Characterization of the Possible Roles for B Class MADS Box Genes in Regulation of Perianth Formation in Orchid

To investigate sepal/petal/lip formation in Oncidium Gower Ramsey, three paleoAPETALA3 genes, O. Gower Ramsey MADS box gene5 (OMADS5; clade 1), OMADS3 (clade 2), and OMADS9 (clade 3), and one PISTILLATA gene, OMADS8, were characterized. The OMADS8 and OMADS3 mRNAs were expressed in all four floral organs as well as in vegetative leaves. The OMADS9 mRNA was only strongly detected in petals and lips. The mRNA for OMADS5 was only strongly detected in sepals and petals and was significantly down-regulated in lip-like petals and lip-like sepals of peloric mutant flowers. This result revealed a possible negative role for OMADS5 in regulating lip formation. Yeast two-hybrid analysis indicated that OMADS5 formed homodimers and heterodimers with OMADS3 and OMADS9. OMADS8 only formed heterodimers with OMADS3, whereas OMADS3 and OMADS9 formed homodimers and heterodimers with each other. We proposed that sepal/petal/lip formation needs the presence of OMADS3/8 and/or OMADS9. The determination of the final organ identity for the sepal/petal/lip likely depended on the presence or absence of OMADS5. The presence of OMADS5 caused short sepal/petal formation. When OMADS5 was absent, cells could proliferate, resulting in the possible formation of large lips and the conversion of the sepal/petal into lips in peloric mutants. Further analysis indicated that only ectopic expression of OMADS8 but not OMADS5/9 caused the conversion of the sepal into an expanded petal-like structure in transgenic Arabidopsis (Arabidopsis thaliana) plants.The ABCDE model predicts the formation of any flower organ by the interaction of five classes of homeotic genes in plants (Yanofsky et al., 1990; Jack et al., 1992; Mandel et al., 1992; Goto and Meyerowitz, 1994; Jofuku et al., 1994; Pelaz et al., 2000, 2001; Theißen and Saedler, 2001; Pinyopich et al., 2003; Ditta et al., 2004; Jack, 2004). The A class genes control sepal formation. The A, B, and E class genes work together to regulate petal formation. The B, C, and E class genes control stamen formation. The C and E class genes work to regulate carpel formation, whereas the D class gene is involved in ovule development. MADS box genes seem to have a central role in flower development, because most ABCDE genes encode MADS box proteins (Coen and Meyerowitz, 1991; Weigel and Meyerowitz, 1994; Purugganan et al., 1995; Rounsley et al., 1995; Theißen and Saedler, 1995; Theißen et al., 2000; Theißen, 2001).The function of B group genes, such as APETALA3 (AP3) and PISTILLATA (PI), has been thought to have a major role in specifying petal and stamen development (Jack et al., 1992; Goto and Meyerowitz, 1994; Krizek and Meyerowitz, 1996; Kramer et al., 1998; Hernandez-Hernandez et al., 2007; Kanno et al., 2007; Whipple et al., 2007; Irish, 2009). In Arabidopsis (Arabidopsis thaliana), mutation in AP3 or PI caused identical phenotypes of second whorl petal conversion into a sepal structure and third flower whorl stamen into a carpel structure (Bowman et al., 1989; Jack et al., 1992; Goto and Meyerowitz, 1994). Similar homeotic conversions for petal and stamen were observed in the mutants of the AP3 and PI orthologs from a number of core eudicots such as Antirrhinum majus, Petunia hybrida, Gerbera hybrida, Solanum lycopersicum, and Nicotiana benthamiana (Sommer et al., 1990; Tröbner et al., 1992; Angenent et al., 1993; van der Krol et al., 1993; Yu et al., 1999; Liu et al., 2004; Vandenbussche et al., 2004; de Martino et al., 2006), from basal eudicot species such as Papaver somniferum and Aquilegia vulgaris (Drea et al., 2007; Kramer et al., 2007), as well as from monocot species such as Zea mays and Oryza sativa (Ambrose et al., 2000; Nagasawa et al., 2003; Prasad and Vijayraghavan, 2003; Yadav et al., 2007; Yao et al., 2008). This indicated that the function of the B class genes AP3 and PI is highly conserved during evolution.It has been thought that B group genes may have arisen from an ancestral gene through multiple gene duplication events (Doyle, 1994; Theißen et al., 1996, 2000; Purugganan, 1997; Kramer et al., 1998; Kramer and Irish, 1999; Lamb and Irish, 2003; Kim et al., 2004; Stellari et al., 2004; Zahn et al., 2005; Hernandez-Hernandez et al., 2007). In the gymnosperms, there was a single putative B class lineage that duplicated to generate the paleoAP3 and PI lineages in angiosperms (Kramer et al., 1998; Theißen et al., 2000; Irish, 2009). The paleoAP3 lineage is composed of AP3 orthologs identified in lower eudicots, magnolid dicots, and monocots (Kramer et al., 1998). Genes in this lineage contain the conserved paleoAP3- and PI-derived motifs in the C-terminal end of the proteins, which have been thought to be characteristics of the B class ancestral gene (Kramer et al., 1998; Tzeng and Yang, 2001; Hsu and Yang, 2002). The PI lineage is composed of PI orthologs that contain a highly conserved PI motif identified in most plant species (Kramer et al., 1998). Subsequently, there was a second duplication at the base of the core eudicots that produced the euAP3 and TM6 lineages, which have been subject to substantial sequence changes in eudicots during evolution (Kramer et al., 1998; Kramer and Irish, 1999). The paleoAP3 motif in the C-terminal end of the proteins was retained in the TM6 lineage and replaced by a conserved euAP3 motif in the euAP3 lineage of most eudicot species (Kramer et al., 1998). In addition, many lineage-specific duplications for paleoAP3 lineage have occurred in plants such as orchids (Hsu and Yang, 2002; Tsai et al., 2004; Kim et al., 2007; Mondragón-Palomino and Theißen, 2008, 2009; Mondragón-Palomino et al., 2009), Ranunculaceae, and Ranunculales (Kramer et al., 2003; Di Stilio et al., 2005; Shan et al., 2006; Kramer, 2009).Unlike the A or C class MADS box proteins, which form homodimers that regulate flower development, the ability of B class proteins to form homodimers has only been reported in gymnosperms and in the paleoAP3 and PI lineages of some monocots. For example, LMADS1 of the lily Lilium longiflorum (Tzeng and Yang, 2001), OMADS3 of the orchid Oncidium Gower Ramsey (Hsu and Yang, 2002), and PeMADS4 of the orchid Phalaenopsis equestris (Tsai et al., 2004) in the paleoAP3 lineage, LRGLOA and LRGLOB of the lily Lilium regale (Winter et al., 2002), TGGLO of the tulip Tulipa gesneriana (Kanno et al., 2003), and PeMADS6 of the orchid P. equestris (Tsai et al., 2005) in the PI lineage, and GGM2 of the gymnosperm Gnetum gnemon (Winter et al., 1999) were able to form homodimers that regulate flower development. Proteins in the euAP3 lineage and in most paleoAP3 lineages were not able to form homodimers and had to interact with PI to form heterodimers in order to regulate petal and stamen development in various plant species (Schwarz-Sommer et al., 1992; Tröbner et al., 1992; Riechmann et al., 1996; Moon et al., 1999; Winter et al., 2002; Kanno et al., 2003; Vandenbussche et al., 2004; Yao et al., 2008). In addition to forming dimers, AP3 and PI were able to interact with other MADS box proteins, such as SEPALLATA1 (SEP1), SEP2, and SEP3, to regulate petal and stamen development (Pelaz et al., 2000; Honma and Goto, 2001; Theißen and Saedler, 2001; Castillejo et al., 2005).Orchids are among the most important plants in the flower market around the world, and research on MADS box genes has been reported for several species of orchids during the past few years (Lu et al., 1993, 2007; Yu and Goh, 2000; Hsu and Yang, 2002; Yu et al., 2002; Hsu et al., 2003; Tsai et al., 2004, 2008; Xu et al., 2006; Guo et al., 2007; Kim et al., 2007; Chang et al., 2009). Unlike the flowers in eudicots, the nearly identical shape of the sepals and petals as well as the production of a unique lip in orchid flowers make them a very special plant species for the study of flower development. Four clades (1–4) of genes in the paleoAP3 lineage have been identified in several orchids (Hsu and Yang, 2002; Tsai et al., 2004; Kim et al., 2007; Mondragón-Palomino and Theißen, 2008, 2009; Mondragón-Palomino et al., 2009). Several works have described the possible interactions among these four clades of paleoAP3 genes and one PI gene that are involved in regulating the differentiation and formation of the sepal/petal/lip of orchids (Tsai et al., 2004; Kim et al., 2007; Mondragón-Palomino and Theißen, 2008, 2009). However, the exact mechanism that involves the orchid B class genes remains unclear and needs to be clarified by more experimental investigations.O. Gower Ramsey is a popular orchid with important economic value in cut flower markets. Only a few studies have been reported on the role of MADS box genes in regulating flower formation in this plant species (Hsu and Yang, 2002; Hsu et al., 2003; Chang et al., 2009). An AP3-like MADS gene that regulates both floral formation and initiation in transgenic Arabidopsis has been reported (Hsu and Yang, 2002). In addition, four AP1/AGAMOUS-LIKE9 (AGL9)-like MADS box genes have been characterized that show novel expression patterns and cause different effects on floral transition and formation in Arabidopsis (Hsu et al., 2003; Chang et al., 2009). Compared with other orchids, the production of a large and well-expanded lip and five small identical sepals/petals makes O. Gower Ramsey a special case for the study of the diverse functions of B class MADS box genes during evolution. Therefore, the isolation of more B class MADS box genes and further study of their roles in the regulation of perianth (sepal/petal/lip) formation during O. Gower Ramsey flower development are necessary. In addition to the clade 2 paleoAP3 gene OMADS3, which was previously characterized in our laboratory (Hsu and Yang, 2002), three more B class MADS box genes, OMADS5, OMADS8, and OMADS9, were characterized from O. Gower Ramsey in this study. Based on the different expression patterns and the protein interactions among these four orchid B class genes, we propose that the presence of OMADS3/8 and/or OMADS9 is required for sepal/petal/lip formation. Further sepal and petal formation at least requires the additional presence of OMADS5, whereas large lip formation was seen when OMADS5 expression was absent. Our results provide a new finding and information pertaining to the roles for orchid B class MADS box genes in the regulation of sepal/petal/lip formation.     

