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.     

A Member of the Heavy Metal P-Type ATPase OsHMA5 Is Involved in Xylem Loading of Copper in Rice

Heavy metal-transporting P-type ATPase (HMA) has been implicated in the transport of heavy metals in plants. Here, we report the function and role of an uncharacterized member of HMA, OsHMA5 in rice (Oryza sativa). Knockout of OsHMA5 resulted in a decreased copper (Cu) concentration in the shoots but an increased Cu concentration in the roots at the vegetative stage. At the reproductive stage, the concentration of Cu in the brown rice was significantly lower in the mutants than in the wild-type rice; however, there was no difference in the concentrations of iron, manganese, and zinc between two independent mutants and the wild type. The Cu concentration of xylem sap was lower in the mutants than in the wild-type rice. OsHMA5 was mainly expressed in the roots at the vegetative stage but also in nodes, peduncle, rachis, and husk at the reproductive stage. The expression was up-regulated by excess Cu but not by the deficiency of Cu and other metals, including zinc, iron, and manganese, at the vegetative stage. Analysis of the transgenic rice carrying the OsHMA5 promoter fused with green fluorescent protein revealed that it was localized at the root pericycle cells and xylem region of diffuse vascular bundles in node I, vascular tissues of peduncle, rachis, and husk. Furthermore, immunostaining with an antibody against OsHMA5 revealed that it was localized to the plasma membrane. Expression of OsHMA5 in a Cu transport-defective mutant yeast (Saccharomyces cerevisiae) strain restored the growth. Taken together, OsHMA5 is involved in loading Cu to the xylem of the roots and other organs.Plants require nutrient elements to maintain normal growth and development. A number of different transporters, such as Cation Diffusion Facilitator, Natural resistance-associated macrophage protein, ATP-Binding Cassette, Zinc- and Iron-regulated-like Protein, and P-type ATPase, have been reported to be involved in the uptake, translocation, distribution, and homeostasis of nutrients (Hall and Williams, 2003; Krämer et al., 2007; Palmer and Guerinot, 2009). Among them, heavy metal-transporting P-type ATPase (HMA), the P_1B subfamily of the P-type ATPase superfamily, has been implicated in heavy metal transport (Williams and Mills, 2005; Grotz and Guerinot, 2006; Argüello et al., 2007; Burkhead et al., 2009). There are eight and nine members of P_1B-ATPase in Arabidopsis thaliana and rice (Oryza sativa), respectively (Williams and Mills, 2005). They are divided into two groups: zinc (Zn)/cadmium (Cd)/cobalt/lead (Pb) and copper (Cu)/silver transporters (Williams and Mills, 2005). AtHMA1 to AtHMA4 in A. thaliana and OsHMA1 to OsHMA3 in rice belong to the former group, while AtHMA5 to AtHMA8 and OsHMA4 to OsHMA9 belong to the latter group, although AtHMA1 has also been shown to transport Zn, Cu, and calcium (Axelsen and Palmgren, 2001; Williams and Mills, 2005; Seigneurin-Berny et al., 2006; Moreno et al., 2008; Kim et al., 2009).All members of HMAs in A. thaliana have been functionally characterized. AtHMA1 is involved in delivering Cu to the stroma, exporting Zn²⁺ from the chloroplast, or as a Ca²⁺/heavy metal transporter to the intracellular organelle (Seigneurin-Berny et al., 2006; Moreno et al., 2008; Kim et al., 2009). AtHMA2 and AtHMA4 localized at the pericycle are partially redundant and responsible for the release of Zn into the xylem (xylem loading) as well as Cd (Hussain et al., 2004; Verret et al., 2004; Wong and Cobbett, 2009; Wong et al., 2009), while AtHMA3 localized at the tonoplast plays a role in the detoxification of Zn/Cd/cobalt/Pb by mediating them into the vacuole (Morel et al., 2009; Chao et al., 2012). On the other hand, AtHMA5 is involved in the Cu translocation from roots to shoots or Cu detoxification of roots (Andrés-Colás et al., 2006; Kobayashi et al., 2008). AtHMA6 (PAA1, for P-type ATPase of Arabidopsis1) localized at the chloroplast periphery has been proposed to transport Cu over the chloroplast envelope, whereas AtHMA8 (PAA2) localized at the thylakoid membranes most likely transports Cu into the thylakoid lumen to supply plastocyanin (Shikanai et al., 2003; Abdel-Ghany et al., 2005). Finally, AtHMA7 (RESPONSIVE-TO-ANTAGONIST1) is responsible for delivering Cu to ethylene receptors and Cu homeostasis in the seedlings (Hirayama et al., 1999; Woeste and Kieber, 2000; Binder et al., 2010).By contrast, only three out of nine P-type ATPase members have been functionally characterized in rice. OsHMA2 was recently reported to be involved in the root-shoot translocation of Zn and Cd (Satoh-Nagasawa et al., 2012; Takahashi et al., 2012; Yamaji et al., 2013). Furthermore, OsHMA2 at the node is required for preferential distribution of Zn to young leaves and panicles (Yamaji et al., 2013). OsHMA3 is localized to the tonoplast of the root cells and responsible for the sequestration of Cd into the vacuoles (Ueno et al., 2010; Miyadate et al., 2011). On the other hand, OsHMA9 was mainly expressed in vascular tissues, including the xylem and phloem (Lee et al., 2007). The knockout lines accumulated more Zn, Cu, Pb, and Cd, suggesting its role in the efflux of these metals from the cells (Lee et al., 2007).Some members of P-type ATPase have also been identified in other plant species, including barley (Hordeum vulgare), wheat (Triticum aestivum), Thlaspi caerulescens (Noccaea caerulescens), and Arabidopsis halleri. HvHMA1 from barley might be involved in mobilizing Zn and Cu during the stage of grain filling (Mikkelsen et al., 2012). HvHMA2 from barley and TaHMA2 from wheat showed similar functions as OsHMA2 in rice (Mills et al., 2012; Tan et al., 2013). AhHMA3 in A. halleri, a Zn hyperaccumulator, is probably involved in high Zn accumulation (Becher et al., 2004; Chiang et al., 2006). Furthermore, AhHMA4 for Zn translocation showed a higher expression level (Chiang et al., 2006; Hanikenne et al., 2008). On the other hand, TcHMA3 from ecotype Ganges of T. caerulescens, a Cd hyperaccumulator, plays an important role in the detoxification of Cd by sequestering Cd into the vacuole of the leaves (Ueno et al., 2011). High expression of TcHMA4 (NcHMA4) was also reported in T. caerulescens (Bernard et al., 2004; Papoyan and Kochian, 2004; Craciun et al., 2012).In this study, we investigated the function and role of an uncharacterized member of P-type ATPase in rice, OsHMA5. We found that OsHMA5 is involved in the xylem loading of Cu at both the vegetative and reproductive growth stages.     

Dynamic Changes in the Distribution of Minerals in Relation to Phytic Acid Accumulation during Rice Seed Development

Phytic acid (inositol hexakisphosphate [InsP₆]) is the storage compound of phosphorus in seeds. As phytic acid binds strongly to metallic cations, it also acts as a storage compound of metals. To understand the mechanisms underlying metal accumulation and localization in relation to phytic acid storage, we applied synchrotron-based x-ray microfluorescence imaging analysis to characterize the simultaneous subcellular distribution of some mineral elements (phosphorus, calcium, potassium, iron, zinc, and copper) in immature and mature rice (Oryza sativa) seeds. This fine-imaging method can reveal whether these elements colocalize. We also determined their accumulation patterns and the changes in phosphate and InsP₆ contents during seed development. While the InsP₆ content in the outer parts of seeds rapidly increased during seed development, the phosphate contents of both the outer and inner parts of seeds remained low. Phosphorus, calcium, potassium, and iron were most abundant in the aleurone layer, and they colocalized throughout seed development. Zinc was broadly distributed from the aleurone layer to the inner endosperm. Copper localized outside the aleurone layer and did not colocalize with phosphorus. From these results, we suggest that phosphorus translocated from source organs was immediately converted to InsP₆ and accumulated in aleurone layer cells and that calcium, potassium, and iron accumulated as phytic acid salt (phytate) in the aleurone layer, whereas zinc bound loosely to InsP₆ and accumulated not only in phytate but also in another storage form. Copper accumulated in the endosperm and may exhibit a storage form other than phytate.The transport of nutrients into developing seeds has received considerable attention. During the grain-filling stage, plants remobilize and transport nutrients distributed throughout the vegetative source organs into seeds. Plant seeds contain large amounts of phosphorus (P) in organic form, which supports growth during the early stages of seedling development. Most of the P in seeds is stored in the form of phytic acid (inositol hexakisphosphate [InsP₆]). Seeds also accumulate mineral nutrients such as potassium (K), magnesium (Mg), calcium (Ca), iron (Fe), zinc (Zn), copper (Cu), and manganese (Mn), which are used in seedling growth. Phytic acid acts as a strong chelator of metal cations and binds them to form phytate, a salt of InsP₆ (Lott et al., 2002; Raboy, 2009). During germination, phytate is decationized and hydrolyzed by phytases, and then inorganic phosphates, inositol, and various minerals are released from the phytate (Loewus and Murthy, 2000). Phytate accumulates within protein bodies, generally of vacuolar origin, in seed storage cells and is usually concentrated in spherical inclusions called globoids. Many studies of the elemental composition of phytate in seeds have been published. Energy-dispersive x-ray microanalyses of many plant species have revealed that, other than P, globoids contain mainly K and Mg as well as low levels of Ca, Mn, Fe, and Zn (Lott, 1984; Lott et al., 1995; Wada and Lott, 1997). This indicates that phytate is a mixed salt of these cations.Whether all storage metal elements can bind equally to InsP₆ is not known, although most elements are thought to exist in seeds in the form of phytate (Raboy, 2009). To form phytate, P and the other elements must be present in the same place. Therefore, determination of the precise locations of P and other elements in seed tissues makes it possible to judge whether an element exists in the form of phytate. Differences in metal distribution with P might suggest a storage form other than phytate. For determining distributions, synchrotron-based x-ray microfluorescence (µ-XRF) imaging utilizing an x-ray microbeam is a powerful tool. The microbeam excites the elements, thereby revealing the details of their spatial distribution. The development of focusing optics for high-energy x-rays using a Kirkpatrick-Baez mirror raises the imaging resolution of elements in µ-XRF analysis. A focal spot size smaller than 1 µm with x-ray energy as high as 100 keV enables detection of the subcellular distribution of elements in plant tissues (Fukuda et al., 2008; Takahashi et al., 2009).Whether there is an order in the affinity of elements for phytic acid in plant cells remains unknown. The stability of InsP₆-metal complexes has been estimated by in vitro titration (Maddaiah et al., 1964; Vohra et al., 1965; Persson et al., 1998). The binding strength of InsP₆ with metal is stronger for Zn and Cu than for Fe, Mn, and Ca. We also do not know if the mineral composition of phytate in seeds is determined by the relative abundance of these elements in the seed or by their biochemical characteristics. As a first step to address these issues, we examined the simultaneous changes in the distribution of P and metal elements during seed development using µ-XRF imaging analysis.Our objective in this study was to observe the dynamic changes in the distribution of some nutritionally important minerals (P, Ca, K, Fe, Zn, and Cu) in relation to the accumulation of phytic acid during rice (Oryza sativa) seed development.     

