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J.I. Schroeder, G. Allen, V. Hugouvieux, J. Kwak, D. Waner (2001) Guard Cell Signal Transduction Annu Rev Plant Physiol Plant Mol Biol 52: 627-658 |
| INTRODUCTION LIGHT SIGNAL TRANSDUCTION AND STOMATAL OPENING Early Events in Blue Light Signal Transduction Cytosolic Factors that Regulate Inward-Rectifying K+in Channels Role of Actin Filaments in Stomatal Movements Cloned Guard Cell K+in Channel Genes MULTIPLE PHYSIOLOGICAL AND ABIOTIC STIMULI INDUCE STOMATAL CLOSING Model for Roles of Ion Channels in ABA-Induced Stomatal Closing CO2-Induced Stomatal Closing STIMULI THAT INCREASE CYTOSOLIC CALCIUM IN GUARD CELLS Roles of [Ca2+]cyt in Stomatal Closing Roles of [Ca2+]cyt in Stomatal Opening SECOND MESSENGER SYSTEMS REGULATING [Ca2+]cyt IN GUARD CELLS Cyclic ADP-Ribose and Vacuolar Ca2+ Release Phospholipase C and Inositol 1,4,5 Trisphosphate Phospholipase D and Phosphatidic Acid Amplifying Calcium Signals by Calcium-Induced Calcium Release Plasma Membrane Calcium Channels and Calcium Influx in Guard Cells Cytosolic [Ca2+] Oscillations are Necessary for Stomatal Closing in Guard Cells ABA-INSENSITIVE PP2C MUTANTS ANION CHANNELS AND ABA-INDUCED STOMATAL CLOSURE Regulation of Guard Cell ABA Signaling by ABC Proteins Protein Kinases Function in ABA Signaling and Anion Channel Regulation Okadaic Acid Sensitive Phosphatases Regulate Anion Channels and ABA Signaling ACTIVITY OF PLASMA MEMBRANE K+OUT CHANNELS IN STOMATAL MOVEMENTS Regulation of K+out by Phosphorylation Syntaxins and ABA Signaling Osmolarity and Temperature Sensitivity Transient K+ Efflux Currents NEW GUARD CELL SIGNALING MUTANTS AND GENETIC APPROACHES New Genetic Screens and Reverse Genetics FUTURE OUTLOOK LITERATURE CITED |
Introduction Opening and closing of stomatal pores is mediated by turgor and volume changes in guard cells. During stomatal opening guard cells accumulate potassium, anions, and sucrose (MacRobbie, 1983; Raschke, 1979; Talbott and Zeiger, 1998). Osmotic water uptake leads to guard cell swelling and stomatal opening. Stomatal closing is mediated by potassium and anion efflux from guard cells, sucrose removal, and metabolism of malate to osmotically inactive starch. Guard cells have become a popular system for dissecting the functions of individual genes and proteins within signaling cascades for the following reasons. 1. Guard cells control CO2 influx and water loss and thus critically affect whole plant growth and physiology.The central role of guard cells in regulating gas exchange is of importance for ecological and biotechnological applications. Agricultural and horticultural use of plants in climates to which these plants are not adapted, as well as short-term climate changes, lead to dramatic crop losses or freshwater consumption under stress conditions such as drought. Recent studies in Arabidopsis have demonstrated that stomatal responses can be manipulated by modifying guard cell signal transduction elements to reduce transpirational water loss and desiccation during drought periods (Gosti et al., 1999; Hugouvieux et al., 2001; Pei et al., 1998). One might ask: Are guard cells different from many other plant cell types (aside from the obvious specializations) in terms of harboring many signal receptors in a single cell? Most likely not, because most individual plant cells respond to many classical hormones, pathogens, and light signals. The combination of the above listed attributes (1 to 4), however, renders guard cells a well-developed model system for interdisciplinary and time-resolved characterizations of mechanisms or segments of early plant signaling cascades. In the present review we focus mainly on recent dissections of signaling transduction mechanisms in guard cells. Of further importance for stomatal movements is signal-dependent modulation of starch-malate metabolism. The mechanisms by which signaling cascades described here tie into metabolic networks is an important frontier of future research and recent reviews on guard cell metabolic pathways can be found elsewhere (Jarvis and Davies, 1998; Netting, 2000; Talbott and Zeiger, 1998). Furthermore, several reviews on aspects of guard cell signal transduction have appeared in recent years (Allen and Sanders, 1997; Assmann, 1999; Assmann and Shimazaki, 1999; Blatt, 1999; Leung and Giraudat, 1998; MacRobbie, 1998; MacAinsh et al., 1997; Müller-Röber et al., 1998; Schroeder et al., 2000; Schroeder et al., 1998; Ward et al., 1995; Zeiger, 2000). Stomatal opening is driven by H+ extrusion through plasma membrane H+ ATPases that are activated by auxins (Lohse and Hedrich, 1992), red light (Serrano, 1998), and blue light (Assmann et al., 1985, Shimazaki et al., 1986). Cell-autonomous light receptors in guard cells induce stomatal opening (Zeiger and Hepler, 1977). Light-induced stomatal opening requires activation of plasma membrane proton (H+)-ATPases (Assmann et al., 1985; Goh et al., 1996; Kinoshita and Shimazaki, 1999; Serrano et al., 1988; Shimazaki et al., 1986), causing plasma membrane hyperpolarization proposed to drive K+ uptake into guard cells via inward-rectifying K+ (K+in) channels (Schroeder et al., 1987; Thiel et al., 1992). Anion (Cl-) influx into stomatal guard cells is thought to occur via H+/anion symporters or anion/OH- antiporters in the plasma membrane. In parallel, starch metabolism leads to accumulation of osmotically active malate in guard cells. In addition, sucrose levels in guard cells increase during light-induced stomatal opening (Lu et al., 1995; Poffenroth et al., 1992; Ritte et al., 1999; Talbott and Zeiger, 1996; Talbott and Zeiger, 1998). Studies with intact leaves show that the main solute supporting stomatal opening at the beginning of a daily cycle was K+, whereas sucrose became predominant later in the daily cycle as guard cell K+ content decreased (Talbott and Zeiger, 1996; Talbott and Zeiger, 1998). Early Events in Blue Light Signal Transduction Cytosolic Ca2+ elevation reversibly inhibits blue light activation of the H+-ATPases (Kinoshita et al., 1995) (K1/2 = 0.3 uM Ca2+). Furthermore, inhibitors of PP1- or PP2A-type (PP1/PP2A) protein phosphatases such as calyculin A and okadaic acid inhibited blue light–dependent H+ pumping and light-induced stomatal opening in Vicia, suggesting that PP1/PP2A phosphatases are positive regulators of light-induced stomatal opening (Kinoshita and Shimazaki, 1997). Abscisic acid (ABA) also inhibits blue light–dependent H+ pumping activity of Vicia guard cell protoplasts (Goh et al., 1996; Roelfsema et al., 1998; Shimazaki et al., 1986). ABA inhibition of apoplastic acidification was not observed in the ABA-insensitive Arabidopsis mutants abi1-1 and abi2-1. Interestingly, the PP1/PP2A inhibitor okadaic acid partially restored ABA inhibition of proton pumping in abi1-1 guard cells, whereas the protein kinase inhibitor K-252a partially restored ABA inhibition of proton pumping in abi2-1 guard cells (Roelfsema et al., 1998). 14-3-3 proteins bind directly to the C-terminal domain and thus activate H+-ATPases, and fusicoccin stabilizes the 14-3-3–H+-ATPase complex (Baunsgaard et al., 1998; Fullone et al., 1998; Jahn et al., 1997). The direct mechanism of guard cell plasma membrane H+-ATPase regulation was characterized in an elegant biochemical study. Blue light activates H+-ATPases via phosphorylation of the C-terminus (Kinoshita and Shimazaki, 1999). Coprecipitation of H+-ATPases with endogenous guard cell 14-3-3 proteins and binding of recombinant 14-3-3 proteins only to the phosphorylated H+-ATPase C-terminus provide evidence for a role of 14-3-3 proteins as a positive regulator in physiological blue light signal transduction. Although two different types of blue light receptors, CRY proteins and NPH1, have been isolated from plants (Ahmad and Cashmore, 1993; Cashmore et al., 1999; Christie et al., 1998), the blue light receptors in guard cells have been proposed to include unique components. Stomata from the zeaxanthin deficient Arabidopsis mutant, npq1 (Niyogi et al., 1998), are impaired in blue light–induced stomatal opening (Frechilla et al., 1999), and stomata from the blue light photoreceptor mutants, cry1, cry2, nph1, nph3, nph4, cry1cry2, and nph1cry1, showed a wild-type blue light response (Frechilla et al., 1999; Lasceve et al., 1999). These results have led to the proposal of a model in which zeaxanthins function as blue light receptors or receptor pigments in guard cells (Frechilla et al., 1999). Subsequently, however, the phototropin PHOT1 and its paralogue PHOT2 were shown to be the guard cell photoreceptors mediating stomatal opening in response to blue light (Kinoshita et al. 2001). PHOT1 was originally identified as photoreceptor mediating blue-light phototropism and