arabidopsis ht1 kinase controls stomatal movements in ......wt plants and no significant response in...

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NATURE CELL BIOLOGY VOLUME 8 | NUMBER 4 | APRIL 2006 391

Arabidopsis HT1 kinase controls stomatal movements in response to CO2Mimi Hashimoto1, Juntaro Negi1, Jared Young2, Maria Israelsson2, Julian I. Schroeder2 and Koh Iba1,3

Guard cells, which form stomata in leaf epidermes, sense a multitude of environmental signals and integrate this information to regulate stomatal movements1,2. Compared with the advanced understanding of light and water stress responses in guard cells2–4, the molecular mechanisms that underlie stomatal CO2 signalling have remained relatively obscure. With a high-throughput leaf thermal imaging CO2 screen, we report the isolation of two allelic Arabidopsis mutants (high leaf temperature 1; ht1-1 and ht1-2) that are altered in their ability to control stomatal movements in response to CO2. The strong allele, ht1-2, exhibits a markedly impaired CO2 response but shows functional responses to blue light, fusicoccin and abscisic acid (ABA), indicating a role for HT1 in stomatal CO2 signalling. HT1 encodes a protein kinase that is expressed mainly in guard cells. Phosphorylation assays demonstrate that the activity of the HT1 protein carrying the ht1-1 or ht1-2 mutation is greatly impaired or abolished, respectively. Furthermore, dominant-negative HT1(K113W) transgenic plants, which lack HT1 kinase activity, show a disrupted CO2 response. These findings indicate that the HT1 kinase is important for regulation of stomatal movements and its function is more pronounced in response to CO2 than it is to ABA or light.

Stomata in the leaf epidermis provide the major pathway for gas exchange between plants and their environment. The primary role of stomata is to optimize the exchange of CO2 and water vapour between the intra-cellular spaces in leaves and the atmosphere under changing environ-mental conditions. CO2 is a physiological signal that regulates stomatal movements1. Stomata open at low CO2 concentrations and close at high CO2 concentrations (ref. 5). Day/night changes in leaf tissue CO2 con-centrations are caused by photosynthesis and respiration. Furthermore, atmospheric CO2 is predicted to double in the present century

6, which will have unpredictable ecological consequences as changes in CO2 concen-tration affect stomatal regulation5,7. Biochemical and electrophysiologi-cal studies have provided insights concerning important factors in CO2 signalling, including cytosolic Ca2+, as well as K+ and anion channels8–15.

However, CO2 signal-transduction mechanisms that function upstream of guard-cell ion channels are largely unknown. One reason may be that mutants with strongly impaired stomatal responses to CO2 have, so far, not been isolated. Thermal imaging facilitates the estimation of stomatal aperture because leaf temperature correlates with water transpiration rate due to evaporative cooling16.

To study the molecular processes that mediate CO2 signalling in guard cells, we have screened for mutants with altered CO2 responses using thermal imaging. Plantlets derived from an ethyl methanesulfonate (EMS)-mutagenized population in the Arabidopsis ecotype Columbia (Col-0) were grown for 3 weeks in a growth chamber, and were then subjected to low CO2 concentration (100 ppm) for 1 h before analysis by thermography. Low CO2 concentration causes stomatal opening (Fig. 1c) and, therefore, measurable leaf cooling in wild-type (WT) plants (Fig. 1a, b). By screening approximately 40,000 M2 plants, two mutant lines, ht1-1 and ht1-2 (high leaf temperature 1), were isolated; these mutants exhibited higher leaf temperatures than the other plants under low CO2 concen-trations (Fig. 1a, b; n = 12, P < 0.02, Student’s t-test). WT plants showed significant leaf temperature changes in response to changes from ambi-ent CO2 concentration (350 ppm) to low CO2 concentration (100 ppm; WT; Fig. 1a, b; n = 12, P < 0.001) or doubled CO2 concentration (700 pp; WT; Fig. 1a, b; n = 12, P < 0.02). The ability to respond to CO2 concentra-tion changes was reduced (ht1-1; Fig. 1a, b; n = 12, P > 0.6, for 350 ppm to 700 ppm CO2) or absent (ht1-2; Fig. 1a, b; n = 12, P > 0.9) in the ht1 mutants. Representative time-courses of changing leaf surface tempera-tures that were measured using an alternative approach in the WT and mutant plants can be seen in Supplementary Information, Fig. S1.

