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.     

An Optimal Frequency in Ca2+ Oscillations for Stomatal Closure Is an Emergent Property of Ion Transport in Guard Cells

Oscillations in cytosolic-free Ca²⁺ concentration ([Ca2+]_i) have been proposed to encode information that controls stomatal closure. [Ca2+]_i oscillations with a period near 10 min were previously shown to be optimal for stomatal closure in Arabidopsis (Arabidopsis thaliana), but the studies offered no insight into their origins or mechanisms of encoding to validate a role in signaling. We have used a proven systems modeling platform to investigate these [Ca2+]_i oscillations and analyze their origins in guard cell homeostasis and membrane transport. The model faithfully reproduced differences in stomatal closure as a function of oscillation frequency with an optimum period near 10 min under standard conditions. Analysis showed that this optimum was one of a range of frequencies that accelerated closure, each arising from a balance of transport and the prevailing ion gradients across the plasma membrane and tonoplast. These interactions emerge from the experimentally derived kinetics encoded in the model for each of the relevant transporters, without the need of any additional signaling component. The resulting frequencies are of sufficient duration to permit substantial changes in [Ca2+]_i and, with the accompanying oscillations in voltage, drive the K⁺ and anion efflux for stomatal closure. Thus, the frequency optima arise from emergent interactions of transport across the membrane system of the guard cell. Rather than encoding information for ion flux, these oscillations are a by-product of the transport activities that determine stomatal aperture.Stomata in the leaf epidermis are the main pathway both for CO₂ entry for photosynthesis and for foliar water loss by transpiration. Guard cells surround the stomatal pore and regulate the aperture, balancing the often conflicting demands for CO₂ and water conservation. Guard cells open and close the pore by expanding and contracting through the uptake and loss, respectively, of osmotic solutes, notably of K⁺, Cl⁻, and malate²⁻ (Mal²⁻; Pandey et al., 2007; Kim et al., 2010; Roelfsema and Hedrich, 2010; Lawson and Blatt, 2014). These transport processes comprise the final effectors of a regulatory network that coordinates transport across the plasma membrane and tonoplast, and maintains the homeostasis of the guard cell. A number of well-defined signals—including light, CO₂, drought and the water stress hormone abscisic acid (ABA)—act on this network, altering transport, solute content, turgor and cell volume, and ultimately stomatal aperture.Much research has focused on stomatal closure, underscoring both Ca²⁺-independent and Ca²⁺-dependent signaling. Of the latter, elevated cytosolic-free Ca²⁺ concentration ([Ca2+]_i) inactivates inward-rectifying K⁺ channels (I_K,in) to prevent K⁺ uptake and activates Cl⁻ (anion) channels (I_Cl) at the plasma membrane to depolarize the membrane and