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RESEARCH PAPER |
Altered stomatal dynamics in ascorbate oxidase over-expressing tobacco plants suggest a role for dehydroascorbate signalling
1Group of Biotechnology of Pharmaceutical Plants, Laboratory of Pharmacognocy, Department of Pharmaceutical Sciences, Aristotle University of Thessaloniki, 541 24 Thessaloniki, Greece
2Dipartimento di Biologia e Patologia Vegetale, Universita‘ di Bari, Via E. Orabona 4, I-70125 Bari, Italia
3Institute for Research on the Environment and Sustainability, Devonshire Bldg., Newcastle University, Newcastle upon Tyne NE1 7RU, UK
* To whom correspondence should be addressed. E-mail: kanellis{at}pharm.auth.gr
Received 26 September 2007; Revised 11 December 2007 Accepted 12 December 2007
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Abstract |
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Control of stomatal aperture is of paramount importance for plant adaptation to the surrounding environment. Here, we report on several parameters related to stomatal dynamics and performance in transgenic tobacco plants (Nicotiana tabacum L., cv. Xanthi) over-expressing cucumber ascorbate oxidase (AO), a cell wall-localized enzyme of uncertain biological function that oxidizes ascorbic acid (AA) to monodehydroascorbic acid which dismutates yielding AA and dehydroascorbic acid (DHA). In comparison to WT plants, leaves of AO over-expressing plants exhibited reduced stomatal conductance (due to partial stomatal closure), higher water content, and reduced rates of water loss on detachment. Transgenic plants also exhibited elevated levels of hydrogen peroxide and a decline in hydrogen peroxide-scavenging enzyme activity. Leaf ABA content was also higher in AO over-expressing plants. Treatment of epidermal strips with either 1 mM DHA or 100 µM hydrogen peroxide resulted in rapid stomatal closure in WT plants, but not in AO-over-expressing plants. This suggests that signal perception and/or transduction associated with stomatal closure is altered by AO over-expression. These data support a specific role for cell wall-localized AA in the perception of environmental cues, and suggest that DHA acts as a regulator of stomatal dynamics.
Key words: ABA, apoplast, ascorbic acid, ascorbate oxidase, dehydroascorbic acid (DHA), hydrogen peroxide, Nicotiana tabacum L., cv. Xanthi, stomata, transgenic plants, water stress
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Introduction |
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Like other living organisms, plants face a continuously changing environment. They must therefore be capable of perceiving and reacting (often rapidly, given their sessile nature) to changes in environmental conditions. This is achieved via a combination of sensors and receptors that have evolved to monitor the physical and chemical signals triggered by external cues, ultimately delivering a suite of responses at the molecular level aimed at maintaining cellular homeostasis and/or changing the developmental programme (Pastori and Foyer, 2002). Stomata must be particularly responsive to external cues, as the control of stomatal aperture is of central importance for plant survival. Consequently, mechanisms underlying stomatal movement and regulation have been much-studied (Hetherington and Woodward, 2003). The identification of the role of abscisic acid (ABA) in governing stomatal closure (Hiron and Wright, 1973) opened new perspectives to research, but the detailed sequence of events acting upstream and downstream of ABA synthesis remain a matter of conjecture. In the past few years, a role for ABA-dependent hydrogen peroxide and nitric oxide production in mediating stomatal closure has been identified (Zhang et al., 2001; Bright et al., 2006). Moreover, tobacco plants with altered expression of dehydroascorbate (DHA) reductase, the enzyme catalysing conversion of DHA to the reduced form ascorbate (AA), have been reported to exhibit abnormal stomatal behaviour; transgenic plants with increased DHA reductase expression showing increased stomatal opening and reduced capacities to control water loss, whereas antisense plants with low DHA reductase expression exhibited partially closed stomata and improved control over transpiration (Chen and Gallie, 2004). Several lines of evidence suggest a crucial role for apoplast redox status in controlling plant response to environmental conditions (Pignocchi and Foyer, 2003; Noctor and Foyer, 2005). Due to the lack of NAD(P)H and glutathione, the redox buffering capacity of the apoplast is much weaker than inside the cell (Pignocchi et al., 2003). It has been hypothesized that the apoplast enzyme ascorbate oxidase (AO) could be a key regulator of extracellular redox status (Noctor and Foyer, 2005). Indeed, tobacco plants over-expressing AO are characterized by the complete oxidation of the cell wall-localized AA pool (Sanmartin et al., 2003) plus changes in the expression and activities of several AA-related enzymes (Fotopoulos et al., 2006). Therefore, AO over-expressing plants were used to probe the role of apoplastic AA in the regulation of stomatal dynamics. These data reveal altered stomatal behaviour in AO over-expressing plants, and suggest a putative role for apoplast DHA accumulation in the regulation of stomatal aperture.
