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.     

Calcineurin B-Like Protein-Interacting Protein Kinase CIPK21 Regulates Osmotic and Salt Stress Responses in Arabidopsis

The role of calcium-mediated signaling has been extensively studied in plant responses to abiotic stress signals. Calcineurin B-like proteins (CBLs) and CBL-interacting protein kinases (CIPKs) constitute a complex signaling network acting in diverse plant stress responses. Osmotic stress imposed by soil salinity and drought is a major abiotic stress that impedes plant growth and development and involves calcium-signaling processes. In this study, we report the functional analysis of CIPK21, an Arabidopsis (Arabidopsis thaliana) CBL-interacting protein kinase, ubiquitously expressed in plant tissues and up-regulated under multiple abiotic stress conditions. The growth of a loss-of-function mutant of CIPK21, cipk21, was hypersensitive to high salt and osmotic stress conditions. The calcium sensors CBL2 and CBL3 were found to physically interact with CIPK21 and target this kinase to the tonoplast. Moreover, preferential localization of CIPK21 to the tonoplast was detected under salt stress condition when coexpressed with CBL2 or CBL3. These findings suggest that CIPK21 mediates responses to salt stress condition in Arabidopsis, at least in part, by regulating ion and water homeostasis across the vacuolar membranes.Drought and salinity cause osmotic stress in plants and severely affect crop productivity throughout the world. Plants respond to osmotic stress by changing a number of cellular processes (Xiong et al., 1999; Xiong and Zhu, 2002; Bartels and Sunkar, 2005; Boudsocq and Lauriére, 2005). Some of these changes include activation of stress-responsive genes, regulation of membrane transport at both plasma membrane (PM) and vacuolar membrane (tonoplast) to maintain water and ionic homeostasis, and metabolic changes to produce compatible osmolytes such as Pro (Stewart and Lee, 1974; Krasensky and Jonak, 2012). It has been well established that a specific calcium (Ca²⁺) signature is generated in response to a particular environmental stimulus (Trewavas and Malhó, 1998; Scrase-Field and Knight, 2003; Luan, 2009; Kudla et al., 2010). The Ca²⁺ changes are primarily perceived by several Ca²⁺ sensors such as calmodulin (Reddy, 2001; Luan et al., 2002), Ca²⁺-dependent protein kinases (Harper and Harmon, 2005), calcineurin B-like proteins (CBLs; Luan et al., 2002; Batistič and Kudla, 2004; Pandey, 2008; Luan, 2009; Sanyal et al., 2015), and other Ca²⁺-binding proteins (Reddy, 2001; Shao et al., 2008) to initiate various cellular responses.Plant CBL-type Ca²⁺ sensors interact with and activate CBL-interacting protein kinases (CIPKs) that phosphorylate downstream components to transduce Ca²⁺ signals (Liu et al., 2000; Luan et al., 2002; Batistič and Kudla, 2004; Luan, 2009). In several plant species, multiple members have been identified in the CBL and CIPK family (Luan et al., 2002; Kolukisaoglu et al., 2004; Pandey, 2008; Batistič and Kudla, 2009; Weinl and Kudla, 2009; Pandey et al., 2014). Involvement of specific CBL-CIPK pair to decode a particular type of signal entails the alternative and selective complex formation leading to stimulus-response coupling (D’Angelo et al., 2006; Batistič et al., 2010).Several CBL and CIPK family members have been implicated in plant responses to drought, salinity, and osmotic stress based on genetic analysis of Arabidopsis (Arabidopsis thaliana) mutants (Zhu, 2002; Cheong et al., 2003, 2007; Kim et al., 2003; Pandey et al., 2004, 2008; D’Angelo et al., 2006; Qin et al., 2008; Tripathi et al., 2009; Held et al., 2011; Tang et al., 2012; Drerup et al., 2013; Eckert et al., 2014). A few CIPKs have also been functionally characterized by gain-of-function approach in crop plants such as rice (Oryza sativa), pea (Pisum sativum), and maize (Zea mays) and were found to be involved in osmotic stress responses (Mahajan et al., 2006; Xiang et al., 2007; Yang et al., 2008; Tripathi et al., 2009; Zhao et al., 2009; Cuéllar et al., 2010).In this report, we examined the role of the Arabidopsis CIPK21 gene in osmotic stress response by reverse genetic analysis. The loss-of-function mutant plants became hypersensitive to salt and mannitol stress conditions, suggesting that CIPK21 is involved in the regulation of osmotic stress response in Arabidopsis. These findings are further supported by an enhanced tonoplast targeting of the cytoplasmic CIPK21 through interaction with the vacuolar Ca²⁺ sensors CBL2 and CBL3 under salt stress condition.     

