This Article Abstract Full Text (PDF) All Versions of this Article: 16/5/1143    most recent tpc.021584v1 Alert me when this article is cited Alert me if a correction is posted Services Email this article to a friend Similar articles in this journal Similar articles in ISI Web of Science Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Citing Articles Citing Articles via HighWire Citing Articles via CrossRef Citing Articles via ISI Web of Science (72) Citing Articles via Google Scholar Google Scholar Articles by Chen, Z. Articles by Gallie, D. R. Search for Related Content PubMed PubMed Citation Articles by Chen, Z. Articles by Gallie, D. R. Agricola Articles by Chen, Z. Articles by Gallie, D. R.

The Ascorbic Acid Redox State Controls Guard Cell Signaling and Stomatal Movement

Department of Biochemistry, University of California, Riverside, California 92521-0129

¹ To whom correspondence should be addressed. E-mail drgallie{at}citrus.ucr.edu; fax 909-787-3590.

	ABSTRACT

TOP ABSTRACT INTRODUCTION RESULTS DISCUSSION METHODS REFERENCES

 
H₂O₂ serves an important stress signaling function and promotes stomatal closure, whereas ascorbic acid (Asc) is the major antioxidant that scavenges H₂O₂. Dehydroascorbate reductase (DHAR) catalyzes the reduction of dehydroascorbate (oxidized ascorbate) to Asc and thus contributes to the regulation of the Asc redox state. In this study, we observed that the level of H₂O₂ and the Asc redox state in guard cells and whole leaves are diurnally regulated such that the former increases during the afternoon, whereas the latter decreases. Plants with an increased guard cell Asc redox state were generated by increasing DHAR expression, and these exhibited a reduction in the level of guard cell H₂O₂. In addition, a higher percentage of open stomata, an increase in total open stomatal area, increased stomatal conductance, and increased transpiration were observed. Guard cells with an increase in Asc redox state were less responsive to H₂O₂ or abscisic acid signaling, and the plants exhibited greater water loss under drought conditions, whereas suppressing DHAR expression conferred increased drought tolerance. Our analyses suggest that DHAR serves to maintain a basal level of Asc recycling in guard cells that is insufficient to scavenge the high rate of H₂O₂ produced in the afternoon, thus resulting in stomatal closure.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION RESULTS DISCUSSION METHODS REFERENCES

 
Of the antioxidants found in plants, ascorbic acid (Asc) is the most abundant. Asc is present in plant species in millimolar concentrations that range from 10 to 300 mM (Smirnoff, 2000) and serves as the major contributor to the cellular redox state. In its antioxidant role, Asc is used by ascorbate peroxidase (APX) to convert H₂O₂ to water, and Asc can directly scavenge superoxide, hydroxyl radicals, and singlet oxygen. Asc can serve as an enzyme cofactor, for example, for violaxanthin de-epoxidase (VDE) (Eskling et al., 1997), which catalyzes the conversion of violaxanthin to zeaxanthin (the xanthophyll cycle), which is required for the dissipation of excess excitation energy during nonphotochemical quenching. Asc also has been implicated in the regulation of cell elongation and progression through the cell cycle (reviewed in Horemans et al., 2000).

Reactive oxygen species serve an important signaling function, providing information about changes in the external environment. H₂O₂ has been implicated to play a signaling role in guard cells that permit gas exchange (e.g., entry of CO₂ and loss of water through the stomatal pore). Stomatal pores in many species open in the morning but close in the afternoon to limit water loss (Assmann, 1993; Assmann and Wang, 2001). Stomatal pores also close in response to water stress through the action of abscisic acid (ABA), which causes an increase in cytosolic Ca²⁺ concentration from H₂O₂-activated Ca²⁺ channels and from release from intracellular stores (Price et al., 1994; McAinsh et al., 1996; Grabov and Blatt, 1998; Guan et al., 2000; Hamilton et al., 2000; Pei et al., 2000; Zhang et al., 2001; Kohler and Blatt, 2002). Activation of plasma membrane–localized anion channels results in guard cell depolarization, potassium efflux, and loss of guard cell turgor and volume, which causes stomatal closure (Ishikawa et al., 1983; Thiel et al., 1992; Blatt and Armstrong, 1993; MacRobbie, 2000; Blatt, 2000; Schroeder et al., 2001a, 2001b). ABA causes an increase in H₂O₂ production, which serves as a signaling intermediate to promote stomatal closure (Price et al., 1994; Pei et al., 2000; Murata et al., 2001; Schroeder et al., 2001a, 2001b; Zhang et al., 2001), although a recent study has suggested that H₂O₂ may also function in a divergent pathway that controls stomatal movement (Kohler et al., 2003). H₂O₂-induced stomatal closure was reversed by exogenous application of ascorbate, which is consistent with its role as a scavenger of H₂O₂ (Zhang et al., 2001). Consequently, plants with increased ascorbate might be predicted to exhibit reduced responsiveness to ABA or H₂O₂ signaling.

Asc biosynthesis differs from that in mammals and is made after the oxidation of L-galactose to L-galactono-1,4-lactone, which in turn is oxidized to Asc. Once used, Asc is oxidized to the monodehydroascorbate radical that is reduced to Asc by monodehydroascorbate reductase (MDHAR) or disproportionates to Asc and dehydroascorbate (DHA). DHA undergoes irreversible hydrolysis to 2,3-diketogulonic acid or is reduced to Asc, a reaction catalyzed by dehydroascorbate reductase (DHAR), which requires glutathione. Thus, DHAR allows the plant to recycle DHA, thereby recapturing the Asc before it is lost. Because Asc is the major reductant in plants, DHAR serves to regulate the intracellular redox state. DHAR is expressed in rate-limiting amounts as overexpression of DHAR in tobacco (Nicotiana tabacum) leaves increased the Asc redox state (Chen et al., 2003). Plants overexpressing DHAR grew and flowered normally under well-watered conditions. However, given the role of Asc as a scavenger of H₂O₂, changes in the Asc redox state might be expected to alter stomatal movement, particularly under conditions in which ABA or H₂O₂ signaling activates stomatal closure.

Water stress results in the depletion of the Asc pool and triggers ABA-induced stomatal closure (Smirnoff and Pallanca, 1996; Leung and Giraudat, 1998; Pastori and Foyer, 2002). Such closure limits CO₂ assimilation and increases the concentration of NADPH as a consequence of a reduction in Calvin cycle activity. Under normal growth conditions, photoactivated chlorophyll transfers its excitation energy to the photosynthetic reaction centers. However, under conditions in which NADP is limiting (e.g., after stomatal closure in a water-stressed leaf), the excitation energy of the photoactivated chlorophyll is transferred to triplet (i.e., ground state) oxygen and excites it to the singlet form. Consequently, water stress increases the production of activated oxygen species (Bowler et al., 1992; Scandalios, 1993).

In this study, we have examined the role that the Asc redox state plays in regulating stomatal function. Plants in which DHAR expression was increased exhibited an increase in the Asc redox state in the leaf as a whole, in the apoplast, and in guard cells, whereas suppression of DHAR expression lead to the opposite effect. Guard cells with increased Asc redox state exhibited greater stomatal opening, both in the percentage of stomata that were open and in the degree of openness. They also exhibited reduced levels of H₂O₂, whereas those with a decreased Asc redox state had an elevated level of H₂O₂. Increasing the Asc redox state resulted in an increase in transpiration rate and stomatal conductance under normal growth conditions. Guard cells with a higher Asc redox state were less responsive to H₂O₂ or ABA signaling, and the plants exhibited enhanced water loss after the imposition of drought conditions. Decreasing the Asc redox state by repressing DHAR expression enhanced stomatal closure under normal growth conditions and after water stress. These observations suggest that the redox state of ascorbate plays an important role in controlling H₂O₂-mediated stomatal closure.

	RESULTS

TOP ABSTRACT INTRODUCTION RESULTS DISCUSSION METHODS REFERENCES

 
H₂O₂ and the Asc Redox State Are Diurnally Regulated
H₂O₂ has been implicated to serve as a signaling intermediate that promotes stomatal closure (McAinsh et al., 1996; Pei et al., 2000; Chen et al., 2003). Given that stomata close in the afternoon, the concentration of foliar H₂O₂ might be predicted to be higher in the afternoon than in the morning when stomata are open. To investigate this, the level of H₂O₂ was determined from expanded tobacco leaves at 2-h intervals during the course of a day. The concentration of H₂O₂ was low in early morning and increased during the afternoon, peaking in midafternoon before decreasing again throughout late afternoon and evening (Figure 1A). The diurnal increase in H₂O₂ concentration was approximately twofold.

View larger version (20K): [in this window] [in a new window]   Figure 1. Diurnal Regulation of H2O2, Asc, DHA, GSH, and GSSG. Leaf samples were taken at 2-h intervals from 6 AM to 10 PM and assayed for H2O2 (A), Asc (B), GSH (open circles), and DHA and GSSG (closed circles). Each was assayed in three independent replicates of leaves pooled from three independent plants, and the average and standard deviation were reported. The Asc and GSH redox state was determined from the Asc/DHA and GSH/GSSG ratios. The bar below each graph indicates dark and light periods.

