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Review

Blackwell Publishing, Ltd.

Tansley review

In the light of stomatal opening: new

insights into ‘the Watergate’

M. Rob G. Roelfsema and Rainer Hedrich

Molecular Plant Physiology and Biophysics, Julius-von-Sachs Institute for Biosciences, Biocenter,

Würzburg University, Julius-von-Sachs-Platz 2, D-97082 Würzburg, Germany

Contents

Summary 665

I. Introduction 665

II. The hydrodynamic valve 666

III. Regulation of ion transport in guard cells 678

IV. Interaction between guard cell signaling pathways 682

V. Outlook 684

Acknowledgements 685

References 685

Author for correspondence:

Rainer Hedrich Tel: +49 931 8886100 Fax: +49 931 8886157 Email: [email protected]

Received:

7 December 2004

Accepted:

14 March 2005

Key words:

abscisic acid (ABA), blue light, CO

2

, crosstalk, guard cell, ion channel, signaling pathway, stomata.

Summary

Stomata can be regarded as hydraulically driven valves in the leaf surface, whichopen to allow CO

2

uptake and close to prevent excessive loss of water. Movementof these ‘Watergates’ is regulated by environmental conditions, such as light, CO

2

and humidity. Guard cells can sense environmental conditions and function as motorcells within the stomatal complex. Stomatal movement results from the transport ofK

+

salts across the guard cell membranes. In this review, we discuss the biophysicalprinciples and mechanisms of stomatal movement and relate these to ion transportat the plasma membrane and vacuolar membrane. Studies with isolated guard cells,combined with recordings on single guard cells in intact plants, revealed that lightstimulates stomatal opening via blue light-specific and photosynthetic-active radiation-dependent pathways. In addition, guard cells sense changes in air humidity and thewater status of distant tissues via the stress hormone abscisic acid (ABA). Guard cellsthus provide an excellent system to study cross-talk, as multiple signaling pathwaysinduce both short- and long-term responses in these sensory cells.

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I. Introduction

Stomata can be regarded as hydraulically driven valves thatoperate in the aerial parts of land plants. These pores enablethe diffusion of gases between the plant’s interior and theatmosphere. As a result of the swelling and shrinking of two

guard cells that border the pore, stomata can open and closeand thus form a ‘Watergate’ that controls the loss of water(for uptake and transport of water in plants see review byZimmerman

et al

., 2004). The regulation of stomatal movementby endogenous and environmental signals has fascinated plantbiologists for centuries (Meidner, 1987). In 1812 the German

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botanist, Moldenhauer, wrote about the stomata ofmaize:

… nur am frühen Morgen, wenn die Sonne auf die nochthauigen Blätter schien, oder wenn ich sonst das Blatt, ohnees vom Stengel zu trennen, oder es zu beschädigen, in einGefäß mit Wasser bog, und es so dem Sonnenlicht aussetzte,wenn also die Ausdünstung der Blätter zurückgehaltenwurde, sah ich sie geöffnet. … ,

Translated, this states that he only observed open stomatain the early morning with the sun shining on still-dewy leaves,or when he put the leaf, without excision from the stem, intoa jar with water and let the sun shine on it, in other wordswhen transpiration was prevented. Moldenhauer (1812) rec-ognized three important signals affecting stomatal movement:light, humidity, and time of day. The pathways coupling thesesignals to stomatal movements have, to date, only beenpartially resolved and are still the subject of intensive research.

The central function of stomata is to balance the uptake ofCO

2

, which is essential for photosynthesis, with loss of waterfrom the plant. During evolution, land plants developed acuticle on aerial tissues to prevent dehydration. This gas-impermeable waxy layer, however, also limits the uptake ofCO

2

. To circumvent the latter restriction, the leaf epidermisharbors a large number of pores, named stomata. The degreeof opening of these stomata is determined by two guard cellsthat surround the stomatal pore. Apart from leaves, stomataare, in certain plants, also found on the stem, petioles, primaryroots (Christodoulakis

et al

., 2002) or in nectaries (Horner

et al

.,2003). In these organs, stomata presumably have functionsother than facilitating CO

2

uptake, such as the extrusion of nectarfrom active nectaries (Horner

et al

., 2003). Little is known aboutthe regulation of stomatal movement in these nonphotosyn-thetic organs. In this review, we will therefore focus on thecontrol of stomatal movement in photosynthetic-active leaves.

In general, light and drought act in an antagonistic manneron stomatal movement. Light induces the opening of stomatato enhance CO

2

uptake, while drought causes stomata toclose, thereby limiting water loss through transpiration. Innature, stomata will often receive both signals at the sametime, as sunny periods frequently coincide with drought.Under these circumstances, plants will have to make a tradeoff – either open the stomata for optimal photosynthesis, orclose them to save H

2

O. The fine-tuning of this process prob-ably differs between plant species and will also depend onother growth conditions and the developmental stage of theplant. However, the principal mechanisms used to processthese signals are probably conserved within the plant king-dom. An exception to this rule is the group of plants withcrassulacean acid (CAM) metabolism. These plants open theirstomata at night and close them during the day, which allowsthem to grow in extremely dry environments. Although theregulation of stomatal movement by light is inversed in CAMplants, the signaling pathways may still have features in

common with C

3

and C

4

plants (Section III, heading 1). Formore details on CAM metabolism see previous excellent reviewsby Black & Osmond (2003) and Lüttge (2002).

This present review will attempt to couple recent advancesin the understanding of guard cell ion-transport with changesin the osmotic pressure, turgor and cell volume. Guard cellshave often been studied in isolation from neighboring cells,and the results were seldom tested for significance in intactplants. At the 13th international workshop on Plant Mem-brane Biology (2004), in Montpellier, this led to the remarkthat: ‘Guard cells seem to have gained life independent of theplant’. Here, we will attempt to put guard cells back in theirnatural context and discuss the current knowledge of guardcell physiology in relation to stomatal function in the intactplant. In this context, the interaction of guard cells withneighboring cells, and their responses to signals from moredistant cell types, will be addressed.

Although stomata are found on most land plants, a few spe-cies have been favored for research on stomata. In this reviewwe will mainly discuss the guard cells of broadbean (

Vicia faba

L.), as guard cells of this species have been studied by using gasexchange, ion-uptake measurements, microprobe analysis,biochemistry, electrophysiology and the pressure probe. Formolecular biology and signal transduction research, guardcells of

Arabidopsis thaliana

are often the system of choice.The stomata of the latter two species differ considerably in size(Langer

et al

., 2004), but also have many features in common.The stomatal complexes of these two dicot species consistonly of two kidney-shaped guard cells and they are distributedon the leaf surface without an obvious patterning. In additionto these dicot species, two nongrass monocot species –

Commelina communis

and

Trandescantia virigiana

– have beenimportant in stomatal research. The epidermis of these speciesis easily detached from the leaf (Weyers & Travis, 1981) andhas been used to study ion uptake (MacRobbie & Lettau,1980a,b; MacRobbie, 1987), turgor pressure (Franks, 2003)and cytoplasmic Ca

2+

signaling (McAinsh

et al

., 1995). Thestomatal complexes of these monocots consist of guard cellsthat are bordered by subsidiary cells and they are distributedparallel to each other on the leaf surface. Stomata of Gramineaeare even more distinct because their guard cells are dumbbellshaped. The mechanics of stomata of these species may differto some extent; however, the hypothesis put forward by VonMohl (1856) – ‘die Porenzellen erweitern durch ihre Turgeszenzdie Spaltöffnung und verengen sie durch Collabiren’ – whichmeans that guard cells open the stomatal pore with their turgor andclose it by collapsing, seems to hold for stomata of all plant species.

II. The hydrodynamic valve

1. Osmotic motor

Water potential

Like most motions exerted by plants,stomatal movements are hydraulically driven. Stomatal

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movements, however, differ from many other motions exertedby plants because they are completely reversible. Movementsrelated to growth, like hypocotyl bending, involve both celldivision and expansion and therefore are only partiallyreversible. Water movement into the guard cell is driven byosmosis and can be best understood considering the chemicalpotential of water. Accumulation of solutes in the guard cellcytoplasm lowers the chemical potential of water (µ

w

) and itsderivative, ‘water potential’ (

Ψ

). The water potential (Eqn 1)is equivalent to the chemical potential of a solution, relativeto the chemical potential of pure water ( ) at the sametemperature and atmospheric pressure, divided by the partialmole volume of water,

◊

w

(Nobel, 1970), as follows:

(Eqn 1)

The water potential of pure water thus is 0 and it becomesmore negative when solutes accumulate. The role of the waterpotential in stomatal movement has been reviewed byRaschke

et al

. (1988) and is based on the relationship (Eqn 2)of the water potential (

Ψ

), turgor pressure (

p

) and osmoticpressure (

π

):

Ψ

=

p

−

π

. (Eqn 2)

This means that an increase in the osmotic potential of theguard cell leads to an initial decrease in the water potential.Given a high hydraulic conductivity of the plasma membrane,water will flow into the guard cell and the

Ψ

of the guard cellwill equilibrate with that of the apoplast. The inflow of waterwill cause the turgor pressure to rise and the guard cells to

swell. The increase in volume of both guard cells causesopening of the stomatal pore.

As a result of the radial arrangement of cellulose fibrils inthe cell wall, the expansion of kidney-shaped guard cellsoccurs mainly along the longitudinal axis (Sharpe

et al

., 1987;Shope

et al

., 2003). As a result, the guard cells bend, keepingthe length of the stomatal complex virtually unchanged. Inaddition to bending, the cross section of guard cells changesfrom a flattened oval to a circular shape during stomatal open-ing (Fig. 1) (Von Mohl, 1856; Raschke & Dickerson, 1972;Franks

et al

., 2001). Changes in guard cell shape and volumeduring stomatal movements were recently quantified forguard cells of

V. faba

by using a pressure probe and confocalmicroscopy (Franks

et al

., 2001; Shope

et al

., 2003). Shope

et al

. (2003) found that the volume of guard cells in openedstomata ranges from 5 to 8 pl and decreases, during stomatalclosure, to 3.5–4.5 pl. This indicates that guard cells of

V. faba

lose

≈

40% of their volume during stomatal closure.

Guard cell volume and turgor

The increase in cell sizeduring stomatal opening also leads to an increase of the guardcell surface. These processes were studied with fluorescentdyes that accumulate in the plasma membrane (Shope

et al

., 2003). Images obtained with a confocal laser-scanningmicroscope revealed an approximately equal change in surfacearea and guard cell volume during stomatal movement (Shope

et al

., 2003). This implies that the guard cells need to expandtheir plasma membrane surface by

≈

40% during stomatalopening. Such a large change in surface area is probablyaccomplished by fusion of vesicles with the plasma membrane,while vesicles are budding from the plasma membrane during

Fig. 1 Changes in the shape of guard cells during stomatal movements. During stomatal opening the guard cell volume increases, which causes them to bend and changes their cross section from an flat oval (right drawing), to a circle (left drawing). Redrawn after Raschke & Dickerson (1972) and Wanner (2004).

µw*

Ψ *=−µ µw w

w◊

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stomatal closure. The latter process was supported with fluid-phase endocytosis (Diekmann

et al

., 1993) and confirmedby emerging fluorescent vesicles derived from the plasmamembrane during stomatal closure (Shope

et al

., 2003; Meckel

et al

., 2004). Vesicle fusion and fission of plant protoplastscan be studied by using the patch clamp technique, as thecapacitance of the protoplast is linearly related to its plasmamembrane surface (Homann, 1998). Application of hydrostaticpressure to the guard cell protoplasts reversibly alters themembrane capacitance. In addition, changes in the plasmamembrane conductance occur as ion-channels are incorporatedinto or removed from the plasma membrane (Homann &Thiel, 2002; Hurst

et al

., 2004). Hydro-passive volume changesin guard cell protoplasts thus provide an elegant systemto study vesicle trafficking and proteins involved this process,such as the syntaxins (Leyman

et al

., 1999).The turgor pressure necessary to provoke an increase in

guard cell volume and surface area has been recorded forintact guard cells by using the pressure probe (Franks

et al

.,1998; Franks

et al

., 2001). A pressure of 4.5 MPa was requiredfor a maximal stomatal opening, a value well in agreementwith the data of Raschke

et al.

