Ozone-triggered rapid stomatal response involves theproduction of reactive oxygen species, and is controlledby SLAC1 and OST1

Triin Vahisalu1,2, Irina Puzorjova2, Mikael Brosche1, Ervin Valk2, Martin Lepiku2, Heino Moldau2, Priit Pechter2,

Yuh-Shuh Wang2, Ove Lindgren2, Jarkko Salojarvi1, Mart Loog2, Jaakko Kangasjarvi1 and Hannes Kollist2,*

1Division of Plant Biology, Department of Biosciences, University of Helsinki, PO Box 65 (Viikinkaari 1), FI-00014 Helsinki,

Finland, and2Institute of Technology, University of Tartu, Nooruse 1, 50411 Tartu, Estonia

Received 26 November 2009; revised 16 January 2010; accepted 20 January 2010; published online 12 March 2010.*For correspondence (fax +37 2737 4900; e-mail hannes.kollist@ut.ee).

SUMMARY

The air pollutant ozone can be used as a tool to unravel in planta processes induced by reactive oxygen species

(ROS). Here, we have utilized ozone to study ROS-dependent stomatal signaling. We show that the ozone-

triggered rapid transient decrease (RTD) in stomatal conductance coincided with a burst of ROS in guard cells.

RTD was present in 11 different Arabidopsis ecotypes, suggesting that it is a genetically robust response. To

study which signaling components or ion channels were involved in RTD, we tested 44 mutants deficient in

various aspects of stomatal function. This revealed that the SLAC1 protein, essential for guard cell plasma

membrane S-type anion channel function, and the protein kinase OST1 were required for the ROS-induced fast

stomatal closure. We showed a physical interaction between OST1 and SLAC1, and provide evidence that

SLAC1 is phosphorylated by OST1. Phosphoproteomic experiments indicated that OST1 phosphorylated

multiple amino acids in the N terminus of SLAC1. Using TILLING we identified three new slac1 alleles where

predicted phosphosites were mutated. The lack of RTD in two of them, slac1-7 (S120F) and slac1-8 (S146F),

suggested that these serine residues were important for the activation of SLAC1. Mass-spectrometry analysis

combined with site-directed mutagenesis and phosphorylation assays, however, showed that only S120 was a

specific phosphorylation site for OST1. The absence of the RTD in the dominant-negative mutants abi1-1 and

abi2-1 also suggested a regulatory role for the protein phosphatases ABI1 and ABI2 in the ROS-induced

activation of the S-type anion channel.

Keywords: stomata, signaling, SLAC1, OST1, ozone, reactive oxygen species.

INTRODUCTION

Stomata, small pores on the aerial parts of plants, control

CO2 influx for photosynthesis and water vapor loss. They

also restrict the entry of ozone (O3) – a major air pollutant

with an increasingly negative impact on crop yields, global

carbon fixation (Hopkin, 2007) and climate change (Sitch

et al., 2007). Ozone degrades immediately to reactive

oxygen species (ROS) in the apoplastic space of plant cells,

and has therefore been used as a tool to study the signaling

role of the apoplastic ROS (Kangasjarvi et al., 2005;

Wrzaczek et al., 2009). ROS are involved in the regulation of

abscisic acid (ABA)- (Lee et al., 1999; Pei et al., 2000),

ethylene- (Desikan et al., 2006), methyl jasmonate-

(Munemasa et al., 2007) and salicylic acid-mediated (Mori

et al., 2001) stomatal signaling. The rapid induction of ROS

during CO2-induced stomatal closure has also been shown

(Kolla et al., 2007). Collectively, ROS are central intermediate

signaling components in plant guard cells, and it is likely that

ozone is a useful tool for the study of ROS-dependent

stomatal signaling.

Stomatal guard cells are among the most studied and best

understood plant signaling systems, yet there remain con-

siderable gaps in the understanding of the signaling that

leads to stomatal movements in response to different

stimuli. For example, the importance of guard cell anion

channels as central regulators of stomatal closure was

demonstrated 20 years ago (Keller et al., 1989; Schroeder

442 ª 2010 The AuthorsJournal compilation ª 2010 Blackwell Publishing Ltd

The Plant Journal (2010) 62, 442–453 doi: 10.1111/j.1365-313X.2010.04159.x

and Hagiwara, 1989), but the protein essential for guard cell

anion channel functioning, SLAC1, was identified only very

recently (Negi et al., 2008; Vahisalu et al., 2008). SLAC1 is

essential for stomatal closure in response to ABA, CO2, O3,

light–dark transitions and humidity change, and by second-

ary messengers Ca2+, H2O2 and NO. However, the signaling

cascades upstream of SLAC1, which require the capturing of

very early and probably transient responses, are as yet

unexplored.

Most experiments addressing the molecular details of

guard cell signaling have been performed with epidermal

peels or isolated guard cells. The preparation procedure(s)

are likely to introduce unwanted effects, including an

elevated production of ROS as a result of damage. Thus,

using intact plants and minimum handling of the plant

would help to define the function of ROS in stomatal

regulation more clearly. The simple application of ozone to

intact plants offers the possibility to control the concentra-

tion and duration of the exposure precisely. We have

constructed a gas-exchange system where eight soil-grown

Arabidopsis plants can be enclosed in individual flow-

through exposure vessels, in a non-invasive manner (Kollist

et al., 2007). Using this system we have shown that as little

as 150 nl l)1 of O3 triggers a rapid transient decrease (RTD)

in stomatal conductance (Kollist et al., 2007). The decrease

was induced within a few minutes of O3 exposure, but the

stomata reopened again despite the continuous presence of

ozone. The recovery suggests that the closure was not a

result of physical ozone damage, but instead reflects the

biological action of ROS formed from ozone breakdown in

the apoplast, transduced through a signaling cascade.

