Download Guard cell photosynthesis and stomatal function

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Guard cell photosynthesis and
stomatal function
Author for correspondence:
Tracy Lawson
Tel: +44 (0) 1206 873327
Fax: +44 (0) 1206 873416
Email: [email protected]
Tracy Lawson
Department of Biological Sciences, University of Essex, Wivenhoe Park, Colchester CO4 3SQ, UK
Received: 4 June 2008
Accepted: 18 September 2008
Contents
Summary
13
I.
Introduction
14
II.
Osmoregulation in guard cells
16
III.
Role of guard cell chloroplasts in
stomatal function
18
V.
VI.
IV.
Linking stomatal behaviour to mesophyll
photosynthesis
23
Stomata in relation to water use/manipulation
of behaviour
26
VII. Concluding remarks and future direction
Chlorophyll a fluorescence studies to examine
guard cell photosynthesis
22
27
Acknowledgements
28
References
29
Summary
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.
© The Author (2008).
Journal compilation © New Phytologist (2008)
Chloroplasts are a key feature of most guard cells; however, the function of these
organelles in stomatal responses has been a subject of debate. This review examines
evidence for and against a role of guard cell chloroplasts in stimulating stomatal
opening. Controversy remains over the extent to which guard cell Calvin cycle
activity contributes to stomatal regulation. However, this is only one of four possible
functions of guard cell chloroplasts; other roles include supply of ATP, blue-light signalling
and starch storage. Evidence exists for all these mechanisms, but is highly dependent
upon species and growth/measurement conditions, with inconsistencies between
different laboratories reported. Significant plasticity and extreme flexibility in guard
cell osmoregulatory, signalling and sensory pathways may be one explanation. The use
of chlorophyll a fluorescence analysis of individual guard cells is discussed in assessing
guard and mesophyll cell physiology in relation to stomatal function. Developments
in transgenic and molecular techniques have recently provided interesting, albeit
contrasting, data regarding the role of these highly conserved organelles in stomatal
function. Recent studies examining the link between mesophyll photosynthesis and
stomatal conductance are discussed. An enhanced understanding of these processes
may be fundamental in generating crop plants with greater water use efficiencies,
capable of combating future climatic changes.
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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; Fq′ / Fm′ ,
quantum efficiency of photosystem II (PSII) photochemistry; MAP, Mehler-ascorbate
peroxidase; NADPH, nicotinamide adenine dinucleotide phosphate; OAA, oxaloacetate; PEPc, phosphoenolpyruvate carboxylase; PGA, 3-phosphoglyceric acid;
Rubisco, ribulose-1,5-bisphosphate carboxylase/oxygenase; RuBPC, ribulose-1,5bisphosphate carboxylase.
I. Introduction
Stomata are small adjustable pores found in large numbers on
the surface of most aerial parts of higher plants and have been
documented in the fossil record from as early as the late
Silurian, 411 Myr ago (Edwards et al., 1992, 1998). A stoma is
formed from two specialized cells in the epidermis (guard cells)
which are morphologically distinct from general epidermal
cells and are responsible for controlling stomatal aperture
(Franks & Farqhuar, 2007). Paired guard cells, in some species
together with epidermal subsidiary cells, form the stomatal
complex (Fig. 1). Subsidiary cells can play a role in stomatal
movements either mechanically or as ion reserves (Raschke &
Fellows, 1971). In most plants stomata can be found on both
the upper (adaxial) and lower (abaxial) leaf surfaces, such leaves
being termed amphistomatous, with the majority of stomata
found on the lower surface (Tichà, 1982). In some species
(particularly trees) stomata are found only on the lower
surface (i.e. the leaf is hypostomatous), whilst some aquatic
plants (such as water lilies) have stomata only on the upper surface
(i.e. the leaf is epistomatous) (Morison, 2003). As the leaf cuticle
is almost impermeable to water and CO2, the central role of
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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stomata is regulation of gas exchange between the inside of the
leaf and the external environment (Cowan & Troughton, 1971;
Jones, 1992). Through their role in controlling transpiration,
stomata also aid in leaf cooling, metabolite fluxes, and longdistance signalling (Brownlee, 2001; Lake et al., 2001; Jia &
Zhang, 2008) as well as acting as a barrier to harmful pollutants
such as ozone and pathogens (Meidner & Mansfield, 1968;
Mansfield & Majernik, 1970).
Plants require sufficient CO2 to enter the leaf for photosynthesis whilst conserving water to avoid dehydration and
metabolic disruption. When fully open, stomatal pores only
occupy between 0.5 and 5% of the leaf surface (Hetherington
& Woodward, 2003; Morison, 2003); however, almost all the
water transpired as well as CO2 absorbed passes through these
pores. For this reason, stomatal function has significant implications for global hydrological and carbon cycles. The quest to
understand stomatal control of photosynthetic CO2 fixation
and plant water relations is becoming increasingly important
with changing climatic conditions. Knowledge of stomatal
function is critical to determine plant responses to environmental
stresses, particularly reduced water availability, and is necessary
to identify plants with decreased water use that are capable of
high yields in more extreme environments (Morison et al.,
2007). On a global scale, drought causes more yield losses
than any other single biotic or abiotic factor (Boyer, 1982),
resulting in increasing pressure for agronomists and plant
breeders to identify crop varieties that are drought tolerant for
sustainable production of food and biofuels on droughtsusceptible land. Increased knowledge of stomatal function
could provide the key to such crop improvements (Jones, 1987;
Wang et al., 2007).
The aims of this review are to examine some of the potential
functions of the guard cell chloroplasts and how these are
linked to stomatal behaviour. Contrasting evidence has led to
a number of controversies regarding guard cell chloroplast
function, in particular concerning the contribution of guard
cell photosynthesis to stomatal regulation, and several opposing
views are discussed in the following sections. Whilst there are
still gaps in our knowledge regarding stomatal regulation, as
well as sensory and signalling mechanisms, a wealth of evidence
exists on stomatal responses to various environmental stimuli,
guard cell osmoregulation and mechanisms of movement. A
© The Author (2008).
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historical account of the various osmoregulatory pathways
and solutes used in stomatal movements is given, which
demonstrates the extreme plasticity in guard cell function
and illustrates the difficulties faced by stomatal researchers
attempting to elucidate specific stomatal mechanisms. The
review also briefly examines the use of recent advancements in
modern techniques (such as antisense technology and the use
of DNA mutation) to address the function of chloroplasts
within guard cells. Recent studies using such techniques have
already revealed interesting and unexpected results, paving
the way for future research, which may allow us to fill gaps in
our understanding of the role of the guard cell chloroplasts in
stomatal regulation.
1. Stomatal regulation
Despite the controversies mentioned above, decades of research
have provided a substantial amount of information regarding
stomatal responses to various environmental stimuli. The details
of these responses, along with mechanisms of pore opening and
closing, are now widely accepted. Stomatal aperture is regulated
by 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 cell
movements is stimulated by illumination with light in the
photosynthetically effective waveband, low CO2 concentrations
and high humidity, whilst closure is promoted by darkness, low
humidity, high temperature and high CO2 concentrations
(see reviews by Assmann, 1993; Willmer & Fricker, 1996;
Outlaw, 2003; Vavasseur & Raghavendra, 2005; Shimazaki
et al., 2007) as well as plant hormones such as abscisic acid
(see review by Weyers & Paterson, 2001). However, there
are exceptions, the most obvious being crassulacean acid
metabolism (CAM) plants which in general maintain closed
stomata during the light period and open stomata in darkness
(Osmond, 1978; see Black & Osmond, 2003). Additionally,
stomata of Gunnera (Osborne, 1989) and Lemna spp. (Park
et al., 1990) are unresponsive to a number of environmental
stimuli, whilst there are also examples of several other species
that do not respond to numerous plant hormones (e.g. Ridolfi
et al., 1996; see Weyers & Paterson, 2001).
It is recognized that stomatal responses to light have at least
two components. One component is the photosynthesisindependent, specific blue-light response that saturates at low
fluence rates, and is often associated with rapid stomatal opening
(see Zeiger et al., 2002) believed to involve the activation of
a plasma membrane H+-ATPase (Kinoshita & Shimazaki,
1999; Shimazaki et al., 2007). The other component, a
photosynthesis-mediated response (termed the red-light response
here and in many other publications), saturates at high fluence
rates similar to those that saturate guard and mesophyll cell
photosynthesis and is inhibited by 3-(3,4-dichlorophenyl)-
© The Author (2008).
