Blackwell Publishing Ltd Tansley revie€¦ · Blackwell Publishing Ltd Tansley review Guard cell photosynthesis and stomatal function Tracy Lawson Department of Biological Sciences,

Review

© The Author (2008). New Phytologist (2009) 181: 13–34 13Journal compilation © New Phytologist (2008) www.newphytologist.org 13

Blackwell Publishing Ltd

Tansley review

Guard cell photosynthesis and stomatal function

Tracy LawsonDepartment of Biological Sciences, University of Essex, Wivenhoe Park, Colchester CO4 3SQ, UK

Contents

Summary 13

I. Introduction 14

II. Osmoregulation in guard cells 16

III. Role of guard cell chloroplasts in stomatal function 18

IV. Chlorophyll a fluorescence studies to examine guard cell photosynthesis 22

V. Linking stomatal behaviour to mesophyll photosynthesis 23

VI. Stomata in relation to water use/manipulation of behaviour 26

VII. Concluding remarks and future direction 27

Acknowledgements 28

References 29

Author for correspondence:Tracy LawsonTel: +44 (0) 1206 873327Fax: +44 (0) 1206 873416Email: [email protected]

Received: 4 June 2008Accepted: 18 September 2008

Summary

Chloroplasts are a key feature of most guard cells; however, the function of theseorganelles in stomatal responses has been a subject of debate. This review examinesevidence for and against a role of guard cell chloroplasts in stimulating stomatalopening. Controversy remains over the extent to which guard cell Calvin cycleactivity contributes to stomatal regulation. However, this is only one of four possiblefunctions of guard cell chloroplasts; other roles include supply of ATP, blue-light signallingand starch storage. Evidence exists for all these mechanisms, but is highly dependentupon species and growth/measurement conditions, with inconsistencies betweendifferent laboratories reported. Significant plasticity and extreme flexibility in guardcell osmoregulatory, signalling and sensory pathways may be one explanation. The useof chlorophyll a fluorescence analysis of individual guard cells is discussed in assessingguard and mesophyll cell physiology in relation to stomatal function. Developmentsin transgenic and molecular techniques have recently provided interesting, albeitcontrasting, data regarding the role of these highly conserved organelles in stomatalfunction. Recent studies examining the link between mesophyll photosynthesis andstomatal conductance are discussed. An enhanced understanding of these processesmay be fundamental in generating crop plants with greater water use efficiencies,capable of combating future climatic changes.

New Phytologist (2009) 181: 13–34 doi: 10.1111/j.1469-8137.2008.02685.x

Key words: Calvin cycle, guard cells, light responses, metabolism, osmoregulation, photosynthesis, stomata.

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Abbreviations: 3-PGA, 3-phosphoglycerate; ATP, adenosine-5-triphosphate; Ci,intercellular CO2 concentration; DCMU, 3-(3,4-dichlorophenyl)-1,1-dimethylurea;DHAP, dihydroxyacetone phosphate; F′, steady-state fluorescences; ,quantum efficiency of photosystem II (PSII) photochemistry; MAP, Mehler-ascorbateperoxidase; NADPH, nicotinamide adenine dinucleotide phosphate; OAA, oxaloac-etate; PEPc, phosphoenolpyruvate carboxylase; PGA, 3-phosphoglyceric acid;Rubisco, ribulose-1,5-bisphosphate carboxylase/oxygenase; RuBPC, ribulose-1,5-bisphosphate carboxylase.

I. Introduction

Stomata are small adjustable pores found in large numbers onthe surface of most aerial parts of higher plants and have beendocumented in the fossil record from as early as the lateSilurian, 411 Myr ago (Edwards et al., 1992, 1998). A stoma isformed from two specialized cells in the epidermis (guard cells)which are morphologically distinct from general epidermalcells and are responsible for controlling stomatal aperture(Franks & Farqhuar, 2007). Paired guard cells, in some speciestogether with epidermal subsidiary cells, form the stomatalcomplex (Fig. 1). Subsidiary cells can play a role in stomatalmovements either mechanically or as ion reserves (Raschke &Fellows, 1971). In most plants stomata can be found on boththe upper (adaxial) and lower (abaxial) leaf surfaces, such leavesbeing termed amphistomatous, with the majority of stomatafound on the lower surface (Tichà, 1982). In some species(particularly trees) stomata are found only on the lowersurface (i.e. the leaf is hypostomatous), whilst some aquaticplants (such as water lilies) have stomata only on the upper surface(i.e. the leaf is epistomatous) (Morison, 2003). As the leaf cuticleis almost impermeable to water and CO2, the central role of

stomata is regulation of gas exchange between the inside of theleaf and the external environment (Cowan & Troughton, 1971;Jones, 1992). Through their role in controlling transpiration,stomata also aid in leaf cooling, metabolite fluxes, and long-distance signalling (Brownlee, 2001; Lake et al., 2001; Jia &Zhang, 2008) as well as acting as a barrier to harmful pollutantssuch as ozone and pathogens (Meidner & Mansfield, 1968;Mansfield & Majernik, 1970).

Plants require sufficient CO2 to enter the leaf for photosyn-thesis whilst conserving water to avoid dehydration andmetabolic disruption. When fully open, stomatal pores onlyoccupy between 0.5 and 5% of the leaf surface (Hetherington& Woodward, 2003; Morison, 2003); however, almost all thewater transpired as well as CO2 absorbed passes through thesepores. For this reason, stomatal function has significant impli-cations for global hydrological and carbon cycles. The quest tounderstand stomatal control of photosynthetic CO2 fixationand plant water relations is becoming increasingly importantwith changing climatic conditions. Knowledge of stomatalfunction is critical to determine plant responses to environmentalstresses, particularly reduced water availability, and is necessaryto identify plants with decreased water use that are capable ofhigh yields in more extreme environments (Morison et al.,2007). On a global scale, drought causes more yield lossesthan any other single biotic or abiotic factor (Boyer, 1982),resulting in increasing pressure for agronomists and plantbreeders to identify crop varieties that are drought tolerant forsustainable production of food and biofuels on drought-susceptible land. Increased knowledge of stomatal functioncould provide the key to such crop improvements (Jones, 1987;Wang et al., 2007).

The aims of this review are to examine some of the potentialfunctions of the guard cell chloroplasts and how these arelinked to stomatal behaviour. Contrasting evidence has led toa number of controversies regarding guard cell chloroplastfunction, in particular concerning the contribution of guardcell photosynthesis to stomatal regulation, and several opposingviews are discussed in the following sections. Whilst there arestill gaps in our knowledge regarding stomatal regulation, aswell as sensory and signalling mechanisms, a wealth of evidenceexists on stomatal responses to various environmental stimuli,guard cell osmoregulation and mechanisms of movement. A

′ ′F Fq m/

Fig. 1 Stomatal complex illustrating the pair of guard cells, complete with chloroplasts and subsidiary cells in which calcium oxalate crystals are often found. Calcium oxalate is believed to play a role in regulation of stomatal aperture (see Ruiz & Mansfield, 1994).

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historical account of the various osmoregulatory pathwaysand solutes used in stomatal movements is given, whichdemonstrates the extreme plasticity in guard cell functionand illustrates the difficulties faced by stomatal researchersattempting to elucidate specific stomatal mechanisms. Thereview also briefly examines the use of recent advancements inmodern techniques (such as antisense technology and the useof DNA mutation) to address the function of chloroplastswithin guard cells. Recent studies using such techniques havealready revealed interesting and unexpected results, pavingthe way for future research, which may allow us to fill gaps inour understanding of the role of the guard cell chloroplasts instomatal regulation.

1. Stomatal regulation

Despite the controversies mentioned above, decades of researchhave provided a substantial amount of information regardingstomatal responses to various environmental stimuli. The detailsof these responses, along with mechanisms of pore opening andclosing, are now widely accepted. Stomatal aperture is regulatedby both internal physiological and external environmental factors(Farquhar & Sharkey, 1982; Morison, 1987; Mansfield et al.,1990; Hetherington & Woodward, 2003; Buckley, 2005)and can respond in time-scales of seconds to hours (Assmann &Wang, 2003). In general, pore opening through guard cellmovements is stimulated by illumination with light in thephotosynthetically effective waveband, low CO2 concentrationsand high humidity, whilst closure is promoted by darkness, lowhumidity, high temperature and high CO2 concentrations(see reviews by Assmann, 1993; Willmer & Fricker, 1996;Outlaw, 2003; Vavasseur & Raghavendra, 2005; Shimazakiet al., 2007) as well as plant hormones such as abscisic acid(see review by Weyers & Paterson, 2001). However, thereare exceptions, the most obvious being crassulacean acidmetabolism (CAM) plants which in general maintain closedstomata during the light period and open stomata in darkness(Osmond, 1978; see Black & Osmond, 2003). Additionally,stomata of Gunnera (Osborne, 1989) and Lemna spp. (Parket al., 1990) are unresponsive to a number of environmentalstimuli, whilst there are also examples of several other speciesthat do not respond to numerous plant hormones (e.g. Ridolfiet al., 1996; see Weyers & Paterson, 2001).

It is recognized that stomatal responses to light have at leasttwo components. One component is the photosynthesis-independent, specific blue-light response that saturates at lowfluence rates, and is often associated with rapid stomatal opening(see Zeiger et al., 2002) believed to involve the activation ofa plasma membrane H+-ATPase (Kinoshita & Shimazaki,1999; Shimazaki et al., 2007). The other component, aphotosynthesis-mediated response (termed the red-light responsehere and in many other publications), saturates at high fluencerates similar to those that saturate guard and mesophyll cellphotosynthesis and is inhibited by 3-(3,4-dichlorophenyl)-

1,1-dimethylurea (DCMU, an inhibitor of photosystem II(PSII)), indicating that it is photosynthesis-dependent (e.g.Kuiper, 1964; Sharkey & Raschke, 1981a; Tominaga et al.,2001; Olsen et al., 2002; Zeiger et al., 2002; Messinger et al.,2006) and suggesting that chlorophyll is the receptor (Assmann& Shimazaki, 1999; Zeiger et al., 2002). This photosynthesis-dependent response can be observed under either blue or redlight capable of driving photosynthesis (Sharkey & Raschke,1981a) and is often believed to operate through mesophyll-drivenconsumption of CO2 reducing the internal CO2 concentration(Ci) (Roelfsema et al., 2002, 2006), to which stomata are knownto respond (Mott, 1988). However, there is also evidencefor a direct guard cell red-light response, independent ofmesophyll photosynthesis, as discussed in Section V.

Stomatal opening is brought about by the accumulation ofions and/or solutes (Imamura, 1943; Fujino, 1967; Outlaw &Manchester, 1979; Outlaw, 1983) in guard cells, which increasesthe osmotic potential, thus lowering the water potential,causing water uptake from the apoplast (Weyers & Meidner,1990; Willmer & Fricker, 1996). Increases in guard cell volumeand hence turgor pressure widen the stomatal pore (Franks &Farqhuar, 2007; Shimazaki et al., 2007). Closure is broughtabout by the reverse, a loss or release of solutes, accompaniedby a loss of water and consequently turgor pressure. Both poreopening and closure are energy-dependent processes (Willmer& Fricker, 1996).

However, our understanding of the perception of, andprecise response of stomata to, different environmental stimuliis not complete. The fact that stomata in isolated epidermalpeels respond to various environmental factors suggests thatpart of the sensory mechanisms is located in the epidermis(Willmer & Fricker, 1996; Frechilla et al., 2002).

It is also well established that there is a strong positivecorrelation between stomatal conductance and mesophyllphotosynthesis (e.g. Wong et al., 1979; Zeiger & Field, 1982),and a close correlation between photosynthetic efficiencies inguard and mesophyll cells has also been observed (Lawsonet al., 2002, 2003). The majority of guard cells have chloroplasts,and these would therefore provide an ideal and convenientlocation for sensory or regulatory mechanisms. Although guardcell chloroplasts are a characteristic feature of most plants, therole of these highly conserved organelles in osmoregulation andtheir importance in stomatal function largely remain unclear.

2. Guard cell chloroplasts

In most species studied, guard cells contain chloroplasts,which vary in number depending upon the species (Willmer& Fricker, 1996; Lawson et al., 2003; Fig. 2). Most speciestypically contain 10–15 chloroplasts per guard cell (Humble& Raschke, 1971), compared with 30–70 in a palisademesophyll cell. However, numbers of chloroplasts per guardcell range from 3–6 in Selaginella (Allaway & Milthorpe,1976) to up to 100 in Polypodium vulgare (Stevens & Martin,

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1978), and guard cells of Paphiopedilum species entirely lackchloroplasts (Nelson & Mayo, 1975; Rutter & Willmer,1979; D’Amelio & Zeiger, 1988) but still maintainfunctional stomata (Nelson & Mayo, 1975). Guard cellsare formed from epidermal cells, which notably also lackchloroplasts (again there are exception such as Polypodiumspecies; Fig. 2).

Guard cell chloroplasts are often smaller, with less granalstacking, and some are less well developed than those inmesophyll cells (Sack, 1987; Shimazaki & Okayama, 1990),although these features vary across plant families (see reviews byPemadasa, 1981; Willmer & Fricker, 1996). Another noticeablefeature of most guard cell chloroplasts is that starch accumulatesin the dark and disappears in the light (Willmer & Fricker,1996), the reverse of the situation in mesophyll cells. However,this may not be the case for all species, as work by Stadleret al. (2003) has revealed that guard cells of Arabidopsis are

practically free of starch in the morning and accumulatestarch during the day.

Before examining guard cell photosynthesis and its possiblerole in stomatal behaviour, including the controversial topicof guard cell Calvin cycle activity, it is essential to provide abrief account of the osmoregulatory pathways that occur inguard cells.

II. Osmoregulation in guard cells

Many decades of research have focused on the osmoregulatorymechanisms found in guard cells. To put this review intocontext, a brief history of stomatal osmoregulation is given inthis section; however, this is by no means exhaustive (I referreaders to the following comprehensive reviews: Talbott &Zeiger, 1998; Zeiger et al., 2002; Outlaw, 2003; Roelfsema &Hedrich, 2005; Vavasseur & Raghavendra, 2005).

Fig. 2 Images of stomata from intact leaves. A reflected light image from Commelina communis (a) and steady-state fluorescence images from Commelina communis (b), Vicia faba (c), Nicotiana tabacum (d), Polypodium vulgare (e). Chlorophyll fluorescence image of epidermal tissue from Polypodium vulgare (f) showing similar photosynthetic efficiency of epidermal and guard cell chloroplasts. Bars, 10 µm.

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In 1908, Lloyd (Lloyd, 1908) observed that stomatacontained more starch when closed in the dark than whenopen during the day, which led to the starch–sugar hypothesis.This theory relies on the interconversion of starch to sugars,which results in osmotic changes, leading to alterations inguard cell turgor, and became the most widely accepted theoryregarding the osmoregulatory mechanism for many decades(Meidner & Mansfield, 1968). In the 1960s Fischer and co-workers highlighted the importance of potassium (K) uptakein stomatal opening (Fischer, 1968; Fischer & Hsiao, 1968)(although a paper on this by Imamura had already beenpublished; Imamura, 1943), and the starch–sugar hypothesiswas effectively replaced with the K+-malate2− theory. Thiswork demonstrated that K+ uptake (Yamashita, 1952; Fischer,1968; see reviews by Raschke, 1975, 1979) in guard cells wascorrelated with stomatal opening, with malate2− and/or chloride(Cl−) (Allaway, 1973; Schnabl, 1977; Schnabl & Raschke,1980; Outlaw, 1983; Willmer & Fricker, 1996; Asai et al.,2000) acting as the counterion(s). The general view is thatmalate2− is the major counterion balancing K+ uptake, althoughspecies such as Allium cepa (in which starch is absent) exclusivelyuse Cl− (Schnabl & Zeiger, 1977; Schnabl & Raschke, 1980).Uptake of K+, driven by a H+ gradient activated by protonATPase (Zeiger, 1983; Shimazaki & Kondo, 1987), was shownto be correlated with malate accumulation (Allaway, 1973)and stomatal opening (see review by Outlaw, 1983). A role forsucrose in guard cell osmoregulation was nearly forgottenuntil several studies suggested that K+ and its counterions couldnot provide all the osmoticum required to support stomatalapertures in Commelina communis (MacRobbie & Lettau,1980a,b), and led to the suggestion that soluble sugarsaccount for additional osmoticum to support opening(MacRobbie, 1987; Talbott & Zeiger, 1993). Evidence existsthat sugars as well as K+-malate2− can act as osmotica for guardcell osmoregulation (Outlaw & Manchester, 1979; Outlaw,1983; Reddy & Rama Das, 1986; Talbott & Zeiger, 1993,1998). For example, Commelina benghalensis accumulatessugars (60% of required osmoticum) and malate2− whentreated with fusicoccin (Reddy et al., 1983), a fungal toxinthat activates the plasma membrane H+-ATPase (Johanssonet al., 1993).