Temperature Responses of C4 Photosynthesis: Biochemical Analysis of Rubisco,Phosphoenolpyruvate Carboxylase,and Carbonic Anhydrase in Setaria viridis

The photosynthetic assimilation of CO₂ in C₄ plants is potentially limited by the enzymatic rates of Rubisco, phosphoenolpyruvate carboxylase (PEPc), and carbonic anhydrase (CA). Therefore, the activity and kinetic properties of these enzymes are needed to accurately parameterize C₄ biochemical models of leaf CO₂ exchange in response to changes in CO₂ availability and temperature. There are currently no published temperature responses of both Rubisco carboxylation and oxygenation kinetics from a C₄ plant, nor are there known measurements of the temperature dependency of the PEPc Michaelis-Menten constant for its substrate HCO₃⁻, and there is little information on the temperature response of plant CA activity. Here, we used membrane inlet mass spectrometry to measure the temperature responses of Rubisco carboxylation and oxygenation kinetics, PEPc carboxylation kinetics, and the activity and first-order rate constant for the CA hydration reaction from 10°C to 40°C using crude leaf extracts from the C₄ plant Setaria viridis. The temperature dependencies of Rubisco, PEPc, and CA kinetic parameters are provided. These findings describe a new method for the investigation of PEPc kinetics, suggest an HCO₃⁻ limitation imposed by CA, and show similarities between the Rubisco temperature responses of previously measured C₃ species and the C₄ plant S. viridis.Biochemical models of photosynthesis are often used to predict the effect of environmental conditions on net rates of leaf CO₂ assimilation (Farquhar et al., 1980; von Caemmerer, 2000, 2013; Walker et al., 2013). With climate change, there is increased interest in modeling and understanding the effects of changes in temperature and CO₂ concentration on photosynthesis. The biochemical models of photosynthesis are primarily driven by the kinetic properties of the enzyme Rubisco, the primary carboxylating enzyme of the C₃ photosynthetic pathway, catalyzing the reaction of ribulose-1,5-bisphosphate (RuBP) with either CO₂ or oxygen. However, the CO₂-concentrating mechanism in C₄ photosynthesis utilizes carbonic anhydrase (CA) to help maintain the chemical equilibrium of CO₂ with HCO₃⁻ and phosphoenolpyruvate carboxylase (PEPc) to catalyze the carboxylation of phosphoenolpyruvate (PEP) with HCO₃⁻. These reactions ultimately provide the elevated levels of CO₂ to the compartmentalized Rubisco (Edwards and Walker, 1983). In C₄ plants, it has been demonstrated that PEPc, Rubisco, and CA can limit rates of CO₂ assimilation and influence the efficiency of the CO₂-concentrating mechanism (von Caemmerer, 2000; von Caemmerer et al., 2004; Studer et al., 2014). Therefore, accurate modeling of leaf photosynthesis in C₄ plants in response to future climatic conditions will require temperature parameterizations of Rubisco, PEPc, and CA kinetics from C₄ species.Modeling C₄ photosynthesis relies on the parameterization of both PEPc and Rubisco kinetics, making it more complex than for C₃ photosynthesis (Berry and Farquhar, 1978; von Caemmerer, 2000). However, the activity of CA is not included in these models, as it is assumed to be nonlimiting under most conditions (Berry and Farquhar, 1978; von Caemmerer, 2000). This assumption is implemented by modeling PEPc kinetics as a function of CO₂ partial pressure (pCO₂) and not HCO₃⁻ concentration, assuming CO₂ and HCO₃⁻ are in chemical equilibrium. However, there are questions regarding the amount of CA activity needed to sustain rates of C₄ photosynthesis and if CO₂ and HCO₃⁻ are in equilibrium (von Caemmerer et al., 2004; Studer et al., 2014).The most common steady-state biochemical models of photosynthesis are derived from the Michaelis-Menten models of enzyme activity (von Caemmerer, 2000), which are driven by the V_max and the K_m. Both of these parameters need to be further described by their temperature responses to be used to model photosynthesis in response to temperature. However, the temperature response of plant CA activity has not been completed above 17°C, and there is no known measured temperature response of K_m HCO₃⁻ for PEPc (K_P). Alternatively, Rubisco has been well studied, and there are consistent differences in kinetic values between C₃ and C₄ species at 25°C (von Caemmerer and Quick, 2000; Kubien et al., 2008), but the temperature responses, including both carboxylation and oxygenation reactions, have only been performed in C₃ species (Badger and Collatz, 1977; Jordan and Ogren, 1984; Bernacchi et al., 2001, 2002; Walker et al., 2013).Here, we present the temperature dependency of Rubisco carboxylation and oxygenation reactions, PEPc kinetics for HCO₃⁻, and CA hydration from 10°C to 40°C from the C₄ species Setaria viridis (succession no., A-010) measured using membrane inlet mass spectrometry. Generally, the 25°C values of the Rubisco parameters were similar to previous measurements of C₄ species. The temperature response of the maximum rate of Rubisco carboxylation (V_cmax) was high compared with most previous measurements from both C₃ and C₄ species, and the temperature response of the K_m for oxygenation (K_O) was low compared with most previously measured species. Taken together, the modeled temperature responses of Rubisco activity in S. viridis were similar to the previously reported temperature responses of some C₃ species. Additionally, the temperature response of the maximum rate of PEPc carboxylation (V_pmax) was similar to previous measurements. However, the temperature response of K_P was lower than what has been predicted (Chen et al., 1994). For CA, deactivation of the hydration activity was observed above 25°C. Additionally, models of CA and PEPc show that CA activity limits HCO₃⁻ availability to PEPc above 15°C, suggesting that CA limits PEP carboxylation rates in S. viridis when compared with the assumption that CO₂ and HCO₃⁻ are in full chemical equilibrium.     

The Operation of Two Decarboxylases,Transamination, and Partitioning of C4 Metabolic Processes between Mesophyll and Bundle Sheath Cells Allows Light Capture To Be Balanced for the Maize C4 Pathway

The C₄ photosynthesis carbon-concentrating mechanism in maize (Zea mays) has two CO₂ delivery pathways to the bundle sheath (BS; via malate or aspartate), and rates of phosphoglyceric acid reduction, starch synthesis, and phosphoenolpyruvate regeneration also vary between BS and mesophyll (M) cells. The theoretical partitioning of ATP supply between M and BS cells was derived for these metabolic activities from simulated profiles of light penetration across a leaf, with a potential 3-fold difference in the fraction of ATP produced in the BS relative to M (from 0.29 to 0.96). A steady-state metabolic model was tested using varying light quality to differentially stimulate M or BS photosystems. CO₂ uptake, ATP production rate (J_ATP; derived with a low oxygen/chlorophyll fluorescence method), and carbon isotope discrimination were measured on plants under a low light intensity, which is considered to affect C₄ operating efficiency. The light quality treatments did not change the empirical ATP cost of gross CO₂ assimilation (J_ATP/GA). Using the metabolic model, measured J_ATP/GA was compared with the predicted ATP demand as metabolic functions were varied between M and BS. Transamination and the two decarboxylase systems (NADP-malic enzyme and phosphoenolpyruvate carboxykinase) were critical for matching ATP and reduced NADP demand in BS and M when light capture was varied under contrasting light qualities.Interest in the C₄ pathway has been increased by the potential for enhancing crop productivity and maintaining yield stability in the face of global warming and population pressure (Friso et al., 2010; Zhu et al., 2010; Covshoff and Hibberd, 2012). Maize (Zea mays), a C₄ plant of the NADP-malic enzyme (ME) subtype, is a leading grain production cereal (www.fao.org). C₄ photosynthesis is a shared activity between mesophyll (M; abbreviations are listed in BS) cells, coupled to allow the operation of a biochemical carbon-concentrating mechanism (CCM). The CCM effectively minimizes photorespiration by increasing the CO₂ concentration in the bundle sheath (C_BS), where Rubisco is exclusively expressed. Since BS and M are connected by plasmodesmata, some CO₂ retrodiffuses. The refixation of that escaping CO₂ by the CCM increases the activity of the CCM and the total ATP demand (ATP_BS + ATP_M) for gross CO₂ assimilation (GA; [ATP_BS + ATP_M]/GA), from a theoretical minimum of five ATPs (Furbank et al., 1990). Leakiness (Φ), the amount of CO₂ retrodiffusing relative to phosphoenolpyruvate (PEP) carboxylation rate, is therefore a proxy for the coordination between the CCM and assimilatory activity (Henderson et al., 1992; Tazoe et al., 2008; Kromdijk et al., 2010; Ubierna et al., 2011; Bellasio and Griffiths, 2013).

Table I.