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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 photosynthesisdependent response can be observed under either blue or red
light capable of driving photosynthesis (Sharkey & Raschke,
1981a) and is often believed to operate through mesophyll-driven
consumption of CO2 reducing the internal CO2 concentration
(Ci) (Roelfsema et al., 2002, 2006), to which stomata are known
to respond (Mott, 1988). However, there is also evidence
for a direct guard cell red-light response, independent of
mesophyll photosynthesis, as discussed in Section V.
Stomatal opening is brought about by the accumulation of
ions and/or solutes (Imamura, 1943; Fujino, 1967; Outlaw &
Manchester, 1979; Outlaw, 1983) in guard cells, which increases
the osmotic potential, thus lowering the water potential,
causing water uptake from the apoplast (Weyers & Meidner,
1990; Willmer & Fricker, 1996). Increases in guard cell volume
and hence turgor pressure widen the stomatal pore (Franks &
Farqhuar, 2007; Shimazaki et al., 2007). Closure is brought
about by the reverse, a loss or release of solutes, accompanied
by a loss of water and consequently turgor pressure. Both pore
opening and closure are energy-dependent processes (Willmer
& Fricker, 1996).
However, our understanding of the perception of, and
precise response of stomata to, different environmental stimuli
is not complete. The fact that stomata in isolated epidermal
peels respond to various environmental factors suggests that
part 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 positive
correlation between stomatal conductance and mesophyll
photosynthesis (e.g. Wong et al., 1979; Zeiger & Field, 1982),
and a close correlation between photosynthetic efficiencies in
guard and mesophyll cells has also been observed (Lawson
et al., 2002, 2003). The majority of guard cells have chloroplasts,
and these would therefore provide an ideal and convenient
location for sensory or regulatory mechanisms. Although guard
cell chloroplasts are a characteristic feature of most plants, the
role of these highly conserved organelles in osmoregulation and
their 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 species
typically contain 10–15 chloroplasts per guard cell (Humble
& Raschke, 1971), compared with 30–70 in a palisade
mesophyll cell. However, numbers of chloroplasts per guard
cell range from 3–6 in Selaginella (Allaway & Milthorpe,
1976) to up to 100 in Polypodium vulgare (Stevens & Martin,
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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.
1978), and guard cells of Paphiopedilum species entirely lack
chloroplasts (Nelson & Mayo, 1975; Rutter & Willmer,
1979; D’Amelio & Zeiger, 1988) but still maintain
functional stomata (Nelson & Mayo, 1975). Guard cells
are formed from epidermal cells, which notably also lack
chloroplasts (again there are exception such as Polypodium
species; Fig. 2).
Guard cell chloroplasts are often smaller, with less granal
stacking, and some are less well developed than those in
mesophyll cells (Sack, 1987; Shimazaki & Okayama, 1990),
although these features vary across plant families (see reviews by
Pemadasa, 1981; Willmer & Fricker, 1996). Another noticeable
feature of most guard cell chloroplasts is that starch accumulates
in 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 Stadler
et al. (2003) has revealed that guard cells of Arabidopsis are
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practically free of starch in the morning and accumulate
starch during the day.
Before examining guard cell photosynthesis and its possible
role in stomatal behaviour, including the controversial topic
of guard cell Calvin cycle activity, it is essential to provide a
brief account of the osmoregulatory pathways that occur in
guard cells.
II. Osmoregulation in guard cells
Many decades of research have focused on the osmoregulatory
mechanisms found in guard cells. To put this review into
context, a brief history of stomatal osmoregulation is given in
this section; however, this is by no means exhaustive (I refer
readers to the following comprehensive reviews: Talbott &
Zeiger, 1998; Zeiger et al., 2002; Outlaw, 2003; Roelfsema &
Hedrich, 2005; Vavasseur & Raghavendra, 2005).
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In 1908, Lloyd (Lloyd, 1908) observed that stomata
contained more starch when closed in the dark than when
open 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 in
guard cell turgor, and became the most widely accepted theory
regarding the osmoregulatory mechanism for many decades
(Meidner & Mansfield, 1968). In the 1960s Fischer and coworkers highlighted the importance of potassium (K) uptake
in stomatal opening (Fischer, 1968; Fischer & Hsiao, 1968)
(although a paper on this by Imamura had already been
published; Imamura, 1943), and the starch–sugar hypothesis
was effectively replaced with the K+-malate2− theory. This
work demonstrated that K+ uptake (Yamashita, 1952; Fischer,
1968; see reviews by Raschke, 1975, 1979) in guard cells was
correlated 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 that
malate2− is the major counterion balancing K+ uptake, although
species such as Allium cepa (in which starch is absent) exclusively
use Cl− (Schnabl & Zeiger, 1977; Schnabl & Raschke, 1980).
Uptake of K+, driven by a H+ gradient activated by proton
ATPase (Zeiger, 1983; Shimazaki & Kondo, 1987), was shown
to be correlated with malate accumulation (Allaway, 1973)
and stomatal opening (see review by Outlaw, 1983). A role for
sucrose in guard cell osmoregulation was nearly forgotten
until several studies suggested that K+ and its counterions could
not provide all the osmoticum required to support stomatal
apertures in Commelina communis (MacRobbie & Lettau,
1980a,b), and led to the suggestion that soluble sugars
account for additional osmoticum to support opening
(MacRobbie, 1987; Talbott & Zeiger, 1993). Evidence exists
that sugars as well as K+-malate2− can act as osmotica for guard
cell osmoregulation (Outlaw & Manchester, 1979; Outlaw,
1983; Reddy & Rama Das, 1986; Talbott & Zeiger, 1993,
1998). For example, Commelina benghalensis accumulates
sugars (60% of required osmoticum) and malate2− when
treated with fusicoccin (Reddy et al., 1983), a fungal toxin
that activates the plasma membrane H+-ATPase (Johansson
et al., 1993).
It is easy to see how these changes in osmoregulatory theories
may have been problematic for researchers determining the
role of guard cell chloroplasts (see Fig. 3). For example, if
sucrose is not considered to be an important solute in water
movements, a role for guard cell photosynthetic carbon reduction
is virtually redundant.
1. Multi-osmoregulatory pathways in guard cells
To resolve the differences reported in the literature among results
obtained using different experimental procedures and different
species, Talbott & Zeiger (1996, 1998) outlined three distinct
pathways (see Fig. 3) involved in guard cell osmoregulation
© The Author (2008).
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that incorporate K+, Cl−, malate2− and sucrose, also involving
guard cell chloroplasts. They suggested that the importance of
these 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− from
carbon skeletons derived from starch (Outlaw & Lowry, 1977;
Outlaw & Manchester, 1979), and is believed to be involved
in the early morning opening response and under blue
light. In the second pathway, which is insensitive to DCMU
(Poffenroth et al., 1992), sucrose is supplied from the breakdown
of starch (Outlaw, 1982) and is also thought to play a role in
blue-light responses (Tallman & Zeiger, 1988; Poffenroth
et al., 1992; Talbott & Zeiger, 1993). In the third, DCMUsensitive pathway, sucrose is supplied as a product of guard cell
photosynthetic carbon reduction (Talbott & Zeiger, 1998).
In summary, K+ accumulation is used primarily for rapid opening
in the morning, whereas turgor maintenance in the afternoon
primarily uses sucrose (Talbott & Zeiger, 1993, 1996, 1998).