It is easy to see how these changes in osmoregulatory theoriesmay have been problematic for researchers determining therole of guard cell chloroplasts (see Fig. 3). For example, ifsucrose is not considered to be an important solute in watermovements, a role for guard cell photosynthetic carbon reductionis virtually redundant.

1. Multi-osmoregulatory pathways in guard cells

To resolve the differences reported in the literature among resultsobtained using different experimental procedures and differentspecies, Talbott & Zeiger (1996, 1998) outlined three distinctpathways (see Fig. 3) involved in guard cell osmoregulation

that incorporate K+, Cl−, malate2− and sucrose, also involvingguard cell chloroplasts. They suggested that the importance ofthese pathways may change depending upon time of day, species,and growth and experimental conditions (Talbott & Zeiger,1996, 1998). The first pathway describes the uptake of K+

and Cl− from the apoplast and/or the synthesis of malate2− fromcarbon skeletons derived from starch (Outlaw & Lowry, 1977;Outlaw & Manchester, 1979), and is believed to be involvedin the early morning opening response and under bluelight. In the second pathway, which is insensitive to DCMU(Poffenroth et al., 1992), sucrose is supplied from the breakdownof starch (Outlaw, 1982) and is also thought to play a role inblue-light responses (Tallman & Zeiger, 1988; Poffenrothet al., 1992; Talbott & Zeiger, 1993). In the third, DCMU-sensitive pathway, sucrose is supplied as a product of guard cellphotosynthetic carbon reduction (Talbott & Zeiger, 1998).In summary, K+ accumulation is used primarily for rapid openingin the morning, whereas turgor maintenance in the afternoonprimarily uses sucrose (Talbott & Zeiger, 1993, 1996, 1998).Talbott & Zeiger (1993) also demonstrated that the majorsolutes change depending upon lighting regimes and the durationof opening. In Vicia faba peels, the initial (30-min) stomatalopening in response to blue-light illumination resulted in a173% increase in the concentration of malate, which thendecreased, and the concentration of sucrose (from starchbreakdown) rose continuously, reaching 215% after 2 h. Underred light, there was little increase in organic acid or maltoseconcentrations, but the sucrose concentration increased to208% (Talbott & Zeiger, 1993), with no evidence of starchbreakdown (Tallman & Zeiger, 1988; Poffenroth et al., 1992;Talbott & Zeiger, 1993). As this was observed in epidermalpeels, sucrose must be supplied from guard cell photosyntheticcarbon reduction. Such observations support the hypothesisof multi-osmoregulatory pathways that are regulated bymeasurement (Talbott & Zeiger, 1998) and growth conditionssuch as humidity (Talbott et al., 2003) and CO2 concentration(Talbott et al., 1996, 1998; Frechilla et al., 2002, 2004; seealso Zeiger et al., 2002). The majority of this work was carriedout using V. faba and it is possible that different mechanismsare used by different species or groups of plants; for example,Arabidopsis lacks starch in the morning (Stadler et al., 2003).

A lack of evidence for significant carbon reduction in theguard cells (Outlaw, 1989; Tarczynski et al., 1989; Reckmannet al., 1990; Gautier et al., 1991) led Outlaw and co-workersto propose an alternative source of sucrose (Lu et al., 1995,1997; Ritte et al., 1999; Outlaw & De Vleighere-He, 2001;Outlaw, 2003; Kang et al., 2007a). Based on the work of Hiteet al. (1993), who suggested that guard cells act as carbonsinks, taking up sucrose via plasma membrane transporters(e.g. Stadler et al., 2003), Outlaw and colleagues suggestedthat apoplastic sucrose recently fixed in the mesophyll cellswas a source for guard cell symplastic sucrose and acted as anosmoticum for stomatal opening or replacing guard cellcarbon stores (Lu et al., 1997; Ewert et al., 2000; Outlaw &

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De Vleighere-He, 2001). However, this mechanism appears tobe dependent on the amount of sucrose in the apoplast, withconcentrations lower than 4 mM unable to support stomatalopening (Ritte et al., 1999). The guard cell apoplastic sucrosecan also exert an osmotic effect, which can drive stomatalclosure, acting as a possible signal between mesophyll assimila-tion rate and transpiration (Kang et al., 2007a). It was alsopostulated that sucrose concentrations near the guard cellregulate gene expression, as has been shown in many othertissues (e.g. Baiser et al., 2004). However, the majority of thesestudies were conducted on V. faba, which is an apoplasticphloem loader, and different mechanisms may regulate stomatalmovements in symplastic loaders (Kang et al., 2007b).

The above research emphasizes the importance of environ-mental growth and experimental conditions, as well asexperimental incubation periods, and provides possiblearguments for the involvement of K+, malate2− and sucrose instomatal function. Additionally, it provides a feasible explana-tion as to why so many conflicting results are reported in theliterature (Zeiger et al., 2002).

III. Role of guard cell chloroplasts in stomatal function

By the early 1990s a consensus was reached that in fact littlewas known about guard cell metabolism (Mansfield et al.,

Fig. 3 Schematic diagram showing possible osmoregulatory pathways in guard cells for solute accumulation. Blue lines represent pathways that are believed to be stimulated mostly by blue illumination, and red lines indicate pathways relating to red light- or photosynthesis-dependent pathways. These pathways may not be mutually exclusive. The diagram is not to scale. (Redrawn from information provided by Talbott & Zeiger, 1998; Vavasseur & Raghavendra, 2005; Shimazaki et al., 2007).

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1990) and, despite over a decade of research, in 2003 Ritte &Raschke (2003) reiterated that there was a lack of informationabout the physiology and role of the guard cell chloroplasts.

There are four primary ways in which guard cell chloroplastscould contribute to stomatal function (see Outlaw, 1983;Tominaga et al., 2001), with experimental evidence supportingall of these functions:• electron transport in guard cells produces ATP and/orreductants used in osmoregulation (Schwartz & Zeiger, 1984;Shimazaki & Zeiger, 1985);• chloroplasts are involved in blue-light signalling and response(Frechilla et al., 1999; Zeiger, 2000);• starch stored in the chloroplasts (either produced from carbonassimilated in the guard cell chloroplasts, or imported from themesophyll) is available to synthesize malate as a counter ion toK+ (Willmer & Fricker, 1996) or is broken down into sucrose;• photosynthetic carbon assimilation within guard cells producesosmotically active sugars (Tallman & Zeiger, 1988; Talbott &Zeiger, 1993, 1998; Zeiger et al., 2002).

1. Guard cell electron transport

Guard cells have a pigment composition similar to that ofthe mesophyll, along with functional photosystem I (PSI)and PSII (Zeiger et al., 1980; Outlaw et al., 1981; Shimazakiet al., 1982; Hipkins et al., 1983). Several researchers haveprovided evidence for linear electron transport, oxygen evolutionand photophosphorylation (Hipkins et al., 1983; Shimazaki& Zeiger, 1985; Willmer & Fricker, 1996; Tsionsky et al., 1997)which can be modulated by CO2 concentration (Melis &Zeiger, 1982) and blue light (Mawson & Zeiger, 1991;Srivastava & Zeiger, 1992). However, studies on fluorescencetransients have suggested that induction profiles in guard cellscan differ from those in mesophyll cells (Zeiger et al., 1980;Shimazaki et al., 1982; Mawson & Zeiger, 1991; Srivastava& Zeiger, 1992). Changes in the fine structure observed influorescence transients are believed to reflect changes in Calvincycle activity throughout the day or with different wavelengthsof light (Mawson & Zeiger, 1991; Srivastava et al., 1998; Zeigeret al., 2002). Additionally, the light-harvesting chlorophyllprotein is found in the phosphorylated state in the dark andis dephosphorylated by red light, the opposite situation tothat in mesophyll (Kinoshita et al., 1993). This reversal hasbeen suggested to be related to high rates of cyclic electronflow observed in guard cell protoplasts of V. faba, supportedby high PSI activity compared with the mesophyll (Lurie,1977), which could enhance ATP production driven by increaseddevelopment of the thylakoid proton gradient. However,Shimazaki & Zeiger (1985) did not observe any unusuallyhigh PSI activity in guard cells of V. faba but showed linearelectron flow to be c. 80% that of the mesophyll. This isconsistent with the values of PSII operating efficiency reportedlater by Lawson et al. (2002, 2003) using high-resolutionchlorophyll a (chla) fluorescence.

In the absence of any CO2 fixation, such electron transportrates could provide sufficient ATP to drive ion exchange duringstomatal opening (Shimazaki & Zeiger, 1985; Fig. 3), dependingon the light wavelength (Schwartz & Zeiger, 1984). Red lightinduced stomatal opening was shown to be DCMU sensitiveand potassium cyanide (KCN) (respiratory poison) resistantwhilst under blue light the reverse was observed (Schwartz& Zeiger, 1984). Patch clamp techniques established that achloroplast modulated red-light response stimulated a protonpump at the plasma membrane in guard cells, suggesting thatguard cell photosynthesis may regulate stomatal aperture, throughthe provision of energy (ATP) and photosynthetic signallingproducts, such as NADPH (Serrano et al., 1988; see also Wu& Assmann, 1993). However, later studies could not confirmthese findings (Roelfsema et al., 2001; Taylor & Assmann, 2001).Experiments conducted under red light with and without theinhibitors oligomycin (an inhibitor of oxidative phosphoryla-tion) and DCMU (an inhibitor of PSII) demonstrated thatguard cells supplied ATP to the cytosol under red light, whichwas utilized by the plasma membrane H+-ATPase for H+

pumping and stomatal opening (Tominaga et al., 2001). Analternative theory for the utilization of photosynthetic electrontransport products suggested that ATP and redox powerprovided from electron transport are used for the reductionof oxaloacetate (OAA) and 3-phosphoglycerate (3-PGA) (fromguard cell CO2 fixation or imported from the cytosol; Fig. 3)and exported to the cytosol via a 3-PGA-triose phosphateshuttle (Shimazaki et al., 1989; Ritte & Raschke, 2003).

Sugar as well as K+ accumulation during red light-inducedstomatal opening has been reported (Talbott & Zeiger, 1998;Olsen et al., 2002), with sugar production possible either fromstarch breakdown (Talbott & Zeiger, 1998) or from the utiliza-tion of end products of electron transport in photosyntheticcarbon reduction within the guard cells themselves (Fig. 3;Talbott & Zeiger, 1998; Olsen et al., 2002). Blue light-inducedstomatal opening (see next section) is generally believed not tobe dependent on products of guard cell electron transport, asit has been observed in the presence of DCMU (Sharkey &Raschke, 1981a; Schwartz & Zeiger, 1984; see also Roelfsema& Hedrich, 2005), and at low fluence rates (Zeiger, 2000;Shimazaki et al., 2007). The energy is thought to be suppliedmostly from mitochondrial respiration (Shimazaki et al., 1982,2007; Schwartz & Zeiger, 1984), although there is also evidencefor energy supply from guard cell chloroplasts (Mawson, 1993a).Stomatal opening in response to weak blue light is greatlyenhanced with a background of red light (Shimazaki et al., 2007),although red light is not essential (Sharkey & Raschke, 1981a).In additon to the Ci-driven response under red light, Shi-mazaki et al. (2007) proposed that guard cell chloroplaststranslocate NADH and ATP into the cytocol under red light,which are then used for malate synthesis under blue light.

The above studies demonstrate a direct role for the productsof guard cell photosynthetic electron transport in stomatalresponses, which is particularly evident under red light, but

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also possibly plays a minor role in the specific blue-light response.Energy and/or redox power can be used for CO2 fixation(through either phosphoenolpyruvate carboxylase (PEPc) orribulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco)),carbohydrate export or ion uptake (Gautier et al., 1991).

2. Role of guard cell chloroplasts in blue-light signalling and response

A recent comprehensive review by Shimazaki et al. (2007)discusses blue-light regulation of stomatal movement in greatdepth and I refer readers to this for full details. Briefly, bluelight induces rapid and highly sensitive stomatal openingcorrelated with the phosphorylation of a plasma membraneH+-ATPase pump and increased H+ pumping, which resultsin the activation of voltage-gated K+ channels by membranehyperpolorization (see Shimazaki et al., 2007 for details), alongwith the inhibition of s-type anion channels in Arabidopsis andV. faba (see Marten et al., 2007). Inhibition of blue light-inducedopening with KCN suggests that ATP for proton pumping issupplied mostly from mitchondrial respiration (Schwartz &Zeiger, 1984; Assmann & Zeiger, 1987; Parvathi & Raghavendra,1995). However, partial inhibition with DCMU (Mawson,1993a) implies a role for guard cell photosynthetic electrontransport in ATP supply, suggesting a possible metabolicco-ordination between photophosphorylation and oxidativephosphorylation in guard cells (Mawson, 1993b). H+ pumpingresults in K+ uptake correlated with malate2− synthesis and/orCl− uptake. Malate2− is the result mostly of starch breakdown(Outlaw & Manchester, 1979), as Arabidopsis mutants that donot accumulate starch lack a proper blue-light response(Lasceve et al., 1997). However, sucrose accumulation from starchbreakdown as an additional osmoticum in blue light-stimulatedopening in isolated V. faba stomata has also been demonstrated(Fig. 3; Tallman & Zeiger, 1988; Talbott & Zeiger, 1993).

Zeaxanthin (Zeiger & Zhu, 1998; Frechilla et al., 1999;Talbott et al., 2002) and phototropins (Kinoshita et al., 2001;Doi et al., 2004; Inoue et al., 2008) have both been suggested asthe blue-light receptor. Support for zeaxanthin as the specificblue-light receptor came from experiments conducted onepidermal peels of Arabidopsis mutants lacking zeaxanthin(non photochemical quenching 1), which failed to respond toblue light (Frechilla et al., 1999), although these results couldnot be confirmed when experiments were conducted onwhole leaves (Eckert & Kaldenhoff, 2000; Kinoshitaet al., 2001). Furthermore, stomata in V. faba treated withan inhibitor of carotenoid biosynthesis maintained a blue-light response, ruling out zeaxanthin as the only blue-lightreceptor (Roelfsema et al., 2006).

Strong evidence for phototropins as blue-light receptorswas provided by Kinoshita et al. (2001), who demonstrated(in both epidermal strips and intact plant material) that doublemutants for phot1 and phot2 proteins (serine/threonine proteinkinase) failed to respond to blue light. They established that

phot1 and phot2 act redundantly as the blue-light receptorsin stomatal responses to blue light, as single mutants showeda typical wild-type blue-light response (Kinoshita et al.,2001), which led to phototropins becoming widely acceptedas the main blue-light receptor.

