The Clickable Guard Cell, Version II: Interactive Model of
Guard Cell Signal Transduction Mechanisms and Pathways

June M. KwakA,D, Pascal MäserB, Julian I. SchroederC,D

ADepartment of Cell Biology and Molecular Genetics, University of Maryland, College Park, MD 20742
BInstitute of Cell Biology, University of Bern, CH-3012 Bern, Switzerland
CDivision of Biological Sciences, Cell and Developmental Biology Section and Center for Molecular Genetics, University of California, San Diego, La Jolla CA 92093-0116, USA
DCorresponding authors, email:, phone: 858-534-7759, fax: 858-534-7108; email:, phone: 301-405-9726, fax: 301-314-1248

The Arabidopsis Book, eds. C. Chang, I. Graham, R. Last, M. McClung & C. Weinig, American Society of Plant Biologists

Guard cells are located in the leaf epidermis and pairs of guard cells surround and form stomatal pores, which regulate CO2 influx from the atmosphere into the leaves for photosynthetic carbon fixation. Stomatal guard cells also regulate water loss of plants via transpiration to the atmosphere. Signal transduction mechanisms in guard cells integrate a multitude of different stimuli to modulate stomatal aperture. Stomata open in response to light. Stomata close in response to drought stress, elevated CO2, ozone and low humidity. In response to drought, plants synthesize the hormone abscisic acid (ABA) that triggers the closing of stomatal pores. Guard cells have become a highly developed model system for dissecting signal transduction mechanisms in plants and for elucidating how individual signaling mechanisms can interact within a network in a single cell. Many new findings have been made in the last few years. This chapter is an update of an electronic interactive chapter in the previous edition of The Arabidopsis Book (Mäser et al. 2003). Here we focus on mechanisms for which genes and mutations have been characterized, including signaling components for which there is substantial signaling, biochemical and genetic evidence. Ion channels have been shown to represent targets of early signal transduction mechanisms and provide functional signaling and quantitative analysis points to determine where and how mutations affect branches within the guard cell signaling network. Although a substantial number of genes and proteins that function in guard cell signaling have been identified in recent years, there are many more left to be identified and the protein-protein interactions within this network will be an important subject of future research. This interactive clickable resource includes the following features:

Figure 1.  Model for the roles of ion channels in ABA signaling.
Figure 2.  Blue light signaling pathways in guard cells.
Figure 3.  ABA signaling pathways in guard cells.

Figure 1 is linked to explanations that appear upon mouse-over. Figures 2 and 3 are clickable and linked to info boxes, which in turn are linked to TAIR, to relevant abstracts in PubMed, and to background explanations from Schroeder et al. (2001), used with permission of Annual Reviews of Plant Biology. A list of the Arabidopsis genes shown in the figures and their accession numbers is available on-line or for download.

          2a. Vacuolar Channels
          3a. K+in Channels Mediate K+ Uptake During Stomatal Opening
          5a. ABA-Regulated Calcium Signaling
          5b. Essential Functions of Anion Channels in Stomatal Closing
          5c. Modulators of ABA Signaling Strength
          5d. Reactive Oxygen Signaling Mechanisms in Guard Cells
          5e. Protein Kinases and Phosphatases in ABA Signaling
          5f. mRNA Processing and ABA Signaling


Being rooted in one place, plants are required to respond to changes in environmental conditions and stresses. Hormone and light signal transduction pathways form complex networks that control plant responses to the environment. Although key mechanisms and genes have been identified in these signal transduction pathways, the protein-protein interaction networks and the functions of individual interactions largely remain to be explored. In guard cells, a network of signal transduction mechanisms integrates water status, hormone responses, light, CO2 and other environmental conditions to regulate stomatal apertures in leaves, for optimal plant growth and survival under diverse conditions. Stomatal guard cells have developed into a favorite model system for understanding how various signaling components can interact within a network in a single plant cell, creating graded bi-directional outputs (stomatal ‘opening’ or ‘closing’ responses). Moreover, cell signaling and genetic studies have shown that guard cell ion channels are targets of signal transduction. Thus detailed insights into the signaling machinery are gained through eletrophysiological recordings of ion channel currents (Schroeder and Hagiwara, 1989; Blatt and Armstrong, 1993; Ward and Schroeder, 1994; Li et al., 2000; Kwak et al., 2003; Mori et al., 2006). Detailed overviews of guard cell signal transduction can be found in several recent reviews (Kuhn and Schroeder, 2003; Pei and Kuchitsu, 2005; Roelfsema and Hedrich, 2005; Israelsson et al., 2006; Pandey et al., 2007; Shimazaki et al., 2007).

Guard cells allow quantitative dissection of the functions of individual genes and proteins within signaling cascades because:

  1. Guard cells can respond to physiological stimuli even after they are isolated, allowing cell biological and time-resolved analyses of stomatal opening and closure in response to various stimuli.

  2. Guard cells respond to several of the classically known hormonal and light stimuli in plants, illustrating that plant receptors and early signaling mechanisms function in these cells.

  3. A network of ion channels in the plasma membrane and tonoplast of guard cells has been characterized that, together with metabolic responses, controls stomatal movements (see Figure 1; Ward et al., 1995). Being targets of early signaling branches, these ion channels provide effective functional signaling and quantitative analysis points to identify and characterize upstream regulators and identify the intermediate targets and signaling branches that are affected either directly or indirectly by these regulators.

  4. Stomatal aperture regulates plant CO2 intake and water loss, thus critically influencing growth and water stress responsiveness (Pei et al., 1998, Hugouvieux et al., 2001).

  5. Patch clamp, Ca2+ imaging and genomic techniques have been developed and adapted to Arabidopsis guard cell signal transduction studies. This allows for molecular genetic, cell biological, biophysical, physiological, and functional genomics analyses of single cell signaling responses (Pei et al., 1997; Pei et al., 1998; Allen et al., 1999; Wang et al., 2001; Hosy et al., 2003; Leonhardt et al., 2004).