engage K⁺ efflux through outward-rectifying K⁺ channels (I_K,out; Keller et al., 1989; Blatt et al., 1990; Thiel et al., 1992; Lemtiri-Chlieh and MacRobbie, 1994). ABA, and most likely CO₂ (Kim et al., 2010), elevate [Ca2+]_i by facilitating Ca²⁺ entry at the plasma membrane to trigger Ca²⁺ release from endomembrane stores, a process often described as Ca²⁺-induced Ca²⁺ release (Grabov and Blatt, 1998, 1999). The hormone promotes Ca²⁺ influx by activating Ca²⁺ channels (I_Ca) at the plasma membrane, even in isolated membrane patches (Hamilton et al., 2000, 2001), which is linked to reactive oxygen species (Kwak et al., 2003; Wang et al., 2013). In parallel, cADP-ribose and nitric oxide promote endomembrane Ca²⁺ release and [Ca2+]_i elevation (Leckie et al., 1998; Neill et al., 2002; Garcia-Mata et al., 2003; Blatt et al., 2007). Best estimates indicate that endomembrane release accounts for more than 95% of the Ca²⁺ entering the cytosol to raise [Ca2+]_i (Chen et al., 2012; Wang et al., 2012).One feature of stomatal response to ABA, and indeed to a range of stimuli both hormonal as well as external, is its capacity for oscillations both in membrane voltage and [Ca2+]_i. Guard cell [Ca2+]_i at rest is typically around 100 to 200 nm, as it is in virtually all living cells. In response to ABA, [Ca2+]_i can rise above 1 μm—and locally, most likely above 10 μm—often in cyclic transients of tens of seconds to several minutes’ duration in association with oscillations in voltage and stomatal closure (Gradmann et al., 1993; McAinsh et al., 1995; Webb et al., 1996; Grabov and Blatt, 1998, 1999; Staxen et al., 1999; Allen et al., 2001). In principle, cycling in voltage and [Ca2+]_i arises as closure is accelerated with a controlled release of K⁺, Cl⁻, and Mal²⁻ from the guard cell and is subject to extracellular ion concentrations (Gradmann et al., 1993; Chen et al., 2012). However, it has been proposed that these, and similar oscillations in a variety of plant cell models, serve as physiological signals in their own right (McAinsh et al., 1995; Ehrhardt et al., 1996; Taylor et al., 1996). In support of such a signaling role, experiments designed to impose [Ca2+]_i (and voltage) oscillations in guard cells have yielded an optimal frequency for closure with a period near 10 min (Allen et al., 2001). Nonetheless, the studies offer no mechanistic explanation for this optimum that could validate a causal role in signaling, and none has been forthcoming since. Here we address questions of how such optimal frequencies in [Ca2+]_i oscillation arise and their relevance for stomatal closure, using quantitative systems analysis of guard cell transport and homeostasis. Our findings indicate that oscillations in voltage and [Ca2+]_i, and their optima associated with stomatal closure, are most simply explained as emerging from the interactions between ion transporters that drive stomatal closure. Thus, we conclude that these oscillations do not control, but are a by-product of the transport that determines stomatal aperture.     