Focus on Ethylene: Ethylene and the Regulation of Physiological and Morphological Responses to Nutrient Deficiencies

To cope with nutrient deficiencies, plants develop both morphological and physiological responses. The regulation of these responses is not totally understood, but some hormones and signaling substances have been implicated. It was suggested several years ago that ethylene participates in the regulation of responses to iron and phosphorous deficiency. More recently, its role has been extended to other deficiencies, such as potassium, sulfur, and others. The role of ethylene in so many deficiencies suggests that, to confer specificity to the different responses, it should act through different transduction pathways and/or in conjunction with other signals. In this update, the data supporting a role for ethylene in the regulation of responses to different nutrient deficiencies will be reviewed. In addition, the results suggesting the action of ethylene through different transduction pathways and its interaction with other hormones and signaling substances will be discussed.When plants suffer from a mineral nutrient deficiency, they develop morphological and physiological responses (mainly in their roots) aimed to facilitate the uptake and mobilization of the limiting nutrient. After the nutrient has been acquired in enough quantity, these responses need to be switched off to avoid toxicity and conserve energy. In recent years, different plant hormones (e.g. ethylene, auxin, cytokinins, jasmonic acid, abscisic acid, brassinosteroids, GAs, and strigolactones) have been implicated in the regulation of these responses (Romera et al., 2007, 2011, 2015; Liu et al., 2009; Rubio et al., 2009; Kapulnik et al., 2011; Kiba et al., 2011; Iqbal et al., 2013; Zhang et al., 2014).Before the 1990s, there were several publications relating ethylene and nutrient deficiencies (cited in Lynch and Brown [1997] and Romera et al. [1999]) without establishing a direct implication of ethylene in the regulation of nutrient deficiency responses. In 1994, Romera and Alcántara (1994) published an article in Plant Physiology suggesting a role for ethylene in the regulation of Fe deficiency responses. In 1999, Borch et al. (1999) showed the participation of ethylene in the regulation of P deficiency responses. Since then, evidence has been accumulating in support of a role for ethylene in the regulation of both Fe (Romera et al., 1999, 2015; Waters and Blevins, 2000; Lucena et al., 2006; Waters et al., 2007; García et al., 2010, 2011, 2013, 2014; Yang et al., 2014) and P deficiency responses (Kim et al., 2008; Lei et al., 2011; Li et al., 2011; Nagarajan and Smith, 2012; Wang et al., 2012, 2014c). Both Fe and P may be poorly available in most soils, and plants develop similar responses under their deficiencies (Romera and Alcántara, 2004; Zhang et al., 2014). More recently, a role for ethylene has been extended to other deficiencies, such as K (Shin and Schachtman, 2004; Jung et al., 2009; Kim et al., 2012), S (Maruyama-Nakashita et al., 2006; Wawrzyńska et al., 2010; Moniuszko et al., 2013), and B (Martín-Rejano et al., 2011). Ethylene has also been implicated in both N deficiency and excess (Tian et al., 2009; Mohd-Radzman et al., 2013; Zheng et al., 2013), and its participation in Mg deficiency has been suggested (Hermans et al., 2010).In this update, we will review the information supporting a role for ethylene in the regulation of different nutrient deficiency responses. For information relating ethylene to other aspects of plant mineral nutrition, such as N₂ fixation and responses to excess of nitrate or essential heavy metals, the reader is referred to other reviews (for review, see Maksymiec, 2007; Mohd-Radzman et al., 2013; Steffens, 2014).     

Subunit Q Is Required to Stabilize the Large Complex of NADPH Dehydrogenase in Synechocystis sp. Strain PCC 6803