Diurnal regulation of antioxidant enzymes also has been reported. Expression from Cat3, one member of the catalase gene family, exhibits circadian rhythm in maize (Zea mays) and Arabidopsis (Redinbaugh et al., 1990; Zhong and McClung, 1996; Polidoros and Scandalios, 1998). Circadian regulation from Arabidopsis Cat2 also was observed and required light as the synchronizer (Zhong et al., 1998). Diurnal changes in expression of superoxide dismutase (SOD) and glutathione reductase (GR) also has been reported in Aster tripolium (Erdei et al., 1998). Moreover, L-galactono--lactone dehydrogenase, which catalyzes the final step during Asc biosynthesis, is diurnally regulated in Arabidopsis (Tamaoki et al., 2003). To investigate whether the activity of antioxidant enzymes exhibits diurnal regulation, the foliar activities of DHAR, MDHAR, APX, GR, catalase (CAT), and SOD were determined at 2-h intervals. The activity of DHAR, MDHAR, GR, and CAT was low in the morning, rose as the day progressed to a maximum during the afternoon, and decreased again during the evening (Figure 2). APX activity was high from early morning to midafternoon, after which it declined. In contrast with these changes, SOD activity remained largely unchanged throughout the day (Figure 2). The increase in activity of some of these antioxidant enzymes was unlikely a daily stress response because the plants were watered twice a day and experienced a maximum daytime temperature of 25.9 ± 0.6°C and an afternoon light intensity of 1191 ± 244 µmol^–2 s^–1. Leaf water content was unchanged between the morning and the afternoon (Table 1), supporting the conclusion that growth of the plants under these conditions did not result in water stress. The diurnal increase of DHAR, MDHAR, GR, CAT, and APX activities, all of which are involved in H₂O₂ scavenging either directly or indirectly, suggests that the afternoon increase in H₂O₂ is not a consequence of a reduction in H₂O₂-scavenging enzymes. Moreover, the increase in these enzyme activities was not sufficient to prevent the diurnal increase in H₂O₂ during the afternoon.

View larger version (33K): [in this window] [in a new window]   Figure 2. Diurnal Regulation of DHAR, GR, CAT, APX, and SOD Activities. Leaf samples were taken at 2-h intervals from 6 AM to 10 PM and assayed for DHAR, MDHAR, GR, CAT, APX, and SOD. Enzyme activities were assayed in three independent replicates of leaves pooled from three independent plants, and the average and standard deviation were reported. The bar below each graph indicates dark and light periods.

View larger version (31K): [in this window] [in a new window]   Figure 3. Generation of DHAR-Overexpressing and DHAR-Suppressed Tobacco. (A) Protein gel blot analysis of DHAR in tobacco suppressed for endogenous DHAR (Ri11), vector-only control (Con), and overexpressing wheat DHAR (D1, C4, and B4). The positions of the His-tag wheat DHAR (His10-DHAR) and endogenous tobacco DHAR are indicated. DHAR activity measured in each leaf type is represented below each lane. (B) The amount of Asc and DHA was measured in expanding leaves, expanded leaves (which exhibited maximal photosynthetic activity), and presenescent leaves of DHAR-suppressed, control, and DHAR-overexpressing transgenic lines. Measurements were made in the afternoon from three independent replicates of leaves pooled from three independent plants, and the average and standard deviation were reported. The Asc redox state is indicated as the Asc/DHA ratio. FW, fresh weight. (C) MDHAR activity (10–8 mol NAPDH oxidized/min/mg protein) was measured in expanding leaves (black histograms), expanded leaves (gray histograms), and presenescent leaves (white histograms) of DHAR-suppressed (Ri11), control (Con), and DHAR-overexpressing (D1) plants. The enzyme activity was assayed in three independent replicates of leaves pooled from three independent plants, and the average and standard deviation were reported.

Reducing DHAR expression would be expected to limit its ability to reduce DHA to Asc and thus result in a decrease in the Asc redox state from either an increase in DHA and/or a reduction in Asc. This prediction was borne out in DHAR-suppressed plants in which a significant reduction in the Asc redox state was observed in expanding, expanded, and presenescent leaves of DHAR-suppressed plants compared with the control (P < 0.001, P < 0.05, P < 0.05, respectively). No significant difference was observed in Asc in expanding, expanded, and presenescent leaves of DHAR-suppressed plants relative to the control (P = 0.472, P = 0.109, P = 0.582, respectively) but was significantly lower than DHAR-overexpressing plants (P < 0.05, P < 0.05, P < 0.005, respectively). DHA was significantly higher in expanding and presenescent leaves of DHAR-suppressed plants relative to the control (P < 0.05 and P < 0.05, respectively) and DHAR-overexpressing plants (P < 0.005 and P < 0.01, respectively) but was not significantly different in control or DHAR-overexpressing expanded leaves (P = 0.777 and P = 0.082, respectively), suggesting that DHA might be lost more quickly in expanded leaves than in expanding and presenescent leaves of DHAR-suppressed plants.

Decreasing DHAR expression resulted in an increase in MDHAR activity (Figure 3C), whereas no change was observed in DHAR-overexpressing plants. The increase in MDHAR activity in DHAR-suppressed plants may be in response to the decrease in DHAR-mediated Asc recycling and the reduction in Asc redox state, which would increase this alternative pathway for the recycling of oxidized ascorbate. These data confirm that DHAR expression controls the Asc redox state and that its wild-type level of expression is rate limiting.

The leaf water potentials, measured for control, DHAR-overexpressing, and DHAR-suppressed leaves in the morning and the afternoon, were all well within nonstress values (Table 2), supporting the conclusion that the plants did not experience water stress under the growth conditions used and that changes in antioxidant enzyme activity observed in Figure 2 were a result of diurnal regulation.

During the morning when stomatal pores are open, no significant difference in the average stomatal aperture was observed between expanding leaves of DHAR-overexpressing (D1 in Figure 4) and vector-only control plants (Con in Figure 4), which was supported by quantitative measurements (Figure 5) (n = 32, P = 0.305). In addition, the average stomatal aperture in expanding leaves of DHAR-suppressed plants (R_i11 in Figure 5) was not significantly different from that of control plants (n = 34, P = 0.089), whereas it was significantly smaller than DHAR-overexpressing plants (n = 34, P < 0.05). The morning stomatal aperture in expanded and presenescent leaves of DHAR-overexpressing plants was significantly greater than that of control plants (n = 47, P < 0.005 and n = 53, P < 0.01, respectively), but the stomatal aperture in expanded and presenescent leaves of DHAR-suppressed plants was not significantly different from that of control plants (n = 47, P = 0.275 and n = 53, P = 0.546, respectively) (Figure 5). Greater than 80% of all stomata were open in expanding and expanded leaves in the morning, with only small differences in the percentage of stomata that were open among leaves of control, DHAR-overexpressing, or DHAR-suppressed plants. Fewer stomata were open in presenescent leaves in the morning compared with expanding or expanded leaves, but more stomata were open in presenescent DHAR-overexpressing leaves and fewer open in presenescent DHAR-suppressed leaves relative to the control (Figure 5).

View larger version (124K): [in this window] [in a new window]   Figure 4. Stomata in Tobacco with Altered Asc Redox State. Abaxial epidermis was collected in the morning and afternoon from expanding leaves of control (Con), DHAR-overexpressing (D1), and DHAR-suppressed (Ri11) tobacco and stained with 0.2% toluidine blue O before imaging using a compound microscope. A representative region from each line is presented.

View larger version (36K): [in this window] [in a new window]   Figure 5. The Asc Redox State Controls Stomatal Movement. Stomatal bioassay experiments were performed as described (Pei et al., 1997). Abaxial epidermis was collected from expanding, expanded, and presenescent leaves of control (Con), DHAR-overexpressing (D1), and DHAR-suppressed (Ri11) tobacco leaves. Epidermal strips were stained and imaged as described in Figure 2. The width and length of at least 30 stomatal apertures were measured and used to determine the stomatal aperture (width/length) with the standard deviation indicated. The percentage of stomata that were open (defined as having a width >1 µm) was determined from at least 400 stomata. The total stomatal open area was calculated from the average stomatal aperture area and percentage of stomata that were open and reported as the total area (µm2) per 100 stomata.