Self-Incompatibility-Induced Programmed Cell Death in Field Poppy Pollen Involves Dramatic Acidification of the Incompatible Pollen Tube Cytosol

Self-incompatibility (SI) is an important genetically controlled mechanism to prevent inbreeding in higher plants. SI involves highly specific interactions during pollination, resulting in the rejection of incompatible (self) pollen. Programmed cell death (PCD) is an important mechanism for destroying cells in a precisely regulated manner. SI in field poppy (Papaver rhoeas) triggers PCD in incompatible pollen. During SI-induced PCD, we previously observed a major acidification of the pollen cytosol. Here, we present measurements of temporal alterations in cytosolic pH ([pH]_cyt); they were surprisingly rapid, reaching pH 6.4 within 10 min of SI induction and stabilizing by 60 min at pH 5.5. By manipulating the [pH]_cyt of the pollen tubes in vivo, we show that [pH]_cyt acidification is an integral and essential event for SI-induced PCD. Here, we provide evidence showing the physiological relevance of the cytosolic acidification and identify key targets of this major physiological alteration. A small drop in [pH]_cyt inhibits the activity of a soluble inorganic pyrophosphatase required for pollen tube growth. We also show that [pH]_cyt acidification is necessary and sufficient for triggering several key hallmark features of the SI PCD signaling pathway, notably activation of a DEVDase/caspase-3-like activity and formation of SI-induced punctate actin foci. Importantly, the actin binding proteins Cyclase-Associated Protein and Actin-Depolymerizing Factor are identified as key downstream targets. Thus, we have shown the biological relevance of an extreme but physiologically relevant alteration in [pH]_cyt and its effect on several components in the context of SI-induced events and PCD.Programmed cell death (PCD) in plants is relatively well documented and characterized (Jones and Dangl, 1996; van Doorn, 2011; van Doorn et al., 2011). There is considerable biochemical evidence for the involvement of caspase-like activities in plant PCD (van Doorn and Woltering, 2005). For example, the vacuolar processing enzyme has YVADase (caspase-1-like) activity (Hatsugai et al., 2004; Rojo et al., 2004; Hara-Nishimura et al., 2005), DEVDase (caspase-3-like) and YVADases are associated with PCD in several plant systems (del Pozo and Lam, 1998; Korthout et al., 2000; Danon et al., 2004), and VEIDase (caspase-6-like) is the main caspase-like activity involved in embryonic pattern formation (Bozhkov et al., 2004). However, because plants have no caspase gene homologs (Sanmartín et al., 2005), the nature of their caspase-like enzymes is the subject of considerable debate. Vacuolar cell death is one of two major classes of PCD in plants (van Doorn et al., 2011). It is thought that collapse of the vacuole is a key irreversible step in several plant PCD systems, including during tissue and organ formation, such as the classic differentiation of tracheary elements (Hara-Nishimura and Hatsugai, 2011). Exactly how this is achieved and what processes are involved remain unknown. Until very recently, it was generally thought that the rupturing vacuole releases proteases, hydrolases, and nucleases, allowing cellular disassembly by an autophagy-like process. Some PCD systems cannot be assigned to either class; these include PCD triggered by the hypersensitive response to biotrophic pathogens, PCD in cereal endosperm, and self-incompatibility (SI)-induced PCD (van Doorn et al., 2011).SI is a genetically controlled pollen-pistil cell-cell recognition system. Self-pollen is recognized by the stigma as being genetically identical, resulting in inhibition of pollen tube growth. Most SI systems use tightly linked polymorphic genes: the pollen (male) and pistil (female) S-determinants. In field poppy (Papaver rhoeas), the S-determinants are a 14-kD signaling ligand field poppy stigma S (PrsS) and a unique transmembrane protein field poppy pollen S (PrpS; Foote et al., 1994; Wheeler et al., 2010). These interact in an S-specific manner, and increases in cytosolic free calcium ([Ca2+]_cyt) are triggered in incompatible pollen tubes (Franklin-Tong et al., 1993), resulting in phosphorylation of soluble inorganic pyrophosphatases (sPPases; Rudd et al., 1996; de Graaf et al., 2006), activation of a Mitogen-Activated Protein Kinase (MAPK; Rudd et al., 2003), and increases in reactive oxygen species (ROS) and nitric oxide (Wilkins et al., 2011, 2014). Most of these components are integrated into a signaling network leading to PCD (Bosch et al., 2008; Wilkins et al., 2014). The actin cytoskeleton is a key target in the field poppy SI response, undergoing