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Logo of nihpaAbout Author manuscriptsSubmit a manuscriptHHS Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
Nature. Author manuscript; available in PMC 2010 April 23.
Published in final edited form as:
PMCID: PMC2858982

SLAC1 is required for plant guard cell S-type anion channel function in stomatal signalling


Stomatal pores, formed by two surrounding guard cells in the epidermis of plant leaves, allow influx of atmospheric carbon dioxide in exchange for transpirational water loss. Stomata also restrict the entry of ozone—an important air pollutant that has an increasingly negative impact on crop yields, and thus global carbon fixation1 and climate change2. The aperture of stomatal pores is regulated by the transport of osmotically active ions and metabolites across guard cell membranes3,4. Despite the vital role of guard cells in controlling plant water loss3,4, ozone sensitivity1,2 and CO2 supply2,57, the genes encoding some of the main regulators of stomatal movements remain unknown. It has been proposed that guard cell anion channels function as important regulators of stomatal closure and are essential in mediating stomatal responses to physiological and stress stimuli3,4,8. However, the genes encoding membrane proteins that mediate guard cell anion efflux have not yet been identified. Here we report the mapping and characterization of an ozone-sensitive Arabidopsis thaliana mutant, slac1. We show that SLAC1 (SLOW ANION CHANNEL-ASSOCIATED 1) is preferentially expressed in guard cells and encodes a distant homologue of fungal and bacterial dicarboxylate/malic acid transport proteins. The plasma membrane protein SLAC1 is essential for stomatal closure in response to CO2, abscisic acid, ozone, light/dark transitions, humidity change, calcium ions, hydrogen peroxide and nitric oxide. Mutations in SLAC1 impair slow (S-type) anion channel currents that are activated by cytosolic Ca2+ and abscisic acid, but do not affect rapid (R-type) anion channel currents or Ca2+ channel function. A low homology of SLAC1 to bacterial and fungal organic acid transport proteins, and the permeability of S-type anion channels to malate9 suggest a vital role for SLAC1 in the function of S-type anion channels.

Stomatal aperture is regulated by light, plant water status, CO2 concentration, relative air humidity, and among other stresses, drought and ozone (O3)3,4. A number of signalling compounds, including abscisic acid (ABA), reactive oxygen species (ROS), nitric oxide (NO) and Ca2+ ions are involved in the regulation of stomatal aperture3,4. Adjustment of stomatal apertures is achieved by controlled transport of osmotically active ions and organic metabolites, including potassium (K+), chloride (Cl) and malate across guard cell membranes3,8,10,11, resulting in changes in osmotic potential. Anion channels have been proposed to function as central regulators of stomatal closure8,11 by mediating anion efflux and causing membrane depolarization, which controls K+ efflux through K+ channels. So far, none of the candidates for plant anion channels — the plant homologues to the animal CLC chloride channels — has been localized to the plasma membrane10, and the first plant CLC channel that was functionally characterized encodes a central vacuolar proton/nitrate exchanger12, rather than an anion channel. Thus, despite their proposed importance in several physiological and stress responses in plants8,10,11, the molecular identity of the guard cell plasma membrane proteins that mediate anion channel activity has remained unknown.

In a mutant screen for O3 sensitivity, a series of Arabidopsis ethyl methanesulphonate (EMS) mutants called radical-induced cell death (rcd) was identified13,14. One of them, a recessive mutant originally referred to as rcd3 (ref. 14) and here renamed slac1 (slow anion channel-associated 1), showed constitutively higher stomatal conductance than the wild type (Columbia, Col-0) (Fig. 1a). Interestingly, both rapid transient15 and long-term O3-induced decreases in stomatal conductance were abolished in slac1 (Fig. 1a). Water loss from excised slac1 leaves resulted in 70–80% fresh weight loss after 90 min, whereas in the wild type, fresh weight loss was only 30% after 90 min (Fig. 1b). These differences in fresh weight loss were not a result of variation in stomatal number because slac1 and wild-type leaves have similar stomatal density (Supplementary Fig. 1). Microarray analyses using messenger RNAs from 3-week-old rosette leaves did not reveal any significant differences in gene expression between slac1 and the wild type when grown under optimal conditions. Furthermore, no other phenotypic differences have been observed between slac1 and the wild type. Together these data suggested that the defect in slac1 lies in defective stomatal regulation and that the O3 damage of slac1 leaves (Supplementary Fig. 2) is a result of increased O3 flux into leaves through more open stomata.