A pair of guard cells surrounds each stomatal pore, located in the leaf epidermis of plants, and controls the aperture of this pore. Stomatal movements are regulated by turgor pressure leading to water flux. Stomatal opening is achieved by uptake of solutes into the cytosol and vacuoles resulting in water uptake into the guard cells and, thus, turgor increase drives opening of the stomatal pore. During stomatal opening, H+-ATPases in the plasma membrane mediate proton efflux from the cytoplasm, leading to membrane hyperpolarization (Assmann et al., 1985; Shimazaki et al., 1986), which in turn activates inward-rectifying K+ channels (Schroeder et al., 1984). Moreover, malate is produced from starch and functions as one of the major osmotica causing an increase in turgor pressure in guard cells. Anions including Cl- and NO3- are transported into the guard cell to further increase turgor pressure.
         Stomatal closure is mediated by turgor reduction in guard cells, which is caused by efflux of K+ and anions from guard cells, sucrose removal, and a parallel conversion of the organic acid malate to osmotically inactive starch (MacRobbie, 1998; Schroeder et al., 2001). Membrane depolarization during stomatal closure is mediated by anion channel activation and inhibition of H+-ATPases in the plasma membrane (Keller et al., 1989; Schroeder and Hagiwara, 1989; Shimazaki et al., 2007). An increase in cytosolic Ca2+ concentration ([Ca2+]cyt) occurs via Ca2+ influx through plasma membrane Ca2+ permeable channels and Ca2+ release from internal stores, leading to activation of anion channels (Schroeder and Hagiwara, 1989) that cause positive shifts (depolarization) in the membrane potential of guard cells. This change in membrane potential down-regulates inward-rectifying K+ (K+in) channels (Schroeder et al., 1987; Pilot et al., 2001) and activates outward-rectifying K+ (K+out) channels (Schroeder et al., 1987; Hosy et al., 2003), resulting in a net K+ efflux from guard cells (Figure 1). In addition, during stomatal closure ABA causes an alkalization of the guard cell cytosol (Blatt and Armstrong, 1993) which directly enhances K+out channel activity (Blatt, 1992; Blatt and Armstrong, 1993; Ilan et al., 1994; Miedema and Assmann, 1996), but also down-regulates the transient R-type anion channels (Schulz-Lessdorf et al., 1996). The efflux of both anions and K+ from guard cells via anion and K+out channels contributes to loss of guard cell turgor, which leads to stomatal closing (Schroeder and Hagiwara, 1989; Pei et al., 1997; Vahisalu et al., 2008). Figure 1 shows an extension of early models of the roles of ion channels and transporters in stomatal opening and closing (Schroeder and Hedrich, 1989; McAinsh et al., 1990; Ward et al., 1995).
         The contribution of plasma membrane H+-ATPases to stomatal movements was assessed by studies with transgenic and mutant plants. Fusicoccin-induced stomatal opening is significantly reduced in the transgenic tobacco plants in which the tobacco H+-ATPase PMA4 is co-suppressed (Zhao et al., 2000; Figure 2). Recently, two dominant mutations in the OST2 gene that encodes the AHA1 H+-ATPase were isolated from a genetic screen in Arabidopsis (Merlot et al., 2007; Figure 3). This study found that these dominant mutations caused constitutive activation of the AHA1/OST2 H+-ATPase, and thus, impaired the stomatal closure induced by ABA. These new findings helped shape the model that continuous membrane hyperpolarization by constitutively-activated H+-ATPase mutants counteracts ABA-induced stomatal closure (Merlot et al., 2007; Figure 3).

Vacuolar Channels
As vacuoles can take up over 90% of the guard cell’s volume, over 90% of the ions exported from the cell during stomatal closing must first be transported from vacuoles into the cytosol (MacRobbie, 1995, 1998). [Ca2+]cyt elevation activates vacuolar K+ (VK) channels, which are proposed to provide a pathway for Ca2+-induced K+ release from the vacuole (Ward and Schroeder, 1994; Figure 3). At resting [Ca2+]cyt, K+ efflux from guard cell vacuoles can be mediated by fast vacuolar (FV) channels, allowing for versatile vacuolar K+ efflux pathways (Allen and Sanders, 1996). The Arabidopsis genome contains five TPK/KCO genes which encode “two-pore” K+ channel proteins. A recent study with tpk1 knockout mutants showed that VK channel currents are absent in the mutant indicating that TPK1/KCO1 encodes a major component of the vacuolar VK channels (Gobert et al., 2007). Expression of TPK1 in yeast led to appearance of VK-like channels in yeast vacuoles (Bihler et al., 2005). 14-3-3 proteins physically interact with TPK1/KCO1 and activate TPK1/KCO1 in a dose-dependent manner (Latz et al., 2007). Furthermore, ABA-induced stomatal closure was impaired by the tpk1 mutation (Gobert et al., 2007), providing genetic evidence for the model that VK channels function in mediating vacuolar K+ release for stomatal closing (Ward and Schroeder, 1994; Gobert et al., 2007).
         It has been proposed that calcium release is induced by inositol hexakisphosphate (Lemtiri-Chlieh et al., 2003), and that a protein phosphorylation event upstream of Ca2+ release is required for ABA signaling (Sokolovski et al., 2005). Slow vacuolar (SV) channels are voltage-dependent and slowly activated at elevated [Ca2+]cyt (Hedrich and Neher, 1987). SV channels were first thought to mediate anion transport (Hedrich et al., 1986), but were later shown to be cation channels that are permeable to Ca2+ ions (Ward and Schroeder, 1994; Ward et al., 1995; Allen and Sanders, 1996; Figure 3). The vacuolar Ca2+-dependent channel TPC1 has been identified as an essential subunit of the vacuolar SV channels, because SV type currents are absent in the tpc1 knockout mutant (Peiter et al., 2005). ABA-induced stomatal closure was not affected in tpc1 mutants, whereas external Ca2+-inhibition of stomatal opening was impaired in tpc1 mutants (Peiter et al., 2005). Absence of Ca2+-activated SV currents was confirmed in an independent study (Ranf et al., 2008). SV channels are down-regulated by vacuolar Ca2+ (Pottosin et al., 1997; Pei et al., 1999), but can allow Ca2+ transport from the vacuole to te cytoplasm (Bewell et al., 1999), which is for example illustrated by "tail" currents recorded in pure CaCl2 gradients (Ward et al., 1995). SV channels are ubiquitous and have been found in all plant vacuoles, but in spite of their broad distribution in plant vacuoles, their physiological functions are not yet understood. Further studies are required to address the physiological role for this SV channel protein in plants.
         CLC family genes function as vacuolar anion (nitrate) uptake transporters (De Angeli et al., 2006), and Cl- channels and malate channels can mediate low-affinity vacuolar anion uptake (Pei et al., 1996; Hafke et al., 2003; Kovermann et al., 2007), but the pathways for anion release from vacuoles still remain elusive.