depolymerization (Snowman et al., 2002) followed by polymerization into highly stable F-actin foci decorated with the actin binding proteins (ABPs) Actin-Depolymerizing Factor (ADF) and Cyclase-Associated Protein (CAP; Poulter et al., 2010, 2011), with both processes being involved in mediating PCD (Thomas et al., 2006). A major player in SI-mediated PCD is a caspase-3-like/DEVDase-like activity (Thomas and Franklin-Tong, 2004; Bosch and Franklin-Tong, 2007). The SI-induced caspase-3-like/DEVDase exhibits maximum substrate cleavage in vitro at pH 5, with peak activity 5 h after SI induction in vivo (Bosch and Franklin-Tong, 2007). The low pH optimum for this caspase-3-like/DEVDase activity is unusual, because most of the cytosolic plant caspase-like activities identified to date have optimal activity close to normal physiological pH (approximate pH, 6.5–7.0; Korthout et al., 2000; Bozhkov et al., 2004; Coffeen and Wolpert, 2004). Because the SI-induced cytosolic-located DEVDase requires a low pH for activity, this suggested that, during SI, the pollen tube cytosol undergoes dramatic acidification. In vivo pH measurements of the cytosol at 1 to 4 h after SI induction confirmed this, when cytosolic pH ([pH]_cyt) had dropped from pH 6.9 to pH 5.5 (Bosch and Franklin-Tong, 2007). This fits the in vitro pH optimum of the caspase-3-like/DEVDase almost exactly, implicating pollen cytosolic acidification as playing a vital role in creating optimal conditions for the activation of the caspase-3-like/DEVDase-like activity and progression of PCD.Under normal cellular conditions, [pH]_cyt is between approximately 6.9 and 7.5 (Kurkdjian and Guern, 1989; Felle, 2001). Pollen tubes, like other tip-growing cells, have [pH]_cyt gradients (Gibbon and Kropf, 1994; Feijó et al., 1999). The [pH]_cyt of the pollen tube shank is an approximate pH of 6.9 to 7.11 (Fricker et al., 1997; Messerli and Robinson, 1998). There has been much debate about the [pH]_cyt gradient, comprising an apical domain with an approximate pH of 6.8 and a subapical alkaline band with an approximate pH of 7.2 to 7.8 in Lilium longiflorum and Lilium formosanum pollen tubes (Fricker et al., 1997; Messerli and Robinson, 1998; Feijó et al., 2001; Lovy-Wheeler et al., 2006). Oscillations of [pH]_cyt between approximate pH values of 6.9 and 7.3 have been linked to tip growth in L. formosanum pollen tubes (Lovy-Wheeler et al., 2006). The vacuole and the apoplast have a highly acidic pH between pH 5 and pH 6 (Katsuhara et al., 1989; Feijó et al., 1999). The majority of studies of pH changes in plant cells reports modest, transient changes in [pH]_cyt of approximately 0.4 and 0.7 pH units during development, gravitropic responses, decreases in light intensity, and addition of elicitors, hormones, and other treatments. For example, during root hair development in Arabidopsis (Arabidopsis thaliana), root [pH]_cyt was elevated from an approximate pH of 7.3 to 7.7 (Bibikova et al., 1998). Root gravitropic responses stimulate small transient [pH]_cyt alterations (Scott and Allen, 1999; Fasano et al., 2001; Johannes et al., 2001). More recently, it has been shown that the [pH]_cyt drops during PCD controlling root cap development; however, exactly how many units the [pH]_cyt decreased was not measured (Fendrych et al., 2014). Other studies investigating [pH]_cyt in response to physiologically relevant signals also report small transient alterations. Light-adapted cells respond to a decrease in light intensity with a rapid transient cytosolic acidification by approximately 0.3 pH units (Felle et al., 1986). Addition of nodulation factors resulted in an increase of 0.2 pH units in root hairs (Felle et al., 1998), and abscisic acid increased the [pH]_cyt of guard cells by 0.3 pH units (Blatt and Armstrong, 1993). Changes in [pH]_cyt are thought to activate stress responses (Felle, 2001). Elicitor treatments resulted in a [pH]_cyt drop of between 0.4 and 0.7 pH units in suspension cells (Mathieu et al., 1996; Kuchitsu et al., 1997), a drop of 0.2 pH units in Nitellopsis obtusa cells treated with salt (Katsuhara et al., 1989), and a drop of 0.3 to 0.7 pH units in Eschscholzia californica (Roos et al., 1998).Here, we investigate SI-induced acidification of the cytosol, providing measurements of physiologically relevant temporal alterations in [pH]_cyt, and identify key targets of this, providing mechanistic insights into these events. The SI-induced acidification plays a pivotal role in the activation of a caspase-3-like/DEVDase activity, the formation of punctate F-actin foci, and ABP localization during SI PCD. We investigate the vacuole as a potential contributor to SI-induced [pH]_cyt acidification.     