Two major complexes of NADPH dehydrogenase (NDH-1) have been identified in cyanobacteria. A large complex (NDH-1L) contains NdhD1, NdhF1, and NdhP, which are absent in a medium size complex (NDH-1M). They play important roles in respiration, NDH-1-dependent cyclic electron transport around photosystem I, and CO₂ uptake. Two mutants sensitive to high light for growth and impaired in cyclic electron transport around photosystem I were isolated from the cyanobacterium Synechocystis sp. strain PCC 6803 transformed with a transposon-bearing library. Both mutants had a tag in an open reading frame encoding a product highly homologous to NdhQ, a single-transmembrane small subunit of the NDH-1L complex, identified in Thermosynechococcus elongatus by proteomics strategy. Deletion of ndhQ disassembled about one-half of the NDH-1L to NDH-1M and consequently impaired respiration, but not CO₂ uptake. During prolonged incubation of the thylakoid membrane with n-dodecyl-β-d-maltoside at room temperature, the rest of the NDH-1L in ΔndhQ was disassembled completely to NDH-1M and was much faster than in the wild type. In the ndhP-deletion mutant (ΔndhP) background, absence of NdhQ almost completely disassembled the NDH-1L to NDH-1M, similar to the results observed in the ΔndhD1/ΔndhD2 mutant. We therefore conclude that both NdhQ and NdhP are essential to stabilize the NDH-1L complex.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., 2011a) and participate in a variety of bioenergetic reactions, such as respiration, cyclic electron transport around PSI, 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 few years, significant achievements have been made in resolving the subunit compositions and functions of the multiple NDH-1 complexes in several cyanobacterial strains (for review, see Battchikova and Aro, 2007; Ogawa and Mi, 2007; Ma, 2009; Battchikova et al., 2011b; Ma and Ogawa, 2015). Four types of NDH-1 have been identified in the cyanobacterium Synechocystis sp. strain PCC 6803 (hereafter, Synechocystis 6803), and all four types of NDH-1 are involved in NDH-1-dependent cyclic electron transport (CET) around PSI (NDH-CET; Bernát et al., 2011). 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 high light-sensitive growth phenotypes (Endo et al., 1999; Battchikova et al., 2011a; Dai et al., 2013; Zhang et al., 2014; Zhao et al., 2014). Therefore, high light strategy can help in identifying the proteins essential to NDH-CET.Proteomics studies revealed the presence of three major NDH-1 complexes in cyanobacteria: a large complex (NDH-1L), a medium size complex (NDH-1M), and a small complex (NDH-1S) with molecular masses of about 460, 350, and 200 kD, respectively (Herranen et al., 2004). NDH-1M consists of 14 subunits (i.e. NdhA–NdhC, NdhE, NdhG–NdhO, and NdhS). In addition to these subunits, the NDH-1L complex contains NdhD1, NdhF1, NdhP, and NdhQ (Prommeenate et al., 2004; Battchikova et al., 2005, 2011b; Zhang et al., 2005, 2014; Nowaczyk et al., 2011; Wulfhorst et al., 2014; Ma and Ogawa, 2015) and is involved in respiration (Zhang et al., 2004). NDH-1S is composed of NdhD3, NdhF3, CO₂ uptake A (CupA), and CupS (Ogawa and Mi, 2007) and is considered to be associated with NDH-1M in the cells as a functional complex NDH-1MS (Zhang et al., 2004, 2005) participating in CO₂ uptake. Among the several copies of ndhD and ndhF genes found in cyanobacterial genomes, ndhD1 and ndhF1 show the highest homology to chloroplast ndhD and ndhF genes, respectively, and CupA and CupS subunits of the cyanobacteria have no counterparts in higher plants. These facts suggest that the structure and composition of NDH-1L, but not the NDH-1MS complex, are similar to those of the chloroplast NDH-1 complex (Battchikova and Aro, 2007; Ogawa and Mi, 2007; Shikanai, 2007; Ma, 2009; Suorsa et al., 2009; Battchikova et al., 2011b; Ifuku et al., 2011; Peng et al., 2011a; Ma and Ogawa, 2015). Despite their similarity, a large number of subunits that constitute the chloroplast NDH-1 complex, including ferredoxin-binding subcomplex subunits NdhT and NdhU and all the subunits of subcomplex B and lumen subcomplex, are absent in the cyanobacterial NDH-1L complex (Battchikova et al., 2011b; Ifuku et al., 2011; Peng et al., 2011a). This implies that the stabilization strategies for the cyanobacterial NDH-1L complex and chloroplastic NDH-1 complex might be significantly different.Recently, a new oxygenic photosynthesis-specific small subunit NdhQ was identified in the NDH-1L complex purified by Ni²⁺ affinity chromatography from Thermosynechococcus elongatus (Nowaczyk et al., 2011). NdhQ is extensively present in cyanobacteria, but its homolog is absent in higher plants (Nowaczyk et al., 2011). In this study, we demonstrate that deletion of NdhQ disassembled the NDH-1L into NDH-1M, but not NDH-1MS, in Synechocystis 6803 and consequently impaired respiration, but not CO₂ uptake. NdhQ and NdhP stabilize the NDH-1L complex. Thus, the stabilization strategy of cyanobacterial NDH-1L is distinctly different from that of the chloroplastic NDH-1 complex.     

Surveying Rubisco Diversity and Temperature Response to Improve Crop Photosynthetic Efficiency

The threat to global food security of stagnating yields and population growth makes increasing crop productivity a critical goal over the coming decades. One key target for improving crop productivity and yields is increasing the efficiency of photosynthesis. Central to photosynthesis is Rubisco, which is a critical but often rate-limiting component. Here, we present full Rubisco catalytic properties measured at three temperatures for 75 plants species representing both crops and undomesticated plants from diverse climates. Some newly characterized Rubiscos were naturally “better” compared to crop enzymes and have the potential to improve crop photosynthetic efficiency. The temperature response of the various catalytic parameters was largely consistent across the diverse range of species, though absolute values showed significant variation in Rubisco catalysis, even between closely related species. An analysis of residue differences among the species characterized identified a number of candidate amino acid substitutions that will aid in advancing engineering of improved Rubisco in crop systems. This study provides new insights on the range of Rubisco catalysis and temperature response present in nature, and provides new information to include in models from leaf to canopy and ecosystem scale.In a changing climate and under pressure from a population set to hit nine billion by 2050, global food security will require massive changes to the way food is produced, distributed, and consumed (Ort et al., 2015). To match rising demand, agricultural production must increase by 50 to 70% in the next 35 years, and yet the gains in crop yields initiated by the green revolution are slowing, and in some cases, stagnating (Long and Ort, 2010; Ray et al., 2012). Among a number of areas being pursued to increase crop productivity and food production, improving photosynthetic efficiency is a clear target, offering great promise (Parry et al., 2007; von Caemmerer et al., 2012; Price et al., 2013; Ort et al., 2015). As the gatekeeper of carbon entry into the biosphere and often acting as the rate-limiting step of photosynthesis, Rubisco, the most abundant enzyme on the planet (Ellis, 1979), is an obvious and important target for improving crop photosynthetic efficiency.Rubisco is considered to exhibit comparatively poor catalysis, in terms of catalytic rate, specificity, and CO₂ affinity (Tcherkez et al., 2006; Andersson, 2008), leading to the suggestion that even small increases in catalytic efficiency may result in substantial improvements to carbon assimilation across a growing season (Zhu et al., 2004; Parry et al., 2013; Galmés et al., 2014a; Carmo-Silva et al., 2015). If combined with complimentary changes such as optimizing other components of the Calvin Benson or photorespiratory cycles (Raines, 2011; Peterhansel et al., 2013; Simkin et al., 2015), optimized canopy architecture (Drewry et al., 2014), or introducing elements of a carbon concentrating mechanism (Furbank et al., 2009; Lin et al., 2014a; Hanson et al., 2016; Long et al., 2016), Rubisco improvement presents an opportunity to dramatically increase the photosynthetic efficiency of crop plants (McGrath and Long, 2014; Long et al., 2015; Betti et al., 2016). A combination of the available strategies is essential for devising tailored solutions to meet the varied requirements of different crops and the diverse conditions under which they are typically grown around the world.Efforts to engineer an improved Rubisco have not yet produced a “super Rubisco” (Parry et al., 2007; Ort et al., 2015). However, advances in engineering precise changes in model systems continue to provide important developments that are increasing our understanding of Rubisco catalysis (Spreitzer et al., 2005; Whitney et al., 2011a, 2011b; Morita et al., 2014; Wilson et al., 2016), regulation (Andralojc et al., 2012; Carmo-Silva and Salvucci, 2013; Bracher et al., 2015), and biogenesis (Saschenbrecker et al., 2007; Whitney and Sharwood, 2008; Lin et al., 2014b; Hauser et al., 2015; Whitney et al., 2015).A complementary approach is to understand and exploit Rubisco natural diversity. Previous characterization of Rubisco from a limited number of species has not only demonstrated significant differences in the underlying catalytic parameters, but also suggests that further undiscovered diversity exists in nature and that the properties of some of these enzymes could be beneficial if present in crop plants (Carmo-Silva et al., 2015). Recent studies clearly illustrate the variation possible among even closely related species (Galmés et al., 2005, 2014b, 2014c; Kubien et al., 2008; Andralojc et al., 2014; Prins et al., 2016).Until recently, there have been relatively few attempts to characterize the consistency, or lack thereof, of temperature effects on in vitro Rubisco catalysis (Sharwood and Whitney, 2014), and often studies only consider a subset of Rubisco catalytic properties. This type of characterization is particularly important for future engineering efforts, enabling specific temperature effects to be factored into any attempts to modify crops for a future climate. In addition, the ability to coanalyze catalytic properties and DNA or amino acid sequence provides the opportunity to correlate sequence and biochemistry to inform engineering studies (Christin et al., 2008; Kapralov et al., 2011; Rosnow et al., 2015). While the amount of gene sequence information available grows rapidly with improving technology, knowledge of the corresponding biochemical variation resulting has yet to be determined (Cousins et al., 2010; Carmo-Silva et al., 2015; Sharwood and Whitney, 2014; Nunes-Nesi et al., 2016).This study aimed to characterize the catalytic properties of Rubisco from diverse species, comprising a broad range of monocots and dicots from diverse environments. The temperature dependence of Rubisco catalysis was evaluated to tailor Rubisco engineering for crop improvement in specific environments. Catalytic diversity was analyzed alongside the sequence of the Rubisco large subunit gene, rbcL, to identify potential catalytic switches for improving photosynthesis and productivity. In vitro results were compared to the average temperature of the warmest quarter in the regions where each species grows to investigate the role of temperature in modulating Rubisco catalysis.     