named NPH1 for "non-phototropic hypocotyl" (Christie et al. 1998). PHOT1 and PHOT2 are plasma membrane-associated serine/threonine kinases that undergo autophosphorylation upon exposure to blue light (Sakamoto and Briggs, 2002); light-sensing is performed by two flavine mononucleotide moieties bound via LOV (light, oxygen or voltage) domains (Huala et al. 1997). Guard cells of an Arabidopsis phot1,2 double mutant were unresponsive to blue light, while single deletion of either gene did not impair stomatal opening (Kinoshita et al. 2001). Thus PHOT1 and PHOT2 act redundantly in blue-light signal transduction in Arabidopsis guard cells. Cytosolic Factors That Regulate Inward-Rectifying K+in Channels K+in channels in guard cells have been proposed to contribute to K+ uptake during stomatal opening (Schroeder et al., 1984), Schroeder et al., 1987). Reviews on the role of K+in channels in guard cells and on the structure and function of these K+ channels have appeared elsewhere (Czempinski et al., 1999; Dreyer et al., 1999; Hedrich and Dietrich, 1996; Maathuis et al., 1997; Schroeder et al., 1994). Extracellular acidification increases the activity of guard cell K+in channels (Blatt, 1992) and of cloned plant K+in channels expressed in Xenopus oocytes (Ichida and Schroeder, 1996; Müller-Röber et al., 1995; Very et al., 1995). Cytosolic Ca2+ ([Ca2+]cyt) elevation inhibits K+in channels, thus limiting K+ uptake (Lemtiri-Chlieh and MacRobbie, 1994; Schroeder and Hagiwara, 1989). Almost complete inhibition of K+in channels has been measured in Vicia guard cells when [Ca2+]cyt was buffered to about 1 uM (Grabov and Blatt, 1997; Grabov and Blatt, 1999; Kelly et al., 1995; Lemtiri-Chlieh and MacRobbie, 1994; Schroeder and Hagiwara, 1989), or when InsP3 is uncaged in the guard cell cytoplasm to raise [Ca2+]cyt (Blatt et al., 1990). Interestingly, in Vicia, K+in channels in abaxial guard cells were [Ca2+]cyt inhibited and stomatal movements were modulated by external Ca2+ and ABA, whereas K+in channels in adaxial guard cells were insensitive to [Ca2+]cyt and stomatal movements were less sensitive to external Ca2+ and ABA. In addition, protein phosphorylation has been suggested to play an important role in modulation of K+in channel activity. Inhibitors of calcineurin (PP2B), a calcium-dependent protein phosphatase, maintained K+in channel activity in spite of elevated [Ca2+]cyt in Vicia guard cells (Luan et al., 1993). Cyclosporine A, an inhibitor of animal PP2B-type phosphatases, inhibits stomatal closure and reduces ABA inhibition of stomatal opening in Pisum, suggesting that PP2Bs might be negative regulators of stomatal opening (Hey et al., 1997). Note that PP2B homologs have not yet been identified in plants. In contrast to PP2B inhibitors, inhibitors of PP1/PP2A protein phosphatases downregulate K+in channel activity in guard cells, suggesting that PP1 or PP2As are positive regulators of K+in channels (Li et al., 1994; Thiel and Blatt, 1994). Biochemical approaches identified a Vicia guard cell Ca2+-dependent protein kinase (CDPK) that phosphorylates the Arabidopsis guard cell K+in channel subunit KAT1 (Li et al., 1998), and heterologous expression of KAT1 and a soybean CDPK in Xenopus oocytes shows a reduction in KAT1-mediated K+ currents (Berkowitz et al., 2000), suggesting a role of CDPK in Ca2+-mediated K+in channel inhibition. Combined genetic and cell biological analyses will be required to identify the kinases that regulate K+in channels. Stomatal aperture measurements show that linolenic acid and arachidonic acid promote stomatal opening and inhibit stomatal closing (Lee et al., 1994). Furthermore, patch clamp analyses show that these fatty acids activate K+in currents and inhibit outward rectifying K+out currents (Lee et al., 1994). These results are interesting as only a few agents have been found that enhance K+in channel activity in guard cells. Furthermore, phospholipase A2 inhibitors decrease stomatal responses to light (Suh et al., 1998). Together these data suggest that phospholipase A2 may be a positive regulator of the guard cell light response pathway. Role of Actin Filaments in Stomatal Movements Cytochalasin D, an actin filament-depolymerizing agent, activates K+in channels and enhances light-induced stomatal opening (Hwang et al., 1997). In contrast, inhibition of K+in channel currents and light-induced stomatal opening was observed when an actin filament stabilizer, phalloidin, was tested (Hwang et al., 1997). These results imply a possible interaction of the actin cytoskeleton with guard cell plasma membrane K+in channels in signal transduction. ABA treatments reorganize the actin structure from a radial pattern to a randomly oriented and short-fragmented pattern (Eun and Lee, 1997). A small GTP-binding protein, AtRac1, can function as a negative regulator in ABA-induced actin reorganization (Lemichez et al., 2000). ABA causes AtRac1 inactivation. This ABA response was impaired in abi1-1 (Lemichez et al., 2000). In addition, transgenic Arabidopsis plants expressing a dominant negative AtRac1 mutant mimic constitutive ABA-induced actin reorganization in guard cells and increase ABA sensitivity of stomatal closure (Lemichez et al., 2000), suggesting that inactivation of AtRac1 is required for actin reorganization and stomatal closure. Together these studies show important roles of actin and small G-proteins in stomatal movements. Cloned Guard Cell K+in Channel Genes The Arabidopsis K+in channel gene KAT1 was cloned by complementation of a yeast mutant defective in K+ transport (Anderson et al., 1992) and shown to mediate K+ currents in Xenopus oocytes with typical properties of plant K+in currents (Hedrich et al., 1995; Hoshi, 1995; Ichida and Schroeder, 1996; Schachtman et al., 1992; Very et al., 1995). Expression studies with Arabidopsis KAT1 and the potato ortholog KST1 show that these plant K+in channel genes are predominantly expressed in guard cells (Müller-Röber et al., 1995; Nakamura et al., 1995). In animals, functional K+ channel proteins are composed of four alpha–subunits (Jan and Jan, 1992) and additional regulatory beta–subunits (Fink et al., 1996). An Arabidopsis cDNA encoding a beta–subunit homolog of K+ channels has been isolated and binding to KAT1 has been reported (Tang et al., 1996). The model that K+in channels contribute to K+ uptake during stomatal opening (Schroeder et al., 1987) has been analyzed using molecular genetic approaches. In a study with transgenic Arabidopsis expressing KAT1 mutants with a reduced sensitivity to Cs+ block, transgenic plants exhibited partial light-induced stomatal opening in the presence of Cs+ concentrations that inhibit stomatal opening in wildtype (Ichida et al., 1997). In another study, transgenic Arabidopsis expressing a dominant negative mutant form of the guard cell K+in channel KAT1 showed 75% reduction in the activity of guard cell K+in channel currents and a reduction in light-induced stomatal opening (Kwak et al., 2001). As redundancy likely exists in K+ channel subunits in guard cells, these data show that dominant negative kat1 can disrupt K+in channels. These data support the model that K+ channels constitute a central mechanism for K+ uptake during stomatal opening ((Schroeder et al., 1984), Schroeder et al., 1987). These studies do not exclude the likely model that other partially redundant (Schroeder et al., 1994) K+ uptake transporters function in parallel in guard cells. For example, there are 13 isoforms of the KT-KUP-HAK K+ transporter gene family (Fu and Luan, 1998; Kim et al., 1998; Quintero and Blatt, 1997; Santa-Maria et al., 1997) in the Arabidopsis genome and some of these are likely to be expressed in guard cells. Furthermore, patch-clamp studies have shown the activity of additional inward-conducting cation channels (Henriksen et al., 1996; Very et al., 1998, Wu, 1995). In conclusion, many positive and negative regulators of H+-pumps, K+in channels, and stomatal opening have been found. A combination of genetic, cell biological, and biochemical studies will allow further testing and expansion of these models. Abscisic acid, produced in response to water deficit, causes stomatal closing. Furthermore, in C3 and C4 plants stomatal closing is induced by darkness and by elevated CO2 concentrations in the intercellular spaces in leaves, arising from respiration (Assmann, 1999, Mansfield et al., 1990). Stomatal closing and reduced transpiration also leads to elevated temperatures inside leaves. Pathogen elicitors also cause stomatal closing, enabling plants to reduce access of pathogens to the inside of leaves (Lee et al., 1994). Aerial pollutants such as ozone and sulfur dioxide cause stomatal closing at high concentrations (Torsethaugen et al., 1999), thus