The lack of ozone-triggered RTD in the ABA-insensitive

mutant abi2-1 (Kollist et al., 2007), carrying a dominant-

negative mutation in the type-2C protein phosphatase ABI2,

suggests a role for protein phosphorylation in O3/ROS-

induced stomatal signaling. Murata et al. (2001) have also

shown that H2O2-induced stomatal closure was impaired in

the abi2-1 mutant. H2O2-induced stomatal closure was also

disrupted in the recessive ABA-insensitive mutant gca2

(Pei et al., 2000). On the contrary, in abi1-1, another ABA-

insensitive dominant-negative mutant of the protein phos-

phatase ABI1 (Murata et al., 2001), and in mutants of the

protein kinase OPEN STOMATA 1 (OST1) (also referred to as

SRK2E and Snf1-related protein kinase 2.6, SnRK2.6), a

positive regulator of ABA-induced stomatal closure (Mustilli

et al., 2002), H2O2-induced stomatal closure was not dis-

rupted, suggesting a role for these proteins between ABA

perception and ROS production.

We have shown that activation of S-type anion channels is

required for ROS-induced stomatal signaling, as ozone-

triggered RTD was absent in the S-type anion channel

mutant slac1 (Vahisalu et al., 2008). However, other struc-

tural and signaling components involved in relaying the

ROS signal from apoplast to stomatal movements have not

been identified. During recent years, the molecular identities

of many other guard cell transport proteins have been

established (for a review see Pandey et al., 2007; Ward et al.,

2008). Testing the characteristics of ozone-triggered RTD in

plant lines carrying mutated versions of proteins involved

in stomatal regulation could help to understand their role in

ROS-dependent processes.

Here we have explored the ozone-triggered RTD further.

We show the time and concentration dependence of the

process, and provide evidence that stomatal closure coin-

cides with the elevated burst of ROS in guard cells. This

suggests that RTD is induced by the ROS triggered by the

application of ozone. By analyzing RTD in several mutants

carrying mutations in proteins shown to be involved in

stomatal regulation, we show that OST1, ABI1, ABI2 and

SLAC1 are regulators of the ROS-induced rapid stomatal

closure. We demonstrate physical interaction between

SLAC1 and OST1, and provide evidence that SLAC1 is

phosphorylated by OST1.

RESULTS

The ozone-triggered RTD is dependent on exposure time

and ozone concentration

Applying 250 nl l)1 of ozone induced a 40% decrease in

stomatal conductance in wild-type (WT) Arabidopsis Col-0

plants within 5–10 min of exposure, followed by reopening

to the pre-exposure level within the next 40 min (Figure 1a).

The commonly observed sustained ozone-induced decrease

in stomatal conductance (Ahlfors et al., 2004) was visible

90 min after ozone onset. The same stomatal behavior was

seen in the 10 other ecotypes tested (Table S1). To further

elucidate the relationship between the duration of ozone

exposure and the decrease in stomatal conductance, Ara-

bidopsis Col-0 plants were treated with 250 nl l)1 of ozone

for 30, 70, 180, 360 and 720 s. Already a 30-s pulse of

250 nl l)1 ozone caused a clearly detectable decrease in

stomatal conductance (Figure 1b). The decrease reached a

maximum with 180 s of ozone, and longer exposures did not

decrease the conductance any further. To address the effect

of ozone concentration, we applied 3-min ozone pulses with

concentrations ranging from 50 to 600 nl l)1 (Figure 1c). The

response can be separated into three segments: ‘no

response’, ‘response’ and ‘saturation’. In the ‘no response’

segment, essentially no decrease in stomatal conductance

was observed. The threshold for the ‘response’ segment was

80 nl l)1 of ozone (95% confidence interval, shown by the

dashed lines). After the threshold, the decrease in stomatal

conductance increased approximately linearly, by 0.15% per

additional nl l)1 of ozone, with 95% confidence intervals of

�0.04% per nl l)1. The ‘saturated’ segment, where the

decrease in stomatal conductance reached its maximum,

was obtained with concentrations higher than 434 nl l)1 of

ozone.

Ozone-triggered stomatal responses 443

ª 2010 The AuthorsJournal compilation ª 2010 Blackwell Publishing Ltd, The Plant Journal, (2010), 62, 442–453

To test the responsiveness of guard cells to ozone

during the recovery period (20–60 min after the first ozone

pulse), we applied four additional 3-min (250 nl l)1) pulses

of ozone with 9-min intervals after the initial pulse

(Figure 2a). These successive pulses had no effect on

R 2 = 0.81

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Figure 1. Ozone-triggered rapid transient decrease (RTD) in stomatal con-

ductance is dependent on the time and concentration of ozone applied.

(a) Time course of stomatal conductance (n = 3, �SEM) of Col-0 plants after

the onset of 250 nl l)1 ozone exposure, indicated by the gray bar.

(b) Ozone-triggered decrease in stomatal conductance after the application of

250 nl l)1 ozone for 30, 70, 180, 360 and 720 s (n = 3, �SEM).

(c) Decrease in stomatal conductance in response to increasing ozone

concentrations. Each point represents an independent experiment. The

response is divided into three segments, separated by two vertical solid

lines: ‘no response’, ‘response’ and ‘saturation’. The solid curve shows a

bootstrap aggregate fit of data using 1000 bootstrap samples, each fitted with

a logistic function. Dashed curves mark the 95% confidence intervals. The

segment borders are identified by computing the maxima of the third

derivative (i.e. the maximum of the change of acceleration).

Sto

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Figure 2. Guard cells are temporarily desensitized to ozone by a primary

3-min ozone pulse.

Time courses of stomatal conductance of Col-0 plants after the onset of

250 nl l)1 ozone shared between: (a) five 3-min pulses with 9-min intervals,

(b) two 3-min pulses with a 90-min interval, (c) ten 3-min pulses with 12-min

intervals. The experiments were repeated between three and five times, with

similar results.