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Materials and methods |
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Plant material and growth conditions
Seeds of wild-type tobacco (Nicotiana tabacum cv. Xanthi) and three transgenic lines over-expressing AO (R50, R60, and R68, selected on the basis of AO expression and single transgene copy number; Sanmartin et al., 2003) were sown in seed trays containing John Innes No. 2 potting compost. After sowing, the seeds were stratified for 2 d at 4 °C and then grown at 23/18 °C day/night in a controlled environment chamber supplying a photosynthetic photon flux density of 150 µmol m–2 s–1 at plant height as a 16 h photoperiod. Relative humidity was maintained at 65–75%. After 10 d, seedlings were transferred to individual pots containing 5 dm3 of the same compost. Plants destined for stomatal conductance measurements were transferred to controlled environment chambers described in detail elsewhere (Sanmartin et al., 2003), in which the temperature ranged from a midday maximum of 28 °C to a night-time minimum of 17 °C, and plants were subject to a photosynthetic photon flux density (PPFD) of 350 µmol m–2 s–1 (at the height of the leaves employed for measurements), in a 16 h photoperiod. RH was maintained at 65–70%. Plants were watered as required and supplied with fertilizer at 14 d intervals (Phostrogen 14N:10P:27K; Monsanto, High Wycombe, Bucks., UK). Fully-expanded leaves from 8–10-week-old plants were used for all analyses. Assays were repeated at least three times using pooled samples (each ‘replicate’ measurement comprising leaves from a minimum of three independent plants, unless otherwise stated). To avoid rhythmic effects on stomatal aperture, experiments were always started at the same time of day (12.00 noon).
Histochemical detection of reactive oxygen species
The detection of H2O2 was performed according to Thordal-Christensen et al. (1997) with the following modifications. Detached leaves were incubated in 1 mg ml–1 3,3'-diaminobenzidine (DAB)-HCl, pH 3.8 (Fluka, Buch, Switzerland) in the dark at room temperature for 8–10 h, then chlorophyll was removed by boiling in ethanol (96%, v/v) for 10 min. The assay was based on the instant polymerization of DAB (to form a reddish-brown complex which is stable in most solvents), as soon as it comes into contact with H2O2 in the presence of peroxidases. To determine the specificity of DAB staining, leaves were also stained in the presence of 200 U ml–1 catalase (bovine liver, Sigma-Aldrich, St Louis, MO, USA), resulting in clear, unstained samples. Deposits of H2O2 were quantified by pixel counts on scanned images using the ImageJ 1.33 program (National Institutes of Health, USA); as described by Romero-Puertas et al. (2004). Results were expressed on a projected area basis [(spot area/total projected leaf area)x100]. Four leaves from independent plants were used per genotype. Detection of H2O2 in guard cells was examined by loading freehand epidermal peels with DAB as described above, with minor modifications. To reduce variation, epidermal strips from control and AO over-expressing leaves were placed on a microscope slide loaded with 500 µl of 2 mg ml–1 DAB-HCl (pH 3.8) for 1 h. After rinsing with dH2O, H2O2 deposits were quantified by measuring the pixel intensity within a defined area (encompassing a guard cell). A minimum of 20 guard cell pairs per genotype were measured. Representative background pixel intensity was obtained from the average of five measurements of similarly sized areas adjacent to the guard cells. The average background value was then subtracted from the average guard cell value, and the difference was presented as the mean pixel intensity per guard cell.
Enzyme assays
Tobacco leaves were ground in buffer (50 mM TRIS–HCl, pH 7.4, 100 mM NaCl, 2 mM EDTA, and 1 mM MgCl2), then subject to 30 min centrifugation at 15 000 g. CAT (10–6 mol H2O2 reduced min–1 mg–1 protein), and APX (10–8 mol AA oxidized min–1 mg–1 protein) activity was determined as described by Aebi (1984) and de Pinto et al. (2000), respectively. For the APX assay, 2 mM AA was present in the extract buffer. Protein was estimated by the method of Bradford (1976), using BSA as a standard.
