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.     

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.     

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

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

Chloroplast Phosphoglycerate Kinase Is Involved in the Targeting of Bamboo mosaic virus to Chloroplasts in Nicotiana benthamiana Plants

The Bamboo mosaic virus (BaMV) is a positive-sense, single-stranded RNA virus. Previously, we identified that the chloroplast phosphoglycerate kinase (chl-PGK) from Nicotiana benthamiana is one of the viral RNA binding proteins involved in the BaMV infection cycle. Because chl-PGK is transported to the chloroplast, we hypothesized that chl-PGK might be involved in viral RNA localization in the chloroplasts. To test this hypothesis, we constructed two green fluorescent protein (GFP)-fused mislocalized PGK mutants, the transit peptide deletion mutant (NO TRANSIT PEPTIDE [NOTP]-PGK-GFP) and the nucleus location mutant (nuclear location signal [NLS]-PGK-GFP). Using confocal microscopy, we demonstrated that NOTP-PGK-GFP and NLS-PGK-GFP are localized in the cytoplasm and nucleus, respectively, in N. benthamiana plants. When NOTP-PGK-GFP and NLS-PGK-GFP are transiently expressed, we observed a reduction in BaMV coat protein accumulation to 47% and 27% that of the wild-type PGK-GFP, respectively. To localize viral RNA in infected cells, we employed the interaction of NLS-GFP-MS2 (phage MS2 coat protein) with the modified BaMV RNA containing the MS2 coat protein binding sequence. Using confocal microscopy, we observed that BaMV viral RNA localizes to chloroplasts. Furthermore, elongation factor1a fused with the transit peptide derived from chl-PGK or with a Rubisco small subunit can partially restore BaMV accumulation in NbPGK1-knockdown plants by helping BaMV target chloroplasts.Bamboo mosaic virus (BaMV) is a single-stranded, positive-sense RNA virus. The genomic RNA of BaMV contains five open reading frames (ORFs) and is 6,366 nucleotides in length with a 5′ cap and a 3′ poly(A) tail (Lin et al., 1994; Yang et al., 1997). ORF1 encodes a 155-kD replicase comprised of a capping enzyme domain that exhibits S-adenosylmethionine-dependent guanylyltransferase activity (Li et al., 2001a; Huang et al., 2004), a helicase-like domain with RNA 5′-triphosphatase activity (Li et al., 2001b), and an RNA-dependent RNA polymerase domain (Li et al., 1998; Cheng et al., 2001). The three overlapping ORFs (i.e. ORF2, ORF3, and ORF4) are known as the triple gene block. They encode for proteins involved in viral movement (Lin et al., 2004, 2006; Vijaya Palani et al., 2006). ORF5 encodes the viral capsid protein (CP), required for virion assembly and viral movement (Cruz et al., 1998).The genomes of positive-strand RNA viruses are templates for both translation and replication. Viral replication complexes are likely to be assembled using host factors to synthesize the minus-strand RNA and then the plus-strand progeny RNA. Recent studies have shown that host factors play important roles in assembling the viral RNA replication complex, selecting and recruiting viral replication templates, activating the complex for RNA synthesis, and other steps (Ahlquist et al., 2003; Patarroyo et al., 2012). The translation and the minus-strand RNA synthesis of poliovirus are regulated by host poly(C) and poly(A) binding proteins and viral polymerase precursor 3CD (Waggoner and Sarnow, 1998; Herold and Andino, 2001; Walter et al., 2002). A number of host genes required for Brome mosaic virus replication have been identified systemically by the yeast (Saccharomyces cerevisiae) genetic approach (Ishikawa et al., 1997; Kushner et al., 2003; Mas et al., 2006; Gancarz et al., 2011). The same approach was used to identify the host factors involved in the replication of Tomato bushy stunt virus (TBSV; Panavas et al., 2005; Li et al., 2009b). A heat shock protein90 homolog (Huang et al., 2012) and the Nicotiana benthamiana glutathione transferase U4 (NbGSTU4; Chen et al., 2013), were identified to interact with the 3′ untranslated region (UTR) of BaMV RNA and enhanced the minus-strand RNA synthesis at the early replication step. The Ser/Thr kinase-like protein localized on cell membrane facilitates the BaMV intercellular movement (Cheng et al., 2013).Previously, we have identified two host proteins (i.e. p51 and p43) interacting specifically with the 3′ UTRs of BaMV by using electrophoretic mobility shift assay (EMSA) and the UV cross-linking competition technique. The results of liquid chromatography-tandem mass spectrometry (LC-MS/MS) and BLAST indicate that the protein sequences of p43 and p51 match the chloroplast phosphoglycerate kinase (chl-PGK) and elongation factor1a (EF1a) of Nicotiana benthamiana, respectively (Lin et al., 2007). Phosphoglycerate kinase is an ATP-generating enzyme that acts in the glycolytic, gluconeogenic, and photosynthetic pathways (Banks et al., 1979; McHarg et al., 1999). chl-PGK is encoded in the nucleus and translated to produce a 50-kD precursor protein and is then processed into mature 43 kD in the chloroplast. In a knockdown experiment through virus-induced gene silencing, the reduction of PGK decreased the accumulation of BaMV coat protein (Lin et al., 2007).Eukaryotic EF1a has been shown to play a role in binding to the tRNA-like structure and upstream pseudoknot in the 3′ UTR of Tobacco mosaic virus to regulate the gene expression and viral replication (Pathak et al., 2008). EF1a has also been involved in the recruitment of viral RNA and has facilitated the replicase complex assembly of TBSV (Pogany et al., 2008). The 3′ UTR of BaMV cannot only bind its replicase but also the EF1a and has been proposed to regulate viral RNA replication (Lin et al., 2007).In this study, we transiently expressed two mislocalized PGK mutants to study the possible functions of chl-PGK that is involved in viral RNA replication. In addition, we used confocal microscopy to investigate the localization of BaMV RNA. Finally, we provided evidence that the down-regulation of BaMV accumulation in PGK-knockdown plants can be restored by the expression of the BaMV RNA binding protein EF1a that is fused to a chloroplast transit peptide.     

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.     

The Cellulase KORRIGAN Is Part of the Cellulose Synthase Complex

Plant growth and organ formation depend on the oriented deposition of load-bearing cellulose microfibrils in the cell wall. Cellulose is synthesized by a large relative molecular weight cellulose synthase complex (CSC), which comprises at least three distinct cellulose synthases. Cellulose synthesis in plants or bacteria also requires the activity of an endo-1,4-β-d-glucanase, the exact function of which in the synthesis process is not known. Here, we show, to our knowledge for the first time, that a leaky mutation in the Arabidopsis (Arabidopsis thaliana) membrane-bound endo-1,4-β-d-glucanase KORRIGAN1 (KOR1) not only caused reduced CSC movement in the plasma membrane but also a reduced cellulose synthesis inhibitor-induced accumulation of CSCs in intracellular compartments. This suggests a role for KOR1 both in the synthesis of cellulose microfibrils and in the intracellular trafficking of CSCs. Next, we used a multidisciplinary approach, including live cell imaging, gel filtration chromatography analysis, split ubiquitin assays in yeast (Saccharomyces cerevisiae NMY51), and bimolecular fluorescence complementation, to show that, in contrast to previous observations, KOR1 is an integral part of the primary cell wall CSC in the plasma membrane.Cellulose microfibrils are synthesized by a hexameric multiprotein complex at the plasma membrane called the cellulose synthase complex (CSC). Genetic analysis, expression data, and coimmunoprecipitation experiments have demonstrated that a functional CSC contains at least three different nonredundant cellulose synthase (CESA) isoforms (Höfte et al., 2007). CESA1, CESA3, and CESA6-like are involved in cellulose biosynthesis during primary cell wall deposition, whereas CESA4, CESA7, and CESA8 are essential for cellulose synthesis in the secondary cell wall (Taylor et al., 1999, 2000, 2003; Desprez et al., 2007; Persson et al., 2007). CSCs labeled by fluorescently tagged CESA proteins migrate in the plasma membrane along cortical microtubules (CMTs), propelled by the polymerization of the β-1,4-glucans (Paredez et al., 2006). Partial depolymerization of CMTs using oryzalin showed that the organized trajectories of CSCs depend on the presence of an intact CMT array. The CSC-microtubule interaction is mediated at least in part by a large protein, POMPOM2/CELLULOSE SYNTHASE INTERACTING1, that binds to both CESAs and microtubules (Lei et al., 2014). Interestingly, complete depolymerization of CMTs does not alter the velocity of the complexes, illustrating that CMTs are necessary for the guidance of CSCs but not for their movement (Paredez et al., 2006). The microtubule cytoskeleton also has a role in the secretion and internalization of CSCs (Crowell et al., 2009; Gutierrez et al., 2009)KORRIGAN1 (KOR1) is a membrane-bound endo-1,4-β-d-glucanase (EGase) that is also required for cellulose synthesis (Nicol et al., 1998). Enzymatic analysis of a recombinant and soluble form of the Brassica napus KOR1 homolog showed substrate specificity for low-substituted carboxymethyl cellulose and