Integrating Image-Based Phenomics and Association Analysis to Dissect the Genetic Architecture of Temporal Salinity Responses in Rice

Salinity affects a significant portion of arable land and is particularly detrimental for irrigated agriculture, which provides one-third of the global food supply. Rice (Oryza sativa), the most important food crop, is salt sensitive. The genetic resources for salt tolerance in rice germplasm exist but are underutilized due to the difficulty in capturing the dynamic nature of physiological responses to salt stress. The genetic basis of these physiological responses is predicted to be polygenic. In an effort to address this challenge, we generated temporal imaging data from 378 diverse rice genotypes across 14 d of 90 mm NaCl stress and developed a statistical model to assess the genetic architecture of dynamic salinity-induced growth responses in rice germplasm. A genomic region on chromosome 3 was strongly associated with the early growth response and was captured using visible range imaging. Fluorescence imaging identified four genomic regions linked to salinity-induced fluorescence responses. A region on chromosome 1 regulates both the fluorescence shift indicative of the longer term ionic stress and the early growth rate decline during salinity stress. We present, to our knowledge, a new approach to capture the dynamic plant responses to its environment and elucidate the genetic basis of these responses using a longitudinal genome-wide association model.Nearly one-third of the 54 million ha of the highly saline soils in the world are located in South and Southeast Asia. Rice (Oryza sativa), which is the primary source of calories and protein for these two regions, is very sensitive to salinity stress, with even moderate salinity levels known to decrease yields by 50% (Zeng et al., 2002). Projected sea level rise due to climate change is expected to increase saltwater ingress in coastal rice-growing regions of South and Southeast Asia. Therefore, development of salt-tolerant rice cultivars is essential to maintain rice productivity in the salinity-affected regions globally.Salt tolerance, defined as the ability to maintain growth and productivity in saline conditions, is a complex polygenic trait that may be influenced by distinct physiological mechanisms (Munns et al., 1982; Munns and Termaat, 1986; Cheeseman, 1988; Munns and Tester, 2008; Horie et al., 2012; for a comprehensive review of genes involved in salinity tolerance in rice, see Negrão et al., 2011) At the cellular level, plants respond to saline conditions in two phases, namely an osmotic (shoot ion independent) and an ionic stress phase, which can occur in an overlapping manner with varying intensity during the course of salinity stress (Munns and Termaat, 1986; Munns, 2002; Munns and James, 2003; Munns and Tester, 2008; Horie et al., 2012). During the osmotic stress phase, which occurs soon after the onset of salinity, the reduction in external osmotic potential disrupts water uptake and impedes cell expansion, which, at the whole plant level, leads to reduced growth rate (Matsuda and Riazi, 1981; Munns and Passioura, 1984; Rawson and Munns, 1984; Azaizeh and Steudle, 1991; Fricke and Peters, 2002; Fricke, 2004; Boursiac et al., 2005). As salinity stress persists over several days and weeks, sodium ions (Na⁺) accumulate to toxic levels, resulting in cell death and precocious leaf senescence (Lutts and Bouharmont, 1996; Munns, 2002; Munns and James, 2003; Ghanem et al., 2008). This is typically observed during the ionic phase of the salinity response (Munns, 2002; Munns and James, 2003; Munns and Tester, 2008). Plants possess distinct mechanisms to adapt to these osmotic and ionic stresses that are controlled by a suite of genes (Apse et al., 1999; Carvajal et al., 1999; Halfter et al., 2000; Ishitani et al., 2000; Shi et al., 2000; Zeng and Shannon, 2000; Rus et al., 2001; Berthomieu et al., 2003; Martínez-Ballesta et al., 2003; Boursiac et al., 2005, 2008; Ren et al., 2005; Huang et al., 2006; Davenport et al., 2007; Obata et al., 2007; Székely et al., 2008; Horie et al., 2011; Rivandi et al., 2011; Asano et al., 2012; Munns et al., 2012; Latz et al., 2013; Schmidt et al., 2013; Campo et al., 2014; Choi et al., 2014; Liu et al., 2014). The genetic basis of temporal adaptive responses to salinity stress remains to be explored in rice and other crops. This is primarily due to challenges in capturing the dynamic physiological responses to salinity for a large number of genotypes in a nondestructive manner. Manual phenotyping to detect incremental changes in growth rate during the osmotic stress and slight shifts in leaf color due to ionic stress is difficult to quantify for a large number of genotypes.In rice, at least one major quantitative trait loci (QTL; saltol) for salinity tolerance has been characterized based on end point measurements of biomass, senescence/injury, and Na⁺ and K⁺ concentrations (Bonilla et al., 2002; Lin et al., 2004; Thomson et al., 2010). SHOOT K⁺ CONTENT1 (SKC1) is the causative gene underlying the saltol region. SKC1 encodes a Na⁺-selective high-affinity potassium transporter that regulates Na⁺/K⁺ homeostasis during salinity stress (Ren et al., 2005). High levels of Na⁺ displace cellular K⁺, an essential element for several enzymatic reactions and physiological processes (Gierth and Mäser, 2007). The ability to maintain cellular K⁺ levels during salinity through the action of Na⁺-selective potassium transporters or Na⁺/H⁺ antiporters is a well-characterized tolerance mechanism in cereals including rice (Ren et al., 2005; Sunarpi et al., 2005; Huang et al., 2006; Møller et al., 2009; Mian et al., 2011; Munns et al., 2012).Numerous studies have utilized conventional linkage mapping to identify QTL for morphological and physiological responses to salinity in rice using discrete end point measurements (Bonilla et al., 2002; Lin et al., 2004; Ren et al., 2005; Negrão et al., 2011; Wang et al., 2012). However, the physiological adaptation to saline conditions is a complex continuous process that develops over time. While some accessions will exhibit similar end point phenotypic values, the genetic and physiological mechanisms giving rise to the similar phenotypes may be very different and the growth trajectories throughout the experiment may be distinct. Although single time point studies have yielded important information regarding the genetic basis of salinity tolerance, such approaches are too simple to reveal the genetic architecture of stress adaptation. With the advent of high-throughput image-based phenotyping platforms, it is now feasible to quantify dynamic responses during the stress treatment for a large number of genotypes (Berger et al., 2010; Golzarian et al., 2011; Chen et al., 2014; Honsdorf et al., 2014).Image-based phenotyping has been combined with genome-wide association studies (GWAS) and linkage mapping to examine the genetic basis of complex developmental processes (Busemeyer et al., 2013; Moore et al., 2013; Topp et al., 2013; Slovak et al., 2014; Würschum et al., 2014; Yang et al., 2014; Bac-Molenaar et al., 2015). Moreover, the introduction of the time axis provides a better understanding of the physiological processes underlying complex stress and developmental responses compared with single end point measurements (Zhang et al., 2012; Moore et al., 2013; Brown et al., 2014; Chen et al., 2014; Slovak et al., 2014; Bac-Molenaar et al., 2015). However, to date, no studies have implemented an association mapping approach using image-derived phenotypes to address the genetic basis of dynamic stress responses in plants. Image-based phenotyping offers several advantages over conventional phenotyping: (1) quantitative measurements can be recorded over discrete time points to capture morphological and physiological responses in a nondestructive manner, and (2) the use of various types of spectral imaging address phenotypes that are not detectable to the human eye such as chlorophyll fluorescence and leaf water content. Integrating dynamic phenotypic data and association mapping has the potential to query genetic diversity across hundreds of accessions for complex traits and provides much higher resolution compared with conventional linkage mapping. Here, we explored the dynamic growth and chlorophyll responses to salinity of a diverse set of rice accessions using high-throughput visible and fluorescence imaging. To assess the genetic basis of plant growth in saline conditions, a logistic model was used to describe the temporal growth responses and was incorporated into the statistical framework necessary for association mapping. Coupled with temporal fluorescence imaging, we present, to our knowledge, new insights into the genetic architecture of osmotic and ionic responses during salinity stress in rice.     