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

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

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

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

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.     

NdhV Is a Subunit of NADPH Dehydrogenase Essential for Cyclic Electron Transport in Synechocystis sp. Strain PCC 6803

Two mutants sensitive to heat stress for growth and impaired in NADPH dehydrogenase (NDH-1)-dependent cyclic electron transport around photosystem I (NDH-CET) were isolated from the cyanobacterium Synechocystis sp. strain PCC 6803 transformed with a transposon-bearing library. Both mutants had a tag in the same sll0272 gene, encoding a protein highly homologous to NdhV identified in Arabidopsis (Arabidopsis thaliana). Deletion of the sll0272 gene (ndhV) did not influence the assembly of NDH-1 complexes and the activities of CO₂ uptake and respiration but reduced the activity of NDH-CET. NdhV interacted with NdhS, a ferredoxin-binding subunit of cyanobacterial NDH-1 complex. Deletion of NdhS completely abolished NdhV, but deletion of NdhV had no effect on the amount of NdhS. Reduction of NDH-CET activity was more significant in ΔndhS than in ΔndhV. We therefore propose that NdhV cooperates with NdhS to accept electrons from reduced ferredoxin.Cyanobacterial NADPH dehydrogenase (NDH-1) complexes are localized in the thylakoid membrane (Ohkawa et al., 2001, 2002; Zhang et al., 2004; Xu et al., 2008; Battchikova et al., 2011b) and participate in a variety of bioenergetic reactions, such as respiration, cyclic electron transport around photosystem I (NDH-CET), and CO₂ uptake (Ogawa, 1991; Mi et al., 1992; Ohkawa et al., 2000). Structurally, the cyanobacterial NDH-1 complexes closely resemble energy-converting complex I in eubacteria and the mitochondrial respiratory chain regardless of the absence of homologs of three subunits in cyanobacterial genomes that constitute the catalytically active core of complex I (Friedrich et al., 1995; Friedrich and Scheide, 2000; Arteni et al., 2006). Over the past decade, new subunits of NDH-1 complexes specific to oxygenic photosynthesis have been identified in several cyanobacterial strains. They are NdhM to NdhQ and NdhS (Prommeenate et al., 2004; Battchikova et al., 2005, 2011b; Nowaczyk et al., 2011; Wulfhorst et al., 2014; Zhang et al., 2014; Zhao et al., 2014b, 2015), in addition to NdhL first identified in the cyanobacterium Synechocystis sp. strain PCC 6803 (hereafter Synechocystis 6803) about 20 years ago (Ogawa, 1992). Among them, NdhS possesses a ferredoxin (Fd)-binding motif and was shown to bind Fd, which suggested that Fd is one of the electron donors to NDH-1 complexes (Mi et al., 1995; Battchikova et al., 2011b; Ma and Ogawa, 2015). Deletion of NdhS strongly reduced the activity of NDH-CET but had no effect on respiration and CO₂ uptake (Battchikova et al., 2011b; Ma and Ogawa, 2015). The NDH-CET plays an important role in coping with various environmental stresses regardless of its elusive mechanism. For example, this function can greatly alleviate heat-sensitive growth phenotypes (Wang et al., 2006a; Zhao et al., 2014a). Thus, heat treatment strategy can help in identifying the proteins essential to NDH-CET.Here, a new oxygenic photosynthesis-specific (OPS) subunit NdhV was identified in Synechocystis 6803 with the help of heat treatment strategy, and its deletion did not influence the assembly of NDH-1L and NDH-1MS complexes and the activities of CO₂ uptake and respiration but impaired the NDH-CET activity. We give evidence that NdhV interacts with NdhS and is another component of Fd-binding domain of cyanobacterial NDH-1 complex. A possible role of NdhV on the NDH-CET activity is discussed.     