In plants grown under atmospheric CO2 concentration for 3 weeks in a growth chamber, the stomatal density on the abaxial surface of leaves was similar in WT and ht1 plants (125 ± 15.8, 124 ± 14.3 and 122 ± 10.2 stomata/mm2 in WT, ht1-1 and ht1-2, respectively; means ± SD; n = 18). Stomatal size, growth rate, and morphology of vegetative and floral organs were identical for the three genotypes (data not shown). The dependence of stomatal aperture on CO2 concentration in the WT and ht1 mutants (Fig. 1c) paralleled that of leaf temperature (Fig. 1a, b). Our results indi-cated that the higher leaf temperatures in ht1 were due to a reduction

1Department of Biology, Faculty of Sciences, Kyushu University, Fukuoka 812-8581, Japan. 2Cell and Developmental Biology Section, Division of Biological Sciences, and Center for Molecular Genetics, University of California, San Diego, La Jolla, CA 92093-0116, USA.3Correspondence should be addressed to K.I. (e-mail: [email protected])

Received 15 December 2005; accepted 15 February 2006; published online 5 March 2006; DOI: 10.1038/ncb1387

print ncb1387.indd 391print ncb1387.indd 391 14/3/06 4:54:17 pm14/3/06 4:54:17 pm

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392 NATURE CELL BIOLOGY VOLUME 8 | NUMBER 4 | APRIL 2006

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of stomatal aperture rather than to a reduced stomatal density, and that differences in leaf temperature CO2 concentrations responses in ht1-1 and ht1-2 compared with WT responses correlated well with differential stomatal aperture responses to CO2 concentration changes (Fig. 1).

We investigated the response of stomatal conductance in ht1 and WT control plants to changes in the CO2 content in the air (Fig. 1d, e). In intact leaves, increases in CO2 concentration from 365 ppm to 800 ppm induced a pronounced decrease in stomatal conductance in the WT plants and no significant response in the ht1-1 (P = 0.4) and ht1-2 (P = 0.13) mutants. Raising CO2 concentration to 2000 ppm induced a further reduction in stomatal conductance in the WT plants, no sig-nificant response in ht1-1 (P = 0.6) and, intriguingly, a small increase in ht1-2 (P < 0.01; Fig. 1d). Decreasing CO2 concentration from 365 to 100 ppm induced a large increase in stomatal conductance in the WT plants, a smaller one in ht1-1 mutants (P = 0.05) and no significant response in ht1-2 mutants (P = 0.1; Fig. 1e). An additional drop to 0 ppm induced a further small increase in WT plants, an increase in the ht1-1

mutants and a small decrease in the ht1-2 mutants (P < 0.05; Fig. 1e). In summary, ht1-1 stomatal conductance did not change in response to increases in CO2 concentration above ambient levels (Fig. 1d), and the sensitivity of ht1-1 to reductions in CO2 concentration was shifted to lower CO2 concentrations relative to WT plants, reflecting a pos-sible CO2 hypersensitive response (Fig. 1e), which correlates with leaf temperature and stomatal movement responses to CO2 concentration elevation (Fig. 1, a–c). The slight inverse responses that are found in the ht1-2 mutant (Fig. 1d, e) may be the result of the intact activity of a counter-balancing regulator of the normal CO2-induced stomatal response in the absence of the HT1 regulator.

The responses of ht1 mutants to light and dark exposures with CO2 concentration held at ambient levels were analyzed to determine whether the mutant alleles are insensitive to light changes. Darkness caused drops in stomatal conductance down to similar levels in the three lines (Fig. 2a). Increasing light intensity resulted in a large increase in stomatal conductance in the WT plants and smaller responses in the ht1-1 and

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Figure 1 Changes in leaf temperature of wild-type plants and ht1 mutants in response to CO2 concentration. Leaf temperature changes correlate with CO2-induced stomatal movement responses. (a) Three-week-old plants (images on the right) were subjected to low (100 ppm), normal (350 ppm) and high (700 ppm) CO2 concentration. Thermal images of wild-type (WT), ht1-1 and ht1-2 seedlings. ht1-1:HT1, ht1-1 mutant plant harbouring a transgene with the HT1 genomic sequence. ht1-2:35SHT1, ht1-2 mutant plant harbouring

a transgene with the HT1 cDNA. (b) Leaf temperature calculated from the quantification of infrared images (means ± SD of measurements on ~5,000 square pixels; n = 6 plants per condition). (c) CO2 response of stomatal apertures in ht1 mutants. Data represent means ± SEM (n = 60) of three independent experiments. (d, e) Time courses of stomatal conductance in response to changes in CO2 concentration in ht1 mutants and WT leaves. Data represent the mean ± SEM of three leaves from 5–6-week-old plants.