Cavitation Resistance in Seedless Vascular Plants: The Structure and Function of Interconduit Pit Membranes

Plant water transport occurs through interconnected xylem conduits that are separated by partially digested regions in the cell wall known as pit membranes. These structures have a dual function. Their porous construction facilitates water movement between conduits while limiting the spread of air that may enter the conduits and render them dysfunctional during a drought. Pit membranes have been well studied in woody plants, but very little is known about their function in more ancient lineages such as seedless vascular plants. Here, we examine the relationships between conduit air seeding, pit hydraulic resistance, and pit anatomy in 10 species of ferns (pteridophytes) and two lycophytes. Air seeding pressures ranged from 0.8 ± 0.15 MPa (mean ± sd) in the hydric fern Athyrium filix-femina to 4.9 ± 0.94 MPa in Psilotum nudum, an epiphytic species. Notably, a positive correlation was found between conduit pit area and vulnerability to air seeding, suggesting that the rare-pit hypothesis explains air seeding in early-diverging lineages much as it does in many angiosperms. Pit area resistance was variable but averaged 54.6 MPa s m⁻¹ across all surveyed pteridophytes. End walls contributed 52% to the overall transport resistance, similar to the 56% in angiosperm vessels and 64% in conifer tracheids. Taken together, our data imply that, irrespective of phylogenetic placement, selection acted on transport efficiency in seedless vascular plants and woody plants in equal measure by compensating for shorter conduits in tracheid-bearing plants with more permeable pit membranes.Water transport in plants occurs under tension, which renders the xylem susceptible to air entry. This air seeding may lead to the rupture of water columns (cavitation) such that the air expands within conduits to create air-vapor embolisms that block further transport. (Zimmermann and Tyree, 2002). Excessive embolism such as that which occurs during a drought may jeopardize leaf hydration and lead to stomatal closure, overheating, wilting, and possibly death of the plant (Hubbard et al., 2001; Choat et al., 2012; Schymanski et al., 2013). Consequently, strong selection pressure resulted in compartmentalized and redundant plant vascular networks that are adapted to a species habitat water availability by way of life history strategy (i.e. phenology) or resistance to air seeding (Tyree et al., 1994; Mencuccini et al., 2010; Brodersen et al., 2012). The spread of drought-induced embolism is limited primarily by pit membranes, which are permeable, mesh-like regions in the primary cell wall that connect two adjacent conduits. The construction of the pit membrane is such that water easily moves across the membrane between conduits, but because of the small membrane pore size and the presence of a surface coating on the membrane (Pesacreta et al., 2005; Lee et al., 2012), the spread of air and gas bubbles is restricted up to a certain pressure threshold known as the air-seeding pressure (ASP). When xylem sap tension exceeds the air-seeding threshold, air can be aspirated from an air-filled conduit into a functional water-filled conduit through perhaps a large, preexisting pore or one that is created by tension-induced membrane stress (Rockwell et al., 2014). Air seeding leads to cavitation and embolism formation, with emboli potentially propagating throughout the xylem network (Tyree and Sperry, 1988; Brodersen et al., 2013). So, on the one hand, pit membranes are critical to controlling the spread of air throughout the vascular network, while on the other hand, they must facilitate the efficient flow of water between conduits (Choat et al., 2008; Domec et al., 2008; Pittermann et al., 2010; Schulte, 2012). Much is known about such hydraulic tradeoffs in the pit membranes of woody plants, but comparatively little data exist on seedless vascular plants such as ferns and lycophytes. Given that seedless vascular plants may bridge the evolutionary transition from bryophytes to woody plants, the lack of functional data on pit membrane structure in early-derived tracheophytes is a major gap in our understanding of the evolution of plant water transport.In woody plants, pit membranes fall into one of two categories: the torus-margo type found in most gymnosperms and the homogenous pit