Abscisic Acid Deficiency Causes Changes in Cuticle Permeability and Pectin Composition That Influence Tomato Resistance to Botrytis cinerea

A mutant of tomato (Solanum lycopersicum) with reduced abscisic acid (ABA) production (sitiens) exhibits increased resistance to the necrotrophic fungus Botrytis cinerea. This resistance is correlated with a rapid and strong hydrogen peroxide-driven cell wall fortification response in epidermis cells that is absent in tomato with normal ABA production. Moreover, basal expression of defense genes is higher in the mutant compared with the wild-type tomato. Given the importance of this fast response in sitiens resistance, we investigated cell wall and cuticle properties of the mutant at the chemical, histological, and ultrastructural levels. We demonstrate that ABA deficiency in the mutant leads to increased cuticle permeability, which is positively correlated with disease resistance. Furthermore, perturbation of ABA levels affects pectin composition. sitiens plants have a relatively higher degree of pectin methylesterification and release different oligosaccharides upon inoculation with B. cinerea. These results show that endogenous plant ABA levels affect the composition of the tomato cuticle and cell wall and demonstrate the importance of cuticle and cell wall chemistry in shaping the outcome of this plant-fungus interaction.Plant defense against pathogens often involves the induction of mechanisms after pathogen recognition, including defense signaling, cell wall strengthening, and localized cell death, but plants also have preformed chemical and structural defense barriers. Fungal pathogens that penetrate the plant tissue directly through the outer surface, rather than via natural plant openings or wounds, must pass through the plant cuticle and epidermal cell wall. Penetration of the host surface happens either by physical means (i.e. by a highly localized pressure in the appressorium) or by chemical means (i.e. by the release of hydrolyzing enzymes). Necrotrophic plant pathogens like Botrytis cinerea typically use the latter strategy. During penetration, they produce cutinases and pectinolytic enzymes such as pectin methylesterases, endopolygalacturonases, and exopolygalacturonases (van Kan, 2006).The cuticle is a hydrophobic barrier that covers the aerial surfaces of the plant. It is mainly composed of cutin, a polyester matrix, and soluble waxes, a complex mixture of hydrophobic material containing very-long-chain fatty acids and their derivatives, embedded into and deposited onto the cutin matrix. It plays an important role in organ development and protection against water loss (Yephremov et al., 1999; Sieber et al., 2000; Kurata et al., 2003; Jung et al., 2006). The cuticle is generally considered as a mere passive physical barrier against pathogen invasion, but it has also been recognized as a potential source of signaling and elicitor molecules (Jenks et al., 1994; Reina-Pinto and Yephremov, 2009). Plant cutin monomers trigger cutinase secretion in pathogenic fungi (Woloshuk and Kolattukudy, 1986), and cutin and wax components initiate appressorium formation and penetration in appressorium-forming pathogens (Kolattukudy et al., 1995; Francis et al., 1996; Gilbert et al., 1996; Fauth et al., 1998; Dickman et al., 2003). In plants, cutin monomers induce pathogenesis-related gene expression and elicit hydrogen peroxide (H₂O₂) synthesis (Fauth et al., 1998; Kim et al., 2008; Park et al., 2008). Transgenic tomato (Solanum lycopersicum) plants expressing the yeast Δ-9 desaturase gene had high levels of cutin monomers that inhibited powdery mildew (Erysiphe polygoni) spore germination, leading to enhanced resistance (Wang et al., 2000). Arabidopsis (Arabidopsis thaliana) plants expressing a fungal cutinase or mutants with a defective cuticle, such as long-chain acyl-CoA synthetase2 and bodyguard, are generally more susceptible to bacteria and equally susceptible to biotrophic fungi but are surprisingly resistant to B. cinerea (Bessire et al., 2007; Chassot et al., 2007; Tang et al., 2007). It has been postulated that a defective or thin cuticle encourages these plants to constitutively express defense-related mechanisms and to secrete antifungal compounds to the plant surface, thereby inhibiting B. cinerea growth (Bessire et al., 2007; Chassot et al., 2007). In addition, cuticle metabolic pathways might directly modulate plant-pathogen interactions by interacting with hormonally regulated defense pathways (Fiebig et al., 2000; Garbay et al., 2007; Mang et al., 2009) or with complex lipid signaling pathways leading to hypersensitive cell death (Raffaele et al., 2008).Once plant pathogens have penetrated the cuticle, they secrete hydrolases that target the plant cell wall (ten Have et al., 1998; Oeser et al., 2002; Vogel et al., 2002; Jakob et al., 2007) that is mainly composed of cellulose, hemicellulose, and pectin (35% of total dry weight). Pectin consists mainly of the polysaccharides homogalacturonan and rhamnogalacturonan I and II. Homogalacturonans are linear chains of α-(1–4)-linked d-GalA residues that can be methylesterified at C-6. Rhamnogalacturonan I and II are more complex, branched polysaccharides. B. cinerea is typically regarded as a pectinolytic pathogen because it possesses an efficient pectinolytic machinery, including a variety of polygalacturonases and pectin methylesterases (PMEs), some of which are important virulence factors (ten Have et al., 1998, 2001; Valette-Collet et al., 2003; Kars et al., 2005). Pectins are a rich source of oligogalacturonides (OGAs), biologically active signaling molecules that can activate plant defense mechanisms (Hahn et al., 1981; Côté and Hahn, 1994; Messiaen and Van Cutsem, 1994; Ridley et al., 2001). The eliciting capacity of the OGAs was shown to depend on their size, which in turn is influenced by the methylesterification pattern of the homogalacturonan fraction (Mathieu et al., 1991; Messiaen and Van Cutsem, 1994). To counteract the activity of fungal pectinases, many plants express polygalacturonase-inhibiting proteins and PME inhibitors, which are localized in the cell wall. The role of these proteins in plant defense against B. cinerea has been extensively demonstrated (Powell et al., 2000; Ferrari et al., 2003; Sicilia et al., 2005; Joubert et al., 2006, 2007; Lionetti et al., 2007). The interaction with the inhibitors not only limits the destructive potential of polygalacturonases but also leads to the accumulation of elicitor-active OGAs (De Lorenzo and Ferrari, 2002). How OGAs are perceived by the plant is still unclear, but in view of the diversity of biological activities and structure requirements, they are thought to be recognized through different proteins, including receptor-like kinases, wall-associated kinases, arabinogalactan proteins, and Pro-rich proteins (Côté and Hahn, 1994; Showalter, 2001; Humphrey et al., 2007).Over the past years, the role of abscisic acid (ABA) in plant-pathogen interactions has gained increased attention. ABA is mostly negatively correlated with resistance against phytopathogens through down-regulation of defense responses orchestrated by salicylic acid, jasmonic acid, and ethylene (Mohr and Cahill, 2001; Audenaert et al., 2002; Mauch-Mani and Mauch, 2005; Asselbergh et al., 2008). In tomato, the ABA-deficient mutant sitiens has an enhanced resistance to B. cinerea (Audenaert et al., 2002) that depends on a timely, localized oxidative burst leading to rapid epidermal cell wall fortification and a faster and higher induction of defense-related gene expression upon infection compared with the wild type (Asselbergh et al., 2007). Moreover, basal defense gene expression is higher in this mutant than in the wild type. As this early response is of vital importance for the resistant reaction of tomato against B. cinerea, we investigated whether alterations in cuticle and/or cell wall, which form the first barrier to the invading pathogen, affect resistance. We demonstrate that the sitiens cuticle is more permeable and that permeability is positively correlated with resistance to B. cinerea. Furthermore, differences in pectin composition and rate of methylesterification occur. Together, these data hint at an unanticipated role for extracellular matrix components in the resistance of tomato against B. cinerea and thus shed new light on the largely unexplored interrelationship between the extracellular matrix and plant-pathogen interactions.     