amorphous cellulose but no activity on crystalline cellulose, xyloglucans, or short cellulose oligomers (Mølhøj et al., 2001; Master et al., 2004). Fractionation of microsomes demonstrated that KOR1 is primarily present in plasma membrane fractions but also at low levels in a tonoplast-enriched fraction (Nicol et al., 1998). Similarly, the KOR1 ortholog from tomato (Solanum lycopersicum) was found in the plasma membrane and fractions enriched for the Golgi apparatus (Brummell et al., 1997). A GFP-KOR1 fusion protein expressed with the Cauliflower mosaic virus 35S promoter accumulated in the Golgi apparatus and post-Golgi compartments and the tonoplast (Robert et al., 2005). Surprisingly for an enzyme involved in cellulose synthesis, the protein could not be detected at the plasma membrane. Using this construct, it was also shown that KOR1 undergoes regulated intracellular cycling (Robert et al., 2005).Although numerous genetic studies indicate that KOR1 is required for cellulose synthesis in primary and secondary cell walls and during cell plate formation (Nicol et al., 1998; Peng et al., 2000; Zuo et al., 2000; Lane et al., 2001; Sato et al., 2001; Szyjanowicz et al., 2004), its precise role in the cellulose synthesis process remains unclear. It has been suggested that KOR1 might be a component of the CSC (Read and Bacic, 2002). However, until now there has been no experimental evidence for this in Arabidopsis (Arabidopsis thaliana), either with coprecipitation experiments or with localization studies (Szyjanowicz et al., 2004; Robert et al., 2005; Desprez et al., 2007). Numerous hypotheses have been proposed to explain the paradoxical role of KOR1 in cellulose synthesis (Robert et al., 2004). KOR1 might have a proofreading activity involved in hydrolyzing disordered amorphous cellulose to relieve stress generated during the assembly of glucan chains in cellulose microfibrils (Mølhøj et al., 2002). Alternatively, KOR1 may determine the length of individual cellulose chains, either during cellulose synthesis or once the microfibril has been incorporated in the wall. A third hypothesis is that KOR1 releases the cellulose microfibril from the CSC before the complex is internalized from the plasma membrane (Somerville, 2006). Studies in cotton (Gossypium hirsutum) fiber extracts identified sitosterol glucoside as a primer for the cellulose synthesis and suggested that KOR1 could be involved in their cleavage from the nascent glucan chain (Peng et al., 2002). However, this scenario is unlikely, since, at least for the bacterial CESA, which is homologous to plant CESAs, there is no evidence for the existence of lipid-linked precursors, as shown by the three-dimensional structure of an active complex (Morgan et al., 2013).In this study, we first confirmed previous observations (Paredez et al., 2008) that, in the leaky kor1-1 mutant, the velocity of the CSCs is reduced compared with that in a wild-type background but that, in addition, the mutation affects the ability of the cellulose synthesis inhibitor CGA325′615 (hereafter referred to as CGA) to induce the accumulation of GFP-CESA3 in a microtubule-associated compartment (MASC/small compartments carrying cellulose synthase complexes [SmaCCs]; Crowell et al., 2009; Gutierrez et al., 2009). This indicates that KOR1 plays a role both in the synthesis of cellulose and in the intracellular trafficking of the CSC. Using gel filtration approaches, we identified KOR1 in fractions of high molecular mass, suggesting that KOR1 is present in membranes as part of a protein complex. We next analyzed the dynamics of GFP-KOR1 expressed in the kor1-1 mutant background under the control of its endogenous promoter. GFP-KOR1 is found in discrete particles at the plasma membrane in the same cells as GFP-CESAs (Crowell et al., 2009). GFP-KOR1 plasma membrane particles migrate along linear trajectories with comparable velocities to those observed for GFP-CESAs. The organization of GFP-KOR1 at the plasma membrane also requires the presence of an intact microtubule array, suggesting that KOR1 and CESA trajectories in the plasma membrane are regulated in the same manner. GFP-KOR1 and mCherry-CESA1 partially colocalize in the plasma membrane, Golgi, and post-Golgi compartments. Finally, we provide evidence for direct interaction between KOR1 and primary cell wall CESA proteins using the membrane-based yeast (Saccharomyces cerevisiae NMY51) two-hybrid (MbYTH) system (Timmers et al., 2009) and bimolecular fluorescence complementation (BiFC). Our data support a new model in which KOR1 is an integral part of the CSC, where it plays a role not only in the synthesis of cellulose but also in the intracellular trafficking of the CSC.     