444 Triin Vahisalu et al.

ª 2010 The AuthorsJournal compilation ª 2010 Blackwell Publishing Ltd, The Plant Journal, (2010), 62, 442–453

stomatal conductance, indicating that the induction of the

RTD was blocked during the recovery period. The respon-

siveness to ozone reappeared when a second pulse of

ozone was applied at 100 min, when stomatal conductance

had fully recovered to the pre-exposure value (Figure 2b).

However, applying nine additional 3-min ozone pulses

with 12-min intervals throughout the recovery phase did

not induce RTD (Figure 2c), suggesting the importance of a

resting period for the guard cells to sense and respond to

ozone again.

Ozone-triggered RTD involves a rapid burst of ROS

To address whether ozone induces an intrinsic ROS burst in

guard cells, we analyzed early ROS production using fluo-

rescence dye and confocal microscopy. The duration of

ozone application was chosen according to the time course

of stomatal conductance upon ozone exposure (Figure 1a):

when RTD was induced (3 min); when stomata had reopened

to the pre-exposure level (45 min); and when the sustained

decrease in stomatal conductance was initiated (90 min). In

addition to the Col-0 WT, we used various mutants: slac1-1,

where ozone-triggered RTD is absent (Vahisalu et al., 2008);

ost1-3 (also referred to as srk2e; Yoshida et al., 2002); and

atrbohD and atrbohD/F, with mutations in NADPH oxidase

catalytic subunit genes, previously shown to regulate ROS

production in guard cells (Torres et al., 1998; Kwak et al.,

2003). At the times indicated, plants were removed from the

O3 treatment, epidermal peels were isolated, stained with

100 lM H2DCFDA and visualized by confocal microscopy

8 min after the removal of the plants from the treatment.

Images were processed and ROS production in guard cells

was quantified as fluorescence brightness.

A low background level of ROS was visible in the

untreated control plants (Figure 3a, 0 min). In all plants

studied, ozone exposure caused a bi-phasic ROS accumula-

tion: elevated ROS signal was detected after 3 min of ozone

exposure, after 45 min ROS levels were close to those of the

untreated plants, and after 90 min of ozone exposure a

second increase in ROS accumulation was evident. In all the

mutants studied the first burst was lower than in the Col-0

plants. In the ost1-3 and the atrbohD mutants, the second

increase in ROS accumulation (90 min) was higher than the

first increase (Figure 3a; Tukey’s honestly significant differ-

ence test results for the data presented in Figure 3a are

shown in Table S2). Untreated atrbohD and atrbohD/F

mutants showed lower ROS levels than Col-0 WT, and in

atrbohD/F, the second ROS peak (90 min) was lower than in

all the other lines.

To localize the subcellular sites of ROS production during

ozone-triggered RTD, we analyzed the confocal microscopy

images in more detail (Figure 3b). At the earliest time point,

3 min from the beginning of ozone exposure, ROS accumu-

lation was visible in the chloroplasts, from where ROS

seemed to diffuse to the other parts of the cell. A more

diffused, transient cytoplasmic ROS accumulation was

visible at 12 min, which disappeared by 45 min, when ROS

accumulation was again at low levels, and only found in

chloroplasts. After 90 min of ozone exposure, a second peak

of ROS accumulation, again spatially co-localized with

chloroplasts, was visible.

0Col-0 atrbohD atrbohD/Fost1-3slac1-1

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Figure 3. Ozone induces an intrinsic burst of reactive oxygen species (ROS) in

Arabidopsis guard cells.

(a) Col-0, slac1-1, ost1-3, atrbohD and atrbohD/F Arabidopsis plants were

exposed to 350 nl l)1 of ozone for 0, 3, 45, 90 min, and epidermal peels were

isolated and stained with 100 lM H2DCFDA, and visualized by confocal

microscopy. ROS production in guard cells, indicated in brightness units, was

quantified by IMAGEQUANT software (n = 12, �SEM).

(b) Col-0 plants were exposed to to 350 nl l)1 of ozone for 0, 3, 12, 45 and

90 min, epidermal peels were isolated and treated as described in (a).

Ozone-triggered stomatal responses 445

ª 2010 The AuthorsJournal compilation ª 2010 Blackwell Publishing Ltd, The Plant Journal, (2010), 62, 442–453

Rapid ozone-triggered RTD is a specific process

A 3-min pulse of ozone triggered RTD in all 11 Arabidopsis

ecotypes tested (Table S1), suggesting that RTD is a genet-

ically robust response. Many mutants deficient in ABA sig-

naling and ROS production in guard cells, in response to

different stimuli, have been identified (Li et al., 2006). We

tested several of them to elucidate the role of these proteins

in the apoplastic ROS-induced RTD (Table S1).

The a and b subunits of the heterotrimeric G protein have

been shown to be necessary for the ozone-induced oxidative

burst in guard cells (Joo et al., 2005). The a subunit is also

involved in regulating stomatal closure in response to ABA

(Wang et al., 2001). Stomata of the Ga mutant gpa1-4, the Gbmutant agb1-2 and the double mutant agb1/gpa1 responded

to ozone like the WT (Table S1), suggesting that heterotri-

meric G proteins were not involved in the signaling from

apoplastic ROS to the activation of anion fluxes under our

conditions.

The AtrbohD, AtrbohE and AtrbohF NADPH oxidase

subunits have a role in guard cell ABA signal transduction

(Kwak et al., 2003). However, atrbohD, atrbohE and atrbohF

single mutants, and all double mutant combinations,

responded to ozone pulse like the WT (Table S1), suggesting

that the O3-derived apoplastic ROS formation can function-

ally mimic the apoplastic ROS production by NADPH

oxidases in guard cells.