The Asc Redox State Governs the Level of H₂O₂ in Guard Cells
The diurnal decrease in the Asc redox state observed in Figure 1 may result from the inability of DHAR to recycle Asc at a rate rapid enough to scavenge the increased rate of H₂O₂ production that occurs in the afternoon as a consequence of excess light. To determine whether alterations in DHAR expression affect the level of guard cell H₂O₂, the level of H₂O₂ was determined in guard cells of DHAR-overexpressing, DHAR-suppressed, and control plants. Epidermal strips from leaves were loaded with 2',7'-dichlorofluorescin diacetate (H₂DCF-DA), and after washing, fluorescence emitted specifically from guard cells was detected at 510 to 530 nm and quantified using confocal microscopy. Much of the H₂O₂ present in guard cells was localized in chloroplasts, in good agreement with previous reports (Zhang et al., 2001). The level of H₂O₂ in guard cells from expanding leaves of DHAR-overexpressing plants collected during the morning was significantly lower than in control guard cells (n = 70, P < 0.001), whereas it was significantly higher in DHAR-suppressed guard cells (n = 70, P < 0.001) (Figure 6, Table 3). These differences in H₂O₂ level could not be explained by changes in APX activity, which was not significantly different from the level in DHAR-overexpressing guard cells (P = 0.075) or in DHAR-suppressed guard cells (P = 0.295) (Table 3). As expected, the level of guard cell H₂O₂ increased during the afternoon. However, the level of H₂O₂ in DHAR-overexpressing guard cells remained significantly lower (n = 70, P < 0.001 in expanding leaves) than that in the control, and H₂O₂ in DHAR-suppressed guard cells remained significantly higher (n = 70, P < 0.001 in expanding leaves) (Figure 6, Table 3). As observed in the morning, the changes in guard cell H₂O₂ could not be explained by changes in APX activity (Table 3). APX activity in expanding leaves of DHAR-overexpressing or DHAR-suppressed plants was not significantly different from that in control plants in the afternoon (P = 0.061 and P = 0.507, respectively). The level of guard cell H₂O₂ in presenescent leaves was higher than that in guard cells of expanding leaves, consistent with previous reports that the level of active oxidative stress increases with leaf age (Havaux et al., 2000; Munne-Bosch and Alegre, 2002). Although the level of guard cell H₂O₂ increased with leaf age, the same differences in guard cell H₂O₂ were observed among the DHAR lines regardless of leaf age (Figure 6, Table 3).

View larger version (59K): [in this window] [in a new window]   Figure 6. The Asc Redox State Controls H2O2 Concentration in Guard Cells. Abaxial epidermis was collected in the morning from control (Con), DHAR-overexpressing (D1), and DHAR-suppressed (Ri11) tobacco leaves. H2O2 production was revealed after loading with H2DCF-DA. Fluorescence (excitation 465 to 495 nm, emission 510 to 530 nm) was recorded using confocal microscopy. A representative region from each line is presented.

View larger version (30K): [in this window] [in a new window]   Figure 7. Alteration of the Asc Redox State in Guard Cells. The level of Asc (A), DHA (B), GSH (D), and GSSG (E) was measured from guard cells isolated from control (Con), DHAR-overexpressing (D1), and DHAR-suppressed (Ri11) tobacco leaves collected in the morning (AM) and afternoon (PM). The Asc redox state (C) determined by [Asc]/[DHA] and the GSH redox state (F) determined by [GSH]/[GSSG] also were determined. FW, fresh weight.

Asc is exported to the apoplast where it functions in cell wall synthesis, protects against exposure to external ROS, and is reimported as DHA. To determine whether the guard cell Asc redox state is similar to that of the apoplast, the level of Asc and DHA in the apoplast was measured. No glucose-6-phosphate dehydrogenase activity, used as a cytosolic marker, was detected, suggesting that the apoplastic fluid was contaminated with little if any cytoplasm. DHA (Figure 8B) was present in the apoplast at a concentration substantially higher than that of Asc (Figure 8A), resulting in an Asc redox state of only 0.36 in control leaves (Figure 8C). This is in good agreement with previous reports (Veljovic-Jovanovic et al., 2001) and is in marked contrast with the redox state of 1.58 in control guard cells (Figure 7C). The level of Asc and DHA in the apoplast increased in DHAR-overexpressing leaves, and the Asc redox state increased to 0.7, whereas the apoplastic level of Asc and DHA decreased in DHAR-suppressed leaves, resulting in a redox state of 0.3 (Figure 8C). No significant change in apoplastic-localized ascorbate oxidase activity was observed between DHAR-overexpressing and control plants, and only a small decrease was observed in DHAR-suppressed plants (Figure 8D), neither of which could account for the observed changes in the apoplastic Asc redox state. These observations demonstrate that the redox state of guard cells differs substantially from that of the apoplast, suggesting that the guard cell redox state is established autonomously.

View larger version (13K): [in this window] [in a new window]   Figure 8. Alteration of the Apoplastic Asc Redox State. The level of Asc (A) and DHA (B) was measured from apoplastic fluid from control (Con), DHAR-overexpressing (D1), and DHAR-suppressed (Ri11) tobacco leaves collected in the morning (AM) and afternoon (PM). The Asc redox state (C) determined by [Asc]/[DHA] also was determined. FW, fresh weight. Ascorbate oxidase (AOX) was also measured (D).

View larger version (38K): [in this window] [in a new window]   Figure 9. Expression Analysis of DHAR in Guard Cells and Leaves. (A) Total RNA was isolated from leaves (L) and guard cell (GC) protoplasts isolated from control tobacco leaves collected in the morning (AM) and afternoon (PM). The relative abundance of transcript amounts for DHAR, MDHAR, SOD, and actin were determined using RT-PCR. The number of cycles of amplification required to detect a transcript while maintaining its amplification in the linear range was 28 for DHAR, 35 for MDHAR, 32 for SOD, and 32 for actin for both whole leaves and isolated guard cells. (B) Total RNA was isolated from leaves and guard cell protoplasts isolated from control (Con), DHAR-overexpressing (D1), and DHAR-suppressed (Ri11) tobacco leaves collected in the morning (AM) and afternoon (PM). The number of cycles of amplification required to detect a transcript while maintaining its amplification in the linear range was 28 and 32 for DHAR, 30 and 35 for MDHAR, 28 and 32 for SOD, and 32 and 28 for actin for whole leaves and isolated guard cells, respectively.

After treatment to open the stomata, no significant difference in the aperture or percentage of stomata that were open was observed among DHAR-overexpressing, DHAR-suppressed, and control leaves as expected (Figure 10). Treatment with H₂O₂ or ABA promoted stomatal closure in all leaf types (Figure 10). However, a greater percentage of stomata from leaves with an increased Asc redox state remained open and of these, they exhibited a larger average aperture after treatment with H₂O₂ or ABA than did stomata from control leaves (for H₂O₂, n = 383, P < 0.001; for ABA, n = 341, P < 0.001). These changes resulted in approximately twice as much total open stomatal area relative to control leaves (Figure 10). The response of guard cells with reduced Asc redox state to H₂O₂ or ABA was similar to that of the control (Figure 10). These results support the conclusion that an increase in the Asc redox state reduces guard cell responsiveness to H₂O₂ and ABA signaling.

View larger version (35K): [in this window] [in a new window]   Figure 10. The Asc Redox State Controls Guard Cell Responsiveness to H2O2 and ABA Signaling. Abaxial epidermis was collected in the morning from expanded leaves of control (Con), DHAR-overexpressing (D1), and DHAR-suppressed (Ri11) tobacco leaves. After incubation in CO2-free buffer under a photon flux density of 200 µmol–2 s–1 to promote stomatal opening, stomatal closure was induced by 50 µM ABA or 1 mM H2O2. Epidermal strips were stained with toluidine blue O or loaded with fluorescence dye to determine the production of H2O2. The width and length of at least 70 stomatal apertures were measured and used to determine aperture area with the standard deviation indicated. The percentage of stomata that were open was determined from at least 400 stomata. The total stomatal open area was calculated by multiplying the area of the average stomatal aperture by the percentage of stomata that were open and reported as the total area (µm2) per 100 stomata.

View larger version (18K): [in this window] [in a new window]   Figure 11. The Asc Redox State Controls Transpiration and Stomatal Conductance. In situ rates of transpiration (A) and stomatal (B) conductance were measured in the afternoon from control (open circles), DHAR-overexpressing (closed circles), and DHAR-suppressed (closed triangles) tobacco leaves using a TPS-1 portable photosynthesis system. Transpiration and stomatal conductance were assayed in every other leaf in three independent replicates of leaves pooled from three independent plants, and the average and standard deviation were reported.

View larger version (46K): [in this window] [in a new window]   Figure 12. The Asc Redox State Controls Water Loss from Tobacco Leaves. Expanded leaves were detached from well-watered plants in the afternoon and immediately weighed. (A) Images of representative expanded leaves were taken at 30 and 120 min after their detachment. (B) Water loss from the detached leaves held at room temperature was followed by determining leaf weight every 5 min for 40 min. Four leaves from separate plants were used. The rate of water loss as loss in leaf weight was plotted against time, and the average and standard deviation were reported.

View larger version (18K): [in this window] [in a new window]   Figure 13. The Asc Redox State Controls Drought Tolerance. CO2 assimilation was measured in every other leaf from control (open circles), DHAR-overexpressing (closed squares), and DHAR-suppressed (closed triangles) tobacco leaves (A) under well-watered conditions and after the imposition of a severe drought (B). CO2 assimilation was measured using a TPS-1 portable photosynthesis system. Drought stress was evident in control and DHAR-overexpressing plants by their leaf wilting.