The Stromal Chloroplast Deg7 Protease Participates in the Repair of Photosystem II after Photoinhibition in Arabidopsis

Light is the ultimate source of energy for photosynthesis; however, excessive light leads to photooxidative damage and hence reduced photosynthetic efficiency, especially when combined with other abiotic stresses. Although the photosystem II (PSII) reaction center D1 protein is the primary target of photooxidative damage, other PSII core proteins are also damaged and degraded. However, it is still largely unknown whether degradation of D1 and other PSII proteins involves previously uncharacterized proteases. Here, we show that Deg7 is peripherally associated with the stromal side of the thylakoid membranes and that Deg7 interacts directly with PSII. Our results show that Deg7 is involved in the primary cleavage of photodamaged D1, D2, CP47, and CP43 and that this activity is essential for its function in PSII repair. The double mutants deg5 deg7 and deg8 deg7 showed no obvious phenotypic differences under normal growth conditions, but additive effects were observed under high light. These results suggest that Deg proteases on both the stromal and luminal sides of the thylakoid membranes are important for the efficient PSII repair in Arabidopsis (Arabidopsis thaliana).Chloroplasts of higher plants carry out one of the most important biochemical reactions: the capture of light energy and its conversion into chemical energy. Although light is the ultimate source of energy for photosynthesis, it can also be harmful to plants. Light-induced loss of photosynthetic efficiency, which is generally termed as photoinhibition, limits plant growth and lowers productivity, especially when combined with other abiotic stresses.The main target of photoinhibition is PSII, which catalyzes the light-dependent water oxidation concomitantly with oxygen production (for review, see Prasil et al., 1992; Aro et al., 1993; Adir et al., 2003). In higher plants, PSII consists of more than 20 subunits, including the reaction center D1 and D2 proteins, cytochrome (Cyt) b₅₅₉, the light-harvesting chlorophyll a-binding proteins CP47 and CP43, the oxygen-evolving 33-kD protein (PsbO), and several low molecular mass proteins (Nelson and Yocum, 2006). The PSII reaction center D1 protein has been identified among PSII proteins as the primary target of light-induced damage (Kyle et al., 1984; Mattoo et al., 1984; Ohad et al., 1984; Adir et al., 1990), but several studies have shown that the D2, CP47, and CP43 proteins are degraded under photoinhibitory conditions (Schuster et al., 1988; Yamamoto and Akasaka, 1995; Jansen et al., 1999; Adir et al., 2003). Moreover, several small PSII subunits, such as PsbH, PsbW, and Cyt b₅₅₉, were also found to be frequently replaced within PSII (Hagman et al., 1997; Ortega et al., 1999; Bergantino et al., 2003). Evidence for the involvement of two families of proteases, FtsH and Deg, in the degradation of the D1 protein in thylakoids of higher plants has been recently described (Lindahl et al., 1996, 2000; Bailey et al., 2002; Sakamoto et al., 2003; Silva et al., 2003; Kapri-Pardes et al., 2007; Sun et al., 2007a, 2007b). However, it is still largely unknown whether degradation of D1 and other PSII proteins involves previously uncharacterized proteases.DegP (or HtrA) proteases were initially identified based on the fact that they are required for the survival of Escherichia coli at high temperatures and for the degradation of abnormal periplasmic proteins (Lipinska et al., 1988; Strauch and Beckwith, 1988). DegP is an ATP-independent Ser endopeptidase, and it contains a trypsin-like protease domain at the N terminus, followed by two PDZ domains (Gottesman, 1996; Pallen and Wren, 1997; Clausen et al., 2002). PDZ domains appear to be important for complex assembly and substrate binding through three or four residues in the C terminus of their target proteins (Doyle et al., 1996; Harris and Lim, 2001). DegP switches between chaperone and protease functions in a temperature-dependent manner. The chaperone function dominates at low temperatures, and DegP becomes proteolytically active at elevated temperatures (Spiess et al., 1999). Crystal structures of different members of the DegP protein family (Krojer et al., 2002; Li et al., 2002; Kim et al., 2003; Wilken et al., 2004) have revealed the structure-function relationship of these PDZ-containing proteases. Trimeric DegP is the functional unit, and the hexameric DegP is formed via the staggered association of trimers (Clausen et al., 2002; Kim and Kim, 2005). At normal growth temperatures, the active site of the protease is located within the chamber of hexameric DegP, which is not accessible to the substrates. However, at high temperatures, conformational changes induce the activation of the protease function (Krojer et al., 2002). Recent studies have shed light on the substrate binding-induced formation of larger oligomeric complexes of DegP (Jiang et al., 2008; Krojer et al., 2008).In Arabidopsis (Arabidopsis thaliana), 16 genes coding for DegP-like proteases have been identified, and at least seven gene products are predicted to be located in chloroplasts (Kieselbach and Funk, 2003; Huesgen et al., 2005; Adam et al., 2006; Sakamoto, 2006; Kato and Sakamoto, 2009). Based on proteomic data, four Deg proteases have been shown to be localized to the chloroplast (Peltier et al., 2002; Schubert et al., 2002) and functionally characterized. Deg1, Deg5, and Deg8 are located in thylakoid lumen, and Deg2 is peripherally associated with the stromal side of thylakoid membranes (Itzhaki et al., 1998; Haußühl et al., 2001; Sun et al., 2007a). Recombinant DegP1, now renamed Deg1, has been shown to be proteolytically active toward thylakoid lumen proteins such as plastocyanin and PsbO of PSII in vitro (Chassin et al., 2002). A 5.2-kD C-terminal fragment of the D1 protein was detected in vitro after incubation of recombinant Deg1 with inside-out thylakoid membranes. In transgenic plants with reduced levels of Deg1, fewer of its 16- and 5.2-kD degradation products were observed (Kapri-Pardes et al., 2007). Deg5 and Deg8 form a dodecameric complex in the thylakoid lumen, and recombinant Deg8 is able to degrade the photodamaged D1 protein of PSII in an in vitro assay (Sun et al., 2007a). The 16-kD N-terminal degradation fragment of the D1 protein was detected in wild-type plants but not in a deg5 deg8 double mutant after high-light treatment. The deg5 deg8 double mutant showed increased sensitivity to high light and high temperature in terms of growth and PSII activity compared with the single mutants deg5 and deg8, suggesting that Deg5 and Deg8 have overlapping functions in the primary cleavage of the CD loop of the D1 protein (Sun et al., 2007a, 2007b). In vitro analysis has demonstrated that recombinant stroma-localized Deg2 was also shown to be involved in the primary cleavage of the DE loop of the D1 protein (Haußühl et al., 2001). However, analysis of a mutant lacking Deg2 suggested that Deg2 may not be involved in D1 degradation in vivo (Huesgen et al., 2006).Here, we have expressed and purified a recombinant DegP protease, His-Deg7. In vitro experiments showed that His-Deg7 is proteolytically active toward the PSII proteins D1, D2, CP43, and CP47. In vivo analyses of a deg7 mutant revealed that the mutant is more sensitive to high light stress than the wild-type plants. We demonstrated that Deg7 is a chloroplast stroma protein associated with the thylakoid membranes and that it interacts with PSII, which suggests that it can cleave the stroma-exposed region of substrate proteins. Our results also provide evidence that Deg7 is important for maintaining PSII function.     