Phototropins Function in High-Intensity Blue Light-Induced Hypocotyl Phototropism in Arabidopsis by Altering Cytosolic Calcium

Phototropins (phot1 and phot2), the blue light receptors in plants, regulate hypocotyl phototropism in a fluence-dependent manner. Especially under high fluence rates of blue light (HBL), the redundant function mediated by both phot1 and phot2 drastically restricts the understanding of the roles of phot2. Here, systematic analysis of phototropin-related mutants and overexpression transgenic lines revealed that HBL specifically induced a transient increase in cytosolic Ca²⁺ concentration ([Ca2+]_cyt) in Arabidopsis (Arabidopsis thaliana) hypocotyls and that the increase in [Ca2+]_cyt was primarily attributed to phot2. Pharmacological and genetic experiments illustrated that HBL-induced Ca²⁺ increases were modulated differently by phot1 and phot2. Phot2 mediated the HBL-induced increase in [Ca2+]_cyt mainly by an inner store-dependent Ca²⁺-release pathway, not by activating plasma membrane Ca²⁺ channels. Further analysis showed that the increase in [Ca2+]_cyt was possibly responsible for HBL-induced hypocotyl phototropism. An inhibitor of auxin efflux carrier exhibited significant inhibitions of both phototropism and increases in [Ca2+]_cyt, which indicates that polar auxin transport is possibly involved in HBL-induced responses. Moreover, PHYTOCHROME KINASE SUBSTRATE1 (PKS1), the phototropin-related signaling element identified, interacted physically with phototropins, auxin efflux carrier PIN-FORMED1 and calcium-binding protein CALMODULIN4, in vitro and in vivo, respectively, and HBL-induced phototropism was impaired in pks multiple mutants, indicating the role of the PKS family in HBL-induced phototropism. Together, these results provide new insights into the functions of phototropins and highlight a potential integration point through which Ca²⁺ signaling-related HBL modulates hypocotyl phototropic responses.Blue light (BL) is a key factor controlling plant growth and morphogenesis. Recent genetics investigations using Arabidopsis (Arabidopsis thaliana) have revealed that the BL receptors phototropin1 (phot1) and phot2 mediate BL-induced plant movements such as phototropism, chloroplast relocation, stomatal opening, leaf flattening, and leaf positioning responses (Inoue et al., 2010). Most of these responses are mediated redundantly by both phot1 and phot2 (Kinoshita et al., 2001; Sakamoto and Briggs, 2002), but some responses are mediated by either phot1 or phot2 (Sakai et al., 2001; Suetsugu et al., 2005). In addition, several lines of evidence have indicated that phot2 might negatively regulate the phot1-mediated response (de Carbonnel et al., 2010) and vice versa (Harada et al., 2003, 2013).One of the numerous physiological processes controlled by BL is phototropism. Phototropism enables plants to bend toward incident light by perceiving the direction, wavelength, and intensity of incident light so that they are able to obtain optimum light. Genetic evidence has shown that both phot1 and phot2 redundantly function to regulate hypocotyl phototropism in a fluence-dependent manner (Sakai et al., 2001). Phot1 functions at both low (0.01–1 μmol m⁻² s⁻¹) and high (greater than 1 μmol m⁻² s⁻¹) fluence rates to mediate phototropic responses, but phot2 functions only at high fluence rates (Inada et al., 2004). The functional specification of phot1 and phot2 could be attributed to the differences in signal intermediates between phot1 and phot2 signaling pathways.Genetic analysis has illustrated that phot1 mediates hypocotyl phototropism via its downstream signal transducers NONPHOTOTROPIC HYPOCOTYL3 (NPH3; Motchoulski and Liscum, 1999), ROOT PHOTOTROPISM2 (RPT2; Sakai et al., 2000), and NONPHOTOTROPIC HYPOCOTYL4/AUXIN RESPONSE FACTOR7 (NPH4/ARF7; Harper et al., 2000), resulting in the asymmetric distribution of auxin and the induction of a phototropic response in higher plants. Recently, studies