reducing further damage of leaf tissues by these pollutants. Thus multiple stimuli elicit stomatal closing. Although the reception mechanisms for these stimuli remain unknown, the signaling pathways need to converge on central guard cell ion channel and metabolic pathways. Model for Roles of Ion Channels in ABA-Induced Stomatal Closing Stomatal closing requires ion efflux from guard cells. Models for roles of ion channels during ABA-induced stomatal closing have been used as a basis for dissecting upstream ABA signal transduction mechanisms (MacRobbie, 1998; Schroeder and Hagiwara, 1989; Schroeder and Hedrich, 1990; Ward et al., 1995). In brief, ABA induces cytosolic Ca2+ increases (McAinsh et al., 1990). Cytosolic calcium elevations, in turn, inhibit plasma membrane proton pumps (Kinoshita et al., 1995) and K+in channels and activate two types or modes (Dietrich and Hedrich, 1994) of plasma membrane anion channels that mediate anion release from guard cells (Keller et al., 1989, Schroeder and Hagiwara, 1989). One of these anion channels shows slow and sustained activation (S-type anion channels) (Linder and Raschke, 1992; Schroeder and Hagiwara, 1989; Schroeder and Keller, 1992), whereas the other anion channel shows rapid transient activation (R-type or GCAC anion channels) (Hedrich et al., 1990). Channel-mediated anion efflux from guard cells causes either transient or sustained anion efflux and depolarization. Depolarization, in turn, deactivates inward-rectifying K+ (K+in) channels and activates outward-rectifying K+ (K+out) channels (Schroeder et al., 1987), resulting in K+ efflux from guard cells. The ensuing long-term efflux of both anions and K+ from guard cells contributes to loss of guard cell turgor and to stomatal closing (Schroeder and Hagiwara, 1989). Recent studies have shown a requirement for rapid ABA-induced Ca2+ influx and S-type anion channel activation for RAB18 expression in Arabidopsis suspension culture cells (Ghelis et al., 2000; Ghelis et al., 2000) indicating that these mechanisms are of general importance for early ABA signaling in other cell types. Most ions released across the plasma membrane of guard cells need first to be released into the cytosol from guard cell vacuoles (8; MacRobbie, 1983; MacRobbie, 1995; Ward et al., 1995; Ward and Schroeder, 1994). Models for the roles of vacuolar K+ and anion channels during stomatal regulation have been proposed previously and are described elsewhere (Allen et al., 1996; Allen et al., 1997; MacRobbie, 1995; Pei et al., 1996; Ward et al., 1995; Ward and Schroeder, 1994). However, these models have not yet been tested as stringently as those for plasma membrane ion channels. Combined molecular genetic (e.g. gene disruption) and cell biological approaches are needed to directly analyze these vacuolar ion channel models. CO2-Induced Stomatal Closing Elevated CO2 concentrations arising from respiration in darkness stimulate stomatal closing (Mansfield et al., 1990). Increases in atmospheric CO2 concentrations are also predicted to reduce stomatal apertures and affect gas exchange (Assmann, 1999; Drake et al., 1997). CO2 signaling mechanisms in guard cells have been reviewed recently (Assmann, 1999; Drake et al., 1997). Here, we briefly summarize some recent findings and signaling models. Elevated CO2 concentrations trigger rises in [Ca2+]cyt (Webb et al., 1996), activate S-type anion and outward-rectifying K+ channel currents (Brearley et al., 1997), and modulate R-type anion channels (K Raschke, personal communication). These data show that the ion channel targets of early CO2 signaling are to a degree shared with ABA signaling. However, upstream CO2 sensing and transduction mechanisms have been reported to differ from ABA signaling, because the abi1-1 and abi2-1 mutants show wild-type CO2-induced stomatal closing (Leymarie et al., 1998). A CO2-induced increase in cell wall malate concentration has been proposed to cause stomatal closing (Hedrich et al., 1994). However, >20 mM external malate was required to produce stomatal closing in two independent studies under the same experimental conditions as the above study (Cousson, 2000; Esser et al., 1997). Furthermore, external malate counteracted CO2-induced stomatal closing (Cousson, 2000), thus calling into question the malate as CO2 sensor hypothesis. An alternative hypothesis suggests that the CO2 sensor is located in guard cell chloroplasts and functions via a CO2-induced decrease in zeaxanthin levels (Zhu et al., 1998). Further understanding of CO2 sensing and signaling in guard cells will help in finding crucial links between the signaling pathways reviewed here and guard cell metabolic pathways. Foremost, future research should have important implications for manipulating gas exchange and carbon fixation in the face of rising atmospheric CO2 levels. Roles of [Ca2+]cyt in Stomatal Closing Many stimuli that result in a change in stomatal aperture have been shown, at least in part, to utilize signal transduction pathways involving changes in guard cell [Ca2+]cyt. ABA-induced stomatal closing is Ca2+-dependent (DeSilva et al., 1985; Gilroy et al., 1990; Schroeder and Hagiwara, 1989; Schwartz, 1985). Note that a Ca2+-independent pathway appears to also exist (Allan et al., 1994). ABA induces repetitive, transient increases or oscillations in guard cell [Ca2+]cyt (5; Gilroy et al., 1991; Grabov and Blatt, 1998; McAinsh et al., 1992; Schroeder and Hagiwara, 1989; Staxen et al., 1999 ). Genetic support for the importance of ABA-induced [Ca2+]cyt elevations in guard cells has been obtained recently, as the ABA-insensitive Arabidopsis mutants abi1-1 and abi2-1 show greatly reduced ABA-induced [Ca2+]cyt elevations, and the abi anion channel regulation and stomatal movement phenotypes are suppressed by experimentally elevating [Ca2+]cyt (Allen et al., 1999). Stomatal closure and guard cell [Ca2+]cyt oscillations can also be induced by increases in external (apoplastic) [Ca2+] (Allen et al., 1999; Allen et al., 1999; McAinsh et al., 1995). These Ca2+-induced [Ca2+]cyt oscillations include a repetitive Ca2+ influx across the plasma membrane coupled to Ca2+ release from intracellular stores for each separate Ca2+ transient (Grabov and Blatt, 1998; McAinsh et al., 1995). Why [Ca2+]cyt regulation is so sensitive to changes in apoplastic Ca2+ is not fully understood, although many species limit Ca2+ accumulation in the apoplast surrounding guard cells to prevent aberrant stomatal regulation (DeSilva et al., 1998; Ruiz and Mansfield, 1994). Increases in CO2 cause stomatal closure and [Ca2+]cyt elevations (Webb et al., 1996). Cold shock (Allen et al., 2000; Wood et al., 2000) and oxidative stress, induced by application of H2O2 or methyl viologen, increase [Ca2+]cyt and result in stomatal closure (McAinsh et al., 1996; Pei et al., 2000). The removal of extracellular Ca2+ using EGTA abolishes CO2- and H2O2-induced [Ca2+]cyt elevations, indicating that plasma membrane Ca2+ influx occurs (Pei et al., 2000; Webb et al., 1996). Roles of [Ca2+]cyt in Stomatal Opening Interestingly, stimuli that result in stomatal opening also induce [Ca2+]cyt elevations. Auxin promotes stomatal opening and direct [Ca2+]cyt measurements (Irving et al., 1992), and pharmacological studies (Cousson and Vavsseur, 1998) suggest a role for [Ca2+]cyt elevations. Furthermore, Ca2+-dependent protein kinases (CDPK) activate guard cell vacuole Cl- channels and malate uptake currents that have been implicated in vacuolar anion uptake during stomatal opening (Pei et al., 1996). Blue light promotes stomatal opening, and pharmacological experiments suggest the involvement of [Ca2+]cyt/calmodulin as a second messenger in this process (Curvetto et al., 1994, Shimazaki et al., 1992). However, increasing external Ca2+ can inhibit light-induced stomatal opening (Allen et al., 1999; Parvathi and Raghavendra, 1997; Shimazaki et al., 1999). Cyclic nucleotides may also act in a Ca2+ dependent stomatal opening pathway as stomatal opening can be stimulated by cAMP (Curvetto et al., 1994) or the membrane-permeable cGMP analog 8-Br-cGMP (Cousson and Vavsseur, 1998). Cyclic GMP-induced stomatal opening is inhibited by chelation of external Ca2+ or by inhibitors of intracellular Ca2+ release (Cousson and Vavsseur, 1998). How a single second messenger such as [Ca2+]cyt can control many diverse and opposing responses in a single cell type remains unknown but likely depends on the Ca2+ channels and Ca2+ regulatory systems activated by each stimulus, the downstream response elements expressed at a given time, and the characteristics and dynamics of the elicited [Ca2+]cyt change (the Ca2+ signature) (MacAinsh and Hetherington, 1998). The recent demonstration that GFP-based “cameleon” calcium indicators (Miyawaki et al., 1999; Miyawaki et al., 1997) function in Arabidopsis will allow addressing of these questions by combined genetic and [Ca2+]cyt signaling studies (Allen et al., 1999). Second Messenger Systems regulating [Ca2+]cyt in Guard Cells Cyclic ADP-Ribose and Vacuolar Ca2+ Release Recent experiments have implicated several second messenger systems in ABA and [Ca2+]cyt signaling in guard cells. In animal cells, cyclic ADP ribose (cADPR) is produced from NAD via the action of the enzyme ADP-Ribosyl cyclase, and it mobilizes Ca2+ from intracellular stores by activation of an endomembrane ion channel known as the ryanodine receptor (RYR) (Lee, 1997). In plant vacuoles, nanomolar cADPR concentrations can activate a Ca2+ permeable current (Allen et al., 1995). Microinjection of cADPR into tomato hypocotyl cells shows that cADPR can function in ABA signaling (Wu et al., 1997). In Commelina guard cells cADPR causes [Ca2+]cyt increases and elicits stomatal closing (Leckie et al., 1998). However, microinjection of the inactive cADPR analog 8-NH2-cADPR or noncyclic ADPR does not elicit [Ca2+]cyt increases or guard cell turgor loss. Ryanodine treatment of guard cells also reduces [Ca2+]cyt increases (Grabov and Blatt, 1999). Note that ABA-induced stomatal closure is only partly inhibited by either microinjection of 8-NH2-cADPR (Leckie et al., 1998) or by nicotinamide, an inhibitor of cADPR production (Jacob et al., 1999; Leckie et al., 1998), suggesting that additional parallel [Ca2+]cyt elevation mechanisms are needed in the ABA signaling cascade. Phospholipase C and Inositol 1,4,5 Trisphosphate Various lines of evidence suggest that phospholipase C (PLC) is a component of ABA signal transduction in guard cells. Early experiments showed that release of caged InsP3 into the cytosol of guard cells could cause [Ca2+]cyt increases and stomatal closure (Gilroy et al., 1990) and inhibit K+in channels (Blatt et al., 1990). Treating guard cell protoplasts with ABA slightly elevates InsP3 levels (Lee et al., 1996; Parmar and Brearley, 1995). Additionally, the PLC inhibitor U-73122 (but not the inactive analog U-73343) inhibits the activity of recombinant PLC from tobacco (Staxen et al., 1999). ABA-induced stomatal closure was also inhibited by U-73122, but only by 20%. However, complete inhibition of ABA-induced stomatal closure can be achieved by treating stomata with a combination of U-73122 and nicotinamide (Jacob et al., 1999; MacRobbie, 2000), suggesting that both cADPR and PLC signaling systems function in ABA signaling. Note that other inositol-phosphates can also act as second messengers in ABA signal transduction in guard cells. In a recent study, myo-inositol hexakisphosphate (InsP6) was identified as an intermediary of guard cell signal transduction. ABA stimulates production of InsP6 in guard cells, and InsP6 perfused into the cytosol via a patch pipette inhibited K+in channels in potato guard cell protoplasts in a Ca2+ dependent manner. These data suggest that InsP6 production is also an important component of ABA signaling (Lemtiri-Chlieh et al., 2000). Phospholipase D and Phosphatidic Acid Phospholipase D (PLD) has been implicated in ABA signaling in aleurone cells (Ritchie and Gilroy, 1998) and in guard cells (Jacob et al., 1999). PLD generates phosphatidic acid (PtdOH), and ABA treatment of Vicia guard cells caused PtdOH levels to transiently increase 2.5-fold (Jacob et al., 1999). PtdOH also promotes stomatal closure and inactivates K+in currents. Guard cell [Ca2+]cyt did not increase following PtdOH treatment, suggesting that PLD acts in a parallel pathway or downstream of Ca2+ mobilizing second messenger systems. An inhibitor of PLD activity, 1-butanol, caused only a partial inhibition of ABA-induced stomatal closure whereas near-complete inhibition of stomatal closure resulted from adding 1-butanol together with nicotinamide (Jacob et al., 1999), suggesting a parallel action of PLD to the cADPR-mediated pathway. Amplifying Calcium Signals by Calcium-Induced Calcium Release Release of caged Ca2+ into the guard cell cytosol can induce [Ca2+]cyt increases or oscillations that are larger than can be accounted for solely by the Ca2+ released by photolysis (Gilroy et al., 1990; McAinsh et al., 1995), suggesting that a [Ca2+]cyt-induced calcium release (CICR) mechanism exists in guard cells. Ca2+-permeable SV channels have been proposed to amplify and propagate [Ca2+]cyt signals in guard cells by CICR from vacuoles (Ward and Schroeder, 1994). Cytosolic Mg2+ sensitizes SV channels to physiological [Ca2+]cyt elevations (Pei et al., 1999) and shifts the voltage-dependence of SV channels (Carpaneto et al., 2001) such that SV channel activity is enhanced. The ability of the SV channel to mediate CICR has been questioned because in mesophyll vacuoles increasing the transvacuolar Ca2+ gradient shifts the voltage-dependence to prevent channel opening under conditions that would otherwise allow Ca2+ to enter the cytosol (Potossin et al., 1997). However, the voltage-dependence and pharmacology of radiolabeled Ca2+ release in vitro from vacuolar vesicles suggests that CICR can be mediated by SV channels (Bewell et al., 1999). The role of SV channels in CICR in guard cells remains an important issue in signal transduction, analysis of which best requires manipulation of SV channel genes. Plasma Membrane Calcium Channels and Calcium Influx in Guard Cells Experimental application of repeated hyperpolarizations negative of -120 mV induced repetitive [Ca2+]cyt transients in guard cells (Grabov and Blatt, 1998). ABA application shifted the threshold of hyperpolarization-activated Ca2+ elevations to -80 mV, indicating that an early event in ABA signaling is the sensitization of Ca2+ influx to membrane potential. Furthermore, in 50 mM KCl buffers [in which the cells are depolarized (Roelfsema and Prins, 1998; Thiel et al., 1992)] ABA only induces [Ca2+]cyt increases in 30–60% of guard cells (Allen et al., 1994 ;Allen et al., 1999; Gilroy et al., 1991; McAinsh et al., 1992; Schroeder and Hagiwara, 1989). Transient ABA activation of Ca2+-permeable channels was found during depolarizations in 37% of guard cells (Schroeder and Hagiwara, 1989), which could contribute to ABA-induced [Ca2+]cyt elevations in depolarized guard cells. Interestingly, in cells maintained in 5 mM KCl, guard cells are hyperpolarized and ABA induces repetitive [Ca2+]cyt transients or oscillations in a higher proportion of cells (80–90%) (Allen et al., 1999; Grabov and Blatt, 1998; Staxen et al., 1999). These data indicate that changes in plasma membrane potential are a central component in ABA signaling. The upstream second messenger mechanisms that activate guard cell plasma membrane Ca2+ channels remained unknown. A recent study in Arabidopsis guard cells shows that reactive oxygen species (ROS) can activate a hyperpolarization-activated Ca2+ influx current (ICa) and that ROS can act as a second messenger in ABA signaling. ABA treatment enhances ROS production in Arabidopsis guard cells (Pei et al., 2000). Interestingly, ABA activates guard cell ICa (Pei et al., 2000) only when NADPH is present in the cytosol, implicating NADPH oxidases in ABA signaling (Murata et al., 2001). Activation of ICa by hydrogen peroxide is impaired in the recessive ABA-insensitive mutant gca2, whereas ABA-induced ROS production remains intact in gca2 (Pei et al., 2000), providing genetic evidence for roles of ROS and ICa in ABA signaling. ABA-induced [Ca2+]cyt oscillations in gca2 stomata are of increased frequencies and shorter transient duration than in wild-type, and imposing [Ca2+]cyt oscillations of wild-type characteristics to gca2 stomata restored closure (Allen et al., 2001). These data lead to a model in which ROS production, GCA2, Ca2+ channel activation, and [Ca2+]cyt oscillations represent a new signaling “cassette” in guard cells. These data further indicate that GCA2 functions in feedback regulation of [Ca2+] oscillation frequency (Fig. 3). A study in maize embryos has also shown that ABA enhances ROS production (Guan et al., 2000). The question whether ROS function as second messengers in ABA signaling in planta and the mechanisms leading to ABA-dependent ROS increases remained unknown. A recent study identified the AtrbohD and AtrbohF NADPH oxidase subunits as being essential for ABA-induced stomatal closing and ABA-induced ICa calcium channel activation in guard cells (Kwak et al., 2003). The atrbohD,F double knock-out mutant impairs ABA-induced ROS production in guard cells. An additional reactive oxygen, the second messenger nitric oxide (NO) was shown to be involved in guard cell signal transduction as well, inducing stomatal closure in V. faba and P. sativum (Mata and Lamattina 2001; Neill