Guard cells respond to a multitude of signals including temperature, partial CO2 pressure, light, humidity, and hormones. For the majority of signals the molecular identity of the sensors is not known, with the notable exception of blue light: The phototropins PHOT1 and PHOT2 were shown to function as blue light receptors in Arabidopsis guard cells (Kinoshita et al., 2001). PHOT1 in de-etiolated seedlings and PHOT2 in leaves were reported to mediate blue light-induced [Ca2+]cyt increases (Baum et al., 1999; Harada et al., 2003). Furthermore, PHOT1 was recently shown to genetically interact with an inositol polyphosphate 5-phosphatase that negatively regulates PHOT1-mediated [Ca2+]cyt increases in root hairs (Chen et al., 2008). Thus, it would be interesting to test whether or not PHOT1 and PHOT2 receptors lead to an increase in cytosolic calcium in guard cells in response to blue light (Figure 2). The RPT2 (Root Phototropism 2) protein is a plant specific protein containing a putative protein-protein interaction domain (Sakai et al., 2000). PHOT1 was shown to form a complex with RPT2 in vivo and to mediate stomatal opening in a RTP2-dependent manner (Inada et al., 2004; Figure 2).
         In guard cells, blue light activates the plasma membrane H+-ATPase by phosphorylation (Kinoshita and Shimazaki, 1999). Because of their importance in stomatal opening, H+-ATPases seem to be a target of multiple modulators. 14-3-3 proteins are conserved in all eukaryotic cells and bind to a variety of signaling proteins. In guard cells, 14-3-3 proteins bind to the phosphorylated plasma membrane H+-ATPase. A 14-3-3 protein was also shown to bind to light-activated, autophosphorylated phototropins in Vicia guard cells (Kinoshita et al., 2003). Thus, it appears that phototropins form a complex with the H+-ATPase via 14-3-3 proteins that possibly maintain an active state (Figure 2). In addition, another protein, VfPIP, interacting with Vicia phototropin was identified from a yeast two-hybrid screen (Emi et al., 2005). This protein shares a high similarity with the dynein light chain, yet its role remains to be determined. Recently, type 1 protein phosphatase (PP1) genes that are expressed in Vicia faba guard cells have been identified as positive regulators of blue light-induced stomatal opening (Takemiya et al., 2006; Figure 2). Introduction of a dominant non-functional mutant PP1 into guard cells inhibited light-induced stomatal opening whereas wild type PP1 had no effect. Furthermore, the PP1 inhibitor tautomycin impaired blue light-induced H+ pumping, phosphorylation of the H+-ATPase, and stomatal opening without affecting the blue light-induced phosphorylation of phototropins. This suggests that PP1 functions downstream of phototropins and upstream of H+-ATPases (Takemiya et al., 2006).
         During light-induced stomatal opening, activation of the plasma membrane H+-ATPase results in membrane hyperpolarization, which cannot be achieved without inactivation of anion channels that provide a large conductance leading to membrane depolarization. Recently it was shown that blue light inhibits the S-type anion channels that are activated by 700 ppm CO2 (Marten et al., 2007a). Furthermore, blue light inhibition of anion channels was not seen in guard cells of the phot1phot2 double mutants, indicating that phototropins also function in this response (Marten et al., 2007a).
         In addition to phototropins, cryptochromes (CRY1 and CRY2), COP1, and NPQ1 also appear to participate in blue light-regulation of stomatal movements (Mao et al., 2005). Compared to wild type, cry1cry2, and phot1phot2 double mutant guard cells are less responsive to blue light, whereas guard cells of a cry1cry2phot1phot2 quadruple mutant did not respond to blue light at all (Mao et al., 2005). Guard cells of COP1 over-expressing plants are hypersensitive to blue light (Mao et al., 2005).

K+in Channels Mediate K+ Uptake During Stomatal Opening
Inward rectifying K+ channels were characterized and proposed to contribute to K+ influx during stomatal opening (Schroeder et al., 1984; Schroeder et al., 1987). The guard cell K+ channel subunit KAT1 and the root-expressed AKT1 homologue were the first cloned Arabidopsis K+in channel genes (Anderson et al., 1992; Sentenac et al., 1992). KAT1 was characterized as the first eukaryotic inward-rectifying K+ channel gene (Schachtman et al., 1992). KAT1 is expressed in guard cells (Nakamura et al., 1995) and expression of dominant negative mutants, as well as knock-out mutant analyses support the model that KAT1 and its homologues play an important role in stomatal opening (Kwak et al., 2001; Lebaudy et al., 2008). KAT2 is also expressed in guard cells (Pilot et al., 2001). Furthermore, expression analysis in Xenopus oocytes and yeast suggests that KAT1 and KAT2 form heteromultimeric channel proteins that are probably responsible for K+ influx into guard cells (Pilot et al., 2001). K+in channel activity is regulated by various factors including [Ca2+]cyt and protein de/phosphorylation. In addition, spermidine was shown to inhibit stomatal opening and induce stomatal closure (Liu et al., 2000; Figure 2). Spermidine is a polyamine that functions as a growth signal in both eukaryotes and prokaryotes. The levels of polyamines increase under stress conditions, suggesting their roles in stress responses (Slocum and Flores, 1991). Addition of spermidine inhibited K+in channel currents in guard cells in a dose-dependent manner, and also inward K+ currents in mesophyll cells expressing KAT1, suggesting a link between stress response and the regulation of K+in channel activity (Liu et al., 2000).
         Phopholipid kinases also modulate stomatal opening. Phosphatidylinositol (PI) 4,5-bisphosphate produced from either PI4P or PI5P was reported to inhibit both light-induced stomatal opening and ABA-induced stomatal closure (Jung et al., 2002; Figure 2). The PI3 kinase and PI4 kinase inhibitors wortmannin and LY294002 impaired ABA-induced [Ca2+]cyt increases and stomatal closure (Jung et al., 2002). PI 4,5-bisphosphate was also shown to inhibit S-type anion channel current in guard cells (Lee et al., 2007). Furthermore, a mutation in the Arabidopsis PIP5 kinase 4 causes reduced stomatal opening in the light (Lee et al., 2007). Stomatal opening of the pip5k4 mutant was rescued by exogenous PI 4,5-bisphosphate, indicating that PIP5K4 is a positive regulator of light-induced stomatal opening (Lee et al., 2007; Figure 2).
         The Arabidopsis genome encodes 11 ROP-type small G proteins that function as a molecular switch in various cellular responses in plants. It was recently shown that ROP2 is expressed in guard cells and translocates to the plasma membrane upon illumination (Jeon et al., 2008). Expression of a constitutively active form of ROP2 negatively modulated stomatal opening in both Vicia and Arabidopsis (Jeon et al., 2008; Figure 2). Furthermore, the guard cell-expressed RIC7 (ROP-Interactive CRIB motif-containing protein 7) physically interacts with ROP2 and translocates to the plasma membrane and cytosol from the nucleus upon illumination, which is dependent on ROP2 activation (Jeon et al., 2008). Expression of RIC7 in Vicia faba guard cells suppresses light-induced stomatal opening (Jeon et al., 2008). These results suggest that RIC7 negatively regulates light-induced stomatal opening by working downstream of ROP2 (Figure 2).
         Transcriptional modulation contributes to the regulation of guard cell signaling cascades, positively or negatively. Two MYB transcription factors have been identified as regulators of stomatal opening. MYB60 was shown to be specifically expressed in Arabidopsis guard cells, and the expression level of MYB60 was reduced by ABA and desiccation (Cominelli et al., 2005). A null mutation in MYB60 leads to a reduction in light-induced stomatal opening and to an enhanced drought tolerance, without affecting ABA sensitivity of guard cells (Cominelli et al., 2005). Furthermore, only 36 genes were shown to be down- or up-regulated in the atmyb60 mutant, suggesting that MYB60 specifically regulates a limited number of genes (Cominelli et al., 2005; Figure 2). In contrast, another MYB transcriptional factor, MYB61, was reported to be highly expressed in guard cells (Liang et al., 2005). Overexpression of MYB61 results in a higher leaf surface temperature (indicative of smaller stomatal apertures), whereas atmyb61 null mutants have a lower leaf surface temperature (larger stomatal apertures), compared to wild type plants (Liang et al., 2005). Similar to the situation with atmyb60, neither overexpression or knockout of MYB61 appears to alter ABA sensitivity of the plants in stomatal movements (Liang et al., 2005). However, dark-induced stomatal closure is partially impaired in atmyb61 (Liang et al., 2005), suggesting that MYB61 is a negative regulator of light-induced stomatal opening and/or a positive regulator of dark-induced stomatal closure (Figure 2).
         In addition to the cytosolic factors, protein trafficking plays a role in regulating K+in channel activity during stomatal movements. During swelling of guard cell protoplasts, an increase in the membrane surface area was reported to be associated with an increase in both K+in channel and K+out channel currents without affecting the activation kinetics of K+ channels (Homann and Thiel, 2002). This result suggests that vesicles containing both K+in and K+out channels are delivered to the plasma membrane during guard cell swelling, resulting in the increase in the membrane surface area and K+ currents. SYP121 is the Arabidopsis homolog of the tobacco syntaxin SYP121/Syr1 that was previously shown to function in ABA-activation of Cl- currents and ABA inhibition of KAT1-mediated K+ currents in oocytes (Leyman et al., 1999). A cell biological study using the dominant-negative Sp2 fragment of SYP121 showed that the Arabidopsis SYP121 protein is specifically responsible for delivery of KAT1 K+in channel to the plasma membrane (Sutter et al., 2006). Furthermore, it was shown that ABA induces specific endocytosis of KAT1 K+in channels in an endosomal pool in guard cells leading to a reduction in the number of KAT1 K+in channel subunits residing in the plasma membrane (Sutter et al., 2007). The endocytosed KAT1 K+in channels were almost entirely recycled back to the plasma membrane 8 hr after removal of ABA (Sutter et al., 2007). These studies indicate that mechanical regulation of K+ channels is involved in stomatal movements (Figure 2).
         In addition to K+in channels, other transporters have been suggested to contribute to stomatal opening. AtCHX20, a proton antiporter that localizes to endomembranes, was reported to positively regulate light-induced stomatal opening (Padmanaban et al., 2007; Figure 2). The CHX family is comprised of 28 members, and their functions are largely unknown (Sze et al., 2004). AtCHX20 is highly and preferentially expressed in guard cells, and Atchx20 T-DNA insertional mutants exhibited a significant reduction in light-induced stomatal opening, implying that the endomembrane cation/H+ exchanger contributes to K+ uptake into vacuoles during stomatal opening (Padmanaban et al., 2007). During stomatal opening, K+ influx occurs which balances an influx of anions, mainly Cl-. Because nitrate is also abundant in soil and transported to plant cells by the nitrate transporter AtNRT1.1/CHL1, it was tested whether AtNRT1.1/CHL1 has a role in stomatal movements (Guo et al., 2003). chl1 mutant plants have reduced stomatal opening, reduced nitrate accumulation, and impaired nitrate-induced membrane depolarization in the light, suggesting that nitrate uptake is accompanying K+ influx and required for stomatal opening (Guo et al., 2003; Figure 2).
         Figure 2 illustrates the signaling cascade from activation of PHOT1, 2 to stomatal opening in terms of known genes and their relative position in the signal transduction pathway. The figure is clickable and linked to an updated version of (Schroeder et al., 2001).