Talbott & Zeiger (1993) also demonstrated that the major
solutes change depending upon lighting regimes and the duration
of opening. In Vicia faba peels, the initial (30-min) stomatal
opening in response to blue-light illumination resulted in a
173% increase in the concentration of malate, which then
decreased, and the concentration of sucrose (from starch
breakdown) rose continuously, reaching 215% after 2 h. Under
red light, there was little increase in organic acid or maltose
concentrations, but the sucrose concentration increased to
208% (Talbott & Zeiger, 1993), with no evidence of starch
breakdown (Tallman & Zeiger, 1988; Poffenroth et al., 1992;
Talbott & Zeiger, 1993). As this was observed in epidermal
peels, sucrose must be supplied from guard cell photosynthetic
carbon reduction. Such observations support the hypothesis
of multi-osmoregulatory pathways that are regulated by
measurement (Talbott & Zeiger, 1998) and growth conditions
such as humidity (Talbott et al., 2003) and CO2 concentration
(Talbott et al., 1996, 1998; Frechilla et al., 2002, 2004; see
also Zeiger et al., 2002). The majority of this work was carried
out using V. faba and it is possible that different mechanisms
are 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 the
guard cells (Outlaw, 1989; Tarczynski et al., 1989; Reckmann
et al., 1990; Gautier et al., 1991) led Outlaw and co-workers
to 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 Hite
et al. (1993), who suggested that guard cells act as carbon
sinks, taking up sucrose via plasma membrane transporters
(e.g. Stadler et al., 2003), Outlaw and colleagues suggested
that apoplastic sucrose recently fixed in the mesophyll cells
was a source for guard cell symplastic sucrose and acted as an
osmoticum for stomatal opening or replacing guard cell
carbon stores (Lu et al., 1997; Ewert et al., 2000; Outlaw &
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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).
De Vleighere-He, 2001). However, this mechanism appears to
be dependent on the amount of sucrose in the apoplast, with
concentrations lower than 4 mM unable to support stomatal
opening (Ritte et al., 1999). The guard cell apoplastic sucrose
can also exert an osmotic effect, which can drive stomatal
closure, acting as a possible signal between mesophyll assimilation rate and transpiration (Kang et al., 2007a). It was also
postulated that sucrose concentrations near the guard cell
regulate gene expression, as has been shown in many other
tissues (e.g. Baiser et al., 2004). However, the majority of these
studies were conducted on V. faba, which is an apoplastic
phloem loader, and different mechanisms may regulate stomatal
movements in symplastic loaders (Kang et al., 2007b).
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The above research emphasizes the importance of environmental growth and experimental conditions, as well as
experimental incubation periods, and provides possible
arguments for the involvement of K+, malate2− and sucrose in
stomatal function. Additionally, it provides a feasible explanation as to why so many conflicting results are reported in the
literature (Zeiger et al., 2002).
III. Role of guard cell chloroplasts in stomatal
function
By the early 1990s a consensus was reached that in fact little
was known about guard cell metabolism (Mansfield et al.,
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1990) and, despite over a decade of research, in 2003 Ritte &
Raschke (2003) reiterated that there was a lack of information
about the physiology and role of the guard cell chloroplasts.
There are four primary ways in which guard cell chloroplasts
could contribute to stomatal function (see Outlaw, 1983;
Tominaga et al., 2001), with experimental evidence supporting
all of these functions:
• electron transport in guard cells produces ATP and/or
reductants 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 carbon
assimilated in the guard cell chloroplasts, or imported from the
mesophyll) is available to synthesize malate as a counter ion to
K+ (Willmer & Fricker, 1996) or is broken down into sucrose;
• photosynthetic carbon assimilation within guard cells produces
osmotically 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 of
the mesophyll, along with functional photosystem I (PSI)
and PSII (Zeiger et al., 1980; Outlaw et al., 1981; Shimazaki
et al., 1982; Hipkins et al., 1983). Several researchers have
provided evidence for linear electron transport, oxygen evolution
and 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 fluorescence
transients have suggested that induction profiles in guard cells
can 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 in
fluorescence transients are believed to reflect changes in Calvin
cycle activity throughout the day or with different wavelengths
of light (Mawson & Zeiger, 1991; Srivastava et al., 1998; Zeiger
et al., 2002). Additionally, the light-harvesting chlorophyll
protein is found in the phosphorylated state in the dark and
is dephosphorylated by red light, the opposite situation to
that in mesophyll (Kinoshita et al., 1993). This reversal has
been suggested to be related to high rates of cyclic electron
flow observed in guard cell protoplasts of V. faba, supported
by high PSI activity compared with the mesophyll (Lurie,
1977), which could enhance ATP production driven by increased
development of the thylakoid proton gradient. However,
Shimazaki & Zeiger (1985) did not observe any unusually
high PSI activity in guard cells of V. faba but showed linear
electron flow to be c. 80% that of the mesophyll. This is
consistent with the values of PSII operating efficiency reported
later by Lawson et al. (2002, 2003) using high-resolution
chlorophyll a (chla) fluorescence.
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In the absence of any CO2 fixation, such electron transport
rates could provide sufficient ATP to drive ion exchange during
stomatal opening (Shimazaki & Zeiger, 1985; Fig. 3), depending
on the light wavelength (Schwartz & Zeiger, 1984). Red light
induced stomatal opening was shown to be DCMU sensitive
and potassium cyanide (KCN) (respiratory poison) resistant
whilst under blue light the reverse was observed (Schwartz
& Zeiger, 1984). Patch clamp techniques established that a
chloroplast modulated red-light response stimulated a proton
pump at the plasma membrane in guard cells, suggesting that
guard cell photosynthesis may regulate stomatal aperture, through
the provision of energy (ATP) and photosynthetic signalling
products, such as NADPH (Serrano et al., 1988; see also Wu
& Assmann, 1993). However, later studies could not confirm
these findings (Roelfsema et al., 2001; Taylor & Assmann, 2001).
Experiments conducted under red light with and without the
inhibitors oligomycin (an inhibitor of oxidative phosphorylation) and DCMU (an inhibitor of PSII) demonstrated that
guard cells supplied ATP to the cytosol under red light, which
was utilized by the plasma membrane H+-ATPase for H+
pumping and stomatal opening (Tominaga et al., 2001). An
alternative theory for the utilization of photosynthetic electron
transport products suggested that ATP and redox power
provided from electron transport are used for the reduction
of oxaloacetate (OAA) and 3-phosphoglycerate (3-PGA) (from
guard cell CO2 fixation or imported from the cytosol; Fig. 3)
and exported to the cytosol via a 3-PGA-triose phosphate
shuttle (Shimazaki et al., 1989; Ritte & Raschke, 2003).
Sugar as well as K+ accumulation during red light-induced
stomatal opening has been reported (Talbott & Zeiger, 1998;
Olsen et al., 2002), with sugar production possible either from
starch breakdown (Talbott & Zeiger, 1998) or from the utilization of end products of electron transport in photosynthetic
carbon reduction within the guard cells themselves (Fig. 3;
Talbott & Zeiger, 1998; Olsen et al., 2002). Blue light-induced
stomatal opening (see next section) is generally believed not to
be dependent on products of guard cell electron transport, as
it 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 supplied
mostly from mitochondrial respiration (Shimazaki et al., 1982,
2007; Schwartz & Zeiger, 1984), although there is also evidence
for energy supply from guard cell chloroplasts (Mawson, 1993a).
Stomatal opening in response to weak blue light is greatly
enhanced 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, Shimazaki et al. (2007) proposed that guard cell chloroplasts
translocate 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 products
of guard cell photosynthetic electron transport in stomatal
responses, 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) or
ribulose-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 great
depth and I refer readers to this for full details. Briefly, blue
light induces rapid and highly sensitive stomatal opening
correlated with the phosphorylation of a plasma membrane
H+-ATPase pump and increased H+ pumping, which results
in the activation of voltage-gated K+ channels by membrane
hyperpolorization (see Shimazaki et al., 2007 for details), along
with the inhibition of s-type anion channels in Arabidopsis and
V. faba (see Marten et al., 2007). Inhibition of blue light-induced
opening with KCN suggests that ATP for proton pumping is
supplied 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 electron
transport in ATP supply, suggesting a possible metabolic
co-ordination between photophosphorylation and oxidative
phosphorylation in guard cells (Mawson, 1993b). H+ pumping
results in K+ uptake correlated with malate2− synthesis and/or
Cl− uptake. Malate2− is the result mostly of starch breakdown
(Outlaw & Manchester, 1979), as Arabidopsis mutants that do
not accumulate starch lack a proper blue-light response
(Lasceve et al., 1997). However, sucrose accumulation from starch
breakdown as an additional osmoticum in blue light-stimulated
opening 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 as
the blue-light receptor. Support for zeaxanthin as the specific
blue-light receptor came from experiments conducted on
epidermal peels of Arabidopsis mutants lacking zeaxanthin
(non photochemical quenching 1), which failed to respond to
blue light (Frechilla et al., 1999), although these results could
not be confirmed when experiments were conducted on
whole leaves (Eckert & Kaldenhoff, 2000; Kinoshita
et al., 2001). Furthermore, stomata in V. faba treated with
an inhibitor of carotenoid biosynthesis maintained a bluelight response, ruling out zeaxanthin as the only blue-light
receptor (Roelfsema et al., 2006).