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ht1-2 mutants (Fig. 2b). We further analyzed whether the ht1-2 mutant exhibits a general inability to open its stomata. As guard cells have a cell-autonomous blue-light response4, we analyzed blue-light-induced changes in stomatal conductance in intact leaves of the ht1 mutants. In both ht1 mutants and WT plants, blue light caused significant induction of stomatal opening, with similar kinetics being present in mutants and WT plants (Fig. 2c; P < 0.005 for ht1-1 from darkness to 30 min blue light and P < 0.005 for ht1-2 for the same conditions). Blue light and the fungal phytotoxin fusicoccin (FC) induce stomatal opening by activation of the

plasma membrane H+-ATPase17. Stomatal opening in response to FC was also observed in ht1 mutants (ht1-1 or ht1-2, Fig. 2d; n = 120, P < 0.01, for controls vs. FC). These results demonstrate that the ht1-2 mutant retains the ability to respond to blue light and exhibits stomatal opening, although ht1-2 did not exhibit stomatal opening in response to low CO2 concentration (Figs 1c, e and 2c, d). One reason for the reduced white-light response in ht1 mutants (Fig. 2a, b) might be that stomatal opening by light is induced, to a significant extent, by a decrease in CO2 through photosynthesis18,19. Furthermore, a degree of crosstalk among stomatal

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Figure 2 Light responses in ht1 mutants at ambient external CO2 concentration. (a, b) Time courses of stomatal conductance in response to changes in white-light intensities in ht1 mutants and wild-type (WT) leaves. Data represent the mean ± SEM of three leaves from 5–6-week-old plants. (c) ht1 mutants show blue-light responses. Data represent the mean ± SEM of five leaves from 5–7-week-old plants. Inset, relative conductance normalized to darkness data point just before blue-light radiation. Scale bar, 30 min. (d) Fusicoccin (FC; 10 µM) induces stomatal opening in ht1 mutants. Data presented are the means of 60 stomatal apertures ± SEM.

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Figure 3 ht1 mutants show abscisic acid responses. As an abscisic acid (ABA) hypersensitive control, era1-1 or era1-2 mutants were used. (a) Comparison of stomatal closing responses induced by ABA (10 µM) in wild-type (WT) and the ht1 mutants. Data presented are the means of 63 stomatal apertures ± SEM. Inset, ABA-induced stomatal closure normalized to stomatal apertures without added ABA in the four lines. (b) Wilting during drought stress in ht1 mutants. Plants are shown after 16 days without watering. WT and ht1 plants were grown under normal watering conditions in a growth chamber for 18 d. WT and ht1 mutant plants were selected at identical developmental stages and size, and were subjected to drought stress by completely terminating irrigation. (c) Effect of ABA on ht1 seed germination. Seeds were plated on a medium supplemented with the indicated concentrations of ABA, and incubated for 4 d at 22 ºC under constant light. The number of germinated seeds was expressed as the percentage of the total number of seeds plated. Data represent the mean ± SD of three independent experiments (60 seeds were plated per data point).




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movement signals is expected, as all signals regulate downstream ion transport and metabolic processes2,8.