membrane characteristic of angiosperms (Choat et al., 2008; Choat and Pittermann, 2009). In conifers, water moves from one tracheid to another through the margo region of the membrane, with the torus sealing the pit aperture should one conduit become embolized. Air seeding occurs when water potential in the functional conduit drops low enough to dislodge the torus from its sealing position, letting air pass through the pit aperture into the water-filled tracheid (Domec et al., 2006; Delzon et al., 2010; Pittermann et al., 2010; Schulte, 2012; but see Jansen et al., 2012). Across north-temperate conifer species, larger pit apertures correlate with lower pit resistance to water flow (r_pit; MPa s m⁻¹), but it is the ratio of torus-aperture overlap that sets a species cavitation resistance (Pittermann et al., 2006, 2010; Domec et al., 2008; Hacke and Jansen, 2009). A similar though mechanistically different tradeoff exists in angiosperm pit membranes. Here, air seeding reflects a probabilistic relationship between membrane porosity and the total area of pit membranes present in the vessel walls. Specifically, the likelihood of air aspirating into a functional conduit is determined by the combination of xylem water potential and the diameter of the largest pore and/or the weakest zone in the cellulose matrix in the vessel’s array of pit membranes (Wheeler et al., 2005; Hacke et al., 2006; Christman et al., 2009; Rockwell et al., 2014). As it has come to be known, the rare-pit hypothesis suggests that the infrequent, large-diameter leaky pore giving rise to that rare pit reflects some combination of pit membrane traits such as variation in conduit membrane area (large or small), membrane properties (tight or porous), and hydrogel membrane chemistry (Hargrave et al., 1994; Choat et al., 2003; Wheeler et al., 2005; Hacke et al., 2006; Christman et al., 2009; Lee et al., 2012; Plavcová et al., 2013; Rockwell et al., 2014). The maximum pore size is critical because, per the Young-Laplace law, the larger the radius of curvature, the lower the air-water pressure difference under which the contained meniscus will fail (Jarbeau et al., 1995; Choat et al., 2003; Jansen et al., 2009). Consequently, angiosperms adapted to drier habitats may exhibit thicker, denser, smaller, and less abundant pit membranes than plants occupying regions with higher water availability (Wheeler et al., 2005; Hacke et al., 2007; Jansen et al., 2009; Lens et al., 2011; Scholz et al., 2013). However, despite these qualitative observations, there is no evidence that increased cavitation resistance arrives at the cost of higher r_pit. Indeed, the bulk of the data suggest that prevailing pit membrane porosity is decoupled from the presence of the single largest pore that allows air seeding to occur (Choat et al., 2003; Wheeler et al., 2005 Hacke et al., 2006, 2007).As water moves from one conduit to another, pit membranes offer considerable hydraulic resistance throughout the xylem network. On average, r_pit contributes 64% and 56% to transport resistance in conifers and angiosperms, respectively (Wheeler et al., 2005; Pittermann et al., 2006; Sperry et al., 2006). In conifers, the average r_pit is estimated at 6 ± 1 MPa s m⁻¹, almost 60 times lower than the 336 ± 81 MPa s m⁻¹ computed for angiosperms (Wheeler et al., 2005; Hacke et al., 2006; Sperry et al., 2006). Presumably, the high porosity of conifer pits compensates for the higher transport resistance offered by a vascular system composed of narrow, short, single-celled conduits (Pittermann et al., 2005; Sperry et al., 2006).Transport in seedless vascular plants presents an interesting conundrum because, with the exception of a handful of species, their primary xylem is composed of tracheids, the walls of which are occupied by homogenous pit membranes (Gibson et al., 1985; Carlquist and Schneider, 2001, 2007; but see Morrow and Dute, 1998, for torus-margo membranes in Botrychium spp.). At first pass, this combination of traits appears hydraulically maladaptive, but several studies have shown that ferns can exhibit transport capacities that are on par with more recently evolved plants (Wheeler et al., 2005; Watkins et al., 2010; Pittermann et al., 2011, 2013; Brodersen et al., 2012). Certainly, several taxa possess large-diameter, highly overlapping conduits, some even have vessels such as Pteridium aquilinum and many species have