Figure 1
Membrane protein SLAC1 controls leaf ozone and water-loss responses

The slac1-1 mutation was identified in the gene At1g12480 by a combination of mapping, candidate gene expression in guard cell microarrays, and analyses of transfer DNA (T-DNA) insertion mutants (see Supplementary Information). SLAC1 encodes a predicted membrane protein of 556 amino acids with a calculated molecular weight of 63.2 kDa and a predicted isoelectric point of 9.58. SLAC1 has hydrophilic amino- and carboxy-terminal tails (189 and 60 amino acids, respectively) and 10 predicted transmembrane helices (Fig. 1c, Supplementary Fig. 3), which contain a C4-dicarboxylate transporter/malic acid transport protein domain (InterPro: IPR004695) defined from the Escherichia coli TehA and Schizosaccharomyces pombe Mae1 proteins. Mae1 is involved in malate uptake16. TehA and Mae1 lack the long hydrophilic tail present in the N terminus of SLAC1, but show a weak, 15–20% amino-acid identity over the transmembrane region with SLAC1 (Supplementary Fig. 4a). SLAC1 shows no homology to the aluminium-activated malate transporters that function in plant aluminium resistance17. Homozygous T-DNA insertion lines (SALK_099139 and SALK_137265, referred to as slac1-3 and slac1-4, respectively; Fig. 1c) both showed similar recessive inheritance, and exhibited similar fresh weight loss from excised leaves as slac1-1 (Fig. 1b). A genomic copy of SLAC1 complemented the mutant phenotype in stably transformed slac1-1 (Fig. 1b).

SLAC1 belongs to a small family of five proteins in Arabidopsis. Three of the proteins, including SLAC1, have a long hydrophilic N-terminal tail, whereas two have only the transmembrane domains. Rice has nine orthologous proteins. The SLAC1 protein is more similar to its rice orthologue Os04g48530 than to the four other Arabidopsis SLAC1 homologues (Supplementary Fig. 4a, b). The transmembrane domains of Arabidopsis and rice SLAC1 homologues and orthologues have several highly conserved amino acids. However, SLAC1 and Os04g48530 also differ from the rest of the proteins in several amino-acid residues (Supplementary Fig. 4a). For example, the amino acid that is mutated in slac1-1 is a serine in SLAC1 and Os04g48530, whereas other family members have an alanine residue in the same position. The serine mutated in slac1-1 is surrounded by three conserved threonine residues, suggesting that this region is significant for either the structure, function or regulation of the protein. Additionally, the predicted intracellular loops between the transmembrane domains have several conserved, positively charged amino-acid residues (Supplementary Figs 3, 4a), also suggesting functional significance.

When 1,582 base pairs (bp) of genomic sequence upstream of the SLAC1 translation start were fused to the reporter gene uidA, the resulting β-glucuronidase (GUS) activity in transgenic plants was localized predominantly to guard cells (Fig. 1d), and occasionally to the vascular strands close to the leaf margins (Fig. 1e). No GUS activity was detected in other parts of the plants. Expression data at the Genevestigator database18 and comparison of gene expression between guard cell and mesophyll cell microarrays also suggest strong preferential guard cell expression of SLAC1.

To study the subcellular location of the SLAC1 protein, green fluorescence protein (GFP) fused to the SLAC1 C terminus was transiently expressed in onion epidermal cells (Fig. 1f–k) and in tobacco protoplasts (Supplementary Fig. 5). Fluorescence and confocal imaging showed that in onion epidermal cells, fluorescence from the SLAC1::GFP fusion protein (Fig. 1f) and the membrane-specific stain FM 4-64 (Fig. 1g) colocalized in merged images (Fig. 1h). GFP fluorescence was observed between the cell wall and the nucleus (Fig. 1j; Supplementary Movie), and was connected to the cell wall through Hechtian strands in plasmolysed cells (Fig. 1k), correlating with plasma membrane localization. Expression in tobacco protoplasts showed results that are consistent with plasma membrane localization (Supplementary Fig. 5).

Stomatal aperture is under environmental and hormonal control. We analysed stimulus responses in stomatal conductance by comparing intact15 slac1 with wild-type plants. Stomatal conductance in slac1 was about 1.5-fold higher during the light period (Fig. 2a). Also, the decline in stomatal conductance at the beginning of the dark period took more than 1 h longer in slac1 compared with the wild type (Fig. 2a). Light/dark transitions during the normal light period caused rapid changes in stomatal conductance in the wild type, whereas slac1 showed a slow and modest response (Fig. 2b). slac1 exhibited a much slower response than the wild type to a decrease in the relative air humidity (Fig. 2c), which is known to cause a rapid reduction of stomatal conductance19. Doubling of [CO2] from 400 p.p.m. to 800 p.p.m. reduced stomatal conductance effectively in the wild type, whereas slac1 showed no responses (Fig. 2d). Thus, slac1 stomata show only a slow and modest response to changes in light and air humidity, and are completely insensitive to O3 stress (Fig. 1a) and elevated [CO2] (Fig. 2d).