The Endoplasmic Reticulum Is a Reservoir for WAVE/SCAR Regulatory Complex Signaling in the Arabidopsis Leaf

During plant cell morphogenesis, signal transduction and cytoskeletal dynamics interact to locally organize the cytoplasm and define the geometry of cell expansion. The WAVE/SCAR (for WASP family verprolin homologous/suppressor of cyclic AMP receptor) regulatory complex (W/SRC) is an evolutionarily conserved heteromeric protein complex. Within the plant kingdom W/SRC is a broadly used effector that converts Rho-of-Plants (ROP)/Rac small GTPase signals into Actin-Related Protein2/3 and actin-dependent growth responses. Although the components and biochemistry of the W/SRC pathway are well understood, a basic understanding of how cells partition W/SRC into active and inactive pools is lacking. In this paper, we report that the endoplasmic reticulum (ER) is an important organelle for W/SRC regulation. We determined that a large intracellular pool of the core W/SRC subunit NAP1, like the known positive regulator of W/SRC, the DOCK family guanine nucleotide-exchange factor SPIKE1 (SPK1), localizes to the surface of the ER. The ER-associated NAP1 is inactive because it displays little colocalization with the actin network, and ER localization requires neither activating signals from SPK1 nor a physical association with its W/SRC-binding partner, SRA1. Our results indicate that in Arabidopsis (Arabidopsis thaliana) leaf pavement cells and trichomes, the ER is a reservoir for W/SRC signaling and may have a key role in the early steps of W/SRC assembly and/or activation.The W/SRC (for WASP family verprolin homologous/suppressor of cAMP receptor regulatory complex) and Actin-Related Protein (ARP)2/3 complex are part of an evolutionarily conserved Rho-of-Plants (ROP)/Rac small GTPase signal transduction cascade that controls actin-dependent morphogenesis in a wide variety of tissues and developmental contexts (Smith and Oppenheimer, 2005; Szymanski, 2005; Yalovsky et al., 2008). Many of the components and regulatory relationships among the complexes were discovered based on the stage-specific cell-swelling and -twisting phenotypes of the distorted class of Arabidopsis (Arabidopsis thaliana) trichome mutants (Szymanski et al., 1999; Zhang et al., 2005, 2008; Djakovic et al., 2006; Le et al., 2006; Uhrig et al., 2007). However, in both maize (Zea mays) and Arabidopsis, W/SRC and/or ARP2/3 are required for normal pavement cell morphogenesis (Frank and Smith, 2002; Mathur et al., 2003b; Brembu et al., 2004). Compared with other Arabidopsis pavement cell mutants, the shape defects of the distorted group are relatively mild. However, the distorted mutants and spike1 (spk1) differ from most other morphology mutants in that they display gaps in the shoot epidermis, most frequently at the interface of pavement cells and stomata (Qiu et al., 2002; Le et al., 2003; Li et al., 2003; Mathur et al., 2003b; Zhang et al., 2005; Djakovic et al., 2006). The cell gaps may reflect either uncoordinated growth between neighboring cells or defective cortical actin-dependent secretion of polysaccharides and/or proteins that promote cell-cell adhesion (Smith and Oppenheimer, 2005; Hussey et al., 2006; Leucci et al., 2007).In tip-growing cells, there is a strict requirement for actin to organize the trafficking and secretion activities of the cell to restrict growth to the apex. In Arabidopsis, the W/SRC-ARP2/3 pathway is not an essential tip growth component, because null alleles of both W/SRC and ARP2/3 subunits do not cause noticeable pollen tube or root hair phenotypes (Le et al., 2003; Djakovic et al., 2006). However, reverse genetic analysis of the W/SRC subunit BRK1 and ARP2/3 in the tip-growing protonemal cells of Physcomitrella patens revealed the obvious importance of this pathway (Harries et al., 2005; Perroud and Quatrano, 2008). Along similar lines, in two different legume species, W/SRC subunits are required for a normal root nodulation response to symbiotic bacteria (Yokota et al., 2009; Miyahara et al., 2010), indicating a conditional importance for this pathway in root hair growth. These genetic studies centered on the W/SRC and ARP2/3 pathways, in addition to those that involve a broader collection of actin-based morphology mutants (Smith and Oppenheimer, 2005; Blanchoin et al., 2010), are defining important cytoskeletal proteins and new interactions with the endomembrane system during morphogenesis. However, it