Variables and acronyms described in the text

Functional Characterization of Proanthocyanidin Pathway Enzymes from Tea and Their Application for Metabolic Engineering

A reevaluation of flux data for Arabidopsis mutants reveals that nitrate uptake through AtNRT1.1 conforms to a single low-affinity transport system that makes virtually no contribution to high-affinity nitrate uptake.In papers by Wang et al. (1998), Liu et al. (1999), and Liu and Tsay (2003), it was proposed that Arabidopsis thaliana Nitrate Transporter1.1 (AtNRT1.1; CHL1) encodes a dual-affinity nitrate transporter that “plays a major role in high-affinity nitrate uptake.” Here, we evaluate this concept by reexamining the uptake kinetics of Arabidopsis (Arabidopsis thaliana) mutant lines defective in NRT1.1 or other nitrate transporters.The uptake of inorganic ions by plant roots conforms to a pattern of biphasic kinetics. At low external ion concentration, ions are absorbed by saturable high-affinity transport systems (HATS), while at high concentrations, nonsaturating low-affinity transport systems (LATS) operate. Such is the case for K⁺, NH₄⁺, NO₃⁻, and ClO₃⁻ (a NO₃⁻ analog; Kochian and Lucas, 1982; Ullrich et al., 1984; Pace and McClure, 1986; Guy et al., 1988; Siddiqi et al., 1990; Aslam et al., 1992). The LATS for ³⁶ClO₃⁻ uptake was linear at [ClO₃⁻] down to 200 μm in tobacco (Nicotiana tabacum; Guy et al., 1988) and for nitrate uptake by barley (Hordeum vulgare) down to 100 μm NO₃⁻ (Aslam et al., 1992). These concentrations were the lowest examined by the latter authors. In the studies by Pace and McClure (1986), Guy et al. (1988), Siddiqi et al. (1990), and Aslam et al. (1992), LATS fluxes were extremely small at low external [NO₃⁻] and linear at both low and high [NO₃⁻].In barley, both constitutive HATS (CHATS) and inducible HATS (IHATS) were demonstrated at low [NO₃⁻], while a constitutive LATS (CLATS) failed to saturate even at 50 mm NO₃⁻ (Siddiqi et al., 1990). Likewise, CHATS and IHATS for nitrate have been demonstrated in Arabidopsis, as well as CLATS and inducible LATS (ILATS; Tsay et al., 1993; Huang et al., 1999).Doddema and Telkamp (1979) isolated an Arabidopsis B1 mutant that was defective in the LATS for nitrate (but not the HATS) by screening for survival on ClO₃⁻. Tsay et al. (1993) isolated the nitrate-inducible AtNRT1.1 gene that encodes the ILATS. Interestingly, Touraine and Glass (1997) were unable to detect reduced LATS or HATS influxes in AtNRT1.1 mutants grown on KNO₃, while Muños et al. (2004) reported increased HATS influx in AtNRT1.1 mutants. Likewise, Remans et al. (2006) failed to detect reduced uptake rates at low (0.5 mm) or high (10 mm) nitrate in AtNRT1.1 mutants.Among eukaryotes, genes encoding IHATS for nitrate were first isolated from Aspergillus nidulans (Unkles et al., 1991) and subsequently from Chlamydomonas reinhardtii (Quesada et al., 1994) and several higher plants (Glass, 2009), and based on the correlations between AtNRT2.1 expression and IHATS influx, it became accepted that IHATS was encoded by AtNRT2.1. This conclusion was supported by the demonstration that transfer DNA mutants disrupted in both AtNRT2.1 and AtNRT2.2 exhibited 67% reduction of HATS but no reduction in LATS function (Filleur et al., 2001). A gene encoding CHATS has not yet been identified, although a mutant with defective CHATS has been isolated (Wang and Crawford, 1996). In summary, it was held that in Arabidopsis, AtNRT2.1 was responsible for IHATS, while AtNRT1.1 and AtNRT1.2 encoded ILATS and CLATS, respectively (Forde, 2000; Li et al., 2007).In papers by Wang et al. (1998), Liu et al. (1999), and Liu and Tsay (2003), it was demonstrated that AtNRT1.1 mutants of Arabidopsis exhibited reduced nitrate uptake even at 10 μm nitrate. The authors concluded that AtNRT1.1 fluxes exhibited saturation kinetics in planta and in Xenopus laevis oocytes and proposed that NRT1.1 encodes a dual-affinity nitrate transporter that “plays a major role in high-affinity nitrate uptake” (Wang et al., 1998). Liu and Tsay (2003) demonstrated that the AtNRT1.1 protein was capable of switching between high- and low-affinity states by phosphorylation of Thr residue 101; under low-nitrogen (N) conditions, phosphorylation mediated via the activation of protein kinase CIPK23 generated a high-affinity transporter (Ho et al., 2009), whereas high-N favored the dephosphorylated low-affinity configuration.     

Coordination of Leaf Photosynthesis,Transpiration, and Structural Traits in Rice and Wild Relatives (Genus Oryza)

The genus Oryza, which includes rice (Oryza sativa and Oryza glaberrima) and wild relatives, is a useful genus to study leaf properties in order to identify structural features that control CO₂ access to chloroplasts, photosynthesis, water use efficiency, and drought tolerance. Traits, 26 structural and 17 functional, associated with photosynthesis and transpiration were quantified on 24 accessions (representatives of 17 species and eight genomes). Hypotheses of associations within, and between, structure, photosynthesis, and transpiration were tested. Two main clusters of positively interrelated leaf traits were identified: in the first cluster were structural features, leaf thickness (Thick_leaf), mesophyll (M) cell surface area exposed to intercellular air space per unit of leaf surface area (S_mes), and M cell size; a second group included functional traits, net photosynthetic rate, transpiration rate, M conductance to CO₂ diffusion (g_m), stomatal conductance to gas diffusion (g_s), and the g_m/g_s ratio. While net photosynthetic rate was positively correlated with g_m, neither was significantly linked with any individual structural traits. The results suggest that changes in g_m depend on covariations of multiple leaf (S_mes) and M cell (including cell wall thickness) structural traits. There was an inverse relationship between Thick_leaf and transpiration rate and a significant positive association between Thick_leaf and leaf transpiration efficiency. Interestingly, high g_m together with high g_m/g_s and a low S_mes/g_m ratio (M resistance to CO₂ diffusion per unit of cell surface area exposed to intercellular air space) appear to be ideal for supporting leaf photosynthesis while preserving water; in addition, thick M cell walls may be beneficial for plant drought tolerance.Leaves have evolved in different environments into a multitude of sizes and shapes, showing great variation in morphology and anatomy (Evans et al., 2004). However, all leaf typologies share common functions associated with chloroplasts, namely to intercept sunlight, take up CO₂ and inorganic nitrogen, and perform photosynthesis as a primary process for growth and reproduction.Investigating relationships between leaf anatomy and photosynthetic features (CO₂ fixation, which involves physical and biochemical processes and loss of water by transpiration) could lead to the identification of structural features for enhancing crop productivity and improve our understanding of plant evolution and adaptation (Evans et al., 2004).Stomata, through which CO₂ and water vapor diffuse into and out of the leaf, are involved in the regulation and control of photosynthetic and transpiration responses (Jarvis and Morison, 1981; Farquhar and Sharkey, 1982). Besides stomata distribution patterns between the abaxial and adaxial lamina surfaces (Foster and Smith, 1986), stomatal density and size are leaf anatomical traits contributing to build the leaf stomatal conductance to gas diffusion (g_s). This is calculated as the reciprocal of the stomatal resistances to gas diffusion; stomatal control results in a lower concentration of CO₂ in the leaf mesophyll (M) intercellular air space (C_i) than in the atmosphere (C_a; Nobel, 2009).Leaf M architecture greatly contributes to the pattern of light attenuation profiles within the lamina (Terashima and Saeki, 1983; Woolley, 1983; Vogelmann et al., 1989; Evans, 1999; Terashima et al., 2011) and affects CO₂ diffusion from the intercellular air space (IAS) to the chloroplast stroma. Therefore, it influences photosynthetic activity (Flexas et al., 2007, 2008) and can have effects on leaf hydrology and transpiration (Sack et al., 2003; Brodribb et al., 2010; Ocheltree et al., 2012). In addition, M architecture sets boundaries for leaf photosynthetic responses to changing environmental conditions (Nobel et al., 1975).Fortunately, several methodologies are currently available (Flexas et al., 2008; Pons et al., 2009) to determine M conductance to CO₂ diffusion (g_m), expressed per unit of leaf surface area. It is calculated as the reciprocal of the cumulated partial resistances exerted by leaf structural traits and biochemical processes from the substomatal cavities to photosynthetic sites (Evans et al., 2009; Nobel, 2009). The resistance to CO₂ diffusion in the liquid phase is 4 orders of magnitude higher than in the gaseous phase (Nobel, 2009); therefore, the changes in CO₂ concentration in the leaf gas phase are small in comparison with the changes in the liquid phase (Niinemets, 1999; Aalto and Juurola, 2002; Nobel, 2009). In the liquid phase, the resistance to CO₂ transfer is built from contributions by the cell walls, the plasmalemma, cytoplasm, chloroplast membranes, and stroma (Tholen and Zhu, 2011; Tholen et al., 2012); in addition, it involves factors associated with the carboxylation reaction (Kiirats et al., 2002; Evans et al., 2009). Thus, the concentration of CO₂ in the chloroplasts (C_c) is lower than C_i and can limit photosynthesis.At steady state, the relationships between the leaf net photosynthetic rate (A), the concentrations of CO₂, and the stomatal conductance to CO₂ diffusion (g_{s_CO2}) and g_m are modeled based on Fick’s first law of diffusion (Nobel, 2009) as:(1)where C_a, C_i, and C_c are as defined above (Flexas et al., 2008).The magnitude of g_m has been found to correlate with certain leaf structural traits in some species, in particular with the M cell surface area exposed to IAS per (one side) unit of leaf surface area (S_mes) and its extent covered by chloroplasts (S_chl; Evans and Loreto, 2000; Slaton and Smith, 2002; Tholen et al., 2012). From a physical modeling perspective, increasing S_mes provides more pathways acting in parallel for CO₂ diffusion (to and from the chloroplasts) per unit of leaf surface area; thus, it tends to reduce the resistance to CO₂ movement into the M cells and to increase g_m (Evans et al., 2009; Nobel, 2009). A number of leaf structural traits affect S_mes, including leaf thickness, cell density, cell volume and shape, and the fraction of the M cell walls in contact with the IAS (Terashima et al., 2001, 2011), and the degree they are linked to S_mes can vary between species (Slaton and Smith, 2002; Terashima et al., 2006). In particular, the presence of lobes on M cells, which are prominent in some Oryza species, may contribute to g_m through increasing S_mes (Sage and Sage, 2009; Terashima et al., 2011; Tosens et al., 2012). The M cell wall can provide resistance in series for M CO₂ diffusion (Nobel, 2009); thicker cell walls may increase resistance to CO₂ movement into the M cells and decrease g_m (Terashima et al., 2006, 2011; Evans et al., 2009).Other leaf traits, such as M porosity (the fraction of M volume occupied by air spaces [Vol_IAS]), has been shown to have a positive correlation with g_m in some species (Peña-Rojas et al., 2005), but the association may be mediated by light availability (Slaton and Smith, 2002). Leaf thickness (Thick_leaf) tends to be negatively linked to g_m, and it may set an upper limit for the maximum g_m, according to Terashima et al. (2006), Flexas et al. (2008), and Niinemets et al. (2009).With respect to leaf structural traits and water relations, Thick_leaf may increase the apoplast path length (resistances in series; Nobel, 2009) in the extra-xylem M (Sack and Holbrook, 2006; Brodribb et al., 2007) for water to reach the evaporation sites, which could decrease the conductance of water through the M and lower the transpiration rate. Interestingly, while thicker M cell walls may reduce g_m, they can enable the development of higher water potential gradients between the soil and leaves, which can be decisive for plant survival and longevity under drought conditions (Steppe et al., 2011).The purpose of this study was to provide insight into how the diversity of leaf structure relates to photosynthesis and transpiration among representative cultivated species and wild relatives in the genus Oryza. This includes, in particular, identifying leaf structural features associated with the diffusion of CO₂ from the atmosphere to the chloroplasts, photosynthesis, transpiration efficiency (A/E), and drought tolerance. The genus consists of 10 genomic groups and is composed of approximately 24 species (the number depending on taxonomic preferences; Kellogg, 2009; Brar and Singh, 2011), including the cultivated species Oryza sativa and Oryza glaberrima. Oryza species are distributed around the world, and they exhibit a wide range of phenotypes, with annual versus perennial life cycles and sun- versus shade-adapted species (Vaughan, 1994; Vaughan et al., 2008; Brar and Singh, 2011; Jagadish et al., 2011). This diversity in the genus is an important resource, which is being studied to improve rice yield, especially under unfavorable environmental conditions. In particular, O. glaberrima, Oryza australiensis, and Oryza meridionalis are of interest as drought-tolerant species (Henry et al., 2010; Ndjiondjop et al., 2010; Scafaro et al., 2011, 2012), while Oryza coarctata is salt tolerant (Sengupta and Majumder, 2010). In this study, a total of 43 leaf functional and structural parameters were collected on 24 accessions corresponding to 17 species within eight genomes (Brar and Singh (2011). Life cycle is as follows: A = annual; B = biennial; P = poliennial. Habitat is as follows: S = shade; S-Sh = sun-shade.