high conduit density, all of which could contribute to increased hydraulic efficiency (Wheeler et al., 2005; Pittermann et al., 2011, 2013). But how do the pit membranes of seedless vascular plants compare? Scanning electron micrographs of fern and lycopod xylem conduits suggest that they are thin, diaphanous, and susceptible to damage during specimen preparation (Carlquist and Schneider 2001, 2007). Consistent with such observations, two estimates of r_pit imply that r_pit in ferns may be significantly lower than in angiosperms; Wheeler et al. (2005) calculated r_pit in the fern Pteridium aquilinum at 31 MPa s m⁻¹, while Schulte et al. (1987) estimated r_pit at 1.99 MPa s m⁻¹ in the basal fern Psilotum nudum. The closest structural analogy to seedless vascular plant tracheids can be found in the secondary xylem of the early-derived vesselless angiosperms, in which tracheids possess homogenous pit membranes with r_pit values that at 16 MPa s m⁻¹ are marginally higher than those of conifers (Hacke et al., 2007). Given that xylem in seedless vascular plants is functionally similar to that in vesselless angiosperms, we expected convergent r_pit values in these two groups despite their phylogenetic distance. We tested this hypothesis, as well as the intrinsic cavitation resistance of conduits in seedless vascular plants, by scrutinizing the pit membranes of ferns and fern allies using the anatomical and experimental approaches applied previously to woody taxa. In particular, we focused on the relationship between pit membrane traits and cavitation resistance at the level of the individual conduit.     

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).     

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.     

Tyloses and Phenolic Deposits in Xylem Vessels Impede Water Transport in Low-Lignin Transgenic Poplars: A Study by Cryo-Fluorescence Microscopy

Of 14 transgenic poplar genotypes (Populus tremula × Populus alba) with antisense 4-coumarate:coenzyme A ligase that were grown in the field for 2 years, five that had substantial lignin reductions also had greatly reduced xylem-specific conductivity compared with that of control trees and those transgenic events with small reductions in lignin. For the two events with the lowest xylem lignin contents (greater than 40% reduction), we used light microscopy methods and acid fuchsin dye ascent studies to clarify what caused their reduced transport efficiency. A novel protocol involving dye stabilization and cryo-fluorescence microscopy enabled us to visualize the dye at the cellular level and to identify water-conducting pathways in the xylem. Cryo-fixed branch segments were planed in the frozen state on a sliding cryo-microtome and observed with an epifluorescence microscope equipped with a cryo-stage. We could then distinguish clearly between phenolic-occluded vessels, conductive (stain-filled) vessels, and nonconductive (water- or gas-filled) vessels. Low-lignin trees contained areas of nonconductive, brown xylem with patches of collapsed cells and patches of noncollapsed cells filled with phenolics. In contrast, phenolics and nonconductive vessels were rarely observed in normal colored wood of the low-lignin events. The results of cryo-fluorescence light microscopy were supported by observations with a confocal microscope after freeze drying of cryo-planed samples. Moreover, after extraction of the phenolics, confocal microscopy revealed that many of the vessels in the nonconductive xylem were blocked with tyloses. We conclude that reduced transport efficiency of the transgenic low-lignin xylem was largely caused by blockages from tyloses and phenolic deposits within vessels rather than by xylem collapse.Secondary xylem in woody plants is a complex vascular tissue that functions in mechanical support, conduction, storage, and protection (Carlquist, 2001; Tyree and Zimmermann, 2002). The xylem must provide a sufficient and safe water supply throughout the entire pathway from roots to leaves for transpiration and photosynthesis. It is well established that enhanced water conductivity of xylem can increase total plant carbon gain (Domec and Gartner, 2003; Santiago et al., 2004; Brodribb and Holbrook, 2005a). According to the Hagen-Poiseuille equation, xylem conductivity should scale with vessel lumen diameter to the fourth power (Tyree and Zimmermann, 2002). Indeed, xylem conductivity largely