is not completely clear how unstable actin filaments and actin bundle networks dictate the growth patterns of cells (Staiger et al., 2009).The difficulty of understanding the functions of specific actin arrays can be explained, in part, by the fact that plant cells that employ a diffuse growth mechanism have highly unstable cortical actin filaments and large actin bundles that do not have a geometry that obviously relates to the direction of growth or a specific subcellular activity (Blanchoin et al., 2010). This is in contrast to the cortical endocytic actin patches in yeast (Saccharomyces cerevisiae; Evangelista et al., 2002; Kaksonen et al., 2003) and cortical meshworks in the lamellipodia of crawling cells (Pollard and Borisy, 2003) that reveal subcellular locations where actin works to locally control membrane dynamics. In thick-walled plant cells, the magnitude of the forces that accompany turgor-driven cell expansion exceed those that could be generated by actin polymerization by orders of magnitude (Szymanski and Cosgrove, 2009). Localized cell wall loosening or the assembly of an anisotropic cell wall generates asymmetric yielding responses to turgor-induced stress (Baskin, 2005; Cosgrove, 2005). Therefore, the actin-based control of cell boundary dynamics is indirect, and the actin cytoskeleton influences cell shape change, in part, by actin and/or myosin-dependent trafficking of hormone transporters (Geldner et al., 2001) and organelles (Prokhnevsky et al., 2008), including those that control the localized delivery of protein complexes and polysaccharides that pattern the cell wall (Leucci et al., 2007; Gutierrez et al., 2009). In this scheme for actin-based growth control, the actin network dynamically rearranges at spatial scales that span from approximately 1- to 10-µm subcellular domains that may locally position organelles (Cleary, 1995; Gibbon et al., 1999; Szymanski et al., 1999) to the more than 100-µm actin bundle networks that operate at the spatial scales of entire cells (Gutierrez et al., 2009; Dyachok et al., 2011). It is clear from the work of several laboratories that the W/SRC and ARP2/3 protein complexes are required to organize cortical actin and actin bundle networks in trichomes (Szymanski et al., 1999; Le et al., 2003; Deeks et al., 2004; Zhang et al., 2005) and cylindrical epidermal cells (Mathur et al., 2003b; Dyachok et al., 2008, 2011). A key challenge now is to understand how plant cells deploy these approximately 10- to 20-nm heteromeric protein complexes to influence the patterns of growth at cellular scales.The genetic and biochemical control of ARP2/3 is complicated, but this is a tractable problem in plants, because the pathway is relatively simple compared with most other species in which it has been characterized. For example, in organisms ranging from yeast to humans, there are multiple types of ARP2/3 activators, protein complexes, and pathways that activate ARP2/3 (Welch and Mullins, 2002; Derivery and Gautreau, 2010). However, the maize and Arabidopsis genomes encode only WAVE/SCAR homologous proteins that can potently activate ARP2/3 (Frank et al., 2004; Basu et al., 2005). Detailed genetic and biochemical analyses of the WAVE/SCAR gene family in Arabidopsis demonstrated that the plant activators function interchangeably within the context of the W/SRC and define the lone pathway for ARP2/3 activation (Zhang et al., 2008). Bioinformatic analyses are consistent with a prominent role for W/SRC in the angiosperms, because in general, WASH complex subunits, which are structurally similar to WAVE/SCAR proteins, are largely absent from the higher plant genomes, while WAVE/SCAR genes are highly conserved (Kollmar et al., 2012).The components and regulatory schemes of the W/SRC-ARP2/3 pathway in Arabidopsis and P. patens are conserved compared with vertebrate species that employ these same protein complexes (Szymanski, 2005). For example, mutant complementation tests indicate that human W/SRC and ARP2/3 complex subunits can substitute for the Arabidopsis proteins (Mathur et al., 2003b). Furthermore, biochemical assays of Arabidopsis W/SRC (Basu et al., 2004; El-Assal et al., 2004; Frank et al., 2004; Le et al., 2006; Uhrig et al., 2007) and ARP2/3 assembly (Kotchoni et al., 2009) have shown that the binary interactions among W/SRC subunits and ARP2/3 complex assembly mechanisms are indistinguishable from those that have been observed