Leaf water content and weight loss following detachment
Leaves were collected and immediately weighed (FW), then rehydrated for 24 h at 4 °C in darkness to determine the turgid weight (TW). Oven-dry weight (DW) was recorded after drying leaf tissue to constant weight at 85 °C. Leaf relative water content (RWC) was determined as:
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Fully-expanded leaves were detached from well-watered plants and immediately weighed (FW). Water loss was then recorded following the incubation of detached leaves under stable environmental conditions at room temperature for 60 min. Measurements were made on four comparable leaves from separate plants. Leaf water loss was plotted as the percentage loss in weight with respect to FW.
Chlorophyll leaching assay
Chlorophyll leaching assays were performed as described by Aharoni et al. (2004). In brief, fully-expanded leaves were detached and rinsed with tap water, weighed, and placed in tubes containing 30 ml of 80% ethanol at room temperature (gently agitating in the dark). One microlitre was removed from each sample after 60 min and 120 min and the absorbance of samples recorded at 664 nm and 647 nm. Chlorophyll concentration was determined using the formulae described by Aharoni et al. (2004).
Stomatal measurements
Abaxial stomatal conductance was recorded on comparable fully-expanded leaves of 12 WT and 12 R60 transformants at regular intervals over the course of a diurnal cycle. All measurements were made with the same porometer, calibrated at frequent intervals during the day (Model AP4, Delta-T Devices, Ltd, Cambridge, UK). The experiment was repeated in its entirety with similar outcomes. Only one of the experimental data sets is presented, since baseline conductance varied at the measurement dates selected, due to differences in physiological age, growing conditions, and plant maturity between experiments.
Stomatal aperture was determined by introducing 200 µl of 40 µg ml–1 3,3'-dihexyloxacarbocyanin iodide (DiO6) to leaf discs mounted (abaxial side up) on glass slides for 5 min in the dark. The stained leaves were rinsed with dH2O, blotted to remove surface liquid, and imaged using a Zeiss epifluorescent microscope fitted with a 450–490 nm excitation filter and a 526 nm barrier filter. The percentage of open stomata (defined as having a width >1 µm) was determined from records of at least 190 stomata observed on eight independent leaves per genotype. The width and length of 190 pores (i.e. stomatal aperture) were measured and this was used to calculate average stomatal aperture (width/length) (Chen and Gallie, 2004). The width and length was also used to calculate stomatal aperture area
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Induction of stomatal closure by DHA and H2O2 was investigated using epidermal strips according to Pei et al. (2000). Strips, taken from leaves of similar size and age, were incubated in Petri dishes containing 30 mM KCl, 10 mM MES–KOH pH 6.0. After 2 h equilibration at room temperature and low PPFD (25 µmol m–2 s–1), strips were transferred to fresh Petri dishes containing buffer (controls), 1 mM DHA in buffer, or 100 µM H2O2 in buffer. 1 mM DHA was chosen after preliminary experiments, showing that it was the most effective concentration without visible signs of damage (5 mM or higher induced some brown spots). After incubating for 2 h, strips were placed onto microscope slides and observed under a light microscope. Opening (width) of at least 200 stomata per treatment were measured using a micrometric grid. To avoid any potential rhythmic effects, all sampling was conducted at midday.
Determination of leaf ABA content
ABA content was determined as described by Lovisolo et al. (2002) with minor modifications. In brief, 1 g of freeze-dried homogenized leaf tissue was extracted in 80% v/v methanol. Extracts were centrifuged twice at 4000 g for 20 min and the supernatant recovered. Methanol was removed under vaccuum and the aqueous residue partitioned three times against ethyl acetate at pH 3.0. Ethyl acetate of the combined organic fractions was removed under vaccuum. The residue was resuspended in TRIS-buffered saline (150 mM NaCl, 1 mM MgCl2 and 50 mM TRIS, pH 7.8) and ABA detected using an immunological approach (Phytodetek kit supplied by Agdia, Elkhart, USA).
RNA blot analysis
Total RNA was isolated from tobacco leaves as described by Verwoerd et al. (1989). Total RNA (10 µg) was fractionated on formaldehyde denaturing agarose gels, transferred to nylon membranes (Nytran® 0.45, Schleicher and Schuell, GmbH, Dassel, Germany), stained with 0.04% methylene blue (to observe equality of loading) and hybridized with a 32P-labelled probe for Cistus creticus NCED cDNA. Probe labelling was carried out with RadPrime DNA Labelling System (Invitrogen, Life Technologies, Madison, WI, USA) according to the manufacturer's instructions. Blot hybridization and membrane washing were performed as described by Church and Gilbert (1984). Experiments were performed twice, with similar results. A partial-length cDNA fragment showing high sequence homology to known NCED genes that was originally isolated from a C. creticus trichome cDNA library (V Falara and AK Kanellis, unpublished data) was used for the construction of the probe.