Strong evidence for phototropins as blue-light receptors
was provided by Kinoshita et al. (2001), who demonstrated
(in both epidermal strips and intact plant material) that double
mutants for phot1 and phot2 proteins (serine/threonine protein
kinase) failed to respond to blue light. They established that
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phot1 and phot2 act redundantly as the blue-light receptors
in stomatal responses to blue light, as single mutants showed
a typical wild-type blue-light response (Kinoshita et al.,
2001), which led to phototropins becoming widely accepted
as the main blue-light receptor.
The magnitude of the blue-light response decreases from
morning to afternoon (Doi et al., 2004), consistent with early
morning stomatal opening, when light is enriched in the blue
wavelengths (Assmann & Shimazaki, 1999), and also consistent
with the theory of varying osmoregulatory pathways (Talbott
& Zeiger, 1998), and changes in guard cell fluorescence transients through the day (Srivastava et al., 1998). It should be
noted, however, that the stomatal response to blue light is not
universal, with several species lacking blue light-induced
stomatal opening. Stomata of the fern Adiantum capillus-veneris
do not open in response to blue light, despite having functional
phototropins and plasma membrane H+-ATPase (Doi et al.,
2006). Additionally, facultative CAM plants displayed bluelight-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 cell
photosynthetic electron transport is malic acid production via
PEPc, 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 relationship
between malate accumulation and starch loss. Light-stimulated
increases in PEPc activity have been demonstrated together with
increased NADP- or NAD-dependent malate dehydrogenase
activity, which catalyses the reduction of OAA (Rao &
Anderson, 1983; Scheibe et al., 1990), and malate accumulation
has been correlated with stomatal aperture (Allaway, 1973;
Pearson, 1973; Pearson & Milthorpe, 1974; Vavasseur &
Raghavendra, 2005). It is widely accepted that guard cells
contain high concentrations of starch and PEP carboxylase
(Willmer et al., 1973; Willmer & Rutter, 1977; Raschke,
1977, 1979; Outlaw & Kennedy, 1978) compared with
mesophyll cells (Cotelle et al., 1999) and many reports have
suggested that this is the major or only pathway for CO2
fixation in guard cells (e.g. Willmer et al., 1973; Reckmann
et al., 1990; see also Vavasseur & Raghavendra, 2005) into
malate and aspartate (Ogawa et al., 1978). The importance
of malate accumulation in light-induced stomatal opening
(Asai et al., 2000) has been demonstrated using 3,3-dichlorodihydroxyphophinoyl-methyl-2-propenoate (DCDP), an inhibitor
of PEPc (Parvathi & Raghavendra, 1997). PEPc activity and
malate formation have also been linked to CO2 response
movements in stomatal guard cells (Outlaw & Lowry, 1977;
Raschke, 1979; Schnabl et al., 1982; Hedrich & Marten,
© The Author (2008).
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1993; Hedrich et al., 1994; Cousins et al., 2007). Differences
in stomatal opening between adaxial and abaxial stomata have
been closely associated with differential starch hydrolysis, malate
synthesis and K+ uptake (Pemadasa, 1983) as well as light
wavelength (Wang et al., 2008), highlighting again the
flexibility of stomatal osmoregulation and behaviour depending
upon the environment. Further support for the importance
of PEPc activity in stomatal opening comes from recent
work conducted on PEPc-deficient mutants of the C4 dicot
Amaranthus edulis, which showed reduced rates of both
stomatal opening and final conductance compared with wildtype controls (Cousins et al., 2007). This recent work is in
agreement with earlier studies on potato (Solanum tuberosum)
plants which demonstrated greater rates of opening when
PEPc was over-expressed and reduced rates of opening in
plants 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 stomatal
opening. In dual light experiments, Tallman & Zeiger (1988)
found substantial starch degradation under blue-light illumination, but only a small amount of K+ uptake. From these observations they suggested that, if starch was converted to malate
(Schnabl, 1980), adequate uptake of K+ would be necessary to
counterbalance the anion. These results are consistent with the
observation that, in 10 μmol m−2 s−1 blue light, the rate of malate
synthesis in V. faba guard cells was only 25% of their maximum
(Ogawa et al., 1978). From such studies it was concluded that
starch breakdown under blue light can also result in accumulation 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 metabolism
has spanned several decades, but as yet there is no general
consensus. There are conflicting reports in the literature
concerning the capacity of photosynthetic carbon reduction in
guard 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 activity
in 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 and
Commelina in the light, 14CO2 was fixed into malate and
aspartate. This was validated later by Raschke & Dittrich
(1977), who showed that neither radioactive 3-PGA nor
Rubisco activity was present in epidermal peels of the same
tissues when they were exposed to 14CO2. Subsequent
experiments demonstrated that guard cell chloroplasts lacked
ribulose-1,5-bisphosphate carboxylase (RuBPC) and ribulose5-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
© The Author (2008).
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Review
cycle products in V. faba guard cell protoplasts in the light
(Schnabl, 1980). Screening 41 species using indirect immunofluorescence, Madavhan & Smith (1982) reported no evidence
of Rubisco in the guard cells of C4 plants and only negligible
detection in C3 plants, but appreciable amounts in onethird of CAM species surveyed. The size of the guard cell
phosphoglycerate pool was unaffected by light, suggesting
that PCRP is not involved in aperture regulation (Outlaw &
Tarczynski, 1984). Reckmann et al. (1990) determined
that only 2% of the solute required for stomatal opening was
provided by Rubisco activity in Pisum sativum, and concluded
that there was insignificant Rubisco activity, confirming the
conclusion of Hampp et al. (1982) that photoreduction of
CO2 by guard cells was absent.
In contrast to the above findings, the presence of Rubisco
in guard cells of V. faba has been unequivocally shown with
immunocytochemical localization (Zemel & Gepstein, 1985).
Numerous studies have also shown that guard cells contain
several 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 at
40–50% compared with mesophyll cells. Shimazaki et al.
(1989) validated these figures, showing RuBPC activity in
guard cells of the same species to be 40% of that in mesophyll
chloroplasts (on a chlorophyll basis). However, they suggested
that the low ratio of CO2 fixation to O2 evolution implied
that the major proportion of ATP and reducing equivalents
was used for reactions other than photosynthetic CO2 fixation.
These authors also pointed out that in many previous studies
values had been calculated on a cell rather than a chlorophyll
basis, and that recalculation would significantly increase previously obtained values in line with their observations.
The results of Gotow et al.’s study (1988) contradicted
earlier findings (Raschke & Dittrich, 1977) and showed that
feeding radio-labelled CO2 to guard cell protoplasts under
red light resulted in incorporation of radioactivity into 3phosphoglycerate, ribulose bisphosphate (RuBP), fructose and
sedoheptulose. Medium alkalinization indicating CO2 uptake
and oxygen evolution by guard cell protoplasts was shown
under white (Gotow et al., 1988) and red light (Shimazaki &
Zeiger, 1987). A photosynthetic dependence of sucrose
accumulation was illustrated using DCMU in epidermal peels
of V. faba under red light by Poffenroth et al. (1992). However,
reduced CO2 concentration triggered K+ uptake rather than
sucrose accumulation under red light (Olsen et al., 2002).