The magnitude of the blue-light response decreases frommorning to afternoon (Doi et al., 2004), consistent with earlymorning stomatal opening, when light is enriched in the bluewavelengths (Assmann & Shimazaki, 1999), and also consistentwith the theory of varying osmoregulatory pathways (Talbott& Zeiger, 1998), and changes in guard cell fluorescence tran-sients through the day (Srivastava et al., 1998). It should benoted, however, that the stomatal response to blue light is notuniversal, with several species lacking blue light-inducedstomatal opening. Stomata of the fern Adiantum capillus-venerisdo not open in response to blue light, despite having functionalphototropins and plasma membrane H+-ATPase (Doi et al.,2006). Additionally, facultative CAM plants displayed blue-light-specific stomatal opening in C3 but not in CAM mode(Lee & Assmann, 1992; Talbott et al., 1997).

3. PEPc activity, malate synthesis and starch breakdown

An alternative sink for the end products of guard cellphotosynthetic electron transport is malic acid production viaPEPc, and CO2 fixation (Willmer & Dittrich, 1974; Raschke& Dittrich, 1977; Schnabl et al., 1982; Willmer, 1983; Outlaw,1990) using carbon skeletons provided by starch breakdown(Pallas & Wright, 1973; see also Asai et al., 2000). Outlaw &Manchester (1979) demonstrated a quantitative relationshipbetween malate accumulation and starch loss. Light-stimulatedincreases in PEPc activity have been demonstrated together withincreased NADP- or NAD-dependent malate dehydrogenaseactivity, which catalyses the reduction of OAA (Rao &Anderson, 1983; Scheibe et al., 1990), and malate accumulationhas been correlated with stomatal aperture (Allaway, 1973;Pearson, 1973; Pearson & Milthorpe, 1974; Vavasseur &Raghavendra, 2005). It is widely accepted that guard cellscontain high concentrations of starch and PEP carboxylase(Willmer et al., 1973; Willmer & Rutter, 1977; Raschke,1977, 1979; Outlaw & Kennedy, 1978) compared withmesophyll cells (Cotelle et al., 1999) and many reports havesuggested that this is the major or only pathway for CO2fixation in guard cells (e.g. Willmer et al., 1973; Reckmannet al., 1990; see also Vavasseur & Raghavendra, 2005) intomalate and aspartate (Ogawa et al., 1978). The importanceof malate accumulation in light-induced stomatal opening(Asai et al., 2000) has been demonstrated using 3,3-dichloro-dihydroxyphophinoyl-methyl-2-propenoate (DCDP), an inhibitorof PEPc (Parvathi & Raghavendra, 1997). PEPc activity andmalate formation have also been linked to CO2 responsemovements in stomatal guard cells (Outlaw & Lowry, 1977;Raschke, 1979; Schnabl et al., 1982; Hedrich & Marten,

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1993; Hedrich et al., 1994; Cousins et al., 2007). Differencesin stomatal opening between adaxial and abaxial stomata havebeen closely associated with differential starch hydrolysis, malatesynthesis and K+ uptake (Pemadasa, 1983) as well as lightwavelength (Wang et al., 2008), highlighting again theflexibility of stomatal osmoregulation and behaviour dependingupon the environment. Further support for the importanceof PEPc activity in stomatal opening comes from recentwork conducted on PEPc-deficient mutants of the C4 dicotAmaranthus edulis, which showed reduced rates of bothstomatal opening and final conductance compared with wild-type controls (Cousins et al., 2007). This recent work is inagreement with earlier studies on potato (Solanum tuberosum)plants which demonstrated greater rates of opening whenPEPc was over-expressed and reduced rates of opening inplants with decreased amounts of PEPc (Gehlen et al., 1996).This work is discussed in greater detail in Section V.

Starch degradation to sucrose is also involved in stomatalopening. In dual light experiments, Tallman & Zeiger (1988)found substantial starch degradation under blue-light illumina-tion, but only a small amount of K+ uptake. From these observa-tions they suggested that, if starch was converted to malate(Schnabl, 1980), adequate uptake of K+ would be necessary tocounterbalance the anion. These results are consistent with theobservation that, in 10 μmol m−2 s−1 blue light, the rate of malatesynthesis in V. faba guard cells was only 25% of their maximum(Ogawa et al., 1978). From such studies it was concluded thatstarch breakdown under blue light can also result in accumula-tion of sucrose rather than malate (Tallman & Zeiger, 1988).

4. Guard cell photosynthesis and sucrose production via the Calvin cycle

Research into guard cell photosynthesis and carbon metabolismhas spanned several decades, but as yet there is no generalconsensus. There are conflicting reports in the literatureconcerning the capacity of photosynthetic carbon reduction inguard cell chloroplasts and its importance in stomatal function(see reviews by Shimazaki et al., 1989; Outlaw, 1989).

Early studies provided little evidence of Calvin cycle activityin guard cell chloroplasts (Outlaw et al., 1979, 1982; Outlaw,1982, 1987, 1989; Tarczynski et al., 1989). Willmer &Dittrich (1974) showed that, in the epidermis of Tulipa andCommelina in the light, 14CO2 was fixed into malate andaspartate. This was validated later by Raschke & Dittrich(1977), who showed that neither radioactive 3-PGA norRubisco activity was present in epidermal peels of the sametissues when they were exposed to 14CO2. Subsequentexperiments demonstrated that guard cell chloroplasts lackedribulose-1,5-bisphosphate carboxylase (RuBPC) and ribulose-5-phosphate kinase (Ru5PK) activity (Outlaw et al., 1979)and other key enzymes (Outlaw et al., 1979; Schnabl, 1981)for the photosynthetic carbon reduction pathway (PCRP).This was in agreement with a lack of phosphorylated Calvin

cycle products in V. faba guard cell protoplasts in the light(Schnabl, 1980). Screening 41 species using indirect immun-ofluorescence, Madavhan & Smith (1982) reported no evidenceof Rubisco in the guard cells of C4 plants and only negligibledetection in C3 plants, but appreciable amounts in one-third of CAM species surveyed. The size of the guard cellphosphoglycerate pool was unaffected by light, suggestingthat PCRP is not involved in aperture regulation (Outlaw &Tarczynski, 1984). Reckmann et al. (1990) determinedthat only 2% of the solute required for stomatal opening wasprovided by Rubisco activity in Pisum sativum, and concludedthat there was insignificant Rubisco activity, confirming theconclusion of Hampp et al. (1982) that photoreduction ofCO2 by guard cells was absent.

In contrast to the above findings, the presence of Rubiscoin guard cells of V. faba has been unequivocally shown withimmunocytochemical localization (Zemel & Gepstein, 1985).Numerous studies have also shown that guard cells containseveral of the other main Calvin cycle enzymes (Shimazaki &Zeiger, 1985; Gotow et al., 1988; Shimazaki, 1989). Zemel &Gepstein (1985) quantified Rubisco on a chlorophyll basis at40–50% compared with mesophyll cells. Shimazaki et al.(1989) validated these figures, showing RuBPC activity inguard cells of the same species to be 40% of that in mesophyllchloroplasts (on a chlorophyll basis). However, they suggestedthat the low ratio of CO2 fixation to O2 evolution impliedthat the major proportion of ATP and reducing equivalentswas used for reactions other than photosynthetic CO2 fixation.These authors also pointed out that in many previous studiesvalues had been calculated on a cell rather than a chlorophyllbasis, and that recalculation would significantly increase pre-viously obtained values in line with their observations.

The results of Gotow et al.’s study (1988) contradictedearlier findings (Raschke & Dittrich, 1977) and showed thatfeeding radio-labelled CO2 to guard cell protoplasts underred light resulted in incorporation of radioactivity into 3-phosphoglycerate, ribulose bisphosphate (RuBP), fructose andsedoheptulose. Medium alkalinization indicating CO2 uptakeand oxygen evolution by guard cell protoplasts was shownunder white (Gotow et al., 1988) and red light (Shimazaki &Zeiger, 1987). A photosynthetic dependence of sucroseaccumulation was illustrated using DCMU in epidermal peelsof V. faba under red light by Poffenroth et al. (1992). However,reduced CO2 concentration triggered K+ uptake rather thansucrose accumulation under red light (Olsen et al., 2002).

It is now widely accepted that the Calvin cycle enzymes arepresent in guard cell chloroplasts; however, the debate overtheir activity, function and role in stomatal behaviour remains(Outlaw, 1996, 2003). Although guard cell photosyntheticcarbon reduction has been shown in epidermal peels (Tallman& Zeiger, 1988; Poffenroth et al., 1992), guard cell chloroplasts(Shimazaki et al., 1982; Gotow et al., 1988; Shimazaki, 1989)and isolated guard cell pairs (Tarczynski et al., 1989), thecontribution to osmotic requirements for stomatal opening

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ranges from 2% (Reckmann et al., 1990) to 40% (Poffenrothet al., 1992) (see Wu & Assmann, 1993). Many reports havesuggested that rates are too low for any functional significance(Outlaw, 1989; Outlaw et al., 1982), whilst others have pro-posed the Calvin cycle to be a major sink for the products ofphotosynthetic electron transport (Cardon & Berry, 1992;Zeiger et al., 2002; Lawson et al., 2002, 2003). Evidence forguard cell production of sucrose has been obtained duringred light-induced stomatal opening in V. faba, where nostarch breakdown was observed and sugar import was ruledout as a result of the use of epidermal peels (Tallman & Zeiger,1988; Talbott & Zeiger, 1993). Parvathi & Raghavendra(1997) also showed that Calvin cycle activity increased withapplication of DCDP, an inhibitor of PEPc activity, suggest-ing that this pathway may become important when PEPc isrestricted.

5. Evidence for all four mechanisms in stomatal function

In conclusion, there is evidence for all four of the abovemechanisms being involved in stomatal function. The blue-light stomatal response is believed to be mostly independent ofguard cell electron transport, as stomatal blue-light responseshave been observed in albino leaves (Karlsson et al., 1983;Roelfsema et al., 2006), with energy for the activation of a plasmamembrane H+-ATPase supplied mostly by the mitochondria(Fig. 3; Schwartz & Zeiger, 1984; Assmann & Zeiger, 1987;Parvathi & Raghavendra, 1995), although there is also evidencefor chloroplastic supply (Mawson, 1993a) and red-lightenhancement (Shimazaki et al., 2007). There is support forboth zeaxanthin and phototropins as the blue-light receptors,with phototropins the most widely accepted. Evidence existsfor the direct use of ATP and/or reductants (produced byguard cell electron transport) in red light-induced stomatalopening (Fig. 3; e.g. Shimazaki et al., 1989; Tominaga et al.,2001; Olsen et al., 2002; Ritte & Raschke, 2003), as well asin sugar production by photosynthetic carbon reductionwithin the guard cells (Fig. 3; Talbott & Zeiger, 1996, 1998).Starch stored in the guard cells can be broken down into eithermalate2− (as a counterion for K+ uptake; see Fig. 3) (Willmer& Dittrich, 1974; Raschke & Dittrich, 1977; Outlaw &Manchester, 1979; Schnabl et al., 1982) or sugars, which actas osmotica for stomatal opening (Outlaw, 1982; Tallman &Zeiger, 1988; Poffenroth et al., 1992). It appears that guardcell chloroplasts can be involved in all four of the pathwaysdescribed above, and that the pathway used is conditional on thespecies, time of day, and experimental protocols.

IV. Chlorophyll a fluorescence studies to examine guard cell photosynthesis

Chlorophyll a fluorescence is a powerful technique to probeand elucidate photosynthetic metabolism in guard cells. The

early pioneering work of Zeiger and co-workers (Zeigeret al., 1980; Melis & Zeiger, 1982; Mawson & Zeiger, 1991)measuring Kautsky kinetics (Kautsky & Franck, 1943) inindividual guard cells showed distinct features associatedwith Calvin cycle activity. The majority of early chlorophyllfluorescence work was restricted to epidermal peels (Ogawaet al., 1982), protoplasts (Outlaw et al., 1981; Goh et al., 1999),single guard cell pairs or the white areas of variegated tissue(Zeiger et al., 1980; Melis & Zeiger, 1982; Cardon & Berry,1992). Cardon & Berry (1992) examined changes in steady-state chlorophyll fluorescence (F ′) from guard cells in thewhite areas of intact leaves of Tradescantia albiflora underdifferent CO2 and O2 concentrations and attributed changesto photochemical and nonphotchemical quenching. Fromthese observations they concluded that both the carboxylationand oxygenation of RuBP were major sinks for the endproducts of photosynthetic electron transport. However, cautionshould be applied when interpreting steady-state fluorescencemeasurements as it is difficult to distinguish betweenphotochemical and nonphotochemical quenching components(Baker, 2008). The report by Cardon & Berry (1992) was thefirst research to provide physiological evidence for Rubisco-mediated CO2 fixation and photorespiration and led the wayfor many subsequent studies.

Advances in fluorescence methodology (see Goh et al., 1999),with the development of the saturation pulse method offluorescence quenching analysis (Bradbury & Baker, 1981;Schreiber et al., 1986) and pulse amplitude modulation (PAM)fluorimetry (Schreiber et al., 1986) in conjunction withtechnological developments in microfluorimetry (Goh et al.,1999) and high-resolution imaging (Oxborough & Baker,1997), made it possible to parametrize measurements ofchlorophyll fluorescence at the cellular (Oxborough & Baker,1997; Goh et al., 1999) and subcellular levels (Baker et al.,2001). With such advancements, measurements of PSIIoperating efficiency ( , where is the differencebetween maximum fluorescence in the light adapted state( ) and steady state fluorescence in the light (F ′ )) could beobtained for individual cells and protoplasts (Goh et al., 1999).The PSII operating efficiency estimates the efficiency at whichlight absorbed by PSII is used for the reduction of the plasto-quinone QA, and can provide an estimate of the quantum yieldof linear electron flux through PSII (Baker, 2008). Goh et al.(1999) first used such techniques to compare fluorescencequenching characteristics in guard and mesophyll cell protoplasts(in V. faba and Arabidopsis). Light induction curves dis-played very similar characteristics, indicating similar functionalorganization of the thylakoid membranes, although guard cellswere saturated at lower light intensities and mesophyll cellprotoplasts had a higher capacity for photosynthetic electrontransport. In the same study, anaerobic conditions suppressedphotosynthetic electron flow in guard cells compared withmesophyll cells. The O2-dependent electron flow suggesteda role for the Mehler-ascorbate peroxidase (MAP) cycle or a

′ ′F Fq m/ ′Fq

′Fm

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close metabolic coupling between photosynthetic electrontransport and export of reducing equivalents via a 3-phosphoglycericacid/dihydroxyacetone phosphate (PGA/DHAP) shuttle andoxidative phosphorylation in the mitochondria (Goh et al.,1999). However, again this work was restricted to measurementsof protoplasts or white areas of variegated plant tissue and wasnot in agreement with later studies conducted on intact greenmaterial (Lawson et al., 2002, 2003).

The first study that simultaneously examined the PSIIoperating efficiencies ( ) of guard and mesophyll cells inintact green tissue revealed guard cell photosynthetic efficiencyto be 70–80% that of mesophyll chloroplasts (Lawson et al.,2002). However, electron transport rates for the two cell typescould not be calculated because of uncertainties in the exactlight absorption and the contribution of PSI fluorescence inguard and mesophyll chloroplasts. In the same study theseresearchers measured at different CO2/O2 concentra-tions in guard cells of intact green leaves of Tradescantia albifloraand Commelina communis and confirmed that Rubisco was amajor sink for the products of photosynthetic electron transport.Later this was confirmed in the guard cells of several other species,including the C4 plant Amaranthus caudatus (Lawson et al.,2003), and was consistent with the results of immunogoldlabelling studies, which found weak labelling of PEPc butsignificant Rubisco labelling in guard cells of Amaranthusviridis (Ueno, 2001). The fact that the same CO2/O2 responsewas observed in guard cells and in mesophyll cells suggeststhat a major proportion of the end products of electron trans-port are being used by Rubisco and the Calvin cycle. Guardcells contain 20–50-fold less chlorophyll than the underlyingmesophyll (Willmer & Fricker, 1996) and therefore, at similarphotosynthetic rates, extrapolation to the whole-cell level wouldresult in much lower guard cell photosynthesis compared withthe mesophyll. However, the small volume of guard cells(one-tenth that of the mesophyll) means that the guard cellCO2 assimilation rate could be one-third to one-tenth that ofthe mesophyll, and therefore guard cell chloroplasts couldprovide a significant energy source for these cells.