Statistical analyses
Statistical analyses were performed using SPSS v.11 (SPSS Inc., Chicago, USA). Biochemical and physiological measurements were subject to ANOVA, with RM-ANOVA employed where appropriate, then significant differences between individual means determined using Tukey's pairwise comparison test at the 5% confidence level.
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Results |
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The effect of AO over-expression on stomatal conductance
Measurements made in situ on one of the AO over-expressing lines (R60) revealed a consistent and significant (P <0.001) reduction in conductance over the photoperiod in R60 compared with WT plants (Fig. 1). The adoption of a novel guard cell staining method that is rapid and requires no fixation steps, employing 3,3'-dihexyloxacarbocyanin iodide (DiO6), a lipophilic cationic fluorescent dye previously used to stain mitochondria and the endoplasmic reticulum in living plant and animal cells, as well as fungal structures (Duckett and Read, 1991), revealed effects on stomatal aperture consistent with conductance data (Fig. 2A, B). Measurements revealed average stomatal aperture, average stomatal area µm–2 leaf and total open pore area µm–2 leaf to be significantly (P <0.05) lower in fully-expanded leaves of AO over-expressing plants compared with WT plants (Fig. 2C–E). AO over-expression resulted in no significant change in stomatal density per unit surface area (Table 1).
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) profile for fully-expanded leaf of well-watered wild-type and AO over-expressing plants. Bars represent SE (n=12 during photoperiod; n=6 during night-time).
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Effect of AO over-expression on leaf water management
Leaves of transgenic AO tobacco plants exhibited significantly reduced water loss following detachment, and this was reflected in the maintenance of leaf turgor compared with wild-type plants (Fig. 3A–C). This suggests that the decline in stomatal conductance associated with in vivo lowering of apoplast AA redox state by over-expression of AO may yield benefits in terms of improved leaf water management. Chlorophyll leaching assays showed no differences between wild-type and transgenic plants (Fig. 3D), suggesting that AO over-expression yielded no significant changes in cuticular resistance to water loss.
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Effect of dehydroascorbate on stomatal closure
Previous studies have shown that, although the total (AA+DHA) content remains unchanged in the apoplast of the AO transformants, the AA/DHA ratio is strongly affected, as only the oxidized form (DHA) can be detected in the apoplast of transgenic plants (Sanmartin et al., 2003). This prompted the investigatation as to whether the partial stomatal closure observed in AO over-expressing plants could be a consequence of increased DHA content. Stomatal aperture was measured in epidermal strips incubated either in buffer or in 1 mM DHA. As an additional control, epidermal strips were also incubated in 100 µM hydrogen peroxide. Data in Fig. 4 show that, at the end of the treatment, 100% of stomata in buffer-incubated strips from WT plants had opened, with a maximum width ranging from 5–10 µm (A). As expected, 100 µM H2O2 induced closure in
80% of stomata (C). Stomatal closure was also observed in DHA-incubated strips (E). Interestingly, stomata from AO over-expressing plants of the R50 transgenic line appeared insensitive to 100 µM H2O2 (D), and only a fraction of the stomata responded to 1 mM DHA (F) as compared to the WT. Higher H2O2 concentration (50 mM) induced complete stomatal closure in transgenic plants (data not shown).
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Histochemical localization of hydrogen peroxide in AO-overexpressing plants
To investigate hydrogen peroxide content and distribution in AO over-expressing plants further, whole leaves (Fig. 5A) and epidermal strips (Fig. 5C) of wild-type and AO over-expressing plants were incubated with DAB, in order to establish the in vivo effect of AO over-expression on H2O2 content. Quantification of the brown precipitate indicative of H2O2 showed significantly (P <0.05) enhanced H2O2 accumulation in both whole leaves (Fig. 5B) and guard cells (Fig. 5D). Much of the H2O2 present in guard cells was localized in chloroplasts, in agreement with previous reports (Zhang et al., 2001). Leaves of the same transgenic plants have previously been treated with H2O2, and this resulted in equal DAB staining levels, suggestive of similar peroxidase activity between R lines and WT plants (Fotopoulos et al., 2006).