CO2 and humidity are environmental stimuli that regulate stomatal movements. The continual rise in atmospheric pCO2 causes an increase in leaf CO2 and a reduction of stomatal apertures in many plant species on a global scale (Medlyn et al., 2001). However, the mechanisms mediating CO2-induced stomatal closing are still less understood than those triggered by ABA, and until recently no genetic mutants had been identified that robustly impaired high pCO2-induced stomatal closing. CO2-induced stomatal closure is mediated by [Ca2+]cyt (Schwartz et al., 1988; Webb et al., 1996; Young et al., 2006). Moreover, the ABA-insensitive mutant gca2 was identified as a first recessive mutation that shows a strong impairment in high CO2-induced stomatal closing (Young et al., 2006; Figure 3). Mutations in the SLAC1 gene encoding a plasma membrane protein with similarity to a bacterial malate transporter exhibit impairment in CO2-induced stomatal closing (Negi et al., 2008; Vahisalu et al., 2008). SLAC1 is highly expressed in guard cells and localized at the plasma membrane (Negi et al., 2008; Vahisalu et al., 2008). SLAC1 shows homology to malate transporters and S-type anion currents are strongly impaired in slac1 alleles (Vahisalu et al., 2008) and consistent with this, malate contents are significantly higher in guard cells of slac1 null mutant compared to wild type guard cells (Negi et al., 2008). Thus gca2 and slac1 mutants represent first recessive CO2-insensitive mutants that are strongly impaired in stomatal closing (Young et al., 2006; Negi et al., 2008; Vahisalu et al., 2008). These mutants are also impaired in their ABA response, suggesting that the encoded proteins function downstream of a convergence point between CO2- and ABA-induced stomatal closing (Figure 3).
         ABC (ATP-binding cassette) transporters are a large family of multi-spanning membrane proteins. An ABC transporter AtABCB14 was shown to mediate malate uptake when expressed in E. coli and HeLa cells (Lee et al., 2008). AtABCB14 is predominantly expressed in guard cells and localized to the plasma membrane. Atabcb14 mutant plants show slightly more rapid high CO2-induced reduction in stomatal conductance (Lee et al., 2008). This study suggests that AtABCB14 functions during stomatal opening in mediating malate uptake into guard cells, which was released via anion channels during stomatal closing responses and possibly also from mesophyll cells. AtABCB14 is proposed to osmotically enhance stomatal opening and to remove extracellular malate which is known to enhance activation of anion channels (Lee et al., 2008).
         The HT1 protein kinase was identified as a signaling component that mediates a strong negative regulation of CO2-induced stomatal closing (Hashimoto et al., 2006). Thus ht1 mutant alleles that show impaired HT1 kinase activity cause a constitutive high CO2 stomatal closing response, even when CO2 concentrations are clamped to low levels (Hashimoto et al., 2006; Figure 3). ht1 mutant alleles exhibit largely functional responses to blue light and ABA (Hashimoto et al., 2006). In this respect, it is interesting to note that the CO2-hypersensitive response in ht1 mutant alleles, continued to show blue light-induced stomatal opening and ABA-induced stomatal closing, even though the degree of stomatal responses differed quantitatively from wild type (Hashimoto et al., 2006), suggesting that HT1 functions as a central negative regulator of early CO2 signaling (Figure 3). Note that cross talk occurs among stomatal movement responses and therefore any mutation that greatly impairs one pathway is expected to have a secondary effect on other responses. For example a blue-light insensitive mutant will show reduced stomatal opening under many conditions and therefore may be expected to quantitatively affect the degree of ABA-induced stomatal closing. Thus, the blue light receptors are presently not added as mechanisms to the primary ABA signaling cascade. Several genes that affect CO2-regulated stomatal movements have been identified, but the CO2 sensing mechanisms remain unknown, in spite of their importance for global plant gas exchange regulation in light of the continuing increase in atmospheric CO2 (Medlyn et al., 2001).
         The humidity response of stomata is interesting for a number of reasons: (i) Stomatal closure occurs very rapidly in response to a reduction in relative humidity. (ii) A previous study with abi1-1, abi2-1, and aba1 suggests that humidity signaling is not ABA-dependent (Assmann et al., 2000). (iii) High humidity enhances the stomatal sensitivity to CO2 (Talbott et al., 2003). Despite its role in stomatal movements, molecular components of the humidity signaling cascade largely remain unknown. Thermal imaging of leaf temperatures allows one to directly measure leaf surface temperature and thus identify mutants with altered stomatal behavior because plants with larger stomatal apertures have a lower leaf surface temperature due to a higher transpiration rate (Mustilli et al., 2002). In a forward genetic screen, mutants were screened for reduced stomatal closing in response to stepwise reductions in relative humidity of the air surrounding plants (Xie et al., 2006). Interestingly, OST1 and ABA2 were identified as positive modulators of relative humidity-controlled stomatal closure (Xie et al., 2006). This result suggests, in addition to a proposed ABA-independent pathway (Assmann et al., 2000), that humidity signaling is mediated by ABA biosynthesis and signaling, indicating a convergence between ABA and humidity signaling (Xie et al., 2006; Figure 3). Further research on this response should be interesting to understand how plants sense humidity changes and then turn on the above signaling mechanisms.
         Furthermore, it was recently shown that Arabidopsis slac1 mutants are impaired in rapid stomatal closure when induced by low humidity, suggesting an important role of the SLAC1-associated anion channel in low humidity-induced stomatal closing (Vahisalu et al., 2008; Figures 1, 2). The residual and slowed stomatal closing response in slac1 plants may be due to R-type anion channels, which remained intact in slac1 guard cells (Vahisalu et al., 2008; Figures 1, 2). Thus initial insights have been obtained into genetic and physiological mechanisms that mediate low-humidity induced stomatal closing, but more information regarding this important response is needed and early humidity sensing mechanisms remain unknown.