The plant stress hormones ethylene, jasmonic acid,

salicylic acid and ABA are important regulators of ozone

responses (Kangasjarvi et al., 2005), and have also been

shown to be regulators of guard cell signaling (Lee et al.,

1999; Pei et al., 2000; Mori et al., 2001; Desikan et al., 2006;

Munemasa et al., 2007). We tested mutants deficient in

biosynthesis and/or essential signal components for each of

these hormones – none of them were required for RTD

(Table S1). Exceptionally, some components of ABA signal-

ing, but not ABA biosynthesis, were required for RTD (see

below).

A mutant deficient in the protein kinase HT1, known to

control stomatal movements in response to CO2 (Hashim-

oto et al., 2006), responded to ozone like the WT (Table

S1). The mutant of the plasma membrane-localized ATP

binding cassette transporter AtMRP5, shown to have

impaired Ca2+ activation of guard cell anion channels

(Suh et al., 2007), had a normal response to ozone (Table

S1), as did the cpk3 and cpk6 (calcium-dependent protein

kinase 3 and 6) mutants required for the activation of

S-type anion currents by ABA and calcium (Mori et al.,

2006) (Table S1). The ABA-insensitive mutant gca2 (Pei

et al., 2000) also responded to ozone like the WT (Table

S1), indicating that the activity of GCA2 is not required for

the apoplastic ROS-induced stomatal closure. These results

suggest that the apoplastic ROS-induced stomatal move-

ments did not operate through the same set of regulatory

components through which CO2, and Ca2+-dependent

signaling act.

There were, however, mutants that did not display the

O3-triggered RTD. We have previously shown that the ABA-

insensitive protein phosphatase type-2C mutant abi2-1

completely lacked the ozone-triggered RTD (Kollist et al.,

2007). Here (Table S1), we show that, in addition to abi2-1,

the ozone-triggered RTD was also absent in abi1-1, ost1-1

and ost1-3, and in two K+-channel mutants the kinetics of

RTD was altered.

Ozone-triggered RTD is modulated by K+ channels, and

requires functional SLAC1, OST1, ABI1 and ABI2

Stomatal movements are facilitated by the activity of ion

channels and transporters in the plasma membrane and

vacuolar membrane of guard cells (Pandey et al., 2007). In

addition to the S-type ion channel mutant slac1-1, where the

ozone-triggered RTD was absent (Figure 4a), the patterns of

RTD were also different in two potassium channel mutants:

gork-1, where the guard cell plasma membrane K+ outward

rectifying channel GORK activity is fully suppressed (Hosy

et al., 2003), and kincless, where the inward rectifying K+

current is abolished (Lebaudy et al., 2008) (Figure 4b). In the

Ws-2 WT, ozone triggered an RTD of 30% within 4 min of

exposure. The response was strongly delayed in gork-1,

where the RTD was only 5% within 4 min, and a maximal

decrease of 23% was only achieved 16 min after the begin-

ning of the exposure (Figure 4b). In kincless, ozone caused a

22% RTD within 4 min, but reopening was almost com-

pletely absent; after 40 min, conductance had fully recov-

ered in the WT, whereas in kincless no recovery was seen

(Figure 4b). These results suggest that, in addition to initial

anion currents (SLAC1), subsequent GORK-mediated K+ flux

is required for the rapid decrease in stomatal conductance.

During the recovery period, additional ozone pulses had no

effect (Figure 2a). This implies that the activity of the inward-

rectifying potassium channel was not directly affected by the

apoplastic ROS induced by O3.

In order to elucidate the role of protein phosphorylation in

the ozone-triggered RTD upstream of SLAC1, we analyzed

RTD in abi1-1, abi2-1 and ost1-1 in more detail (Figure 5).

Whereas a 3-min O3 pulse triggered RTD in both Col-0 and

Ler WTs, stomata of all three protein phosphorylation

mutants were insensitive to the O3 pulse, suggesting that

the protein kinase OST1 and the phosphatases ABI1/ABI2 are

required for the SLAC1-dependent RTD.

Protein interaction, genetic and phosphoproteomic analysis

suggest that SLAC1 is regulated by OST1

The absence of ozone-triggered RTD in ost1 mutants

immediately suggested a model where SLAC1 is regulated

by OST1. Therefore, we first addressed whether SLAC1 and

OST1 interact in a split-ubiquitin membrane yeast two-

hybrid system (Johnsson and Varshavsky, 1994). SLAC1 was

446 Triin Vahisalu et al.

ª 2010 The AuthorsJournal compilation ª 2010 Blackwell Publishing Ltd, The Plant Journal, (2010), 62, 442–453

fused to the C terminal of ubiquitin (Cub) and the LexA-VP16

transcription factor as bait, and the OST1 was fused to the

N terminal of ubiquitin with an I3G mutation (NubG) as prey

(Figure S1a). Growth on selective media, and strong acti-

vation of the lacZ reporter gene in yeast co-transformed with

the bait and prey (Figure S1b), indicated an interaction

between SLAC1 and OST1. To confirm this result in planta,

we used a bimolecular fluorescence complementation

(BiFC) assay. Co-infiltration of 35S:SLAC1-YFPC and

35S:OST1-YFPN in Nicotiana benthamina leaves yielded YFP

signals in the plasma membrane and nucleus, with the latter

possibly being a result of high expression levels from the

35S promoter (Figure 6a). No fluorescence signal was

observed when 35S:YFPC and 35S:YFPN, 35S:SLAC1-YFPC

and 35S:YFPN, or 35S:YFPC and 35S:OST1-YFPN were

co-infiltrated. The data supports the split-ubiquitin yeast

two-hybrid analysis, and provides evidence for a physical

interaction between SLAC1 and OST1.

The prediction programs NetPhos and Scansite (http://

www.cbs.dtu.dk/services/NetPhos and http://scansite.mit.

edu) suggested several putative phosphorylation sites in

the N-terminal tail of SLAC1. To analyze the importance of

these sites for the regulation of SLAC1 in vivo, we used

TILLING (Till et al., 2003), and identified mutants where three

of the predicted phosphorylation sites were mutated: S38F

(slac1-6), S120F (slac1-7) and S146F (slac1-8). Two of these

mutants, slac1-7 and slac1-8 were deficient in RTD, similar to

slac1-1, whereas slac1-6 responded to ozone like the WT

(Figure 6c).