Abscisic acid (ABA) response analyses of stomatal movements showed that the stomata of ht1 mutants respond to ABA, a phytohormone that triggers stomatal closure, to a similar extent as those of WT plants (Fig. 3a). Stomatal apertures of ht1-1 and ht1-2 plants were generally smaller than WT apertures at ambient CO2 concentration (Fig. 3a), as also shown in Fig. 1c. To further analyze whether the smaller stomatal apertures of ht1 are due to ABA hypersensitivity, we used era1 mutants as a control. Mutants in the ERA1 gene have been shown to cause ABA-hypersensitive stomatal closing and prolonged seed dormancy20,21 (Fig. 3). Normalization to non-ABA control stomatal apertures showed that there were no significant differences in stomatal closing responses to ABA in WT plants and the ht1 mutants (Fig. 3a, inset; Generalized Linear Models, family=quasi and link=log, P > 0.3, n = 504). On the contrary, the stomatal closing responses to ABA in era1-2 mutants were significantly different from those of WT plants or ht1 mutants (P < 0.02). This was supported by data showing that ht1 mutants exhibited severe wilting, similar to that seen in WT plants, whereas era1 mutants showed reduced wilting during drought stress (Fig. 3b). Furthermore, both ht1-1 and ht1-2 seeds germinated at similar rates as WT seeds on plates containing 0–1 µM ABA (Fig. 3c). We conclude that ht1 mutants are normal with respect to their ABA signalling responses. These results also indicate

that further stomatal closing can occur in the ht1 mutants in response to ABA (Fig. 3a) or darkness (Fig. 2a), although they have smaller stomatal apertures at ambient CO2 concentration and the ht1 mutants could not close their stomata further in response to high CO2 concentra-tion (Fig. 1). Thus, HT1 mediates a primarily CO2-dependent reaction and does not have a purely structural or a general physiological function that would impair all stomatal movement responses.

The HT1 locus was mapped to a location between two Cereon sin-gle nucleotide polymorphism markers, CER451254 and CER460537, on chromosome 1 (Fig. 4a). Sequencing of this 46-kb region revealed that a putative kinase gene, At1g62400, harboured a point mutation at nucleotide 796 in ht1-1 that resulted in the exchange of Arg 211 for Lys (Fig. 4b). In the ht1-2 mutant allele, a single base-pair (bp) substitution was detected at the donor splice site of the first intron at nucleotide 448 (Fig. 4b). This resulted in an in-frame 42-bp deletion at positions 406–447, which corresponded to the deletion of 14 amino-acid residues, Val 136 to Gln 149.

We transformed ht1 plants with a 4.4-kb genomic fragment, includ-ing At1g62400 (ht1:HT1). Six independent homozygous ht1:HT1 lines showed low leaf temperature under low CO2 concentration and a depend-ence of leaf temperature on CO2 concentration, which resembled the CO2 responses of WT plants, showing ht1 complementation (ht1-1:HT1, Fig. 1a, b). The ht1-2 mutant was also complemented by the genomic HT1 vector; the ht1-2:HT1 gave rise to similar results (data not shown). These

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Figure 4 HT1 is a protein kinase expressed in guard cells. (a) Map-based cloning of the HT1 gene. Genetic mapping of 1,074 chromosomes localized HT1 (white rectangle) to the 46-kb region between the markers CER451254 on BAC F24O1and CER460537 on BAC T3P18. (b) Structure of the HT1 gene. HT1 consists of three exons (open boxes); black boxes highlight the 5′ and 3′ untranslated regions, respectively. Conserved subdomains of the protein kinase family are indicated by roman numerals. ht1-1 and ht1-2 indicate the mutation sites. In subdomain II, there is an invariant lysine (K113). Arrows indicate regions used in reverse transcription polymerase

chain reaction (RT-PCR) (Fig. 4c). (c) RT-PCR analysis of HT1 mRNA. Expression in young leaf (y) from the 10-day-old wild-type (WT) seedlings; mature leaf (m) and roots (r) from 3-week-old WT plants; and stems (s) and flowers (f) from 4-week-old WT plants (left) is shown. Expression in aerial parts of 3-week-old WT, mutants (ht1-1, ht1-2) and transgenic (35SHT1(K113W)) plants (right). The EF1α gene was used as an internal standard for cDNA amounts. (d) HT1 expression pattern of HT1–promoter–GUS (pHT1::GUS) transgenic plants. Scale bars: in whole plant, 1 cm; leaf tissue, 200 µm; guard cell, 20 µm.




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data suggest that At1g62400 corresponds to HT1. A similar result was obtained in plants of ht1-2:35SHT1 lines, in which the HT1 cDNA was expressed under the control of the CaMV 35S promoter (ht1-2:35SHT1, Fig. 1a, b; ht1-1:35SHT1 showed similar results, data not shown). A slight difference in leaf temperature between WT and complemented mutant plants can be observed (Fig. 1a, b), which is likely to occur due to differ-ences in the expression levels of HT1 protein; nevertheless, the comple-mented lines show a clear recovery of CO2 responsiveness.