It is now widely accepted that the Calvin cycle enzymes are
present in guard cell chloroplasts; however, the debate over
their activity, function and role in stomatal behaviour remains
(Outlaw, 1996, 2003). Although guard cell photosynthetic
carbon 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), the
contribution to osmotic requirements for stomatal opening
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ranges from 2% (Reckmann et al., 1990) to 40% (Poffenroth
et al., 1992) (see Wu & Assmann, 1993). Many reports have
suggested that rates are too low for any functional significance
(Outlaw, 1989; Outlaw et al., 1982), whilst others have proposed the Calvin cycle to be a major sink for the products of
photosynthetic electron transport (Cardon & Berry, 1992;
Zeiger et al., 2002; Lawson et al., 2002, 2003). Evidence for
guard cell production of sucrose has been obtained during
red light-induced stomatal opening in V. faba, where no
starch breakdown was observed and sugar import was ruled
out 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 with
application of DCDP, an inhibitor of PEPc activity, suggesting that this pathway may become important when PEPc is
restricted.
5. Evidence for all four mechanisms in stomatal
function
In conclusion, there is evidence for all four of the above
mechanisms being involved in stomatal function. The bluelight stomatal response is believed to be mostly independent of
guard cell electron transport, as stomatal blue-light responses
have been observed in albino leaves (Karlsson et al., 1983;
Roelfsema et al., 2006), with energy for the activation of a plasma
membrane H+-ATPase supplied mostly by the mitochondria
(Fig. 3; Schwartz & Zeiger, 1984; Assmann & Zeiger, 1987;
Parvathi & Raghavendra, 1995), although there is also evidence
for chloroplastic supply (Mawson, 1993a) and red-light
enhancement (Shimazaki et al., 2007). There is support for
both zeaxanthin and phototropins as the blue-light receptors,
with phototropins the most widely accepted. Evidence exists
for the direct use of ATP and/or reductants (produced by
guard cell electron transport) in red light-induced stomatal
opening (Fig. 3; e.g. Shimazaki et al., 1989; Tominaga et al.,
2001; Olsen et al., 2002; Ritte & Raschke, 2003), as well as
in sugar production by photosynthetic carbon reduction
within the guard cells (Fig. 3; Talbott & Zeiger, 1996, 1998).
Starch stored in the guard cells can be broken down into either
malate2− (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 act
as osmotica for stomatal opening (Outlaw, 1982; Tallman &
Zeiger, 1988; Poffenroth et al., 1992). It appears that guard
cell chloroplasts can be involved in all four of the pathways
described above, and that the pathway used is conditional on the
species, time of day, and experimental protocols.
IV. Chlorophyll a fluorescence studies to
examine guard cell photosynthesis
Chlorophyll a fluorescence is a powerful technique to probe
and elucidate photosynthetic metabolism in guard cells. The
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early pioneering work of Zeiger and co-workers (Zeiger
et al., 1980; Melis & Zeiger, 1982; Mawson & Zeiger, 1991)
measuring Kautsky kinetics (Kautsky & Franck, 1943) in
individual guard cells showed distinct features associated
with Calvin cycle activity. The majority of early chlorophyll
fluorescence work was restricted to epidermal peels (Ogawa
et 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 steadystate chlorophyll fluorescence (F′) from guard cells in the
white areas of intact leaves of Tradescantia albiflora under
different CO2 and O2 concentrations and attributed changes
to photochemical and nonphotchemical quenching. From
these observations they concluded that both the carboxylation
and oxygenation of RuBP were major sinks for the end
products of photosynthetic electron transport. However, caution
should be applied when interpreting steady-state fluorescence
measurements as it is difficult to distinguish between
photochemical and nonphotochemical quenching components
(Baker, 2008). The report by Cardon & Berry (1992) was the
first research to provide physiological evidence for Rubiscomediated CO2 fixation and photorespiration and led the way
for many subsequent studies.
Advances in fluorescence methodology (see Goh et al., 1999),
with the development of the saturation pulse method of
fluorescence quenching analysis (Bradbury & Baker, 1981;
Schreiber et al., 1986) and pulse amplitude modulation (PAM)
fluorimetry (Schreiber et al., 1986) in conjunction with
technological developments in microfluorimetry (Goh et al.,
1999) and high-resolution imaging (Oxborough & Baker,
1997), made it possible to parametrize measurements of
chlorophyll fluorescence at the cellular (Oxborough & Baker,
1997; Goh et al., 1999) and subcellular levels (Baker et al.,
2001). With such advancements, measurements of PSII
operating efficiency ( Fq′ / Fm′ , where Fq′ is the difference
between maximum fluorescence in the light adapted state
( Fm′ ) and steady state fluorescence in the light (F ′ )) could be
obtained for individual cells and protoplasts (Goh et al., 1999).
The PSII operating efficiency estimates the efficiency at which
light absorbed by PSII is used for the reduction of the plastoquinone QA, and can provide an estimate of the quantum yield
of linear electron flux through PSII (Baker, 2008). Goh et al.
(1999) first used such techniques to compare fluorescence
quenching characteristics in guard and mesophyll cell protoplasts
(in V. faba and Arabidopsis). Light induction curves displayed very similar characteristics, indicating similar functional
organization of the thylakoid membranes, although guard cells
were saturated at lower light intensities and mesophyll cell
protoplasts had a higher capacity for photosynthetic electron
transport. In the same study, anaerobic conditions suppressed
photosynthetic electron flow in guard cells compared with
mesophyll cells. The O2-dependent electron flow suggested
a role for the Mehler-ascorbate peroxidase (MAP) cycle or a
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close metabolic coupling between photosynthetic electron
transport and export of reducing equivalents via a 3-phosphoglyceric
acid/dihydroxyacetone phosphate (PGA/DHAP) shuttle and
oxidative phosphorylation in the mitochondria (Goh et al.,
1999). However, again this work was restricted to measurements
of protoplasts or white areas of variegated plant tissue and was
not in agreement with later studies conducted on intact green
material (Lawson et al., 2002, 2003).
The first study that simultaneously examined the PSII
operating efficiencies ( Fq′ / Fm′ ) of guard and mesophyll cells in
intact green tissue revealed guard cell photosynthetic efficiency
to be 70–80% that of mesophyll chloroplasts (Lawson et al.,
2002). However, electron transport rates for the two cell types
could not be calculated because of uncertainties in the exact
light absorption and the contribution of PSI fluorescence in
guard and mesophyll chloroplasts. In the same study these
researchers measured Fq′ / Fm′ at different CO2/O2 concentrations in guard cells of intact green leaves of Tradescantia albiflora
and Commelina communis and confirmed that Rubisco was a
major 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 immunogold
labelling studies, which found weak labelling of PEPc but
significant Rubisco labelling in guard cells of Amaranthus
viridis (Ueno, 2001). The fact that the same CO2/O2 response
was observed in guard cells and in mesophyll cells suggests
that a major proportion of the end products of electron transport are being used by Rubisco and the Calvin cycle. Guard
cells contain 20–50-fold less chlorophyll than the underlying
mesophyll (Willmer & Fricker, 1996) and therefore, at similar
photosynthetic rates, extrapolation to the whole-cell level would
result in much lower guard cell photosynthesis compared with
the mesophyll. However, the small volume of guard cells
(one-tenth that of the mesophyll) means that the guard cell
CO2 assimilation rate could be one-third to one-tenth that of
the mesophyll, and therefore guard cell chloroplasts could
provide a significant energy source for these cells.
1. Alternative sources of energy
Although the aim of this review is to concentrate on guard cell
chloroplasts and their possible role in stomatal function,
guard cells also contain numerous mitochondria (Willmer &
Fricker, 1996; Vavasseur & Raghavendra, 2005), about one-third
the number of those in the mesophyll (Allaway & Setterfield,
1972), and several reports have suggested that these are the
most important organelle in guard cells. Hampp et al. (1982)
originally proposed that there was an absence of photoreduction
of CO2 in guard cells but a high metabolic flux through the
catabolic pathway. High respiration rates were observed by
Raghavendra & Vani (1989), suggesting that ATP produced
through oxidative phosphorylation was important for stomatal
movements (Parvathi & Raghavendra, 1997).
© The Author (2008).