Finally, we performed in vitro phosphoproteomic exper-

iments to examine whether SLAC1 is a substrate of OST1. As

shown earlier (Belin et al., 2006), recombinant 6xHis

N-terminal-tagged OST1 protein produced in Escherichia

coli displayed autophosphorylating activity, and effectively

phosphorylated a generic substrate, such as histone III

(Figure 6d). To address phosphorylation of SLAC1 by

OST1, we used a SLAC1 N-terminal fragment (SLAC1–186)

as a substrate. The phosphorylation of SLAC11–186 by OST1

was rapid and efficient (Figure 6d). Prediction programs and

experiments with slac1-7 and slac1-8 suggested the pres-

ence of several functionally important phosphorylation sites

in SLAC1. To address this, we separated the OST1 phos-

phorylated SLAC1–186 on SDS-PAGE gel, supplemented with

Mn2+-Phos-tag, which incorporates a phosphate-binding

0 10 20 30 40 500.3

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Time (min)

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Figure 4. Ozone-triggered rapid transient decrease (RTD) in stomatal con-

ductance is absent in slac1-1, and is altered in gork-1 and kincless.

Time courses of stomatal conductance (n = 3, �SEM) after the onset of a

3-min 250 nl l)1 ozone pulse, indicated by the gray bar. (a) Col-0 and slac1-1.

(b) Ws, kincless and gork-1.

0 10 20 30 40 500.3

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Time (min)

Figure 5. Ozone-triggered rapid transient decrease (RTD) in stomatal

conductance is absent in abi1-1, abi2-1 and ost1-1.

Time courses of stomatal conductance (n = 3, �SEM) in Ler, abi1-1, abi2-1

and ost1-1, after the onset of a 3-min 250 nl l)1 ozone pulse, indicated by the

gray bar.

Ozone-triggered stomatal responses 447

ª 2010 The AuthorsJournal compilation ª 2010 Blackwell Publishing Ltd, The Plant Journal, (2010), 62, 442–453

compound that retards the mobility of phosphorylated

proteins in proportion to their degree of phosphorylation

(Kinoshita et al., 2006). Multiple differentially migrating

bands in Phos-tag gels indicated that several amino acids

in SLAC1–186 were phosphorylated by OST1 (Figure 6e). To

address whether S120 and S146 of SLAC1 are targets of

OST1, we used SLAC1–186 fragments where S120 and S146

were mutated to alanine. Clearly, the different multiphosph-

orylation pattern of S120A, compared with the WT SLAC1–186,

indicates that S120 was phosphorylated by OST1 (Fig-

ure 6e). On the contrary, S146 was not phosphorylated by

OST1, as the SLAC11–186 fragment containing the mutation

35S:SLAC1-YFPC/35S:OST1-YFPN

YFP Chl Merge

(a)

N

C

Plasma membrane

S38Fslac1-6

S120Fslac1-7

S146Fslac1-8

(b)

(c)

(d)

(e)

(f)

Sto

mat

al c

ondu

ctan

ce

(rel

ativ

e un

its)

Time (min)

0.2

0.4

0.6

0.8

1.0

1.2

0 10 20 30 40 50

O3

slac1-1

slac1-7

Col-er

slac1-6

slac1-8

SLAC1–186

WTSLAC1–186

S120ASLAC1–186

S146A SLAC1–186

S120A/S146A

Time

S59S86

S120S113TKSTLSK

QVSLETGFSVLNRSequence of identified peptide Phosphprylation site

SFGGFESGGIINGGDGRK

QKSLLPSIIR

Time 2’ 8’4’ 2’ 8’4’ 2’ 8’4’

2’ 8’4’2’ 8’4’2’ 8’4’2’ 8’4’

OST1 OST1 1:250 OST1 1:250

H3

OST1autoph.

CBB

SLAC11-186

MERKQSNAHSTFADINEVEDEAEQELQQQENNNNKRFSGNRGPNRGKQRPFRGFSRQVS59LETGFSVLNRESRE

RDDKKSLPRSGRS86FGGFESGGIINGGDGRKTDFSMFR

TKS113TLSKQKS120LLPSIIRERDIENSLRTEDGETKDDSIN

ENVSAGRYFAALRGPELDEVKDNEDILLPKEEQWPFLL

SLAC1–186H3

Figure 6. Interaction, phosphoproteomic and mutant data suggest that SLAC1 is regulated by OST1 kinase.

(a) Bimolecular fluorescence complementation assays were performed with Nicotiana benthamiana leaves infiltrated with 35:SLAC1-YFPC and 35S:OST1-YFPN. YFP

signal (YFP), chlorophyll autofluorescence (Chl) and overlay of YFP and chlorophyll autofluorescence (Merge) are shown. A similar result was observed in �20

individual cells in three independent repeats.

(b) Schematic diagram of the SLAC1 protein, indicating the positions of the mutations in slac1-6, slac1-7 and slac1-8.

(c) Time courses of stomatal conductance (n = 5, �SEM) in Col-er and alleles of slac1 after the onset of a 3-min 250 nl l)1 ozone pulse (indicated by the gray bar).

(d) Recombinant protein kinase OST1 has autophosphorylation activity (left panel), phosphorylates SLAC11–186 (middle panel) and a generic substrate

histone III (right panel).

(e) Different SLAC1–186 fragments phosphorylated by OST1 and separated on SDS-PAGE gel supplemented with Mn2+-Phos-tag. The bands visible represent the

degree of phosphorylation by OST1.

(f) SLAC1–186 phosphorylated by OST1 was analyzed with nanoLC-MS/MS system. The upper table shows the sequence of identified peptides phosphorylated by

OST1. The lower table shows the protein sequence of SLAC11–186. Phosphorylated serines are indicated in red. The results were confirmed through several

replications.