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Assay of H2O2-scavenging enzyme activity
CAT and APX activities were measured in leaves of WT and transgenic plants. A statistically significant (P <0.05) reduction in APX activity was recorded in AO over-expressing lines R60 and R68 compared with wild-type plants (Fig. 6). No differences were found in CAT activity between transgenic and wild-type plants. This suggests that the observed increase in H2O2 content could, at least in part, be the result of reduced detoxifying enzymatic capacity in AO over-expressing plants.
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Detection of foliar ABA levels and NCED gene expression
Leaves of transgenic plants displayed a 4-fold increase in ABA levels compared with wild-type plants, suggesting a constitutive shift in ABA metabolism as a result of AO over-expression (Fig. 7A). To determine whether the increased foliar ABA content in AO over-expressing plants may be the result of a constitutive increase in the expression of NCED, a gene encoding a 9-cis-epoxycarotenoid dioxygenase that catalyses the key step in ABA biosynthesis (Tan et al., 1997), RNA blot analysis was carried out using a heterologous NCED probe from Cistus creticus on RNA isolated from leaves of plants grown under normal conditions. However, both wild-type and transgenic AO plants exhibited similar NCED mRNA abundance levels (Fig. 7B).
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Discussion |
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Results presented here show that the targeted over-expression of AO (Sanmartin et al., 2003) reduces stomatal aperture and, as a consequence, rates of leaf water loss are reduced upon detachment. This is consistent with previous measurements of stomatal conductance reported for additional AO over-expressing transformants as well as the data reported by Chen and Gallie (2004) for plants with reduced expression of DHA reductase (DHAR), which proved more resistant to water stress than WT plants. Both our AO over-expressing plants and Chen and Gallie's (2004) DHAR antisense plants exhibit higher apoplast DHA content than the WT. On the basis of these considerations, epidermal strips of WT tobacco plants were challenged with DHA, in order to understand whether this oxidized form of AA could be responsible for the observed stomatal closure. Notably, stomata of AO over-expressing plants were more closed than the WT in leaf disc assays (Fig. 2) compared with unchallenged epidermal strips (Fig. 4A, B). This apparent contradiction can be explained if one considers that buffer-incubated strips of both WT and over-expressing plants must have similar apoplast DHA content, due to dilution occurring in the free space. Our results clearly show that 2 h incubation in a buffered solution containing 1 mM DHA induces closure of approximately 80% of stomata in WT plants, implying that DHA is the active metabolite causing stomatal closure. This value is comparable to the effect of incubation (under the same conditions) in 100 µM hydrogen peroxide, a well-known inducer of stomatal closure (Bright et al., 2006). This observation opens the possibility that H2O2 may simply cause stomatal closure by oxidizing apoplastic ascorbate.
Many stresses (including pathogens, soil moisture deficit, salinity, and air pollutants) are known directly or indirectly to cause AA oxidation in the leaf apoplast. The extracellular matrix of epidermal cells and guard cells represents the first line of contact between the plant and its external environment, and there is growing evidence that the redox balance of this compartment plays a key role in regulating plant growth and development, plus perception and response to adverse conditions (Pignocchi and Foyer, 2003; Pignocchi et al., 2003, 2006). Our results lend support to this hypothesis, and also highlight DHA as a potential factor in the signalling pathway. The chemical structure of DHA, with its vicinal carbonyl group, gives the molecule its peculiar reactivity (Deutsch, 2000) with, amongst other compounds, thiols, whereby it induces oxidative protein folding (see for example, Banhegyi et al., 2003). Reversible modification of specific proteins by DHA could be important in cell signalling. Although possible DHA-regulated target proteins are still to be identified, it is conceivable that AA oxidation to DHA constitutes a vital signal transduction module governing plant reaction to stressful environmental conditions.