et al. 2002). Plants can produce NO not only from arginine by NO synthase but also from nitrate via nitrite by nitrate reductase (NR) (Sakihama et al. 2002). An Arabidopsis double mutant (non-null allele) in the NR genes NIA1 and NIA2 did not produce NO in guard cells upon exposure to ABA, and nia1,2 stomata did not respond to ABA, providing genetic evidence for the requirement of NR activity in ABA-mediated stomatal closure (Desikan et al. 2002). Interestingly, Arabidopsis ABA-insensitive abi1-1 and abi1-2 mutants still produced NO in response to ABA, placing ABI1 and ABI2 downstream of NIA1 and NIA2 in ABA signaling (Desikan et al. 2002) or in a parallel branch. Single channel recordings in Vicia guard cell protoplasts have identified a Ca2+ channel that is hyperpolarization-activated and shows a 250-fold increased open probability following addition of 20 uM ABA (Hamilton et al., 2000). Currents exhibited by this channel are very similar to ICa. The activation by ABA occurs in isolated patches, suggesting that ABA perception and channel activation are closely associated in Vicia. Interestingly, the single channel open probability was reduced tenfold when buffering [Ca2+]cyt from 200 nM to 2 uM, indicating that the channel is downregulated during [Ca2+]cyt elevation and is therefore subject to negative feedback control during [Ca2+]cyt signaling (Hamilton et al., 2000). Both at the single-channel level (Hamilton et al., 2000) and in whole cells (Pei et al., 2000 ), the activity of the Ca2+ influx channel can show spontaneous oscillatory behavior without exogenous ABA addition. Background ICa activity was inhibited by addition of 0.1 mM DTT in Arabidopsis guard cells (Pei et al., 2000). The spontaneous ICa activity may contribute to spontaneous [Ca2+]cyt elevations found in hyperpolarized guard cells (Allen et al., 1999; Grabov and Blatt, 1998; Staxen et al., 1999). Because ICa channels are regulated by ROS, various stress signals may control Ca2+ influx by regulating the oxidative state of guard cells. For example, pathogen elicitors trigger ROS production in guard cells and stomatal closing (Lee et al., 1994), and ozone, which closes stomatal pores (Torsethaugen et al., 1999), might modulate ICa. In this regard, ROS production and ICa have been proposed to function as a shared “signaling cassette” of multiple stress signaling pathways (Pei et al., 2000). Additional Ca2+ influx pathways may be provided by plasma membrane stretch-activated Ca2+ permeable channels (Cosgrove and Hedrich, 1991) or via a Ca2+ permeability of transient and sustained K+out channels (Pei et al., 1998; Romano et al., 1998). Clearly, Ca2+ influx, Ca2+ release, and second messengers are integrated in guard cells to produce a [Ca2+]cyt signal that controls stomatal movements. How these separate processes may be integrated to produce [Ca2+]cyt signals that encode information necessary for stomatal closure is an important focus of ongoing studies (see MacRobbie, 2000). Cytosolic [Ca 2+ ] Oscillations Are Necessary for Stomatal Closing in Guard Cells In a few cases in animal cells, the frequency of [Ca2+]cyt oscillations has been shown to control the efficiency and specificity of cellular responses (DeKoninck and Schulman, 1998; Dolmetsch et al., 1998; Li et al., 1998). In plants, it remains to be investigated whether Ca2+ oscillations are an absolute requirement for eliciting physiological responses. Models for membrane potential oscillations in guard cells have been generated (Buschmann and Gradmann, 1997; Gradmann et al., 1993; Kolb et al., 1995 ) to analyze different mechanisms that may contribute to generation of [Ca2+]cyt oscillations. Using guard cells of the Arabidopsis V-ATPase mutant de-etiolated 3 (det3) (Schumacher et al., 1999), which has reduced endomembrane proton pumping and energization, external Ca2+ and oxidative stress elicited prolonged Ca2+ increases (plateaus) that did not oscillate, whereas wild-type cells show [Ca2+]cyt oscillations (Allen et al., 2000). Unexpectedly, steady state stomatal closure was inhibited in det3 in response to these stimuli. Conversely, cold and ABA elicited Ca2+ oscillations in det3, and stomatal closures were not impaired (Allen et al., 2000). Moreover, in det3 guard cells, experimentally imposing external Ca2+-induced oscillations rescued steady state stomatal closure in response to external Ca2+, and imposing Ca2+ plateaus in wild-type guard cells prevented steady-state stomatal closing. These data provide genetic evidence that stimulus-specific Ca2+ oscillations, rather than a mere plateau of [Ca2+]cyt, are necessary for long-term stomatal closure (Allen et al., 2000). These findings suggest that guard cells may provide an excellent genetic system to study [Ca2+]cyt pattern-dependent responses. By correlating stomatal aperture to imposed [Ca2+] oscillations, Allen et al. were able to decipher the signal with respect to [Ca2+] peak number, peak duration, and frequency (Allen et al., 2001a; reviewed in Allen et al., 2001b). The fact that cytosolic [Ca2+] oscillations are necessary for stomatal closure (Allen and Schroeder, 2001) indicates an important role for Ca2+-binding proteins in guard cell ABA signaling. Candidate Ca2+-binders in Arabidopsis are the SOS3-like calcium binding proteins SCaBPs, related to yeast calcineurin B (Liu and Zhu, 1998). Indeed, RNAi-mediated genetic "knock-down" of SCaBP5 rendered guard cells hypersensitive to ABA (Guo et al. 2002). Arabidopsis scabp5 silencing lines were also ABA-hypersensitive in seed germination, seedling growth, and gene expression (Guo et al. 2002). Since ScaBP genes are closely related to one another, individual knock-out will be instrumental in characterizing functions of individual genes. A kinase was identified (PKS3) that interacts with SCaBP5 in the yeast two-hybrid system and that, when knocked-down by RNAi, exhibited the same phenotypes as scabp RNAi lines (Guo et al. 2002). Interestingly, PKS3 interacted also with the phosphatases ABI1 and ABI2 when expressed in yeast (Guo et al. 2002). ABA-insensitive PP2C Mutants Genetic screens for Arabidopsis mutants insensitive to ABA inhibition of seed germination yielded two dominant mutants that are impaired in ABA-induced stomatal closure (Koornneef et al., 1984). The corresponding genes, ABI1 and ABI2, both encode type 2C protein phosphatases and the dominant mutant alleles abi1-1 and abi2-1 have point mutations altering a conserved amino acid (Leung et al., 1994; Leung et al., 1997; Meyer et al., 1994; Rodriguez et al., 1998). Several downstream responses to ABA are impaired in these Arabidopsis mutants including K+out and K+in channel regulation (Armstrong et al., 1995) and anion channel activation (Pei et al., 1997). These mutations also impair ABA-induced [Ca2+]cyt increases (Allen et al., 1999). Furthermore, experimental elevation in [Ca2+]cyt causes anion channel activation and stomatal closure in abi1-1 and abi2-1, thus bypassing the effects of the abi1-1 and abi2-1 mutations (Allen et al., 1999). These data demonstrate that the dominant abi PP2C mutants interfere with very early ABA signaling events that act upstream of [Ca2+]cyt (Allen et al., 1999). Because the only known alleles of these genes were dominant mutations, it has been unclear whether the ABI1 and ABI2 phosphatases are positive or negative regulators of ABA signaling or indeed whether they affect ABA signaling at all in wildtype. Recently, however, intragenic revertants of the abi1-1 and abi2-1 mutants were isolated and shown to have reduced or no phosphatase activity in vitro (Gosti et al., 1999; Merlot et al., 2001). Because a double mutant of both revertants shows hypersensitivity to ABA, ABI1 and ABI2 are likely negative regulators of ABA signaling (Merlot et al., 2001). In correlation, overexpression of wild-type ABI1 in maize mesophyll protoplasts blocks ABA regulation of gene expression (Sheen, 1998). In spite of these advances, ABI1 and ABI2 gene deletion or silencing mutants would be useful for a stringent test of their functions, because all intragenic revertant mutations lie downstream of the dominant mutant site (Gosti et al., 1999; Merlot et al., 2001), which might form an attachment to an essential signaling protein. Anion Channels and ABA-induced Stomatal Closure Anion channel activation at the plasma membrane of guard cells has been proposed as an essential step during stomatal closure (Hedrich et al., 1990; Schroeder and Hagiwara, 1989; Schroeder and Keller, 1992). S-type anion channel currents in guard cells are activated by increases in [Ca2+]cyt in Vicia (Schroeder and Hagiwara, 1989) and Arabidopsis (Allen et al., 1999). R-type anion currents are activated following an increase in external Ca2+ (Hedrich et al., 1990). Whether ABA