for human W/SRC (Gautreau et al., 2004) and yeast ARP2/3 (Winter et al., 1999). After an initial period of controversy concerning the biochemical control of W/SRC, it is now apparent that vertebrate W/SRC (Derivery et al., 2009; Ismail et al., 2009), like the ARP2/3 complex (Machesky et al., 1999), is intrinsically inactive and requires positive regulation by Rac and other factors to fully activate ARP2/3 (Ismail et al., 2009; Lebensohn and Kirschner, 2009; Chen et al., 2010). Although overexpression of dominant negative ROP mutants causes trichome swelling and a reduced trichome branch number (Fu et al., 2002), the involvement of ROPs in trichome morphogenesis has been difficult to prove with a loss-of-function ROP allele because so many ROPs are expressed in this cell type (Marks et al., 2009). Existing reports on ROP loss-of-function mutants demonstrate the importance of pavement cell morphogenesis but do not document a trichome phenotype (Fu et al., 2005; Xu et al., 2010). A recent report describes a clever strategy to generate ROP loss-of-function lines that used the ectopic expression of ROP-specific bacterial toxins. There was a strong association between inducible expression of the toxins and the appearance of trichomes with severe trichome swelling and reduced branch number phenotypes (Singh et al., 2012). Although the exact mechanism of ROP-dependent control of W/SRC remains to be determined, the results described above in combination with the detection of direct interactions between the ROPGEF SPK1, active forms of ROP, and W/SRC subunits (Basu et al., 2004, 2008; Uhrig et al., 2007) strongly suggest that W/SRC is a ROP effector complex.The major challenge in the field now is to better understand the cellular control of W/SRC and how the complex is partitioned into active and inactive pools. In mammalian cells that crawl on a solid substrate, current models propose that a cytosolic pool of inactive WAVE/SCAR proteins and W/SRC is locally recruited and activated at specific plasma membrane surfaces in response to signals from some unknown Rac guanine nucleotide-exchange factor (GEF), protein kinase, and/or lipid kinase (Oikawa et al., 2004; Lebensohn and Kirschner, 2009; Chen et al., 2010). However, in Drosophila melanogaster neurons (Bogdan and Klämbt, 2003) and cultured human melanoma cells (Steffen et al., 2004), there are large pools of W/SRC with a perinuclear or organelle-like punctate localization that has no obvious relationship to cell shape or motility, raising uncertainty about the cellular mechanisms of W/SRC activation and the importance of different subcellular pools of the complex.In plants, cell fractionation experiments indicate that SCAR1 and ARP2/3 have an increased association with membranes compared with their animal counterparts (Dyachok et al., 2008; Kotchoni et al., 2009). In tip-growing moss protonemal cells, both the W/SRC subunit BRK1 and ARP2/3 localize to a population of unidentified organelles within the apical zone (Perroud and Quatrano, 2008). Similar live-cell imaging experiments in Arabidopsis reported a plasma membrane localization for SCAR1 and BRK1 in a variety of shoot epidermal and root cortex, and their accumulation at young trichome branch tips and at three-way cell wall junctions may define subcellular domains for W/SRC-ARP2/3-dependent actin filament nucleation at the plasma membrane (Dyachok et al., 2008). However, to our knowledge, active W/SRC, defined here as the fraction of W/SRC that colocalizes with ARP2/3 or actin, has not been reported in plants, and the plasma membrane is not necessarily the only organelle involved in W/SRC regulation. For example, the reported accumulation of BRK1 and SCAR1 at three-way cell wall junctions has a punctate appearance at the cell cortex that may not simply correspond to the plasma membrane (Dyachok et al., 2008). Also, in young stage 4 trichomes, there was an uncharacterized pool of intracellular SCAR1, but not BRK1, that localized to relatively large punctate structures (Dyachok et al., 2008). The endoplasmic reticulum (ER) may also be involved in W/SRC regulation. The ER-localized DOCK family ROPGEF SPK1 (Zhang et al., 2010) physically associates with multiple W/SRC proteins (Uhrig et al., 2007; Basu et al., 2008) and, based on genetic criteria, is an upstream, positive regulator of the W/SRC-ARP2/3 pathway (Basu et al., 2008). In the leaf, one function of SPK1 is to promote normal trafficking