448 Triin Vahisalu et al.

ª 2010 The AuthorsJournal compilation ª 2010 Blackwell Publishing Ltd, The Plant Journal, (2010), 62, 442–453

S146A, or the combined mutations of S120A and S146A,

revealed similar phosphorylation patterns as the WT SLAC1–186

and SLAC1–186 containing S120A, respectively. Additionally,

we analyzed the SLAC11–186 fragment phosphorylated by

OST1 with mass spectrometry. This indicated that Ser59,

Ser86, Ser113 and Ser120, but not Ser146, were phosphor-

ylated by OST1 (Figure 6f and Figure S2).

Taken together, these data indicate that regulation of

SLAC1 by OST1 may involve the phosphorylation of multi-

ple amino acids, and suggests that Ser120 of SLAC1 is one of

the functionally important phosphosites.

DISCUSSION

Reactive oxygen species have a demonstrated role in guard

cell signaling in response to various external and internal

factors (Pei et al., 2000; Kolla et al., 2007). Apoplastic ROS,

formed as a result of the activity of the NADPH oxidases

(Kwak et al., 2003), are essential components in guard cell

ABA signaling. Ozone is known to degrade to various ROS in

the apoplast, thus it is likely that the ozone-triggered RTD

addressed in this study is a result of the action of the apo-

plastic ROS. This ROS from ozone degradation would act in

a similar manner as the ROS produced by NADPH oxidase

activity in ABA- or methyl jasmonate-induced stomatal

closure (Kwak et al., 2003; Munemasa et al., 2007). Hence,

ozone can be used as a tool to simplify the very complex

regulatory network in guard cells, and enables the study of

the role of ROS alone. The fast kinetics of RTD (Figure 1b)

implies that either the ROS are perceived directly in the

apoplast, followed by a rapid signal transmission to guard

cells, or that the ROS formed in the apoplast translocate

(most likely after dismutation to H2O2, as O�2 is impermeable

through a biological membrane) to the inside of guard cells,

where they immediately elicit the response.

Ozone triggered a biphasic ROS accumulation in guard

cells, where chloroplasts were the major source for ROS

formation (Figure 3). Previously, it has been shown that

ozone-induced ROS production was initiated from guard cell

chloroplasts, followed by ROS production in guard cell

membranes, which required NADPH oxidases encoded by

the AtrbohD and AtrbohF genes (Joo et al., 2005). The time

point studied with atrbohD and atrbohF by Joo et al. (2005)

was 1 h. In our experiments, the first phase of O3-induced

ROS accumulation (detected after 3 min) was significantly

reduced, but not abolished, in atrbohD and atrbohD/F

mutants (Figure 3a). This suggests the presence of several

sources for initial ROS production, for example, cell wall

peroxidases, in addition to the NADPH oxidases. Recently, it

has been shown that OST1 can phosphorylate AtrbohF, and

possibly regulates its activity (Sirichandra et al., 2009). The

ost1-3 mutant had lower initial ROS production (Figure 3a),

suggesting that regulation of Atrboh-mediated ROS produc-

tion by OST1 could be functionally relevant during initial

ozone responses. Interestingly, the second ROS peak was

lower in the atrbohD/F double mutant (Figure 3a), which

suggests that the membrane-bound NADPH oxidases have

an influence on the second peak of ROS production. It is

noteworthy that the timing of the first and second peak of

ROS accumulation (Figure 3) coincided with the fast and

slow decrease in stomatal conductance triggered by ozone

(Figure 1a). The decline in ROS production to control levels

at 45 min, despite the plant being continuously exposed to

ozone, favors a model for enzymatic control of ROS

production. However, whether this temporal coincidence

also has a mechanistic grounding needs to be studied

further, as well as the role of chloroplastic ROS in guard cell

signaling.

After the perception of the apoplastic ROS, the signal is

rapidly transduced to the guard cell anion channel SLAC1, as

RTD was absent in the slac1 mutant. However, RTD is not a

result of the activity of the plasma membrane anion channel

only: in the K+ efflux channel mutant gork-1, the ozone

response was delayed, suggesting that after the activation of

the anion fluxes, K+ fluxes through GORK were also required

for the rapid decrease in stomatal conductance (Figure 4b).

Thus, apparently the signal from apoplastic ozone/ROS is

first passed to SLAC1, and the subsequent activation of

anion fluxes and membrane depolarization are required for

GORK activation. The lack of recovery in stomatal conduc-

tance in the kincless mutant indicates a central role for K+

inward-rectifying channels during the recovery period

(Figure 4b).

An intriguing feature of the ozone-triggered RTD is that

during the recovery period further ozone pulses had no

effect (Figure 2b). This suggests that the primary ozone

pulse changes the status of the voltage-dependent channel

assembly in a manner that temporarily desensitizes stomata

to further ozone pulses. Blocking and desensitization have

been shown to occur for several ion channels under different

treatments (Roelfsema and Prins, 1997; Raschke et al., 2003).

Additionally, blue light treatment rapidly induces Ca2+

transients (Baum et al., 1999), and a 30–120-min recovery

period is required to elicit the response again. Similarly,

both H2O2 (Price et al., 1994) and ozone (Clayton et al., 1999;

Evans et al., 2005) induced rapid transient increases in

cytosolic Ca2+, and a recovery period of a few hours was

required before the response could be elicited again. One of

the possible causes of desensitization may be channel

inactivation (Hedrich et al., 1990; Pei et al., 1998), or if Ca2+

is part of the RTD, it might need to be returned to its original

resting state concentration and/or localization.