One of the best understood plant signaling networks is the one triggered by ABA in guard cells, causing stomatal closure. As shown in Figure 3, the ion channels which control this response and many genes encoding positive as well as negative regulators of guard cell ABA signaling have been identified in Arabidopsis. Recently, three proposed ABA receptor genes have been cloned from Arabidopsis, encoding the RNA-binding protein FCA, a Mg-chelatase H subunit, and the proposed G-protein coupled receptor GCR2, respectively (Razem et al., 2006; Shen et al., 2006; Liu et al., 2007b). The Mg-chelatase H subunit and GCR2 but not FCA were reported to function in ABA regulation of stomatal movements (Razem et al., 2006; Shen et al., 2006; Liu et al., 2007b), although the GCR2 results could not be reproduced and have been questioned (Gao et al., 2007; Johnston et al., 2007; Liu et al., 2007a). In addition, a weak mutant allele in the Mg--chelatase subunit, cch1 (Harper et al., 2004) shows strong recessive ABA insensitivity phenotypes in ABA-induced stomatal closing and in ABA inhibition of seed germination and root elongation (Shen et al., 2006). It is therefore surprising that mutations in this gene have not been identified in conventional forward genetic screens (Koornneef et al., 1984). In contrast to the cch1 allele, known gun5 mutant alleles in the same Mg-chelatase subunit gene did not affect ABA signaling (Shen et al., 2006). It is interesting to note, in contrast to other plant hormone receptors, that these three proposed ABA receptors do not share any primary structural features.

ABA-Regulated Calcium Signaling

ABA triggers cytosolic calcium ([Ca2+]cyt) increases (Figure 3; McAinsh et al., 1990; Allan et al., 1994; Grabov and Blatt, 1998; Allen et al., 2001; Webb et al., 2001; Klusener et al., 2002; Li et al., 2004; Israelsson et al., 2006; Nomura et al., 2008). Cytosolic Ca2+ elevations in guard cells down-regulate both K+in channels (Schroeder and Hagiwara, 1989) and plasma membrane H+-ATPases (Kinoshita et al., 1995), providing a mechanistic basis for ABA and Ca2+ inhibition of K+ uptake during stomatal closure (Figure 1).
         Hyperpolarization-activated Ca2+-permeable “ICa” channels in the plasma membrane contribute to the increase in [Ca2+]cyt (Hamilton et al., 2000; Pei et al., 2000; Kwak et al., 2003; Figure 3). ABA was shown to activate these plasma membrane Ca2+ channels in both Vicia faba and Arabidopsis (Hamilton et al., 2000; Pei et al., 2000; Murata et al., 2001). Protein phosphorylation plays a role in the activation of Ca2+ channels (Kohler and Blatt, 2002). Two Ca2+-dependent protein kinases, CPK3 and CPK6, are required for ABA activation of S-type anion and Ca2+-permeable channels (Mori et al., 2006). Two other CDPKs, CPK4 and CPK11, also help mediate ABA responses (Zhu et al., 2007). In addition, Ca2+ channels are regulated by hydrogen peroxide leading to increases in [Ca2+]cyt (Pei et al., 2000; Kohler et al., 2003; Kwak et al., 2003; Lemtiri-Chlieh and Berkowitz, 2004; Figure 3). A hyperpolarization-activated Ca2+-permeable current was also activated by cyclic AMP (Lemtiri-Chlieh and Berkowitz, 2004). Further research is needed to determine whether the cAMP-activated Ca2+ channels are the same channels as ABA-regulated ICa channels.
         [Ca2+]cyt elevations activate two different types of anion channels in the plasma membrane of guard cells: Slow-activating sustained S-type (Schroeder and Hagiwara, 1989) and rapid transient (R-type; Hedrich et al., 1990; Roelfsema et al., 2004) anion channels. Both mediate anion release from guard cells, causing depolarization (Figure 1).
         Early intracellular Ca2+ imaging studies showed that a significant number of guard cells do not show measurable [Ca2+]cyt elevations in response to ABA (Schroeder and Hagiwara, 1990; Gilroy et al., 1991; McAinsh et al., 1992). These findings led to the model that both Ca2+-dependent and Ca2+-independent pathways mediate ABA responses (Allan et al., 1994). Based on cells showing no ABA-induced Ca2+ increases the Ca2+-independent pathway has also been proposed and discussed in further studies (Allan et al., 1994; Ward et al., 1995; Romano et al., 2000; Levchenko et al., 2005; Marten et al., 2007b). However, the relative contribution of the proposed Ca2+-independent pathway to ABA-induced stomatal closing has not yet been quantified, and questions of whether known ABA signaling mutations transduce or by-pass this pathway remain unanswered. In addition, recent research on CO2 and ABA signaling has led to the hypothesis that these physiological stomatal closing stimuli sensitize guard cells to cytosolic Ca2+ (Young et al., 2006; Yang et al., 2008). This proposed Ca2+ sensitivity priming might also mediate Ca2+-dependent signaling at lower cytosolic Ca2+ levels (Israelsson et al., 2006). Further work, beyond the negative results from multiple laboratories that led to the Ca2+-independent model, is needed to characterize the Ca2+-independent pathway.
        In addition to promotion of stomatal closure, ABA also inhibits stomatal opening by negatively regulating the plasma membrane H+-ATPases in a dose-dependent manner (Goh et al., 1996). Increases in [Ca2+]cyt were shown to inhibit the ATP hydrolysis and H+ pumping by plasma membrane H+-ATPase, suggesting that [Ca2+]cyt mediates this ABA inhibition of H+-ATPase activity (Kinoshita et al., 1995). Increases in [Ca2+]cyt also inhibit K+in channels and thus contribute to ABA inhibition of stomatal opening (Schroeder and Hagiwara, 1989; Wang et al., 1998).