(1972) and Fischer (1972)(Tables 1 and 2). The latter authors used concentrated sucrosesolutions to induce stomatal closure, and achieved completeclosure in solutions with an osmotic pressure of 8 MPa(Raschke

et al

., 1972). Taking into account a decrease in cellvolume of 40% during stomatal closure, the osmotic pressureof guard cells in fully opened stomata was 4.8 MPa. Theseresults led to the conclusion that the turgor of

V. faba

guardcells increases with 0.23 MPa µm

−

1

increase in stomatal aper-ture (Table 2). Fischer (1973) determined a very similar valueof 0.2 MPa µm

−

1

. Apparently, a turgor increase of

≈

0.2 MPais necessary to drive a 1-µm increase in aperture for

V. faba

stomata in epidermal strips.

Interaction of guard cells and epidermal cells

It shouldbe noted that the data of Fischer (1972) and Raschke

et al

.(1972) were obtained using epidermal peels of

V. faba

inwhich the epidermal pavement cells were disrupted. In thelatter preparations, the maximal stomatal aperture was

≈

12 µ

. In the experiments of Franks

et al

. (1998), theepidermal cells remained intact and the maximal aperture wasonly 8 µm. These differences are in agreement with dataof Klein

et al

. (1996), who compared fusicoccin-inducedstomatal opening in epidermal strips of

V. faba

, with viableand disrupted epidermal pavement cells. The stomatalopening was 8 µm in the presence and 16 µm in the absenceof viable epidermal cells. A reduction in the stomatal apertureof

≈

50% by viable subsidiary cells and epidermal cells, overa whole range of apertures, was also found for

T. virgiana

(Franks

et al

., 1998) and for

C. communis

(MacRobbie &Lettau, 1980a). Apparently, viable epidermal cells provide aback pressure that reduces the stomatal aperture by

≈

50%.The absence of back pressure in many epidermal strip pre-

parations also explains why stomata often do not completelyclose, while fully closed stomata are generally observed indark-adapted intact leaves (Felle

et al

., 2000).The interaction of epidermal cells with the stomata was

studied in detail by using epidermal strips of

T. virgiana

(Franks

et al

., 1998). If the epidermal cells were disrupted, thestomata started to open as soon as turgor pressure was applied.When epidermal cells had a turgor pressure of 0.9 MPa, how-ever, no stomatal opening was observed with guard cell turgorpressures of < 1.25 MPa. At higher pressures, stomata startedto open, but viable epidermal cells reduced the final stomatalaperture (Franks

et al

., 1998; Franks, 2003). The increase ofturgor without causing stomatal opening has been describedas the ‘Spannungsphase’, by Stålfelt (1929). At higher turgorpressures, guard cells are in the ‘Motorphase’, in which anincrease in turgor leads directly to stomatal opening. Duringthe ‘Motorphase’ a virtual linear relationship between theguard cell turgor and the stomatal aperture exists, as long asthe aperture is not limited by constrains in the extensibility ofthe cell wall (Franks

et al

., 1998).

Table 1 Biophysical parameters of open and closed Vicia faba stomata in the presence or absence of viable epidermal cells

Closed stomata Open stomata

Guard cell volume(disrupted epidermal cells)

4 pla 6.5 pla

Stomatal aperture(disrupted epidermal cells)

2 µmb 14 µmb

Guard cell turgor(disrupted epidermal cells)

2.0 MPac 4.8 MPac

Stomatal aperture(viable epidermal cells)

0 µmd 8 µmd

Guard cell turgor(viable epidermal cells)

1.0 MPae 4.5 MPae

K+ content (disrupted and viable epidermal cells)

0.3 pmolf 2.5 pmolf

aThe volume of closed stomata ranges from 3.5 to 4.5 pl and that of open stomata from 5 to 8 pl (Shope et al., 2003).bThe minimal opening of stomata in epidermal strips with distrupted epidermal pavement cells is 2 µm according to Raschke et al. (1972), and the maximal opening ranges from 12 µm (Raschke et al., 1972) to 16 µm (Klein et al., 1996).cRaschke et al. (1972) determined incipient plasmolysis at 2 MPa for closed stomata and at 8 MPa for open stomata; the latter value corresponds to a turgor pressure of 4.8 MPa, taking into account a reduction in volume of 40%.dStomata adjacent to intact epidermal cells completely close (Felle et al., 2000) and open to ≈ 8 µm (Klein et al., 1996; Franks et al., 1998).eChanges in stomatal aperture in strips with turgescent epidermal cells occur at guard cell turgor pressures ranging from 1.0 to 4.5 MPa (Franks et al., 1998).fThe closed stomata were found to contain 0.1–0.55 pmol of potassium per guard cell, while open stomata of guard cells were reported to contain 2.1–3.0 pmol (Fischer, 1972; Humble & Raschke, 1972; Allaway & Hsiao, 1973; Outlaw & Lowry, 1977).

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Ion accumulation In intact plants, guard cells need toaccumulate a large volume of solutes to raise their osmoticpotential and force the stomatal pore to open. In the earlyliterature, guard cells were believed to convert starch into sugars,as starch disappears from guard cell chloroplasts during stomatalopening in the light (Strugger & Weber, 1926). However, thisis not the case for guard cells of all plants, as A. thaliana guardcells synthesize starch during the light period and degradethis polymer in the dark (Stadler et al., 2003). In this respect,guard cells are not different from mesophyll cells, whichaccumulate starch during the light period and remobilizecarbohydrates in the dark (Zeeman et al., 1998). Apparently,the storage and breakdown of starch in guard cells does notnecessarily correlate to stomatal movement.

Later reports showed that stomatal opening depends onthe uptake of K+ into guard cells (Fischer, 1968). Guard cellsof V. faba take up ≈ 2 pmol of K+ during stomatal opening(MacRobbie, 1987), which relates to a K+ uptake of0.18 pmol of K+ µm−1 (Table 2); Fischer (1972) determineda value of 0.21 pmol of K+ µm−1. In the long term, guard cellscan only accumulate positively charged ions, when these arecounterbalanced by anions. For guard cells in epidermal stripsof V. faba, K+ uptake was balanced by the accumulation ofequal amounts of Cl– and malate– (Raschke & Schnabl, 1978).Providing that the latter would hold for guard cells in intact

plants, these motor cells accumulate 2 pmol of K+, 1 pmol ofCl– and 1 pmol of malate–. Malate will probably have only asingle negative charge as it is predominantly stored in the guardcell vacuole (Schnabl & Kottmeier, 1984a), which presumablyhas an acidic pH. The maximum increase in the osmotic pressurecan now be calculated (Nobel, 1970) as follows:

, (Eqn 3)

where πs is the osmotic pressure of all solutes, Σ cj is the sumof the concentration of all solutes and RT has a value of2.44 l MPa mol−1 at 20°C. Given a volume of 4 pl, the increase inπs would be 2.4 MPa. Considering a turgor pressure of≈ 1 MPa of guard cells in closed stomata, the final turgorpressure would be ≈ 3.4 MPa. This is less than the 4.5 MPathat would be expected for fully open stomata. In fact, theincrease in osmotic pressure has been overestimated above, asthe increase in volume of the guard cells during stomatalopening was not taken into account. Furthermore, Eqn 3 onlyholds for dilute solutions, as it overestimates the osmoticpressure of concentrated solutions (Nobel, 1970). Apparently,the measured accumulation of K+ salts in guard cells does notcause a rise in osmotic pressure large enough to explain theincrease in stomatal aperture, a discrepancy also observed byFischer (1972) and MacRobbie (1987). This indicates thateither the accumulation of K+ has been systematically under-estimated, or that other solutes accumulate in addition to K+ salts.

Sugar uptake There are several lines of evidence indicatingthat guard cells accumulate sugars, in addition to K+ salts,during stomatal opening. The concentration of sucrose inguard cells of V. faba can increase, from 0.2 to 0.7 pmol percell, during the light period (Talbott & Zeiger, 1996).However, sucrose mainly accumulates during the late lightperiod, when K+ concentrations already decrease again. Theaccumulated sucrose thus may just replace K+ over the courseof the day, instead accelerating stomatal opening. The role ofsugars in stomatal movement is supported by the resultsof Ritte et al. (1999), who found that guard cell protoplastsof Pisum sativum take up significant amounts of glucose,fructose and sucrose. More recently, the A. thaliana hexosetransporters AtSTP1 (Stadler et al., 2003) and AtSTP13 (M.Büttner, pers. comm.) were found to be highly expressed inguard cells. However, the transcript level of the AtSTP1transporter was highest in the dark, which brings its role instomatal opening into question. Mutant plants lackingAtSTP1 also did not show a reduction in stomatal opening,indicating that either other hexose transporters (such asAtSTP13) or sucrose transporters can act as a backup forAtSTP1, or that sugar uptake is not required for stomatalopening.

Although there is no problem in understanding how theuptake of sugars can contribute to stomatal opening, it isunknown how sugars can disappear rapidly from the guard

Table 2 Estimated changes in biophysical parameters of Vicia faba stomata during stomatal movement, in the presence or absence of viable epidermal cells

Change in:Disrupted epidermal cells

Viable epidermal cells

Guard cell volume 0.21 pl µm−1 a b

Guard cell turgor 0.23 MPa µm−1 c 0.46 MPa µm−1 d

K+-content 0.18 pmol µm−1 e 0.36 pmol µm−1 f

aOwing to the approximate linear relationship between stomatal volume and turgor pressure (Shope et al., 2003), as well as between turgor pressure and stomatal aperture (Franks et al., 1998), a volume change of 0.21 pl µm−1 can be calculated for stomata in epidermal peels with disrupted epidermal cells. (Based on the data shown in Table 1.)bIt is unclear how the volume changes of guard cells during stomatal movement is influenced by viable epidermal cells.cThe data in Table 1 relate to a change in guard cell turgor pressure of 0.23 MPa µm−1 for stomata in epidermal strips with disrupted epidermal cells.dViable epidermal cells reduce the stomatal aperture by ≈ 50% and thus the change in guard cell turgor per µm aperture is twice as large. Franks et al. (1998) determined a change of 0.4 MPa µm−1.eBased on the data in Table 1, guard cells accumulate 0.18 pmol of K+ per µm increase in stomatal aperture. Fischer (1972) concluded that guard cells take up 0.21 pmol µm−1, in the absence of viable epidermal cells.fIn the presence of viable epidermal cells the stomatal aperture is reduced by 50%, indicating that guard cells take up twice as much K+ per µm incease in stomatal aperture.

πs jRT c = ∑

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cell cytoplasm during stomatal closure. The latter process canoccur very rapidly, as abscisic acid (ABA) induces stomatalclosure within 10 min (Roelfsema et al., 2004). The rate ofstarch synthesis is regulated by the activity of ADP-glucosepyrophosphorylase, which is ≈ 35 mmol kg−1(dry weight) h−1

in guard cells (Outlaw & Tarczynski, 1984). Based on a dryweight of 3.1 pg per guard cell (Outlaw & Lowry, 1977), thiscorrelates to the conversion of 0.1 pmol of ADP-glucose h−1

per guard cell, which is much too slow to support ABA-induced stomatal closure. It is therefore more likely thatsugars are extruded from guard cells during stomatal closure,which also would explain the accumulation of sucrose in theapoplast of guard cells at low humidity (Outlaw & DeVlieghere-He, 2001). The transport proteins that facilitatesuch an extrusion of sugars, however, have not yet been iden-tified, but it may be that STP- and SUC-transporters can actin the reverse mode (own unpublished data).

Na+-supported stomatal movement Under physiologicalconditions, guard cells will mainly accumulate K+ salts duringstomatal opening because this cation is available atconcentrations of 3–40 m in the guard cell wall (Felle et al.,2000; see review Roelfsema & Hedrich, 2002). However,stomatal opening is also supported by other monovalentcations such as Rb+, Li+, Cs+ and Na+ (Humble & Hsiao,1969). Most of these cations will not accumulate in theapoplast at concentrations high enough to compete with K+.An exception is Na+, for plants growing on salt-enriched soils.Na+ supports stomatal opening in V. faba and C. communis,and in C. communis Na+ is even more efficient than K+

(Willmer & Mansfield, 1969). However, stomatal closureis disabled in stomata that were previously opened in thepresence of Na+ ( Jarvis & Mansfield, 1980; Robinson et al.,1997). Guard cells thus seem to be equipped with a Na+-uptake system, but have limitations considering Na+ release.The inability to extrude Na+ may be caused by Na+ blockageof plasma membrane K+ channels (Thiel & Blatt, 1991).Alternatively, it may be the result of an inability of slowvacuolar (SV) channels to extrude Na+ (Ivashikina &Hedrich, 2005; see also section II, 2). The latter result is wellin agreement with the swollen vacuoles observed for guardcells that were opened in the presence of Na+ (MacRobbie,1983). Halophytes may have developed two mechanisms toovercome this problem: they are either capable of extrudingNa+ or they selectively accumulate K+, but not Na+ (Robinsonet al., 1997). In the halophyte Aster tripolium, the latteradaptation was associated with a Na+-induced change in theactivity of K+-uptake channels (Véry et al., 1998).