Using reverse transcription polymerase chain reaction (RT-PCR) analy-sis, HT1 mRNA was detected in leaves, stems, roots and flowers (Fig. 4c, left). HT1 mRNA was generally of low abundance and was usually not detected in northern blots of these samples. Characterization of trans-genic plants harbouring a transcriptional fusion of the β-glucuronidase (GUS) reporter gene and the HT1 promoter (pHT1::GUS) further indi-cated that HT1 expression occurred in the whole plant (Fig. 4d, left). In leaves, the expression of HT1 was mostly confined to guard cells (Fig. 4d,

right). In agreement with this observation, the expression of HT1 was highest in mature leaves, which have more abundant stomata (Fig. 4c, left). Furthermore, microarray analyses show that the HT1 gene (At1g62400) is preferentially expressed in guard cells compared with mesophyll cells22.

RT-PCR experiments showed that the PCR product (primers, see Fig. 4b) in ht1-2 was smaller than that in the WT plants, probably due to the 42-bp deletion (Fig. 4c, right). Little difference in HT1 mRNA abundance between WT, ht1-1 and ht1-2 plants was observed in mature leaves (Fig. 4c, right). CO2 treatment also had no detectable effect on the level of HT1 mRNA (data not shown).

The HT1 cDNA was cloned by RT-PCR using total RNA prepared from 3-week-old plants. The first in-frame ATG codon of the transcript initi-ated an open reading frame encoding a 390-amino-acid protein with a predicted relative molecular mass of 44,000 (Fig. 4b). The putative HT1 protein has all of the conserved features of the catalytic domains of protein kinases, except for subdomain I23 (Fig. 4b). To determine whether the HT1 protein and its mutants possess phosphorylation activity, we expressed each one as a His-tagged construct in Escherichia coli and performed in vitro kinase assays. The recombinant HT1 protein (His–HT1) exhibited autophosphorylation (Fig. 5a, lanes 2 and 8) and casein phosphorylation (Fig. 5a, lanes 3 and 9) when incubated with 32P-γ-ATP. Site-directed mutagenesis generated an ht1-1 equivalent mutant construct (His–HT1(R211K)), which converted the conserved active-site residue Arg 211 into Lys. The His–HT1(R211K) caused reduction in the phosphorylation activity (Fig. 5a, lanes 4 and 10). No significant levels of kinase activity were detected for the analogous His-construct of ht1-2, which contains a 14-amino-acid deletion corresponding to a part of the kinase subdomains III and IV (His–HT1(∆136–149); Fig. 5a, lanes 5 and 11). Therefore, the kinase activities of the WT HT1 and its two mutants correlated closely with the CO2 response phenotypes that were observed in whole plants. The kinase activities of HT1 and HT1(R211K) were Ca2+ independent (Fig. 5a, lanes 2–4 and 8–10), and the active proteins phosphorylated not only casein but also myelin basic protein (data not shown).

Lys 113 of HT1 corresponds to a highly conserved residue in sub-domain II (Fig. 4b) that is required for activity in most protein kinases23. To verify that the phosphorylation signal was due to HT1 phosphoryla-tion, Lys 113 was changed to Trp by means of site-directed mutagenesis24. The mutant protein (HT1(K113W)) exhibited no kinase activity (Fig. 5a, lanes 6 and 12), resembling denatured HT1 (Fig. 5a, lanes 1 and 7). To further address an in vivo role of HT1, we adopted a dominant-inhibi-tory strategy using the kinase-negative mutant of HT1 (HT1(K113W) gene) as, in the case of kinase-negative mutants24–26, we anticipated that overexpression of the mutant HT1 might interfere with the proper asso-ciation of endogeneous HT1 with its targets. This could potentially result in inhibition of the HT1-mediated signalling pathway. The experiments were carried out using a transgenic line that overexpresses this inactive HT1 isoform, controlled by the CaMV35S promoter in a WT background (WT:35SHT1(K113W), Fig. 5b, c). Transgenic lines showed higher leaf temperatures at low CO2 concentration, and the sensitivity to CO2 was repressed (Fig. 5b, c; for comparisons with WT, ht1-1 and ht1-2 mutant responses, see Fig. 1a, b). These CO2 responses were similar to those observed in ht1-2 plants (Fig. 1a, b). The kinase-negative HT1 protein, which had the same amino-acid sequences as the WT HT1 protein except for one amino acid, could not complement the ht1-2 mutants (Fig. 5b, c; ht1-2:35SHT1(K113W)). Together with data showing that the CO2 sensitivities of WT, mutants or transgenic plants were closely linked to