depends on anatomical features, including conduit diameters and frequencies (Salleo et al., 1985; McCulloh and Sperry, 2005). However, there are hydraulic limits to maximum vessel diameters, because xylem conduits have to withstand the strong negative pressures of the transpiration stream that could cause cell collapse or embolisms within vessels that are structurally inadequate to withstand these forces (Tyree and Sperry, 1989; Lo Gullo et al., 1995; Hacke et al., 2000). To some extent, stomatal regulation of transpiration limits the negative pressures that the xylem experiences (Tardieu and Davies, 1993; Cochard et al., 2002; Meinzer, 2002; Brodribb and Holbrook, 2004; Buckley, 2005; Franks et al., 2007; Woodruff et al., 2007). Nevertheless, plants rely on an array of structural reinforcements of xylem to ensure the safety of water transport. The size of xylem elements, vessel redundancy, intervessel pit and membrane geometries, and the thickness, microstructure, and chemical composition of cell walls are among the features that regulate tradeoffs between efficiency and safety of xylem water transport (Baas and Schweingruber, 1987; Hacke et al., 2001; Domec et al., 2006; Ewers et al., 2007; Choat et al., 2008).The xylem cell wall is made up of cellulose bundles that are hydrogen bonded with hemicelluloses, which are in turn embedded within a lignin matrix (Mansfield, 2009; Salmén and Burgert, 2009). Besides providing this matrix for the cell wall itself, lignin is thought to contribute to many of the mechanical and physical characteristics of wood as well as conferring passive resistance to the spread of pathogens within a plant (Niklas, 1992; Boyce et al., 2004; Davin et al., 2008). Lignin typically represents 20% to 30% of the dry mass of wood and therefore is among the most abundant stores of carbon in the biosphere (Zobel and van Buijtenen, 1989). The complex molecular structure and biosynthetic pathway of various types of lignins have been studied extensively (Boerjan et al., 2003; Ralph et al., 2004, 2007; Higuchi, 2006; Boudet, 2007; Davin et al., 2008). The monomeric composition of lignin varies between different cell types of the same species depending on the functional specialization of the cell (Yoshinaga et al., 1992; Watanabe et al., 2004; Xu et al., 2006). The composition and amount of lignin in wild plants varies in response to climatic conditions (Donaldson, 2002) or gravitational and mechanical demands (Pruyn et al., 2000; Kern et al., 2005; Rüggeberg et al., 2008). It is clear that plants are capable of regulating the lignification pattern in differentiating cells, which provides them with flexibility for responding to environmental stresses (Donaldson, 2002; Koehler and Telewski, 2006; Ralph et al., 2007; for review, see Vanholme et al., 2008).Whereas some level of lignin is a requisite for all vascular plants, it is often an unwanted product in the pulp and paper industry because it increases the costs of paper production and associated water treatments necessary for environmental protection (Chen et al., 2001; Baucher et al., 2003; Peter et al., 2007). Reducing the lignin content of the raw biomass material may allow more efficient hydrolysis of polysaccharides in biomass and thus facilitate the production of biofuel (Chen and Dixon, 2007). With the ultimate goal of development of wood for more efficient processing, much research has been aimed at the production of genetically modified trees with altered lignin biosynthesis (Boerjan et al., 2003; Boudet et al., 2003; Li et al., 2003; Halpin, 2004; Ralph et al., 2004, 2008; Chiang, 2006; Coleman et al., 2008a, 2008b; Vanholme et al., 2008; Wagner et al., 2009). It is now technically possible to achieve more than 50% reductions of lignin content in xylem of poplar (Populus spp.; Leplé et al., 2007; Coleman et al., 2008a, 2008b), but the consequences of such reduction on plant function have received relatively little attention (Koehler and Telewski, 2006). In-depth studies on the xylem structure and functional performance of transgenic plants with low lignin are limited, despite the need to assess their long-term sustainability for large-scale production (Anterola and Lewis, 2002; Hancock et al., 2007; Coleman et al., 2008b, Voelker, 2009; Horvath et al., 2010).Genetically modified plants are suitable models for studying fundamental questions of the physiological role of