	DISCUSSION

TOP ABSTRACT INTRODUCTION RESULTS DISCUSSION METHODS REFERENCES

 
Ascorbate might be expected to be involved in regulating H₂O₂-induced stomatal closure through its ability to scavenge H₂O₂ (Schroeder et al., 2001a, 2001b; Chen et al., 2003). In support of this, the level of H₂O₂ increased and the Asc redox state decreased in guard cells in the afternoon, which correlated with stomatal closure. In this study, we have investigated the role that Asc plays in regulating stomatal movement. Increasing the Asc redox state was achieved through overexpression of DHAR, whereas reducing it was achieved through suppression of DHAR. The increase in the Asc redox state in DHAR-overexpressing plants can be understood through the recycling function of DHAR, which would increase the likelihood that DHA is converted to Asc before being lost through decay. The increase in the level of DHA in DHAR-suppressed plants would be expected from their reduced ability to convert DHA into Asc. Changes in the level of DHAR expression in guard cells resulted in the alteration of their Asc redox state in that an increase was measured in DHAR-overexpressing guard cells, whereas a decrease was measured in DHAR-suppressed guard cells. Increasing foliar DHAR expression also increased the apoplastic Asc redox state, whereas decreasing DHAR expression had the opposite effect. However, the apoplast and guard cell Asc redox states differed substantially, suggesting that the Asc redox state of guard cells is established autonomously.

During the afternoon, when stomatal closure occurs, increasing the Asc redox state increased the percentage of stomata that remained open and the size of the stomatal aperture. The largest change in stomatal behavior was observed in expanding and expanded leaves that decreased as the leaves aged, suggesting the possibility that signaling in guard cells may become less efficient before entry into the senescence program. Decreasing the Asc redox state reduced the average stomatal aperture and reduced the percentage of stomata that were open. The change in stomatal movement was inversely correlated with the level of H₂O₂ present in guard cells, which was lower in guard cells with a high Asc redox state and higher in guard cells with a lower Asc redox state. These differences in H₂O₂ concentration correlated with the observed differences in stomatal behavior in the afternoon. The fact that the observed differences in H₂O₂ concentration in the morning do not result in similar differences in stomatal behavior suggests that the absolute level of H₂O₂ in guard cells of the lines examined may not be high enough at that time to trigger stomatal closure. The observation that treatment with H₂O₂ promoted stomatal closure to a lesser extent in guard cells with a higher Asc redox state indicates that the increase in Asc redox state reduced guard cell responsiveness to H₂O₂ signaling. The finding that zeaxanthin is involved in blue light sensing that results in stomatal opening (Zeiger, 2000) might suggest that changes in Asc, required for the VDE catalyzed synthesis of zeaxanthin, may affect stomatal opening through changes in the xanthophyll cycle. However, reduction of Asc in the Arabidopsis vtc-1 mutant did not affect the xanthophyll cycle (Veljovic-Jovanovic et al., 2001), suggesting that the wild-type concentration of Asc is not rate limiting for VDE. The wild-type level of Asc in the DHAR-suppressed plants suggests that the xanthophyll cycle would be unaffected under normal growth conditions. Moreover, the degree of stomatal opening in DHAR-overexpressing and DHAR-suppressed plants in the morning was less affected by changes in the Asc redox state than was the degree of stomatal closure in the afternoon, suggesting that the effects of altering the Asc redox state may be limited to H₂O₂ signaling. In addition, because H₂O₂ serves as an intermediary of ABA-induced stomatal closure, responses elicited by ABA should be similar to those elicited by H₂O₂. This prediction was borne out as treatment with ABA promoted stomatal closure, but guard cells with a higher Asc redox state were less responsive to ABA than were control guard cells.

The effect that changes in the Asc redox state have on stomatal movement would be expected to affect stomatal conductance and the rate of transpiration. Under well-watered conditions, increasing the Asc redox state increased the rate of transpiration and stomatal conductance, whereas decreasing it reduced transpiration. Because ABA-induced stomatal closure is the primary means by which plants respond to conditions of water stress (Leung and Giraudat, 1998), increasing the Asc redox state would be expected to affect the ability of a plant to reduce transpiration during water stress. This was confirmed using whole plant and detached leaf assays. Increasing the Asc redox state increased the rate of water loss threefold, whereas reducing the Asc redox state decreased water loss up to 30%. In addition, after a severe water stress that caused leaf wilting in DHAR-overexpressing and control plants and in which the rate of CO₂ assimilation was virtually abolished, leaves with a lower Asc redox state retained turgor, and their rate of CO₂ assimilation was only slightly reduced relative to well-watered conditions. It should be noted that decreasing the Asc redox state reduced the rate of CO₂ assimilation compared with control plants, which is consistent with the reduction in total open stomatal area observed for DHAR-suppressed leaves. These data suggest that increasing the Asc redox state results in a higher rate of transpiration under normal growth conditions and reduced responsiveness to water stress. This in turn results in an enhanced rate of water loss as a consequence of the increase in the total open stomatal area. Conversely, decreasing the Asc redox state reduced transpiration and CO₂ assimilation from young leaves under normal growth conditions as a consequence of the reduction in the open stomatal area, but it also reduced water loss resulting in increased drought tolerance. These observations support the conclusion that Asc affects leaf function through alterations in stomatal movement.

Previous studies of plants with altered levels of antioxidants have not reported a change in stomatal function. Plants engineered to express higher levels of GR, the enzyme required in GSH recycling, were shown to have elevated Asc levels (Foyer et al., 1995). Although the plants exhibited little change in CO₂ assimilation, the time of day when the measurements were taken or the age of the leaves, both of which are crucial in observing stomatal behavior, were not indicated, making direct comparisons difficult. The Arabidopsis vtc-1 mutant, which contains only 30% of the wild-type level of Asc but retains an altered Asc redox state, exhibits a normal level of photosynthesis (Veljovic-Jovanovic et al., 2001). This study did not employ plants with elevated Asc. The DHAR-suppressed plants used in this study are not Asc deficient but rather have a decreased Asc redox state that may explain the differences observed between the present plants and the vtc-1 mutant. Moreover, the stomatal function of the vtc-1 mutant was not examined during the course of the day or in response to H₂O₂ or ABA, making it difficult to draw conclusions between a lower level of Asc versus a decrease in Asc redox state. However, one significant difference between the vtc-1 mutant and plants with altered DHAR expression is that the former affects an enzyme in the Asc biosynthetic pathway, whereas DHAR affects Asc recycling, which may be expected to be a more rapid means to respond to the generation of reactive oxygen species. The observation that DHAR-suppressed plants (in which the level of Asc is unchanged but have a decreased Asc redox state) exhibit greater stomatal closure suggests that it is the Asc redox state rather than the level of Asc that affects guard cell function.

The role of Asc in guard cell functioning can thus be understood through its role as a scavenger of H₂O₂ whereby the balance between H₂O₂ production and the Asc redox state establishes whether the H₂O₂ concentration rises to a level that can trigger stomatal closure. The diurnal increase in H₂O₂ during the afternoon is likely a result of photosynthetic-related processes, such as photorespiration and oxygen photoreduction (the Mehler peroxidase reaction), which serves to maintain electron flow through photosystem I and maintains its correct function. The Mehler reaction comprises the transfer of electrons from photosystem I to oxygen to form superoxides that SOD then disproportionates to O₂ and H₂O₂, the latter of which is reduced to water by APX using Asc as the reductant. Together, the Mehler reaction and the Asc-mediated reduction of H₂O₂ form the water-water cycle in which Asc is consumed to help protect against photoinhibition (Asada, 1999). ABA also can elicit H₂O₂ production as part of the signaling required to promote stomatal closure (Price et al., 1994; Pei et al., 2000; Murata et al., 2001; Schroeder et al., 2001a, 2001b; Zhang et al., 2001). Moreover, peroxisomes generate H₂O₂ as part of photorespiration and may contribute to the level of H₂O₂ present in guard cells. Interestingly, although APX activity could be detected in isolated guard cells, CAT expression or activity, which is normally used to detoxify peroxisome-generated H₂O₂, could not be detected (data not shown), suggesting that H₂O₂ may be detoxified in tobacco guard cells without substantial contribution by CAT. Consumption of Asc in detoxifying H₂O₂ generated by these processes as well as others produces DHA. Because DHAR is present in rate-limiting amounts, the diurnal increase in DHA during the afternoon may reflect a rate of H₂O₂ production and consumption of Asc that exceeds the ability of DHAR to efficiently regenerate Asc. Thus, under conditions of nonsaturating light (e.g., during early morning), the existing Asc redox state may be sufficient to scavenge a basal level of H₂O₂ production and prevent stomatal closure, whereas in excess light, the consumption of Asc by the water-water cycle, the inability of a rate-limiting amount of DHAR to regenerate Asc efficiently, and the resulting decrease in the Asc redox state may allow the concentration of H₂O₂ to increase to a level that signals stomatal closure. Therefore, a model integrating these observations can be proposed in which stomatal closure is triggered when conditions of excess light and the accompanying elevated production of H₂O₂ lower the Asc redox state to a level that it can no longer maintain a low concentration of H₂O₂. Overexpression of DHAR provides a larger reservoir of Asc and more efficient regeneration of Asc that can scavenge higher levels of H₂O₂, thereby preventing its accumulation to a level sufficient to trigger stomatal closure. This would explain the greater open stomatal area, increased stomatal conductance, higher transpiration rate, higher rate of water loss, decreased tolerance to water stress, and reduced guard cell responsiveness to ABA and H₂O₂ signaling that are observed in DHAR-overexpressing plants. Conversely, reducing DHAR expression below wild-type levels reduces the Asc redox state to a level similar to that observed in control plants during the afternoon. The less efficient regeneration of Asc would lead to an elevated accumulation of H₂O₂, which in turn would trigger a greater degree of stomatal closure even under nonstress conditions. Therefore, the guard cell Asc redox state would be expected to control stomatal movement through modulating H₂O₂ signaling.