Essential Function of Anion Channels in Stomatal Closing

Anion channels in the plasma membrane of guard cells were proposed to provide a central control mechanism for stomatal closing (Schroeder and Hagiwara, 1989). Genetic evidence for this model was recently obtained. SLAC1 encodes a plasma membrane protein in Arabidopsis guard cells and is required for the function of S-type anion channels and for CO2-, ABA- and ozone-induced stomatal closing (Negi et al., 2008; Vahisalu et al., 2008; Figure 3). The SLAC1 protein has distant similarity to a bacterial malate transporter. Since S-type anion channels have a permeability to malate anions (Schmidt and Schroeder, 1994), and slac1 mutants lack S-type anion currents in guard cells (Vahisalu et al., 2008), it is thought that SLAC1 may encode a long sought subunit of these guard cell anion channels. slac1 mutant alleles show residual stomatal closing responses to low-humidity, light-dark transitions and ABA, which correlates with the finding that R-type anion channels remain functional in slac1 guard cells (Vahisalu et al., 2008), and R-type channels are thus modeled to function in parallel to S-type anion channels (Keller et al., 1989; Schroeder and Keller, 1992) (Figures 1 and 3). slac1 mutant alleles show greatly reduced responses in ABA-, CO2-, low-humidity, ozone- and Ca2+-induced stomatal closing (Negi et al., 2008; Vahisalu et al., 2008), supporting the model that S-type anion channels are major control mechanisms of stomatal closing (Schroeder and Hagiwara, 1989; Schroeder et al., 1993).
         ABC transporters have been suggested to regulate S-type anion channels or to serve as the channel themselves (Leonhardt et al., 1997; Leonhardt et al., 1999). Two ABC transporters AtMRP4 and AtMRP5 were reported to have opposite regulatory roles in stomatal opening. Null mutations in AtMRP4 resulted in enhanced stomatal opening in response to light, increased transpirational water loss, and enhanced drought sensitivity, indicating that AtMRP4 is a negative regulator of stomatal opening (Klein et al., 2004). Another ABC transporter, AtMRP5, is expressed in guard cells and localizes to the plasma membrane (Suh et al., 2007; Figure 3). Atmrp5-1 knockout mutants show reduced stomatal opening, in response to light, when compared to wild type plant responses (Klein et al., 2003). Atmrp5-1 mutants also display other phenotypes consistent with the observed stomatal behavior, including a reduced transpiration rate, reduced waters loss in detached leaves, enhanced drought tolerance, indicating that AtMRP5 positively regulates stomatal opening (Klein et al., 2003). Furthermore, the atmrp5 null mutant shows reduced sensitivity to ABA in stomatal closing (Gaedeke et al., 2001; Klein et al., 2003). atmrp5 gene disruption impairs Ca2+-activated S-type anion channel currents (Suh et al., 2007). However, in contrast to slac1 alleles, atmrp5 mutations impair both S-type anion channel and Ca2+ channel regulation (Suh et al., 2007; Figure 3), indicating that AtMRP5 may function as a transmembrane ion channel regulator, analogous to its mammalian family members, the cystic fibrosis transmembrane conductance regulator (CFTR) and sulfonylurea receptor (SUR) (Ashcroft, 2006; Gadsby et al., 2006). Whether AtMRP5 directly interacts with ion channel proteins remains presently unknown.

Modulators of ABA Signaling Strength
The gene ERA1 encodes a β-subunit of farnesyltransferase, an enzyme that modifies specific target proteins by attaching a hydrophobic prenyl (farnesyl) group to their C terminus. actin reorganization and stomatal closure (Lemichez et al., 2001). The null rop10 mutant shows enhanced ABA response in stomatal closure, and a dominant-negative mutant of ROP10 partially suppresses abi2-1 mutant phenotypes in seed germination and root elongation (Zheng et al., 2002). Cytosolic Ca2+ also mediates ABA-triggered actin reorganization, which in turn activates mechanosensitive Ca2+ channels (Hwang and Lee, 2001; Zhang et al., 2007a).
         The extracellular Ca2+ sensor CAS was cloned from Arabidopsis using a functional expression assay, combined with Ca2+ imaging, in HEK293 mammalian cells (Han et al., 2003). CAS encodes a plant-specific protein that has a single transmembrane domain. The C-terminal region of CAS shares a sequence similarity with a rhodanese-like domain (PF00581) that might be involved in protein-protein interactions, and several acidic amino acids reside in the N-terminus that was shown to bind to Ca2+ (Han et al., 2003). CAS is mainly expressed in aerial parts of plants including guard cells and mediates external Ca2+-induced [Ca2+]cyt increases by increasing IP3 concentration (Han et al., 2003; Tang et al., 2007; Nomura et al., 2008). In cas KO mutants and CAS antisense plants, external Ca2+-induced stomatal closure is impaired whereas CAS overexpression triggers stomatal closure in the absence of external Ca2+ (Han et al., 2003; Nomura et al., 2008). The cellular targeting of CAS is presently a matter of debate, since there is data indicating plasma membrane and/or thylakoid membrane localizations (Han et al., 2003; Nomura et al., 2008; Figure 3).
         Phospholipids, phospholipid kinases, and phospholipid lipases function in guard cell ABA signaling. The phospholipase C (PLC) blocker U-73122 partially inhibits ABA-induced stomatal closure, suggesting that PLC positively regulates ABA signaling (Staxen et al., 1999). Phospholipase PLDα1 was reported to also function as a positive regulator by producing phophatidic acid (PA) that binds to and inhibits the negative regulator PP2C ABI1 (Zhang et al., 2004a; Figure 3). While ABI1 binding of PA would stimulate ABA promotion of stomatal closure, PA and PLDα1 interact with the GPA1 subunit (see below) to mediate ABA inhibition of stomatal opening, indicating a dual function of PA (Mishra et al., 2006). Pldα1 knock out mutants were reported to be disrupted in ABA-induced stomatal closing, and this disruption was alleviated in abi1 and pldα1 double knockouts, suggesting that PLDα1 is not required for ABA responses in the absence of ABI1. The strong suppressor effect of abi1 knockout on the pldα1 phenotype appears interesting and unique, since many close homologues to the ABI1 protein phosphatase have been shown to have similar functions to ABI1 and are co-expressed in guard cells (Leonhardt et al., 2004; Saez et al., 2004; Kuhn et al., 2006; Saez et al., 2006; Yoshida et al., 2006b). Thus further investigation of the reported interaction of PLDα1 and ABI1 would be interesting.
         Phosphatidylinositol 3- and 4-phosphate, sphingosine-1-phosphate (S1P), and phytosphingosine-1-phosphate (phyS1P) are additional phospholipids that positively regulate ABA signaling, indicating that PI3 kinase, PI4 kinase, and sphingosine kinase are positive effectors of the ABA signaling cascade (Ng et al., 2001; Jung et al., 2002; Coursol et al., 2003; Coursol et al., 2005). In the gpa1 mutant, S1P-induced stomatal closure and ion channel activation are impaired, suggesting that S1P and phyS1P function upstream of GPA1 (Coursol et al., 2003). A pharmacological study combined with calcium imaging analysis suggests that PI3P and PI4P function upstream of [Ca2+]cyt increases (Jung et al., 2002).
         Genetic evidence suggests that the heterotrimeric G protein α subunit (GPA1) mediates ABA signaling. Both ABA inhibition of inward K+ currents and stomatal opening are reduced by the gpa1-1 and gpa1-2 T-DNA insertional mutations (Wang et al., 2001; Figure 3). GPA1 physically interacts with GCR1, a G protein coupled receptor (Pandey and Assmann, 2004). In contrast to the gpa1 mutants, gcr1 mutant guard cells are hypersensitive to ABA and S1P, suggesting that GCR1 is a negative regulator of this ABA response (Pandey and Assmann, 2004). Analysis of ABA responses in seed germination, compared to guard cells, show differential effects of G protein mutants. Both gpa1 and gcr1 mutants show ABA hypersensitivity in seed germination and early seedling development, indicating that GPA1 negatively regulates this signaling pathway (Pandey et al., 2006). Thus G proteins affect ABA signaling and modulate ABA responses in a cell- or tissue-specific manner (Pandey et al., 2006; Pandey et al., 2007).