Figure 2
Mutations in SLAC1 impair stomatal responses to changes in environment

The concentration of the plant stress hormone ABA increases under drought and induces stomatal closure through second messengers, including ROS, cytosolic Ca2+ and NO2022. We measured stomatal responses to ABA, hydrogen peroxide (H2O2), NO and repetitive Ca2+ pulses (Fig. 3). Stomata of slac1 mutants showed a strong insensitivity to ABA (Fig. 3a and Supplementary Fig. 6a). Similarly, they showed significantly reduced responses to H2O2 (Fig. 3b) and the NO donor sodium nitroprusside (SNP) (Fig. 3c). Transient addition and removal of Ca2+ to the extracellular solution bathing leaf epidermides, while shifting the K+ equilibrium potential, allows experimental imposition of defined intracellular Ca2+ transients in guard cells, resulting in stomatal closure2325. Four repetitive 5-min pulses of 1 mM external Ca2+ were applied (Fig. 3d; top inset; Supplementary Fig. 7). The imposed intracellular Ca2+ ([Ca2+]i) oscillation pattern of slac1-1 guard cells was similar to that of the wild type (Supplementary Fig. 7). The average amplitudes of imposed [Ca2+ ]i transients and the integrated total [Ca2+]i increases per period were statistically similar in wild-type and slac1-1 guard cells (see Supplementary Information). Imposed [Ca2+]i transients caused the typical downstream Ca2+-induced reactive and programmed 23,24 stomatal closure in the wild type, whereas the response was greatly impaired in slac1-1 and slac1-3 (Fig. 3d). Thus slac1 mutant guard cells do not abrogate imposed cytosolic Ca2+ oscillations, but show a strong impairment in downstream Ca2+ oscillation-induced stomatal closing.

Figure 3
Impaired stomatal responses to ABA, H2O2, NO and Ca2+ in slac1

The activation of S- and R-type anion efflux channels, both of which can transmit Cl and malate efflux from guard cells8,9,11, is proposed to decrease guard cell osmotic potential, leading to stomatal closure3,4,8,10,11. This is consistent with Cl and malate efflux occurring in response to ABA26,27. We therefore applied whole-cell patch clamp techniques to characterize the functioning of S-type and R-type anion channel activities. In wild-type guard cells, elevated cytosolic Ca2+ (2 µM) activated ion currents that were selective for Cl over caesium ions (Cs+) (n = 16 guard cells) and showed a relative permeability ratio for malate to chloride anions of 0.125 (n = 12 guard cells), consistent with previous anion selectivity analyses of S-type anion channel currents9 (Supplementary Fig. 8).

S-type anion currents were readily recorded in wild-type guard cells (Fig. 4a, d). However, only very small combined background whole-cell membrane currents and patch-clamp seal currents were observed in slac1-1 and slac1-3 guard cells (Fig. 4b–d). R-type anion currents11 were activated as described25,28. Interestingly, no significant differences in R-type anion currents between wild-type and slac1 guard cells were observed (Fig. 4e, f). Similarly, ABA activation of Ca2+-permeable ‘ICa’ channel currents21 was not disrupted in slac1 guard cells (Supplementary Fig. 9). However, when ABA activation of S-type anion channels was analysed, slac1 mutants showed only small whole-cell currents (Fig. 4h–j), whereas S-type anion currents were recorded in wild-type guard cells (Fig. 4g, j).

Figure 4
Ca2+ and ABA activations of S-type anion channels are impaired in slac1 guard cells

Continuing increases in ozone concentrations in the troposphere owing to human activities are predicted to have a negative affect on crop yields and global carbon sinks in the future1,2. The ozone sensitivity of slac1 leaves (Supplementary Fig. 2), the predominant guard cell expression of SLAC1 (Fig. 1d, e) and abolishment of O3-induced stomatal closure in slac1 mutants (Fig. 1a) together provide direct genetic evidence for the importance of O3 sensing in guard cells for plant O3 tolerance. Only a few plant mutants are known that show CO2 insensitivity5,6 or a constitutive high CO2 response7 in stomatal movements, but no recessive CO2-insensitive mutant gene has been isolated so far. All slac1 alleles are recessive and show a complete lack of high CO2-induced stomatal closure (Fig. 2), illustrating that the SLAC1 protein is a central positive mediator of CO2-induced stomatal closure.