lignin because of the possibility of controlling lignification without the confounding effects encountered when comparing across plant tissues or stages of development (Koehler and Telewski, 2006; Leplé et al., 2007; Coleman et al., 2008b). Research on Arabidopsis (Arabidopsis thaliana) and tobacco (Nicotiana tabacum) has shown that down-regulation of lignin biosynthesis can have diverse effects on plant metabolism and structure, including changes in the lignin amount and composition (p-hydroxyphenyl/guaiacyl/syringyl units ratio) as well as the collapse of xylem vessel elements (Lee et al., 1997; Sewalt et al., 1997; Piquemal et al., 1998; Chabannes et al., 2001; Jones et al., 2001; Franke et al., 2002; Dauwe et al., 2007). Among temperate hardwoods, poplar has been established as a model tree for genetic manipulations because of its ecological and economic importance, fast growth, ease of vegetative propagation, and its widespread use in traditional breeding programs (Bradshaw et al., 2001; Brunner et al., 2004). The question of how manipulation of lignin can affect the anatomy and physiological function of xylem in poplar has been addressed in part by several research groups (Anterola and Lewis, 2002; Boerjan et al., 2003; Leplé et al., 2007; Coleman et al., 2008b). Some studies that involved large lignin reductions reported no significant alterations in the xylem anatomy (Hu et al., 1999; Li et al., 2003). However, in many other experiments, reduced total lignin content was associated with significant growth retardation, alterations in the lignin monomer composition, irregularities in the xylem structure (Anterola and Lewis, 2002; Leplé et al., 2007; Coleman et al., 2008b), and the patchy occurrence of collapsed xylem cells (Coleman et al., 2008b; Voelker, 2009). Furthermore, severely down-regulated lignin biosynthesis has resulted in greatly reduced xylem water-transport efficiency (Coleman et al., 2008b; Lachenbruch et al., 2009; Voelker, 2009). It is generally assumed that the reduced water transport ability of xylem with very low lignin contents is caused by collapsed conduits and/or increased embolism due to the entry of air bubbles into the water-conducting cells (Coleman et al., 2008b; Wagner et al., 2009), but detailed anatomical investigations of the causes of impaired xylem conductivity of low-lignin trees are lacking. Analysis of the anatomical basis for the properties of xylem conduits in plants with genetically manipulated amounts and composition of lignin can provide a deeper understanding of the physiological role of lignin as well as the lower limit of down-regulation of lignin biosynthesis at which trees can still survive within natural environments.One of the approaches for the suppression of lignin biosynthesis is down-regulation of 4-coumarate:coenzyme A ligase (4CL), an enzyme that functions in phenylpropanoid metabolism by producing the monolignol precursor p-coumaroyl-CoA (Kajita et al.,1997; Allina et al., 1998; Hu et al., 1998; Harding et al., 2002; Jia et al., 2004; Costa et al., 2005; Friedmann et al., 2007; Wagner et al., 2009). In a 2-year-long field trial on the physiological performance of poplar (Populus tremula × Populus alba) transgenic clones, out of 14 genotypes with altered lignin biosynthesis (down-regulated 4CL), five showed dramatically reduced wood-specific conductivity (k_s) compared with that of control trees (Voelker, 2009). Those mutants with the severely reduced k_s were also characterized by having the lowest wood lignin contents (up to an approximately 40% reduction) in the study. Trees with transgenic events characterized by the formation of abnormally brown wood exhibited regular branch dieback at the end of the growing season, despite having been regularly watered (Voelker, 2009). Our objective was to identify the structural features responsible for reduced transport efficiency in the xylem of transgenic poplars with extremely low lignin contents. We employed fluorescence and laser scanning confocal microscopy for anatomical analyses of xylem structure as well as dye-flow experiments followed by cryo-fluorescence microscopy to visualize the functioning water-conductive pathways in xylem at the cellular level. We report the frequent occurrence of tyloses and phenolic depositions in xylem vessels of strongly down-regulated trees that may be the cause of their reduced xylem conductivity.     