2. Guard cell ion transporters

Ion-transport proteins of guard cells have been studied ingreat detail because their activity is linked directly to stomatalmovement. Guard cell ion transport differs from that in most

other plant cells because ion transport in guard cells occurs intwo directions (to enable stomatal opening as well as closure).By contrast, other plant cells (such as mesophyll- or epidermalpavement cells) grow irreversibly. It is therefore highly likelythat the properties of ion transporters, and their regulation byvarious signals, differ between different cell types. Cell type-specificity of ion-transport genes has rarely been documented,but each cell seems to run a unique set of transport proteins.For instance, the expression rate of the K+-channel gene,GORK, is very high in guard cells, but it is virtually absentfrom mesophyll cells (Ache et al., 2000).

The regulation of guard cell ion transport by light andhumidity will be addressed in section III. Below we will reviewthe present knowledge about the properties of ion-transportproteins in the guard cell plasma- and vacuolar membrane.Because of the large data set available for these transportproteins, it is impossible to cover this in every detail (seereview Véry & Sentenac, 2002). Instead, we will focus on theproperties of transporters linked to stomatal function (Fig. 2).

Plasma membrane transport K+ transport The importanceof K+ uptake during stomatal opening has been undisputed,ever since Fischer showed, in 1968, that stomatal movementin epidermal strips depends on the uptake of K+ (Fischer,1968). However, the nature of the proteins facilitating K+

uptake remained unclear until Schroeder et al. (1984) showedthe presence of K+-selective ion channels in the plasmamembrane of V. faba guard cells. Another breakthrough came8 yr later with the isolation of the first plant K+-channel genes(Anderson et al., 1992; Sentenac et al., 1992), which wasfollowed by the guard cell-specific expression of the K+

channel encoded by KST1 and KAT1 (Müller-Röber et al.,1995; Nakamura et al., 1995).

Plant plasma membrane K+-channel types can be dividedinto three groups: outward-rectifying channels, which becomemore active at positive potentials (Fig. 3c); inward-rectifyingchannels, activating at more negative potentials (Fig. 3d); anda third type of channel that is largely voltage-independent.The first two types of K+ channels have been recorded inguard cells of various plant species, such as V. faba (Schroederet al., 1987; Blatt, 1988) A. thaliana (Roelfsema & Prins,1997; Pei et al., 1997), Nicotiana bethamiana and N. tabacum(Armstrong et al., 1995; Dietrich et al., 1998), Zea mays(Fairley-Grenot & Assmann, 1993) and Populus tremula ×P. tremuloides (Langer et al., 2004). Based on their voltagedependence, outward-rectifying K+ channels are, at physio-logical conditions, responsible for K+ extrusion, while inward-rectifying channels mediate K+ uptake. Genes encodingvoltage-independent K+ channels are only expressed at lowrates in guard cells and the activity of these channels hasnot, so far, been recognized for guard cells (Blatt et al., 1990;Blatt, 1992; Roelfsema & Prins, 1997, 1998).

The gating of outward- and inward-rectifying K+ channelsdiffers in a crucial way: activation of outward K+ channels is

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dependent on the extracellular K+ concentration, while thatof inward K+ channels is not. Outward-rectifying K+ channelsrespond to changes in the extracellular K+ concentration insuch a way that the channel always activates at potentialsslightly more positive than the Nernst potential for K+ (Blatt,1988; Roelfsema & Prins, 1997; Gaymard et al., 1998; Acheet al., 2000). In contrast, the activation of inward channels isnot affected by K+ (Blatt, 1992; Roelfsema & Prins, 1997),and these channels do not exclusively facilitate K+ uptake,but can mediate K+ extrusion at very low extracellular K+ con-centrations (Véry et al., 1995; Brüggemann et al., 1999). Inintact plants of V. faba, the K+ concentration in the guard cellwall has been estimated at between 3 and 40 m (Felle et al.,2000; Roelfsema & Hedrich, 2002). The outward K+ channelof guard cells in intact V. faba becomes active at potentialsclose to −50 mV, while the inward rectifier activates at poten-tials negative of −100 mV (Roelfsema et al., 2001). In intactplants of A. thaliana (Fig. 3a,b) and Populus (Langer et al.,2004) the outward K+ channels also activate at −50 mV, butthe threshold potential of inward K+ channels is more negativethan −100 mV. A species-specific threshold potential for

Fig. 2 Overview of ion-transport proteins that have been identified or postulated in the plasma membrane (upper scheme) and vacuolar membrane (lower scheme) of guard cells. Transport proteins, for which one or more genes have been isolated, are shown in black, while those for which only physiological evidence exists are in grey. 1. Outward-rectifying K+ channel, GORK. 2. Inward-rectifying K+ channel, KAT1, KAT2, AKT1, AKT2/3 and AtKC. 3. Hyperpolarization-activated Ca2+ channel. 4. Ca2-ATPase, ACA. 5. R-type anion channel. 6. S-type anion channel. 7. H+-ATPase, AHA. 8. NO3

− transporter, CHL1. 9. Cl– transporter. 10. Slow vacuolar (SV) channel; 11. Fast vacuolar (FV) channel. 12. Vacuolar K+-selective (VK) channel. 13. K+ transporter, NHX. 14. H+-pyrophosphatase. 15. V-type H+-ATPase, V1 and V0-subcomplexes. 16. Ca2+-dependent protein kinase (CDPK)-activated anion channel. 17. Hyperpolarization-activated anion channel. 18. Malate carrier, AttDT. 19. Ca2+-carrier, CAX; ACA. 20. Vacuolar ACA. 21. Voltage-gated Ca2+ channel. 22. Inositol triphosphate (IP3)- and inostitol hexakis-phosphate (IP6)-gated Ca2+ channels. 23. Cyclic ADP ribose-activated Ca2+ channel.

Fig. 3 K+ channel currents in guard cells of intact plants and through ‘Shaker like’ K+ channels expressed in Xenopus oocytes. (a) Plasma membrane current of an Arabidopsis thaliana var. Landsberg erecta guard cell on the abaxial side of a leaf. The plasma membrane was stepped from a holding potential of −80 mV, to test potentials ranging from −180 mV to 20 mV, at 20-mV increments. Note the activation of inward- and outward-rectifying K+ channels with increasing negative- and positive potentials, respectively. (b) Current–voltage relationship of steady-state currents from the same cell as in (a). Note the activation of inward- and outward-rectifying K+ channels at potentials negative of −120 mV and positive of −60 mV, respectively. (c) Whole-oocyte currents of GORK recorded in a bath

solution containing 30 mM KCl, 1 mM CaCl2, 1 mM MgCl2 and 10 mM Tris/Mes, pH 7.5. Currents were elicited from a holding potential of −80 mV to test potentials ranging from −80 to 50 mV, at 10-mV increments. (d) Oocyte currents of KAT1 recorded in 30 mM KCl, 1 mM CaCl2, 1 mM MgCl2 and 10 mM Mes/Tris, pH 5.6. Currents were elicited with test potentials ranging from 20 to −150 mV, at 10-mV increments.

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activation was not found by using patch-clamp recordings(Dietrich et al., 1998), indicating that the threshold potentialis set by cytoplasmic factors that are lost during the whole-cellconfiguration.

The first two plant K+ channels were cloned from Arabidopsis,and denoted KAT1 (Anderson et al., 1992) and AKT1(Sentenac et al., 1992); this was followed by the identificationof the first KAT1 homolog, KST1, from potato (Müller-Röber et al., 1995). All these genes encode inward-rectifyingK+ channels when expressed in Xenopus oocytes (Schachtmanet al., 1992; Müller-Röber et al., 1995; Véry et al., 1995) oryeast cells (Bertl et al., 1995). These and later studies revealedthat all members of the ‘Shaker like’ K+ channels are localizedin the plasma membrane and function as K+-selective ionchannels (Dietrich et al., 2001; Véry & Sentenac, 2002).KST1 and KAT1 are highly expressed in guard cells (Müller-Röber et al., 1995; Nakamura et al., 1995) and therefore soonbecame known as guard cell K+-uptake channels. Mutants ofA. thaliana, with an En-transposon inserted in the openreading frame of KAT1, were therefore expected to lack inwardK+ channels. However, recordings with intracellular micro-electrodes revealed no difference between inward K+ currentsof the KAT1 En-insertion mutant and wild type (Szyrokiet al., 2001). Furthermore, these loss-of-function mutantsdisplayed normal K+-uptake kinetics and stomatal opening.This led to a search for other K+-channel genes expressed inguard cells, which could back up for the loss of KAT1. Indeed,guard cells were found to express the K+-channel genes KAT2(Pilot et al., 2001), AKT1, AKT2/3 and AtKC1, although atlower expression levels than that of KAT1 (Szyroki et al., 2001).Functional K+ channels probably assemble four subunits,encoded by one or more of these genes, to form a homomericor a hetromeric complex (Dreyer et al., 1997). This mightexplain why the introduction of a nonfunctional KAT1 genelowered the inward K+ conductance of guard cells by 70%(Kwak et al., 2001). Nonfunctional KAT1 subunits may formtetrameric channel proteins in combination with all fivewild-type genes, thereby creating nonfunctional K+ channels.

The properties of inward K+ channels are thus not encodedby a single gene, but are probably the result of differentproperties encoded by at least five genes (Szyroki et al., 2001).The subunits KAT1, KAT2 and AKT1 all encode inward-rectifying K+ channels in heterologous expression systems.The fourth gene, AtKC1, probably does not encode a functionalK+ channel itself, but its expression can alter the propertiesof K+ channels formed by the other subunits (Reintanz et al.,2002). The fifth gene expressed in guard cells, AKT2/3, encodesa largely voltage-independent channel when expressed inXenopus oocytes (Marten et al., 1999; Lacombe et al., 2000).This channel probably plays an important role in the phloemof the shoot (Marten et al., 1999), but not in roots (Birnbaumet al., 2003). Voltage-independent K+ channels have not beenrecognized for guard cells, indicating that the properties theAKT2/3 subunits may change through interaction with other

K+-channel subunits (Ivashikina et al., 2003), or as a result ofphosphorylation (Chérel et al., 2002).

In contrast to the inward-rectifying K+ channels, outward-rectifying K+ channels in the guard cells of A. thaliana areencoded by a single gene only, named GORK (Ache et al., 2000).Disruption of the GORK gene causes a complete loss of theoutward-rectifying K+ channels in the plasma membrane ofguard cells (Hosy et al., 2003). The stomata of the GORKloss-of-function mutant close more slowly after treatment withABA than do wild-type stomata (Hosy et al., 2003). However,the fact that stomata still close in response to ABA and darknessin gork-1, indicates that guard cells possess an alternative trans-port system capable of releasing K+ during stomatal closure.

Anion transport The efflux of K+ from guard cells duringstomatal closure is electrically neutralized by a concomitantefflux of anions. Anions pass through the guard cell membranevia channels that conduct a range of small anions (Hedrich& Marten, 1993; Schmidt & Schroeder, 1994). Two types ofchannels conducting anions have been identified in the guardcell membrane: rapid (R)-type (Keller et al., 1989; Hedrichet al., 1990); and slow (S)-type (Schroeder & Hagiwara,1989; Linder & Raschke, 1992) anion channels (Fig. 4). Themost obvious difference between these channels is a strongdecrease in the open probability of R-type channels whencells are clamped to potentials negative of −50 mV and a slowinactivation after returning to more positive potentials(Kolb et al., 1995), while S-type channels exhibit only a weakvoltage-dependent deactivation (Linder & Raschke, 1992;Schroeder & Keller, 1992; Dietrich & Hedrich, 1994).