regulates R-type anion channels remains to be determined. ABA activation of S-type anion channels in the plasma membrane of guard cells has now been demonstrated in Arabidopsis (Pei et al., 1997), tobacco (Grabov et al., 1997), and Vicia (Leonhardt et al., 1999; Li et al., 2000; Schwarz and Schroeder, 1998). This response is disrupted in the Arabidopsis abi1-1 and abi2-1 mutants, providing genetic evidence that activation of S-type channels contributes to stomatal closure (Pei et al., 1997). Putative anion channel genes have been cloned from tobacco (Lurin et al., 1996) and Arabidopsis (Hechenberger et al., 1996) based on sequence homology with the animal voltage-dependent ClC chloride channels [for review see (Barbier-Brygoo et al., 2000)]. Further work is needed to determine whether these genes encode components of native plant plasma membrane anion channels. Regulation of Guard Cell ABA Signaling by ABC Proteins Pharmacological studies have led to the model that guard cell S-type anion channels may be encoded by or regulated by ATP binding cassette (ABC) proteins (Leonhardt et al., 1997; Leonhardt et al., 1999). ABC proteins comprise a large family of membrane proteins that actively translocate a wide spectrum of substrates. In addition, ABC proteins regulate the activity of other unrelated transporters. ABC proteins such as CFTR (cystic fibrosis transmembrane conductance regulator) or SUR (sulfonylurea receptor) show Cl- channel activity and/or regulate other channels [for review see (Theodoulou, 2000)]. Inhibitors of SUR, such as glibenclamide, prevent ABA-induced stomatal closure in Vicia and Commelina. Furthermore, the ABC protein inhibitors DPC and glibenclamide block slow anion currents in Vicia guard cells (Leonhardt et al., 1999). In contrast, cromakalin, an antagonist of glibenclamide, triggers stomatal closing in Commelina and reverses the inhibition of glibenclamide on S-type anion currents in Vicia guard cells (Leonhardt et al., 1999). Protein Kinases Function in ABA Signaling and Anion Channel Regulation Pharmacological approaches, using the serine/threonine protein kinase inhibitor K252a and cytosolic replacement of ATP, showed that phosphorylation events are central positive regulators in ABA-induced stomatal closure in Vicia (Schmidt et al., 1994), as well as in Commelina (Esser et al., 1997), Pisum (Hey et al., 1997), and Arabidopsis (Allen et al., 1999). K252a abolishes both anion channel activity and ABA-induced stomatal closing (Schmidt et al., 1994). In correlation with these results, ABA induction of gene expression (RD29a and KIN2) in tomato hypocotyls is inhibited by K252a Wu et al., 1997), together with other studies, indicating that kinase-dependent transduction of ABA signaling ( Schmidt et al., 1994) is of general significance (Knetsch et al., 1996; Sheen, 1996; Wu et al., 1997). Note that the R-type anion currents are not regulated by phosphorylation events but that nucleotide binding activates these anion channels (Hedrich et al., 1990; Schulz-Lessdorf et al., 1996). Biochemical approaches led to the characterization of a Ca2+-independent, ABA inducible 48-kDa kinase (non-MAP kinase) in Vicia guard cells (Li and Assmann, 1996; Mori and Muto, 1997). The kinase activity was named AAPK or ABR. Recently, an AAPK gene was cloned (Li et al., 2000). Transient expression of a dominant negative allele of AAPK, which abolished kinase activity, prevents ABA activation of S-type anion currents and stomatal closing. In correlation with these findings, in Vicia S-type anion currents are activated at low [Ca2+]cyt and high ATP concentrations, suggesting that a final phosphorylation event in anion channel activation can be Ca2+ independent (Schwarz and Schroeder, 1998). Recessive loss-of-function mutations in AAPK will allow further analysis of AAPK function in guard cells. Two Ca2+-dependent protein kinases of 53 kDa and 58 kDa have been characterized in Vicia guard cells (Li et al., 1998; Mori and Muto, 1997). Removal of Ca2+ with BAPTA in Vicia guard cell protoplast suspensions prevents ABR kinase activation and indicates that Ca2+ is required upstream for ABR activation (Mori and Muto, 1997). These data and overexpression studies on maize protoplasts suggest that CDPKs may be positive regulators in ABA signal transduction (Sheen, 1996). In addition, MAP kinases have been reported to positively control ABA-induced stomatal closure in Pisum (Burnett et al., 2000). ABA causes a transient activation of a 43-kDa MAP kinase named AMBPK. AMBPK exhibits all fundamental MAP kinase properties, including tyrosine phosphorylation (Burnett et al., 2000). The MAPK kinase inhibitor PD98059 abolished ABA-induced stomatal closing and ABA induction of dehydrin mRNA (Burnett et al., 2000). ABA activation of MAP kinases in barley aleurone was previously reported (Heimovaara-Dijkstra et al., 2000; Knetsch et al., 1996). In parallel to the above kinases that transduce ABA signals, other protein kinases have been suggested to act as negative regulators of ABA signaling in tobacco and Arabidopsis guard cells in the abi1-1 background (Armstrong et al., 1995; Pei et al., 1997). Application of kinase inhibitors partially restores ABA activation of anion channels and regulation of K+ channels and stomatal closing in abi1-1 backgrounds (Armstrong et al., 1995; Pei et al., 1997). Okadaic Acid–sensitive Phosphatases Regulate Anion Channels and ABA Signaling Inhibitors of PP1 and PP2A protein phosphatases such as okadaic acid (OA) were found to enhance S-type anion currents and ABA-induced stomatal closure in Vicia (Schmidt et al., 1994; Schwarz and Schroeder, 1998), as well as in tobacco (Grabov et al., 1997), Commelina (Esser et al., 1997), and Pisum (Hey et al., 1997), and enhance ABA-induced gene expression in tomato hypocotyls (Wu et al., 1997), suggesting that PP1s or PP2As act as negative regulators in ABA signaling (PP1/PP2Aneg). In Vicia, either OA or ABA maintain anion current activation without cytosolic ATP, indicating that ABA may indeed downregulate a PP1/PP2Aneg (Schwarz and Schroeder, 1998). In addition to negatively regulating PP1/PP2As, evidence suggests that other PP1/PP2As can also act as positive regulators in ABA signaling (PP1/PP2Apos). In Arabidopsis, OA partially inhibited ABA activation of S-type anion channels and stomatal closing (Pei et al., 1997). A similar inhibitory effect of OA was also observed on ABA signaling during stomatal opening and ABA induction of dehydrin mRNA in Pisum epidermal peels (Hey et al., 1997). OA also inhibits ABA-induced expression of the PHAV1 gene in barley aleurone (Kuo et al., 1996). Experiments in Pisum show that the activity of either PP1/PP2Apos or PP1/PP2Aneg can be resolved depending on the aperture of stomates (Hey et al., 1997). The PP2A protein phospharase inhibitor okadaic acid inhibits ABA signaling in guard cells (Pei et al., 1997). To analyze the hypothesis that PP2As function in ABA signaling, (Kwak et al., 2002) identified from guard cell cDNA libraries a gene encoding a PP2A regulatory A subunit, RCN1. RCN1 was characterized previously as a molecular component affecting auxin transport and gravitropism (Garbers et al., 1996; Rashotte et al., 2001) and a loss-of function rcn1 mutant has been isolated (Deruère et al., 1999). When Kwak et al. (2002) analyzed guard cell responses, they found that the rcn1 mutation impairs ABA-induced stomatal closing and ABA activation of slow anion channels. However, rcn1 did not affect stomatal closure induced by external application of Ca2+ or H2O2, indicating that RCN1 may not function downstream of cytosolic [Ca2+] elevations. Consistent with these findings, ABA-induced cytosolic [Ca2+] elevations are reduced in rcn1 guard cells, suggesting that RCN1 functions upstream of cytosolic [Ca2+] elevations. rcn1 phenotypes in ABA inhibition of seed germination, in ABA-induced stomatal closing and in ABA-induced [Ca2+]cyt elevations can be phenocopied in wild-type by external application of the PP1 and PP2A inhibitor okadaic acid. Together, these observations indicate that RCN1 may act upstream of induced cytosolic [Ca2+] elevations. These different studies bring to light that a complex phosphorylation and dephosphorylation network exists in guard cells and that significant differences can occur in ABA signaling depending on the physiological state of guard cells. Activity of Plasma Membrane K+out Channels in Stomatal Movements Efflux of K+ from the cell during stomatal closing has been proposed to occur through outward rectifying K+ (K+out) channels that are activated by membrane depolarization (Schroeder et al., 1987). Guard cells respond to ABA by enhancing K+out and reducing K+in channel currents (Blatt, 1990; Blatt and Armstrong, 1993). Unlike K+in