Last-Century Increases in Intrinsic Water-Use Efficiency of Grassland Communities Have Occurred over a Wide Range of Vegetation Composition,Nutrient Inputs,and Soil pH

Last-century climate change has led to variable increases of the intrinsic water-use efficiency (W_i; the ratio of net CO₂ assimilation to stomatal conductance for water vapor) of trees and C₃ grassland ecosystems, but the causes of the variability are not well understood. Here, we address putative drivers underlying variable W_i responses in a wide range of grassland communities. W_i was estimated from carbon isotope discrimination in archived herbage samples from 16 contrasting fertilizer treatments in the Park Grass Experiment, Rothamsted, England, for the 1915 to 1929 and 1995 to 2009 periods. Changes in W_i were analyzed in relation to nitrogen input, soil pH, species richness, and functional group composition. Treatments included liming as well as phosphorus and potassium additions with or without ammonium or nitrate fertilizer applications at three levels. W_i increased between 11% and 25% (P < 0.001) in the different treatments between the two periods. None of the fertilizers had a direct effect on the change of W_i (ΔW_i). However, soil pH (P < 0.05), species richness (P < 0.01), and percentage grass content (P < 0.01) were significantly related to ΔW_i. Grass-dominated, species-poor plots on acidic soils showed the largest ΔW_i (+14.7 μmol mol⁻¹). The ΔW_i response of these acidic plots was probably related to drought effects resulting from aluminum toxicity on root growth. Our results from the Park Grass Experiment show that W_i in grassland communities consistently increased over a wide range of nutrient inputs, soil pH, and plant community compositions during the last century.The intrinsic water-use efficiency (W_i) of plants is controlled by photosynthetic carbon assimilation and stomatal conductance via the leaf-level coupling of CO₂ and water fluxes. A general, but variable, increase of W_i under rising atmospheric CO₂ has been observed in long-term studies (Peñuelas et al., 2011; Franks et al., 2013; Saurer et al., 2014), but little is known about other environmental or ecosystem factors, which may interact with the effect of increasing CO₂ on W_i. An improved understanding of putative interactive mechanisms is important because changes in W_i may have significant effects on the global terrestrial carbon and water cycles (Gedney et al., 2006; Betts et al., 2007). This study explores the interactive effects of the increase in atmospheric CO₂ (observed over the last century), nutrient loading, and soil pH together with other related effects on plant species richness and functional group composition on the coupling of plant CO₂ and water fluxes in a seminatural grassland in southeastern England.W_i is a leaf-level efficiency that has also been termed potential water-use efficiency or physiological water-use efficiency, as it excludes the direct influence of vapor pressure deficit (VPD), a parameter determined by environmental conditions, on leaf-level water-use efficiency (Farquhar et al., 1989; Franks et al., 2013). W_i reports the relationship between net CO₂ assimilation rate (A_n) and stomatal conductance for water vapor (g_H2O):(1)According to the first law of Fick, A_n can be given as the product of the stomatal conductance for CO₂ (g_CO2) and the concentration gradient between the atmosphere (c_a) and the leaf internal gas space (c_i): A_n= g_CO2 (c_a – c_i). Using g_CO2 (c_a – c_i) instead of A_n in Equation 1, replacement of g_H2O/g_CO2 by the numerical value of g_H2O/g_CO2 (1.6) and rearrangement yields the following alternative expression of W_i:(2)Equation 2 reveals that past changes of W_i must have been controlled by two parameters: the change of c_a and the concurrent change of 1 – c_i/c_a, the relative gradient for CO₂ diffusion into the leaf (Franks et al., 2013). A change in the relative gradient is determined by the changes in A_n relative to g_H2O, as leaves respond to changing c_a and other environmental factors. In particular, Equation 2 shows that any variation in the climate change response of W_i is determined by the c_i/c_a response, if the comparison is made for vegetation at the same location and in the same period of time.Studies with C₃ vegetation, including trees/forests and C₃ grasslands, have revealed a general increase of W_i in the last century (Bert et al., 1997; Duquesnay et al., 1998; Feng, 1999; Arneth et al., 2002; Saurer et al., 2004; Barbosa et al., 2010; Köhler et al., 2010; Andreu-Hayles et al., 2011). In many cases, c_i/c_a, estimated by ¹³C discrimination (Farquhar et al., 1989), varied relatively little. Indeed, it has been suggested, based on theoretical grounds and empirical evidence from studies over geological/evolutionary to short time scales, that adaptive feedback responses will tend to maintain c_i/c_a approximately constant (Ehleringer and Cerling, 1995; Franks et al., 2013), as plants optimize carbon gain with respect to water loss (Cowan and Farquhar, 1977). Yet, c_i/c_a-dependent variation in the W_i response to climate change has also been noted (Peñuelas et al., 2011; Köhler et al., 2012) over the last century, indicating that additional factors, perhaps including other global change drivers, can modify the W_i response over this time scale, at least transiently. A meta-analysis by Peñuelas et al. (2011) reports c_i/c_a-dependent increases of W_i for different forests between 6% and 36% from the early 1960s to 2000s. A recent study by Saurer et al. (2014) on European forest trees found increases in W_i ranging from 1% to 53% during the last century. The strongest increase of W_i was recorded in regions where summer soil-water availability decreased in the last century. For different grassland communities, the c_i/c_a-dependent increases of W_i varied between 13% and 28% at one site (Köhler et al., 2012) from 1915 to 2009. Evidently, such variation can have important repercussions for the coupling of terrestrial CO₂ and water fluxes. Yet, little is known about the mechanism(s) underlying the variation.At the Park Grass Experiment (PGE) at Rothamsted, England, Köhler et al. (2012) observed a nitrogen supply-dependent enhancement of the W_i response on plots receiving nitrate fertilizer and maintained at a near-neutral soil pH by liming. However, the actual relationship between nitrogen supply and W_i response did not hold when the unlimed control (soil pH approximately 5.2) was included in the comparison. Remarkably, however, there was a significant positive relationship between the grass content of the community and the W_i response of the experimental plots in the investigation. These results suggested that the effect of nutrient supply on the W_i response of the grassland communities was indirect, perhaps working via effects on soil pH and/or vegetation composition (plant species richness or functional group composition).The PGE provides a unique opportunity to study century-scale variation in the c_i/c_a-dependent variation of W_i for a wide range of diverse grassland communities. Much of the extant ecosystem-scale variability of plant species richness and soil pH in temperate grasslands of Europe (Ceulemans et al., 2014) is included in the range of plot-scale plant species richness and soil pH at the PGE (which is reported in this investigation). The different long-term applications of fertilizer and lime over the past century have resulted in substantial changes in soil pH, species richness, and grass content on the experimental plots, but in most cases, within-plot changes over the study period considered here (1915–2009) were comparatively small (Crawley et al., 2005; Silvertown et al., 2006). All experimental plots are located at the same site and are exposed to the same weather conditions. Consequently, trends in climate as a direct driver for differences in W_i between plots can be ruled out.Here, we explore putative mechanisms underlying eventual c_i/c_a-dependent variation of W_i during the last century at the PGE by, first, quantifying the sustained effect of a wide range of contrasting fertilizer treatments (n = 16) on the change of W_i during the last century and, second, analyzing the relationships between the observed W_i response of treatments and the respective nutrient status, soil pH, plant species richness, and plant functional group composition of the grassland communities.     