Several different cellular regulatory and signaling pro-

cesses in stomatal guard cells appear to converge at the

activation of anion fluxes, where SLAC1 is a central compo-

nent. ROS also play a role in several of these regulatory

cascades. The absence or presence of RTD in various guard

cell signaling and ion channel mutants (Table S1) allowed us

to dissect components that are required for apoplastic ROS

Ozone-triggered stomatal responses 449

ª 2010 The AuthorsJournal compilation ª 2010 Blackwell Publishing Ltd, The Plant Journal, (2010), 62, 442–453

responses in guard cells. Collectively, our results suggest

that the signal pathway from apoplastic ROS to guard cell

anion channel activation is at least partly independent from

previously described CO2 and ABA signaling pathways.

The use of protein kinase and phosphatase inhibitors has

previously shown that protein phosphorylation is involved

in the activation of anion channels (Schmidt et al., 1995;

Grabov et al., 1997; MacRobbie, 1998; Pei et al., 2000). Some

protein kinases involved in these responses have been

identified. The protein kinase HT1, a central component in

the regulation of stomatal movements by CO2, was not

required for the ROS activation of SLAC1. The activity of the

Ca2+-dependent protein kinases CPK3 and CPK6, which have

been shown to be involved in the Ca2+-dependent ABA

activation of S-type anion currents (Mori et al., 2006),

showed a normal RTD response. Thus, these protein kinas-

es, and other proteins, such as GCA2, previously shown to

be involved in CO2- or ABA- and Ca2+-dependent stomatal

regulation, are not components required for the apoplastic

ROS-induced regulation of stomatal movement. The OXI1

kinase, necessary for some ROS signaling processes (Rentel

et al., 2004), was not required for RTD either (Table S1). In

addition, ABA itself is not part of the signal cascade, as the

ABA-deficient aba1-3 mutant showed WT responses.

However, we have previously shown that in addition to

slac1 (Vahisalu et al., 2008), the ABA-insensitive protein

phosphatase type 2C mutant, abi2-1, completely lacked the

ozone-induced decrease in stomatal conductance (Kollist

et al., 2007). The ozone-triggered RTD was also missing in a

second dominant-negative mutant of the type-2C protein

phosphatases, ABI1, and in two mutant alleles of the OST1,

demonstrating that these proteins are essential in the

apoplastic ROS-induced RTD (Figure 5). It has been shown

recently that ABI1, ABI2 and OST1 interact physically both in

yeast two-hybrid and in planta assays, and that these and

other phosphatases act as negative regulators of OST1 via

PYR/PYL/RCAR proteins (Fujii et al., 2009; Umezawa et al.,

2009; Vlad et al., 2009). Similarly, our genetic data suggested

that ABI1 and ABI2 could be regulators of OST1 in ROS-

induced stomatal closure through dephosphorylation

(Figures 5 and 7). Using the split-ubiquitin yeast two-hybrid

and BiFC assays, we showed a physical interaction between

OST1 and SLAC1 (Figure S1b and 6a). The physical interac-

tion between SLAC1 and OST1, and the requirement for

OST1-dependent phosphorylation for the activation of

SLAC1, was also very recently demonstrated by two other

groups (Geiger et al., 2009; Lee et al., 2009).

Our phosphoproteomic experiments proved that OST1

was able to phosphorylate multiple amino acids of SLAC1,

including Ser120, but not Ser146 (Figure 6d–f), further

supporting that OST1 is responsible for SLAC1 activation.

This also suggests that Ser146 could be the target for a

different protein kinase. An alternative explanation for the

absence of RTD in slac1-8 (Figure 6c) could be a conforma-

tional change of SLAC1 protein, as in the slac1-8 mutant,

Ser146 is substituted with phenylalanine, which is a hydro-

phobic and considerably larger amino acid. SLAC1 has been

shown to be involved in the regulation of stomatal closure in

response to many factors, such as CO2, darkness, humidity,

ABA and Ca2+ (Vahisalu et al., 2008). Thus, it would be of

great interest to study the stomatal responses of slac1-7 and

slac1-8 to other stimuli, in order to address whether different

phosphorylation patterns of SLAC1 exist in response to

different stimuli.

Collectively, our data suggest a model for RTD (Figure 7)

where OST1 activates SLAC1 via phosphorylation, and

suggests the importance of S120 for this regulation. ABI1

and ABI2 keep OST1 inactive by dephosphorylation (Umez-

awa et al., 2009; Vlad et al., 2009). Possibly, the protein

phosphatase activity of ABI1 and ABI2 could also be directed

towards SLAC1: this should be addressed in further studies.

Recently, OST1 has been shown to phosphorylate and

reduce the activity of the inward K+ channel KAT1 (Sato

et al., 2009). Thus, OST1 would activate SLAC1 and at the

same time inactivate KAT1, leading to faster stomatal

closure.

Using ozone we show that the last steps of the ROS-

induced signaling cascade, leading to the activation of

SLAC1 and the induction of stomatal closure, could be as

Figure 7. Schematic model of events during ozone-triggered rapid transient

decrease (RTD) in stomatal conductance.

Ozone induces reactive oxygen species (ROS) production in the guard cells

(1). This leads to the activation of OST1 (2). The lack of RTD in abi1-1 and

abi2-1 suggests that the protein phosphatases ABI1 and ABI2 regulate OST1

activity (Umezawa et al., 2009). Our data suggest that several amino acid

residues of the N-terminal tail of SLAC1 are phosphorylated by OST1, and

highlight the functional importance of S120 (3). Phosphorylation leads to the

activation of S-type anion channels and anion efflux (A) out) from guard cells

(4), which in turn causes plasma membrane depolarization, and activation of

outward-rectifying K+ channel GORK and K+ efflux from guard cells (5). The

overall efflux of anions and K+ contributes to the loss of guard cell turgor,

leading to stomatal closure. For reopening, the activity of K+ uptake channels

are required (6).