between the ER and Golgi; however, arp2/3 mutants do not share ER-stress phenotypes with spk1 (Zhang et al., 2010), making it unclear if SPK1 and the ER are directly involved in W/SRC signaling.This paper focuses on the localization and control of the W/SRC subunit NAP1/GNARLED/NAPP/HEM1/2. Arabidopsis NAP1 directly interacts with the ROP/Rac effector subunit SRA1/PIROGI/KLUNKER/PIRP (Basu et al., 2004; El-Assal et al., 2004; Uhrig et al., 2007). Based on the equally severe syndrome of nap1 and arp2/3 null phenotypes, and double mutant analyses, the only known function of NAP1 is to positively regulate ARP2/3 (Brembu et al., 2004; Deeks et al., 2004; El-Din El-Assal et al., 2004; Li et al., 2004). The vertebrate SRA1-NAP1 dimer is important for W/SRC assembly (Gautreau et al., 2004) and forms an extended physical surface that trans-inhibits the C-terminal ARP2/3-activating domain of WAVE/SCAR (Chen et al., 2010). The plant NAP1 and SRA1 proteins share end-to-end amino acid conservation with their vertebrate homologs and may form a heterodimer with similar functions (Basu et al., 2004; El-Assal et al., 2004; Uhrig et al., 2007). We report here that Arabidopsis NAP1 is strongly associated with ER membranes. In a detailed series of localization experiments, we detect a complicated intracellular distribution of NAP1 among the ER, the nucleus, and unidentified submicrometer punctae. A large pool of ER-associated NAP1 is inactive, based on the low level of colocalization with actin.Its accumulation on the ER does not require activating signals from either SPK1 or SRA1. These data indicate that the ER is a reservoir for W/SRC signaling and suggest that early steps in the positive regulation of NAP1 and the W/SRC occur on the ER surface.     

Stomatal Blue Light Response Is Present in Early Vascular Plants

Light is a major environmental factor required for stomatal opening. Blue light (BL) induces stomatal opening in higher plants as a signal under the photosynthetic active radiation. The stomatal BL response is not present in the fern species of Polypodiopsida. The acquisition of a stomatal BL response might provide competitive advantages in both the uptake of CO₂ and prevention of water loss with the ability to rapidly open and close stomata. We surveyed the stomatal opening in response to strong red light (RL) and weak BL under the RL with gas exchange technique in a diverse selection of plant species from euphyllophytes, including spermatophytes and monilophytes, to lycophytes. We showed the presence of RL-induced stomatal opening in most of these species and found that the BL responses operated in all euphyllophytes except Polypodiopsida. We also confirmed that the stomatal opening in lycophytes, the early vascular plants, is driven by plasma membrane proton-translocating adenosine triphosphatase and K⁺ accumulation in guard cells, which is the same mechanism operating in stomata of angiosperms. These results suggest that the early vascular plants respond to both RL and BL and actively regulate stomatal aperture. We also found three plant species that absolutely require BL for both stomatal opening and photosynthetic CO₂ fixation, including a gymnosperm, C. revoluta, and the ferns Equisetum hyemale and Psilotum nudum.Stomata regulate gas exchange between plants and the atmosphere (Zeiger, 1983; Assmann, 1993; Roelfsema and Hedrich, 2005; Shimazaki et al., 2007; Kim et al., 2010). Acquisition of stomata was a key step in the evolution of terrestrial plants by allowing uptake of CO₂ from the atmosphere and accelerating the provision of nutrients via the transpiration stream within the plant (Hetherington and Woodward, 2003; McAdam and Brodribb, 2013). Stomatal aperture is regulated by changes in the turgor of guard cells, which are induced by environmental factors and endogenous phytohormones. Light is a major factor in the promotion of stomatal opening, and the opening is mediated via two distinct light-regulated pathways that are known as photosynthesis- and blue light (BL)-dependent responses under photosynthetic active radiation (PAR; Vavasseur and Raghavendra, 2005; Shimazaki et al., 2007; Lawson et al., 2014).The photosynthesis-dependent stomatal opening is induced by a continuous high intensity of light, and the action spectrum for the stomatal opening resembles that of photosynthetic pigments in leaves (Willmer and Fricker, 1996). Both mesophyll and guard cells possess photosynthetically