Experiments with ABA, ROS, NO and Ca2+ suggest that SLAC1 is an essential protein functioning downstream of these messengers in mediating stomatal closure (Figs 3, ,44 and Supplementary Figs 6, 7, 9). The phenotype of slac1 differs from the ATP-binding cassette transporter mutant, atmrp5, which shows partial repression of ABA-induced stomatal closure, partial S-type anion current activity and impaired Ca2+ channel activation29. The strong impairment in S-type anion channel and normal Ca2+ channel activity in slac1 guard cells is consistent with SLAC1 being more closely associated with S-type anion channels than is AtMRP5, and provides direct genetic evidence for the model that these anion channels function as a central control mechanism for stomatal closure8.

R-type anion channel activity was not disrupted in slac1 guard cells (Fig. 4e, f), providing genetic evidence for a molecular separation of the membrane proteins required for S- and R-type anion channels. It remains possible that these anion channel types share other protein subunits30. R-type channels may be responsible for the slow stomatal conductance decrease observed in response to light/dark transitions and decrease in relative humidity (Fig. 2a–c).

The data presented demonstrate that SLAC1 encodes an essential subunit for S-type anion channel function or regulation. The low homology of SLAC1 to bacterial and fungal organic acid transporters indicates a possible role for SLAC1 in contributing to formation of an anion-transporting pore. Further research on SLAC1 and its homologues should increase the general understanding of plasma-membrane anion channel structure and regulation in plants.


Three- to six-week-old A. thaliana plants grown in a controlled environment were used. slac1-1 was isolated from an O3-sensitivity mutant screen13. The mapping population was generated by outcrossing to Ler, and an impaired water-loss phenotype was used as a mapping trait. For water-loss analyses, the weight of the detached leaves was followed. Whole-plant stomatal conductance responses to O3, light/dark transitions, elevated CO2 and lowered humidity were measured using the Arabidopsis whole-rosette gas-exchange system15. For GUS activity and complementation analyses, transgenic SLAC1 promoter-driven GUS expression lines and complementation lines with SLAC1 genomic DNA were analysed. For transient gene-expression studies, a SLAC1::GFP fusion protein under the control of a 35S promoter was delivered into onion epidermides by particle bombardment, and to tobacco protoplasts by electroporation. Images were acquired by confocal microscopy. For stomatal responses to H2O2, NO and ABA, stomatal apertures were measured from extracted epidermal fragments after pre-incubation of leaves in opening buffer. Stomatal responses to Ca2+ transients and Ca2+ imaging experiments were analysed in intact leaf epidermides by imposing extracellular calcium pulses23,25. For electrophysiological analyses, Arabidopsis guard cell protoplasts were isolated enzymatically, and Ca2+ activation of S- and R-type anion currents and ABA activation of S-type anion and ICa Ca2+ currents were recorded as described25,29.

Supplementary Material



We thank M. Uuskallio and I. Puzõrjova for technical help. This research was supported by the Academy of Finland Centre of Excellence programme and Helsinki University Environmental Research Centre (to J.K.), by Estonian Science Foundation and University of Tartu start-up grants (to H.K.), by NIH, NSF and, in part, DOE grants (to J.I.S.), and a Leverhulme Trust Early Career Fellowship (to R.D.)


Full Methods and any associated references are available in the online version of the paper at

Supplementary Information is linked to the online version of the paper at

Author Contributions T.V., H.K. and Y.-F.W. contributed equally to this work. J.K. and H.K. designed the experiments in Figs 1 and and2.2. A.L., H.K. and T.V. identified the SLAC1 gene. T.V. and M.B. performed the expression, complementation and subcellular localization analyses in Fig. 1 and Supplementary Fig. 5. H.K. and H.M. performed experiments in Fig. 2. H.K. performed experiments in Supplementary Figs 1 and 2. R.D. designed and performed experiments in Fig. 3b, c and Supplementary Fig. 6b. J.I.S. and J.K. designed experiments in Figs 3a and d, and and4,4, and Supplementary Figs 6a, 7, 8 and 9. W.-Y.C. and G.V. performed experiments in Fig. 3d and Supplementary Fig. 6a. N.N. performed experiments in Fig. 3a and Supplementary Fig. 7. Y.-F.W. performed experiments in Fig. 4 and Supplementary Figs 8 and 9. J.K. and J.I.S. wrote the paper. All the authors discussed the results, and commented on and edited the manuscript.

The primary microarray data reported has been deposited with the ArrayExpress database under accession number E-MEXP-1388.

Reprints and permissions information is available at


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