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Figure 5 HT1 kinase activity correlates with CO2 sensitivity. (a) Phosphorylation activity of HT1 and the indicated mutant proteins in vitro (see text). Assays were performed in the presence (Ca2+; right 6 lanes) or absence (EGTA; left 6 lanes) of CaCl2. Closed arrowhead indicates signals from autoradiograms of autophosphorylated proteins. Open arrowheads indicate phosphorylation of casein. –c, minus casein. K, relative molecular mass in thousands. (b) Thermal images of transgenic plants overexpressing a dominant-negative HT1 (HT1(K113W)) isoform in the wild-type (WT) background (WT: 35SHT1(K113W)) or ht1-2 background (ht1-2:35SHT1(K113W)). (c) Leaf temperatures calculated from the quantification of infrared images (means ± SD of measurements on ~5,000 square pixels; n = 6 plants per condition).




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their respective kinase activities, these results demonstrate that HT1 kinase activity is important for the CO2 response in plants rather than the structure of the HT1 protein. HT1 is the first functionally and molecu-larly isolated gene and protein that strongly modulates a CO2 signalling pathway that mediates stomatal movements. Leaf temperature, stomatal response and gas-exchange analyses of the ht1 mutants show that ht1 kinase mutant stomata show less stomatal opening at low CO2 concentra-tion and a more closed state prior to exposures to high CO2 concentration (Fig. 1). These data suggest that the ht1 mutation causes a CO2 hypersensi-tive response, thereby removing a major negative regulation mechanism of CO2 signalling and allowing progression of stomatal closing even in the absence of applied CO2. Similarly, the ethylene signal-transduction pathway has been shown to include strong negative regulators, for which loss of function causes constitutive ethylene responses27,28.

Much progress has been made in our understanding of the molecular mechanisms that underlie stomatal responses to stimuli, such as light and ABA2–4,29. However, how CO2 interacts with these factors in the process of stomatal regulation has remained a matter of debate2,9,13,30. Compared with ABA responses (Fig. 3), white-light and dark responses (Fig. 2a, b) were reduced in ht1 mutants; however, blue light and FC stimulation (Fig. 2c, d) continued to cause stomatal opening even in ht1-2 mutants, which were completely impaired in low CO2-induced stomatal opening (Fig. 1). Different stimuli in guard cells regulate the same targets2,3,29 (ion channels, H+-ATPase, metabolism, etc.) and these pathways form a com-plex signalling network. Direct biochemical evidence for the function of the HT1 protein kinase in the CO2 response has not yet been obtained; however, it is interesting that our findings show an impairment of the ht1 mutants in CO2 responses, but a much larger degree of responsiveness to other physiological stimuli. The identification of the HT1 protein kinase as a key molecular regulator of stomatal movements will help to unravel CO2 signal transduction and its interplay with the molecular network of light and ABA signalling in guard cells.

METHODSPlant material. All Arabidopsis lines used in this study were derived from the Columbia (Col-0) background. EMS-mutagenized Col M2 seeds were purchased from Lehle Seeds (Round Rock, TX, USA).

Mutant screen. M2 seeds were grown on half-strength MS medium supplemented with 1% (w/v) sucrose and 0.5% gellan gum at pH 5.7 for 18 d in a growth chamber (22 °C, 70% relative humidity, with constant light of 230 μmol m–2 s–1). CO2 concentra-tions were adjusted in the range of 350–450 ppm, unless otherwise noted. The plants were then transplanted into pots with vermiculite moistened with nutrients. After 3 d in the growth chamber, the pots were incubated under constant light (43 μmol m–2 s–1) in a growth cabinet (23 °C, 43% relative humidity) that was equipped with an auto-matic CO2 control unit (FR-SP, Koito Co., Japan). The plants were subjected to low CO2 conditions (100 ppm) for 1 h before thermal images were obtained using a ther-mography apparatus (TVS-8000MkII, Nippon Avionics, Co. Ltd, Japan) containing an infrared camera equipped with an 18.2° (H) × 13.6° (V) lens. The camera was equipped with a cooled 160 (H) x 120 (V) InSb array detector that is responsive to short infrared waves (3.6–4.6 µm band). The specified temperature resolution was 0.025 °C at room temperature. Leaf emissivity was set to 1 as an absolute measure-ment of leaf temperature was not required. The camera was mounted vertically at approximately 1 m above the leaf canopy for observations and was connected to a colour monitor to facilitate observation of individual plants. Images were fed into a Windows-based computer and were analyzed using beta release 4 of the pub-lic domain image-analysis program, Scion Image (http://www.scioncorp.com). Individual putative mutant plants with a higher leaf temperature than the WT plants were selected using an infrared image. These candidate mutants were grown to matu-rity and their phenotypes were re-tested in the next (M3) generation.

