was found to be inhibited by the animal iGluR modulators 6,7-dinitroquinoxaline-2,3-dione and 6-cyano-7-nitroquinoxaline-2,3-dione (Tapken et al., 2013).The study of these channels has so far been restricted to those members that are located in the plasma membrane and were proved to be functional in the expression systems used. Instead, various localization prediction tools suggest that some of the plant GLRs might have chloroplast and mitochondrial targeting. In general, determining the subcellular localization of a protein is an important step toward understanding its function. We recently reported the localization of GLR3.4 to the inner chloroplast membrane (Teardo et al., 2011), which was also shown to harbor a 6,7-dinitroquinoxaline-2,3-dione-sensitive, calcium-permeable channel activity (Teardo et al., 2010). No other studies have addressed the eventual subcellular localization of other putative Glu receptors.In this work, we show that an isoform of GLR3.5 is efficiently targeted to the mitochondria. Functional expression of the channel in this organelle is indicated by the fact that its absence in knockout plants leads to a dramatically altered ultrastructure of mitochondria that impacts the plant physiology, ultimately leading to an anticipated senescence.