Under challenge by 100 µM H2O2, stomata of AO over-expressing plants are not as responsive as those of WT plants. This is quite surprising, since it is well-known that hydrogen peroxide is a powerful inducer of stomatal closure. Presumably this means that the signalling mechanism is altered in AO over-expressing plants. The observation also suggests that both DHA and H2O2 probably co-operate by reinforcing one another. In vivo localization of H2O2 indicated that AO over-expression in the apoplast resulted in H2O2 accumulation in both whole leaves and guard cells (Fig. 5). Similarly, Yamamoto et al. (2005) demonstrated increased H2O2 accumulation in the symplast and apoplast of transgenic tobacco plants over-expressing tobacco AO, while Chen and Gallie reported that in vivo lowering of the AA redox state, by manipulation of DHAR expression in the symplast, results in increased H2O2 content in guard cells and leaf apoplast (Chen and Gallie, 2004, 2005, respectively). APX activity was observed to be lower in transgenic plant lines R60 and R68 than in WT plants, grown under optimal conditions (Fig. 6A). Such a constitutive shift in ROS detoxifying enzyme capacity is consistent with the findings of Pignocchi et al. (2006), who reported that in vivo lowering of the AA redox state by AO over-expression resulted in a significant reduction in whole leaf APX activity in tobacco under non-stressed growth conditions. Moreover, these authors also report no differences in CAT activity, which supports our findings. This suggests that the observed increase in H2O2 content may, at least in part, be the result of reduced detoxifying enzymatic capacity. APX and CAT rapidly destroy the majority of H2O2 produced by metabolism, but low steady-state levels are allowed to persist to maintain redox signalling pathways (Noctor and Foyer, 1998). Therefore, higher H2O2 content in AO over-expressing plants may reflect the lowered APX activity. H2O2 can also react directly with AA, yielding DHA. Such self-reinforcement mechanisms are typical of signalling cascades and developmental processes (Meinhardt, 1994). A correlation between AA degradation products and H2O2 generation was also recently suggested by Green and Fry (2005). The authors reported that the pathway for AA degradation is located in the apoplast, demonstrating that AA degradation can occur via DHA, yielding oxalate plus L-threonate. This pathway operates extracellularly, potentially generating H2O2 at several steps in the pathway.
ABA plays a key role in stomatal regulation, both under non-stressed growth conditions and during soil-water deficit (Raschke, 1987). Leaves of AO over-expressing plants exhibited substantially higher ABA content compared with WT plants (Fig. 7). Although no change could be observed in NCED gene expression this is not conclusive. Rossel and colleagues (2006) recently showed no correlation between increased ABA content of alx8 Arabidopsis mutants and NCED3 expression, and there might be other NCED genes that are responsible for regulating the ABA content of plant tissues. Interestingly, López-Carbonell et al. (2006) recently observed a correlation between DHA content and ABA accumulation in foliage of the AA-deficient vtc1 mutant of Arabidopsis. This mutant, defective in the activity of the AA-biosynthetic enzyme GDP-mannose pyrophosphorylase, is characterized by low AA content (approximately 30% of WT) and low AA/DHA ratio (Conklin et al., 1996). The vtc1 mutant shows abnormal ABA accumulation, which correlates with higher DHA content observed in these plants. These data lend additional support to the hypothesis that DHA acts as an early activator of stomatal closure, and acts supposedly upstream of H2O2 production by NADH oxidase and ABA synthesis. ABA leads to stomatal closure via an increase in cytosolic Ca2+ concentration through H2O2-activated Ca2+ channels (Price et al., 1994; Grabov and Blatt, 1998; Pei et al., 2000; Zhang et al., 2001). Notably, AO over-expressing plants show increased expression of a plasma membrane Ca2+ channel-associated gene (Pignocchi et al., 2006), further supporting the view that imbalanced signalling occurs in the AO over-expressing transformants, leading to constitutive activation of mechanisms inducing stomata closure.
In conclusion, the data presented suggest that in vivo shifts in the apoplast AA redox state serve an important role in controlling stomatal aperture. The proposed model in which DHA elicits stomatal closure allows us to accommodate not only our data, but also other observations on stomatal movement (Chen and Gallie, 2004; Bright et al., 2006) within the same theoretical framework. A range of stressful conditions cause AA oxidation to DHA and, in turn, this molecule can modulate plant responses to stress in different ways, regulating ABA synthesis and increasing hydrogen peroxide production. The putative DHA-mediated mechanism of signal transduction is still unclear, but is probably related to DHA capability to cause reversible thiol disulphide exchanges in specific proteins, an aspect deserving of further detailed investigation.
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Acknowledgements |
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We thank Dr A Patakas and Mr A Mpeis for their valuable help with ABA detection, Ms G Tanou for her assistance with enzyme activity assays, and Dr D D'Haese and A Kaminska (Newcastle University, UK) for their experimental contribution. This work was supported through grants awarded to AKK (GR-GSRT-Greek British Council Bilateral Agreement) and JB (NERC and UK-DEFRA).
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