Nitrogen Stress Affects the Turnover and Size of Nitrogen Pools Supplying Leaf Growth in a Grass

The effect of nitrogen (N) stress on the pool system supplying currently assimilated and (re)mobilized N for leaf growth of a grass was explored by dynamic ¹⁵N labeling, assessment of total and labeled N import into leaf growth zones, and compartmental analysis of the label import data. Perennial ryegrass (Lolium perenne) plants, grown with low or high levels of N fertilization, were labeled with ¹⁵NO₃⁻/¹⁴NO₃⁻ from 2 h to more than 20 d. In both treatments, the tracer time course in N imported into the growth zones fitted a two-pool model (r² > 0.99). This consisted of a “substrate pool,” which received N from current uptake and supplied the growth zone, and a recycling/mobilizing “store,” which exchanged with the substrate pool. N deficiency halved the leaf elongation rate, decreased N import into the growth zone, lengthened the delay between tracer uptake and its arrival in the growth zone (2.2 h versus 0.9 h), slowed the turnover of the substrate pool (half-life of 3.2 h versus 0.6 h), and increased its size (12.4 μg versus 5.9 μg). The store contained the equivalent of approximately 10 times (low N) and approximately five times (high N) the total daily N import into the growth zone. Its turnover agreed with that of protein turnover. Remarkably, the relative contribution of mobilization to leaf growth was large and similar (approximately 45%) in both treatments. We conclude that turnover and size of the substrate pool are related to the sink strength of the growth zone, whereas the contribution of the store is influenced by partitioning between sinks.This article examines the nitrogen (N) supply system of growing grass leaves, and it investigates how functional and kinetic properties of this system are affected by N stress. The N supply of growing leaves is a dominant target of whole-plant N metabolism. This is primarily related to the high N demand of the photosynthetic apparatus and the related metabolic machinery of new leaves (Evans, 1989; Makino and Osmond, 1991; Grindlay, 1997; Lemaire, 1997; Wright et al., 2004; Johnson et al., 2010; Maire et al., 2012). The N supply system, as defined here, is an integral part of the whole plant: it includes all N compounds that supply leaf growth. Hence, it integrates all events between the uptake of N from the environment (source), intermediate uses in other processes of plant N metabolism, and the eventual delivery to the leaf growth zone (sink; Fig. 1). N that does not ultimately serve leaf growth is not included in this system; all N that serves leaf growth is included, irrespective of its localization in the plant. Conceptually, two distinct sources supply N for leaf growth: N from current uptake and assimilation that is directly transferred to the growing leaf (“directly transferred N”) and N from turnover/redistribution of organic compounds (“mobilized N”).Open in a separate windowFigure 1.Schematic representation of N fluxes in the leaf growth zone and in the N supply system of leaf growth in a grass plant. A, Scheme of a growing leaf, with its growth zone (including zones of cell division, expansion, and maturation) and recently produced tissue (RPT). N import (I; μg h⁻¹) into the growth zone is mostly in the form of amino acids. Inside the growth zone, the nitrogenous substrate is used in new tissue construction. Then, N export (E; μg h⁻¹) is in the form of newly formed, fully expanded nitrogenous tissue (tissue-bound export with RPT) and is calculated as leaf elongation rate (L_ER; mm h⁻¹) times the lineal density of N in RPT (ρ; μg mm⁻¹): E = L_ER × ρ (Lattanzi et al., 2004). In a physiological steady state, import equals export (I = E) and the N content of the growth zone (G; μg [not shown]) is constant. Labeled N import into the growth zone (I_lab) commences shortly after labeling of the nutrient solution with ¹⁵N. The labeled N content of the growth zone (G_lab; μg) increases over time (dG_lab/dt) until it eventually reaches isotopic saturation (Fig. 2B). Similarly, the lineal density of labeled N in RPT (ρ_lab) increases until it approaches ρ. At any time, the export of labeled N in RPT (E_lab) equals the concurrent ρ_lab × L_ER. The import of labeled N is obtained as I_lab = E_lab + dG_lab/dt (Lattanzi et al., 2005) and considers the increasing label content in the growth zone during labeling. The fraction of labeled N in the import flux (f_{lab I}) is calculated as f_{lab I} = I_lab/I. The time course of f_{lab I} (Fig. 3) reflects the kinetic properties of the N supply system of leaf growth (C). B, Scheme of a vegetative grass plant (reduced to a rooted tiller with three leaves) with leaf growth zone. N import into the growth zone (I) originates from (1) N taken up from the nutrient solution that is transferred directly to the growth zone following assimilation (directly transferred N) and (2) N derived from turnover/redistribution of stores (mobilized N). The store potentially includes proteins in all mature and senescing tissue in the shoot and root of the entire plant. As xylem, phloem, and associated transfer cells/tissue provide for a vascular network that connects all parts of the plant, the mobilized N may principally originate from any plant tissue that exhibits N turnover/mobilization. The fraction of total N uptake that is allocated to the N supply system of the growth zone equals U (see model in C). The fraction of total mobilized N allocated to the growth zone equals M (see model in C). C, Compartmental model of the source-sink system supplying N to the leaf growth zone, as shown by Lattanzi et al. (2005) and used here. Newly absorbed N (U; μg h⁻¹) enters a substrate pool (Q₁); from there, the N is either imported directly into the growth zone (I) or exchanged with a store (Q₂). Q₁ integrates the steps of transport and assimilation that precede the translocation to the growth zone. Q₂ includes all proteins that supply N for leaf growth during their turnover and mobilization. The parameters of the model, including the (relative) size and turnover of pools Q₁ and Q₂, the deposition into the store (D; μg h⁻¹), and the mobilization from the store (M; μg h⁻¹), and the contribution of direct transfer relative to mobilization to the N supply of the growth zone are obtained by fitting the compartmental model to the f_{lab I} data (A) obtained in dynamic ¹⁵N labeling experiments (for details, see “Materials and Methods”). During physiological steady state, the sizes of Q₁ and Q₂ are constant, I = U, and M = D. [See online article for color version of this figure.]Amino acids are the predominant form in which N is supplied for leaf growth in grasses, and incorporation in new leaf tissue occurs mainly in the leaf growth zone (Gastal and Nelson, 1994; Amiard et al., 2004). This is a heterotrophic piece of tissue that includes the zones of cell division and elongation, is located at the base of the leaf, and is encircled by the sheath of the next older leaf (Volenec and Nelson, 1981; MacAdam et al., 1989; Schnyder et al., 1990; Kavanová et al., 2008). As most N is taken up in the form of nitrate but supplied to the growth zone in the form of amino acids, the path of directly transferred N includes a series of metabolic and transport steps. These include transfer to and loading into the xylem, xylem transport and unloading, reduction and ammonium assimilation, cycling through photorespiratory N pools, amino acid synthesis, loading into the phloem, and transport to the growth zone (Hirel and Lea, 2001; Novitskaya et al., 2002; Stitt et al., 2002; Lalonde et al., 2003; Dechorgnat et al., 2011). The time taken to pass through this sequence is unknown at present, as is the effect of N deficiency on that time. Also, it is not known how much N is contained in, and moving through, the different compartments that supply leaf growth with currently assimilated N.At the level of mature organs, mainly leaves, there is considerable knowledge about N turnover and redistribution. Much less is known about the fate of the mobilized N and its actual use in sink tissues like the leaf growth zone. The processes in mature organs are associated with the maintenance metabolism of proteins, organ senescence, and adjustments in leaf protein levels to decreasing irradiance inside growing canopies when leaves become shaded by overtopping newer ones (Evans, 1993; Vierstra, 1993; Hikosaka et al., 1994; Anten et al., 1995; Hirel et al., 2007; Jansson and Thomas, 2008; Moreau et al., 2012). N mobilization in shaded leaves supports the optimization of photosynthetic N use efficiency at plant and canopy scale (Field, 1983; Evans, 1993; Anten et al., 1995), it reduces the respiratory burden of protein maintenance costs (Dewar et al., 1998; Amthor, 2000; Cannell and Thornley, 2000), and it provides a mechanism for the conservation of the most frequently growth-limiting nutrient (Aerts, 1996). Mobilization of N involves protein turnover and net degradation (Huffaker and Peterson, 1974), redistribution in the form of amino acids (Simpson and Dalling, 1981; Simpson et al., 1983; Hörtensteiner and Feller, 2002), and (at least) some of the mobilized N is supplied to new leaf growth (Lattanzi et al., 2005).N fertilizer supply has multiple direct and indirect effects on plant N metabolism (Stitt et al., 2002; Schlüter et al., 2012). In particular, it modifies the N content of newly produced leaves, leaf longevity/senescence, and the dynamics of light distribution inside expanding canopies (Evans, 1983, 1989; Lötscher et al., 2003; Moreau et al., 2012). Thus, N fertilization influences the availability of recyclable N. At the same time, it augments the availability of directly transferable N to leaf growth. The net effect of these factors on the importance of mobilized versus directly transferred N substrate for leaf growth is not known. Also, it is unknown how N fertilization influences the functional characteristics of the N supply system, such as the size and turnover of its component pools.The assessment of the importance of directly transferred versus mobilized N for leaf growth requires studies at the sink end of the system (i.e. investigations of the N import flux into the leaf growth zone). Directly transferred N and mobilized N can be distinguished on the basis of their residence time in the plant, the time between uptake from the environment and import into the leaf growth zone: direct transfer involves a short residence time (fast transfer), whereas mobilized N resides much longer in the plant before it is delivered to the growth zone (slow transfer; De Visser et al., 1997; Lattanzi et al., 2005). Such studies require dynamic labeling of the N taken up by the plant (Schnyder and de Visser, 1999) and monitoring of the rate and isotopic composition/label content of N import into the leaf growth zone (Lattanzi et al., 2005). For grass plants in a physiological steady state, N import and the isotopic composition of the imported N are calculated from the leaf elongation rate and the lineal density of N in newly formed tissue (Fig. 1A; Lattanzi et al., 2004) and the change of tracer content in the leaf growth zone and recently produced leaf tissue over time (Lattanzi et al., 2005). Such data reveal the temporal change of the fraction of labeled N in the N import flux (f_{lab I}), which then can be used to characterize the N supply system of leaf growth via compartmental modeling. So far, there is only one study that has partially characterized this system (Lattanzi et al., 2005): this work was conducted with a C₃ grass, perennial ryegrass (Lolium perenne), and a C₄ grass, Paspalum dilatatum, growing in mixed stands and indicated that two interconnected N pools supplied the leaf growth zone in both species: a “substrate pool” (Q₁), which provided a direct route for newly absorbed and assimilated N import into the leaf growth zone (directly transferred N), and a mobilizing “store” (Q₂), which supplied N to the leaf growth zone via the substrate pool (Fig. 1C). The relative contribution of mobilization from the store was least important in the fast-growing, dominant individuals and most important in subordinate, shaded individuals. That work did not address the role of N deficiency, and the limited short-term resolution of the study (labeling intervals of 24 h or greater) precluded an analysis of the fast-moving parts of the system.Accordingly, this work addresses the following questions. How does N deficiency influence the substrate supply system of the leaf growth sink in terms of the number, size, and turnover (half-life) of its kinetically distinct pools? How does N deficiency affect the relationship between directly transferred and mobilized N for leaf growth? And what additional insight on the compartmental structure of the supply system is obtained when the short-term resolution of the analysis is increased by 1 order of magnitude? The work was performed with vegetative plants of perennial ryegrass grown in constant conditions with either a low (1.0 mm; termed low N) or high (7.5 mm; high N) nitrate concentration in the nutrient solution. In both treatments, a large number of plants were dynamically labeled with ¹⁵N over a wide range of time intervals (2 h to more than 20 d). The import of total N and ¹⁵N tracer into growth zones was estimated at the end of each labeling interval. Tracer data were analyzed with compartmental models following principles detailed by Lattanzi et al. (2005, 2012) and Lehmeier et al. (2008) to address the specific questions. Previous articles reported on root and shoot respiration (Lehmeier et al., 2010) and cell division and expansion in leaf growth zones (Kavanová et al., 2008) in the same experiment.     