1. Alternative sources of energy

Although the aim of this review is to concentrate on guard cellchloroplasts and their possible role in stomatal function,guard cells also contain numerous mitochondria (Willmer &Fricker, 1996; Vavasseur & Raghavendra, 2005), about one-thirdthe number of those in the mesophyll (Allaway & Setterfield,1972), and several reports have suggested that these are themost important organelle in guard cells. Hampp et al. (1982)originally proposed that there was an absence of photoreductionof CO2 in guard cells but a high metabolic flux through thecatabolic pathway. High respiration rates were observed byRaghavendra & Vani (1989), suggesting that ATP producedthrough oxidative phosphorylation was important for stomatalmovements (Parvathi & Raghavendra, 1997).

Fumarase activity (which is involved in the tricarboxylicacid (TCA) cycle) has been shown to be high in guard cells ofV. faba and P. sativum (Hampp et al., 1982; see also Outlaw,2003), and trangenic tomato (Solanum lycopersicum) plants withconsiderable reductions in mitochondrial fumarate hydratase(fumarase) activity showed substantial reductions in stomatalaperture, resulting in CO2 limitation of photosynthesis(Nunes-Nesi et al., 2007). Application of inhibitors ofphotophosphorylation (DCMU) and oxidative phosphoryla-tion (KCN) has shown that both mechanisms are used forlight-induced opening, but depend on wavelength (Schwartz& Zeiger, 1984). Their relative importance alters when eitherof these pathways is restricted (Parvathi & Raghavendra, 1997),suggesting that both organelles (chloroplasts and mitochondria)play a role in stomatal function (Asai et al., 2000).

V. Linking stomatal behaviour to mesophyll photosynthesis

Stomatal conductance is well co-ordinated with mesophyllphotosynthetic CO2 fixation (Wong et al., 1979; Farquhar &Wong, 1984; Mansfield et al., 1990). Numerous studies havedemonstrated a strong correlation between photosynthesisand stomatal conductance under a variety of different lightintensities and nutrient and CO2 concentrations (Radin et al.,1988; Hetherington & Woodward, 2003). This relationshipcauses, or is a consequence of, a constant Ci:Ca ratio (where Cais the external CO2 concentration), which has been observedto remain constant over the long term (Wong et al., 1979,1985), although short-term variations have often been apparent(Sharkey & Raschke, 1981; Morison, 1987). However, itshould also be noted that this relationship has easily beenbroken in transgenic plants with various modifications tophotosynthetic metabolism (e.g. Hudson et al., 1992; Lauereret al., 1993; Stitt & Schulze, 1994; von Caemmerer et al.,2004; Cousins et al., 2007; Baroli et al., 2008). The closerelationship between photosynthesis and stomatal behaviourled to the hypothesis that guard cell responses may be linkedto mesophyll photosynthetic capacity via a mesophyll signal orthat guard cell photosynthesis itself may provide a metabolitesignal (Wong et al., 1979). Chloroplast ATP pool size was putforward by Farquhar & Wong (1984) as a possible metabolite,a theory that was built upon later by Buckley et al. (2003),whilst zeaxanthin has been put forward by Zeiger & Zhu(1998) in view of a close correlation between zeaxanthinconcentration and stomatal apertures (in response to light andCO2; see Zhu et al., 1998).

The debate over whether guard cell chloroplasts and/orguard cell photosynthesis plays a direct role in the co-ordinationof stomatal movements in relation to mesophyll photosyntheticCO2 demand remains unresolved. Recently, Roelfsema et al.(2002, 2006) have argued against a direct role for guard cellchloroplasts in red light-induced stomatal movements. Theseresearchers used albino areas of variegated plant tissue of

′ ′F Fq m/

′ ′F Fq m/

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V. faba treated with norflurazon (nf; inhibits carotenoidbiosynthesis), and showed stomatal opening in these two tissuetypes in response to blue light but not red (Roelfsema et al.,2006). They concluded that a lack of red-light response isconsistent with intercellular CO2 concentration as the inter-mediate signal in the stomatal red-light response. Moreover,stomatal opening in response to red light was only apparentwhen light was applied to a large area of the leaf, and not when itwas applied to individual guard cells, supporting a mesophyll-driven Ci response (Roelfsema et al., 2002).

These obervations are in good agreement with earlier studiesby Karlsson (1986), who showed that lowering the atmosphericCO2 concentration had a similar effect to red light andenhanced the blue-light response. It should, however, bementioned that stomata in the albino portion of the leaf cannotbe considered to be completely indicative of responses ingreen tissue (Scarth & Shaw, 1951; Lawson et al., 2002) astheir movements are much slower than those in green areas(Scarth, 1932). However, Scarth (1932) also noted under redlight that stomata located near the green tissue tend to openfurther than those at greater distances, adding support to thetheory of an indirect effect of red light on stomatal movementsthrough the action of mesophyll photosynthesis. Furtherevidence for a CO2-mediated red light-induced stomatalopening response has been provided by the Arabidopsis hightemperature 1 (HT1) mutant which carries a mutation in thegene encoding a protein kinase (Hashimoto et al., 2006). Thesemutants lack both a guard cell CO2 response and a red-lightresponse, but respond to blue light, supporting the notion thatred light-driven stomatal opening is promoted by reduced Ci,although to corroborate this it would be necessary to presentstomatal conductance against Ci rather than CO2. Additionalsupport for Ci-driven stomatal opening has been provided inNicotiana tabacum in which a MAP kinase gene (NtMPK4)involved in the activation of anion channels was silenced.These plants did not close in elevated atmospheric CO2 andshowed a reduced response to red light (Marten et al., 2008).

A recent publication by Mott et al. (2008) has suggested(as have many other studies; see above) that most stomatalresponses to light and CO2 occur in response to an unknownmesophyll-generated signal. Epidermal peels of Tradescantiapallida, V. faba and P. sativum showed no stomatal response tolight or CO2, but when T. pallida and P. sativum peels weregrafted back onto mesophyll (either their own correspondingmesophyll or that of a different species), stomatal responseswere restored, although this was not the case for V. faba. Theauthors argued against a direct effect of mesophyll-drivenchanges in Ci, as increasing ambient CO2 from 120 to540 μmol mol−1 did not induce stomatal closure, whereasdarkness resulted in complete closure, but Ci was only200 μmol mol−1. In agreement with this, a further recentstudy reported that abaxial stomata of Helianthus annuus weremore sensitive to light transmitted through the leaf (self-transmitted light) than to direct illumination, highlighting a

photosynthesis-dependent involvement in stomatal responsesto light. This was attributed to an unknown photosyntheticmetabolite and not a Ci-driven effect, as Ci was maintained ata constant value (Wang et al., 2008).

Several studies have also argued against a direct effect of Cion stomatal-driven responses to red light; for example,stomata were shown to respond to light even when Ci was heldconstant (Messinger et al., 2006; Lawson et al., 2008; Wanget al., 2008). Additionally, stomatal responses to Ci and Ciresponses to light are believed to be too small to account forthe large changes in stomatal conductance that are oftenobserved in response to light (Sharkey & Raschke, 1981b).Furthermore, red-light responses have been documented inseveral studies conducted in epidermal peels (Tallman &Zeiger, 1988; Olsen et al., 2002) and protoplasts (Raschke &Dittrich, 1977; Goh et al., 1999) isolated from the mesophyll.

Evidence for a direct role of guard cell chloroplasts in redlight-induced stomatal opening has been reported, althoughthis may be species dependent. Recently, Doi & Shimazaki(2008) examined stomatal responses in the fern Adiantumcapillus-veneris to CO2 in darkness and found the stomata tobe unresponsive to low or high CO2 concentrations but toopen in response to red light. The fact that they observed asynergistic effect of red and far-red light on stomatal opening,and greater sensitivity when light was applied directly to thelower surface along with a lack of response to Ci, led theseauthors to conclude that opening in this species is driven byphotosynthetic electron transport in guard cell chloroplasts.This is probably driven by K+ uptake, as CsCl (a K+ channelblock) inhibited the response (Doi & Shimazaki, 2008). Inthe same experiment, Arabidopsis was used as a control andshowed ‘typical’ Ci responses in the dark, highlighting thepossibility that different species may use alternative signallingpathways and mechanisms. In contrast to these findings, thestomata of an Arabidopsis mutant that lacks a functionalSLOW ANION CHANNEL-ASSOCIATED 1 (SLAC1) gene,which encodes a plasma membrane anion channel, were foundto fail to close at a high CO2 concentration in the dark and inthe light in one study (Negi et al., 2008), and to open in thelight and close more slowly in the dark in another (Vahisaluet al., 2008). These studies support the involvement of iontransport mechanisms in light-dependent stomatal move-ments, that are not dependent solely on Ci-driven responses.

To address stomatal behaviour in relation to mesophyllphotosynthesis, Messinger et al. (2006) suggested that thebalance between photosynthetic carbon reduction by Rubiscoand electron transport capacity was the key mechanism linkingstomatal response to light and CO2 concentration. This work wasbased on alterations in the amount of ATP and/or zeaxanthinresulting from a change in the balance of guard cell electrontransport (and energy states of the thylakoid membrane) inrelation to photosynthetic carbon reduction, determined bylight and Ci. Variations in the concentration of zeaxanthin inturn would alter guard cell aperture in response to blue light

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and CO2 (Zeiger & Zhu, 1998; Zhu et al., 1998; Zeiger et al.,2002). As with zeaxanthin, ATP should increase with light anddecrease with Ci with increased Calvin cycle activity. IncreasedATP in guard cells could be used in the cytosol for protonpumping at the plasmalemma (Tominaga et al., 2001), oralternatively the energy may be utilized to produce sucrose asan osmoticum for stomatal opening through photosyntheticcarbon reduction (Talbott & Zeiger, 1993). This work alsosuggested that there are at least two mechanisms by whichstomata respond to CO2, one dependent on photosynthesis,and the other photosynthetically independent (Messinger et al.,2006). Multiple CO2 response mechanisms have previouslybeen suggested by Assmann (1999). However, studies conductedon transgenic plants have demonstrated similar conductancesin wild-type and trangenic plants (see next section), despite thelatter having an alteration in the balance of electron transportrates relative to carboxylation capacity (von Caemmerer et al.,2004; Baroli et al., 2008).

1. Progress using transgenic and mutant plants

The choice of an ideal experimental system is critical whenattempting to determine the role of mesophyll or guard cellphotosynthesis in stomatal function and in the past has reliedon epidermal peels or guard cell protoplasts. Such systems areoften criticized because of possible mesophyll contamination(Weyers & Travis, 1981; Outlaw, 1983). However, the removalof the mesophyll could prevent any mesophyll signalling andmay induce other mechanistic responses (Lee & Bowling,1995; Lawson et al., 2002). In the late 1970s, Outlaw andco-workers introduced the technique of dissecting individualcells (Outlaw, 1980; Hampp & Outlaw, 1987; Outlaw & Zhang,2001), highlighting its advantage in controlling contamination.Improved molecular and transgenic techniques have providedmodern powerful tools (Webb & Baker, 2002) with which toaddress many questions regarding photosynthesis in relationto stomatal function and have already provided some invaluableinformation (see above).

Early studies conducted on transgenic plants with impairedphotosynthesis revealed some surprising results. The effects ofRubisco concentration on photosynthesis were studied inde-pendently by several groups, all of which studies suggestedlittle effect on stomatal behaviour (Quick et al., 1991; Stittet al., 1991; Hudson et al., 1992). Specifically, these studiesshowed similar stomatal conductance values in transgenicplants compared with wild-type controls, despite a severereduction in photosynthesis and higher Ci concentrations.Studies on tobacco (Nicotiana tabacum) plants with reducedphosphoribulokinase (Paul et al., 1995) or Rieske FeS protein(Price et al., 1998) also showed no effect on stomatal conduct-ance, and Price et al. (1998) concluded that ‘stomata are notstrongly reliant on photosynthetic electron transport for settingconductance’. However, work on transgenic antisense PEPcplants supported a role for malate and PEPc activity in guard

cells, with delays in stomatal opening responses in potato withreduced PEPc activity, whilst over-expressors showed acceleratedopening (Gehlen et al., 1996). These findings are supportedby recent work on Amaranthus edulis mutants deficient in PEPc,which show both reduced rates of opening and also reducedfinal stomatal conductances (Cousins et al., 2007). Stomata inplants with 12% wild-type fructose-1, 6-bisphatase (FBPase)activity showed significantly faster opening responses andhigher final conductances with increasing irradiance, despitelower photosynthetic rates and elevated Ci. However, this wasdependent upon humidity and external CO2 concentration(Muschak et al., 1999).

However, the aims of the above studies were not specifically todetermine stomatal responses. Certain assumptions were made:firstly, because Cauliflower mosaic virus (CAMv) promotors wereused in the majority of the studies, it was assumed that all cellswere antisensed in a similar manner, and, secondly, it was assumedthat the observed response was equivalent to steady-state stomatalconditions. To directly address the influence of reducedphotosynthetic capacity using antisense technology, vonCaemmerer et al. (2004) used high-resolution chlorophyllfluorescence imaging to show for the first time that photosyn-thetic efficiency was reduced to a similar extent in the guardcells as in the mesophyll cells in tobacco plants with reducedconcentrations of Rubisco. Decreasing Rubisco activity resultedin an imbalance between chloroplast electron transport andthe photosynthetic carbon reduction capacity, which in turncould lead to an increased amount of ATP and/or conversionof xanthophyll pigments to zeaxanthin. Increased nonphoto-chemical quenching was observed in the antisense plants,which could be interpreted as an increase in the amount ofzeaxanthin. Both ATP and zeaxanthin have been implicated asplaying a key role in stomatal opening responses. However, astep change in irradiance revealed similar stomatal responsesin terms of opening rates and final conductances in antisenseRubisco plants and wild-type controls, despite significantlylower photosynthetic rates in the former. This led to elevatedinternal CO2 concentrations within these plants, which initiallywas interpreted as a reduced sensitivity to Ci (von Caemmereret al., 2004). However, manipulation of Ci through changesin Ca resulted in stomatal closure, suggesting that stomatamay response to Ca and not Ci (von Caemmerer et al., 2004).The overall conclusion from this work was that neither mesophyllnor guard cell photosynthesis was necessary for stomatalopening responses.

The fact that the majority of studies used white light or amixture of red and blue light does not rule out the possibilitythat blue light-stimulated opening, independent of photosyn-thesis, overrides any mesophyll/guard cell signal (Talbott &Zeiger, 1993). To resolve this issue, a recent study of red-lightresponses by Baroli et al. (2008) distinguished betweenantisense Rubisco tobacco plants with 10–15% wild-typeRubisco activity, which have major reductions in the carbox-ylation capacity of photosynthesis, and antisense tobacco

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plants with impaired rates of electron transport via reductions inthe cytochrome b6f complex. No changes in stomatal openingwere observed in either of the transgenic plants in response toa step change in red light at ambient CO2 concentrations,leading to the conclusion that this response was independentof guard or mesophyll cell photosynthesis. The fact that nophenotypic stomatal responses were observed despite a decreasein sucrose concentration also strongly suggests that somethingother than sucrose acts as osmoregulator during opening.However, recent work conducted on antisense sedopheptulose-1,7-bisphosphatase (SBPase) tobacco plants has shown aminor regulatory role for photosynthetic electron transport inresponse to red light (Lawson et al., 2008). A step change inred illumination resulted in an increased rate of stomatalopening which was not observed under a blue/red light mix.ATP concentrations in the antisense SBPase plants may beincreased because of the reduced ATP consumption bythe Calvin cycle. These authors suggested the possibility ofincreased stomatal opening under red light, as a result of anincrease in ATP available for proton pumping (Tominagaet al., 2001). However, Baroli et al. (2008) reported little effectof reduced ATP on red light-induced stomatal opening intransgenic tobacco plants with reduced cytochrome b6f complex(Baroli et al., 2008).