450 Triin Vahisalu et al.

ª 2010 The AuthorsJournal compilation ª 2010 Blackwell Publishing Ltd, The Plant Journal, (2010), 62, 442–453

simple as the recently described minimal signaling pathway

for regulating ABA-induced gene expression (Fujii et al.,

2009). Furthermore, the two pathways share several com-

mon components: ABI1, ABI2 and OST1. Our next challenge

is to find out how the cells perceive ROS and translate this

into activation of OST1.

EXPERIMENTAL PROCEDURES

Plant material and growth conditions

For the whole-plant gas-exchange experiments, 24–26-day-oldplants were used. Plants were grown as described previously(Kollist et al., 2007). The ost1-1 mutant is in the Ler background(Mustilli et al., 2002). The ost1-3 is a T-DNA knock-out of OST1kinase in the Col-0 background (originally referred to as srk2e;Yoshida et al., 2002). For clarity, we refer to srk2e as ost1-3throughout this report. The source and identity of other mutantsused in the study are given in Table S1.

Whole-plant stomatal conductance measurements and

fluorescence microscopy

The Arabidopsis whole-rosette gas-exchange measurement detailswere described previously (Kollist et al., 2007). Prior to ozoneexposure, plants were acclimated in the measuring cuvettes for atleast 1 h. Plants were exposed to 350 nl l)1 ozone for 3, 12, 45 or90 min. Abaxial epidermal peels were isolated and loaded with100 lM H2DCFDA in 10 mM Tris–HCl, pH 7.2, for 5 min in darkness,and were washed with 10 mM Tris–HCl, pH 7.2. Clean-air controlpeels were isolated and loaded after acclimation in gas-exchangecuvettes.

ROS production was visualized by Nikon TE2000-U C1 confocalmicroscope (Nikon, http://www.nikon.com) using excitation at488 nm and emission at 530 nm. Images were processed usingNikon EZ-C1 FREEVIEWER software (gold version 3.30; Nikon).Brightness values of individual guard cell pairs were obtained aftercorrecting for the brightness of epidermal cells with IMAGEQUANT -v4.2a (Molecular Dynamics, now part of GE Healthcare, http://www.gelifesciences.com).

Isolation of TILLING lines

New ethyl methanesulphonate mutants slac1-6 (S38F), slac1-7(S120F) and slac1-8 (S146F) were identified through TILLING (Tillet al., 2003; http://tilling.fhcrc.org). Details are described in Appen-dix S1.

Split-ubiquitin membrane yeast two-hybrid assay

The split-ubiquitin yeast two-hybrid assay was conducted using theDUALmembrane kit 3 (Dualsystems Biotech, http://www.dualsystems.com). Details are described in Appendix S1.

BiFC interaction experiments

The cDNA of SLAC1 and OST1 was cloned into the pSPYNE andpSPYCE vectors. BiFC experiments were performed using transienttransfection of N. benthamiana leaves with Agrobacteriumtumefaciens, as described by Voinnet et al. (2003), images wereacquired 48–72 h after transfection by confocal microscopy (alsosee Appendix S1).

Protein expression and purification

OST1 and SLAC1 N-terminal fragments encoding amino acids 1–186(SLAC11–186) cDNAs were cloned into the pQE-30 UA (Qiagen,

http://www.qiagen.com) and pET28a (Novagen, now part of Merck,http://www.merck4biosciences.com) vectors, respectively. 6xHIS-OST1 was expressed using the XL-1 blue E. coli strain (Strata-gene, http://www.stratagene.com). 6xHIS-SLAC11–186 variants wereexpressed in Rosetta (DE3) pLysS cells (Novagen). Recombinantproteins were purified using a Chelating Sepharose� FastFlow (Amersham, now part of GE Healthcare, http://www.gelifesciences.com) column chelated with CoCl2 (for details, see Appen-dix S1).

In vitro kinase assays and mass spectrometry

Proteins were separated by SDS-PAGE using a 12% (w/v) acrylam-ide gel or 10% (w/v) acrylamide gel supplemented with Phos-Tag(Kinoshita et al., 2006). Gels were stained with Coomassie brilliantblue R-250 (Sigma-Aldrich, http://www.sigmaaldrich.com), andincorporation of 32P to the proteins was detected and visualized byautoradiography (for further details see Appendix S1). NanoLC-MS/MS analysis for the mapping of OST1 phosphorylation sites inSLAC11–186 was carried out by using LTQ-Orbitrap (Thermo FisherScientific, http://www.thermofisher.com), equipped with a nano-spray source (Proxeon, http://www.proxeon.com) and a 1200 Seriesnano-LC system (Agilent Technologies, http://www.agilent.com)(for details, see Appendix S1).

ACKNOWLEDGEMENTS

We acknowledge several labs (listed in Table S1) for sharing theirmutants. Work in the labs of HK and ML was funded by the EstonianScience Foundation (grants 7763, 6766, 7869, 7361 and EMP24)and Estonian Ministry of Education and Research (themeSF0180071s07). Work in the JK’s lab was supported by the Academyof Finland Centre of Excellence program (2006–2011), MB wassupported by an Academy of Finland Post-Doctoral grant (deci-sion# 108760), and TV was supported by the Finnish GraduateSchool in Plant Biology.

SUPPORTING INFORMATION

Additional Supporting Information may be found in the onlineversion of this article:Figure S1. The split-ubiquitin yeast two-hybrid assay suggests thatSLAC1 interacts with OST1 kinase.Figure S2. Mass spectrometry analysis suggests that OST1 phos-phorylates at least four serines in the SLAC1–186 fragment.Table S1. List of mutants tested for ozone-triggered rapid transientdecrease (RTD) in stomatal conductance.Table S2. Statistical analysis of data presented in Figure 3a (Tukey’shonestly significant difference test).Appendix S1. Supplementary experimental procedures.Please note: As a service to our authors and readers, this journalprovides supporting information supplied by the authors. Suchmaterials are peer-reviewed and may be re-organized for onlinedelivery, but are not copy-edited or typeset. Technical supportissues arising from supporting information (other than missingfiles) should be addressed to the authors.

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