Brassinosteroid Regulates Cell Elongation by Modulating Gibberellin Metabolism in Rice

Brassinosteroid (BR) and gibberellin (GA) are two predominant hormones regulating plant cell elongation. A defect in either of these leads to reduced plant growth and dwarfism. However, their relationship remains unknown in rice (Oryza sativa). Here, we demonstrated that BR regulates cell elongation by modulating GA metabolism in rice. Under physiological conditions, BR promotes GA accumulation by regulating the expression of GA metabolic genes to stimulate cell elongation. BR greatly induces the expression of D18/GA3ox-2, one of the GA biosynthetic genes, leading to increased GA₁ levels, the bioactive GA in rice seedlings. Consequently, both d18 and loss-of-function GA-signaling mutants have decreased BR sensitivity. When excessive active BR is applied, the hormone mostly induces GA inactivation through upregulation of the GA inactivation gene GA2ox-3 and also represses BR biosynthesis, resulting in decreased hormone levels and growth inhibition. As a feedback mechanism, GA extensively inhibits BR biosynthesis and the BR response. GA treatment decreases the enlarged leaf angles in plants with enhanced BR biosynthesis or signaling. Our results revealed a previously unknown mechanism underlying BR and GA crosstalk depending on tissues and hormone levels, which greatly advances our understanding of hormone actions in crop plants and appears much different from that in Arabidopsis thaliana.     

S-Nitrosylation of Ascorbate Peroxidase Is Part of Programmed Cell Death Signaling in Tobacco Bright Yellow-2 Cells

Nitric oxide (NO) is a small redox molecule that acts as a signal in different physiological and stress-related processes in plants. Recent evidence suggests that the biological activity of NO is also mediated by S-nitrosylation, a well-known redox-based posttranslational protein modification. Here, we show that during programmed cell death (PCD), induced by both heat shock (HS) or hydrogen peroxide (H₂O₂) in tobacco (Nicotiana tabacum) Bright Yellow-2 cells, an increase in S-nitrosylating agents occurred. NO increased in both experimentally induced PCDs, although with different intensities. In H₂O₂-treated cells, the increase in NO was lower than in cells exposed to HS. However, a simultaneous increase in S-nitrosoglutathione (GSNO), another NO source for S-nitrosylation, occurred in H₂O₂-treated cells, while a decrease in this metabolite was evident after HS. Consistently, different levels of activity and expression of GSNO reductase, the enzyme responsible for GSNO removal, were found in cells subjected to the two different PCD-inducing stimuli: low in H₂O₂-treated cells and high in the heat-shocked ones. Irrespective of the type of S-nitrosylating agent, S-nitrosylated proteins formed upon exposure to both of the PCD-inducing stimuli. Interestingly, cytosolic ascorbate peroxidase (cAPX), a key enzyme controlling H₂O₂ levels in plants, was found to be S-nitrosylated at the onset of both PCDs. In vivo and in vitro experiments showed that S-nitrosylation of cAPX was responsible for the rapid decrease in its activity. The possibility that S-nitrosylation induces cAPX ubiquitination and degradation and acts as part of the signaling pathway leading to PCD is discussed.Nitric oxide (NO) is a gaseous and diffusible redox molecule that acts as a signaling compound in both animal and plant systems (Pacher et al., 2007; Besson-Bard et al., 2008). In plants, NO has been found to play a key role in several physiological processes, such as germination, lateral root development, flowering, senescence, stomatal closure, and growth of pollen tubes (Beligni and Lamattina, 2000; Neill et al., 2002; Correa-Aragunde et al., 2004; He et al., 2004; Prado et al., 2004; Carimi et al., 2005). In addition, NO has been reported to be involved in plant responses to both biotic and abiotic stresses (Leitner et al., 2009; Siddiqui et al., 2011) and in the signaling pathways leading to programmed cell death (PCD; Delledonne et al., 1998; de Pinto et al., 2006; De Michele et al., 2009; Lin et al., 2012; Serrano et al., 2012).The cellular environment may greatly influence the chemical reactivity of NO, giving rise to different biologically active NO-derived compounds, collectively named reactive nitrogen species, which amplify and differentiate its ability to activate physiological and stress-related processes. Many of the biological properties of NO are due to its high affinity with transition metals of metalloproteins as well as its reactivity with reactive oxygen species (ROS; Hill et al., 2010). However, recent evidence suggests that protein S-nitrosylation, due to the addition of NO to reactive Cys thiols, may act as a key mechanism of NO signaling in plants (Wang et al., 2006; Astier et al., 2011). NO is also able to react with reduced glutathione (GSH), the most abundant cellular thiol, thus producing S-nitrosoglutathione (GSNO), which also acts as an endogenous trans-nitrosylating agent. GSNO is also considered as a NO store and donor and, as it is more stable than NO, acts as a long-distance NO transporter through the floematic flux (Malik et al., 2011). S-Nitrosoglutathione reductase (GSNOR), which is an enzyme conserved from bacteria to humans, has been suggested to play a role in regulating S-nitrosothiols (SNO) and the turnover of S-nitrosylated proteins in plants (Liu et al., 2001; Rusterucci et al., 2007).A number of proteins involved in metabolism, stress responses, and redox homeostasis have been identified as potential targets for S-nitrosylation in Arabidopsis (Arabidopsis thaliana; Lindermayr et al., 2005). During the hypersensitive response (HR), 16 proteins were identified to be S-nitrosylated in the seedlings of the same species (Romero-Puertas et al., 2008); in Citrus species, S-nitrosylation of about 50 proteins occurred in the NO-mediated resistance to high salinity (Tanou et al., 2009).However, while the number of candidate proteins for S-nitrosylation is increasing, the functional significance of protein S-nitrosylation has been explained only in a few cases, such as for nonsymbiotic hemoglobin (Perazzolli et al., 2004), glyceraldehyde 3-phosphate dehydrogenase (Lindermayr et al., 2005; Wawer et al., 2010), Met adenosyltransferase (Lindermayr et al., 2006), and metacaspase9 (Belenghi et al., 2007). Of particular interest are the cases in which S-nitrosylation involves enzymes controlling ROS homeostasis. For instance, it has been reported that S-nitrosylation of peroxiredoxin IIE regulates the antioxidant function of this enzyme and might contribute to the HR (Romero-Puertas et al., 2007). It has also been shown that in the immunity response, S-nitrosylation of NADPH oxidase inactivates the enzyme, thus reducing ROS production and controlling HR development (Yun et al., 2011).Recently, S-nitrosylation has also been shown to be involved in PCD of nitric oxide excess1 (noe1) rice (Oryza sativa) plants, which are mutated in the OsCATC gene coding for catalase (Lin et al., 2012). In these plants, which show PCD-like phenotypes under high-light conditions, glyceraldehyde 3-phosphate dehydrogenase and thioredoxin are S-nitrosylated. This suggests that the NO-dependent regulation of these proteins is involved in plant PCD, similar to what occurs in animal apoptosis (Sumbayev, 2003; Hara et al., 2005; Lin et al., 2012). The increase in hydrogen peroxide (H₂O₂) after exposure to high light in noe1 plants is responsible for the production of NO required for leaf cell death induction (Lin et al., 2012). There is a strict relationship between H₂O₂ and NO in PCD activation (Delledonne et al., 2001; de Pinto et al., 2002); however, the mechanism of this interplay is largely still unknown (for review, see Zaninotto et al., 2006; Zhao, 2007; Yoshioka et al., 2011). NO can induce ROS production and vice versa, and their reciprocal modulation in terms of intensity and timing seems to be crucial in determining PCD activation and in controlling HR development (Delledonne et al., 2001; Zhao, 2007; Yun et al., 2011).In previous papers, we demonstrated that heat shock (HS) at 55°C and treatment with 50 mm H₂O₂ promote PCD in tobacco (Nicotiana tabacum) Bright Yellow-2 (BY-2) cells (Vacca et al., 2004; de Pinto et al., 2006; Locato et al., 2008). In both experimental conditions, NO production and decrease in cytosolic ascorbate peroxidase (cAPX) were observed as early events in the PCD pathway, and cAPX decrease has been suggested to contribute to determining the redox environment required for PCD (de Pinto et al., 2006; Locato et al., 2008).In this study, the production of nitrosylating agents (NO and GSNO) in the first hours of PCD induction by HS or H₂O₂ treatment in tobacco BY-2 cells and their role in PCD were studied. The possibility that S-nitrosylation could be a first step in regulating cAPX activity and turnover as part of the signaling pathway leading to PCD was also investigated.     

Structural Basis of Efficient Electron Transport between Photosynthetic Membrane Proteins and Plastocyanin in Spinach Revealed Using Nuclear Magnetic Resonance

In the photosynthetic light reactions of plants and cyanobacteria, plastocyanin (Pc) plays a crucial role as an electron carrier and shuttle protein between two membrane protein complexes: cytochrome b₆f (cyt b₆f) and photosystem I (PSI). The rapid turnover of Pc between cyt b₆f and PSI enables the efficient use of light energy. In the Pc-cyt b₆f and Pc-PSI electron transfer complexes, the electron transfer reactions are accomplished within <10⁻⁴ s. However, the mechanisms enabling the rapid association and dissociation of Pc are still unclear because of the lack of an appropriate method to study huge complexes with short lifetimes. Here, using the transferred cross-saturation method, we investigated the residues of spinach (Spinacia oleracea) Pc in close proximity to spinach PSI and cyt b₆f, in both the thylakoid vesicle–embedded and solubilized states. We demonstrated that the hydrophobic patch residues of Pc are in close proximity to PSI and cyt b₆f, whereas the acidic patch residues of Pc do not form stable salt bridges with either PSI or cyt b₆f, in the electron transfer complexes. The transient characteristics of the interactions on the acidic patch facilitate the rapid association and dissociation of Pc.     