Other suggested functions of chloroplasts in guard cells inregulation of stomatal behaviour include the production ofreactive oxygen species such as H2O2 (possibly via Mehleractivity at PSI) which may play a role in abscisic acid (ABA)signal transduction (Zhang et al., 2001). A role for ascorbicacid (Asc) redox state has also been postulated in guard cellregulation. Plants with increased guard cell Asc redox state(through increased expression of dehydroascorbate reductase(DHAR)) exhibited a reduced concentration of guard cellH2O2 and consequently higher stomatal conductances (Chen& Gallie, 2004).

As highlighted above, the use of transgenic and mutantplants has provided significant information regarding guardcell mechanisms (von Caemmerer et al., 2004; Baroli et al.,2008; Lawson et al., 2008), sensory molecules (e.g. Eckert &Kaldenhoff, 2000) and signal transduction cascades (Inoueet al., 2008), as well as ion uptake and ion channel regulation(Serna, 2008), all of which play key roles in stomatal sensitivityand behaviour.

VI. Stomata in relation to water use/manipulation of behaviour

One of the important outcomes of understanding how guardcells function is the potential to engineer drought-tolerantplants. This prospect has received increasing attention fromthe wider scientific community, with several reports publishedrecently suggesting that stomatal metabolism may hold thekey (Nilson & Assmann, 2007). For example, maize (Zeamays) plants with increased amounts of NADP-malic enzyme

(ME), which converts malate and NADP to pyruvate,NADPH and CO2, had altered stomatal behaviour. ME-transformed plants had decreased stomatal conductance,showing signs of drought avoidance associated with guard cellmalate metabolism. A negative aspect of this drought-toleranceengineering was that, following exposure to drought, thedevelopment of necrosis was more rapid in leaves from plantswith the highest ME expression (Laporte et al., 2002). Masleet al. (2005) reported the isolation of a ‘transpiration efficiencygene’, ERECTA, which acts on cell expansion and cell division,amongst other processes, resulting in modification of leafdiffusive properties and mesophyll capacity for photosynthesis,leading to greater water use efficiency, in Arabidopsis. Increasesin drought resistance have also been reported in Arabidopsismutants, with alterations or disruptions of guard cell membranetransporters (Klein et al., 2004), calcium-dependent proteinkinases (Ma & Wu, 2007), and the expression of aquaporin genes(Cui et al., 2008) and genes involved in ABA biosynthesis,expression or sensitivity (Jakab et al., 2005; Wang et al., 2005;Yang et al., 2005). Such studies are not restricted to Arabidopsis;for example, over-expression of the stress-responsive geneSNAC1 (STRESS-RESPONSIVE NAC1) enhanced droughttolerance in rice (Oryza sativa) (Hu et al., 2006).

In the attempt to produce plants with increased water useefficiency or drought tolerance, genetic engineering or mutationsprovide an opportunity to alter not only stomatal physiologyand function (Nilson & Assmann, 2007) but also anatomicalfeatures, such as stomatal density and size (originally proposedin the 1970s; Jones, 1976, 1977) and amounts of leaf cuticularwax (Aharoni et al., 2004). A recent review by Wang et al.(2007) highlights the importance of, and recent progressmade in, identifying genes controlling stomatal density orpatterning, and how such genetic manipulations may increaseplant water use efficiency.

Altering the stomatal density does not automatically alterstomatal conductance (Fig. 4; Lawson, 1997; Weyers & Lawson,1997; Lawson & Morison, 2004). Figure 4 shows a model ofpredicted stomatal conductance with changes in variousstomatal characters. From this model it is obvious that stomatalaperture has the greatest control over stomatal conductance,with stomatal density being secondary. An example of this can befound in experiments conducted on Arabidopsis over-expressingthe STOMATAL DENSITY AND DISTRIBUTION 1 (SDD1)gene, resulting in plants with a 40% reduction in stomataldensity, and the Arabidopsis sdd1-1 mutant (Berger & Altmann,2000), in which stomatal density is increased to 300% of thatof wild type. Under growth conditions, no differences instomatal conductance or assimilation rate were observed inthe over-expressers and the sdd1-1 mutants compared withwild type. Lower stomatal density was compensated for by anincrease in aperture and, conversely, reduced stomatal aperturecompensated for increased stomatal density (Bussis et al.,2006). It should be mentioned that, although mutants maybe identified as ‘drought resistant’ or with ‘increased water use

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efficiency’, such traits may not be evident when they aregrown in competitive environments. Basco et al. (2008) recentlyreported that Arabidopsis ABA oversensitive mutants, whichdisplay enhanced stomatal closure, could not compete withwild type for water when the plants were grown together. Suchfindings also have significant implications for screeningprotocols when attempting to identify mutants (Basco et al.,2008). It is also important to note that screening plants forincreased water use efficiency should include measurements ofphotosynthetic performance in relation to stomatal behaviour,as reduced stomatal conductance can decrease water use butalso limit photosynthetic carbon assimilation.

In conjunction with advances in molecular biology,substantial progress has been made in technology andmethodology. The use of thermal imagery (Jones, 1999, 2004;Jones et al., 2002) in combination with chla fluorescence(Chaerle et al., 2007) has the potential to determine instanta-neous water use efficiency, and is not only a potential screeningtool allowing determination of both photosynthetic performanceand stomatal behaviour but also a powerful approach toelucidating correlations between stomatal behaviour andphotosynthetic capacity. An example of combined chlorophyllfluorescence imaging and thermography is shown in Fig. 5.Extreme treatments, in which stomata in one area of the leaf

were blocked with grease to prevent stomatal conductance anda vein was severed to prevent uptake of DCMU, were used toshow the relationship between PSII operating efficiency andstomatal conductance. Images of showing spatial andtemporal resolution of PSII operating efficiency were com-pared with thermal images of leaf temperature, which is modu-lated by stomatal behaviour and other environmental factors(i.e. in general, the greater the stomatal conductance the greaterthe evaporative cooling of the leaf and the lower the leaf tempera-ture). From the images it is apparent where stomatal behaviouris influencing PSII operating efficiency and vice versa.

VII. Concluding remarks and future direction

Stomatal research over the past few decades has revealed acomplicated network of osmoregulatory and signalling pathwaysin guard cells (e.g. Li et al., 2006). It appears that these highlyplastic cells have the capability to alter mechanisms of responsedepending upon environmental growth and experimentalconditions, complicated further by time of day and pretreat-ments (Zeiger et al., 2002), all of which appear to be speciesdependent. Such flexibility gives stomata the necessarycapability to maintain a regulatory role in plant water statusand photosynthetic capacity. This review has concentrated onguard cell chloroplast photosynthesis and in particular Calvincycle function, a highly controversial topic (Outlaw, 1989,2003), with evidence for and against functional guard cellphotosynthetic regulation of stomatal behaviour. Recent researchconducted on antisense SBPase plants suggests guard cellphotosynthesis and/or carbon reduction may play a role instomatal responses to red light (Lawson et al., 2008). However,at the same time, work on antisense Rubisco and b6f plantscasts doubt on any role for guard cell photosynthesis, includingthe production of ATP, in red light-induced opening (e.g vonCaemmerer et al., 2004; Baroli et al., 2008). Discrepanciesin results and conclusions regarding the role of guard cellchloroplasts in stomatal function are probably attributable to theunique plasticity of guard cells, which can make interpretationsdifficult, with often opposing conclusions in different laboratoriesin which research was conducted under different conditions(see Zeiger et al., 2002). Stomatal research in the future shouldtherefore take into account the time of day experiments areconducted, the conditions under which the plants are grownand the type of material used, as all of these factors can impacton stomatal responses, signalling pathways, and solutes requiredfor osmoregulation of stomatal aperture.

The current transition towards using mutants and transgenicplants along with the identification of gene trap lines (Galbiatiet al., 2008) opens a new window of opportunity to pursuedifferent avenues of research to answer some of the manyquestions that still remain regarding guard cell metabolism.To date, attention has focused on photosynthetic pathways inguard and mesophyll cells, and to a certain extent the oxidativephosphorylation pathway has been neglected. Transgenic plants

Fig. 4 Predicted sensitivity of stomatal conductance (gs) to changes in pore dimension and frequency within empirically derived ranges. Effects on gs of adjusting each anatomical character within its estimated range were calculated following equations of Lawson (1997), Weyers & Lawson, (1997) and Lawson & Morison (2004). The analysis uses typical ranges of values derived from observations of Phaseolus vulgaris (Lawson, 1997): stomatal aperture, 0–15 µm; stomatal density, 35–65 mm−2; pore length, 33–40 µm; pore depth, 15–25 µm. Values within each range were used to calculate stomatal conductance using the following equation: 1/rs = (d + 2c)/(Dw × As × SD), where rs is stomatal resistance, d is pore depth (mm), c is an end correction (see Weyers & Meidner, 1990), Dw is water diffusivity in air (mm2 s−1), As is pore area (mm2), and SD is stomatal density (mm−2). The vertical lines represent the gs obtained using the median values for each variable and was calculated at 346 mmol m−2 s−1.

′ ′F Fq m/

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and mutants provide an ideal opportunity to determine therole of this pathway in stomatal sensory and response mecha-nisms. The development and discovery of guard cell specificpromoters (see Yang et al., 2008) will allow manipulation ofguard cell metabolism without disruption of mesophyllphotosynthetic metabolism. Such systems will hopefullyprovide a probe that will help to fully elucidate the linkbetween mesophyll photosynthesis and stomatal conductance.Microarray and proteomic technology allows gene expressionpatterns involved in signal transduction pathways to be identifiedand assessed under different environmental conditions andstresses (Coupe et al., 2006). Leonhardt et al. (2004) havedemonstrated the power of microarray technology comparingthe expression profiles of guard and mesophyll cells. They notedthat, when leaves were sprayed with ABA, there was repressionof many of the enzymes involved in guard cell metabolism,including a decrease in PEPc transcript, which agrees withearlier work reporting decreased PEPc activity under drought(Kopka et al., 1997). Transcriptomic analysis can also identifytranscription factors that are necessary for stomatal movementmediating stomatal responses to light and darkness (Gray,2005; see review by Casson & Gray, 2008).

To date, most stomatal research has concentrated on plantspecies very familiar to stomatal biologists, but there are still

numerous gaps in our knowledge regarding stomatal behaviourin CAM and grass species. C3/CAM intermediates may providean ideal opportunity to uncover light and CO2 responses aswell as induction of specific genes or signalling pathways.Stomata respond to numerous environmental stimuli, yetmost studies are conducted in isolation. There is a desperateneed to determine the hierarchy of stomatal responses, andthe influence of combined factors on stomatal behaviour,response and signalling mechanisms.

Although significant advances in the understanding ofguard cell function and stomatal responses have been madeover the last century, many gaps in our knowledge remainregarding guard cell metabolism and its role in stomatalbehaviour. The use of antisense techniques in conjunction withguard cell-specific promoters, and modifications of guard cellchloroplast metabolism, coupled with in situ measurements ofphotosynthetic performance, stomatal function and responsesto various stimuli, may provide the key to ascertaining theroles of these highly conserved organelles.

Acknowledgements

I would like to thank several colleagues for their contributions,ideas and discussion over the course of my past and current

Fig. 5 Chlorophyll fluorescence (a, b) and thermal (c, d) images of a sycamore leaf fed with 3-(3,4-dichlorophenyl)-1,1-dimethylurea (DCMU) through the transpiration stream. An area of stomata was blocked on one half of the leaf by applying a patch of grease, and a major vein was severed on the other half. The patch increased leaf temperature (c), and reduced the quantum efficiency of photosystem II (PSII) photochemistry (F′q/F′m, where F′q is the difference between maximum fluorescence in the light adapted state (F′m) and steady state fluorescence in the light (F′) (a). After DCMU feeding (b, d), F′q/F′m was reduced and leaf temperature was increased. However, DCMU was not distributed where the vein was severed so F′q/F′m remained high and the leaf temperature was lower. Under the patch, there was little transpiration and DCMU uptake, and therefore F′q/F′m remained high even though the CO2 supply was limited, indicating that photorespiration was the sink for the products of electron transport Scale bars represent 20 mm (unpublished data of T. Lawson, J. I. L. Morison and N. R. Baker).

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research. In particular Dr James I. L. Morison and ProfessorNeil R. Baker are acknowledged for the opportunity to workin their laboratory imaging chlorophyll fluorescence in guardcells. My initial interest in the stomatal regulation of gasexchange was inspired during work for a PhD with DrJonathan Weyers (Dundee University), at which time I wasencouraged to carry out my first research on this subject.Professors Christine Raines and Susanne von Caemmererhave provided many useful discussions. Dr Tanja Hofmann isgratefully acknowledged for critical reading of the manuscript.I would also like to thank three anonymous reviewers for theircomments, which have greatly enhanced the manuscript.Work on chlorophyll fluorescence imaging in guard cells wasfunded by a BBSRC grant awarded to Dr James I. L. Morisonand Professor Neil R. Baker, whilst recent work on transgenicplants was funded by the University of Essex.

References

Aharoni A, Dixit S, Jetter R, Thoenes E, van Arkel G, Pereira A. 2004. The SHINE clade of AP2 domain transcription factors activates was biosynthesis, alters cuticle properties and confers drought tolerance when over expressed in Arabidopsis. Plant Cell 16: 2463–2480.

Allaway WG. 1973. Accumulation of malate in guard cells of Vicia faba during stomatal opening. Planta 110: 63–70.

Allaway WG, Milthorpe FL. 1976. Structure and functioning of stomata. In: Kozlowski TT, ed. Water deficits and plant growth, Vol. IV. New York, NY, USA: Academic Press, 57–102.

Allaway WG, Setterfield G. 1972. Ultrastructural observations on guard cells of Vicia faba and Allium porrum. Canadian Journal of Botany 50: 1405–1413.

Asai N, Nakajima N, Tamaoki M, Kamada H, Kondo N. 2000. Role of malate synthesis mediated by phophoenolpyruvate carboxylase in guard cell in the regulation of stomata movement. Plant and Cell Physiology 41: 10–15.

Assmann SM. 1993. Signal transduction in guard cells. Annual Review of Cell Biology 9: 345–375.

Assmann SM. 1999. The cellular basis of guard cell sensing of rising CO2. Plant, Cell & Environment 22: 629–637.

Assmann SM, Shimazaki K. 1999. The multisensory guard cell: stomatal responses to blue light and abscisic acid. Plant Physiology 119: 809–815.

Assmann SM, Wang X-Q. 2003. From milliseconds to millions of years: guard cells and environmental responses. Current Opinions in Plant Biology 4: 421–428.

Assmann SM, Zeiger E. 1987. Guard cell bioenergetics. In: Zeiger E, Farquhar G, Cown IR, eds. Stomatal function. Stanford, CA, USA: Stanford University Press, 163–194.

Baiser M, Hemmann G, Holman R, Corke F, Card R, Smith C, Rook F, Bevan MW. 2004. Characterisation of mutants in Arabidopsis showing increased sugar-specific gene expression, growth and development responses. Plant Physiology 134: 81–91.

Baker NR. 2008. Chlorophyll fluorescence: a probe of photosynthesis in vivo. Annual Review of Plant Biology 59: 89–113.

Baker NR, Oxborough K, Lawson T, Morison JIL. 2001. High resolution imaging of photosynthetic activities of tissues, cells and chloroplasts in leaves. Journal of Experimental Botany 52: 615–621.