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Fumarase activity (which is involved in the tricarboxylic
acid (TCA) cycle) has been shown to be high in guard cells of
V. faba and P. sativum (Hampp et al., 1982; see also Outlaw,
2003), and trangenic tomato (Solanum lycopersicum) plants with
considerable reductions in mitochondrial fumarate hydratase
(fumarase) activity showed substantial reductions in stomatal
aperture, resulting in CO2 limitation of photosynthesis
(Nunes-Nesi et al., 2007). Application of inhibitors of
photophosphorylation (DCMU) and oxidative phosphorylation (KCN) has shown that both mechanisms are used for
light-induced opening, but depend on wavelength (Schwartz
& Zeiger, 1984). Their relative importance alters when either
of 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 mesophyll
photosynthetic CO2 fixation (Wong et al., 1979; Farquhar &
Wong, 1984; Mansfield et al., 1990). Numerous studies have
demonstrated a strong correlation between photosynthesis
and stomatal conductance under a variety of different light
intensities and nutrient and CO2 concentrations (Radin et al.,
1988; Hetherington & Woodward, 2003). This relationship
causes, or is a consequence of, a constant Ci:Ca ratio (where Ca
is the external CO2 concentration), which has been observed
to 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, it
should also be noted that this relationship has easily been
broken in transgenic plants with various modifications to
photosynthetic metabolism (e.g. Hudson et al., 1992; Lauerer
et al., 1993; Stitt & Schulze, 1994; von Caemmerer et al.,
2004; Cousins et al., 2007; Baroli et al., 2008). The close
relationship between photosynthesis and stomatal behaviour
led to the hypothesis that guard cell responses may be linked
to mesophyll photosynthetic capacity via a mesophyll signal or
that guard cell photosynthesis itself may provide a metabolite
signal (Wong et al., 1979). Chloroplast ATP pool size was put
forward 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 zeaxanthin
concentration and stomatal apertures (in response to light and
CO2; see Zhu et al., 1998).
The debate over whether guard cell chloroplasts and/or
guard cell photosynthesis plays a direct role in the co-ordination
of stomatal movements in relation to mesophyll photosynthetic
CO2 demand remains unresolved. Recently, Roelfsema et al.
(2002, 2006) have argued against a direct role for guard cell
chloroplasts in red light-induced stomatal movements. These
researchers used albino areas of variegated plant tissue of
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V. faba treated with norflurazon (nf; inhibits carotenoid
biosynthesis), and showed stomatal opening in these two tissue
types in response to blue light but not red (Roelfsema et al.,
2006). They concluded that a lack of red-light response is
consistent with intercellular CO2 concentration as the intermediate signal in the stomatal red-light response. Moreover,
stomatal opening in response to red light was only apparent
when light was applied to a large area of the leaf, and not when it
was applied to individual guard cells, supporting a mesophylldriven Ci response (Roelfsema et al., 2002).
These obervations are in good agreement with earlier studies
by Karlsson (1986), who showed that lowering the atmospheric
CO2 concentration had a similar effect to red light and
enhanced the blue-light response. It should, however, be
mentioned that stomata in the albino portion of the leaf cannot
be considered to be completely indicative of responses in
green tissue (Scarth & Shaw, 1951; Lawson et al., 2002) as
their movements are much slower than those in green areas
(Scarth, 1932). However, Scarth (1932) also noted under red
light that stomata located near the green tissue tend to open
further than those at greater distances, adding support to the
theory of an indirect effect of red light on stomatal movements
through the action of mesophyll photosynthesis. Further
evidence for a CO2-mediated red light-induced stomatal
opening response has been provided by the Arabidopsis high
temperature 1 (HT1) mutant which carries a mutation in the
gene encoding a protein kinase (Hashimoto et al., 2006). These
mutants lack both a guard cell CO2 response and a red-light
response, but respond to blue light, supporting the notion that
red light-driven stomatal opening is promoted by reduced Ci,
although to corroborate this it would be necessary to present
stomatal conductance against Ci rather than CO2. Additional
support for Ci-driven stomatal opening has been provided in
Nicotiana 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 and
showed 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 stomatal
responses to light and CO2 occur in response to an unknown
mesophyll-generated signal. Epidermal peels of Tradescantia
pallida, V. faba and P. sativum showed no stomatal response to
light or CO2, but when T. pallida and P. sativum peels were
grafted back onto mesophyll (either their own corresponding
mesophyll or that of a different species), stomatal responses
were restored, although this was not the case for V. faba. The
authors argued against a direct effect of mesophyll-driven
changes in Ci, as increasing ambient CO2 from 120 to
540 μmol mol−1 did not induce stomatal closure, whereas
darkness resulted in complete closure, but Ci was only
200 μmol mol−1. In agreement with this, a further recent
study reported that abaxial stomata of Helianthus annuus were
more sensitive to light transmitted through the leaf (selftransmitted light) than to direct illumination, highlighting a
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photosynthesis-dependent involvement in stomatal responses
to light. This was attributed to an unknown photosynthetic
metabolite and not a Ci-driven effect, as Ci was maintained at
a constant value (Wang et al., 2008).
Several studies have also argued against a direct effect of Ci
on stomatal-driven responses to red light; for example,
stomata were shown to respond to light even when Ci was held
constant (Messinger et al., 2006; Lawson et al., 2008; Wang
et al., 2008). Additionally, stomatal responses to Ci and Ci
responses to light are believed to be too small to account for
the large changes in stomatal conductance that are often
observed in response to light (Sharkey & Raschke, 1981b).
Furthermore, red-light responses have been documented in
several 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 red
light-induced stomatal opening has been reported, although
this may be species dependent. Recently, Doi & Shimazaki
(2008) examined stomatal responses in the fern Adiantum
capillus-veneris to CO2 in darkness and found the stomata to
be unresponsive to low or high CO2 concentrations but to
open in response to red light. The fact that they observed a
synergistic effect of red and far-red light on stomatal opening,
and greater sensitivity when light was applied directly to the
lower surface along with a lack of response to Ci, led these
authors to conclude that opening in this species is driven by
photosynthetic electron transport in guard cell chloroplasts.
This is probably driven by K+ uptake, as CsCl (a K+ channel
block) inhibited the response (Doi & Shimazaki, 2008). In
the same experiment, Arabidopsis was used as a control and
showed ‘typical’ Ci responses in the dark, highlighting the
possibility that different species may use alternative signalling
pathways and mechanisms. In contrast to these findings, the
stomata of an Arabidopsis mutant that lacks a functional
SLOW ANION CHANNEL-ASSOCIATED 1 (SLAC1) gene,
which encodes a plasma membrane anion channel, were found
to fail to close at a high CO2 concentration in the dark and in
the light in one study (Negi et al., 2008), and to open in the
light and close more slowly in the dark in another (Vahisalu
et al., 2008). These studies support the involvement of ion
transport mechanisms in light-dependent stomatal movements, that are not dependent solely on Ci-driven responses.
To address stomatal behaviour in relation to mesophyll
photosynthesis, Messinger et al. (2006) suggested that the
balance between photosynthetic carbon reduction by Rubisco
and electron transport capacity was the key mechanism linking
stomatal response to light and CO2 concentration. This work was
based on alterations in the amount of ATP and/or zeaxanthin
resulting from a change in the balance of guard cell electron
transport (and energy states of the thylakoid membrane) in
relation to photosynthetic carbon reduction, determined by
light and Ci. Variations in the concentration of zeaxanthin in
turn would alter guard cell aperture in response to blue light
© The Author (2008).
Journal compilation © New Phytologist (2008)
Tansley review
and CO2 (Zeiger & Zhu, 1998; Zhu et al., 1998; Zeiger et al.,
2002). As with zeaxanthin, ATP should increase with light and
decrease with Ci with increased Calvin cycle activity. Increased
ATP in guard cells could be used in the cytosol for proton
pumping at the plasmalemma (Tominaga et al., 2001), or
alternatively the energy may be utilized to produce sucrose as
an osmoticum for stomatal opening through photosynthetic
carbon reduction (Talbott & Zeiger, 1993). This work also
suggested that there are at least two mechanisms by which
stomata respond to CO2, one dependent on photosynthesis,
and the other photosynthetically independent (Messinger et al.,
2006). Multiple CO2 response mechanisms have previously
been suggested by Assmann (1999). However, studies conducted
on transgenic plants have demonstrated similar conductances
in wild-type and trangenic plants (see next section), despite the
latter having an alteration in the balance of electron transport
rates 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 when
attempting to determine the role of mesophyll or guard cell
photosynthesis in stomatal function and in the past has relied
on epidermal peels or guard cell protoplasts. Such systems are
often criticized because of possible mesophyll contamination
(Weyers & Travis, 1981; Outlaw, 1983). However, the removal
of the mesophyll could prevent any mesophyll signalling and
may induce other mechanistic responses (Lee & Bowling,
1995; Lawson et al., 2002). In the late 1970s, Outlaw and
co-workers introduced the technique of dissecting individual
cells (Outlaw, 1980; Hampp & Outlaw, 1987; Outlaw & Zhang,
2001), highlighting its advantage in controlling contamination.