Transgenic plants. Transgenic Arabidopsis plants were generated by transforma-tion that was mediated by Agrobacterium tumefaciens. For functional complemen-tation of the ht1 mutants, the HT1 genomic region (nucleotides 54586–58950 of BAC F24O1) containing At1g62400 was amplified by PCR from genomic DNA using the oligonucleotides 5′-CTTCTCTAAGCTTTCGATGCAAACCA-3′ and 5′-GATGTATTGCAAGAGCTGATCAATTGGGTCATGAGACGAC-3′ and was then inserted into the pGEM-T Easy Vector (Promega, Madison, WI). A SalI-MunI fragment, including the HT1 genomic sequences, was cloned into the SalI-EcoRI site of the T-DNA vector pBI101. The HT1 cDNA was obtained using the SMART RACE PCR kit (Clontech, CA) according to the manufac-turer’s instructions. The full-length cDNA was then amplified using Pfu DNA polymerase (Stratagene, La Jolla, CA) with the oligonucleotides 5′- TACCCGGGATTTGTTTCCTTCTCTGTTTCTGC-3′ and 5′-TAGAGCTCTCAATAAGTATCATTATATATCATAC-3′; the amplified DNA was inserted into the SmaI-SacI site of the T-DNA vector pBI121. A 35SHT1(K113W) construct was created by introduction of the HT1(K113W) fragment, which was produced by site-directed mutagenesis and PCR as described previously24, into PshAI-BamHI of the HT1 cDNA followed by cloning into pBI121.

Gas exchange. Recordings of stomatal conductance were made using a Li-Cor 6400 Portable Photosynthesis system (LI-COR Inc., NE). Leaves were kept at a constant humidity of 32 ± 2% relative humidity, a constant temperature of 26.5 ºC and a constant light level of 75 µmol m–2 s–1 or a constant CO2 concentration of 365 ppm, unless otherwise noted. For analyses of blue-light responses, plants were kept at a constant temperature of 25 ºC, and blue-light stimulation was obtained by using light-emitting diodes with a maximum wavelength of 470 nm and a ± 10-nm half-bandwidth (Li-Cor 6400 Leaf Chamber Fluorometer, LI-COR Inc.).

Stomatal density measurements and aperture response analyses. Three-week-old plants that had been grown at ambient CO2 concentration were used for sto-matal density measurements and for aperture measurements in response to CO2. Abaxial epidermal peels of the plants were taken from the sixth or seventh leaf and were used for measurements of stomatal number and aperture determina-tion immediately after incubation at the indicated CO2 concentration. Eighteen random parts (each part consisted of 0.12 mm2) from four leaves per genotype were counted for determination of stomatal density. Stomatal aperture meas-urements in response to ABA or FC were conducted as described previously31, with minor modifications. Leaves from 4- to 5-week-old plants were floated in solutions containing 30 mM KCl, 5 mM Mes-KOH, pH 6.15 and 1 mM CaCl2, and were incubated in the growth chamber. ABA or FC from a stock solution in dimethyl sulfoxide (DMSO) was added to the solution after 2 h of illumination and stomatal apertures were measured 2 h later. Stomatal apertures were ana-lyzed in epidermal peels with a digital camera attached to a microscope (BH2, Olympus, Tokyo, Japan).