Baroli I, Price D, Badger MR, von Caemmerer S. 2008. The contribution of photosynthesis to the red light response of stomatal conductance. Plant Physiology 146: 737–747.

Basco R, Janda T, Galiba G, Papp I. 2008. Restricted transpiration may not result in improved drought tolerante in a competitive environment for water. Plant Science 174: 200–204.

Berger D, Altmann T. 2000. A subtilisin-like serine protease involved in the regulation of stomatal density and distribution in Arabidopsis thaliana. Genes and Development 14: 1119–1131.

Black CC, Osmond CB. 2003. Crassulacean acid metabolism photosynthesis: ‘Working the night shift’. Photosynthesis Research 76: 329–341.

Boyer JS. 1982. Plant productivity and environment. Science 218: 443–448.Bradbury M, Baker NR. 1981. Analysis of the slow phases of the in vivo

chlorophyll fluorescence induction curve. Changes in the redox state of photosystem II electron acceptors and fluorescence emission from photosystem I and II. Biochimica et Biophysica Acta 63: 542–551.

Brownlee C. 2001. The long and the short of stomatal density signals. Trends in Plant Science 6: 441–442.

Buckley TN. 2005. The control of stomata by water balance. New Phytologist 168: 275–292.

Buckley TN, Mott KA, Farquhar GD. 2003. A hydromechanical and biochemical model of stomatal conductance. Plant, Cell & Environment 26: 1767–1785.

Bussis D, von Groll U, Fisahn J, Altmann T. 2006. Stomatal aperture can compensate altered stomatal density in Arabidopsis thaliana at growth light conditions. Functional Plant Biology 33: 1037–1043.

von Caemmerer S, Lawson T, Oxborough K, Baker NR, Andrews TJ, Raines CA. 2004. Stomatal conductance does not correlate with photosynthetic capacity in transgenic tobacco with reduced amounts of Rubisco. Journal of Experimental Botany 55: 1157–1166.

Cardon ZG, Berry J. 1992. Effects of O2 and CO2 concentration on the steady-state fluorescence yield of single guard-cell pairs in intact leaf-disks of Tradescantia albiflora – evidence for Rubisco-mediated CO2 fixation and photorespiration in guard-cells Plant Physiology 99: 1238–1244.

Casson S, Gray JE. 2008. Influence of environmental factors on stomatal development. New Phytologist 178: 9–23.

Chaerle L, Leinonen I, Jones HG, van Der Staeten D. 2007. Monitoring and screening plant populations with combined thermal and chlorophyll fluorescence imaging. Journal of Experimental Botany 58: 773–784.

Chen Z, Gallie DR. 2004. The ascorbic acid redox state controls guard cell signalling and stomatal movement. The Plant Cell 16: 1143–1162.

Cotelle V, Pierre JN, Vavasseur A. 1999. Potential strong regulation of guard cell phosphoenolpyruvate carboxylase through phosphorylation. Journal of Experimental Botany 50: 777–783.

Coupe SA, Palmer BG, Lake JA, Overy SA, Oxborough K, Woodward FI, Gray JE, Quick WP. 2006. Systemic signalling of environmental cues in Arabidopsis leaves. Journal of Experimental Botany 57: 329–341.

Cousins AB, Baroli I, Badger MR, Ivakov A, Lea PJ, Leegood RC, von Caemmerer S. 2007. The role of phosphoenolpyruvate carboxylase during C4 photosynthetic isotope exchange and stomatal conductance. Plant Physiology 145: 1006–1017.

Cowan IR, Troughton JH. 1971. The relative role of stomatal in transpiration and assimilation. Planta 97: 325–336.

Cui XH, Hai FS, Chen H, Chen J, Wang XC. 2008. Expression of the Vicia faba VfPIP1 gene in Arabidopsis thaliana plants improves their drought resistance. Journal of Plant Research 121: 207–214.

D’Amelio ED, Zeiger E. 1988. Diversity of guard cell plastids of the Orchidaceae: a structural and functional study. Canadian Journal of Botany 66: 257–271.

Doi M, Shigenaga A, Emi T, Kinoshita T, Shimazaki K. 2004. A transgene encoding a blue-light receptor, phot1, restores blue-light responses in the Arabidopsis phot1 phot2 double mutant. Journal of Experimental Botany 55: 517–523.

Doi M, Shimazaki K-I. 2008. The stomata of the fern Adiantum capillus-veneris do not respond to CO2 in dark and open by photosynthesis in guard cells. Plant Physiology 147: 922–930.

Doi M, Wada M, Shimazaki K-I. 2006. The fern Adiantum capillus-veneris lacks stomatal responses to blue light. Plant Cell Physiology 47: 748–755.

Tansley review


Review30

Eckert M, Kaldenhoff R. 2000. Light-induced stomatal movement of selected Arabidopsis thaliana mutants. Journal of Experimental Botany 51: 1435–1442.

Edwards D, Davies KL, Axe L. 1992. A vascular conducting strand in the early land plant Cooksonia. Nature 357: 683–685.

Edwards D, Kerp H, Hess H. 1998. Stomata in early land plants: an anatomical and ecophysiology approach. Journal of Experimental Botany 49: 255–278.

Ewert MS, Outlaw WH Jr, Zhang S, Aghoram K, Riddle KA. 2000. Accumulation of an apoplastic solute in the guard cell wall is sufficient to exert a significant effect on transpiration in Vicia faba leaflets. Plant Cell and Environment 23: 195–203.

Farquhar GD, Sharkey TD. 1982. Stomatal conductance and photosynthesis. Annual Review of Plant Physiology 33: 317–345.

Farquhar GD, Wong SC. 1984. An empirical model of stomatal conductance. Australian Journal of Plant Physiology 11: 191–210.

Fischer RA. 1968. Stomatal opening: role of potassium uptake by guard cells. Science 160: 784–785.

Fischer RA, Hsiao TC. 1968. Stomatal opening: in isolated epidermal strips of Vicia faba. II. Responses to KCL concentration and the role of potassium absorption. Plant Physiology 43: 1953–1958.

Franks P, Farqhuar GD. 2007. The mechanical diversity of stomata and its significance in gas-exchange control. Plant Physiology 143: 78–87.

Frechilla S, Talbott LD, Zeiger E. 2002. The CO2 response of Vicia faba guard cells acclimates to growth environment. Journal of Experimental Botany 53: 545–550.

Frechilla S, Talbott LD, Zeiger E. 2004. The blue light-specific response of Vicia faba stomata acclimates to growth environment. Plant Cell Physiology 45: 1709–1714.

Frechilla S, Zhu J, Talbott LD, Zeiger E. 1999. Stomata from npq1, a zeaxanthin-less Arabidopsis mutant, lack a specific response to blue light. Plant Cell Physiology 40: 949–954.

Fujino M. 1967. Role of adenosine triphosphate and adenosine triphosphatase in stomatal movements. Science Bulletin of the Faculty of Education, Nagasaki University 18: 1–47.

Galbiati M, Simoni L, Pavesi G, Cominelli E, Francia P, Vavasseur A, Nelson T, Bevan M, Tonelli C. 2008. Gene trap lines identifiy Arabidopsis genes expressed in stomatal guard cells. Plant Journal 53: 750–762.

Gautier H, Vavasseur A, Pierre G, Lasceve G. 1991. Relationship between respiration and photosynthesis in guard cell and mesophyll cell protoplasts of Commelina communis L. Plant Physiology 95: 636–641.

Gehlen J, Panstruga R, Smets H, Merkelbach S, Kleines M, Porsch P, Fladung M, Becker I, Rademacher T, Hausler RE et al. 1996. Effects of altered phosphoenolpyruvate carboxylase activities on transgenic C3 plant Solanum tuberosum. Plant Molecular Biology 32: 831–848.

Goh C-H, Schreiber U, Hedrich R. 1999. New approach of monitoring changes in chlorophyll a fluorescence of single guard cells and protoplasts in response to physiological stimuli. Plant, Cell & Environment 22: 1057–1070.

Gotow K, Taylor S, Zeiger E. 1988. Photosynthetic carbon fixation in guard cell protoplasts of Vicia faba L. Plant Physiology 86: 700–705.

Gray JE. 2005. Guard cells: transcription factors regulate stomatal movements. Current Biology 15: 593–595.

Hampp R, Outlaw WH Jr. 1987. Microanalysis in plant biochemistry. Naturwissenschaften 74: 431–438.

Hampp R, Outlaw WH Jr, Tarczynski MC. 1982. Profile of basic carbon pathways in guard cells and other leaf cells of Vicia faba L. Plant Physiology 70: 1582–1585.

Hashimoto M, Negi J, Young J, Israelsson M, Schroeder JI, Iba K. 2006. Arabidopsis HT1 kinase controls stomatal movements in response to CO2. Nature Cell Biology 8: 391–397.

Hedrich R, Marten I. 1993. Malate induced feedback regulation of plasma membrane anion channels would provide a CO2 sensor to guard cells. EMBO 12: 897–901.

Hedrich R, Marten I, Lohse G, Dietrich P, Winter H, Lohaus G, Heldt H-W. 1994. Malate-sensitive anion channels enable guard cells to sense changes in the ambient CO2 concentration. Plant Journal 6: 741–748.

Hetherington AM, Woodward FI. 2003. The role of stomata in sensing and driving environmental change. Nature 424: 901–908.

Hipkins ME, Fitzimons PJ, Weyers JDB. 1983. The primary processes of photosystem II in purified guard-cell and mesophyll-cell protoplasts from Commelina communis L. Planta 159: 554–560.

Hite DRC, Outlaw WH Jr, Tarczynski MC. 1993. Elevated levels of both sucrose-phosphate synthase and sucrose synthase in Vicia guard cells indicate cell-specific carbohydrate interconversion. Plant Physiology 101: 1217–1221.

Hu HH, Dai MQ, Yao JL, Xiao BZ, Li XH, Zhang QF, Xiong LZ. 2006. Overexpressing a NAM, ATAF, and CUC (NAC) transcription factor enhances drought resistance and salt tolerance in rice. Proceedings of the National Academy of Sciences, USA103: 12987–12992.

Hudson GS, Evans JR, von Caemmerer S, Arvidsson YBC, Andrews TJ. 1992. Reduction of ribulose-1,5-bisphosphate carboxylase/oxygenase content by antisense RNA reduces photosynthesis in transgenic tobacco plants. Plant Physiology 98: 294–302.

Humble GD, Raschke K. 1971. Stomatal opening quantitatively related to potassium transport. Evidence from electron probe analysis. Plant Physiology 48: 447–453.

Imamura S. 1943. Untersuchungen uber den mechanismus der turgorschwandung der spaltoffnungesschliesszellen. Japanese Journal of Botany 12: 82–88.

Inoue S-I, Kinoshita T, Matsumoto M, Nakayama K, Doi M, Shimazaki K-I. 2008. Blue light-induced autophosphorylation of phototropin is a primary step for signaling. PNAS 105: 5626–5631.

Jakab G, Ton J, Flors V, Zimmerli L, Metraux J-P, Mauch-Mani B. 2005. Enhancing Arabidopsis salt and drought stress tolerance by chemical priming for its abscisic acid responses. Plant Physiology 139: 267–274.

Jia W, Zhang J. 2008. Stomatal movement and long-distance signaling in plants. Plant Signaling & Behaviour 10: 772–777.

Johansson F, Sommarin M, Larsson C. 1993. Fusicoccin activates the plasma membrane H+-ATPase by a mechanism involving the C-terminal inhibitory domain. The Plant Cell 5: 321–327.

Jones HG. 1976. Crop characteristics and the ratio between assimilation and transpiration. Journal of applied ecology 13: 605–622.

Jones HG. 1977. Transpiration in barley lines with differing stomatal frequency. Journal of Experimental Botany 28: 162–168.

Jones HG. 1987. Breeding for stomatal characters. In: Zeiger E, Farquhar GD, Cowan IR, eds. Stomatal function. Stanford, CA, USA: Stanford University Press, 431–443.

Jones HG. 1992. Plants and microclimate: a quantitative approach to environmental plant physiology. Cambridge, UK: Cambridge University Press.

Jones HG. 1999. Use of thermography for quantitative studies of spatial and temporal variation of stomatal conductance over leaf surfaces. Plant, Cell & Environment 22: 1043–1055.

Jones HG. 2004. Application of thermal imaging and infrared sensing in plant physiology and ecophysiology. Advances in Botanical Research 41: 107–163

Jones HG, Stoll M, Santos T, de Sousa C, Chaves MM, Grant OM. 2002. Use of infrared thermography for monitoring stomatal closure in the field: application to grapevine. Journal of Experimental Botany 53: 2249–2260.

Kang Y, Outlaw WH Jr, Anderson PC, Fiore GB. 2007a. Guard cell apoplastic sucrose concentration – a link between leaf photosynthesis and stomatal aperture size in apoplastic phloem loader Vicia faba L. Plant, Cell & Environment 30: 551–558.

Kang Y, Outlaw WH Jr, Fiore GB, Riddle KA. 2007b. Guard cell apoplastic photosynthate accumulation corresponds to a phloem-loading mechanisms. Journal of Experimental Botany 58: 4061–4070.

Karlsson PE. 1986. Blue light regulation of stomata in (Triticum aestivum) seedlings: I. Influence of red background illumination and initial conductance level. Physiologia Plantarum 66: 202–206.

Tansley review


Review 31

Karlsson PE, Hoglund H-O, Klockare R. 1983. Blue light induces stomatal transpiration in wheat seedlings with chlorophyll deficiency caused by SAN 9789. Physiologia Plantarum 57: 417–421.

Kautsky H, Franck U. 1943. Chlorophyllfluoreszenz und Kohlensaureassimilation. Biochemische Zeitschrift 315: 139–232.

Kinoshita T, Doi M, Suetsugu N, Kagawa T, Wada M, Shimazaki K. 2001. Phot1 and phot2 mediate blue light regulation of stomatal opening. Nature 414: 656–660.

Kinoshita T, Shimazaki K. 1999. Blue light activates the plasma membrane H+-ATPase by phosphorylation of the C-terminus in stomatal guard cells. EMBO Journal 18: 5548–5558.

Kinoshita T, Shimazaki K, Nishimura M. 1993. Phosphorylation and dephosphorylation of guard-cell proteins from Vicia faba L. in response to light and dark. Plant Physiology 102: 917–923.

Klein M, Geiser M, Suh SJ, Kolukisaoglu HU, Azevedo L, Plaza S, Curtis MD, Richter A, Weder B, Schulz B et al. 2004. Disruption of AtMRP4, a guard cell plasma membrane ABCC-type ABC transporter, leads to deregulation of stomatal opening and increased drought susceptibility. Plant Journal 39: 219–236.

Kopka J, Provart NJ, Muller-Rober B. 1997. Potato guard cells respond to drying soil by a complex change in the expression of genes related to carbon metabolism and turgor regulation. Plant Journal 11: 871–882.

Kuiper PJC. 1964. Dependence upon wavelength of stomatal movement in epidermal tissue of Senecio odoris. Plant Physiology 39: 952–955.

Lake JA, Quick WP, Beerling DJ, Woodward FI. 2001. Signals from mature to new leaves. Nature 411: 154–155.

Laporte MM, Shen B, Tarczynski C. 2002. Engineering for drought avoidance: expression of maize NADP-malic enzyme in tobacco results in altered stomatal function. Journal of Experimental Botany 53: 699–705.

Lasceve G, Leymarie J, Vasasseur A. 1997. Alterations in light-induced stomatal opening in the starch-deficient mutant of Arabidopsis thaliana L. deficient in chloroplasts phophoglucomutase activity. Plant, Cell & Environment 20: 350–358.

Laurerer M, Saftic D, Quick WP, Labate C, Fichtner K, Schulze E-D, Rodermel SR, Bogorad L, Stitt M. 1993. Decreased ribulose-1,5-bisphosphate carboxylase-oxygenase in transgenic tobacco transformed with ‘antisense’ rbcS. VI. Effect on photosynthesis in plants grown at different irradiance. Planta 190: 332–345.