Guard cells in intact plants often maintain membranepotentials negative of −100 mV (Roelfsema et al., 2001; 2002)and the difference in gating between S-type and R-type anionchannels may thus be of physiological importance. Anion-channel activation will normally depolarize the guard cellplasma membrane (see II. 3). However, in cells with a membranepotential negative of −120 mV, a depolarization cannot beinitiated by R-type channels. Indeed, we have observed guardcells in intact plants with a negative membrane potential anda high activity of R-type channels (own unpublished results).In these cells a depolarization can be initiated only throughchanges in the voltage dependence of R-type channels orthrough the activation of S-type anion channels. A change inthe voltage dependence of R-type anion channels can becaused by anions at the extracellular side of the channels.Organic anions, such as malate, acetate and proprionate, shiftthe half-maximal activation potential of R-type channels tomore negative values (Hedrich & Marten, 1993; Dietrich &Hedrich, 1998) and thus increase the potency of the channelto initiate a depolarization. The effect of malate is probablythe most significant, as this anion accumulates at high con-centrations in guard cells during stomatal opening (Outlaw &Lowry, 1977; Raschke & Schnabl, 1978). Anion channelsare permeable to malate (Hedrich et al., 1994; Schmidt &

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Schroeder, 1994) and thus will extrude this organic anion dur-ing stomatal closure. The apoplastic concentration of malatewas found to rise in response to high CO2 concentrations(Hedrich et al., 1994) and during prolonged illumination(Lohaus et al., 2001).

Apart from a difference in gating characteristics, R- andS-type channels also differ in their sensitivity to anion-channelblockers. R-type channels are blocked by stilbene derivates,such as DIDS (4,4′-diisothiocyanatostilbene-2,2′-disulfonicacid; Hedrich & Marten, 1993), while S-type channels areinsensitive to DIDS (Schwartz et al., 1995, own unpublishedresults). S-type channels are blocked by NPPB (5-nitro-2-(3-phenylpropylamio)benzoic acid), 9-AC (anthracene-9-carboxylicacid) and niflumic acid (Schroeder et al., 1993; Schwartzet al., 1995), but these blockers are not specific because theyalso inhibit R-type anion channels (Marten et al., 1992) andoutward-rectifying K+ channels (Garrill et al., 1996).

Apart from the difference in gating properties and theirsensitivity to stilbene derivatives, R- and S-type anion channelshave many features in common. Their single-channel con-ductance and ion selectivity are similar, which indicates thatboth channel types have a common pore structure or are justtwo gating modes of a single protein (Dietrich & Hedrich,1994). Both channel types were also found in A. thalianaguard cells (Pei et al., 1997; Pei et al., 2000); however, the genesencoding these plasma-membrane anion channels are stillunknown. Several genes homologous to animal ClC channelswere found in the Arabidopsis genome, but these genes probablyencode anion channels of intracellular membranes (Geelenet al., 2000; Lurin et al., 2000). In agreement with these results,ClC channels were not identified in purified plasma mem-branes of cell-suspension cultures of A. thaliana (Marmagneet al. 2004). Instead, voltage-dependent anion channel (VDAC)proteins were present in the plasma membrane, an anionchannel normally associated with the mitochondria. Futureexperiments need to provide unequivocal evidence for VDACproteins as genuine plasma-membrane anion channels thatharbor the properties of R- and S-type anion channels.

Compared to the anion-release system of the guard cellmembrane, little is known about the anion-uptake transport-ers. During stomatal opening guard cells accumulate anions,which electrically neutralize the uptake of K+. Anion uptakeis a flexible process that depends on the extracellular presenceof anions. As mentioned in II. 1, guard cells accumulate Cl– andmalate in equal concentrations when stomata, in epidermalstrips, are supplied with Cl– (Raschke & Schnabl, 1978).However, if no plasma membrane-permeable anions areapplied, K+ uptake is completely balanced by malate synthesis(Raschke & Schnabl, 1978). Other anions, such as NO3

– canalso support stomatal opening (Guo et al., 2003). Arabidopsismutants, with reduced levels of the NO3

− transporter, CHL1,displayed smaller stomatal apertures than wild type, whenNO3

− was supplied to detached leaves (Guo et al., 2003).Stomatal opening, however, was similar to that of wild-typeArabidopsis in the presence of Cl–, indicating that the CHL1transporter enables the uptake of NO3

− into guard cells, butdoes not represent the Cl– uptake system. Most likely, guardcells also possess a carrier that couples the uptake of Cl– withthat of H+, but this transporter still remains to be identified.

Ca2+ transport The interaction between extracellular K+ andCa2+ on stomatal movement has been studied extensively (seereviews Raschke, 1975a; MacRobbie, 1987). In general, highextracellular K+ and Ca2+ concentrations act antagonistically,as stomatal opening is stimulated by K+ and inhibited by Ca2+.The sensitivity to extracellular Ca2+, however, differs betweenspecies. Whereas the presence of 0.25 m extracellular Ca2+

reduced stomatal opening in C. communis by > 50% (DeSilvaet al., 1985a), it reduced stomatal apertures to < 25% in V. faba(Fischer, 1972) and A. thaliana (Roelfsema & Prins, 1995).The latter data reflect an effect of Ca2+ at an extracellular K+

Fig. 4 R- and S-type anion channels in the guard cell plasma membrane. (a) Current traces of R-type anion channels of a far depolarized guard cell in an intact Vicia faba plant (Roelfsema et al., 2001). The currents were elicited from a holding potential of −140 mV to test potentials ranging from −120 to 80 mV at 20-mV increments. (b) Current traces of S-type anion channels of a V. faba guard cell in an epidermal strip bathed in 5 mM CsCl, 1 mM CaCl2 and 1 mM Mes/BTP, pH 6.0. The currents were elicited from a holding potential of 20 mV to test potentials ranging from 0 to −180 mV, at −20-mV increments. Note the slow deactivation of S-type anion channels at the most negative membrane potentials. (c) Current–voltage relationship of the same cell as in (a) sampled during the last 100 ms of the test pulses. Note the maximum conductance at −80 mV. (d) Current–voltage relationship of the same cell as in (b) sampled during the last 500 ms of the test pulses. The reversal potential at 0 mV indicates an incomplete block of outward-rectifying K+ channels under the conditions applied.

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concentration of 50 m; at lower K+ concentrations the Ca2+

effects become even more apparent (Fischer, 1972). ExtracellularCa2+ probably inhibits stomatal opening by evoking increasesin the intracellular Ca2+ concentration (McAinsh et al., 1995).It has long been thought that high external Ca2+ concentrationswould enhance leak Ca2+ currents across the plasma membrane.Such leak currents of Ca2+ would be supported by the highconcentration gradient for Ca2+ across the plasma membrane.The apoplastic Ca2+ concentration is ≈ 100 µ (Felle et al.,2000; Roelfsema & Hedrich, 2002), while the cytoplasmicconcentration ranges from 100 to 300 n (Webb et al., 2001;Levchenko et al., 2005). Recently, it was shown that guard cellsdo not simply leak Ca2+ but instead possess a plasma-membraneCa2+ sensor (CAS) (Han et al., 2003). The CAS sensor triggersa rise in the intracellular Ca2+ concentration, in response to anincrease in the extracellular Ca2+ concentration. Guard cellsexpressing antisense constructs of CAS were no longerresponsive to a rise in the extracellular Ca2+ concentration.

Calcium may enter guard cells via nonselective Ca2+ chan-nels in the plasma membrane. These nonselective channelshave been found in a large number of plant cells and are, ingeneral, permeable to monovalent and divalent cations(Demidchik et al., 2002). In guard cell protoplasts, Ca2+-permeable channels were identified that are activated by stretch(Cosgrove & Hedrich, 1991) or ABA (Schroeder & Hagiwara,1990). Nonselective cation channels that activate uponhyperpolarization were suggested by Lohse & Hedrich (1992)(Fig. 5b,c). The physiological relevance of these channels in

guard cells became apparent after Grabov & Blatt (1998)showed hyperpolarization-induced rises in the cytoplasmicCa2+ concentration. Subsequent studies showed that thesechannels are stimulated by 50 µ H2O2 in A. thaliana (Peiet al., 2000) and through phosphorylation and ABA in V. faba(Hamilton et al. 2000; Köhler & Blatt, 2002).

The Ca2+ homeostasis in plant cells is maintained by Ca2+

transporters in the plasma membrane, vacuolar membraneand endoplasmic reticulum (ER) (Sanders et al., 2002). Theplasma membrane harbors P-type ATPases (ACAs) that cantransport Ca2+ against steep concentration gradients. TheseCa2+ ATPases are characterized by an autoinhibitory domainat the N-terminus that binds Ca2+/calmodulin (Sze et al.,2000; Axelsen & Palmgren, 2001). At high cytoplasmic Ca2+

concentrations the N-terminus can be released from the coreprotein, thereby activating the Ca2+ pump. The latter mecha-nism provides a negative-feedback system that automaticallycounteracts a rise in cytoplasmic Ca2+. Ca2+ regulates the ACApumps also via an additional pathway involving Ca2+-dependentprotein kinases (CDPK) (Sze et al., 2000). Genes encodingCa2+-permeable channels in the guard cell plasma membranehave not yet been cloned (Hetherington & Brownlee, 2004),but they may be homologous to the ionotropic glutamatereceptors (Lacombe et al., 2001).

H+ transport The guard cell membrane is energized by a P-type ATPase that translocates H+ from the cytoplasm to theguard cell wall (Fig. 5a). This proton pump is the only primaryactive transporter in the plasma membrane, apart from anumber of P-type ATPases that transport Ca2+ or heavy metals(Axelsen & Palmgren, 2001). The guard cell H+-ATPase hasa low activity at pH 7.5 and becomes more active at acidic pHvalues (Becker et al., 1993). This pH dependence indicates arole for the proton pump in maintaining H+ homeostasis.Guard cells maintain a cytoplasmic pH of 7.5–7.8 (Blatt &Armstrong, 1993; Grabov & Blatt, 1997); a shift to moreacidic values thus automatically activates the plasma membraneH+-ATPase. In addition, H+-ATPases are regulated throughan autoinhibitory domain at the C-terminus. This domain isreleased from the catalytic site, after binding of a 14-3-3protein, based on phosphorylation-dependent (Würtele et al.,2003), as well as phosphorylation-independent, mechanisms(Fuglsang et al., 2003). Binding of the phosphorylated C-terminus to 14-3-3 proteins is stabilized by fusicoccin, a fungaltoxin that hyperstimulates stomatal opening (Braunsgaardet al., 1998; Kinoshita & Shimazaki, 1999).

Vacuolar membrane transport A proportion of the ions thataccumulate in the guard cell during stomatal opening willremain in the cytoplasm, while the rest is sequestered in thevacuoles. Guard cells posses several small vacuoles that maybe interconnected to form a reticulum (Palevitz et al., 1981;Faraday et al., 1982). Swollen guard cells contain only few butlarge vacuoles that shrink and become fragmented during

Fig. 5 Electrical properties of Vicia faba guard cell protoplasts after elimination of the plasma membrane K+ and anion currents, redrawn after Lohse & Hedrich (1992). (a) Current–voltage relationship of the plasma membrane H+-ATPase measured in the whole-cell patch clamp mode. A voltage ramp was applied from −240 to 10 mV. (b) Voltage relationship of whole-cell currents, with ATP in the pipette and in the absence (white symbols) or presence (black symbols) of 5 mM LaCl3 in the bath solution. (c) Voltage relationship of whole-cell currents remaining after ATP depletion in the absence of LaCl3.

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stomatal closure (Diekmann et al., 1993). However, a detailedstudy of changes in vacuole size during stomatal movement hasnot been conducted. A vacuolar sequestration is of importancefor ions, such as Ca2+, that also act as second messengers.Vacuoles are an important intracellular store of Ca2+ and areprobably involved in the release of Ca2+ during signalingevents in guard cells (MacRobbie, 1998; Sanders et al., 2002;Hetherington & Brownlee, 2004). Furthermore, sequestrationof malate is important, as this anion negatively feeds back oncarbon fixation, by inhibition of phospho(enol)pyruvate (PEP)carboxylase (Schnabl & Kottmeier, 1984b; Raschke et al., 1988).Indeed, most of the malate in guard cell protoplasts of V. fabais located in the vacuole (Schnabl & Kottmeier, 1984a).

Although a number of channels, carriers and pumps at thevacuolar membrane have been identified (Fig. 3), the regula-tion of vacuolar membrane transport is poorly understood. Ingeneral, transport activities were studied using isolated vacu-oles, devoid of cytoplasmic components that regulate thesetransporters in intact cells. Recording techniques that meas-ure the activity of ion transporters at more physiological con-ditions are technically demanding, but will be essential for abetter understanding of ion transport regulation in the future.Likewise, little is known about the membrane potential of theguard cell vacuole. A potential difference of 20–50 mV, neg-ative at the cytoplasmic site, is often assumed (MacRobbie,1998), but no direct recordings of vacuoles in intact guardcells exist. Furthermore, the vacuolar membrane potentialpresumably changes during stomatal movements, but theextent to which changes in this potential contribute to ionrelease remains unknown.