Reactive oxygen signaling mechanisms in guard cells
Reactive oxygen species (ROS) and the gaseous nitric oxide (NO) function as second messengers in guard cell ABA signaling. ABA induces ROS production in guard cells (Pei et al., 2000; Zhang et al., 2001). ROS lead to activation of hyperpolarization-activated Ca2+ channels and stomatal closure (Pei et al., 2000; Murata et al., 2001). NADPH oxidases are multi spanning membrane proteins that produce extracellular superoxide (Keller et al., 1998; Sagi and Fluhr, 2001; Sagi and Fluhr, 2006). Guard cell microarray experiments showed that of the 10 NADPH oxidase genes in the Arabidopsis genome, AtRbohD and AtRbohF are highly expressed in guard cells (Kwak et al., 2003). Analysis of AtrbohD/F double knock out mutant guard cells, showed that these NADPH oxidases contribute to ABA-triggered ROS production, and function in ABA-activation of plasma membrane Ca2+ channels, and ABA-induced stomatal closure (Kwak et al., 2003; Figure 3). Compared to the mammalian counterpart, one of the unique structural features of plant NADPH oxidases is that there are two EF-hand Ca2+ binding domains in their N-terminus, suggesting the regulation of the protein by Ca2+ (Keller et al., 1998; Torres et al., 1998). Plant NADPH oxidases are activated by Ca2+ (Sagi and Fluhr, 2001; Ogasawara et al., 2008). Furthermore, NADPH oxidases appear to be positively regulated by protein kinases (Kobayashi et al., 2007; Nuhse et al., 2007; Ogasawara et al., 2008).
         The ethylene receptor ETR1 was reported to mediate ethylene- and H2O2-induced stomatal closure, and AtrbohF functions in this pathway (Desikan et al., 2005; Desikan et al., 2006), indicating a signaling convergence between ABA and ethylene signaling pathways. However, ethylene was also shown to counteract and delay ABA-induced stomatal closure (Tanaka et al., 2005). Thus further research would be helpful to illuminate these apparently counter-acting physiological ethylene responses in stomatal regulation.
         Hydrogen peroxide also functions in ABA inhibition of blue light-induced H+ pumping by triggering dephosphorylation of H+-ATPase in the plasma membrane (Zhang et al., 2004b). Hydrogen peroxide was reported to inhibit ATP hydrolysis by the plasma membrane H+-ATPase and blue light-induced phosphorylation of the H+-ATPase, whereas the cellular H2O2 scavenger ascorbate partially rescues ABA inhibition of H+ pumping, indicating that H2O2 is another molecular player mediating ABA inhibition of stomatal closure (Zhang et al., 2004b). Cytoplasmic alkalization and PI3P appear to function upstream of ROS, whereas ABA-induced NO production and stomatal closure rely on ABA-triggered ROS production (Park et al., 2003; Suhita et al., 2004; Bright et al., 2006). NO triggers Ca2+ release from intracellular stores contributing to an increase in the cytosolic Ca2+ concentration (Figure 3), resulting in inhibition of K+in channels and activation of anion channels (Garcia-Mata et al., 2003). The glutathione peroxidase AtGPX3 physically interacts with ABI1 and ABI2 and contributes to the regulation of redox states in guard cells (Miao et al., 2006; Figure 3). In addition, the redox state of the H2O2 scavenger ascorbate plays a role in stomatal movements. Overexpression of dehydroascorbate reductase results in increased ascorbate reduction, a lower H2O2 level in guard cells, and increased stomatal conductance and water transpiration (Chen and Gallie, 2004). Together, these studies show a key role for ROS in modulating the mechanism that mediate stomatal closure, including Ca2+ channel activation (Pei et al., 2000; Murata et al., 2001; Kwak et al., 2003; Kwak et al., 2006), H+ pump inhibition (Zhang et al., 2004b), and, via NO production, K+in channel de-activation and anion channel activation (Garcia-Mata et al., 2003).

Protein kinases and phosphatases in ABA signaling

Protein de/phosphorylation plays a key regulatory role in guard cell ABA signaling. The dominant ABA insensitive abi1-1 and abi2-1 loci were isolated (Koornneef et al., 1984) and identified as closely related type 2C protein phosphatases (Leung et al., 1994; Meyer et al., 1994; Leung et al., 1997; Rodriguez et al., 1998). Hydrogen peroxide inhibits the activities of the negative regulatory type 2C protein phosphatases ABI1 and ABI2 (Meinhard and Grill, 2001; Meinhard et al., 2002). Localization of the mutant abi1-1 protein into the nucleus is required for the ABA-insensitive response (Moes et al., 2008). When the nuclear localization sequence in abi1-1 is disrupted, the transcription of genes regulated by ABA is rescued to wild-type levels, thus suggesting that abi1-1 functions as a hypermorphic allele (Moes et al., 2008). Two additional PP2Cs, HAB1/AtP2C-HA and AtPP2CA, were identified to function as negative regulators of ABA signaling (Leonhardt et al., 2004; Kuhn et al., 2006; Yoshida et al., 2006b; Figure 3). Moreover, the stomatal response to ABA is enhanced when the two negatively regulating PP2C ABI1 and HAB1 are inactivated simultaneously (Saez et al., 2006). The dominant mutations abi1-1 and abi2-1 impair ABA activation of ICa Ca2+ channels (Murata et al., 2001) and S-type anion channels (Pei et al., 1997). ABI2 physically interacts with the SOS2 protein kinase, and this interaction is disrupted by the abi2-1 mutation (Ohta et al., 2003; Figure 3). Note that no pharmacological inhibitors for PP2Cs are known. Therefore, the dominant natures of the abi1-1 and abi2-1 alleles (Koornneef et al., 1984) do not allow to distinguish whether the phenotypes of these mutants occur through direct protein-protein interactions in early ABA signaling, or via less direct, possibly neomorphic or hypermorphic modulation of the early ABA signaling pathway (for detailed previous discussions, see (Pei et al., 1997; Murata et al., 2001; Yoshida et al., 2006b). A recent study showed that ABI1 physically interacts with the OST1/SnRK2E protein kinase, and ABA activation of OST1/SRK2E is inhibited in abi1-1 but not in abi2-1 mutants (Yoshida et al., 2006a). The PKS3 kinase interacts with the calcium binding protein ScaBP5/CBL1 and the PP2C ABI2 and ABI1, and RNAi-mediated silencing of ScaBP5 causes ABA hypersensitivity (Guo et al., 2002; Figure 3).
         In addition to PP2Cs, type2A protein phosphatases also function in ABA signaling. The RCN1 gene encoding the PP2A regulatory subunit A is a positive regulator of the ABA signaling cascade in guard cells and seed germination (Kwak et al., 2002). RCN1 was previously shown to increase PP2A enzyme activity (Deruère et al., 1999). T-DNA insertional mutation in RCN1 causes impairment in both ABA-induced stomatal closure and ABA activation of anion channels, a reduced sensitivity of ABA-triggered cytosolic Ca2+ increases, ABA-insensitive gene expression, and ABA-insensitive seed germination (Kwak et al., 2002; Figure 3). Furthermore, the PP2A inhibitor okadaic acid phenocopies the rcn1 phenotypes in wild type plants (Kwak et al., 2002).
         The putative protein tyrosine phosphatase gene PHS1 was also suggested to negatively regulate ABA signaling (Quettier et al., 2006), suggesting a role for protein tyrosine phosphatases in stomatal movements (MacRobbie, 2002).