have demonstrated that PHYTOCHROME KINASE SUBSTRATE (PKS) proteins are required for hypocotyl phototropism and that PKS1 binds PHOT1 and NPH3 in vivo (Lariguet et al., 2006). In addition, ATP-BINDING CASSETTE B19 (ABCB19), a newly identified auxin transporter, has been reported to interact with phot1 to regulate the BL-dependent phototropism (Christie et al., 2011). However, little is known about phot2-mediated phototropism for functional specialization, especially under high fluence rates of blue light (HBL), although several lines of evidence have shown that phot2- and phot1-mediated signaling pathways share some intermediates in BL responses (Kimura and Kagawa, 2006; Christie, 2007). Previous researches have suggested that phot1 acts not only positively in the presence of RPT2 but also negatively in its absence during the phototropic response of hypocotyls at high fluence rates, suggesting that RPT2 modulates the function of phot1. However, RPT2 does not act in the phot2-mediated pathway (Inada et al., 2004). More recently, RCN1-1, the A1 subunit of Ser/Thr PROTEIN PHOSPHATASE2A (PP2A), has been identified to interact with phot2. While reduced PP2A activity enhances the activity of phot2, it does not enhance either phot1 dephosphorylation or the activity of phot1 in mediating phototropism (Tseng and Briggs, 2010).Besides these signal intermediates noted above, phototropins may also confer their effects through the change of ion homeostasis. Ca²⁺ is a case in point. Recent reports have demonstrated that phototropins mediate the mobilization of Ca²⁺ in response to BL and that phot1 and phot2 mediate Ca²⁺ increases with distinctive mechanism in leaf cells according to the changes of ambient light intensity (Harada and Shimazaki, 2007). Under low fluence rates of BL, phot1 solely mediated Ca²⁺ influx through the channels in the plasma membrane. Under HBL, the increase in cytosolic Ca²⁺ concentration ([Ca2+]_cyt) is primarily attributed to phot2-dependent Ca²⁺ release from the internal calcium stores as well as the plasma membrane Ca²⁺ channels. Interestingly, the inhibitory effects of phospholipase C (PLC) inhibitors on the BL-induced responses in the wild type are larger than those in the phot1 single mutant, which indicates that there are some functional interactions between phot1 and phot2 to induce the elevation of cytosolic Ca²⁺ (Harada et al., 2003).However, until now, the function of Ca²⁺ in the phototropin-mediated phototropism signaling process has remained largely unknown. Pharmacological experiments indicate that changes in [Ca2+]_cyt are required for the phot1-mediated inhibition of hypocotyl growth but not for phot1-mediated phototropism (Folta et al., 2003). Otherwise, electrophysiological studies indicate that phototropic bending involves changes in ion fluxes, including calcium (Babourina et al., 2004). Such divergent responses show that the link between phototropins and calcium has not been firmly established in the case of hypocotyl phototropism. In phototropism, the phot1-dependent relocalization of the auxin efflux carrier PIN-FORMED1 (PIN1) is required for auxin redistribution (Blakeslee et al., 2004), and the PINOID kinase influences the relocalization of PIN1 (Friml et al., 2004). Given that both the calmodulin-related protein TCH3 and the calcium-binding protein AtPBP1 can bind to the PINOID kinase (Benjamins et al., 2003), it would appear that the cross talk among phototropins, auxin, and calcium is an important event for phototropism.Here, we show that HBL induces increases in [Ca2+]_cyt, which are mostly attributed to the function of phot2, and that the increases in [Ca2+]_cyt are required for HBL-induced phototropism in Arabidopsis hypocotyls. We also demonstrate that PKS1 may integrate phototropins with auxin transport in phot2-dependent Ca²⁺ signaling, and we discuss the possible molecular link between phototropins and other potential signal elements in HBL-induced phototropism.     