Focus Issue on Calcium Signaling: Identification of Cyclic GMP-Activated Nonselective Ca2+-Permeable Cation Channels and Associated CNGC5 and CNGC6 Genes in Arabidopsis Guard Cells

Cytosolic Ca²⁺ in guard cells plays an important role in stomatal movement responses to environmental stimuli. These cytosolic Ca²⁺ increases result from Ca²⁺ influx through Ca²⁺-permeable channels in the plasma membrane and Ca²⁺ release from intracellular organelles in guard cells. However, the genes encoding defined plasma membrane Ca²⁺-permeable channel activity remain unknown in guard cells and, with some exceptions, largely unknown in higher plant cells. Here, we report the identification of two Arabidopsis (Arabidopsis thaliana) cation channel genes, CNGC5 and CNGC6, that are highly expressed in guard cells. Cytosolic application of cyclic GMP (cGMP) and extracellularly applied membrane-permeable 8-Bromoguanosine 3′,5′-cyclic monophosphate-cGMP both activated hyperpolarization-induced inward-conducting currents in wild-type guard cells using Mg²⁺ as the main charge carrier. The cGMP-activated currents were strongly blocked by lanthanum and gadolinium and also conducted Ba²⁺, Ca²⁺, and Na⁺ ions. cngc5 cngc6 double mutant guard cells exhibited dramatically impaired cGMP-activated currents. In contrast, mutations in CNGC1, CNGC2, and CNGC20 did not disrupt these cGMP-activated currents. The yellow fluorescent protein-CNGC5 and yellow fluorescent protein-CNGC6 proteins localize in the cell periphery. Cyclic AMP activated modest inward currents in both wild-type and cngc5cngc6 mutant guard cells. Moreover, cngc5 cngc6 double mutant guard cells exhibited functional abscisic acid (ABA)-activated hyperpolarization-dependent Ca²⁺-permeable cation channel currents, intact ABA-induced stomatal closing responses, and whole-plant stomatal conductance responses to darkness and changes in CO₂ concentration. Furthermore, cGMP-activated currents remained intact in the growth controlled by abscisic acid2 and abscisic acid insensitive1 mutants. This research demonstrates that the CNGC5 and CNGC6 genes encode unique cGMP-activated nonselective Ca²⁺-permeable cation channels in the plasma membrane of Arabidopsis guard cells.Plants lose water via transpiration and take in CO₂ for photosynthesis through stomatal pores. Each stomatal pore is surrounded by two guard cells, and stomatal movements are driven by the change of turgor pressure in guard cells. The intracellular second messenger Ca²⁺ functions in guard cell signal transduction (Schroeder and Hagiwara, 1989; McAinsh et al., 1990; Webb et al., 1996; Grabov and Blatt, 1998; Allen et al., 1999; MacRobbie, 2000; Mori et al., 2006; Young et al., 2006; Siegel et al., 2009; Chen et al., 2010; Hubbard et al., 2012). Plasma membrane ion channel activity and gene expression in guard cells are finely regulated by the intracellular free calcium concentration ([Ca2+]_cyt; Schroeder and Hagiwara, 1989; Webb et al., 2001; Allen et al., 2002; Siegel et al., 2009; Kim et al., 2010; Stange et al., 2010). Ca²⁺-dependent protein kinases (CPKs) function as targets of the cytosolic Ca²⁺ signal, and several members of the CPK family have been shown to function in stimulus-induced stomatal closing, including the Arabidopsis (Arabidopsis thaliana) CPK3, CPK4, CPK6, CPK10, and CPK11 proteins (Mori et al., 2006; Zhu et al., 2007; Zou et al., 2010; Brandt et al., 2012; Hubbard et al., 2012). Further research found that several CPKs could activate the S-type anion channel SLAC1 in Xenopus laevis oocytes, including CPK21, CPK23, and CPK6 (Geiger et al., 2010; Brandt et al., 2012). At the same time, the Ca²⁺-independent protein kinase Open Stomata1 mediates stomatal closing and activates the S-type anion channel SLAC1 (Mustilli et al., 2002; Yoshida et al., 2002; Geiger et al., 2009; Lee et al., 2009; Xue et al., 2011), indicating that both Ca²⁺-dependent and Ca²⁺-independent pathways function in guard cells.Multiple essential factors of guard cell abscisic acid (ABA) signal transduction function in the regulation of Ca²⁺-permeable channels and [Ca2+]_cyt elevations, including Abscisic Acid Insensitive1 (ABI1), ABI2, Enhanced Response to Abscisic Acid1 (ERA1), the NADPH oxidases AtrbohD and AtrbohF, the Guard Cell Hydrogen Peroxide-Resistant1 (GHR1) receptor kinase, as well as the Ca²⁺-activated CPK6 protein kinase (Pei et al., 1998; Allen et al., 1999, 2002; Kwak et al., 2003; Miao et al., 2006; Mori et al., 2006; Hua et al., 2012). [Ca2+]_cyt increases result from both Ca²⁺ release from intracellular Ca²⁺ stores (McAinsh et al., 1992) and Ca²⁺ influx across the plasma membrane (Hamilton et al., 2000; Pei et al., 2000; Murata et al., 2001; Kwak et al., 2003; Hua et al., 2012). Electrophysiological analyses have characterized nonselective Ca²⁺-permeable channel activity in the plasma membrane of guard cells (Schroeder and Hagiwara, 1990; Hamilton et al., 2000; Pei et al., 2000; Murata et al., 2001; Köhler and Blatt, 2002; Miao et al., 2006; Mori et al., 2006; Suh et al., 2007; Vahisalu et al., 2008; Hua et al., 2012). However, the genetic identities of Ca²⁺-permeable channels in the plasma membrane of guard cells have remained unknown despite over two decades of research on these channel activities.The Arabidopsis genome includes 20 genes encoding cyclic nucleotide-gated channel (CNGC) homologs and 20 genes encoding homologs to animal Glu receptor channels (Lacombe et al., 2001; Kaplan et al., 2007; Ward et al., 2009), which have been proposed to function in plant cells as cation channels (Schuurink et al., 1998; Arazi et al., 1999; Köhler et al., 1999). Recent research has demonstrated functions of specific Glu receptor channels in mediating Ca²⁺ channel activity (Michard et al., 2011; Vincill et al., 2012). Previous studies have shown cAMP activation of nonselective cation currents in guard cells (Lemtiri-Chlieh and Berkowitz, 2004; Ali et al., 2007). However, only a few studies have shown the disappearance of a defined plasma membrane Ca²⁺ channel activity in plants upon mutation of candidate Ca²⁺ channel genes (Ali et al., 2007; Michard et al., 2011; Laohavisit et al., 2012; Vincill et al., 2012). Some CNGCs have been found to be involved in cation nutrient intake, including monovalent cation intake (Guo et al., 2010; Caballero et al., 2012), salt tolerance (Guo et al., 2008; Kugler et al., 2009), programmed cell death and pathogen responses (Clough et al., 2000; Balagué et al., 2003; Urquhart et al., 2007; Abdel-Hamid et al., 2013), thermal sensing (Finka et al., 2012; Gao et al., 2012), and pollen tube growth (Chang et al., 2007; Frietsch et al., 2007; Tunc-Ozdemir et al., 2013a, 2013b). Direct in vivo disappearance of Ca²⁺ channel activity in cngc disruption mutants has been demonstrated in only a few cases thus far (Ali et al., 2007; Gao et al., 2012). In this research, we show that CNGC5 and CNGC6 are required for a cyclic GMP (cGMP)-activated nonselective Ca²⁺-permeable cation channel activity in the plasma membrane of Arabidopsis guard cells.     

A Signaling Pathway Linking Nitric Oxide Production to Heterotrimeric G Protein and Hydrogen Peroxide Regulates Extracellular Calmodulin Induction of Stomatal Closure in Arabidopsis

