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.     

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

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

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.     

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

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

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.     

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

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

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.     

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.     

Responses of Arabidopsis and Wheat to Rising CO2 Depend on Nitrogen Source and Nighttime CO2 Levels

A major contributor to the global carbon cycle is plant respiration. Elevated atmospheric CO₂ concentrations may either accelerate or decelerate plant respiration for reasons that have been uncertain. We recently established that elevated CO₂ during the daytime decreases plant mitochondrial respiration in the light and protein concentration because CO₂ slows the daytime conversion of nitrate (NO₃−) into protein. This derives in part from the inhibitory effect of CO₂ on photorespiration and the dependence of shoot NO₃− assimilation on photorespiration. Elevated CO₂ also inhibits the translocation of nitrite into the chloroplast, a response that influences shoot NO₃− assimilation during both day and night. Here, we exposed Arabidopsis (Arabidopsis thaliana) and wheat (Triticum aestivum) plants to daytime or nighttime elevated CO₂ and supplied them with NO₃− or ammonium as a sole nitrogen (N) source. Six independent measures (plant biomass, shoot NO₃−, shoot organic N, ¹⁵N isotope fractionation, ¹⁵NO₃⁻ assimilation, and the ratio of shoot CO₂ evolution to O₂ consumption) indicated that elevated CO₂ at night slowed NO₃− assimilation and thus decreased dark respiration in the plants reliant on NO₃−. These results provide a straightforward explanation for the diverse responses of plants to elevated CO₂ at night and suggest that soil N source will have an increasing influence on the capacity of plants to mitigate human greenhouse gas emissions.The CO₂ concentration in Earth’s atmosphere has increased from about 270 to 400 µmol mol^–1 since 1800, and may double before the end of the century (Intergovernmental Panel on Climate Change, 2013). Plant responses to such increases are highly variable, but plant nitrogen (N) concentrations generally decline under elevated CO₂ (Cotrufo et al., 1998; Long et al., 2004). One explanation for this decline is that CO₂ inhibits nitrate (NO₃−) assimilation into protein in the shoots of C₃ plants during the daytime (Bloom et al., 2002, 2010, 2012, 2014; Cheng et al., 2012; Pleijel and Uddling, 2012; Myers et al., 2014; Easlon et al., 2015; Pleijel and Högy, 2015). This derives in part from the inhibitory effect of CO₂ on photorespiration (Foyer et al., 2009) and the dependence of shoot NO₃− assimilation on photorespiration (Rachmilevitch et al., 2004; Bloom, 2015).A key factor in global carbon budgets is plant respiration at night (Amthor, 1991; Farrar and Williams, 1991; Drake et al., 1999; Leakey et al., 2009). Nighttime elevated CO₂ may inhibit, have a negligible effect on, or stimulate dark respiration, depending on the plant species (Bunce, 2001, 2003; Wang and Curtis, 2002), plant development stage (Wang et al., 2001; Li et al., 2013), experimental approach (Griffin et al., 1999; Baker et al., 2000; Hamilton et al., 2001; Bruhn et al., 2002; Jahnke and Krewitt, 2002; Bunce, 2004), and total N supply (Markelz et al., 2014). The current study is, to our knowledge, the first to examine the influence of N source, NO₃− versus ammonium (NH₄+), on plant dark respiration at elevated CO₂ during the night.Plant organic N compounds account for less than 5% of the total dry weight of a plant, but conversion of NO₃− into organic N expends about 25% of the total energy in shoots (Bloom et al., 1989) and roots (Bloom et al., 1992). During the day, photorespiration supplies a portion of the energy (Rachmilevitch et al., 2004; Foyer et al., 2009), but at night, this energetic cost is borne entirely by the respiration of C substrates (Amthor, 1995) and may divert a substantial amount of reductant from the mitochondrial electron transport chain (Cousins and Bloom, 2004). The relative importance of NO₃− assimilation at night versus the day, however, is still a matter of intense debate (Nunes-Nesi et al., 2010). Here, we estimated NO₃− assimilation using several independent methods and show in Arabidopsis (Arabidopsis thaliana) and wheat (Triticum aestivum), two diverse C₃ plants, that NO₃− assimilation at night can be substantial, and that elevated CO₂ at night inhibits this process.