	METHODS

TOP ABSTRACT INTRODUCTION RESULTS DISCUSSION METHODS REFERENCES

 
DNA Constructs, Plant Transformation, and Protein Gel Blot Analysis
Full-length wheat (Triticum aestivum) and tobacco (Nicotiana tabacum) DHAR cDNAs (accession numbers AY074784 and AY074787, respectively) were isolated as described previously (Chen et al., 2003). Transgenic tobacco (N. tabacum cv Xanthi) expressing the His-tagged wheat DHAR from the CaMV 35S promoter (in the binary vector pBI101) was generated using Agrobacterium tumefaciens as described (Chen et al., 2003). Transgenic tobacco plants suppressed for DHAR were identified after the introduction of a tobacco DHAR construct in pBI101. Plants were grown in a greenhouse supplied with charcoal-filtered air and were used when 13 to 15 weeks old.

Anti-DHAR antiserum raised against DHAR purified from wheat seedlings was used for protein gel blot analysis. Protein extracts were resolved using standard SDS-PAGE, and the protein was transferred to a 0.22-µm nitrocellulose membrane by electroblotting. After transfer, the nitrocellulose membranes were blocked in 5% milk and 0.01% thimerosal in TPBS (0.1% Tween 20, 13.7 mM NaCl, 0.27 mM KCl, 1 mM Na₂HPO₄, and 0.14 mM KH₂PO₄) followed by incubation with primary antibodies diluted typically 1:1000 to 1:2000 in TPBS with 1% milk for 1.5 h. The blots were then washed twice with TPBS and incubated with goat anti-rabbit horseradish peroxidase–conjugated antibodies (Southern Biotechnology Associates, Birmingham, AL) diluted to 1:5000 to 1:10,000 for 1 h. The blots were washed twice with TPBS, and the signal was detected typically between 1 to 15 min using chemiluminescence (Amersham, Buckinghamshire, UK).

Plant Growth Conditions
All plants were grown in 5-gallon pots with commercial soil in a glasshouse supplied with charcoal-filtered air and were not pot bound for the experiments. Experiments were performed and repeated from November through the winter to avoid excessive heat or light. Plants were watered to saturation twice a day (7 AM and 1 PM) to ensure that the soil was never dry. Plants were grown under natural light conditions in a 10-h-light and 14-h-dark cycle. The average temperature during the day was 25.9 ± 0.6°C and during the night was 20.2 ± 0.5°C. The average light intensity in the morning (9 AM) was 514 ± 206 µmol m^–2 s^–1 and in the afternoon (1 PM) was 1191 ± 244 µmol m^–2 s^–1. Plants used in the experiments had produced 20 leaves and had not yet produced an inflorescence.

Enzyme Assays
DHAR activity was assayed essentially as described (Hossain and Asada, 1984). Tobacco leaves were ground in extraction buffer (50 mM Tris-HCl, pH 7.4, 100 mM NaCl, 2 mM EDTA, and 1 mM MgCl₂), and soluble protein was obtained after a 5-min centrifugation at 13,000 rpm. DHAR was assayed from an equal amount of protein as described (Bradford, 1976) in 50 mM K₂HPO₄/KH₂PO₄, pH 6.5, 0.5 mM DHA, and 1 mM GSH and its activity followed by an increase in absorbance at 265 nm. MDHAR (10^–8 mol NAPDH oxidized/min/mg protein), SOD (units to inhibit nitroblue tetrazolium photoreduction by 50%), CAT (10^–6 mol H₂O₂ reduced/min/mg protein), GR (10^–8 mol NAPDH oxidized/min/mg protein), and APX (10^–8 mol Asc oxidized/min/mg protein) activities were determined as described (Gainnopolitis and Pies, 1977; Aebi, 1984; de Pinto et al., 2000). Ascorbate oxidase activity present in apoplastic wash fluid (obtained as described below) was determined from the decrease in A₂₆₅ (extinction coefficient of 14 mM^–1cm^–1) at 25°C in a reaction mixture containing 0.1 M sodium phosphate, pH 5.6, 0.5 mM EDTA, and 100 µM Asc as described (Pignocchi et al., 2003).

Guard Cell Isolation
Guard cells and guard cell protoplasts were isolated essentially as described (Kruse et al., 1989). Pieces collected from expanding leaves of 8-week-old plants were transferred to a blender jar in 100 mL of homogenization buffer (10% Ficoll, 5 mM CaCl₂, and 0.1% polyvinylpyrrolidone 40) and homogenized at high speed for 1 to 2 min. Epidermal fragments were collected using a nylon mesh (220 µm) and rinsed thoroughly with water. To isolate guard cell protoplasts, the epidermal fragments were incubated in 40 mL of digestion solution (0.25 M D-mannitol, 0.7% cellulysin [Behring Diagnostics, La Jolla, CA], 1 mM CaCl₂, 0.1% polyvinylpyrrolidone 40, and 1 µg/mL of pepstatin A, pH 5.5) with shaking for 45 min, collected on nylon mesh, rinsed with 0.25 M mannitol and 1 mM CaCl₂, and incubated in fresh enzyme solution for an additional 45 min. Guard cell protoplasts were then released within 3 h in a 50-mL solution of 0.35 M mannitol, 1 mM CaCl₂, 1% cellulase Onozuka R-10 (Yakult Honsha, Tokyo, Japan), 0.01% pectolyase Y-23 (Seishin Pharmaceutical, Tokyo, Japan), and 1 µg/mL of pepstatin A, pH 5.5. Epidermal debris were removed using nylon mesh (30 µm), and the protoplasts were collected by centrifugation at 60g for 7 min and were washed three times with 0.35 M mannitol and 1 mM CaCl₂.

Asc and DHA Measurements
Asc was measured as described (Foyer et al., 1983). Apoplastic levels were obtained essentially as described (Pignocchi et al., 2003) in which leaves from 8-week-old tobacco were collected, the midvein removed, and 2-cm² pieces vacuum infiltrated at –70 kPa with chilled 10 mM citrate, pH 3.0, for 10 min. Leaves were blotted dry, rolled, and centrifuged in a prechilled syringe at 2000 rpm for 10 min at 4°C. An equal volume of 2.5 M HClO₄ (for the ascorbate assay) or 10% metaphoric acid (for the GSH assay) was added to the apoplastic wash fluid. Guard cells were isolated as described (Kruse et al., 1989). For whole leaf assays, leaves were ground in 2.5 M HClO₄ and centrifuged at 13,000 rpm for 10 min. Two volumes of 1.25 M Na₂CO₃ were added to the supernatant, and after centrifugation, 100 µL was added to 895 µL 100 mM K₂HPO₄/KH₂PO₄, pH 5.6. Asc was determined by the change in absorbence at 265 nm after the addition of 0.25 units of ascorbate oxidase. The total amount of reduced and oxidized ascorbic acid (i.e., Asc and DHA) was determined by reducing DHA to Asc (in a reaction containing 100 mM K₂HPO₄/KH₂PO₄, pH 6.5, 2 mM GSH, and 0.1 µg recombinant wheat DHAR protein incubated at 25°C for 20 min) before measuring Asc. The amount of DHA was determined as the difference between these two assays. The level of GSH and GSSG was determined as described (Shimaoka et al., 2000).

Determination of Leaf Water Potential and Leaf Water Content
Leaf water potential was determined at 10 AM and 2 PM using a pressure chamber (Model 3000; Soilmoisture Equipment, Santa Barbara, CA) as described by Gómez-Cadenas et al. (1996). Data are presented as the average and standard deviation of six replicate leaves. Leaf water content was measured at 10 AM and 2 PM. Leaf fresh weight was measured immediately, and their water saturated weight was determined after allowing the leaf to absorb water until saturation. Leaf dry weight was measured after drying at 80°C. The difference in water weight between fresh and water-saturated leaves was then calculated in the morning and afternoon and expressed on a leaf dry weight basis. Data are presented as the average and standard deviation of six replicate leaves.