channels, however, K+out channels are largely insensitive to increases in [Ca2+]cyt that occur during ABA signaling (Lemtiri-Chlieh and MacRobbie, 1994; Schroeder and Hagiwara, 1989). A guard cell-expressed SKOR K+ channel cDNA homologue, named GORK , was isolated that when expressed in Xenopus oocytes produces outward rectifying K+ channels with properties similar to K+out channels (Ache et al., 2000). A knock-out in the GORK gene leads to lack of K+out channel currents in guard cells (Hosy et al., 2003). ABA induces an increase in the cytosolic pH of guard cells (Blatt and Armstrong, 1993; Grabov and Blatt; 1997; Irving et al., 1992). Experiments in Vicia guard cells show that K+out currents are enhanced by increased cytoplasmic pH (Blatt, 1992). The pH stimulation of K+out channels occurs in isolated membrane patches and is thus membrane delimited (Miedema and Assmann, 1996). ABA-induced increases in K+out currents can be inhibited by acidification or buffering of the guard cell cytoplasmic pH, showing that cytosolic pH has a functional role in ABA signal transduction (Blatt and Armstrong, 1993). Regulation of K+out by Phosphorylation Several lines of evidence suggest that protein phosphorylation plays a role in modulation of K+out channel activity. Guard cells from tobacco plants transformed with the dominant phosphatase mutant allele abi1-1 from Arabidopsis show K+out currents that are two- to fourfold lower than wild type, and both K+in and K+out currents are insensitive to modulation by ABA, suggesting a role for PP2Cs in guard cell signaling (Armstrong et al., 1995). Despite the ABA insensitivity of K+out channels in the abi1-1-transformed guard cells, concurrent measurements of intracellular pH show normal pH increases in response to ABA (Grabov and Blatt, 1998). This suggests that the abi1-1 phosphatase acts downstream of, or parallel to, the cytosolic pH changes. If cytosolic pH directly modulates K+out channels (Miedema and Assmann, 1996), then abi1-1 must act in a parallel pathway to alter the responsiveness of the K+out channel to pH. In addition to the PP2C phosphatases identified genetically, experiments using the phosphatase inhibitor OA implicate PP1- or PP2A-type protein phosphatases in the regulation guard cell K+out channels. OA downregulates both K+in and K+out channel currents in Vicia (Thiel and Blatt, 1994). However, another study showed that OA downregulated only K+in, and not K+out, currents (Li et al., 1994). Despite the experimental differences, one study indicates a putative role for PP1- and/or PP2A-phosphatases as positive regulators of K+out channel activity in guard cells (possibly related to PP1/PP2Apos). Syntaxins and ABA Signaling Syntaxins play important roles in membrane fusion. A tobacco cDNA encoding a homolog of human and yeast syntaxin (NtSyr1) was isolated using heterologous expression of drought-stressed tobacco leaf mRNA in Xenopus oocytes (Leyman et al., 1999). Expression of mRNA pools show ABA activation of endogeneous Ca2+-activated Cl- currents in oocytes (Leyman et al., 1999) and ABA downregulation of KAT1-mediated K+in channels in oocytes (Sutton et al., 2000), suggesting that oocytes will provide an approach for isolating and analyzing putative ABA receptor cDNAs. By subfractionation the NtSyr1 mRNA was isolated, which when expressed in oocytes constitutively activated the Cl- currents without addition of ABA (Leyman et al., 1999). Whether NtSyr1 contributes to the ABA response in oocytes remains to be determined (Leyman et al., 1999). Voltage clamp recordings in tobacco guard cells provide pharmacological evidence for a role of syntaxins in ABA regulation of K+ channels and S-type anion channels. These data suggest that syntaxin acts as a positive regulator of ABA signaling in guard cells (Leyman et al., 1999). Guard cell volume is changed during stomatal movements and is accompanied by changes in membrane surface area (Homann, 1998; Kubitscheck et al., 2000). Syntaxins may link ABA signaling to membrane trafficking. Osmolarity and Temperature Sensitivity Guard cell measurements of 86R+ efflux kinetics show that whereas a rapid transient K+ release response to ABA was dependent on the concentration of ABA and duration of exposure, the end-state or final internal 86R+ concentration reached was not (MacRobbie, 1995). This led to the suggestion that guard cells have some means of sensing their internal osmolarity that is linked to regulation of K+ efflux channels. Vicia guard cell plasma membrane patches exposed to osmotic gradients show K+out channels that are inactivated by hypotonic (guard cell swelling) conditions and activated by hypertonic (guard cell shrinking) conditions (Liu and Luan, 1998). Recent work has suggested that K+out channels may be involved in the response of guard cells to another environmental stimulus: temperature. At moderate temperatures (13° to 20°C), both K+in and K+out channel conductances in Vicia increase with increasing temperature. At temperatures from 20° to 28°C, K+out conductance decreases with increasing temperature, whereas K+in conductance continues to increase (Ilan et al., 1992). This difference in temperature response between K+in and K+out channels at higher temperatures would favor K+ influx and stomatal opening and thus could allow increased transpirational cooling of leaves (Ilan et al., 1992). Transient K + Efflux Currents In addition to the slow activating K+out channels discussed above, a rapidly activated transient outward K+ current, (IAP), has been observed in Arabidopsis and tobacco guard cells (Armstrong et al., 1995; Pei et al., 1998; Roelfsema and Prins, 1997). In addition to being the first inactivating K+out current characterized in plants, IAP has several unusual characteristics. In contrast to the slow K+out channels that are activated by alkaline cytosolic pH (Blatt, 1992), IAP is slightly inhibited by alkaline pHcyt (Pei et al., 1998). IAP is also inhibited by increased [Ca2+]cyt (Pei et al., 1998), whereas guard cell K+out channels are not regulated by small cytoplasmic Ca2+ changes. IAP channels show a significant Ca2+ permeability (Pei et al., 1998). The physiological role of this transient current is unknown, but IAP may contribute to shorter-term adjustments in stomatal aperture or to membrane potential oscillations observed in guard cells (Gradmann et al., 1993; Thiel et al., 1992). New Guard Cell Signaling Mutants and Genetic Approaches Quantitative and mechanistic characterization (Allen et al., 1999; Pei et al., 1997) of new Arabidopsis guard cell signaling mutants is paramount to achieve a molecular understanding of the ABA signaling cascade. Furthermore genes for many of the above proposed cell biologically and pharmacologically derived mechanisms have not yet been identified. Loss-of-function mutations in the Arabidopsis ERA1 farnesyltransferase beta subunit cause an enhanced response to ABA in seeds (Cutler et al., 1996). Moreover, the era1 mutant shows ABA hypersensitive stomatal closing and ABA hypersensitive activation of S-type anion currents (Pei et al., 1998). Furthermore, era1 plants show reduced water loss during drought (Pei et al., 1998). Application of farnesyltransferase inhibitors to wild-type stomata mimics the era1 phenotype, suggesting that ERA1 functions in early guard cell signaling and illustrating the complementarity of “steady-state” gene knockouts and “short-term” inhibitor applications. In mammals and yeast, farnesylation of signaling proteins promotes their membrane location and protein-protein interactions, suggesting that a negative regulator of guard cell ABA signaling is targeted via farnesylation (Pei et al., 1998). ABA hypersensitive [Ca2+]cyt elevations in era1 show that ERA1 functions upstream of [Ca2+]cyt elevations (Allen et al., 2002). A new ABA hypersensitive loss-of-function mutant, abh1, was isolated and characterized by screening for ABA hypersensitivity in seed germination in a primary screen and identifying the subset of mutations that affect guard cell signaling in a secondary screen (Hugouvieux et al., 2001). Stomatal closing is hypersensitive to ABA in abh1 and, consistent with this phenotype, ABA-induced [Ca2+]cyt elevations and guard cell plasma membrane anion currents are enhanced and K+in currents are reduced in abh1 (Hugouvieux et al., 2002). Suprisingly, ABH1 encodes a subunit of a nuclear RNA cap binding complex (Hugouvieux et al., 2001) previously described in mammals and yeast, which regulates RNA processing and growth factor signal transduction (Izaurralde et al., 1995; Izaurralde et al., 1994; Wilson et al., 1999). ABH1 may control the strength of ABA signaling by modulating the expression of an early ABA signal transduction element(s). Recently, there have been major advances