K+ transport Three types of K+-permeable channels have beendetermined in the vacuolar membrane of guard cells (Allenet al., 1998). Based on differences in gating characteristics,these channels were classified as slow vacuolar (SV), fastvacuolar (FV) (Hedrich et al., 1986; Hedrich & Neher, 1987;Schulz-Lessdorf & Hedrich, 1995; Allen & Sanders, 1996)and vacuolar K+-selective (VK) channels (Ward & Schroeder,1994; Allen & Sanders, 1996). SV channels represent themost prominent ion conductance in the vacuolar membraneand slowly activate at depolarizing potentials (more positive atthe cytoplasmic side), conditions that allow K+ release fromthe vacuole. However, at large depolarization, the K+ fluxreverses and K+ is transported into the vacuole via SV channels(Ivashikina & Hedrich, 2005). FV channels are similar in thisrespect, as they also activate with depolarization, but theiractivation is much faster. The Ca2+ dependence of FV and SVchannels differs – FV are inhibited by Ca2+ concentrationsof > 0.1 µ (Hedrich et al., 1986; Allen & Sanders, 1996),whereas SV channels are inactive at low concentrations ofCa2+, but are activated by Ca2+ concentrations of > 0.5 µ(Hedrich & Neher, 1987). Both channels are cation permeableand thus may enable K+ release, probably under differentcytoplasmic conditions. VK channels represent the third type

of K+-selective channel in the vacuole (Ward & Schroeder,1994). These channels are nonrectifying and thus appearas K+-selective leaks that may enable K+ release or uptake atconditions that do not support the activation of SV- or FVchannels (Allen et al., 1998).

Vacuolar K+ channels may be encoded by the KCO (TPK )genes, as all of these channels, apart from TPK4, are targetedto the vacuolar membrane (Czempinski et al., 2002; D. Beckeret al., unpublished). The KCO channels have been renamedTPK (which stands for two pore domain K+ channels) (Beckeret al., 2004), with the exception of KCO3, which has only onepore domain. The pore domains of the TPK channel andKCO3 contain the GYGD motif, suggesting that they encodeion channels with a K+-selective conductance (Czempinskiet al., 1999). This was recently proven for TPK4, the soleArabidopsis TPK channel located in the plasma membrane(Becker et al., 2004). Whether or not this is also the case forvacuolar TPK channels is currently under investigation.

K+ ions are taken up into vacuoles against the vacuolar mem-brane potential, the transport therefore is probably mediatedby secondary active carriers. K+ uptake may be carried out by theNa+/H+ (NHX)-transporters, well known for their ability totransport Na+ (Gaxiola et al., 1999), but which also mediateH+-coupled K+ transport (Venema et al., 2003; Cellier et al.,2004). In addition, less well-characterized transporters arefound in the Arabidopsis genome; these transporters have highsequence similarity to known H+/K+ carriers (Sze et al., 2004).

Anion transport The vacuolar membrane potential supportsa flow of anions from the cytoplasm into the vacuole throughanion channels. For guard cell vacuoles, only a single publicationhas reported on anion channels (Pei et al., 1996). This anionchannel was reported to be activated by CDPK and tofacilitate the flow of Cl– and malate2– into vacuoles. For othercell types, voltage-dependent anion channels have been describedthat open at negative membrane potentials, indicating thatthey indeed support anion sequestration into vacuoles(Pantoja & Smith, 2002; Hafke et al., 2003). This may beimportant for the sequestration of malate, as the cytoplasmicconcentration of this anion has to be kept low in order toprevent the inhibition of PEPcarboxylase (Raschke et al.,1988; Schnabl & Kottmeier, 1984b). For malate, the uptakeprocess into the vacuole is complex, as malate has two negativecharges at neutral pH, but may lose one upon entry into theacidic vacuole. Monovalent malate anions could either betrapped in the vacuole, or flow back into the cytoplasm,depending on the selectivity of anion channels. Alternatively,malate may be transported into the vacuole by carriers, suchas the malate carrier, At tDT (Emmerlich et al., 2003).

Anions can also be released via anion channels, but thisrequires depolarization of the vacuolar membrane. If themembrane potential would approximate 0 mV, anions wouldflow down their concentration gradient into the cytoplasm.Vacuolar anion channels may be encoded by genes homologous

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to the bacterial and animal ClC channels. Disruption of theAtClC-a gene resulted in plants that accumulate less NO3

−

when grown at high levels of NO3− (Geelen et al., 2000).

However, the activity of ClC channels, in general, and their rolein the vacuolar membrane, in particular, still await confirmationby direct measurements. Patch clamp studies, on isolatedvacuoles of plants overexpressing a CLC channel or lackingone, may help to establish the function of these genes.

Ca2+ transport The large vacuole of guard cells plays animportant role in Ca2+ homeostasis (Sanders et al., 2002).Although the Ca2+ concentration of guard cell vacuoles hasnot been studied in detail, it is generally assumed that it is inthe m range, as recorded for rhizoid cells of Riccia fluitantsand root cells of Z. mays (Felle, 1988). Ca2+ accumulates inthe vacuoles as a result of the activity of ATP-dependent ACApumps (Sze et al., 2000; Axelsen & Palmgren, 2001) as wellas of the CAX-encoded Ca2+/H+ antiporters (Hirschi et al.,1996). The ACA pumps are regulated by Ca2+/calmodulin, asare their counterparts at the plasma membrane, while theregulation of the CAX carriers seems to be more indirectand involves several CXIP proteins that closely interact withCAX1 and -4 (Cheng & Hirschi, 2003).

In contrast to the limited information available about Ca2+

sequestration into the vacuoles of guard cells, many reportshave focused on Ca2+-release channels. Because of the steepconcentration gradient, the opening of Ca2+-permeable chan-nels in the vacuolar membrane results in a Ca2+ flux to thecytoplasm. This, in turn, leads to a rise in the cytoplasmicCa2+ concentration, which may encode a signal for downstreamresponses (Evans & Hetherington, 2001; Sanders et al., 2002).Based on patch clamp recordings, plant vacuolar membranesseem to harbor a number of ligand-gated Ca2+-permeablechannels. In beet roots, vacuolar membrane Ca2+ currents areinduced by inositol triphosphate (IP3) and cyclic-ADP ribose(Allen et al., 1995), while guard cell vacuolar membranes havealso been shown to be sensitive for cyclic-ADP ribose (Leckieet al., 1998) and inositol hexakis-phosphate (IP6) (Lemtiri-Chlieh et al., 2003). In addition to these ligand-gated Ca2+

channels, a voltage-gated Ca2+ channel (VVCa-Channel) hasbeen described for the guard cell vacuolar membrane (Allen& Sanders, 1994).

The vacuolar SV channel, initially described as a K+ channel(Hedrich et al., 1986), has been suggested to functionas a Ca2+-induced Ca2+-release (CICR) channel (Ward &Schroeder, 1994; Allen & Sanders, 1995; Pei et al., 1999;Sanders et al., 2002). The latter role of the SV-channels wasinvestigated by Pottosin et al. (1997), who found that the SVchannel was blocked by luminal Ca2+. Later studies of Pottosinet al., 2004) and Ivashikina & Hedrich (2005) also disagreewith a role of SV-channels in Ca2+-induced Ca2+ release.

The single-copy gene, AtTPC (Furuichi et al., 2001), is likelyto encode one of the Ca2+ channels in the vacuolar membrane,as its gene product was localized to this membrane by using a

proteomics approach (Carter et al., 2004). Genes homologousto the animal IP3- and ryanodine-receptors have not beenidentified in the Arabidopsis genome (Schwacke et al., 2003).Apparently, the genes encoding these receptors in plants havedeveloped independently from the animal counterparts, orhave diverged considerably from them during evolution.

H+ transport Two types of H+-translocating pumps – V-typeH+-ATPases (V-ATPases) and pyrophosphatases (PPases) –co-reside in the same vacuolar membrane (Hedrich et al.,1986; 1989; Sze et al., 1999). The V-type ATPase is a proteincomplex similar to the F1/F0-ATPases and consists of a largenumber of different subunits (Sze et al., 2002; Kluge et al.,2003). The V1 subcomplex catalyzes ATP hydrolysis androtates, during H+ translocation, with respect to the V0subcomplex in the membrane. The stoichiometry of thesepumps is variable: values were determined ranging from morethan 3 (Davies et al., 1994) to 1 H+ per hydrolyzed ATP(Müller & Taiz, 2002). A low stoichiometry is probablyimportant for very acidic vacuoles because a high protonmotive force can inverse the proton flux through V-typeATPases (Gambale et al., 1994). For guard cell vacuoles thestoichiometry may be higher, as these compartments containhigh concentrations of K+ and thus may not have such a lowpH. V-type ATPases are probably regulated by the redox stateas well as through changes in the cytoplasmic nucleotideconcentration (Kluge et al., 2003), but no information isavailable about this type of regulation in guard cells. Theimportance of these transporters in guard cell physiology isevident from studies on the det3 mutant (Schumacher et al.,1999). These mutants are affected in subunit C of the V-ATPase and display only 70% of the V-ATPase activitycompared to wild-type cells. Guard cells of det3 did not closein response to an increase in the extracellular Ca2+ concentrationor to the application of H2O2 (Allen et al., 2000). Theabsence of these responses was therefore linked to an alteredCa2+ homeostasis in det3 guard cells.

In addition to the V-ATPases, PPases in Arabidopsiscontribute to the generation of the electrochemical gradientacross vacuolar membranes (Drozdowicz & Rea, 2001).These relatively simple proteins use the hydrolysis of PPi topump H+ into the vacuolar lumen. The activity of thesepumps is highest in young growing tissues, in which PPi isavailable as a by-product of biosynthesis (Maeshima, 2000).Because the synthetic rate of macromolecules is probablyless pronounced in mature guard cells, it is unlikely thatPPases play a major role in energizing the guard cell vacuolarmembrane.

3. Coupling of ion fluxes

Transport of osmotically active compounds across the plasma-and vacuolar membranes is driven by H+-translocatingprimary transporters. Although both plasma- and vacuolar

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membranes also posses Ca2+-ATPases, the latter pumpsprobably serve to compensate for Ca2+ influx. This means thatguard cell transport of solutes against their concentrationgradient is predominantly fueled by H+-ATPases (Fig. 6). Theelectrochemical gradient generated (proton motive force)can be decomposed in an electrical potential and in a H+-concentration difference, both of which are used to drivesolute flow.

Stomatal opening During stomatal opening, the plasmamembrane potential of V. faba guard cells in intact plants is ≈−110 mV (Fig. 7a). The reversal potential of K+ is close to−75 mV and the membrane potential thus can drive the uptakeof potassium. K+ uptake is facilitated by inward-rectifying K+

channels, which activate at membrane potentials negative of

−100 mV (Roelfsema et al., 2001). For each K+ taken up,an H+ ion has to be extruded in order to keep the membraneelectrically charged. The H+ gradient can drive the uptakeof anions, such as NO3

− or Cl–, by an H+-coupled symportsystem (MacRobbie, 1987). The stoichiometry of the lattertransporter is not known for guard cells, but has been foundto range between 1 and 2 for other cell types (Sanders, 1980;Felle, 1994). In addition to carrier-mediated anion uptake,guard cells synthesize considerable amounts of malate(Raschke et al., 1988). Most malate will be stored in the acidicvacuole and thus will have a single negative charge. Synthesisof one malate anion and its transport to the vacuole will thusresult in a surplus of one H+ ion. The H+ ion will be extrudedfrom the guard cell to keep the cytoplasmic pH constant andenable the uptake of one K+ ion. This process is as efficient as

Fig. 6 Coupling of ion-fluxes during stomatal movements. During stomatal opening an electrochemical gradient across the plasma membrane is maintained by active H+-ATPases (1). Based on the hyperpolarized membrane potential (−110 mV), K+ can be taken up via inward-rectifying K+ channels (2). Cl– may be taken up from the apoplast by symport with H+ (3), while malate is synthesized from phosphoenolpyruvate and CO2 (4). V-type ATPases maintain the electrochemical gradient across the vacuolar membrane (5). Cl– can be transported along the electrical potential (≈ −40 mV) into the vacuole via anion channels (6). A malate carrier may be involved in maintaining low cytoplasmic malate levels (7), while K+ is probably taken up against the electrical potential, by an H+-driven antiporter (8). Stomatal closure is associated with efflux of K+ via channels (9), presumably causing a depolarization of the vacuolar membrane (Em ≈ 0 mV). In the event that the vacuolar membrane depolarizes, Cl– may be extruded via an anion channel (10). Activation of plasma-membrane anion channels enables an anion efflux along the electrical potential (11) and causes a depolarization (−50 mV). As a result of the depolarized plasma membrane, K+ can be released via outward-rectifying K+ channels (12).