Improved molecular and transgenic techniques have provided
modern powerful tools (Webb & Baker, 2002) with which to
address many questions regarding photosynthesis in relation
to stomatal function and have already provided some invaluable
information (see above).
Early studies conducted on transgenic plants with impaired
photosynthesis revealed some surprising results. The effects of
Rubisco concentration on photosynthesis were studied independently by several groups, all of which studies suggested
little effect on stomatal behaviour (Quick et al., 1991; Stitt
et al., 1991; Hudson et al., 1992). Specifically, these studies
showed similar stomatal conductance values in transgenic
plants compared with wild-type controls, despite a severe
reduction in photosynthesis and higher Ci concentrations.
Studies on tobacco (Nicotiana tabacum) plants with reduced
phosphoribulokinase (Paul et al., 1995) or Rieske FeS protein
(Price et al., 1998) also showed no effect on stomatal conductance, and Price et al. (1998) concluded that ‘stomata are not
strongly reliant on photosynthetic electron transport for setting
conductance’. However, work on transgenic antisense PEPc
plants supported a role for malate and PEPc activity in guard
© The Author (2008).
Journal compilation © New Phytologist (2008)
Review
cells, with delays in stomatal opening responses in potato with
reduced PEPc activity, whilst over-expressors showed accelerated
opening (Gehlen et al., 1996). These findings are supported
by recent work on Amaranthus edulis mutants deficient in PEPc,
which show both reduced rates of opening and also reduced
final stomatal conductances (Cousins et al., 2007). Stomata in
plants with 12% wild-type fructose-1, 6-bisphatase (FBPase)
activity showed significantly faster opening responses and
higher final conductances with increasing irradiance, despite
lower photosynthetic rates and elevated Ci. However, this was
dependent upon humidity and external CO2 concentration
(Muschak et al., 1999).
However, the aims of the above studies were not specifically to
determine stomatal responses. Certain assumptions were made:
firstly, because Cauliflower mosaic virus (CAMv) promotors were
used in the majority of the studies, it was assumed that all cells
were antisensed in a similar manner, and, secondly, it was assumed
that the observed response was equivalent to steady-state stomatal
conditions. To directly address the influence of reduced
photosynthetic capacity using antisense technology, von
Caemmerer et al. (2004) used high-resolution chlorophyll
fluorescence imaging to show for the first time that photosynthetic efficiency was reduced to a similar extent in the guard
cells as in the mesophyll cells in tobacco plants with reduced
concentrations of Rubisco. Decreasing Rubisco activity resulted
in an imbalance between chloroplast electron transport and
the photosynthetic carbon reduction capacity, which in turn
could lead to an increased amount of ATP and/or conversion
of xanthophyll pigments to zeaxanthin. Increased nonphotochemical quenching was observed in the antisense plants,
which could be interpreted as an increase in the amount of
zeaxanthin. Both ATP and zeaxanthin have been implicated as
playing a key role in stomatal opening responses. However, a
step change in irradiance revealed similar stomatal responses
in terms of opening rates and final conductances in antisense
Rubisco plants and wild-type controls, despite significantly
lower photosynthetic rates in the former. This led to elevated
internal CO2 concentrations within these plants, which initially
was interpreted as a reduced sensitivity to Ci (von Caemmerer
et al., 2004). However, manipulation of Ci through changes
in Ca resulted in stomatal closure, suggesting that stomata
may response to Ca and not Ci (von Caemmerer et al., 2004).
The overall conclusion from this work was that neither mesophyll
nor guard cell photosynthesis was necessary for stomatal
opening responses.
The fact that the majority of studies used white light or a
mixture of red and blue light does not rule out the possibility
that blue light-stimulated opening, independent of photosynthesis, overrides any mesophyll/guard cell signal (Talbott &
Zeiger, 1993). To resolve this issue, a recent study of red-light
responses by Baroli et al. (2008) distinguished between
antisense Rubisco tobacco plants with 10–15% wild-type
Rubisco activity, which have major reductions in the carboxylation capacity of photosynthesis, and antisense tobacco
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plants with impaired rates of electron transport via reductions in
the cytochrome b6f complex. No changes in stomatal opening
were observed in either of the transgenic plants in response to
a step change in red light at ambient CO2 concentrations,
leading to the conclusion that this response was independent
of guard or mesophyll cell photosynthesis. The fact that no
phenotypic stomatal responses were observed despite a decrease
in sucrose concentration also strongly suggests that something
other than sucrose acts as osmoregulator during opening.
However, recent work conducted on antisense sedopheptulose1,7-bisphosphatase (SBPase) tobacco plants has shown a
minor regulatory role for photosynthetic electron transport in
response to red light (Lawson et al., 2008). A step change in
red illumination resulted in an increased rate of stomatal
opening which was not observed under a blue/red light mix.
ATP concentrations in the antisense SBPase plants may be
increased because of the reduced ATP consumption by
the Calvin cycle. These authors suggested the possibility of
increased stomatal opening under red light, as a result of an
increase in ATP available for proton pumping (Tominaga
et al., 2001). However, Baroli et al. (2008) reported little effect
of reduced ATP on red light-induced stomatal opening in
transgenic tobacco plants with reduced cytochrome b6f complex
(Baroli et al., 2008).
Other suggested functions of chloroplasts in guard cells in
regulation of stomatal behaviour include the production of
reactive oxygen species such as H2O2 (possibly via Mehler
activity at PSI) which may play a role in abscisic acid (ABA)
signal transduction (Zhang et al., 2001). A role for ascorbic
acid (Asc) redox state has also been postulated in guard cell
regulation. Plants with increased guard cell Asc redox state
(through increased expression of dehydroascorbate reductase
(DHAR)) exhibited a reduced concentration of guard cell
H2O2 and consequently higher stomatal conductances (Chen
& Gallie, 2004).
As highlighted above, the use of transgenic and mutant
plants has provided significant information regarding guard
cell 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 (Inoue
et al., 2008), as well as ion uptake and ion channel regulation
(Serna, 2008), all of which play key roles in stomatal sensitivity
and behaviour.
VI. Stomata in relation to water use/
manipulation of behaviour
One of the important outcomes of understanding how guard
cells function is the potential to engineer drought-tolerant
plants. This prospect has received increasing attention from
the wider scientific community, with several reports published
recently suggesting that stomatal metabolism may hold the
key (Nilson & Assmann, 2007). For example, maize (Zea
mays) plants with increased amounts of NADP-malic enzyme
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(ME), which converts malate and NADP to pyruvate,
NADPH and CO2, had altered stomatal behaviour. MEtransformed plants had decreased stomatal conductance,
showing signs of drought avoidance associated with guard cell
malate metabolism. A negative aspect of this drought-tolerance
engineering was that, following exposure to drought, the
development of necrosis was more rapid in leaves from plants
with the highest ME expression (Laporte et al., 2002). Masle
et al. (2005) reported the isolation of a ‘transpiration efficiency
gene’, ERECTA, which acts on cell expansion and cell division,
amongst other processes, resulting in modification of leaf
diffusive properties and mesophyll capacity for photosynthesis,
leading to greater water use efficiency, in Arabidopsis. Increases
in drought resistance have also been reported in Arabidopsis
mutants, with alterations or disruptions of guard cell membrane
transporters (Klein et al., 2004), calcium-dependent protein
kinases (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 gene
SNAC1 (STRESS-RESPONSIVE NAC1) enhanced drought
tolerance in rice (Oryza sativa) (Hu et al., 2006).
In the attempt to produce plants with increased water use
efficiency or drought tolerance, genetic engineering or mutations
provide an opportunity to alter not only stomatal physiology
and function (Nilson & Assmann, 2007) but also anatomical
features, such as stomatal density and size (originally proposed
in the 1970s; Jones, 1976, 1977) and amounts of leaf cuticular
wax (Aharoni et al., 2004). A recent review by Wang et al.