in our understanding of the molecular events in ABA signal transduction due to the development of new strategies. One highly elegant and successful approach is a stomatal movement screen using infrared thermography in which small differences in leaf temperature were used to isolate new Arabidopsis guard cell mutants. Using this technique, Mustilli et al. (2002) isolated two allelic Arabidopsis recessive mutants, ost1-1 and ost1-2, in which guard cells are insensitive to ABA. OST1 encodes an ABA-activated Ca2+-independent protein kinase (AAPK) that shares 79% amino acid identity with V. faba AAPK previously isolated and characterized by biochemical approaches (Li et al., 2000). Analogous to AAPK, also called ABR kinase (Mori and Muto, 1997), OST1 is activated by ABA (Li et al., 2000; Mustilli et al., 2002). In another study the same gene was identified (named SnRK2) in reverse genetic analysis as a component that functions in dehydration stress in Arabidopsis (Yoshida et al., 2002). In addition, ABA-induced ROS production was disrupted in ost1 guard cells, whereas applied H2O2 induced similar stomatal closing in wild type and in ost1 suggesting that the ost1 mutation disrupts ABA signaling upstream of ROS production. Moreover, ABA activation of the OST1 kinase was impaired in abi1-1, suggesting that OST1 acts between abi1-1 and ROS production. Using the two-hybrid system which allows a rapid screen for putative partners of a given protein, Himmelbach et al. (2002) isolated the homeobox–leucine zipper transcription factor, ATHB6 as a putative ABI1-interacting protein. Interestingly, the interaction between ATHB6 and ABI1 positively correlated with the PP2C activity of the ABI1 catalytic domain and was completely abolished in a point-mutated, catalytically inactive ABI1. In addition, stable expression analysis documented a > 200-fold ABA mediated induction of reporter expression under the control of the ATHB6 promoter that was dependent on abi1-1. Transgenic lines overexpressing ATHB6 under the control of the 35S promoter exhibited increased water loss in leaves, comparable to the wilty mutant abi1-1. All together these results demonstrate that ATHB6 can physically interact with ABI1 and that the deregulation of ATHB6 decreases the sensitivity towards ABA in responses that are controled by ABI1. New Genetic Screens and Reverse Genetics To date, most of the ABA-insensitive mutations identified in guard cell signaling (abi1-1, abi2-1, and aapk) are dominant (Finkelstein and Somerville, 1990; Leung et al., 1997; Li et al., 2000). This suggests that redundancy in phosphatase and kinase activities may limit the isolation of recessive mutations in such genes. Dominant mutations can also be generated in Arabidopsis by activation tagging or the random overexpression of wild-type genes (Weigel et al., 2000). Such mutants can aid in identifying and isolating redundant genes involved in guard cell signaling pathways. However, dominant mutations can result from interactions with unnatural partners, causing neomorphic responses. To identify unequivocally the functions of such genes, it will be important to isolate loss-of-function mutants (Himmelbach et al., 1998). Disruption or silencing of homologous redundant genes expressed in guard cells may allow more direct characterizations of in vivo functions of redundant genes. Isolating guard cell signaling mutants is not trivial, owing to the lack of easily scorable phenotypes or markers, and is more difficult than isolating stomatal development mutants, which include cell-to-cell signaling mechanisms (Berger and Altmann, 2000; Geisler et al., 2000). Thus the era1 and abh1 guard cell phenotypes were identified in secondary stomatal movement screens (Hugouvieux et al., 2001; Pei et al., 1998). A highly elegant stomatal movement screen has been developed in which small differences in leaf temperature [due to stomatal transpiration (Raskin and Ladyman, 1988)] were used to isolate new Arabidopsis guard cell signaling mutants (J Giraudat & S Merlot, personal communication; Merlot et al., 2002). In a different approach, mutations that affect circadian control of stomatal movements were identified by selecting Arabidopsis mutants with an enhanced sensitivity to sulfur dioxide at specific times of day (R McClung, personal communication). This screen has led to isolation of circadian timing-defective (ctd) and out-of-phase (oop) mutants. In another screen using luciferase as a reporter, many mutants were isolated that are affected in osmotic, ABA, and cold stress–induced signal transduction (Ishitani et al., 1997). A subset of these mutations will likely affect guard cell signaling. Use of novel creative genetic screens will lead to identification of many new mechanisms affecting guard cell signaling. For example, new guard cell signaling mutants in stomatal responses could be isolated based on variation in responses among Arabidopsis ecotypes and use of recombinant inbred lines to map quantitative trait loci (Alonso-Blanco et al., 1998) that affect guard cell signaling. Note that many Arabidopsis light and hormone signaling mutants have been isolated based on whole-plant or whole-tissue phenotypes, which can lead to isolation of genes that indirectly affect a signaling pathway via crosstalk or indirect feedback among signaling cascades. Analyses of ion channel regulation and [Ca2+]cyt signaling in mutants allows one to closely associate mutations with specific elements in early signaling (Allen et al., 1999; Pei et al., 1998; Pei et al., 1997). The completion of the Arabidopsis genome sequence will lead to reverse genetic functional characterizations of new guard cell signaling components using biophysical cell biological (Allen et al., 1999; Allen et al., 1999; Pei et al., 1997) and genomic methods developed and adapted to Arabidopsis guard cells. The identification of the full complement of guard cell– expressed genes is now possible using DNA array and chip technologies, which will have profound influence on future research. Reverse genetic analyses of guard cell–expressed genes will be needed to identify genes and test the function and relative contribution of the many proposed signal transducers reviewed here. Furthermore, many of the signal transducers reviewed here likely form signal transduction complexes consisting of many proteins. Proteomic approaches (Li et al., 2000) will play an increasingly important role for identifying members of guard cell signaling complexes. Future Outlook Guard cell research has revealed many new signal transduction components and led to models of early signal transduction elements and signaling cassettes in plants. Many of the proposed mechanisms summarized in this review can now be directly tested by reverse genetics. Furthermore, this research has led to an initial understanding of how a large number of signaling mechanisms can interact in concert to produce a rapid physiological response. Future research on mutants will define molecular junction points of an integrated network. As discussed in the introduction, most plant cells respond to the classically known hormones and light signals in specific ways. In this sense, each plant cell contains a ucosm of plant signaling cascades with intricate crosstalk and specificity mechanisms. The guard cell system lends itself to functional characterization of many new unknown early signaling mechanisms (see 1 to 4 in Introduction). Interdisciplinary studies using cell biological, genomic, molecular genetic, biophysical, reverse genetic, and proteomic approaches will define much of the future research in this field. Initial examples have shown that manipulation of guard cell signaling genes in Arabidopsis can affect stomatal movements, leading to reduced water loss and slowing of desiccation during transitory drought periods (Gosti et al., 1999; Hugouvieux et al., 2001; Pei et al., 1998). Future research in this area combined with inducible guard cell-specific gene expression or cell-specific gene silencing will lead to identification of mechanisms for engineering improved gas exchange in response to drought, elevated atmospheric CO2, and other environmental stresses (Schroeder et al., 2000). In this respect, guard cell signaling research holds much promise at addressing major environmental and agricultural problems of the twenty-first century. Acknowledgments We thank many colleagues for communicating new and unpublished findings. Majid Ghassemian, Nathalie Leonhardt, Pascal Mäser, Jared Young, and other members of the Schroeder lab are gratefully acknowledged for comments on the manuscript. Research in the authors’ laboratory was supported by NSF, NIH, and DOE grants. Literature Cited |
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