active chloroplasts, and their photosynthesis has been suggested to contribute to stomatal opening in leaves. The decrease in the concentration of intercellular CO₂ (Ci) caused by photosynthetic CO₂ fixation or some unidentified mediators and metabolites from mesophyll cells is supposed to elicit stomatal opening, although the exact nature of the events is unclear (Wong et al., 1979; Vavasseur and Raghavendra, 2005; Roelfsema et al., 2006; Mott et al., 2008; Lawson et al., 2014).BL-dependent stomatal opening requires a strong intensity of PAR as a background: weak BL solely scarcely elicits stomatal opening, but the same intensity of BL induces the fast and large stomatal opening in the presence of strong red light (RL; Ogawa et al., 1978; Shimazaki et al., 2007). Since such stomatal opening requires BL under the RL or PAR, we call the opening reaction a BL-dependent response of stomata. BL-dependent stomatal response takes place and proceeds in natural environments because the sunlight contains both BL and RL and facilitates photosynthetic CO₂ fixation (Assmann, 1988; Takemiya et al., 2013a). In this stomatal response, BL and PAR (BL, RL, and other wavelengths of light) seem to act as a signal and an energy source, respectively.The BL-dependent stomatal opening is initiated by the absorption of BL by phototropin1 and phototropin2 (Kinoshita et al., 2001), the plant-specific BL receptors, in guard cells followed by activation of the plasma membrane proton-translocating adenosine triphosphatase (H+-ATPase; Kinoshita and Shimazaki, 1999). Two newly identified proteins, protein phosphatase1 and blue light signaling1 (BLUS1), mediate the signaling between phototropins and H+-ATPase (Takemiya et al., 2006, 2013a, 2013b). The activated H+-ATPase evokes a plasma membrane hyperpolarization, which drives K⁺ uptake through the voltage-gated, inward-rectifying K⁺ channels (Assmann, 1993; Shimazaki et al., 2007; Kim et al., 2010; Kollist et al., 2014). The accumulation of K⁺ causes water uptake and increases turgor pressure of guard cells, and finally results in stomatal opening. The BL-dependent opening is enhanced by RL, and BL at low intensity is effective in the presence of RL (Ogawa et al., 1978; Iino et al., 1985; Shimazaki et al., 2007; Suetsugu et al., 2014). These stomatal responses by RL and BL are commonly observed in a number of seed plants so far examined.Fine control of stomatal aperture to various environmental factors has been characterized in many angiosperms. Although morphological and mechanical diversity of stomata is widely documented, little is known about the functional diversity (Willmer and Fricker, 1996; Hetherington and Woodward, 2003). Our previous study indicated that BL-dependent stomatal response is absent in the major fern species of Polypodiopsida, including Adiantum capillus-veneris, Pteris cretica, Asplenium scolopendrium, and Nephrolepis auriculata, but the stomata of these species open by PAR including RL (Doi et al., 2006). When the epidermal peels isolated from A. capillus-veneris are treated with photosynthetic electron transport inhibitor 3-(3,4-dichlorophenyl)-1,1dimethylurea (Doi and Shimazaki, 2008), the response is completely inhibited, but the responses in the seed plants of Vicia faba and Commelina communis are relatively insensitive to 3-(3,4-dichlorophenyl)-1,1dimethylurea (Schwartz and Zeiger, 1984). These findings suggest that there is functional diversity in light-dependent stomatal response in different lineages of land plants. In accord with this notion, the different sensitivities of stomatal response to abscisic acid and CO₂ have been reported among the plant species of angiosperm, gymnosperm, ferns, and lycophytes (Mansfield and Willmer, 1969; Brodribb and McAdam, 2011), although the exact responsiveness to abscisic acid and CO₂ is still debated (Chater et al., 2011, 2013; Ruszala et al., 2011; McAdam and Brodribb, 2013).To address the origin and distribution of stomatal light responses, we investigated the presence of a stomatal response using a gas exchange method and various lineages of vascular plants, including euphyllophytes and lycophytes. Unexpectedly, all plant lineages except Polypodiopsida in monilophytes exhibited a stomatal response to BL in the presence of RL, suggesting that the response was present in the early evolutionary stage of vascular plants. We also report the stomatal opening in response to RL in these plant species.