(2007) highlights the importance of, and recent progress
made in, identifying genes controlling stomatal density or
patterning, and how such genetic manipulations may increase
plant water use efficiency.
Altering the stomatal density does not automatically alter
stomatal conductance (Fig. 4; Lawson, 1997; Weyers & Lawson,
1997; Lawson & Morison, 2004). Figure 4 shows a model of
predicted stomatal conductance with changes in various
stomatal characters. From this model it is obvious that stomatal
aperture has the greatest control over stomatal conductance,
with stomatal density being secondary. An example of this can be
found in experiments conducted on Arabidopsis over-expressing
the STOMATAL DENSITY AND DISTRIBUTION 1 (SDD1)
gene, resulting in plants with a 40% reduction in stomatal
density, and the Arabidopsis sdd1-1 mutant (Berger & Altmann,
2000), in which stomatal density is increased to 300% of that
of wild type. Under growth conditions, no differences in
stomatal conductance or assimilation rate were observed in
the over-expressers and the sdd1-1 mutants compared with
wild type. Lower stomatal density was compensated for by an
increase in aperture and, conversely, reduced stomatal aperture
compensated for increased stomatal density (Bussis et al.,
2006). It should be mentioned that, although mutants may
be identified as ‘drought resistant’ or with ‘increased water use
© The Author (2008).
Journal compilation © New Phytologist (2008)
Tansley review
Review
were blocked with grease to prevent stomatal conductance and
a vein was severed to prevent uptake of DCMU, were used to
show the relationship between PSII operating efficiency and
stomatal conductance. Images of Fq′ / Fm′ showing spatial and
temporal resolution of PSII operating efficiency were compared with thermal images of leaf temperature, which is modulated by stomatal behaviour and other environmental factors
(i.e. in general, the greater the stomatal conductance the greater
the evaporative cooling of the leaf and the lower the leaf temperature). From the images it is apparent where stomatal behaviour
is influencing PSII operating efficiency and vice versa.
VII. Concluding remarks and future direction
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.
efficiency’, such traits may not be evident when they are
grown in competitive environments. Basco et al. (2008) recently
reported that Arabidopsis ABA oversensitive mutants, which
display enhanced stomatal closure, could not compete with
wild type for water when the plants were grown together. Such
findings also have significant implications for screening
protocols when attempting to identify mutants (Basco et al.,
2008). It is also important to note that screening plants for
increased water use efficiency should include measurements of
photosynthetic performance in relation to stomatal behaviour,
as reduced stomatal conductance can decrease water use but
also limit photosynthetic carbon assimilation.
In conjunction with advances in molecular biology,
substantial progress has been made in technology and
methodology. 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 instantaneous water use efficiency, and is not only a potential screening
tool allowing determination of both photosynthetic performance
and stomatal behaviour but also a powerful approach to
elucidating correlations between stomatal behaviour and
photosynthetic capacity. An example of combined chlorophyll
fluorescence imaging and thermography is shown in Fig. 5.
Extreme treatments, in which stomata in one area of the leaf
© The Author (2008).
Journal compilation © New Phytologist (2008)
Stomatal research over the past few decades has revealed a
complicated network of osmoregulatory and signalling pathways
in guard cells (e.g. Li et al., 2006). It appears that these highly
plastic cells have the capability to alter mechanisms of response
depending upon environmental growth and experimental
conditions, complicated further by time of day and pretreatments (Zeiger et al., 2002), all of which appear to be species
dependent. Such flexibility gives stomata the necessary
capability to maintain a regulatory role in plant water status
and photosynthetic capacity. This review has concentrated on
guard cell chloroplast photosynthesis and in particular Calvin
cycle function, a highly controversial topic (Outlaw, 1989,
2003), with evidence for and against functional guard cell
photosynthetic regulation of stomatal behaviour. Recent research
conducted on antisense SBPase plants suggests guard cell
photosynthesis and/or carbon reduction may play a role in
stomatal responses to red light (Lawson et al., 2008). However,
at the same time, work on antisense Rubisco and b6f plants
casts doubt on any role for guard cell photosynthesis, including
the production of ATP, in red light-induced opening (e.g von
Caemmerer et al., 2004; Baroli et al., 2008). Discrepancies
in results and conclusions regarding the role of guard cell
chloroplasts in stomatal function are probably attributable to the
unique plasticity of guard cells, which can make interpretations
difficult, with often opposing conclusions in different laboratories
in which research was conducted under different conditions
(see Zeiger et al., 2002). Stomatal research in the future should
therefore take into account the time of day experiments are
conducted, the conditions under which the plants are grown
and the type of material used, as all of these factors can impact
on stomatal responses, signalling pathways, and solutes required
for osmoregulation of stomatal aperture.
The current transition towards using mutants and transgenic
plants along with the identification of gene trap lines (Galbiati
et al., 2008) opens a new window of opportunity to pursue
different avenues of research to answer some of the many
questions that still remain regarding guard cell metabolism.
To date, attention has focused on photosynthetic pathways in
guard and mesophyll cells, and to a certain extent the oxidative
phosphorylation pathway has been neglected. Transgenic plants
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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).
and mutants provide an ideal opportunity to determine the
role of this pathway in stomatal sensory and response mechanisms. The development and discovery of guard cell specific
promoters (see Yang et al., 2008) will allow manipulation of
guard cell metabolism without disruption of mesophyll
photosynthetic metabolism. Such systems will hopefully
provide a probe that will help to fully elucidate the link
between mesophyll photosynthesis and stomatal conductance.
Microarray and proteomic technology allows gene expression
patterns involved in signal transduction pathways to be identified
and assessed under different environmental conditions and
stresses (Coupe et al., 2006). Leonhardt et al. (2004) have
demonstrated the power of microarray technology comparing
the expression profiles of guard and mesophyll cells. They noted
that, when leaves were sprayed with ABA, there was repression
of many of the enzymes involved in guard cell metabolism,
including a decrease in PEPc transcript, which agrees with
earlier work reporting decreased PEPc activity under drought
(Kopka et al., 1997). Transcriptomic analysis can also identify
transcription factors that are necessary for stomatal movement
mediating stomatal responses to light and darkness (Gray,
2005; see review by Casson & Gray, 2008).
To date, most stomatal research has concentrated on plant
species very familiar to stomatal biologists, but there are still
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numerous gaps in our knowledge regarding stomatal behaviour
in CAM and grass species. C3/CAM intermediates may provide
an ideal opportunity to uncover light and CO2 responses as
well as induction of specific genes or signalling pathways.
Stomata respond to numerous environmental stimuli, yet
most studies are conducted in isolation. There is a desperate
need to determine the hierarchy of stomatal responses, and
the influence of combined factors on stomatal behaviour,
response and signalling mechanisms.
Although significant advances in the understanding of
guard cell function and stomatal responses have been made
over the last century, many gaps in our knowledge remain
regarding guard cell metabolism and its role in stomatal
behaviour. The use of antisense techniques in conjunction with
guard cell-specific promoters, and modifications of guard cell
chloroplast metabolism, coupled with in situ measurements of
photosynthetic performance, stomatal function and responses
to various stimuli, may provide the key to ascertaining the
roles 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
© The Author (2008).
Journal compilation © New Phytologist (2008)
Tansley review
research. In particular Dr James I. L. Morison and Professor
Neil R. Baker are acknowledged for the opportunity to work
in their laboratory imaging chlorophyll fluorescence in guard
cells. My initial interest in the stomatal regulation of gas
exchange was inspired during work for a PhD with Dr
Jonathan Weyers (Dundee University), at which time I was
encouraged to carry out my first research on this subject.
Professors Christine Raines and Susanne von Caemmerer
have provided many useful discussions. Dr Tanja Hofmann is
gratefully acknowledged for critical reading of the manuscript.
I would also like to thank three anonymous reviewers for their
comments, which have greatly enhanced the manuscript.
Work on chlorophyll fluorescence imaging in guard cells was
funded by a BBSRC grant awarded to Dr James I. L. Morison
and Professor Neil R. Baker, whilst recent work on transgenic
plants was funded by the University of Essex.
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