Lawson T. 1997. Heterogeneity in stomatal characteristics. PhD thesis. Dundee, UK: University of Dundee.

Lawson T, Lefebvre S, Baker NR, Morison JIL, Raines C. 2008. Reductions in mesophyll and guard cell photosynthesis impact on the control of stomatal responses to light and CO2. Journal of Experimental Botany 59: 3609–3619.

Lawson T, Morison JIL. 2004. Stomatal function and Physiology. In: Hemsley AR, Poole I, eds. The evolution of plant physiology; from whole plants to ecosystem. Cambridge, UK: Elsevier Academic Press, 217–242.

Lawson T, Oxborough K, Morison JIL, Baker NR. 2002. Responses of photosynthetic electron transport in stomatal guard cells and mesophyll cells in intact leaves to light, CO2, and humidity. Plant Physiology 128: 52–62.

Lawson T, Oxborough K, Morison JIL, Baker NR. 2003. The response of guard cell photosynthesis to CO2, O2, light and water stress in a range of species are similar. Journal of Experimental Botany 54: 1734–1752.

Lee DM, Assmann SM. 1992. Stomatal responses to light in the facultative Crassulacean acid metabolism species, Portulacaria afra. Physiologica plantarum 85: 35–42.

Lee J-S, Bowling DJF. 1995. Influence of the mesophyll on stomatal opening. Australian Journal of Plant Physiology 22: 357–363.

Leonhardt N, Kwak JM, Robert N, Waner D, Leonhardt G, Schroeder J. 2004. Microarray expression analyses of Arabidopsis guard cells and isolation of a recessive abscisic acid hypersensitive protein phosphatase 2C mutant. The Plant Cell 16: 596–615.

Li S, Assmann SM, Albert R. 2006. Predicting essential components of signal transduction networks: a dynamic model of guard cell abscisic acid signaling. PLoS Biology 4: 1732–1748.

Lloyd FE. 1908. The physiology of stomata. Publication of the Carnegie Institution of Washington No. 82. Washington, DC, USA: Carnegie Institution of Washington.

Lu P, Outlaw WH Jr, Smith BD, Freed GA. 1997. A new mechanism for the regulation of stomatal aperture size in intact leaves: accumulation of mesophyll-derived sucrose in guard cell walls of Vicia faba. Plant Physiology 114: 109–118.

Lu P, Zhang SQ, Outlaw WH Jr, Riddle KA. 1995. Sucrose: a solute that accumulates in the guard-cell-apoplast and guard cell symplast of open stomata. FEBS Letters 326: 180–184.

Lurie S. 1977. Photochemical properties of guard cell chloroplasts. Plant Science Letters 10: 219–223.

Ma SY, Wu WH. 2007. AtcPK23 functions in Arabidopsis responses to drought and salt stresses. Plant Molecular Biology 65: 511–518.

MacRobbie EAC. 1987. Ionic relations of guard cells. In: Zeiger E, Farquhar G, Cown IR, eds. Stomatal function. Stanford, CA, USA: Stanford University Press, 125–162.

MacRobbie EAC, Lettau J. 1980a. Ion content and aperture in ‘isolated’ guard cells of Commelina communis L. Journal of Membrane Biology 53: 199–205.

MacRobbie EAC, Lettau J. 1980b. Potassium content and aperture of ‘intact’ stomatal and epidermal cells of Commelina communis L. Journal of Membrane Biology 56: 249–256.

Madavhan S, Smith BN. 1982. Localization of ribulose bisphosphate carboxylase in the guard cells by an indirect immunofluorescence technique. Plant Physiology 69: 273–277.

Mansfield TA, Hetherington AM, Atkinson CJ. 1990. Some current aspects of stomatal physiology. Annual Review in Plant Physiology and Plant Molecular Biology 41: 55–75.

Mansfield TA, Majernik O. 1970. Can stomata play a part in protecting plants against air pollutants? Environmental Pollution 1: 149–154.

Marten H, Hedrich R, Roelfsema MRG. 2007. Blue light inhibits guard cell plasma membrane anion channels in a phototropin-dependent manner. Plant Journal 50: 29–39.

Marten H, Hyun T, Gomi K, Seo S, Hedrich R, Roelfsema MRG. 2008. Silencing of NtMPK4 impairs CO2-induced stomatal closure, activation of anion channels and cytosolic Ca2+ signals in Nicotiana tbacum guard cells. Plant Journal 55: 698–708.

Masle J, Gilmour SR, Farquhar GD. 2005. The ERECTA gene regulates plant transpiration efficiency in Arabidopsis. Nature 436: 866–870.

Mawson BT. 1993a. Regulation of blue-light-induced proton pumping by Vicia faba L. guard cell protoplasts: energetic contributions by chloroplastic and mitochondrial activities. Planta 191: 293–301.

Mawson BT. 1993b. Modulation of photosynthesis and respiration in guard and mesophyll cell protoplasts by oxygen concentration. Plant, Cell & Environment 16: 207–214.

Mawson BT, Zeiger E. 1991. Blue light modulation of chlorophyll a fluorescence transient in guard cell chloroplasts. Plant Physiology 96: 753–760.

Meidner H, Mansfield TA. 1968. Physiology of stomata. Glasgow, UK: Blackie.Melis A, Zeiger E. 1982. Chlorophyll a fluorescence transient in mesophyll

and guard cells: modulation of guard cell photophosphorylation by CO2. Plant Physiology 69: 642–647.

Messinger SM, Buckley TN, Mott KA. 2006. Evidence for involvement of photosynthetic processes in the stomatal response to CO2. Plant Physiology 140: 771–778.

Morison JIL. 1987. Intercellular CO2 concentration and stomatal response to CO2. In: Zeiger E, Farquhar G, Cown IR, eds. Stomatal function. Stanford, CA, USA: Stanford University Press, 229–251.

Morison JIL. 2003. Plant water use, stomatal control. In: Stewart BA, Howell TA, ed. Encyclopedia of water science. New York, NY, USA: Marcel Dekker Inc., 680–685.

Tansley review


Review32

Morison JIL, Baker NR, Mullineaux PM, Davies WJ. 2007. Improving water use in crop production. Philosophical Transactions of the Royal Society B 363: 639–658.

Mott KA. 1988. Do stomata respond to CO2 concentrations other than intercellular? Plant Physiology 86: 200–230.

Mott KA, Sibbernsen ED, Shope JC 2008. The role of the mesophyll in stomatal responses to light and CO2. Plant, Cell & Environment 31: 1299–1305.

Muschak M, Willmitzer L, Fisahn J. 1999. Gas-exchange analysis of chloroplastic fructose-1,6-bisphosphatase antisense potatoes at different air humidities and at elevated CO2. Planta 209: 104–111.

Negi J, Matsuda O, Nagasawa T, Oba Y, Takahashi H, Kawai-Yamada M, Uchimiya H, Hashimoto M, Iba K. 2008. CO2 regulation SLAC1 and its homologues are essential for anion homeostasis in plant cells. Nature 452: 483–488.

Nelson SO, Mayo JM. 1975. The occurrence of functional nonchlorophyllous guard cells in Paphiopedilum spp. Canadian Journal of Botany 53: 1–7.

Nilson SE, Assmann SM. 2007. The control of transpiration. Insights from Arabidopsis. Plant Physiology 143: 19–27.

Nunes-Nesi A, Carrari F, Gibon Y, Sulpice R, Lytovchenko A, Fisahn J, Graham J, Ratcliffe RG, Sweetlove LJ, Fernie AR. 2007. Deficiency of mitochondrial fumarase activity in tomato plants impairs photosynthesis via an effect on stomatal function. Plant Journal 50: 1093–1106.

Ogawa T, Grantz D, Boyer J, Govindjee. 1982. Effects of cations and abscisic acid on chlorophyll a fluorescence in guard cells of Vicia faba. Plant Physiology 69: 1140–1144.

Ogawa T, Ishikawa H, Shimada K, Shibata K. 1978. Synergistic action of red and blue light and action for malate formation in guard cells of Vicia faba L. Planta 142: 61–65.

Olsen RL, Pratt RB, Gump P, Kemper A, Tallman G. 2002. Red light activates a chloroplast-dependent ion uptake mechanism for stomatal opening under reduced CO2 concentrations in Vicia spp. New Phytologist 153: 497–508.

Osborne BA. 1989. Comparison of photosynthesis and productivity of Gunnera tinctoria Molina (Mirbel) with and without the phycobiont Nostoc punctiforme L. Plant, Cell & Environment 12: 941–946.

Osmond CB. 1978. Crassulacean acid metabolism: a curiosity in context. Annual Review of Plant Physiology 29: 397–414.

Outlaw WH Jr. 1980. A descriptive evaluation of quantitive histochemical methods based on pyridine nucleotides. Annual Review of Plant Physiology 31: 299–311.

Outlaw WH Jr. 1982. Carbon metabolism in guard cells. Recent Advances in Phytochemistry 17: 185–222.

Outlaw WH Jr. 1983. Current concepts on the role of potassium in stomatal movements. Physiologia plantarum 59: 302–311.

Outlaw WH Jr. 1987. A minireview: comparative biochemistry of photosynthesis in palisade cells, spongy cells and guard cells of C3 leaves. In: Biggins J, eds. Progress in photosynthesis research, Vol. IV. Dordrecht, the Netherlands: Martinus Nijhoff, 265–272.

Outlaw WH Jr. 1989. Critical examination of the quantitative evidence for and against photosynthetic CO2 fixation in guard cells. Physiologia Plantarum 77: 275–281.

Outlaw WH Jr. 1990. Kinetic properties of guard-cell phosphoenolpyruvate carboxylase. Biochemical Physiology Pfanzen 186: 317–325.

Outlaw WH Jr. 1996. Stomata: biophysical and biochemical aspects. In: Baker NR, eds. Photosynthesis and the environment. Dordrecht, the Netherlands: Kluwer Academic Publishers, 241–259.

Outlaw WH. 2003. Integration of cellular and physiological functions of guard cells. Critical Reviews in Plant Science 22: 503–529.

Outlaw WH Jr, De Vleighere-He X. 2001. Transpiration rate: an importance factor in controlling sucrose content of the guard cell apoplast of broad bean. Plant Physiology 126: 1717–51724.

Outlaw WH Jr, Kennedy J. 1978. Enzymic and substrate basis for the anaplerotic step in guard cells. Plant Physiology 62: 648–652.

Outlaw WH Jr, Lowry OH. 1977. Organic acid and potassium accumulation in guard cells during stomatal opening. Proceedings of National Academy of Sciences, USA 74: 4434–4438.

Outlaw WH Jr, Manchester J. 1979. Guard cell starch concentration quantitatively related to stomatal aperture. Plant Physiology 64: 79–82.

Outlaw WH Jr, Manchester J, Di Camelli CA, Randall DD, Rapp B, Veither GM. 1979. Photosynthetic carbon reduction pathway is absent in chloroplasts of Vicia faba guard cells. Proceedings of the National Academy of Sciences, USA 76: 6371–6375.

Outlaw WH, Mayne BC, Zenger VE, Manchester J. 1981. Presence of both photosystems in guard cells of Vicia faba L.: Implications for environmental signaling processing. Plant Physiology 67: 12–16.

Outlaw WH Jr, Tarczynski MC. 1984. Guard cell starch biosynthesis regulated by effectors of ADP-glucose pyrophosphoryase. Plant Physiology 74: 424–429.

Outlaw WH Jr, Tarczynski MC, Anderson LC. 1982. Taxonomic survey for the presence of ribulose-1,5-bisphosphate carboxylase activity in guard cells. Plant Physiology 70: 1218–1220.

Outlaw WH Jr, Zhang S. 2001. Single-cell dissection and microdroplet chemistry. Journal of Experimental Botany 52: 605–614.

Oxborough K, Baker NR. 1997. Resolving chlorophyll a fluorescence images of photosynthetic efficiency into photochemical and nonphotochemical components – calculation of qP and without measuring . Photosynthesis Research 54: 135–142.

Pallas JE Jr, Wright BG. 1973. Organic acid changes in the epidermis of Vicia faba and their implication in stomatal movement. Plant Physiology 51: 588–590.

Park KH, Hahn KW, Kang BG. 1990. Lack of correlation between senescence and stomatal aperture in Lemna gibba G3. Plant Cell Physiology 31: 731–733.

Parvathi K, Raghavendra AS. 1995. Bioenergetic processes in guard cells related to stomatal function. Physiologia Plantarum 93: 146–154.

Parvathi K, Raghavendra AS. 1997. Both Rubisco and phosphoenolpyruvate carboxylase are beneficial for stomatal function in epidermal strips of Commelina benghalensis. Plant Science 124: 153–157.

Paul MJ, Knight, JS, Habash D, Parry MAJ, Lawlor DW, Barnes SA, Loynes A, Gray JC. 1995. Reduction in phosphoribulokinase activity by antisense RNA in transgenic tobacco – effect on CO2 assimilation and growth in low irradiance. Plant Journal 7: 535–542.

Pearson CJ. 1973. Daily changes in stomatal aperture and in carbohydrates and malate within epidermis and mesophyll of leaves of Commelina cyanea and Vicia faba. Australian Journal of Biological Sciences 26: 1035–1044.

Pearson CJ, Milthorpe FL. 1974. Structure, carbon dioxide fixation and metabolism of stomata. Australian Journal of Plant Physiology 1: 221–236.

Pemadasa MA. 1981. Photocontrol of stomatal movements. Biological Review 56: 551–588.

Pemadasa MA. 1983. Effects of benzo-18-crown-6 in antagonism with abscisic acid on abaxial and adaxial stomatal opening. New Phytologist 93: 13–24.

Poffenroth M, Green DB, Tallman G. 1992. Sugar concentrations in guard cells of Vicia faba illuminated with red or blue light: analysis by high performance liquid chromatography. Plant Physiology 98: 1460–1471.

Price GD, von Caemmerer S, Evans J, Chow W, Anderson J, Bader M. 1998. Photosynthesis is strongly reduced by antisense suppression of chloroplastic cytochrome bf complex in transgenic tobacco. Australian Journal of Plant Physiology 25: 445–452.

Quick WP, Schurr U, Scheibe R, Schulze E-D, Rodermel SR, Bogorad L, Stitt M. 1991. Decreased ribulose, 1,5-bisphosphate carboxylase-oxygenase in transgenic tobacco transformed with ‘antisense’ rbcS. I. Impact on photosynthesis in ambient grown conditions. Planta 183: 542–554.

Radin JW, Hartung W, Kimball BA, Mauney JR. 1988. Correlation of stomatal conductance with photosynthetic capacity of cotton only in a CO2-enriched atmosphere: mediation by abscisic acid? Plant Physiology 88: 1058–1062.

′ ′F Fv m/′Fo

Tansley review


Review 33

Raghavendra AS, Vani T. 1989. Respiration in guard cells: pattern and possible role in stomatal function. Journal of Plant Physiology 135: 3–8.

Rao M, Anderson LE. 1983. Light and stomatal metabolism. Plant Physiology 71: 456–459.

Raschke K. 1975. Stomatal action. Annual Review of Plant Physiology 26: 309–340.

Raschke K. 1979. Movement of stomata. In: Haupt W, Feinleib ME, eds. Encyclopedia of plant physiology, Vol. 7: physiology of movements. Berlin: Springer, 387–411.

Raschke K, Dittrich P. 1977. [14C] carbon dioxide fixation by isolated leaf epidermis with stomata closed or open. Planta 134: 69–75.

Raschke K, Fellows MP. 1971. Stomatal movement in Zea mays: shuttle of potassium and chloride between guard cells and subsidiary cells. Planta 101: 296–316.

Reckmann U, Scheibe R, Raschke K. 1990. Rubisco activity in guard cell compared with the solute requirement for stomatal opening. Plant Physiology 92: 246–253.