mRNA processing and ABA signaling
Transcriptional and post-transcriptional regulations provide cellular mechanisms through which ABA signaling is modulated. For reviews on transcription factors that function in ABA signaling, readers are referred to the following reviews (Giraudat, 1995; Finkelstein et al., 2002; Zhu, 2002; Kuhn and Schroeder, 2003; Yamaguchi-Shinozaki and Shinozaki, 2006). Mutations in either of the nuclear mRNA cap binding protein subunits ABH1 and CBP20 confer ABA hypersensitivity in Arabidopsis, indicating that specific mRNA processing events may be needed for expression of negative regulators of ABA signaling (Hugouvieux et al., 2001; Papp et al., 2004; Figure 3). Expression of the negative regulator AtPP2CA is greatly reduced in abh1 mutants (Hugouvieux et al., 2001) and atpp2ca mutants were identified in two independent forward genetic screens for causing ABA hypersensitivity (Kuhn et al., 2006; Yoshida et al., 2006b). Further analyses of AtPP2CA transcripts in abh1 showed that this effect can contribute to the abh1 phenotype, but detailed analyses also demonstrated that additional mechanisms are required for generating the ABA hypersensitivity of abh1 (Kuhn et al., 2006). The recent finding that the nuclear mRNA CAP binding proteins, including ABH1, function in generating small non-coding RNAs, may provide further insights into mechanisms by which abh1 causes ABA hypersensitivity (Gregory et al., 2008). Other mRNA binding proteins, for which mutations cause ABA hypersensitivity, include the dsRNA-binding protein HYL1, the poly(A)-specific RNase AHG2/AtPARN, and an Sm-like protein (Lu and Fedoroff, 2000; Xiong et al., 2001a; Gregory et al., 2008). These results imply that RNA processing and turnover is utilized by plant cells to regulate ABA signaling or that rate-limiting genes that function in ABA signaling are affected in these mRNA processing mutants (for more details, see Kuhn and Schroeder, 2003).
         Another guard cell-expressed RNA-binding protein GRP7 functions in mRNA export to the cytoplasm and positively regulates ABA-induced stomatal closure (Kim et al., 2008; Figure 3). The AP2 type AtERF7 interacts with the PKS3 kinase and a putative transcription repressor AtSin3 to negatively regulate ABA signaling (Song et al., 2005). Studies with transgenic plants showed that over-expression of the MYB44 transcription factor enhances ABA sensitivity in stomatal closure (Jung et al., 2008). Transgenic plants overexpressing MYB44 have reduced levels of negatively acting PP2Cs including ABI1, ABI2, AtPP2CA, HAB1, and HAB2 (Jung et al., 2008). Two calcium-dependent protein kinase genes CPK4 and CPK11 were also reported to positively regulate ABA signaling, possibly through ABF1 and ABF4 transcription factors (Zhu et al., 2007; Figure 3).
         The syntaxin protein OSM1/SYP61 and the RING finger E3 ligase protein SDIR1 were also found to positively control guard cell ABA signaling (Zhu et al., 2002; Zhang et al., 2007b).


Many molecular players of guard cell ABA signaling have been identified, suggesting a complex ABA signaling network in guard cells. Some mechanisms are central to ABA signaling and stomatal closing, such as the different classes of guard cell ion channels and H+ pumps (Ward et al., 1995; Kinoshita and Shimazaki, 1999; Pei et al., 2000), the OST1 kinase (Mustilli et al., 2002; Yoshida et al., 2002), the SLAC1 anion channel associated membrane protein (Negi et al., 2008; Vahisalu et al., 2008), cytosolic Ca2+ (Schroeder and Hagiwara, 1989; McAinsh et al., 1990; Mori et al., 2006; Zhu et al., 2007) and as negative regulators, the clade of PP2Cs that include ABI1 and ABI2. These data indicate that there are signaling branches within ABA signaling, but that there are also essential signaling nodes, including the above mechanisms (Hetherington and Woodward, 2003). Thus it appears likely that some mechanisms more directly mediate ABA signaling, and other mechanisms may be peripheral modulators. However, more research is needed to distinguish such mechanisms. An important aspect in this respect is the detailed quantification of the degree of ABA-induced stomatal closing that occurs in each experimental treatment, which can be best analyzed in “blind” analyses of stomatal signaling (Murata et al., 2001; Mori et al., 2006). Moreover, many genes and mechanisms are waiting to be identified, including the mechanisms that mediate CO2 sensing, humidity sensing as well as understanding the diverse proposed ABA receptors.
         Guard cells use a complex signaling network to create a “graded binary” output that can readily be observed under the microscope: stomatal 'opening' or 'closing'. The study of guard cell signaling provides insights into how the many cellular processes assemble together to create a quantifiable single cell output. Thus, identification and characterization of genes that are highly and specifically expressed in guard cells using various approaches, including gene trap screens and cell-type specific microarray analyses (Leonhardt et al., 2004; Galbiati et al., 2008; Yang et al., 2008), would help to provide a further understanding of ABA signal transduction networks. Furthermore, dynamic modeling and computational simulations of ABA signaling as done by Li et al. (2006) could lead to comprehensive and quantitative information on ABA signaling in guard cells.


We thank Annual Reviews for permission to use the review by Schroeder et al., 2001 for this electronic model and for electronically linked text. We also thank Michelle Turek for critical reading of this manuscript and Daren Miller for help with the webpage. Preparation of this article and research from the authors' laboratories were supported by NSF (MCB0417118), NIH (GM060396) and DOE (DE-FG02-03ER15449) grants to J.I.S, and by NSF (MCB-0614203, MCB-0618402) and USDA (2004-35100-14909, 2007-35100-18377) grants to J.M.K., and a Swiss NSF grant (PP00A-114819) to P.M.


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