Enhanced Photosynthesis and Growth in atquac1 Knockout Mutants Are Due to Altered Organic Acid Accumulation and an Increase in Both Stomatal and Mesophyll Conductance

Stomata control the exchange of CO₂ and water vapor in land plants. Thus, whereas a constant supply of CO₂ is required to maintain adequate rates of photosynthesis, the accompanying water losses must be tightly regulated to prevent dehydration and undesired metabolic changes. Accordingly, the uptake or release of ions and metabolites from guard cells is necessary to achieve normal stomatal function. The AtQUAC1, an R-type anion channel responsible for the release of malate from guard cells, is essential for efficient stomatal closure. Here, we demonstrate that mutant plants lacking AtQUAC1 accumulated higher levels of malate and fumarate. These mutant plants not only display slower stomatal closure in response to increased CO₂ concentration and dark but are also characterized by improved mesophyll conductance. These responses were accompanied by increases in both photosynthesis and respiration rates, without affecting the activity of photosynthetic and respiratory enzymes and the expression of other transporter genes in guard cells, which ultimately led to improved growth. Collectively, our results highlight that the transport of organic acids plays a key role in plant cell metabolism and demonstrate that AtQUAC1 reduce diffusive limitations to photosynthesis, which, at least partially, explain the observed increments in growth under well-watered conditions.Stomata are functionally specialized microscopic pores that control the essential exchange of CO₂ and H₂O with the environment in land plants. Stomata are found on the surfaces of the majority of the aerial parts of plants, rendering them as the main control point regulating the flow of gases between plants and their surrounding atmosphere. Accordingly, the majority of water loss from plants occurs through stomatal pores, allowing plant transpiration and CO₂ absorption for the photosynthetic process (Bergmann and Sack, 2007; Kim et al., 2010). The maintenance of an adequate water balance through stomatal control is crucial to plants because cell expansion and growth require tissues to remain turgid (Sablowski and Carnier Dornelas, 2014), and minor reductions in cell water volume and turgor pressure will therefore compromise both processes (Thompson, 2005). As a result, the high sensitivity of plant tissues to turgor has prompted the use of reverse genetic studies in attempt to engineer plants with improved performance (Cowan and Troughton, 1971; Xiong et al., 2009; Borland et al., 2014; Franks et al., 2015).In most land plants, not only redox signals invoked by shifts in light quality (Busch, 2014) but also the transport of inorganic ions (e.g. K⁺, Cl⁻, and NO⁻₃) as well as metabolites such as the phytohormone abscisic acid (ABA), Suc, and malate, are important players controlling stomatal movements (Hetherington, 2001; Roelfsema and Hedrich, 2005; Pandey et al., 2007; Blatt et al., 2014; Kollist et al., 2014). In this context, although organic acids in plants is known to support numerous and diverse functions both within and beyond cellular metabolism, only recently have we obtained genetic evidence to support that modulation of guard cell malate and fumarate concentration can greatly influence stomatal movements (Nunes-Nesi et al., 2007; Araújo et al., 2011b; Penfield et al., 2012; Medeiros et al., 2015). Notably malate, in particular, has been considered as a key metabolite and one of the most important organic metabolites involved in guard cell movements (Hedrich and Marten, 1993; Fernie and Martinoia, 2009; Meyer et al., 2010). During stomatal aperture, the flux of malate into guard cells coupled with hexoses generated on starch breakdown lead to decreases in the water potential, and consequently, water uptake by the guard cells ultimately opens the stomata pore (Roelfsema and Hedrich, 2005; Vavasseur and Raghavendra, 2005; Lee et al., 2008). On the other hand, during stomatal closure, malate is believed to be converted into