Fig. 7 Light- and CO2-induced changes in the electrical properties of Vicia faba guard cells in intact plants. (a) Guard cell membrane potential hyperpolarizing in the light and depolarizing in the dark. The bar below the graph indicates the time-points at which the microscope lamp was switched on and off. (b) Guard cell hyperpolarizing in the absence of atmospheric CO2 and depolarizing again with 700 µl l−1 of CO2. The bar below the graph indicates when the CO2 supply was shut off (white area) and on (striped area). From Roelfsema et al. (2002). (c) Activation of an inward current measured in a guard cell clamped constantly to −100 mV after supplying CO2, as indicated by the bar below the graph (white area: 0 µl l−1 CO2, striped area: 700 µl l−1 CO2).

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K+ uptake in combination with Cl–, provided that the Cl–/H+

carrier has a stoichiometry of 1. If the stoichiometry of thelatter carrier is > 1, uptake of K+ in combination with Cl–

would thus require higher H+-extrusion rates at the plasmamembrane than with malate synthesis.

As an alternative to ion uptake, guard cells may accumulatesugars. Guard cell protoplasts of P. sativum take up ≈ 15 fmolof glucose per cell h−1 and, at lower rates, also fructoseand glucose (Ritte et al., 1999). These molecules are probablyimported via H+-coupled transport by carriers, such as thehexose transporter, STP1 (Stadler et al., 2003). The stoichi-ometry of the sucrose transporter, SUC1, was found to be onesucrose molecule per H+ (Carpaneto et al., 2005); if this sto-ichiometry would apply also for other sugar transporters,guard cells can take up one sugar molecule per H+ ion extruded.Sugar uptake thus seems to be less efficient than K+ uptakecombined with malate accumulation.

As discussed in section II. 2, most of the solutes taken upduring stomatal opening will be sequestered in the guard cellvacuole (Fig. 6). The vacuolar membrane potential is negativeat the cytoplasmic side, as for the plasma membrane, but nowall solutes are to be transported in the opposite direction.Anions can be transported via ion channels as long as the con-centration difference between the cytoplasm and the vacuoleis compensated by the membrane potential difference.Provided that the vacuolar membrane potential is −40 mVduring stomatal opening, the concentration of monovalentanions in the vacuole should be five times lower than in thecytoplasm. If the anion concentration difference exceeds thisvalue, anion transport into the vacuole is probably coupled toa flow of H+ ions to the cytoplasm. K+ ions are transportedinto the vacuole against the potential difference and thusare probably sequestered into the vacuole by an H+-coupledantiport mechanism. Finally, sugars may be taken up intothe vacuole by a recently identified subfamily of sugar trans-porters localized in this endomembrane and homologous toERD6 (M. Büttner, pers. comm.).

Stomatal closure In V. faba, stomatal closure can occur veryrapidly compared to stomatal opening. Whereas the transpira-tion rate increases with a velocity of 0.7 µmol m−2 s−2 in thelight, it decreases with a velocity of 1.9 µmol m−2 s−2 after theapplication of ABA (Langer et al., 2004). This difference invelocity probably correlates with the transport mechanismsinvolved in both processes. Whereas opening is based onactive transport, closure is based on the release of solutes alongtheir concentration gradients (Fig. 6). Stomatal closure thereforemay be dependent entirely on the activation of ion channelsin the vacuolar and plasma membranes. K+ ions will flow fromthe vacuole to the cytoplasm when K+ channels in the vacuolarmembrane are activated. The activation of K+ channels willalso depolarize the vacuolar membrane and provide a drivingforce for anion efflux. The anions will thus be extruded fromthe vacuole after the activation of anion-permeable channels.

However, it is unclear whether the events described abovereally occur during stomatal closure, as neither the activationof K+ and anion channels, nor the concurrent depolarizationof the vacuolar membrane, have, to date, been recorded.

During dark-induced stomatal closure, the plasma mem-brane of V. faba guard cells depolarizes slowly to ≈ −50 mV(Roelfsema et al., 2001; Fig. 7a). This value is positive of theK+-reversal potential and thus supports the efflux of K+ ionsvia outward-rectifying K+ channels. The dark-induced depo-larization is probably brought about by the deactivation of H+

ATPases in combination with the activation of S-type anionchannels (Roelfsema et al., 2001; 2002). An efflux of K+ viaplasma-membrane channels is thus accompanied by anionextrusion.

It is unclear how sugars are released from the vacuoleand cytoplasm of guard cells. In principle this could occur viasugar-permeable porins but, in Arabidopsis, porins are onlyassociated with mitochondrial and plastid membranes (Schwackeet al., 2003; Clausen et al., 2004). Alternatively, sugars may bereleased by transporters encoded by genes homologous tosugar carriers; such carriers may function in a reverse mode orfunction in a mode uncoupled from H+ translocation.

III. Regulation of ion transport in guard cells

Because of the close connection between ion transport andchanges in the osmotic content of guard cells, stomatalmovement can be traced back to guard cell ion transport.Regulation of ion transport in guard cells thus directly affectsthe leaf conductance, an important determinant for plantgrowth. Nevertheless, guard cells seem to respond more or lessautonomously to environmental signals. This is exemplifiedby the responses of stomata, in isolated epidermal strips,to signals such as light, ABA and CO2. Although the latterresponses mimic the behavior of stomata in intact plants,it has long been recognized that the responses measured inepidermal strips are different from those measured on intactleaves (Willmer & Mansfield, 1969). For instance, light inducesan increase in the transpiration rate of 0.7 µmol m−2 s−2

(Langer et al., 2004) in V. faba leaves, which correlates to anincrease in the stomatal aperture of ≈ 6.5 µm h−1 (Kappenet al., 1987). A much slower rate of opening, of ≈ 2.5 µm h−1,was found for stomata in epidermal strips (Schwartz & Zeiger,1984). Likewise, stomatal closure induced by ABA also occursmore slowly in epidermal strips than in intact leaves. Whereasstomata in intact leaves close with a rate of ≈ 18 µm h−1, basedon the transpiration measurements (Langer et al., 2004), orwith a rate of ≈ 21 µm h−1, as determined by microscopicalobservations (Roelfsema et al., 2001), a velocity of ≈ 8 µm h−1

was found with epidermal strips (Blatt & Armstrong, 1993).This indicates that guard cells, to some extent, lose theirresponsiveness after isolation, which may be caused by slowdedifferentiation in the absence of signals derived from viableneighboring cells. Note that the guard cells of tobacco and

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sugar beet can regenerate complete new plants (Sahgal et al.,1994; Hall et al., 1996). Apparently, guard cells in intactleaves are embedded in a niche, which retains them in a highlyspecialized state. The way in which mature guard cells areinfluenced by other cells within the leaf is not known in detail,but signal transduction pathways causing epidermal leaf cellsto develop into guard cells are probably involved (Bergmann,2004; Larkin et al., 2003; Bergmann et al., 2004).

The loss of guard cell responsiveness in isolated epidermalstrips has led to the development of techniques to study guardcells in the intact plant (Felle et al., 2000; Roelfsema et al.,2001). As expected, guard cell responses recorded using thelatter method differed from those observed with guard cellsin isolated epidermal strips and protoplasts thereof. Below wewill discuss the current knowledge about guard cell signaltransduction and relate these to electrophysiological data andtranspiration measurements. As mentioned in the first sectionof this review, the leaf conductance is mainly influenced bylight, humidity and circadian rhythms. The latter regulatorymechanism has been recently reviewed in this series (Webb,2003) and we will therefore only cover the effect of light andhumidity.

1. Light-induced stomatal opening

The nature of the light receptor in guard cells has been longdiscussed. At the beginning of the 20th century, plant biologistsrealized that stomata open at low CO2 concentrations andpostulated that light acts via a lowered CO2 concentration inthe leaf (Stahl, 1920; Heath, 1959). This hypothesis seemedto fit action spectra obtained for stomatal opening, whichare very similar to the absorption spectrum of chlorophyll(Kuiper, 1964). A closer examination, however, revealedthat blue light was more effective than red light (Sharkey &Raschke, 1981). In fact, blue light can still stimulate stomatalopening when provided on top of saturating intensities of redlight (Iino et al., 1985). This response to blue light can nowbe assigned to the activity of the PHOT1 and PHOT2 blue-light receptors located in the plasma membrane (Kinoshitaet al., 2001; Sakamoto & Briggs, 2002). Stomata of the phot1phot2 double mutant are no longer specifically responsiveto blue light; however, they still open in response tophotosynthetic-active radiation. Guard cells thus have twolight-perception pathways acting parallel to each other: thePHOT1/PHOT2-mediated pathway, which is dependenton blue light, and a second pathway that is dependent onphotosynthetic-active radiation (Assmann & Shimazaki,1999; Roelfsema et al., 2002).

Blue light-dependent pathway The PHOT receptors arereceptor kinases associated with the plasma membrane thatare activated by blue light (Briggs & Christie, 2002). The N-terminal half of the protein contains two LOV-domains thatbind flavin mononucleotides (Christie et al., 1999) and act as

light sensors, while the C-terminal half is a serine/threoninekinase. Blue light triggers autophosphorylation of PHOTreceptors, which leads to the binding of 14-3-3 proteins(Kinoshita et al., 2003). Binding of the 14-3-3 protein mayhelp to keep the PHOT receptors in an active state. A similarmechanism is found for the activation of plasma-membraneH+-ATPases, downstream in the blue light signal pathway. Here,the C-terminus forms an autoinhibitory domain that bindsthe catalytic site of the H+-ATPase (Braunsgaard et al., 1998;Fuglsang et al., 2003). Blue light leads to the phoshorylationof this domain and the binding of 14-3-3 proteins (Kinoshita& Shimazaki, 1999). This releases the C-terminus from thecatalytic site and activates the H+-ATPase.

Although it is tempting to speculate that the PHOT recep-tors directly phosphorylate the H+-ATPase, this has yet to beshown. The signaling pathway seems to be more complicated,as PHOT1-induced stomatal opening depends on a func-tional RPT2 protein (Inada et al., 2004). Based on homology,the RTP2 protein presumably mediates protein–proteininteractions. Furthermore, several lines of evidence indicatethe involvement of changes in the cytoplasmic Ca2+ concen-tration of guard cells (Shimazaki et al., 1999). The blue lightresponse of guard cells is inhibited by verapamil, a Ca2+-chan-nel blocker (Shimazaki et al., 1997), and is stimulated by highconcentrations of external Ca2+ (Roelfsema et al., 1998). Inseedlings of Arabidopsis, blue light induces a rise in the cyto-plasmic Ca2+ level, via a PHOT1-dependent pathway (Baumet al., 1999). Furthermore, PHOT1 and -2 stimulate theactivity of hyperpolarization-activated Ca2+ channels in theplasma membrane of mesophyll cells (Stoelzle et al., 2003).Blue light-activated Ca2+ channels, however, have not yetbeen shown for guard cells, and the effect of blue lighton the cytoplasmic Ca2+ concentration of guard cells is stillunknown.

The blue light-induced activation of H+-ATPases can berecorded as extracellular acidification by guard cell protoplasts(Shimazaki et al., 1986) or guard cells in epidermal strips(Roelfsema et al., 1998). Alternatively, this process can bemeasured as a blue light-stimulated outward current acrossthe guard cell plasma membrane (Assmann et al., 1985). Theextrusion of H+ drives the uptake of K+ via inward-rectifyingK+ channels, at a ratio of 1 : 1 (Section II, 3). In theory, themagnitude of the H+ current thus should be linked to thespeed of the stomatal opening induced by blue light. Contin-uous blue light triggers an increase in the stomatal conduct-ance of V. faba leaves of ≈ 0.08 (mol m−2 s−1) h−1 (Iino et al.,1985). Based on the results of Kappen et al. (1987), such achange in conductance is linked to an increase in stomatalopening of ≈ 4 µm h−1. For stomata in intact leaves this wouldrequire an increase in turgor pressure of ≈ 1.8 MPa h−1

(Table 2). In order to support such a change in the turgorpressure, guard cells should accumulate ≈ 1.4 pmol of K+ h−1

(Table 2), which requires an extrusion of ≈ 1.4 pmol of H+ h−1,correlating to an outward current of 36 pA. In guard cell

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protoplasts measured in the whole-cell configuration, bluelight induces an outward current of ≈ 6 pA (Assmann et al.,1985; Schroeder, 1988; Taylor & Assmann, 2001). Thecurrent measured in these patch-clamp studies is thus muchsmaller, as would be expected from the rate of stomatal open-ing. Loss of cytoplasmic components in the whole-cell modemay explain the relatively small blue light-induced currents, assuggested by the larger current measured in slow-whole-cellrecordings (Schroeder, 1988). In the guard cells of intact plants,the blue light-triggered current was even larger (Roelfsemaet al., 2001); here, on average, an outward current of 60 pAwas recorded 2 min after the onset of blue light. Continuousblue light triggered a transient response; the currents levelledoff after reaching a peak value within 3 min (Fig. 8c).