Reddy AR, Rama Das VS. 1986. Stomatal movement and sucrose uptake by guard cell protoplasts of Commelina benghalensis L. Plant Cell & Physiology 27: 1565–1570.

Reddy KB, Rao CR, Raghavendra AS. 1983. Change in levels of starch and sugars in epidermis of Commelina behghalensis during fusiccoccin stimulated stomatal opening. Journal of Experimental Botany 145: 1018–1025.

Ridolfi M, Fauveau ML, Label P, Garrec JP, Dreyer E. 1996. Responses to water stress in an ABA-unresponsive hybrid poplar (Populus koreana × trichocarpa cv. Peace). I. Stomatal function. New Phytologist 134: 445–454.

Ritte G, Raschke K. 2003. Metabolite export of isolated guard cell chloroplasts of Vicia faba. New Phytologist 159: 195–202.

Ritte G, Rosenfeld J, Rohrig K, Raschke K. 1999. Rates of sugar uptake by guard cell protoplasts of Pisum sativum L. related to the solute requirements for stomatal opening. Plant Physiology 121: 647–655.

Roelfsema MRG, Hanstein S, Fell HH, Hedrich R. 2002. CO2 provides an intermediate link in the red light response of guard cells. Plant Journal 32: 65–75.

Roelfsema MRG, Hedrich R. 2005. In the light of stomatal opening: new insights into ‘the Watergate’. New Phytologist 167: 665–691.

Roelfsema MRG, Konrad KR, Marten H, Psaras GK, Hartung W, Hedrich H. 2006. Guard cells in albino leaf patches do not respond to photosynthetically active radiation, but are sensitive to blue light, CO2 and abscisic acid. Plant, Cell & Environment 29: 1595–1605.

Roelfsema MRG, Steinmeyer R, Staal M, Hedrich R. 2001. Single guard cell recordings in intact plants: light-induced hyperpolarization of the plasma membrane. Plant Journal 26: 1–13.

Ruiz LP, Mansfield TA. 1994. A postulated role for calcium oxalate in the regulation of calcium ions in the vicinity of guard cells. New Phytologist 127: 473–481.

Rutter JM, Willmer CM. 1979. A light and electron microscopy study of the epidermis of Paphiopedilum spp. with emphasis on stomatal ultrastructure. Plant, Cell & Environment 2: 211–219.

Sack FD. 1987. The development and structure of stomata. In: Zeiger E, Farquhar G, Cown IR, eds. Stomatal function. Stanford, CA, USA: Stanford University Press, 59–90.

Scarth GW. 1932. Mechanism of the action of light and other factors on stomatal movements. Plant Physiology 7: 481–504.

Scarth GW, Shaw M. 1951. Stomatal movement and photosynthesis in Pelargonium I. Effect of light and carbon dioxide. Plant Physiology 26: 207–225.

Scheibe R, Reckmann U, Hedrich R, Raschke K. 1990. Malate dehydrogenases in guard cells of Pitsum sativum. Plant Physiology 93: 1358–1364.

Schnabl H. 1977. Isolation and identification of soluble polysaccharides in epidermal tissue of Allium cepa. Planta 135: 307–311.

Schnabl H. 1980. CO2 and malate metabolism in starch-containing and starch-lacking guard cell protoplasts. Planta 149: 52–58.

Schnabl H. 1981. The compartmentation of carboxylating and decarboxylating enzymes in guard cell protoplasts. Planta 152: 307–313.

Schnabl H, Elbert C, Kramer G. 1982. The regulation of the starch-malate balances during volume changes of guard cell protoplasts. Journal of Experimental Botany 33: 996–1003.

Schnabl H, Raschke K. 1980. Potassium chloride as stomatal osmoticum in Allium cepa L., a species devoid of starch in guard cells. Plant Physiology 65: 88–93.

Schnabl H, Zeiger E. 1977. The mechanism of stomatal movement in Allium cepa L., species devoid of starch in guard cells. Planta 136: 37–43.

Schreiber U, Schliwa U, Bilger W. 1986. Continuous recording of photochemical and nonphotochemical chlorophyll fluorescence quenching with a new type of modulation fluorometer. Photosynthesis Research 10: 51–62.

Schwartz A, Zeiger E. 1984. Metabolic energy for stomatal opening. Role of photophosphorylation and oxidative phosphorylation. Planta 161: 129–136.

Serna L. 2008. Coming close to a stoma ion channel. Nature 10: 509–511.Serrano E, Zeiger E, Hagiwara SE.1988. Red light stimulates an electrogenic

proton pump in Vicia guard cell protoplasts. Proceedings of the National Academy of Sciences, USA 85: 436–440.

Sharkey TD, Raschke K. 1981a. Effect of light quality on stomatal opening in leaves of xanthium strumarium L. Plant Physiology 68: 1170–1174.

Sharkey TD, Raschke K. 1981b. Separation and measurement of direct and indirect effects of light on stomata. Plant Physiology 68: 33–40.

Shimazaki K-I. 1989. Ribulosebisphosphate carboxylase activity and photosynthetic O2 evolution rate in Vicia guard cell protoplasts. Plant Physiology 91: 459–463.

Shimazaki K-I, Doi M, Assmann SM, Kinoshita T. 2007. Light regulation of stomatal movements. Annual Review of Plant Biology 58: 219–247.

Shimazaki K-I, Gotow K, Kondo N. 1982. Photosynthetic properties of guard cell protoplasts from Vicia faba L. Plant Cell Physiology 23: 871–879.

Shimazaki K-I, Kondo N. 1987. Plasma membrane H+-ATPase in guard-cell protoplasts from Vicia faba L. Plant Cell Physiology 28: 893–900.

Shimazaki K-I, Okayama S. 1990. Calvin-Benson cycle enzymes in guard cell protoplast and their role in stomatal movements. Biochemie Physiologie Pflanzen 186: 327–331.

Shimazaki K-I, Terada J, Tanaka K, Kondo N. 1989. Calvin-Benson cycle enzymes in guard-cell protoplasts from Vicia faba L. Plant Physiology 90: 1057–1064.

Shimazaki K-I, Zeiger E. 1985. Cyclic and noncyclic phosphorylation in isolated guard cell protoplasts from Vicia faba L. Plant Physiology 78: 211–214.

Shimazaki K-I, Zeiger E. 1987. Red light-dependent CO2 uptake and oxygen evolution in guard cell protoplasts of Vicia faba L.: evidence for photosynthetic CO2 fixation. Plant Physiology 84: 7–9.

Srivastava A, Zeiger E. 1992. Fast fluorescence quenching from isolated guard cell chloroplasts of Vicia faba is induced by blue light and not by red light. Plant Physiology 100: 1562–1566.

Srivastava A, Zeiger E, Strasser RJ. 1998. Light sensing by guard cells chloroplasts: large antenna size and high electron transport rates. In: Garad G, eds. Photosynthetic mechanisms and effects, Vol. 5. New York, NY, USA: Kluwer, 2813–2816.

Stadler R, Buttner M, Ache P, Hedrich R, Ivashikina N, Melzer M, Shearson SM, Smith SM, Sauer N. 2003. Diurnal and light-regulated expression of AtSTP1 in guard cells of Arabidopsis. Plant Physiology 133: 528–537.

Stevens RA, Martin ES. 1978. Structural and functional aspects of stomata. 1. Developmental studies in Polypodium vulgare. Planta 142: 307–316.

Tansley review


Review34

Stitt M, Quick WP, Schurr U, Schulze E-D, Rodermel SR, Bogorad L. 1991. Decreased ribulose-1,5-bisphosphate carboxylase-oxygenase transgenic tobacco transformed with ‘antisense’ rbcS. II. Flux-control coefficients for photosynthesis in varying light, CO2 and air humidity. Planta 183: 555–566.

Stitt M, Schulze D. 1994. Does Rubisco control the rate of photosynthesis and plant growth? An exercise in molecular ecophysiology. Plant, Cell & Environment 17: 465–487.

Talbott LD, Rahveh E, Zeiger E. 2003. Relative humidity is a key factor in the acclimation of the stomatal response to CO2. Journal of Experimental Botany 54: 2141–2147.

Talbott LD, Srivastava A, Zeiger E. 1996. Stomata from growth-chamber grown Vicia faba have an enhanced sensitivity to CO2. Plant, Cell & Environment 19: 1188–1194.

Talbott LD, Zeiger E. 1993. Sugar and organic acid accumulation in guard cells of Vicia faba in response to red and blue light. Plant Physiology 102: 1163–1169.

Talbott LD, Zeiger E. 1996. Central role for potassium and sucrose in guard-cell osmoregulation. Plant Physiology 111: 1051–1057.

Talbott LD, Zeiger E. 1998. The role of sucrose in guard cell osmoregulation. Journal of Experimental Botany 49: 329–337.

Talbott LD, Zhu JX, Han SW, Zeiger E. 2002. Phytochrome and blue-light mediated stomatal opening in the orchid, Paphiopedilum. Plant and Cell Physiology 43: 639–646.

Talbott LD, Zhu J, Mawson BT, Amodeo G, Nouhi Z, Levy K, Zeiger E. 1997. Induction of CAM in Mesembryanthemum crystallinum abolishes the stomatal response to blue light and light-dependent zeaxanthin formation in guard cell chloroplasts. Plant Cell Physiology 38: 236–242.

Tallman G, Zeiger E. 1988. Light quality and osmoregulation in Vicia faba guard cells: evidence for the involvement of three metabolic pathways. Plant Physiology 88: 887–895.

Tarczynski MC, Outlaw WH Jr, Arold N, Neuhoff V, Hampp R. 1989. Electrophoretic assay for ribulose 1, 5-bisphosphate carboxylase/oxygenase in guard cells and other leaf cells of Vicia faba L. Plant Physiology 89: 1088–1093.

Taylor AR, Assmann SM. 2001. Apparent absence of a redox requirement for blue light activation of pump current in broad bean guard cells. Plant Physiology 125: 329–338.

Tichà I. 1982. Photosynthetic characteristics during ontogenesis of leaves. 7. Stomatal density and sizes. Photosynthetica 16: 375–471.

Tominaga M, Kinoshita T, Shimazaki K. 2001. Guard-cell chloroplasts provide ATP required for H+ pumping in the plasma membrane and stomatal opening. Plant Cell Physiology 42: 795–802.

Tsionsky M, Cardon ZG, Bard AJ, Jackson RB. 1997. Photosynthetic electron transport in single guard cells as measured by scanning electrochemical microscopy. Plant Physiology 113: 895–901.

Ueno O. 2001.Ultrastructural localization of photosynthetic and photorespiratory enzymes in epidermal, mesophyll, bundle sheath, and vascular bundle cells of the C4 dicot Amaranthus viridis. Journal of Experimental Botany 52: 1003–1013.

Vahisalu T, Kollist H, Wang Y-F, Nishimura N, Chan W-Y, Valerio G, Lamminmäki A, Brosché M, Moldau H, Desikan R et al. 2008. SLAC1 is required for plant guard cell S-type anion channel function in stomatal signalling. Nature 452: 487–493.

Vavasseur A, Raghavendra AS. 2005. Guard cell metabolism and CO2 sensing. New Phytologist 165: 665–682.

Wang Y, Chen X, Xiang C-B. 2007. Stomatal density and bio-water saving. Journal of Intergrative Plant Biology 49: 1435–1444.

Wang Y, Noguchi KO, Terashima I. 2008. Distinct light responses of adaxial and abaxial stomata in leaves of Helianthus annuus L. Plant, Cell & Environment 31: 1307–1315.

Wang Y, Ying JF, Kuzma M, Chalifoux M, Sample A, McArthur C, Uchacz T, Sarvas C, Wan JX, Dennis DT et al. 2005. Molecular tailoring of farnesylation for plant drought tolerance and yield protection. Plant Journal 43: 413–424.

Webb AR, Baker AJ. 2002. Stomatal biology: new techniques, new challenges. New Phytologist 153: 365–369.

Weyers JDB, Lawson T. 1997. Heterogeneity in stomatal characteristics. Advances in Botanical Research 26: 317–252.

Weyers JDB, Meidner H. 1990. Methods in stomatal research. Harlow UK: Longman Scientific and Technical.

Weyers JDB, Paterson NW. 2001. Plant hormones and the control of physiological processes. New Phytologist 152: 375–407.

Weyers JDB, Travis AJ. 1981. Selection and preparation of leaf epidermis for experiments on stomatal physiology. Journal of Experimental Botany 32: 837–850.

Willmer CM. 1983. Phosphoenolpyruvate carboxylase activity and stomatal operation. Physiology Veg 21: 943–953.

Willmer CM, Dittrich P. 1974. Carbon dioxide fixation by epidermal and mesophyll tissues of Tulipa and Commelina. Planta 117: 123–132.

Willmer CM, Fricker M. 1996. Stomata, 2nd edn. London: Chapman & Hall.

Willmer CM, Pallas JE Jr, Black CC Jr. 1973. Carbon dioxide metabolism in leaf epidermal tissue. Plant Physiology 52: 448–452.

Willmer CM, Rutter JC. 1977. Guard cell malic acid metabolism during stomatal movements. Nature 269: 327–328.

Wong SC, Cowan IR, Farquhar GD. 1979. Stomatal conductance correlates with photosynthetic activity. Nature 282: 424–426.

Wong SC, Cowan IR, Farquhar GD. 1985. Leaf conductance in relation to rate of CO2 assimilation. I. Influence of nitrogen nutrition, phosphorus nutrition, photon flux density and ambient partial pressure of CO2 during ontogeny. Plant Physiology 78: 821–825.

Wu WH, Assmann SM. 1993. Photosynthesis by guard-cell chloroplasts of Vicia faba L. – effects of factors associated with stomatal movement. Plant Cell Physiology 34: 1015–1022.

Yamashita T. 1952. Influences of potassium supply upon various properties and movement of guard cell. Sielboldia Acta Biollogy 1: 51–70.

Yang CY, Chen YC, Jauh GY, Wang CS. 2005. A lily ASR protein involves abscisic acid signaling and confers drought and salt resistance in Arabidopsis. Plant Physiology 139: 836–846.

Yang Y, Costa A, Leonhardt N, Siegel RS, Schroeder JI. 2008. Isolation of strong Arabidopsis guard cell promoter and its potential as a research tool. Plant Methods 4: 1–15.

Zeiger E. 1983. The biology of guard cells. Annual Review of Plant Physiology 34: 411–475.

Zeiger E. 2000. Sensory transduction of blue light in guard cells. Trends in Plant Science 5: 183–185.

Zeiger E, Armond P, Melis P. 1980. Fluorescence properties of guard cell chloroplasts: evidence for linear electron transport and light harvesting pigments of photosystem I and II. Plant Physiology 67: 17–20.

Zeiger E, Field C. 1982. Photocontrol of the functional coupling between stomatal conductance in the intact leaf. Plant Physiology 70: 370–375.

Zeiger E, Talbott LD, Frechilla S, Srivastava A, Zhu J. 2002. The guard cell chloroplast: a perspective for the twenty-first century. New Phytologist 153: 415–424.

Zeiger E, Zhu JX. 1998. Role of zeaxanthin in blue light photoreception and the modulation of light–CO2 interactions in guard cells. Journal of Experimental Botany 49: 433–442.

Zemel E, Gepstein S. 1985. Immunological evidence for the presence of ribulose bisphosphate carboxylase in guard cell chloroplasts. Plant Physiology 78: 586–590.

Zhang X, Zhang L, Dong F, Gao J, Galbraith DW, Song C-P. 2001. Hydrogen peroxide is involved in abscisic acid-induced stomatal closure in Vicia faba. Plant Physiology 126: 1438–1448.

Zhu J, Talbott LD, Jin X, Zeiger E. 1998. The stomatal response to CO2 is linked to changes in guard cell zeaxanthin. Plant, Cell & Environment 21: 813–820.

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