Stomatal Measurements
Stomatal bioassay experiments were performed as described (Pei et al., 1997). Abaxial epidermis strips were loaded onto glass slides with 100 µL of buffer A (50 mM KCl and 10 mM Mes, pH 6.1). After staining with 0.2% toluidine blue O for 20 s, the sample was rinsed twice with distilled water and imaged using a compound microscope (Leica, Wetzlar, Germany). For each sample, the percentage of stomata that were open (defined as having a width >1 µm) was determined from at least 400 stomata. The width and length of the opening (i.e., aperture) of only those stomata that were open were measured and used to calculate the average stomatal aperture (width/length). The width and length of at least 30 stomatal pores were measured. The width and length of open stomatal apertures also were used to calculate the stomatal aperture area [ x (width/2) x (length/2)], which together with the percentage of stomata that remained open, were used to calculate the total open stomatal area per unit leaf area containing 100 stomata [i.e., (average stomatal aperture area) x (percentage of stomata that remained open) x 100].

Induction of stomatal closure by ABA and H₂O₂ was investigated using epidermal strips that were first incubated in CO₂-free buffer A for 2 h at 22 to 25°C under a photon flux density of 200 µmol m^–2 s^–) to promote stomatal opening. ABA (50 µM final concentration, dissolved in 95% ethanol, with equal volume of ethanol used as a control) or H₂O₂ (1 mM final concentration) was added to the buffer. After the indicated time, samples were stained with toluidine blue O for image recording or loaded with fluorescence dye to determine the production of H₂O₂.

Superoxide and H₂O₂ Measurements
Superoxide was assayed as described (Sutherland and Learmonth, 1997). Leaves were abraded with sandpaper, and 20 to 30 0.5-cm-diameter discs were shaken in 5 mL 10 mM citrate buffer, pH 6.0, to which 50 µM of the tetrazolium dye, sodium, 3'-[1-[phenylamino-carbonyl]-3,4-tetrazolium]-bis(4-methoxy-6-nitro)benzene-sulfonic acid hydrate (XTT), was added. The reduction of XTT in 1 mL of the solution was recorded every 5 min at 470 nm, and the amount of superoxide was determined from the rate of XTT reduction.

Leaf discs (100 mg) were ground in liquid nitrogen, extracted in 150 µL of 25 mM HCl, and centrifuged at 5000g for 5 min at 4°C. Pigments were removed by vortexing in the presence of activated charcoal and centrifugation at 12,000g for 15 min. H₂O₂ was determined by measuring relative fluorescence (excitation at 315 nm, emission at 425 nm) in a reaction containing Hepes-NaOH, pH 7.5, 20 µL of extract, 500 µM homovanillic acid, and 0.5 unit horseradish peroxidase against a standard curve as described (Creissen et al., 1999).

H₂O₂ production from guard cells was examined by loading epidermal peels with H₂DCF-DA as described (Lee et al., 1999). To reduce variation, epidermal strips from control, DHAR overexpressors, and DHAR-suppressed leaves were placed on a single microscope slide loaded with 200 µL of 20 µM H₂DCF-DA for 5 min. After washing with buffer A, dye emission (excitation 465 to 495 nm, emission 510 to 530 nm) from at least 70 guard cells was recorded using an Insight confocal microscope (Kent, WA). Fluorescence intensity, measured as the pixel intensity present specifically within a defined area encompassing a guard cell, was obtained using MCID Elite image software (version 7.0; Imaging Research, St. Catharines, Ontario, Canada). A minimum of 70 guard cell pairs was measured. Representative background pixel intensity was obtained from the average of 10 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 fluorescence intensity of guard cell.

RT-PCR Analysis
Total nucleic acid was isolated from whole leaves and guard cell protoplasts, and the DNA was removed using RQ1 RNase free DNase I (Promega, Madison, WI). The absence of DNA in the samples was confirmed after saturating PCR (38 cycles) with actin-specific primers. One microgram of total RNA was used for cDNA synthesis using Omniscript RT (Qiagen, Valencia, CA) with oligo(dT)₂₀ as the primer. PCR reactions contained 1x buffer, 2 µL of the reverse transcription reaction, and 1.5 µg of forward and reverse primers in a total reaction volume of 25 µL using the following conditions: 94°C/5 min; 94°C/50 s, annealing temperature (see below)/50 s; 72°C/1 min, 72°C/5 min. PCR products were visualized on ethidium bromide–stained 1.2% agarose gels.

Primers used were as follows: actin (X63603) (annealing temperature, 52°C), forward, 5'-CGCGAAAAGATGACTCAAATC-3' and reverse, 5'-AGATCCTTTCTGATATCCACG-3'; CAT (U93244) (annealing temperature, 55°C), forward, 5'-CGGATACCTGAGCGTGTTGTTCATG-3' and reverse, 5'-GTGATTATTGTGATGAGCACAC-3'; MDHAR (BQ842867) (annealing temperature, 55°C), forward, 5'-ACTTCAAATAGCCGTTTTTAATCCA-3' and reverse, 5'-AGTTGAACATGTTGATCATTCTC-3'; FeSOD (M55090) (annealing temperature, 53°C), forward, 5'-TGCTTTGGAGCCTCATATGAG-3' and reverse, 5'-AAGTCCAGATAGTAAGCATGC-3'; tobacco DHAR (AY074787) (annealing temperature, 55°C), forward, 5'-AATTGGATCCCTGATTCTGATGT-3' and reverse, 5'-GCGAAACAACGGGATTATAATTATG-3'; wheat DHAR (AY074784) (annealing temperature, 59°C), forward, 5'-AATTGGATCCCTGATTCTGATGT-3' and reverse, 5'-GGATCCAGGGGCTTACGGGTTCACTTTC-3'; to detect wheat and tobacco DHAR (annealing temperature, 59°C), forward, 5'-AATTGGATCCCTGATTCTGATGT-3' and reverse, 5'-AGATGGTA(G/C)AG(C/T)TTCGGAGCCA-3'.

Leaf Water Loss Assay
Expanded leaves were detached from well-watered plants and immediately weighed. Water loss from the detached leaves held at room temperature was followed by determining leaf weight every 5 min for 40 min. Four leaves from separate plants were used. The rate of water loss was plotted as the loss in leaf weight over time. In situ rates of CO₂ assimilation, transpiration, and stomatal conductance were measured with the TPS-1 portable photosynthesis system (PP Systems, Haverhill, MA). Every other leaf on a plant was measured.

Sequence data from this article have been deposited with the EMBL/GenBank data libraries under accession numbers AY074784 and AY074787.

	Acknowledgments

 
The authors thank Patricia Springer for use of the light microscope. This work was supported by Grant NRICGP 02-35100-12469 from the USDA and the University of California Agricultural Experiment Station.

	Footnotes

 
The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantcell.org) is: Daniel R. Gallie (drgallie{at}citrus.ucr.edu).

Article, publication date, and citation information can be found at www.plantcell.org/cgi/doi/10.1105/tpc.021584.

Received February 5, 2004; accepted February 29, 2004.

	REFERENCES

TOP ABSTRACT INTRODUCTION RESULTS DISCUSSION METHODS REFERENCES

 
Aebi, H. (1984). Catalase in vitro. Methods Enzymol. 105, 121–126.[Web of Science][Medline]

Asada, K. (1999). The water-water cycle in chloroplasts: Scavenging of active oxygens and dissipation of excess photons. Annu. Rev. Plant Physiol. Plant Mol. Biol. 50, 601–639.[CrossRef][Web of Science]

Assmann, S.M. (1993). Signal transduction in guard cells. Annu. Rev. Cell Biol. 9, 345–375.[CrossRef][Web of Science][Medline]

Assmann, S.M., and Wang, X.Q. (2001). From milliseconds to millions of years: Guard cells and environmental responses. Curr. Opin. Plant Biol. 4, 421–428.[CrossRef][Web of Science][Medline]

Blatt, M.R. (2000). Cellular signaling and volume control in stomatal movements in plants. Annu. Rev. Cell Dev. Biol. 16, 221–241.[CrossRef][Web of Science][Medline]

Blatt, M.R., and Armstrong, F. (1993). K⁺ channels of stomatal guard cells: Abscisic-acid-evoked control of the outward rectifier mediated by cytoplasmic pH. Planta 191, 330–341.[Web of Science]

Bowler, C., Vanmontagu, M., and Inze, D. (1992). Superoxide dismutase and stress tolerance. Annu. Rev. Plant Physiol. Plant Mol. Biol. 43, 83–116.[CrossRef][Web of Science]

Bradford, M.M. (1976). A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 72, 248–254.[CrossRef][Web of Science][Medline]

Chen, Z., Young, T.E., Ling, J., Chang, S.-C., and Gallie, D.R. (2003). Increasing vitamin C content of plants through enhanced ascorbate recycling. Proc. Natl. Acad. Sci. USA 100, 3525–3530.[Abstract/Free Full Text]

Creissen, G., Firmin, J., Fryer, M., Kular, B., Leyland, N., Reynolds, H., Pastori, G., Wellburn, F., Baker, N., Wellburn, A., and Mullineaux, P. (1999). Elevated glutathione biosynthetic capacity in the chloroplasts of transgenic tobacco plants paradoxically causes increased oxidative stress. Plant Cell 11, 1277–1292.[Abstract/Free Full Text]

de Pinto, M.C., Tommasi, F., and De Gara, L. (2000). Enzymes of the ascorbate biosynthesis and ascorbate-glutathione cycle in cultured cells of tobacco Bright Yellow 2. Plant Physiol. Biochem. 38, 541–550.[CrossRef][Web of Science]

Erdei, L., Szegeletes, Z.S., Barabas, K.N., Pestenacz, A., Fulop, K., Kalmar, L., Kovacs, A., Toth, B., and Der, A. (1998). Environmental stress and the biological clock in plants: Changes of rhythmic behaviour of carbohydrates, antioxidant enzymes and stomatal resistance by salinity. J. Plant Physiol. 152, 265–271.