Photosynthetic-active radiation-dependent pathway Apartfrom the blue light-specific pathway, guard cells are alsosensitive to photosynthetic-active radiation (PAR). Thisis most obvious from the effect of DCMU (3(3,4-dichlorophenyl)-1-1-dimethylurea), an inhibitor of photosystemII, which inhibits red light-induced (Kuiper, 1964), but notblue light-induced, stomatal opening (Sharkey & Raschke, 1981;Schwartz & Zeiger, 1984). This response is often referred toas the red light response, but it can also be activated by bluelight at high intensities. For this pathway, the old hypothesis,postulating that intercellular CO2 may be an intermediatesignal, seems to apply. We found that guard cells in the intactplant did not respond to red light irradiation, given as a smallbeam directly on individual guard cells (Roelfsema et al., 2002).However, switching on and off red light over a large area onthe leaf did alter the plasma membrane potential of guardcells. The broad beam of red light was found to lower theintercellular CO2 concentration, while a small beam of redlight had no effect. CO2 thus probably functions as anintermediate signal in the PAR-dependent pathway.

It has long been debated to what extent guard cells canconduct photosynthesis (Outlaw, 1989). Although the chlo-rophyll content of guard cells is much lower than that ofmesophyll cells, chlorophyll fluorescence recordings indicatethat chloroplasts of guard cells and mesophyll cells do notdiffer much (Goh et al., 1999; Baker et al., 2001). Guard cellsthus seem to operate photosynthetic electron transport andCO2 fixation (Shimazaki et al., 1982; Wu & Assmann, 1993),enabling them to lower their own CO2 concentration, tosome extent. This may explain why guard cells in epidermalstrips open in red light. The diffusion rate of CO2 in bathsolutions covering the epidermal strip will be low, and guardcell photosynthesis may therefore have a significant effect onthe intracellular CO2 concentration. In intact plants, how-ever, guard cell photosynthesis will be less effective becauseguard cells have direct access to the atmosphere and the CO2diffusion rate is much higher.

Guard cells in intact plants respond to high CO2 concen-trations or the offset of red light, with the activation of anionchannels, which causes depolarization of the plasma mem-brane (Brearley et al., 1997; Roelfsema et al., 2002; Fig. 7b).As a result of this depolarization, the flux of K+ across theplasma membrane reverses. Instead of K+ uptake via inward-rectifying channels in the absence of CO2, K

+ is extruded viaoutward-rectifying channels in the presence of CO2. The acti-vation of anion channels is thus sufficient to enable both theefflux of anions and K+. As a result of the loss of osmolitesfrom guard cells, the stomata will close.

Various mechanisms have been suggested that couple a risein the CO2 concentration to changes in the activity of plasma-membrane ion channels (reviewed in Vavasseur & Raghavendra,2005). Hedrich et al. (1994) found that the apoplastic malateconcentration rises at elevated concentrations of CO2 and, inturn, can activate R-type anion channels in guard cells. Similar

Fig. 8 Abscisic acid (ABA) and blue light-triggered changes in the electrical properties of the plasma membrane of Vicia faba guard cells in intact plants. (a) Depolarization of a guard cell elicited by 10 µM ABA applied to an impaled guard cell via the leaf surface. The black bar below the graph indicates time of ABA application. From Roelfsema et al. (2004). (b) ABA-induced changes in the plasma membrane current of a guard cell constantly clamped to a holding potential of −100 mV. The black bar below the trace indicates the time-point of ABA application. From Roelfsema et al. (2004). Note that ABA triggers inward currents in two phases: in phase 1 a large (but transient) increase occurs, while a small (but steady) inward current is present in phase 2. (c) Change in plasma membrane current of a guard cell clamped to −100 mV, triggered by blue light given on a background of red light. The current trace is interrupted by short intervals, in which current-voltage profiles of the cell were recorded.

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observations were made by Raschke et al. (2003), using guardcells in epidermal strips. Alterations of ion-channel activity viaelevated Ca2+ concentrations have been suggested by Webbet al. (1996), while a role for the intracellular pH could beexcluded by Brearley et al. (1997). Finally, Goh et al. (2002)found that both photosynthesis and the activity of inward K+

channels are affected by cytoplasmic ATP (Goh et al., 2002).Photosynthesis and plasma membrane transport may thus becoordinated via changes in the cytoplasmic energy charge ofguard cells.

2. Humidity and water status

Drought stress acts antagonistically to light on stomatalmovement: it induces stomatal closure or inhibits openingto prevent excessive loss of water via the stomata. Stomatarespond directly to changes in humidity and to ABA releasedfrom roots upon the onset of drought stress. The latter signal-transduction pathway has been studied extensively, while thedirect response of guard cells to humidity has received muchless attention. It has been postulated that ABA is released frommesophyll cells during drought and subsequently inducesstomatal closure (Hartung et al., 1988). However, the findingthat the stomata of ABA-insensitive mutants still respondto humidity (Assmann et al., 2000) points to a more directresponse of guard cells to the air water content. Althoughguard cells are capable of responding to changes in airhumidity, they seem poorly constructed to monitor changesin the plant water status directly (Raschke, 1987). Changesin the latter will only cause small changes in the guard cellvolume because of the high osmotic pressure in these cells.Lowering the relative humidity of the air will affect the turgorpressure in guard cells, as well as in epidermal pavement cells.Because of the mechanical advantage of epidermal cells(Sharpe et al., 1987), stomata will initially open after a dropin air humidity, but stomatal closure sets in upon prolongedexposure times (Kappen et al., 1987; Mott et al., 1997).Closure of stomata, observed at low air humidity, apparentlyis not a simple consequence of turgor loss, but is a guardcell-controlled process.

Abscisic acid-induced stomatal closure Guard cells arelocated some distance from the root tips, the tissue that willfirst experience soil drought. The restricted ability of guardcells to monitor such changes in water status is overcome byABA-dependent signaling. Roots in drying soil release ABAinto the xylem stream, which functions as a signal for thestomata to close (Hartung et al., 2002; Wilkinson & Davies,2002). The guard cell receptor for ABA is unknown and itslocation within the guard cell is discussed controversially. Onedata set points to a receptor at the extracellular site of theplasma membrane (Anderson et al., 1994; Yamazaki et al.,2003), while another set of results indicates a localizationinside the cell (Allan et al., 1994; Schwartz et al., 1994;

Levchenko et al., 2005). Downstream of the presumed ABAreceptor, a large number of signal intermediates have beensuggested. We will not go into detail about ABA signaling, asthe receptor remains a black box and suggested signalingpathways have already been the topic of a number of recentreviews (Hetherington, 2001; Schroeder et al., 2001; Finkelsteinet al., 2002; Jones & Assmann, 2004). Instead, we will restrictthe discussion to a small number of intermediates that have beenlinked directly to the transcription-independent regulationof guard cell ion-transporter activity.

ABA-induced rises in cytoplasmic-free Ca2+ The regulationof ion transport in guard cells has been linked for a longtime to changes in the cytoplasmic Ca2+ concentration. Thefirst indications of Ca2+ as a second messenger came fromexperiments with C. communis, which showed that a highlevel of extracellular Ca2+ enhances the ABA inhibition ofstomatal opening (DeSilva et al., 1985a), while Ca2+-channelblockers had the opposite effect (DeSilva et al., 1985b).Further evidence was obtained with Ca2+-reporter dyes,which show that ABA can induce a rise in cytoplasmic Ca2+

(McAinsh et al., 1990; Gilroy et al., 1991). Finally, in guardcells of C. communis that were injected with the Ca2+ buffer,BAPTA (1,2-bis(2-aminophenoxy)ethane-N,N,N’,N’-tetraaceticacid), ABA did not evoke a rise in the cytoplasmic Ca2+

concentration and stomatal closure was prevented (Webb et al.,2001). Despite this evidence for the involvement of Ca2+ inABA action, a Ca2+-independent ABA signal pathway also seemsto exist in guard cells. Several publications have reported apopulation of stomata, in C. communis, that close in responseto ABA without detectable changes in the cytoplasmic Ca2+

concentration (McAinsh et al., 1990; Gilroy et al., 1991; Allanet al., 1994). This indicates that a rise in the cytoplasmic Ca2+

concentration of guard cells is not essential for stomatal closure.Rises in the cytoplasmic Ca2+ concentration were proposed

to be initiated via the ABA activation of hyperpolarization-activated plasma membrane Ca2+ channels (Grabov & Blatt,1998). These plasma-membrane channels were shownto activate through direct interaction with ABA in V. faba(Hamilton et al. 2000) and through the release of H2O2 inArabidopsis (Pei et al., 2000). Alternatively, ABA could induceincreases in the Ca2+ concentration through the stimulationof Ca2+ channels in intracellular membranes, gated by cADPribose (Leckie et al., 1998) or inositol phosphates (Staxenet al., 1999; Lemtiri-Chlieh et al., 2003); see also section II, 2.However, electrophoretic injection of these messengers, intoguard cells of intact V. faba plants, could not mimic ABAaction (Levchenko et al., 2005). Apparently, cADP ribose, IP3and IP6 cannot replace ABA concerning the activation ofanion channels.

ABA not only triggers single rises in the cytoplasmic Ca2+

concentration, it can also induce repetitive Ca2+ oscillations(Staxen et al., 1999; Allen et al., 2000). The oscillation fre-quency was reported to be signal specific and was suggested to

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enable guard cells to differentiate between signals leading eitherto stomatal opening or closure (McAinsh & Hetherington,1998; Allen et al., 2001; Hetherington & Brownlee, 2004).ABA-induced oscillations in cytoplasmic Ca2+ have an averageperiod of 10.3 min in Arabidopsis (Allen et al., 2001), whichis longer than the 5–10 min required for the initiation ofstomatal closure by ABA (Roelfsema et al., 2004). This takesthe frequency of Ca2+ oscillations as a signal that inducesstomatal closure into question. Nevertheless, the Ca2+ signaturemay play a role in other responses, such as the inhibition ofstomatal reopening after removal of ABA (Allen et al., 2001),or ABA-induced changes in gene transcription. Such a role forCa2+ oscillations has been described for gene regulation inanimal cells (Dolmetsch et al., 1998).

Evidence for ABA-induced Ca2+ oscillations was obtainedwith guard cells in epidermal strips, but guard cells in intactplants did not show ABA-induced changes in cytoplasmicCa2+ (Levchenko et al., 2005). This might indicate that Ca2+

oscillations reflect the inability of isolated guard cells tomaintain a steady Ca2+ concentration (own unpublished data).Note that Ca2+ oscillations in isolated guard cells can also occurspontaneously and that, on occasion, these oscillations areturned off by ABA (Klüsener et al., 2002). Apparently, Ca2+

changes triggered by ABA are heterogeneous in nature, andfuture research needs to explain how the variability of Ca2+

responses relates to stomatal function.

ABA regulation of plasma-membrane ion transport The mostprominent change in ion-transport activity at the plasmamembrane of guard cells is the activation of plasma-membrane anion channels (Thiel et al., 1992; Grabov et al.,1997; Pei et al., 1997; Raschke et al., 2003; Roelfsema et al.,2004). The activation of anion channels creates a pathway foranion efflux and thereby depolarizes the plasma membrane.In addition to anion-channel activation, ABA probablyalso reduces the activity of plasma-membrane H+-ATPases(Goh et al., 1996; Zhang et al., 2004), which would favor apronounced depolarization. As a result of plasma-membranedepolarization, K+ ions will be extruded from the guard cellsvia outward-rectifying channels (Thiel et al., 1992; Roelfsemaet al., 2004). Based on the biophysical properties of theplasma membrane, the activation of anion channels is thussufficient to induce an extrusion of anions as well as K+.Nevertheless, ABA was also found to regulate plasma-membrane K+ channels directly. The maximal conductanceof inward K+ channels was found to be reduced by ABA, inguard cells of epidermal strips (Blatt & Armstrong, 1993) aswell as in protoplasts thereof (Schwartz et al., 1994). ABA wasalso reported to stimulate outward-rectifying K+ channels inguard cells of isolated epidermal strips (Blatt et al., 1990), butthis could not be confirmed by measurements on guard cellsin intact plants (Roelfsema et al., 2004). In the discussion ofK+-channel regulation by ABA, it is often forgotten that thesechannels in the first instance are regulated by the membrane

potential. In most cells, ABA will depolarize the plasmamembrane (Fig. 8a) to values at which the outward K+

channel becomes active, but at which the inward K+ channelis no longer activated (Roelfsema et al., 2004). In thesecells, the reduction in maximum conductance of inwardK+ channels will thus have no physiological implications.