Reactive Oxygen Species-Dependent Nitric Oxide Production Contributes to Hydrogen-Promoted Stomatal Closure in Arabidopsis

The signaling role of hydrogen gas (H₂) has attracted increasing attention from animals to plants. However, the physiological significance and molecular mechanism of H₂ in drought tolerance are still largely unexplored. In this article, we report that abscisic acid (ABA) induced stomatal closure in Arabidopsis (Arabidopsis thaliana) by triggering intracellular signaling events involving H₂, reactive oxygen species (ROS), nitric oxide (NO), and the guard cell outward-rectifying K⁺ channel (GORK). ABA elicited a rapid and sustained H₂ release and production in Arabidopsis. Exogenous hydrogen-rich water (HRW) effectively led to an increase of intracellular H₂ production, a reduction in the stomatal aperture, and enhanced drought tolerance. Subsequent results revealed that HRW stimulated significant inductions of NO and ROS synthesis associated with stomatal closure in the wild type, which were individually abolished in the nitric reductase mutant nitrate reductase1/2 (nia1/2) or the NADPH oxidase-deficient mutant rbohF (for respiratory burst oxidase homolog). Furthermore, we demonstrate that the HRW-promoted NO generation is dependent on ROS production. The rbohF mutant had impaired NO synthesis and stomatal closure in response to HRW, while these changes were rescued by exogenous application of NO. In addition, both HRW and hydrogen peroxide failed to induce NO production or stomatal closure in the nia1/2 mutant, while HRW-promoted ROS accumulation was not impaired. In the GORK-null mutant, stomatal closure induced by ABA, HRW, NO, or hydrogen peroxide was partially suppressed. Together, these results define a main branch of H₂-regulated stomatal movement involved in the ABA signaling cascade in which RbohF-dependent ROS and nitric reductase-associated NO production, and subsequent GORK activation, were causally involved.Stomata are responsible for leaves of terrestrial plants taking in carbon dioxide for photosynthesis and likewise regulate how much water plants evaporate through the stomatal pores (Chaerle et al., 2005). When experiencing water-deficient conditions, surviving plants balance photosynthesis with controlling water loss through the stomatal pores, which relies on turgor changes by pairs of highly differentiated epidermal cells surrounding the stomatal pore, called the guard cells (Haworth et al., 2011; Loutfy et al., 2012).Besides the characterization of the significant roles of abscisic acid (ABA) in regulating stomatal movement, the key factors in guard cell signal transduction have been intensively investigated by performing forward and reverse genetics approaches. For example, both reactive oxygen species (ROS) and nitric oxide (NO) have been identified as vital intermediates in guard cell ABA signaling (Bright et al., 2006; Yan et al., 2007; Suzuki et al., 2011; Hao et al., 2012). The key ROS-producing enzymes in Arabidopsis (Arabidopsis thaliana) guard cells are the respiratory burst oxidase homologs (Rboh) D and F (Kwak et al., 2003; Bright et al., 2006; Mazars et al., 2010; Marino et al., 2012). Current available data suggest that there are at least two distinct pathways responsible for NO synthesis involved in ABA signaling in guard cells: the nitrite reductase (NR)- and l-Arg-dependent pathways (Desikan et al., 2002; Besson-Bard et al., 2008). Genetic evidence further demonstrated that removal of the major known sources of either ROS or NO significantly impairs ABA-induced stomatal closure. ABA fails to induce ROS production in the atrbohD/F double mutant (Kwak et al., 2003; Wang et al., 2012) and NO synthesis in the NR-deficient mutant nitrate reductase1/2 (nia1/2; Bright et al., 2006; Neill et al., 2008), both of which lead to impaired stomatal closure in Arabidopsis. Most importantly, ROS and NO, which function both synergistically and independently, have been established as ubiquitous signal transduction components to control a diverse range of physiological pathways in higher plants (Bright et al., 2006; Tossi et al., 2012).The guard cell outward-rectifying K⁺ channel (GORK) encodes the exclusive voltage-gated outwardly rectifying K⁺ channel protein, which was located in the guard cell membrane (Ache et al., 2000; Dreyer and Blatt, 2009). Expression profiles revealed that this gene is up-regulated upon the onset of drought, salinity, and cold stress and ABA exposure (Becker et al., 2003; Tran et al., 2013). Reverse genetic evidence further showed that GORK plays an important role in the control of stomatal movements and allows the plant to reduce transpirational water loss significantly (Hosy et al., 2003) and participates in the regulation of salinity tolerance by preventing salt-induced K⁺ loss (Jayakannan et al., 2013). Due to the high complexity of guard cell signaling cascades, whether and how ABA-triggered GORK up-regulation is attributed to the generation of cellular secondary messengers, such as ROS and NO, is less clear.Hydrogen gas (H₂) was recently revealed as a signaling modulator with multiple biological functions in clinical trails (Ohsawa et al., 2007; Itoh et al., 2009; Ito et al., 2012). It was previously found that a hydrogenase system could generate H₂ in bacteria and green algae (Meyer, 2007; Esquível et al., 2011). Although some earlier studies discovered the evolution of H₂ in several higher plant species (Renwick et al., 1964; Torres et al., 1984), it was also proposed that the eukaryotic hydrogenase-like protein does not metabolize H₂ (Cavazza et al., 2008; Mondy et al., 2014). Since the explosion limit of H₂ gas is about 4% to 72.4% (v/v, in the air), the direct application of H₂ gas in experiments is flammable and dangerous. Regardless of these problems to be resolved, the methodology, such as using exogenous hydrogen-rich water (HRW) or hydrogen-rich saline, which is safe, economical, and easily available, provides a valuable approach to investigate the physiological function of H₂ in animal research and clinical trials. For example, hydrogen dissolved in Dulbecco’s modified Eagle’s medium was found to react with cytotoxic ROS and thus protect against oxidative damage in PC12 cells and rats (Ohsawa et al., 2007). The neuroprotective effect of H₂-loaded eye drops on retinal ischemia-reperfusion injury was also reported (Oharazawa et al., 2010). In plants, corresponding results by using HRW combined with gas chromatography (GC) revealed that H₂ could act as a novel beneficial gaseous molecule in plant responses against salinity (Xie et al., 2012; Xu et al., 2013), cadmium stress (Cui et al., 2013), and paraquat toxicity (Jin et al., 2013). More recently, the observation that HRW could delay the postharvest ripening and senescence of kiwifruit (Actinidia deliciosa) was reported (Hu et al., 2014).Considering the fact that the signaling cascades for salt, osmotic, and drought stresses share a common cascade in an ABA-dependent pathway, it would be noteworthy to identify whether and how H₂ regulates the bioactivity of ABA-induced downstream components and, thereafter, biological responses, including stomatal closure and drought tolerance. To resolve these scientific questions, rbohD, rbohF, nia1/2, nitric oxide associated1 (noa1; Van Ree et al., 2011), nia1/2/noa1, and gork mutants were utilized to investigate the relationship among H₂, ROS, NO, and GORK in the guard cell signal transduction network. By the combination of pharmacological and biochemical analyses with this genetics-based approach, we provide comprehensive evidence to show that H₂ might be a newly identified bioeffective modulator involved in ABA signaling responsible for drought tolerance, that HRW-promoted stomatal closure was mainly attributed to the modulation of ROS-dependent NO generation, and that GORK might be the downstream target protein of H₂ signaling.     

Focus Issue on Calcium Signaling: Calcium-Dependent Protein Kinase CPK6 Positively Functions in Induction by Yeast Elicitor of Stomatal Closure and Inhibition by Yeast Elicitor of Light-Induced Stomatal Opening in Arabidopsis