Extracellular calmodulin (ExtCaM) regulates stomatal movement by eliciting a cascade of intracellular signaling events including heterotrimeric G protein, hydrogen peroxide (H₂O₂), and Ca²⁺. However, the ExtCaM-mediated guard cell signaling pathway remains poorly understood. In this report, we show that Arabidopsis (Arabidopsis thaliana) NITRIC OXIDE ASSOCIATED1 (AtNOA1)-dependent nitric oxide (NO) accumulation plays a crucial role in ExtCaM-induced stomatal closure. ExtCaM triggered a significant increase in NO levels associated with stomatal closure in the wild type, but both effects were abolished in the Atnoa1 mutant. Furthermore, we found that ExtCaM-mediated NO generation is regulated by GPA1, the Gα-subunit of heterotrimeric G protein. The ExtCaM-dependent NO accumulation was nullified in gpa1 knockout mutants but enhanced by overexpression of a constitutively active form of GPA1 (cGα). In addition, cGα Atnoa1 and gpa1-2 Atnoa1 double mutants exhibited a similar response as did Atnoa1. The defect in gpa1 was rescued by overexpression of AtNOA1. Finally, we demonstrated that G protein activation of NO production depends on H₂O₂. Reduced H₂O₂ levels in guard cells blocked the stomatal response of cGα lines, whereas exogenously applied H₂O₂ rescued the defect in ExtCaM-mediated stomatal closure in gpa1 mutants. Moreover, the atrbohD/F mutant, which lacks the NADPH oxidase activity in guard cells, had impaired NO generation in response to ExtCaM, and H₂O₂-induced stomatal closure and NO accumulation were greatly impaired in Atnoa1. These findings have established a signaling pathway leading to ExtCaM-induced stomatal closure, which involves GPA1-dependent activation of H₂O₂ production and subsequent AtNOA1-dependent NO accumulation.Plant guard cells control opening and closure of the stomata in response to phytohormones (e.g. abscisic acid [ABA]) and various environmental signals such as light and temperature, thereby regulating gas exchange for photosynthesis and water status via transpiration (Schroeder et al., 2001). Cytosolic calcium ([Ca²⁺]_i) has been shown to be a key second messenger that changes in response to multiple stimuli in guard cells (McAinsh et al., 1995; Grabov and Blatt, 1998; Wood et al., 2000). A large proportion of Ca²⁺ is localized in extracellular space. It has been shown that external Ca²⁺ concentration ([Ca²⁺]_o) promotes stomatal closure and induces oscillation in [Ca²⁺]_i in guard cells (MacRobbie, 1992; McAinsh et al., 1995; Allen et al., 2001). However, how the guard cells perceive [Ca²⁺]_o concentration and convert [Ca²⁺]_o changes into [Ca²⁺]_i changes was not understood until a calcium-sensing receptor (CAS) in the plasma membrane of guard cells in Arabidopsis (Arabidopsis thaliana) was identified (Han et al., 2003). The external Ca²⁺ (Ca²⁺_o)-induced [Ca²⁺]_i increase is abolished in CAS antisense lines (Han et al., 2003). Both [Ca²⁺]_o and [Ca²⁺]_i show diurnal oscillation that is determined by stomatal conductance, whereas the amplitude of [Ca²⁺]_i oscillation is reduced in CAS antisense lines (Tang et al., 2007). The reduced amplitude of [Ca²⁺]_i diurnal oscillation in response to Ca²⁺_o treatment suggests the potential existence of other [Ca²⁺]_o sensor(s) that may transmit [Ca²⁺]_o information into the [Ca²⁺]_i response in coordination with CAS. Extracellular calmodulin (ExtCaM) could be such an additional [Ca²⁺]_o sensor.Calmodulin is a well-known Ca²⁺ sensor that is activated upon binding of Ca²⁺. It has been shown that calmodulin exists not only intracellularly but also extracellularly in many plant species (Biro et al., 1984; Sun et al., 1994, 1995; Cui et al., 2005). ExtCaM has been implicated in several important biological functions, such as the promotion of cell proliferation, pollen germination, and tube growth (Sun et al., 1994, 1995; Ma and Sun, 1997; Ma et al., 1999; Cui et al., 2005; Shang et al., 2005). ExtCaM is found in the cell wall of guard cells in Vicia faba and in the epidermis of Arabidopsis by immunogold labeling/electron microscopy and western-blot analyses, respectively, and the endogenous CaM in the extracellular space has been shown to regulate stomatal movements (Chen et al., 2003; Xiao et al., 2004). Under natural conditions, once the activity of ExtCaM has been inhibited by its membrane-impermeable antagonist W7-agrose or CaM antibody, stomatal opening under light is enhanced and stomatal closure in darkness is inhibited in V. faba and Arabidopsis (Chen et al., 2003; Xiao et al., 2004). [Ca²⁺]_i and cytosolic hydrogen peroxide (H₂O₂) changes, two events involved in ExtCaM-regulated stomatal movement (Chen et al., 2004), are likely regulated by light/darkness (Chen and Gallie, 2004; Tang et al., 2007), suggesting that ExtCaM plays an important physiological role in the regulation of stomatal diurnal rhythm. Calmodulin-binding proteins have been found in the protoplast of suspension-cultured Arabidopsis cells, supporting the idea that ExtCaM functions as a peptide-signaling molecule (Cui et al., 2005). Furthermore, ExtCaM triggers [Ca²⁺]_i elevation in guard cells of V. faba and Arabidopsis and in lily (Lilium daviddi) pollen (Chen et al., 2004; Xiao et al., 2004; Shang et al., 2005). These observations support the notion that ExtCaM could be a potential [Ca²⁺]_o sensor for external calcium, and this external calcium sensing could subsequently regulate the [Ca²⁺]_i level through a signaling cascade.It is interesting that ExtCaM and ABA induce some parallel changes in second messengers in guard cell signaling. Our previous studies show that ExtCaM induces [Ca²⁺]_i increase and H₂O₂ generation through the Gα-subunit (GPA1) of a heterotrimeric G protein, and increased H₂O₂ further elevates [Ca²⁺]_i (Chen et al., 2004). G protein, Ca²⁺, and H₂O₂ are well-known second messengers in ABA-induced guard cell signaling (McAinsh et al., 1995; Grabov and Blatt, 1998; Pei et al., 2000; Wang et al., 2001; Zhang et al., 2001; Liu et al., 2007). However, the signaling cascade triggered by ExtCaM in guard cells is poorly understood. New ABA signaling components in guard cells could provide a clue in the study of the molecular mechanism of ExtCaM guard cell signaling.Recently, nitric oxide (NO) has been shown to serve as an important signal molecule involved in many aspects of developmental processes, including floral transition, root growth, root gravitropism, adventitious root formation, xylogenesis, seed germination, and orientation of pollen tube growth (Beligni and Lamattina, 2000; Pagnussat et al., 2002; He et al., 2004; Prado et al., 2004; Gabaldón et al., 2005; Stohr and Stremlau, 2006). Increasing evidence points to a role for NO as an essential component in ABA signaling in guard cells (Garcia-Mata and Lamattina, 2001, 2002; Neill et al., 2002). It has been shown that nitrate reductase (NR) reduces nitrite to NO, and the nia1, nia2 NR-deficient mutant in Arabidopsis showed reduced ABA induction of stomatal closure (Desikan et al., 2002; Bright et al., 2006). Although animal nitric oxide synthase (NOS) activity has been detected in plants and inhibitors of mammalian NOS impair NO production in plants (Barroso et al., 1999; Corpas et al., 2001), the gene(s) encoding NOS in plants is still not clear. AtNOS1 in Arabidopsis was initially reported to encode a protein containing NOS activity (Guo et al., 2003). However, recent studies have raised critical questions regarding the nature of AtNOS1 and suggested that AtNOS1 appears not to encode a NOS (Crawford et al., 2006; Zemojtel et al., 2006). However, the originally described Atnos1 mutant is deficient in NO accumulation (Crawford et al., 2006). Consequently, AtNOS1 was renamed AtNOA1 (for NITRIC OXIDE ASSOCIATED1; Crawford et al., 2006). Therefore, the Atnoa1 mutant provides a useful tool for dissecting the function of NO in plants. At present, the molecules that regulate NO generation in ABA-mediated guard cell signaling are not clear. Evidence suggests that H₂O₂, a second messenger important for the regulation of many developmental processes and stomatal movement (Pei et al., 2000; Zhang et al., 2001; Coelho et al., 2002; Demidchik et al., 2003; Kwak et al., 2003), regulates NO generation in guard cells (Lum et al., 2002; He et al., 2005; Bright et al., 2006).Given the parallel signaling events induced by ABA and ExtCaM, we investigated whether NO is involved in the regulation of ExtCaM-induced stomatal closure in Arabidopsis and whether it is linked to G protein and H₂O₂, two key regulators of both ExtCaM and ABA regulation of stomatal movements. Using Arabidopsis mutants (e.g. GPA1 null mutants, the NO-producing mutant Atnoa1, and the guard cell H₂O₂ synthetic enzymatic mutant atrbohD/F) combined with pharmacological analysis, we present compelling evidence to establish a linear functional relationship between Gα, H₂O₂, and NO in ExtCaM guard cell signaling.