RT-PCR analysis. Total RNA extraction was performed, and single-stranded cDNA synthesized from total RNA was used as RT-PCR templates, according to the method described by Sugimoto et al.32. We used 1 µl of the RT reaction as a template in 10 µl PCR reactions. PCR was run for 27 cycles of 30 s at 94 °C, 30 s at 55 °C and 1 min at 72 °C. The RT-PCR primers for HT1 and its mutants were 5′-GGAATCTTGGTCGATGATCC-3′ and 5′-CAATGGTGGTCTTTCGTTCTT-3′. The amplified DNA fragments covered 161–921 bp of HT1. As an internal standard for cDNA, a 700-bp fragment of the EF1α cDNA was amplified, as described previously33.

Accession number. The DDBJ accession number for HT1 sequences is AB221045.

Analysis of GUS activity. A pHT1::GUS construct was obtained by ampli-fying 2 kb of the HT1 promoter region from genomic DNA using the oligonucleotides 5′-CTTCTCTAAGCTTTCGATGCAAACCA-3′ and 5′-CCATATGTCTGGTTTATGTTTCA-3′; the product was then inserted into the pGEM-T Easy Vector. A HindIII-PmeI fragment, including the HT1 promoter sequences, was cloned into the HindIII and SmaI sites of pBI101. GUS activity was assayed on 2-week-old seedlings grown on MS plates after overnight incuba-tion with 5-bromo-4-chloro-3indolyl-d-glucuronide as a substrate. All sequences amplified by PCR were confirmed by sequencing.




L E T T E R S

Preparation of recombinant proteins. An NdeI site was introduced in front of the ATG start codon of HT1 and ht1 mutants by PCR using each cDNA as a template. The constructs were then ligated in-frame into the pET-28a (+) vector (Novagen, Madison, WI), and were confirmed by DNA sequencing. BL21(DE3) cells transformed with pET-28a (+) constructs were induced with 1 mM IPTG for 16 h at 25 °C. His-tagged proteins were purified on nickel columns (Amersham Biosciences, Uppsula, Sweden). Purified His-tagged proteins were recognized specifically by anti-His-probe antibodies (Toyobo, Osaka, Japan) using an immu-noblot analysis (data not shown).

In vitro phosphorylation assay. An autophosphorylation assay was performed by incubating purified recombinant proteins (1 µg) in reaction buffer (25 mM Tris, pH 7.5, 10 mM MgCl2 and 1 mM CaCl2 or EGTA) in the presence of 0.6 µCi

32P-γ-ATP at 30 °C for 15 min. The reaction was stopped by the addition of SDS loading buffer, and kinase activities were detected by autoradiography after proteins had been resolved on a 12% SDS-polyacrylamide gel. Phosphorylation activities of HT1 and its mutants were determined in 10 µl of the kinase reaction buffer using 0.15 µg casein as a substrate under the same conditions.

Note: Supplementary Information is available on the Nature Cell Biology website.

ACKNOWLEDGEMENTSWe thank Y. Machida and K. Harada, K.M. Kawano, E. Kasuya and all of the members of our laboratories for technical assistance and discussion. We also thank the Arabidopsis Biological Resource Center and Cereon Genomics for access to polymorphism information. This research was supported by CREST, JST and the Japan Society of the Promotion of Science (17370019) grants (K.I.), and by National Science Foundation (MCB0417118) and the National Institutes of Health (R01GM060396) grants (J.I.S.). M.I. is a Formas post-doctoral fellow.

COMPETING INTERESTS STATEMENTThe authors declare that they have no competing financial interests.

Published online at http://www.nature.com/naturecellbiology/Reprints and permissions information is available online at http://npg.nature.com/reprintsandpermissions/

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Supplementary Information, Figure 1

Figure S1 CO2 responses of leaf surface temperatures measured by a thermistor. The

leaf temperature at the abaxial surface was monitored using a micro-thermistor probe

(P2L-64, Technol Seven, Japan) held in place with a fly trapping paste (Nisso shoji, Co.,

LTD. Japan). The sensitivity of the thermistor was 0.01 °C, and measurements were

taken at 1 min intervals. Air temperature (a. t.) in close proximity to the plants was

measured concurrently using an additional thermistor probe.

© 2006 Nature Publishing Group

NATURE CELL BIOLOGY ADVANCE ONLINE PUBLICATION 1

E R R AT U M

In the advance online publication of Hashimoto et al. (Nature Cell Biol. 2006; DOI: 10.1038/ncb1387), the following sentence was omitted from the acknowledgements:

M.I. is a Formas postdoctoral fellow.

This error has been corrected online.

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