Eskling, M., Arvidsson, P., and Akerlund, H.E. (1997). The xanthophyll cycle, its regulation and components. Physiol. Plant 100, 806–816.[CrossRef]

Esterbauer, H., Grill, D., and Welt, R. (1990). Der jahreszeitliche Rhytmus des Acorbinsauresystems in Nadeln von Picea abies. Z. Pflanzenphysiol. 98, 393–402.

Foyer, C.H., Rowell, J., and Walker, D. (1983). Measurements of the ascorbate content of spinach leaf protoplasts and chloroplasts during illumination. Planta 157, 239–244.

Foyer, C.H., Souriau, N., Perret, S., Lelandais, M., Kunert, K.J., Pruvost, C., and Jouanin, L. (1995). Overexpression of glutathione reductase but not glutathione synthetase leads to increases in antioxidant capacity and resistance to photoinhibition in poplar trees. Plant Physiol. 109, 1047–1057.[Abstract]

Gainnopolitis, C.N., and Pies, S.K. (1977). Superoxide dismutases. I. Occurrence in higher plants. Plant Physiol. 59, 309–314.[Abstract/Free Full Text]

Gómez-Cadenas, A., Tadeo, F.R., Talon, M., and Eduardo Primo-Millo, E. (1996). Leaf abscission induced by ethylene in water-stressed intact seedlings of Cleopatra mandarin requires previous abscisic acid accumulation in roots. Plant Physiol. 112, 401–408.[Abstract]

Grabov, A., and Blatt, M.R. (1998). Membrane voltage initiates Ca2+ waves and potentiates Ca2+ increases with abscisic acid in stomatal guard cells. Proc. Natl. Acad. Sci. USA 95, 4778–4783.[Abstract/Free Full Text]

Guan, L.M., Zhao, J., and Scandalios, J.G. (2000). Cis-elements and trans-factors that regulate expression of the maize Cat1 antioxidant gene in response to ABA and osmotic stress: H₂O₂ is the likely intermediary signaling molecule for the response. Plant J. 22, 87–95.[CrossRef][Web of Science][Medline]

Hamilton, D.W., Hills, A., Kohler, B., and Blatt, M.R. (2000). Ca2+ channels at the plasma membrane of stomatal guard cells are activated by hyperpolarization and abscisic acid. Proc. Natl. Acad. Sci. USA 97, 4967–4972.[Abstract/Free Full Text]

Havaux, M., Bonfils, J.P., Lutz, C., and Niyogi, K.K. (2000). Photodamage of the photosynthetic apparatus and its dependence on the leaf developmental stage in the npq1 Arabidopsis mutant deficient in the xanthophyll cycle enzyme violaxanthin de-epoxidase. Plant Physiol. 124, 273–284.[Abstract/Free Full Text]

Horemans, N., Foyer, C.H., and Asard, H. (2000). Transport and action of ascorbate at the plant plasma membrane. Trends Plant Sci. 5, 263–267.[CrossRef][Web of Science][Medline]

Hossain, M.A., and Asada, K. (1984). Purification of dehydroascorbate reductase from spinach and its characterization as a thiol enzyme. Plant Cell Physiol. 25, 85–92.[Abstract/Free Full Text]

Ishikawa, H., Aizawa, H., Kishira, H., Ogawa, T., and Sakata, M. (1983). Light-induced changes of membrane potential in guard cells of Vicia faba. Plant Cell Physiol. 24, 769–772.[Abstract/Free Full Text]

Kohler, B., and Blatt, M.R. (2002). Protein phosphorylation activates the guard cell Ca2+ channel and is a prerequisite for gating by abscisic acid. Plant J. 32, 185–194.[CrossRef][Web of Science][Medline]

Kohler, B., Hills, A., and Blatt, M.R. (2003). Control of guard cell ion channels by hydrogen peroxide and abscisic acid indicates their action through alternate signaling pathways. Plant Physiol. 131, 385–388.[Free Full Text]

Kruse, T., Tallman, G., and Zeiger, E. (1989). Isolation of guard cell protoplasts from mechanically prepared epidermis of vicia faba leaves. Plant Physiol. 90, 1382–1386.[Abstract/Free Full Text]

Lee, S., Choi, H., Suh, S., Doo, I.S., Oh, K.Y., Choi, E.J., Schroeder Taylor, A.T., Low, P.S., and Lee, Y. (1999). Oligogalacturonic acid and chitosan reduce stomatal aperture by inducing the evolution of reactive oxygen species from guard cells of tomato and Commelina communis. Plant Physiol. 121, 147–152.[Abstract/Free Full Text]

Leung, J., and Giraudat, J. (1998). Abscisic acid signal transduction. Annu. Rev. Plant Physiol. Plant Mol. Biol. 49, 199–222.[CrossRef][Web of Science]

MacRobbie, E.A. (2000). ABA activates multiple Ca(2+) fluxes in stomatal guard cells, triggering vacuolar K(+)(Rb(+)) release. Proc. Natl. Acad. Sci. USA 97, 12361–12368.[Abstract/Free Full Text]

McAinsh, M.R., Clayton, H., Mansfield, T.A., and Hetherington, A.M. (1996). Changes in stomatal behavior and guard cell cytosolic free calcium in response to oxidative stress. Plant Physiol. 111, 1031–1042.[Abstract]

Munne-Bosch, S., and Alegre, L. (2002). Plant aging increases oxidative stress in chloroplasts. Planta 214, 608–615.[CrossRef][Web of Science][Medline]

Murata, Y., Pei, Z.M., Mori, I.C., and Schroeder, J. (2001). Abscisic acid activation of plasma membrane Ca(2+) channels in guard cells requires cytosolic NAD(P)H and is differentially disrupted upstream and downstream of reactive oxygen species production in abi1-1 and abi2-1 protein phosphatase 2C mutants. Plant Cell 13, 2513–2523.[Abstract/Free Full Text]

Pastori, G.M., and Foyer, C.H. (2002). Common components, networks, and pathways of cross-tolerance to stress. The central role of "redox" and abscisic acid-mediated controls. Plant Physiol. 129, 460–468.[Free Full Text]

Pei, Z.M., Kuchitsu, K., Ward, J.M., Schwarz, M., and Schroeder, J.I. (1997). Differential abscisic acid regulation of guard cell slow anion channels in Arabidopsis wild-type and abi1 and abi2 mutants. Plant Cell 9, 409–423.[Abstract]

Pei, Z.M., Murata, Y., Benning, G., Thomine, S., Klusener, B., Allen, G.J., Grill, E., and Schroeder, J.I. (2000). Calcium channels activated by hydrogen peroxide mediate abscisic acid signaling in guard cells. Nature 406, 731–734.[CrossRef][Medline]

Pignocchi, C., Fletcher, J.M., Wilkinson, J.E., Barnes, J.D., and Foyer, C.H. (2003). Function of ascorbate oxidase in tobacco. Plant Physiol. 132, 1631–1641.[Abstract/Free Full Text]

Polidoros, A.N., and Scandalios, J.G. (1998). Circadian expression of the maize Cat3 catalase gene is highly conserved among diverse maize genotypes with structurally different promoters. Genetics 149, 405–415.[Abstract/Free Full Text]

Price, A.H., Taylor, A., Ripley, S.J., Griffiths, A., Trewavas, A.J., and Knight, M.R. (1994). Oxidative signals in tobacco increase cytosolic calcium. Plant Cell 6, 1301–1310.[Abstract]

Redinbaugh, M.G., Sabre, M., and Scandalios, J.G. (1990). Expression of the maize Cat3 catalase gene is under the influence of a circadian rhythm. Proc. Natl. Acad. Sci. USA 87, 6853–6857.[Abstract/Free Full Text]

Scandalios, J.G. (1993). Oxygen stress and superoxide dismutases. Plant Physiol. 101, 7–12.[Web of Science][Medline]

Schroeder, J.I., Allen, G.J., Hugouvieux, V., Kwak, J.M., and Waner, D. (2001a). Guard cell signal transduction. Annu. Rev. Plant Physiol. Plant Mol. Biol. 52, 627–658.[CrossRef][Web of Science][Medline]

Schroeder, J.I., Kwak, J.M., and Allen, G.J. (2001b). Guard cell abscisic acid signaling and engineering drought hardiness in plants. Nature 410, 327–330.[CrossRef][Medline]