starch, which has no osmotic activity (Penfield et al., 2012) or, alternatively, is released from the guard cells to the surrounding apoplastic space (Lee et al., 2008; Negi et al., 2008; Vahisalu et al., 2008; Meyer et al., 2010).The role of organic acids on the stomatal movements has been largely demonstrated by studies related to malate transport (Lee et al., 2008; Meyer et al., 2010; Sasaki et al., 2010). In the last decade, two protein families were identified and functionally characterized to be directly involved with organic acid transport at the guard cell plasma membrane and to be required for stomatal functioning (Lee et al., 2008; Meyer et al., 2010; Sasaki et al., 2010). In summary, AtABCB14, a member of the ABC (ATP binding cassette) family, which is involved in malate transport from apoplast to guard cells, was described as a negative modulator of stomatal closure induced by high CO₂ concentration; notably, exogenous application of malate minimizes this response (Lee et al., 2008). In addition, members of a small gene family, which encode the anion channels SLAC1 (slow anion channel 1) and four SLAC1-homologs (SLAHs) in Arabidopsis (Arabidopsis thaliana), have been described to be involved in stomatal movements. SLAC1 is a well-documented S-type anion channel that preferentially transports chloride and nitrate as opposed to malate (Vahisalu et al., 2008, 2010; Geiger et al., 2010; Du et al., 2011; Brandt et al., 2012; Kusumi et al., 2012). Lack of SLAC1 in Arabidopsis and rice (Oryza sativa) culminated in a failure in stomatal closure in response to high CO₂ levels, low relative humidity, and dark conditions (Negi et al., 2008; Vahisalu et al., 2008; Kusumi et al., 2012). Although mutations in AtSLAC1 impair S-type anion channel functions as a whole, the R-type anion channel remained functional (Vahisalu et al., 2008). Indeed, a member of the aluminum-activated malate transporter (ALMT) family, AtALMT12, an R-type anion channel, has been demonstrated to be involved in malate transport, particularly at the plasma membrane of guard cells (Meyer et al., 2010; Sasaki et al., 2010). Although AtALMT12 is a member of ALMT family, it is not activated by aluminum, and therefore Meyer et al. (2010) proposed to rename it as AtQUAC1 (quick-activating anion channel 1; Imes et al., 2013; Mumm et al., 2013). Hereafter, we will follow this nomenclature. Deficiency of a functional AtQUAC1 has been documented to lead to changes in stomatal closure in response to high levels of CO₂, dark, and ABA (Meyer et al., 2010). Taken together, these studies have clearly demonstrated that both S- and R-type anion channels are key modulators of stomatal movements in response to several environmental factors.Despite a vast number of studies involving the above-mentioned anion channels, little information concerning the metabolic changes caused by their impairment is currently available. Such information is important to understand stomatal movements, mainly considering that organic acids, especially the levels of malate in apoplastic/mesophyll cells, have been highlighted as of key importance in leaf metabolism (Fernie and Martinoia, 2009; Araújo et al., 2011a, 2011b; Lawson et al., 2014; Medeiros et al., 2015). Here, we demonstrate that a disruption in the expression of AtQUAC1, which leads to impaired stomatal closure (Meyer et al., 2010), was accompanied by increases in mesophyll conductance (g_m), which is defined as the conductance for the transfer of CO₂ from the intercellular airspaces (C_i) to the sites of carboxylation in the chloroplastic stroma (C_c). By further characterization of atquac1 knockout plants, we demonstrated that reduced diffusive limitations resulted in higher photosynthetic rates and altered respiration that, in turn, led to enhanced biomass accumulation. Overall, the results obtained are discussed both in terms of the importance of organic acid transport in plant cell metabolism and with regard to the contribution that it plays in the regulation of both stomatal function and growth.     

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.