Yeast elicitor (YEL) induces stomatal closure that is mediated by a Ca²⁺-dependent signaling pathway. A Ca²⁺-dependent protein kinase, CPK6, positively regulates activation of ion channels in abscisic acid and methyl jasmonate signaling, leading to stomatal closure in Arabidopsis (Arabidopsis thaliana). YEL also inhibits light-induced stomatal opening. However, it remains unknown whether CPK6 is involved in induction by YEL of stomatal closure or in inhibition by YEL of light-induced stomatal opening. In this study, we investigated the roles of CPK6 in induction by YEL of stomatal closure and inhibition by YEL of light-induced stomatal opening in Arabidopsis. Disruption of CPK6 gene impaired induction by YEL of stomatal closure and inhibition by YEL of light-induced stomatal opening. Activation by YEL of nonselective Ca²⁺-permeable cation channels was impaired in cpk6-2 guard cells, and transient elevations elicited by YEL in cytosolic-free Ca²⁺ concentration were suppressed in cpk6-2 and cpk6-1 guard cells. YEL activated slow anion channels in wild-type guard cells but not in cpk6-2 or cpk6-1 and inhibited inward-rectifying K⁺ channels in wild-type guard cells but not in cpk6-2 or cpk6-1. The cpk6-2 and cpk6-1 mutations inhibited YEL-induced hydrogen peroxide accumulation in guard cells and apoplast of rosette leaves but did not affect YEL-induced hydrogen peroxide production in the apoplast of rosette leaves. These results suggest that CPK6 positively functions in induction by YEL of stomatal closure and inhibition by YEL of light-induced stomatal opening in Arabidopsis and is a convergent point of signaling pathways for stomatal closure in response to abiotic and biotic stress.Stomata, formed by pairs of guard cells, play a critical role in regulation of plant CO₂ uptake and water loss, thus critically influencing plant growth and water stress responsiveness. Guard cells respond to a variety of abiotic and biotic stimuli, such as light, drought, and pathogen attack (Israelsson et al., 2006; Shimazaki et al., 2007; Melotto et al., 2008).Elicitors derived from microbial surface mimic pathogen attack and induce stomatal closure in various plant species such as Solanum lycopersicum (Lee et al., 1999), Commelina communis (Lee et al., 1999), Hordeum vulgare (Koers et al., 2011), and Arabidopsis (Arabidopsis thaliana; Melotto et al., 2006; Khokon et al., 2010). Yeast elicitor (YEL) induces stomatal closure in Arabidopsis (Klüsener et al., 2002; Khokon et al., 2010; Salam et al., 2013). Our recent studies showed that YEL inhibits light-induced stomatal opening and that protein phosphorylation is involved in induction by YEL of stomatal closure and inhibition by YEL of light-induced stomatal opening (Salam et al., 2013).Cytosolic Ca²⁺ has long been recognized as a conserved second messenger in stomatal movement (Shimazaki et al., 2007; Roelfsema and Hedrich 2010; Hubbard et al., 2012). Elevation of cytosolic free Ca²⁺ concentration ([Ca2+]_cyt) is triggered by influx of Ca²⁺ from apoplast and release of Ca²⁺ from intracellular stores in guard cell signaling (Leckie et al., 1998; Hamilton et al., 2000; Pei et al., 2000; Garcia-Mata et al., 2003; Lemtiri-Chlieh et al., 2003). The influx of Ca²⁺ is carried by nonselective Ca²⁺-permeable cation (I_Ca) channels that are activated by plasma membrane hyperpolarization and H₂O₂ (Pei et al., 2000; Murata et al., 2001; Kwak et al., 2003). Elevation of [Ca2+]_cyt activates slow anion (S-type) channels and down-regulates inward-rectifying potassium (K_in) channels in guard cells (Schroeder and Hagiwara, 1989; Grabov and Blatt, 1999). The activation of S-type channels is a hallmark of stomatal closure, and the suppression of K_in channels is favorable to stomatal closure but not to stomatal opening (Pei et al., 1997; Kwak et al., 2001; Xue et al., 2011; Uraji et al., 2012).YEL induces stomatal closure with extracellular H₂O₂ production, intracellular H₂O₂ accumulation, activation of I_Ca channels, and transient [Ca2+]_cyt elevations (Klüsener et al., 2002; Khokon et al., 2010). However, it remains to be clarified whether YEL activates S-type channels and inhibits K_in channels in guard cells.Calcium-dependent protein kinases (CDPKs) are regulators in Ca²⁺-dependent guard cell signaling (Mori et al., 2006; Zhu et al., 2007; Geiger et al., 2010, 2011; Zou et al., 2010; Munemasa et al., 2011; Brandt et al., 2012; Scherzer et al., 2012). In guard cells, CDPKs regulate activation of S-type and I_Ca channels and inhibition of K_in channels (Mori et al., 2006; Zou et al., 2010; Munemasa et al., 2011). A CDPK, CPK6, positively regulates activation of S-type channels and I_Ca channels without affecting H₂O₂ production in abscisic acid (ABA)- and methyl jasmonate (MeJA)-induced stomatal closure (Mori et al., 2006; Munemasa et al., 2011). CPK6 phosphorylates and activates SLOW ANION CHANNEL-ASSOCIATED1 expressed in Xenopus spp. oocyte (Brandt et al., 2012; Scherzer et al., 2012). These findings underline the role of CPK6 in regulation of ion channel activation and stomatal movement, leading us to test whether CPK6 regulates the induction by YEL of stomatal closure and inhibition by YEL of light-induced stomatal opening.In this study, we investigated activation of S-type channels and inhibition of K_in channels by YEL and roles of CPK6 in induction by YEL of stomatal closure and inhibition by YEL of light-induced stomatal opening. For this purpose, we examined the effects of mutation of CPK6 on induction by YEL of stomatal closure and inhibition by YEL of light-induced stomatal opening, activation of I_Ca channels, transient [Ca2+]_cyt elevations, activation of S-type channels, inhibition of K_in channels, H₂O₂ production in leaves, and H₂O₂ accumulation in leaves and guard cells.     

Focus on Water: Polarity of Water Transport across Epidermal Cell Membranes in Tradescantia virginiana

Using the automated cell pressure probe, small and highly reproducible hydrostatic pressure clamp (PC) and pressure relaxation (PR) tests (typically, applied step change in pressure = 0.02 MPa and overall change in volume = 30 pL, respectively) were applied to individual Tradescantia virginiana epidermal cells to determine both exosmotic and endosmotic hydraulic conductivity (L_pOUT and L_pIN, respectively). Within-cell reproducibility of measured hydraulic parameters depended on the method used, with the PR method giving a lower average coefficient of variation (15.2%, 5.8%, and 19.0% for half-time, cell volume [V_o], and hydraulic conductivity [L_p], respectively) than the PC method (25.4%, 22.0%, and 24.2%, respectively). V_o as determined from PC and PR tests was 1.1 to 2.7 nL and in the range of optically estimated V_o values of 1.5 to 4.9 nL. For the same cell, V_o and L_p estimates were significantly lower (about 15% and 30%, respectively) when determined by PC compared with PR. Both methods, however, showed significantly higher L_pOUT than L_pIN (L_pOUT/L_pIN ≅ 1.20). Because these results were obtained using small and reversible hydrostatically driven flows in the same cell, the 20% outward biased polarity of water transport is most likely not due to artifacts associated with unstirred layers or to direct effects of externally applied osmotica on the membrane, as has been suggested in previous studies. The rapid reversibility of applied flow direction, particularly for the PR method, and the lack of a clear increase in L_pOUT/L_pIN over a wide range of L_p values suggest that the observed polarity is an intrinsic biophysical property of the intact membrane/protein complex.The conductivity of membranes to water (hydraulic conductivity [L_p]) is an important property of the cells of all organisms, and whether plant cell membranes exhibit a polarity in this property has been debated for a number of decades (Dainty and Hope, 1959; Steudle, 1993). Most early evidence for polarity was based on transcellular osmotic experiments using giant algal cells in the Characeae, in which the relative areas of cell membrane exposed to conditions of osmotic inflow (endosmosis) or outflow (exosmosis) could be varied and, hence, L_p for both directions determined (Tazawa and Shimmen, 2001). Interpretation of these experiments is complicated by unstirred layer (USL) effects (Dainty, 1963), but even after accounting for these, it was concluded that inflow L_p (L_pIN) was higher than outflow L_p (L_pOUT) in these cells, with L_pOUT/L_pIN of about 0.65 (Dainty, 1963). When using osmotic driving forces in algal cells, L_pOUT/L_pIN values of between 0.5 and 0.91 have been reported in many studies (Steudle and Zimmermann, 1974; Steudle and Tyerman, 1983; Tazawa et al., 1996), and the same direction of polarity was also reported using osmotic driving forces in whole roots of maize (Zea mays; Steudle et al., 1987). When applying hydrostatic driving forces in algal cells using the pressure probe (Steudle, 1993), which is less influenced by USL effects (Steudle et al., 1980), L_pOUT/L_pIN has been closer to 1 (0.83–1; Steudle and Zimmermann, 1974; Steudle and Tyerman, 1983). However, in higher plant cells, an analysis of the data presented by Steudle et al. (1980, 1982) and Tomos et al. (1981) indicates the opposite polarity, with L_pOUT/L_pIN averaging from 1.2 to 1.4. Moore and Cosgrove (1991) used two contrasting hydrostatic methods to measure L_p in sugarcane (Saccharum spp.) stem cells: (1) the most commonly used pressure relaxation (PR) method, in which cell turgor pressure (P_cell) changes during the measurement, and (2) the more technically demanding pressure clamp (PC) method, in which P_cell is maintained constant. Consistent with other studies in higher plant cells, Moore and Cosgrove (1991) reported average L_pOUT/L_pIN from 1.15 (PC) to 1.65 (PR). Using the PR method in epidermal cells of barley (Hordeum vulgare), Fricke (2000) reported only a modest L_pOUT/L_pIN (based on reported half-time [T_1/2]) of 1.08. In view of the contribution of proteins (e.g. aquaporins) to overall membrane L_p, Tyerman et al. (2002) suggested that polarity may result either from asymmetry in the pores themselves or from an active regulation of the conductive state of the pores in response to the experimental conditions that cause inflow or outflow. Either of these mechanisms may explain the wide range of values reported in the literature for L_pOUT/L_pIN. Cosgrove and Steudle (1981) reported that a substantial (6-fold) and rapid (within 20 s) reduction in L_p could occur in the same cell, and in hindsight, this presumably reflected the influence of aquaporins. Cosgrove and Steudle (1981) did not consider the lower L_p as indicative of the L_p in situ, and Wan et al. (2004) reported that a reduction in L_p was associated with perturbations to P_cell on the order of 0.1 MPa. Hence, if measured membrane L_p itself can exhibit substantial changes over relatively short periods of time in the same cell, then further study of systematic differences between L_pOUT and L_pIN will require a robust hydrostatic methodology (PC or PR) that can reversibly and reproducibly apply small perturbations in pressure (P) to individual cells over short periods of time.For the PR method, a T_1/2 of water exchange is measured by fitting an exponential curve to the observed decay in P_cell over time following a step change in volume, and membrane L_p can be calculated if cell surface area (A), cell volume (V_o), and volumetric elastic modulus (ε) are known (Steudle, 1993). In practice, A and V_o are typically calculated from optical measurements of individual cell dimensions or estimates using average values, and ε is calculated based on V_o and an empirical change in pressure (dP) to change in volume (dV) relation for each cell (Steudle, 1993; Tomos and Leigh, 1999). In the PC method, first developed by Wendler and Zimmermann (1982), V_o (and, given reasonable assumptions about cell geometry, A) is estimated without the need for optical measurements, and L_p can be measured without the need to determine dP/dV or ε. However, this method is technically more demanding because it requires precise P control as well as a continuous record of the volume flow of water across the cell membrane (as measured by changes in the position of the cell solution/oil meniscus within the glass capillary over time) and has rarely been used (Wendler and Zimmermann, 1982, 1985; Cosgrove et al., 1987; Moore and Cosgrove, 1991; Zhang and Tyerman, 1991; Murphy and Smith, 1998). Since volume (V) is continuously changing over time, this approach may also be influenced by the hydraulic conductance of the capillary tip (K_h) used to make the measurements as well as surface tension effects due to the progressive changes in capillary diameter with meniscus position, and these influences have not been quantitatively addressed.Automation of the pressure probe operation, particularly automatic tracking of the meniscus location in the glass microcapillary tip, would address many of the above-mentioned issues, and to date, several attempts have been made to monitor the meniscus location using electrical resistance (Hüsken et al., 1978) or hardware-based image analysis (Cosgrove and Durachko, 1986; Murphy and Smith, 1998). Recently, Wong et al. (2009) redesigned the automated cell pressure probe (ACPP), originally proposed by Cosgrove and Durachko (1986), using a software-based meniscus detection system and a precise pressure control system. In the new ACPP system, both the position of the meniscus and oil pressure (P_oil) are recorded frequently (typically at 10 Hz), and P_oil is controlled with a resolution of ±0.002 MPa. We have combined the ACPP with a new technique to reproducibly fabricate microcapillary tips of known hydraulic properties (Wada et al., 2011) in order to correct for K_h and surface tension effects in both PC and PR estimates of the water relations parameters of Tradescantia virginiana epidermal cells and have determined the relation of L_pOUT to L_pIN in these cells.