Schupp, R., and Rennenberg, H. (1988). Diurnal changes in the glutathione content of spruce needles (Picea abies L.). Plant Sci. 57, 113–117.[CrossRef]

Shimaoka, T., Yokota, A., and Miyake, C. (2000). Purification and characterization of chloroplast dehydroascorbate reductase from spinach leaves. Plant Cell Physiol. 41, 1110–1118.[Abstract/Free Full Text]

Smirnoff, N. (2000). Ascorbic acid: Metabolism and functions of a multi-faceted molecule. Curr. Opin. Plant Biol. 3, 229–235.[Web of Science][Medline]

Smirnoff, N., and Pallanca, J.E. (1996). Ascorbate metabolism in relation to oxidative stress. Biochem. Soc. Trans. 24, 472–478.[Web of Science][Medline]

Sutherland, M.W., and Learmonth, B.A. (1997). The tetrazolium dyes MTS and XTT provide new quantitative assay for superoxide and superoxide dismutase. Free Radic. Res. 27, 283–289.[Web of Science][Medline]

Tamaoki, M., Mukai, F., Asai, N., Nakajima, N., Kubo, A., Aono, M., and Saji, H. (2003). Light-controlled expression of a gene encoding L-galactono--lactone dehydrogenase which affects ascorbate pool size in Arabidopsis. Plant Sci. 164, 1111–1117.[CrossRef]

Thiel, G., MacRobbie, E.A., and Blatt, M.R. (1992). Membrane transport in stomatal guard cells: The importance of voltage control. J. Membr. Biol. 126, 1–18.[Web of Science][Medline]

Veljovic-Jovanovic, S.D., Pignocchi, C., Noctor, G., and Foyer, C.H. (2001). Low ascorbic acid in the vtc-1 mutant of Arabidopsis is associated with decreased growth and intracellular redistribution of the antioxidant system. Plant Physiol. 127, 426–435.[Abstract/Free Full Text]

Wildi, B., and Lutz, C. (1996). Antioxidant composition of selected high alpine plant species from different altitudes. Plant Cell Environ. 19, 138–146.

Zeiger, E. (2000). Sensory transduction of blue light in guard cells. Trends Plant Sci. 5, 183–185.[CrossRef][Web of Science][Medline]

Zhang, X., Zhang, L., Dong, F., Gao, J., Galbraith, D.W., and Song, C.P. (2001). Hydrogen peroxide is involved in abscisic acid-induced stomatal closure in Vicia faba. Plant Physiol. 126, 1438–1448.[Abstract/Free Full Text]

Zhong, H.H., and McClung, C.R. (1996). The circadian clock gates expression of two Arabidopsis catalase genes to distinct and opposite circadian phases. Mol. Gen. Genet. 251, 196–203.[Web of Science][Medline]

Zhong, H.H., Painter, J.E., Salome, P.A., Straume, M., and McClung, C.R. (1998). Imbibition, but not release from stratification, sets the circadian clock in Arabidopsis seedlings. Plant Cell 10, 2005–2018.[Abstract/Free Full Text]

This article has been cited by other articles:

				J.-H. Li, Y.-Q. Liu, P. Lu, H.-F. Lin, Y. Bai, X.-C. Wang, and Y.-L. Chen A Signaling Pathway Linking Nitric Oxide Production to Heterotrimeric G Protein and Hydrogen Peroxide Regulates Extracellular Calmodulin Induction of Stomatal Closure in Arabidopsis Plant Physiology, May 1, 2009; 150(1): 114 - 124. [Abstract] [Full Text] [PDF]

				M. Zhu, S. Dai, S. McClung, X. Yan, and S. Chen Functional Differentiation of Brassica napus Guard Cells and Mesophyll Cells Revealed by Comparative Proteomics Mol. Cell. Proteomics, April 1, 2009; 8(4): 752 - 766. [Abstract] [Full Text] [PDF]

				H. Gautier, C. Massot, R. Stevens, S. Serino, and M. Genard Regulation of tomato fruit ascorbate content is more highly dependent on fruit irradiance than leaf irradiance Ann. Bot., February 1, 2009; 103(3): 495 - 504. [Abstract] [Full Text] [PDF]

				Z. Zhao, W. Zhang, B. A. Stanley, and S. M. Assmann Functional Proteomics of Arabidopsis thaliana Guard Cells Uncovers New Stomatal Signaling Pathways PLANT CELL, December 1, 2008; 20(12): 3210 - 3226. [Abstract] [Full Text] [PDF]

				Z. Chen and D. R. Gallie Dehydroascorbate Reductase Affects Non-photochemical Quenching and Photosynthetic Performance J. Biol. Chem., August 1, 2008; 283(31): 21347 - 21361. [Abstract] [Full Text] [PDF]

				A. Nunes-Nesi, R. Sulpice, Y. Gibon, and A. R. Fernie The enigmatic contribution of mitochondrial function in photosynthesis J. Exp. Bot., May 1, 2008; 59(7): 1675 - 1684. [Abstract] [Full Text] [PDF]

				V. Fotopoulos, M. C. De Tullio, J. Barnes, and A. K. Kanellis Altered stomatal dynamics in ascorbate oxidase over-expressing tobacco plants suggest a role for dehydroascorbate signalling J. Exp. Bot., March 1, 2008; 59(4): 729 - 737. [Abstract] [Full Text] [PDF]

				F. Moradi and A. M. Ismail Responses of Photosynthesis, Chlorophyll Fluorescence and ROS-Scavenging Systems to Salt Stress During Seedling and Reproductive Stages in Rice Ann. Bot., June 1, 2007; 99(6): 1161 - 1173. [Abstract] [Full Text] [PDF]

				Z. Chen and D. R. Gallie Dehydroascorbate Reductase Affects Leaf Growth, Development, and Function Plant Physiology, October 1, 2006; 142(2): 775 - 787. [Abstract] [Full Text] [PDF]

				J. M. Cheeseman Hydrogen peroxide concentrations in leaves under natural conditions J. Exp. Bot., July 1, 2006; 57(10): 2435 - 2444. [Abstract] [Full Text] [PDF]

				J. M. Kwak, V. Nguyen, and J. I. Schroeder The Role of Reactive Oxygen Species in Hormonal Responses Plant Physiology, June 1, 2006; 141(2): 323 - 329. [Full Text] [PDF]

				C. Pignocchi, G. Kiddle, I. Hernandez, S. J. Foster, A. Asensi, T. Taybi, J. Barnes, and C. H. Foyer Ascorbate Oxidase-Dependent Changes in the Redox State of the Apoplast Modulate Gene Transcript Accumulation Leading to Modified Hormone Signaling and Orchestration of Defense Processes in Tobacco Plant Physiology, June 1, 2006; 141(2): 423 - 435. [Abstract] [Full Text] [PDF]

				C. Barth, M. De Tullio, and P. L Conklin The role of ascorbic acid in the control of flowering time and the onset of senescence J. Exp. Bot., May 1, 2006; 57(8): 1657 - 1665. [Abstract] [Full Text] [PDF]

				B. A. Wolucka, A. Goossens, and D. Inze Methyl jasmonate stimulates the de novo biosynthesis of vitamin C in plant cell suspensions J. Exp. Bot., September 1, 2005; 56(419): 2527 - 2538. [Abstract] [Full Text] [PDF]

				Z. Chen and D. R. Gallie Increasing Tolerance to Ozone by Elevating Foliar Ascorbic Acid Confers Greater Protection against Ozone Than Increasing Avoidance Plant Physiology, July 1, 2005; 138(3): 1673 - 1689. [Abstract] [Full Text] [PDF]

				A. Nunes-Nesi, F. Carrari, A. Lytovchenko, A. M.O. Smith, M. Ehlers Loureiro, R. G. Ratcliffe, L. J. Sweetlove, and A. R. Fernie Enhanced Photosynthetic Performance and Growth as a Consequence of Decreasing Mitochondrial Malate Dehydrogenase Activity in Transgenic Tomato Plants Plant Physiology, February 1, 2005; 137(2): 611 - 622. [Abstract] [Full Text] [PDF]

				I. C. Mori and J. I. Schroeder Reactive Oxygen Species Activation of Plant Ca2+ Channels. A Signaling Mechanism in Polar Growth, Hormone Transduction, Stress Signaling, and Hypothetically Mechanotransduction Plant Physiology, June 1, 2004; 135(2): 702 - 708. [Full Text] [PDF]

This Article Abstract Full Text (PDF) All Versions of this Article: 16/5/1143    most recent tpc.021584v1 Alert me when this article is cited Alert me if a correction is posted Services Email this article to a friend Similar articles in this journal Similar articles in ISI Web of Science Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Citing Articles Citing Articles via HighWire Citing Articles via CrossRef Citing Articles via ISI Web of Science (72) Citing Articles via Google Scholar Google Scholar Articles by Chen, Z. Articles by Gallie, D. R. Search for Related Content PubMed PubMed Citation Articles by Chen, Z. Articles by Gallie, D. R. Agricola Articles by Chen, Z. Articles by Gallie, D. R.