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.     

Photosynthetic Energy Conversion Efficiency: Setting a Baseline for Gauging Future Improvements in Important Food and Biofuel Crops

The conversion efficiency (ε_c) of absorbed radiation into biomass (MJ of dry matter per MJ of absorbed photosynthetically active radiation) is a component of yield potential that has been estimated at less than half the theoretical maximum. Various strategies have been proposed to improve ε_c, but a statistical analysis to establish baseline ε_c levels across different crop functional types is lacking. Data from 164 published ε_c studies conducted in relatively unstressed growth conditions were used to determine the means, greatest contributors to variation, and genetic trends in ε_c across important food and biofuel crop species. ε_c was greatest in biofuel crops (0.049–0.066), followed by C₄ food crops (0.046–0.049), C₃ nonlegumes (0.036–0.041), and finally C₃ legumes (0.028–0.035). Despite confining our analysis to relatively unstressed growth conditions, total incident solar radiation and average growing season temperature most often accounted for the largest portion of ε_c variability. Genetic improvements in ε_c, when present, were less than 0.7% per year, revealing the unrealized potential of improving ε_c as a promising contributing strategy to meet projected future agricultural demand.Substantial increases in yield are needed to feed and fuel the world’s growing human population. With an estimated population of nine billion people by the middle of this century (Lutz and Samir, 2010) and rising affluence resulting in greater consumption of grain-fed animal products (Cirera and Masset, 2010), different studies predict that, by midcentury, global crop production will need to increase 60% to 120% over 2005 levels without the expansion of agricultural land area (Tilman et al., 2011; Alexandratos and Bruinsma, 2012).Doubling yields in major food and fuel crops requires considerable effort, especially as yields are beginning to plateau in many major food crops. Yield increases necessary for doubling productivity by midcentury are estimated at 1.16% to 1.31% each year in all cereals (Hall and Richards, 2013), 1.7% per year in wheat (Triticum aestivum; Rosegrant and Agcaoili, 2010), and 2.4% (noncompounding average per year) across all major grain crops (Ray et al., 2013). However, global mean increases from the past 20 to 30 years suggest that yield gains in rice (Oryza sativa) and wheat are approximately 1% (Lopes et al., 2012; Manès et al, 2012; Ray et al., 2013) and declining in some areas of the world (Cassman et al., 2010; Fischer and Edmeades, 2010; Long and Ort, 2010; Ray et al., 2013). Global yearly increases are estimated at 1.3% in soybean (Glycine max) and 1.6% in maize (Zea mays), with similar concerns that yield trends may also be decreasing in some major growing regions (Lobell and Gourdji, 2012; Ray et al., 2013).Efforts to increase yields in the next few decades must also account for environmental and sustainability goals (Sayer et al., 2013) as well as heightened environmental stresses predicted to occur due to climate change, which are already responsible for some of the stagnation in yield increases. Anthropogenic sources of greenhouse gases have caused an approximately 1°C increase in land surface temperatures since 1900, and global mean surface temperatures are likely to increase by up to 2.4°C to 4.8°C by the end of the century (IPCC, 2013). Drought is also expected to become more frequent and intense in many regions of the world (Dai, 2011; IPCC, 2013). Of the variability present in major food crop yield gains, 30% can be explained by climate change alone (Lobell and Field, 2007), with drastic decreases in barley (Hordeum vulgare), maize, rice, sorghum (Sorghum bicolor), soybean, and wheat yields as average growing season temperatures surpass the temperature optimum for each crop (Lobell and Gourdji, 2012). Current levels of atmospheric CO₂ concentration [CO₂] are the highest they have been in at least 800,000 years (IPCC, 2013). Elevated [CO₂] increases water use efficiency (Ainsworth and Long, 2005, Bernacchi et al., 2007, Leakey et al., 2009), but probably not to an extent that would mitigate the resulting reductions in yield caused by higher temperature and higher vapor pressure deficit (Ort and Long, 2014). Additionally, any fertilization effects on C₃ yields due to elevated [CO₂] would be at least in part negated by drought and temperature stress, leaving yield increases far from optimal (Long et al., 2006a; Lobell and Gourdji, 2012).