In guard cells of intact plants, ABA activates plasma-mem-brane anion channels in a transient manner; anion currentspeak after ≈ 5 min and decay thereafter (Fig. 8b; Roelfsemaet al., 2004; Levchenko et al., 2005). However, residualanion-channel activity remains in the presence of ABAand the current only returns to prestimulus values after ABAremoval. The first phase of this response may thus lead torapid stomatal closure, while the second phase will mainlyprevent the reopening of stomata. The two phases may explainthe differences recognized between ABA induction of stomatalclosure and ABA inhibition of stomatal opening.

It has long been postulated that ABA stimulates anionchannels through a Ca2+-dependent pathway, based on thefollowing observations: ABA can induce rises in the cytoplasmicCa2+ concentration, and both R- and S-type anion channelsare activated by cytoplasmic Ca2+ (Schroeder & Hagiwara,1989; Hedrich et al., 1990). Surprisingly, the activation ofanion channels in guard cells of intact plants occurred withouta concomitant rise in the cytoplasmic Ca2+ concentration(Levchenko et al., 2005). The response was not fully Ca2+

independent, as it was repressed after injection of the Ca2+

buffer, BAPTA. Apparently, Ca2+ signals are not essential forABA activation of anion channels, in agreement with previousresults described for ABA on anion channels, using patch-clamped protoplasts (Schwarz & Schroeder, 1998). A verysimilar situation exits for ABA inhibition of inward-rectifyingK+ channels. These channels display a steep Ca2+ dependence(Grabov & Blatt, 1999), but inhibition of the inward K+

channel by ABA in V. faba protoplasts occurred in a Ca2+-independent manner (Romano et al., 2000). This leads to theconclusion that both anion- and inward K+ channels are sen-sitive to cytoplasmic Ca2+, but are predominantly regulated byABA via a Ca2+-independent pathway.

IV. Interaction between guard cell signaling pathways

The signal pathways for light and humidity have been discussed,so far, as linear chains of events that couple environmentalchanges to a response at the plasma or vacuolar membrane. Innature, however, signals from the environment are seldomso discrete. On a hot summer day, for instance, high lightintensities will often coincide with low humidity. The firstsignal will cause stomata to open, while the second inducesstomatal closure. Guard cells therefore must be able to processthese signals in such a way that a collision of opposingresponses is prevented. The models of guard cell signalingshould also explain why stomatal opening is depressed at

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midday and how stomata of shaded plants in the canopyrespond to sudden sunflecks. Several mechanisms may functionin guard cells to obtain the optimal response with multiplechanges in the environment. Downstream in these pathways,various responses may differ in timing, while upstream signalpathways could converge at certain stages.

1. Timing of guard cell responses

Guard cells in intact plants display responses at the plasmamembrane with a specific timing, depending on the stimulusreceived. This is most obvious for ABA action where, after alag time of ≈ 2 min, ABA induces an inward anion currentthat peaks after ≈ 5 min and than drops to a residual level after≈ 10 min (Roelfsema et al., 2004; Fig. 8b). As discussed insection III. 2, the initial large inward current will cause a rapidclosure of stomata, while the residual anion channel activitymay prevent stomata from reopening. During the secondphase of the ABA response, stomata may become responsiveto light again, although the degree of stomatal opening will besmaller, as in the absence of ABA (Leymarie et al., 1998). Thespecific timing of the ABA response enables plants to respondquickly to drought without completely losing sensitivity toother environmental factors.

Although the timing of ABA responses in guard cells isrobust, the magnitude of responses can differ considerably.With repetitive ABA applications, we found that the respon-

siveness of single guard cells to ABA can change in time(Roelfsema et al., 2004). This heterogeneity in responsivenesscould be beneficial for plants under field conditions, aspointed out by Kaiser & Kappen (2001). During drought, theABA level in the xylem will slowly rise and because of differ-ences in responsiveness it will completely close some stomata,while others will remain open, to some extent. Upon stomatalclosure, transpiration will decrease and less ABA will reach thestomatal complex. This may create a simple feedback systemthat does not involve communication between guard cells andthe root. Such a communication would be very difficult in talltrees because of the large distance the signal would need totravel. In favor of such a feedback system is the observationthat guard cells monitor the amount of ABA reaching thestomatal complex, instead of the ABA concentration (Trejoet al., 1995).

Another feedback system regulating stomatal movementinvolves change in the intracellular CO2 concentration(Roelfsema et al., 2002, Fig. 9). High CO2 concentrationsinduce stomatal closure, which, in turn, lowers CO2 uptakeand thus decreases the intracellular CO2 concentration. Incontrast to the ABA response, the guard cell response to CO2sets in slowly and anion channel activation does not reach amaximum within 15 min (Fig. 7c). This may not apply forguard cells in epidermal strips, as, in this preparation, a peakanion channel activity was recorded 2 min after CO2 applica-tion (Brearley et al., 1997). The absence of a specific timing of

Fig. 9 Schematic presentation of signal pathways leading to light-induced stomatal opening. (a) Epidermal pavement cell, (b) guard cell, (c) mesophyll cell. Photosynthetic-active radiation (red light) reduces the CO2 concentration in the leaf, thereby deactivating anion channels in guard cells. Blue light acts on the PHOT1 and 2 receptors in the plasma membrane and activates the H+-ATPase. Both pathways lead to hyperpolarization of guard cells and provoke K+ uptake, causing the guard cell to swell and the stomatal pore to open. Note that opening of the stomatal pore increases the influx of CO2, which, in turn, can activate guard cell anion channels and induce stomatal closure. The pathway involving CO2 thus provides a negative feedback mechanism.

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the CO2 response, in intact plants, points to the role of CO2in a feedback loop. This response may continue until the CO2concentration in the substomatal cavity decreases to such anextent that the response automatically shuts off (Fig. 9).

Guard cells rapidly respond to blue light, the membranepotential hyperpolarizes after ≈ 60 s (Roelfsema et al., 2001),but the rise in plasma membrane current starts earlier (Fig. 8c).The outward H+-ATPase current peaks 3 min after the onsetof blue light, in agreement with a maximal degree of phos-phorylation of plasma membrane H+-ATPases (Kinoshita &Shimazaki, 1999). Just as with ABA, blue light will causemaximal change in stomatal movement several minutes afterstimulus onset. After prolonged stimulation with blue light,the outward H+ current decays to a steady-state level, suffi-cient to cause prolonged stomatal opening with time.

2. Cross talk of signaling pathways

Cross talk of signaling pathways in plant cells is an emergingtopic in plant biology (Taylor & McAinsh, 2004) and mayexplain the ability of sessile organisms to adapt to changingenvironments. As a result of cross talk, one signal may alter theguard cell’s responsiveness to another signal. Such an interactionhas been proposed for CO2 and ABA (Raschke, 1975b), ABAand blue light (Goh et al., 1996; Roelfsema et al., 1998),and for blue light and CO2 (Assmann, 1988). High CO2concentrations can increase the ABA responsiveness, whileABA, in turn, decreases the blue light responsiveness of thestomata. Finally, the blue light responsiveness is enhancedat low intercellular CO2 concentrations. It is not difficult toimagine what sort of complexity emerges if the interactionof multiple signals are taken into account. Instead of testingvarious signal combinations on different plant species, itmay be more fruitful to search for molecular determinantsat which several signal pathways converge.

Interaction of signaling may occur downstream in thepathways, as distinct signals can regulate the same ion trans-porters. For instance, both ABA and CO2 activate anionchannels, which may explain their synergistic mode of actionconcerning stomatal closure (Raschke, 1975b). Cross talk mayalso occur, to a greater extent, upstream and involve commonregulatory proteins or second messengers. The search for suchinteractions is assisted by the signaling mutants available forArabidopsis. For instance, the stomata of abscisic acidinsensitive (abi1) mutants were found to open more slowlyin response to light than those of wild-type plants (Eckert &Kaldenhoff, 2000). This indicates that ABA signaling inter-acts with the blue light and PAR-response pathways in guardcells.

Many other interactions have also been suggested for theABA signal pathway. ABA signaling may be linked to salttolerance as well as to cold acclimation (Chinnusamy et al.,2004). Furthermore, a close correlation was found betweenthe ABA- and sugar-sensing pathways (Finkelstein & Gibson,

2002). Note, however, that these interactions were recognizedfor whole plants or seedlings, but were not always confirmedfor guard cells. The interconnections found may hold forsome ABA responses, but not necessarily for all. For instance,the ABI4 gene is important for ABA and sugar signaling inseeds, but the gene does not seem to play a role in the regula-tion of stomatal movement. Another mutation (abi8) thataffects both ABA and sugar signaling does interfere with theABA inhibition of stomatal opening (Brocard-Gifford et al.,2004). Apparently it is hard to generalize interconnectionsof pathways. After all, from gene-expression studies we knowthat different cell types are equipped with different hardware,which makes it unlikely that the same software will run in allcell types. Differences in pathways for specific responses, celltypes and species therefore should be taken into account.

V. Outlook

Guard cell research has progressed a long way, from the initialmicroscopic observations of stomatal apertures by Moldenhauer(1812), to the specific regulation of single ion channels. Onthis way, however, it was often forgotten to reflect on whetherthe changes measured in transporter activity had the properamplitude, direction and kinetics, in view of the dramaticchanges in osmotic content of guard cells during stomatalmovement. The calculations in this review indicate that bluelight-induced changes in ion fluxes, recorded for guard cells inthe intact plant, fit changes in the ion content of guard cellsduring stomatal movement. In general, however, the ion fluxesand changes in solute concentrations recorded are somewhatsmaller than the changes calculated in solute concentration.This may point to the involvement of additional transportprocesses, such as the accumulation of sugars. With theidentification of the first sugar transporters in guard cells,these nonionic molecules will come into focus again.

Regulation of stomatal movement by some signals from theenvironment is partially understood at the molecular level, suchas the blue light-dependent activation of plasma-membraneH+-ATPases. For this pathway, both the receptors and thedownstream responses have been identified (Kinoshita &Shimazaki, 1999; Kinoshita et al., 2001) and fit the changesin transport activity measured directly on single guard cells(Roelfsema et al., 2001; Taylor & Assmann, 2001). Identifi-cation of the signaling components that interconnect thereceptor with the H+-ATPase may shed light on the way thatthe blue light signaling is affected by other signals, such asABA. Compared to the blue light-dependent pathway, little isknown about the PAR (CO2)-dependent pathway. Recently,stomatal movement was shown to be related to ascorbatesynthesis (Chen & Gallie, 2004), which indicates that thecytoplasmic redox state could serve as a switch, initiating thisresponse. Compared to the light responses of guard cells, ABAsignaling has received overwhelming attention. A large numberof mutants have been isolated, which are either hyper- or less

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sensitive to the stress hormone (Schroeder et al., 2001;Finkelstein et al., 2002). Despite the attempt to put the mutatedgene products in hierarchal schemes, a common model forABA signal transduction has not yet been obtained. The factthat, in some cases, data have been integrated, irrespective ofthe plant species, cell type or response studied, may haveadded to the degree of complication. Putting the signal com-ponents in place will be a major task for plant cell biology. Inthis context, fast electrical responses and more slowly emerg-ing changes in gene-expression rates should be decomposedfor given cell types. Guard cells will continue to be one ofthe systems of choice, because of the possibility to measureresponses for defined target proteins, within seconds up tohours after stimulus onset. We can thus expect exciting newinsights into ‘the Watergate’, soon again.

Acknowledgements

We wish to thank D. Geiger, V. Levchenko and H. Marten(University of Würzburg) for providing data for the figures,S. Liebig for the drawing in Fig. 1, and I. Marten (University ofWürzburg) for help with the preparation of the manuscript.This work was supported by grants from the DeutscheForschungsgemeinschaft to R.H.

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